FACULTY OF SCIENCE
Strategies of Parasitism in Early
Branching Apicomplexa
Habilitation Thesis
Andrea Bardůnek Valigurová
Department of Botany and Zoology
Brno 2019
© Andrea Bardůnek Valigurová, Masaryk University, 2019
Abstrakt
Tato habilitační práce, jejíž součástí je 16 vědeckých publikací, se věnuje hostitelskoparazitickým
interakcím a pohybu u bazálních linií výtrusovců (Apicomplexa), kteří
představují jednu z nejúspěšnějších skupin eukaryotických jednobuněčných
parazitů. Zaměřuji se na bazální výtrusovce, jako jsou protokokcidie, agamokokcidie,
blastogregariny a gregariny omezené na bezobratlé hostitele a kryptosporidie
parazitující obratlovce včetně člověka. Organizace jejich buněčného kortexu a
cytoskeletu koreluje s různými způsoby motility a je výsledkem strukturálních změn
invazivního stadia během proměny na trofozoita u parazitů s různými strategiemi
parazitismu (epicelulární, extracelulární a intracelulární lokalizace), vedoucím k
významným modifikacím nebo dokonce ztrátě mechanismů buněčné motility.
Výsledky této studie odhalily obrovskou rozmanitost v subcelulární organizaci u
bazálních výtrusovců související s jejich různými strategiemi parazitismu, což
potvrzuje důležitost dalšího výzkumu, který by měl poskytnout hlubší pochopení
biologie a evolučních cest u Apicomplexa obecně.
Abstract
This habilitation thesis, supplemented by 16 research papers, deals with hostparasite
interactions and motility in early branching lineages of Apicomplexa, one of
the most successful groups of eukaryotic unicellular parasites. I focus on basal
apicomplexans such as protococcidia, agamococcidia, blastogregarines and
gregarines restricted to invertebrate hosts and cryptosporidia parasitising
vertebrates, including humans. The organisation of their cell cortex and cytoskeleton
correlates with their diverse modes of motility and results from modifications to the
zoite structure during transformation to the trophozoite stage in parasites with
different parasitism strategies (epicellular, extracellular and intracellular
localisation), leading to significant changes or even loss of cell motility mechanisms.
The results of this study revealed enormous diversity in the subcellular organisation
of deep-branching apicomplexans related to their different parasitism strategies,
thereby confirming the importance of further research, which should provide a
deeper understanding of the biology and evolutionary pathways of Apicomplexa in
general.
Acknowledgement
I would like to thank all colleagues and friends who helped with the research
presented in this thesis.
In particular, Timur G. Simdyanov, Gita G. Paskerova and Joseph Schrével
contributed greatly to the papers dealing with marine apicomplexans from
invertebrate hosts. I am also grateful to Isabelle Florent for her help with the
research on cryptosporidia motility. Great thanks go to Naděžda Vaškovicová, who
led me into the mysteries of freeze etching and taught me to read the replicas. My
special thanks go to my students Magdaléna Kováčiková, Andrei Diakin, Janka
Melicherová, Veronika Mazourová, Jana Ilgová and Lenka Tůmová, whose
enthusiasm and friendly attitude gave me the energy to continue my research under
all circumstances. I am deeply grateful to Milan Gelnar for the possibility to perform
my research at the Department of Botany and Zoology of Masaryk University in Brno.
I am also greatly indebted to the Laboratory of Electron Microscopy, Biology Centre
of the Czech Academy of Sciences in České Budějovice, where the majority of my
ultrastructural investigations were performed.
Finally, I would like to thank my family, in particular my parents for their
endless support and my husband Petr, who was very patient with me during the
thesis completion and took great care of our little son.
Content
1 Introduction......................................................................................................................................................7
2 Early branching lineages of Apicomplexa........................................................................................8
2.1 Recent views on apicomplexan taxonomy and evolution..............................................8
2.2 Introduction to model parasites..................................................................................................9
3 Host-parasite interactions in early branching Apicomplexa.............................................12
3.1 Attachment strategies of epicellular parasites.................................................................12
3.2 Attachment strategies of epicellular parasites enveloped by a
parasitophorous sac.........................................................................................................................19
3.3 Niches of intracellular parasites...............................................................................................25
3.4 Feeding strategies.............................................................................................................................27
3.5 Pathogenicity to the host...............................................................................................................31
4 Motility in early branching Apicomplexa......................................................................................36
4.1 Apicomplexan motility and cytoskeleton ............................................................................36
4.2 Motility and cytoskeleton in archigregarines and blastogregarines....................37
4.3 Motility and cytoskeleton in eugregarines..........................................................................40
4.4 Motility and cytoskeleton in protococcidia and agamococcidia.............................45
4.5 Motility and cytoskeleton in cryptosporidia......................................................................46
5 Conclusions ....................................................................................................................................................48
6 References ......................................................................................................................................................49
7 List of publications included in the thesis ....................................................................................58
7.1 Valigurová A. (2012). Sophisticated adaptations of Gregarina cuneata (Apicomplexa)
feeding stages for epicellular parasitism. PLoS One 7(8), e42606.
7.2 Valigurová A., Vaškovicová V., Musilová N., Schrével J. (2013). The enigma of
eugregarine epicytic folds: where gliding motility originates? Frontiers in Zoology
10, 57.
7.3 Melicherová J., Ilgová J., Kváč M., Sak B., Koudela B., Valigurová A. (2014). Life cycle
of Cryptosporidium muris in two rodents with different responses to parasitization.
Parasitology 141(2), 287-303.
7.4 Valigurová A., Paskerova G.G., Diakin A., Kováčiková M., Simdyanov T.G. (2015).
Protococcidian Eleutheroschizon duboscqi, an unusual apicomplexan interconnecting
gregarines and cryptosporidia. PLoS One 10(4), e0125063.
7.5 Melicherová J., Mazourová V., Valigurová A. (2016). In vitro excystation of
Cryptosporidium muris oocysts and viability of released sporozoites in different
incubation media. Parasitology Research 115(3), 1113-1121.
7.6 Diakin A., Paskerova G.G., Simdyanov T.G., Aleoshin V.V., Valigurová A. (2016).
Morphology and molecular phylogeny of coelomic gregarines (Apicomplexa) with
different types of motility: Urospora ovalis and U. travisiae from the polychaete
Travisia forbesii. Protist 167(3), 279-301.
7.7 Schrével J., Valigurová A., Prensier G., Chambouvet A., Florent I., Guillou L. (2016).
Ultrastructure of Selenidium pendula, the type species of archigregarines, and
phylogenetic relations to other marine Apicomplexa. Protist 167(4), 339-368.
7.8 Chambouvet A., Valigurová A., Pinheiro L.M., Richards T.A., Jirků M. (2016).
Nematopsis temporariae (Gregarinasina, Apicomplexa, Alveolata) is an intracellular
infectious agent of tadpole livers. Environmental Microbiology Reports 8(5), 675-
679.
7.9 Diakin A., Wakeman K.C., Valigurová A. (2017). Description of Ganymedes yurii sp. n.
(Ganymedidae), a new gregarine species from the Antarctic amphipod Gondogeneia
sp. (Crustacea). Journal of Eukaryotic Microbiology 64(1), 56-66.
7.10 Kováčiková M., Simdyanov T.G., Diakin A., Valigurová A. (2017). Structures related to
attachment and motility in the marine eugregarine Cephaloidophora cf. communis
(Apicomplexa). European Journal of Protistology 59, 1-13.
7.11 Valigurová A., Vaškovicová N., Diakin A., Paskerova G.G., Simdyanov T.G., Kováčiková
M. (2017). Motility in blastogregarines (Apicomplexa): Native and drug-induced
organisation of Siedleckia nematoides cytoskeletal elements. PLoS One 12(6),
e0179709.
7.12 Melicherová J., Hofmannová L., Valigurová A. (2018). Response of cell lines to actual
and simulated inoculation with Cryptosporidium proliferans. European Journal of
Protistology 62, 101-121.
7.13 Valigurová A., Pecková R., Doležal K., Sak B., Květoňová D., Kváč M., Nurcahyo W.,
Foitová I. (2018). Limitations in the screening of potentially anti-cryptosporidial
agents using laboratory rodents with gastric cryptosporidiosis. Folia Parasitologica
65, 010.
7.14 Kováčiková M., Vaškovicová N., Nebesářová J., Valigurová A. (2018). Effect of
jasplakinolide and cytochalasin D on cortical elements involved in the gliding
motility of the eugregarine Gregarina garnhami (Apicomplexa). European Journal of
Protistology 66, 97-114.
7.15 Simdyanov T.G., Paskerova G.G., Valigurová A., Diakin A., Kováčiková M., Schrével J.,
Guillou L., Dobrovolskij A.A., Aleoshin V.V. (2018). First ultrastructural and
molecular phylogenetic evidence from the blastogregarines, an early branching
lineage of plesiomorphic Apicomplexa. Protist 169(5), 697-726.
7.16 Paskerova G.G., Miroliubova T.S., Diakin A., Kováčiková M., Valigurová A., Guillou L.,
Aleoshin V.V., Simdyanov T.G. (2018). Fine structure and molecular phylogenetic
position of two marine gregarines, Selenidium pygospionis sp. n. and S. pherusae sp.
n., with notes on the phylogeny of Archigregarinida (Apicomplexa). Protist 169(6),
826-852.
8 Other publications related to the habilitation thesis.............................................................60
7
1 Introduction
Apicomplexa consist entirely of parasitic genera occurring in a wide spectrum of
invertebrates and vertebrates, including humans. Their representatives have
evolved unique adaptations for invading and surviving within hosts. It is assumed
that ancestral apicomplexans parasitised marine annelids and that their adaptation
to a parasitic lifestyle and further radiation took place before the era of vertebrates,
spreading first to other marine invertebrates (crustaceans, turbellarians,
echinoderms, etc.), then freshwater and terrestrial invertebrates, and finally
vertebrates. In contrast to apicomplexan etiologic agents of globally significant
human and animal diseases (e.g. malaria, toxoplasmosis, cryptosporidiosis), which
have been well studied, the enormously diversified basal apicomplexan lineages
restricted to invertebrate hosts remain poorly understood, despite being important
from an evolutionary perspective due to their basal phylogenetic position. New
findings on biodiversity and other biological aspects, such as cell motility, invasion
strategies and host-parasite interactions, in representatives of early branching
lineages will facilitate our understanding of evolutionary pathways in Apicomplexa.
From the beginning of my research activity in the Department of Botany and
Zoology, I have directed my studies on basal Apicomplexa to cryptosporidia and
gregarines, which are thought to be deep-branching apicomplexans. Direct
comparison of host-parasite interactions in both parasite groups revealed that they
exhibit specific morphological and developmental similarities, indicating that
cryptosporidia may represent a missing link between the gregarines and coccidia.
This thesis, summarising observations on apicomplexan representatives of diverse
basal lineages published in the enclosed papers, offers a novel perspective on
parasitism strategies in Apicomplexa, and highlights the importance of deeper
investigations into parasites that have not been attributed as having any economic
or medical significance.
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2 Early branching lineages of Apicomplexa
2.1 Recent views on apicomplexan taxonomy and evolution
Apicomplexa Levine, 1970, emend. Adl et al., 2012, which inhabits almost all
known phyla of multicellular organisms, is one of the most monitored groups of
unicellular eukaryotic parasites. Based largely on phenotypic characteristics,
Apicomplexa were traditionally considered to contain four clearly defined groups:
gregarines, coccidia, haemosporidia and piroplasmids. These groups were originally
designed to be utilitarian rather than phylogenetic (Morrison, 2009). Recently,
phylogenetic evidence provided by molecular data has led to this traditional
taxonomic scheme being questioned. Subsequently, numerous works dealing with
phylogeny and systematic arrangements within the Apicomplexa have been
published (e.g. Adl et al., 2012; Cavalier-Smith, 2014; Cavalier-Smith and Chao, 2004;
Perkins, 2000).
Fig. 1. Hypothetical tree of life for the Apicomplexa (taken from Votýpka et al, 2017).
9
Parasite taxonomy in this thesis follows the non-rank classification of eukaryotes
published by Adl et al. (2012), in which the Apicomplexa, along with protalveolates,
dinoflagellates and ciliates, form the higher order group Alveolata. However,
phylogenetic relationships and the evolutionary history of organisms within the
Apicomplexa remain an open question (Simdyanov et al., 2018). It is generally
accepted that ancestral apicomplexans parasitised marine annelids and that their
adaptation to a parasitic lifestyle took place before the vertebrate period (Cox, 1994;
Théodoridès, 1984). Several early-dispersed apicomplexan branches are
hypothesised, e.g. blastogregarines, archigregarines, eugregarines and
neogregarines, agamococcidia, protococcidia and cryptosporidia. These exhibit an
enormous diversity in both dimension and cell architecture, depending on their
surrounding environment and parasitism strategy, and appear to be perfect
examples of coevolution between parasites and their hosts. It is thought most likely
that an archigregarine stem group gave rise to cryptosporidia, and a lineage uniting
(eu)coccidia with piroplasmids (Leander et al., 2006).
2.2 Introduction to model parasites
Gregarines (Conoidasida Levine, 1988, Gregarinasina Dufour, 1828) are deepbranching
apicomplexan parasites of invertebrates. Based on differences in their life
history, gregarines are traditionally classified into three subgroups: archigregarines
(Archigregarinorida Grassé, 1953; prevailing extracellular development with
merogony), eugregarines (Eugregarinorida Léger, 1900; merogony absent,
predominant extracellular development) and neogregarines (Neogregarinorida
Grassé, 1953; merogony present, intracellular or extracellular development) (Adl et
al., 2012; Perkins, 2000). Studies on the following archigregarines and eugregarines
are included in this thesis: Selenidium pendula Giard, 1884 from the intestine of the
marine polychaete Scolelepis squamata (Müller, 1806); S. pygospionis Paskerova,
Miroliubova, Diakin, Kováčiková, Valigurová, Guillou, Aleoshin and Simdyanov, 2018
from the intestine of the marine polychaete Pygospio elegans Claparède, 1863; S.
pherusae Paskerova, Miroliubova, Diakin, Kováčiková, Valigurová, Guillou, Aleoshin
and Simdyanov, 2018 from the intestine of the marine polychaete Pherusa plumosa
(Müller, 1776); Cephaloidophora cf. communis Mawrodiadi, 1908 from the intestine
of the barnacle Balanus balanus Linnaeus, 1758; Ganymedes yurii Diakin, Wakeman
10
and Valigurová, 2016 from the intestine of the marine amphipod Gondogeneia sp.;
Urospora ovalis Dogiel, 1910 and U. travisiae Dogiel, 1910 from the body cavity of the
marine polychaete Travisia forbesii Johnston, 1840; Gregarina cuneata Stein, 1848,
G. polymorpha (Hammerschmidt, 1838) and G. steini Berndt, 1902 from the intestine
of larval mealworm beetle Tenebrio molitor Linnaeus, 1758; G. garnhami Canning,
1956 from the intestine of the desert locust Schistocerca gregaria (Forskål, 1775);
and the first gregarine reported from a vertebrate host, Nematopsis temporariae
Nöller, 1920 from the liver of tadpoles (Diakin et al., 2014, 2016, 2017; Chambouvet
et al., 2016; Kováčiková et al., 2017a, 2017b, 2018; Paskerova et al., 2018; Schrével
et al., 2016; Valigurová, 2012; Valigurová et al., 2013).
Until recently, blastogregarines (Blastogregarinea Chatton and Villeneuve,
1936, emend), apicomplexan parasites of marine polychaetes with uncertain
taxonomic affiliation and comprising four known species, were poorly studied.
Previous studies considered them to be highly modified gregarines, an intermediate
lineage between gregarines and coccidia or even an isolated group of eukaryotes.
Superficially, they resemble gregarines but lack syzygy and gametocyst stages in
their life cycle, and are characterised by permanent multinuclearity and
gametogenesis by means of budding, which distinguishes them considerably from
other apicomplexans (Caullery and Mesnil, 1898). The studied blastogregarines, the
type species Siedleckia nematoides Caullery and Mesnil, 1898 from the intestine of
Scoloplos armiger (Müller, 1776) and Chattonaria mesnili (Chatton and Dehorne,
1929) from the intestine of Orbinia latreillii (Audouin and Milne Edwards, 1833),
possess both gregarine (folded or smooth pellicle, persisting mucron and apical
complex during the larger part of lifecycle, oocysts with free sporozoites) and
coccidian (gametes associated with two kinetosomes, absence of gametocyst,
pronounced difference in size between male and female gametes, microgametes with
two flagella) features. Phylogenetic analysis indicates that blastogregarines are an
independent, early diverging lineage of Apicomplexa. The traits shared with
archigregarines, i.e. distinctive tegument structure and myzocytotic feeding via a
well-developed apical complex, probably represent the ancestral states of the
corresponding cell structures for Apicomplexa (Simdyanov et al., 2018; Valigurová
et al., 2017).
Cryptosporidia (Conoidasida Levine, 1988, Cryptosporidium Tyzzer, 1907)
11
comprise causative agents of zoonotic diseases of the gastrointestinal and
respiratory tract of vertebrates, including humans. The genus Cryptosporidium was
conceived for a peculiar parasite found in the stomach of laboratory mice, whose
endogenous stages develop attached to the host epithelium with development
including both asexual and sexual developmental stages (Tyzzer, 1907). Based on
Perkins (2000), the family Cryptosporidiidae Léger, 1911, with a single genus
Cryptosporidium, was previously placed within the class Conoidasida Levine, 1988
(subclass Coccidiasina Leuckart, 1879; order Eucoccidiorida Léger and Duboscq,
1910; suborder Eimeriorina Léger, 1911). The first doubts regarding the taxonomic
position of cryptosporidia came following the publication of a work reporting crossreactivity
of an anti-Cryptosporidium monoclonal antibody with a monocystid
gregarine (Bull et al., 1998), leading to speculation that cryptosporidia represent a
missing link between the gregarines and coccidia. Later studies revealed that
Cryptosporidium has closer phylogenetic affinity to gregarines than coccidia sensu
stricto and that it represents an early emerging branch of Apicomplexa (Barta and
Thompson, 2006; Carreno et al., 1999; Leander et al., 2006). Recent systematic work
has placed cryptosporidia within Conoidasida Levine, 1988, comprising Coccidia
Leuckart, 1879, Gregarinasina Dufour, 1828 and the genus Cryptosporidium (Adl et
al., 2012). Publications included in this thesis deal with the gastric species
Cryptosporidium proliferans Kváč, Havrdová, Hlásková, Daňková, Kanděra, Ježková,
Vítovec, Sak, Ortega, Xiao, Modrý, Chelladurai, Prantlová and McEvoy, 2016
(previously known as strain TS03 of C. muris Tyzzer, 1907) originating from a
naturally infected East African mole rat Tachyoryctes splendens (Rüppell, 1835) and
passaged in laboratory BALB/c mice and southern multimammate mice Mastomys
coucha (Smith, 1834) (Melicherová et al., 2014, 2016, 2018; Valigurová et al., 2018).
One of the most poorly explored basal apicomplexans are the protococcidia
(Incertae sedis Apicomplexa: Protococcidiorida Kheisin, 1956), with uncertain
subdivisions and comprising eight genera: Angeiocystis, Coelotropha, Grellia,
Eleutheroschizon, Mackinnonia, Myriosporides, Myriospora and Sawayella. These are
expected to lack merogony, with gamogony and sporogony occurring extracellularly
(Adl et al., 2012; Perkins, 2000). The genus Eleutheroschizon Brasil, 1906 contains
two known intestinal species from marine polychaetes, the type species E. duboscqi
Brasil, 1906 from S. armiger (investigated in this study) and E. murmanicum
12
Awerinzew, 1908 from Ophelia limacina (Rathke, 1843). In these species,
endogenous stages exhibit epicellular development and share the features of
coccidia and gregarines; i.e. cell polarity in advanced developmental stages
(trophozoites, gamonts) characteristic for gregarines, localisation within the
parasitophorous sac similar to that of cryptosporidia and a life cycle lacking
merogony and syzygy stages, despite showing a coccidian type of gametogony
(Brasil, 1906; Chatton and Villeneuve, 1936; Valigurová et al., 2015). Gamonts detach
from the host intestinal epithelium and disperse into the environment, where
gametogenesis and sporogenesis take place.
Agamococcidia (Incertae sedis Apicomplexa: Agamococcidiorida Levine, 1979)
are another small and poorly investigated group of Apicomplexa (comprising two
genera, Gemmocystis and Rhytidocystis) from marine polychaetes, each displaying an
unusual life cycle lacking gametogony and merogony (Adl et al., 2012). Included here
is an unpublished study dealing with a new species of Rhytidocystis Henneguy, 1907,
an intracellular parasite from the intestine of the marine polychaete T. forbesii
(Diakin and Valigurová, 2014).
3 Host-parasite interactions in early branching
Apicomplexa
3.1 Attachment strategies of epicellular parasites
Apicomplexans exhibit highly specific adaptations for invading and surviving
within their hosts. These have evolved under distinct evolutionary pressures,
resulting in diverse host-parasite interactions. Host cell invasion by apicomplexan
zoites is a rapid process depending on a sequence of tightly controlled events
(Dubremetz et al., 1998). First, they have to orient on the surface of the target host
tissue/cell and find an appropriate site for invading. The next steps include attaching
and/or penetrating the host tissue/cell, often accompanied by a parasite-induced
modification of the target cells. These critical steps are facilitated by differentiation
of the parasite into a highly specialised, motile invasive stage, the zoite, equipped
with a set of sophisticated organelles and a cytoskeleton dedicated to reaching and
invading the host cell. The zoite exhibits high cell polarity by having an anterior pole
equipped with a unique invasion apparatus, the apical complex, consisting of
13
specialised secretory organelles (rhoptries, micronemes), polar ring(s) and a conoid.
This apparatus, traditionally used as the best-defined feature for Apicomplexa, can
also be found in other Myzozoa, a group comprising apicomplexans, dinoflagellates
and several lineages of free-living predatory or parasitic flagellates that employ a
myzocytosis-based mode of feeding, i.e. the process by which the predator/parasite
attaches to the prey/target cell and sucks out or feeds permanently on its cytoplasm
via specialised organelles (Cavalier-Smith and Chao, 2004). It is probable that the
evolution of Apicomplexa progressed from myzocytotic predation to myzocytotic
extracellular parasitism, as exhibited by some gregarines and cryptosporidia, and
finally to intracellular parasitism, typical for coccidia. The Apicomplexa demonstrate
two main determinative evolutionary trends: i) the origination of epicellular
parasitism, observed mostly in gregarines, with subsequent modification of the
attachment apparatus and motility mode in the vegetative stage (trophozoite); and
ii) origination of intracellular parasitism in typical coccidia and Aconoidasida,
accompanied by a rejection of trophozoite polarity and motility (Valigurová et al.,
2015).
The attachment apparatus of deep-branching apicomplexans evolved at the
apical end of the sporozoite (the first, invasive stage in the apicomplexan life cycle)
and demonstrates an enormous diversity in architecture. Gregarines and
blastogregarines, for example, exhibit diverse strategies for attachment to the host
tissue, including (i) intratissular or intracellular localisation with or without a
reduced area of attachment in neogregarines; (ii) a mucron in blastogregarines,
archigregarines, monocystid eugregarines and some neogregarines; (iii) a more
advanced mucron-like structure in aseptate eugregarines, losing the apical complex
and strengthening its attachment function, (iv) a simple (Fig. 2) or (v) complicated
epimerite, equipped with various structures (e.g. hooks or spines, digitations, hairs)
in eugregarines (Fig. 3); and (vi) a sucker-like protomerite or modified protomerite
with rhizoids in septate eugregarines (Fig. 4) (Cook et al., 2001; Desportes and
Schrével, 2013; Kováčiková et al., 2017b; Lucarotti, 2000; MacMillan, 1973b;
Paskerova et al., 2018; Schrével et al., 2016; Schrével and Vivier, 1966; Simdyanov
and Kuvardina, 2007; Simdyanov et al., 2018; Tronchin and Schrével, 1977;
Valigurová, 2012; Valigurová et al., 2007, 2009; Valigurová and Koudela, 2005, 2006,
2008).
14
Fig. 2. Schematic diagram of the differing forms of simple epimerite (taken and
modified from Valigurová, 2001). A. Rudimentary. B. Conical papilla. C. Spherical. D.
Button-shaped E. Ovoid. F. Lance-shaped. G. Reverse heart-shaped. e – epimerite, p –
protomerite, d – deutomerite.
Fig. 3. Schematic diagram of the differing forms of complicated epimerite (taken and
modified from Valigurová, 2001). A-C. Various forms with hooks. D. Segmented disc
with a pivot. E. Sucker-shaped with branched papillae. F. Papilla with peripheral
teeth, basally bound by a pillow. G. Spherical with club-shaped protrusions. H.
Spherical with peripheral hooks and a central spike.
Fig. 4. Schematic diagram of modified protomerite. A. Sucker-like protomerite. B.
Protomerite with rhizoids (taken and modified from Valigurová, 2001).
Blastogregarines embed their mucron into the enterocyte’s brush border,
which bears the microcilia and microvilli. The mucronal complex of S. nematoides is
15
equipped with apical organelles, the conoid, internal and external polar rings,
numerous rhoptries and putative micronemes, and a mucronal vacuole (Simdyanov
et al., 2018; Valigurová et al., 2017). The mucron is covered with a trimembrane
pellicle typical of Apicomplexa, except for the region against the conoid where the
mucronal vacuole has a wide inlet opening, considered a cytostome-cytopharyngeal
complex performing myzocytosis (Fig. 5). The cytostome opens into a tiny gap
between the parasite and host cell plasma membranes with the appearance of a
septate cell junction. No significant modifications of the host cell are present, except
for increased electron density of the host plasma membrane directly facing the
cytostome, which bears uniformly spaced dense structures on its external surface,
most likely a perforated or modified host cell coat. The strongly modified mucron of
C. mesnili lacks the conoid and rhoptries, and anchors to the host cell via peripheral
bulges formed by large alveoli between the cytomembranes of the inner membrane
complex (IMC). A gap of varying width is present between the C. mesnili pellicle and
the host cell membrane, with no evidence of a septate cell junction.
Fig. 5. Schematic diagram of the mucronal complex in blastogregarines (taken and
modified from Simdyanov et al., 2018). A-B. Siedleckia nematoides, C-D. Chattonaria
mesnili: the schemes of the longitudinal and cross sections of the mucron and pellicle,
respectively. alv – alveoles between the cytomembranes of the inner membrane
complex, co – conoid, cs – cytostome, hm – host cell plasma membrane, imc – inner
membrane complex, mt – microtubules, mv – mucronal vacuole, pm – parasite
plasma membrane, pr – polar ring (giving rise to the subpellicular microtubules in
Siedleckia); rh – rhoptries, sj – septate cell junction between the parasite and host
cell, smt – subpellicular microtubules.
16
The apical phagotrophy in alveolate free-living predators with open conoid
and rhoptries may be at the origin of the anchoring apparatus of archigregarines. The
mucron of S. pendula appears as a regular mammiliform area and a series of
shortened microvilli are visible at the periphery of the mucron when embedded in
epithelial cells. A tropism for host cells rich in granules can be seen in intestinal
epithelium parasitised by S. pendula. After trophozoite detachment, the trace left by
the mucron in the host tissue is regular, sometimes with a small hole in a subcentral
position (Schrével et al., 2016). The mucronal complex comprises the conoid,
mucronal vacuoles, numerous rhoptries and micronemes (Fig. 6).
Fig. 6. Schematic diagram of the
anterior region of the Selenidium
trophozoite (taken and modified from
Schrével, 1968). Mucronal vacuoles
form at the level of the conoid and,
after detachment, these migrate
posteriorly. Numerous pinocytotic
vesicles are present at the periphery
of the mucronal vacuoles. c – conoid,
cmt – microfibrillar curtain that
covers the microtubular network
forming the conoid, db – dense body,
hc – host cell, imc – inner membrane
complex, m – mitochondrion, mv –
mucronal vacuole, o – opening in the
pellicle, p – pellicle, pm – parasite
plasma membrane, pv – pinocytotic
vesicles, r – rhoptry, v – empty
vesicles, vdb – vesicles containing
dense bodies, vmv – vesicles in the
mucronal vacuole, vmw – vesicles
with multi-membrane whorls.
As S. pendula is the archigregarine type species and an early branching
apicomplexan, its conoid represents a good model for studying the transition
between Apicomplexa with a closed conoid and free-living alveolate ancestors with
an open conoid, as found in the early branching dinoflagellates (Okamoto and
Keeling, 2014; Schrével et al., 2016). The conoids of S. pendula, S. hollandei and S.
orientale appear similar to those in eugregarine sporozoites and Toxoplasma gondii,
but lack the apical polar ring (Desportes, 1969; Sheffield et al., 1971; Schrével, 1968;
Schrével et al., 2016; Simdyanov and Kuvardina, 2007). In contrast, the polar ring,
17
adjacently located to the IMC at the apical conoid end and giving rise to subpellicular
microtubules, has been documented in S. pherusae (Paskerova et al., 2018).
The eugregarine sporozoite develops into a large extracellular vegetative stage,
the trophozoite, equipped with an apical region specialised for attachment to the
host cell (Valigurová, 2012). The epimerites of eugregarines from herbivorous hosts
are usually simple and button-shaped, while they are complicated and equipped with
strong hooks, spines or numerous filaments in carnivorous hosts (Schrével and
Philippe, 1993). The attachment process of septate eugregarines (Fig. 7) from the
intestines of terrestrial hosts has been well studied. After sporozoite attachment to
the host cell, an epimeritic bud arises over the opened apical region and gradually
develops into an epimerite overlain by an electron-translucent cortical vesicle
(Desportes, 1969; Tronchin and Schrével, 1977; Valigurová, 2012; Valigurová et al.,
2007, 2009; Valigurová and Koudela, 2005, 2008). Most likely, this develops from
fused flat vesicles (distributed at the epimerite periphery and originating from the
parasite’s endoplasmic reticulum) that turn into a single large vesicle packed with
microfilaments. Formation of the cortical vesicle appears to be related to the
presence of a flask-shaped organelle in the youngest stages (suggestive of a rhoptry),
which empties its contents during the transformation of the sporozoite into a
trophozoite (Sheffield et al., 1971; Tronchin and Schrével, 1977; Valigurová et al.,
2007; Valigurová and Koudela, 2005). Interestingly, the cortical vesicle in
Didymophyes gigantea has been interpreted as a periparasitic space between the
host and parasite, functioning as a parasitophorous vacuole (Hildebrand, 1976). The
cortical vesicle does indeed resemble the internal space of the parasitophorous
vacuole due to its translucent appearance with traces of an opaque or filamentous
material. In addition, fine tubules passing through the cortical vesicle and attaching
to the epimerite-host cell interface have been documented in G. garnhami
(Valigurová and Koudela, 2008). Hence, it could be speculated that the cortical
vesicle is in fact an incomplete parasitophorous envelope restricted to the embedded
apical region (epimerite) of gregarines (Valigurová et al., 2015). This idea is
supported by the fact that the cortical vesicle appears subsiding and irregular during
the course of epimerite gradual regression before trophozoite detachment from the
host tissue (Valigurová et al., 2009). The epimerite is only covered by a plasma
membrane, while the rest of the eugregarine is covered by a pellicle consisting of the
18
plasma membrane underlain by two cortical cytomembranes of the IMC. Along with
the formation of the epimerite, a membrane fusion site (i.e. osmiophilic ring) forms
connecting the host plasma membrane, the membrane-like structure beneath the
cortical vesicle and the epimerite plasma membrane. The IMC extends only as far as
the top of the protomerite and ends at the membrane fusion site. The trophozoite
remains epicellular and the only part in close contact with the host cell is the
epimerite plasma membrane. The growing epimerite gradually implants into the
host cell, causing a deep invagination of the host cell plasma membrane, but does not
penetrate it (Valigurová 2012; Valigurová et al., 2007, 2009). The trilaminate
interface between the host cell and the epimerite is formed by the epimerite and host
plasma membranes, separated by a dense layer. The accumulation of actin at the base
of the epimerite suggests that the osmiophilic ring is contractile (Ghazali et al., 1989;
Schrével and Vivier, 1966; Tronchin and Schrével, 1977). Phalloidin labelling has
confirmed the presence of filamentous form of actin (F-actin) at the position of the
osmiophilic ring (Valigurová, 2012; Valigurová et al., 2009). A continuous or
discontinuous fibrillar septum separates the epimerite from the protomerite in a few
species, supported by a ring rich in α-tubulin in one documented case (Kováčiková
et al., 2017b; Valigurová and Koudela, 2008).
Two contradictory hypotheses describe trophozoite detachment from host tissue
at the end of development. One describes detachment via epimerite retraction, selfregulated
by the vegetative stage (Lucarotti, 2000; Valigurová and Koudela, 2008;
Valigurová et al., 2009), while the other is based on gradual epimerite constriction
facilitated by the contractility of the osmiophilic ring surrounding the base of the
epimerite, acting as a sphincter during the separation of the epimerite from the rest
of the gregarine (Desportes and Schrével, 2013; Ghazali et al., 1989; Ghazali and
Schrével, 1993; Schrével and Philippe, 1993; Tronchin et al., 1986; Tronchin and
Schrével, 1977). Insofar as the vegetative phase of the eugregarine life cycle usually
lasts longer than four days (Harry, 1970; Valigurová and Koudela, 2005),
trophozoites must be adapted either to keeping the host cell alive during their
development or for eventual reattachment to a younger cell in better physiological
condition after abandoning the senescing cell. The latter could be facilitated by a
retractable epimerite and progressive gliding motility in eugregarines (Lucarotti,
2000; Valigurová, 2012). This is especially true in gregarines with a permanent
19
epimerite, with its architecture reminiscent of a modified protomerite and persisting
in gamonts (Kováčiková et al., 2017b). In this case, the feeding stages of G. cuneata,
which exhibit spectacular adaptations to epicellular parasitism, provide further
support for the epimerite retraction hypothesis. Though the parasite becomes firmly
anchored to the brush border of the epithelium via numerous epimerite digitations
deeply invaginating the host plasma membrane, trophozoites are able to detach and
retain an intact epimerite (Valigurová, 2012). After epimerite retraction in mature
trophozoites, early syzygies (syzygy = pairing up of gamonts before the formation of
a gametocyst) of G. cuneata are often found to be attached to the host tissue via a
modified protomerite of the primite (the anterior, female member of the syzygy), the
apex of which exhibits an undulating pattern. Accumulation of actin in Gregarina
protomerite indicates that attachment via a modified protomerite could be
facilitated by increased flexibility in this region (Heintzelman, 2004; Valigurová,
2012; Valigurová et al., 2009, 2013). Likewise, actinocephalid gamonts lose their
simple globular epimerite and attach by a sucker-like protomerite. The interspace
between the epicytic folds of the attached protomerite and the intestinal epithelium
is packed with host microvilli embedded in a dense adhesive material, likely
produced by exocytic vesicles in the protomerite apical cytoplasm (Cook et al., 2001;
Valigurová, 2012).
3.2 Attachment strategies of epicellular parasites enveloped by a
parasitophorous sac
The peculiar niche of cryptosporidia at the brush border of the gastrointestinal
epithelium has been the subject of extensive debate for decades. While some refer to
cryptosporidia as intracellular though extracytoplasmic parasites, others prefer the
term epicellular to describe the unique localisation of cryptosporidian
developmental stages within a parasitophorous sac of host cell origin (Barta and
Thompson, 2006; Borowski et al., 2010; Cavalier-Smith, 2014; Clode et al., 2015;
Ryan et al., 2016; Valigurová et al., 2008). The parasitophorous sac, introduced for
the first time by Paperna and Vilenkin (1996), is the preferable term for the hostderived
structure enveloping cryptosporidian endogenous stages. The
parasitophorous sac is an epicellular niche enveloping the entire parasite composed
of two continuous host plasma membranes on the outer and inner sides and
20
enclosing a thin layer of host cell cytoplasm (Valigurová et al., 2008). In contrast, the
term ‘parasitophorous vacuole’ previously applied for Cryptosporidium is misleading
as it refers solely to a vacuolar space bordered by a membrane (Scholtyseck, 1979).
Long-term observations of several cryptosporidian species developing in vivo and in
vitro support the term epicellular to reflect their localisation within host tissue as the
invasive stages do not penetrate under the host cell plasma membrane and do not
come into close contact with the host cytoplasm (Jirků et al., 2008; Melicherová et al.,
2014, 2018; Valigurová et al., 2007, 2008; Valigurová, personal observation). The
invading zoite induces modulation of the host plasma membrane, which loses its
microvillous nature and forms a circular membrane fold, tightly encircling the apical
end of the zoite and gradually rising up along the zoite (Fig. 7). This membrane
protrusion and encapsulation of the parasite is actin-dependent (Nelson et al., 2006).
Parasite-induced reorganisation and accumulation of host cell actin at the
attachment site leads to the formation of a dense band supported by the actin plaque,
which is intimately involved in parasite anchoring (Forney et al., 1999; O'Hara et al.,
2008). This dense band in the host cytoplasm, located just beneath the parasite
attachment site and separating the unmodified and modified parts of the host cell,
consists of electron-dense microfibrils interwoven perpendicularly with an adjacent
filamentous network of polymerised actin (Beyer et al., 2000; Landsberg and
Paperna, 1986; O'Hara et al., 2008; Valigurová et al., 2008, 2015). Observations on
tubular structures in oblique sections of the dense band have led to the hypothesis
that it serves as a barrier against deeper penetration of the parasite into the host cell
cytoplasm (Beyer et al., 2000). Vacuolation of the parasite apical cytoplasm during
invasion appears to be the first sign of the incoming anterior vacuole, which is
considered a precursor of the feeder organelle. The anterior vacuole membrane folds
to form the lamellae of the feeder organelle. Later, both the parasite plasma
membrane and the anterior vacuole membrane fuse with the inner membrane of the
parasitophorous sac to form the Y-shaped membrane junction (= annular ring). The
parasite remains attached to the host cell surface, only enveloped by the host
membrane folds. Pore-like structures are occasionally seen on the surface of the sac
(Valigurová et al., 2007, 2008). The parasitophorous sac serves as a protective coat
against the hostile conditions of the vertebrate gastrointestinal tract and reinforces
attachment of the parasite to host epithelium. Simulated parasitisation of cell lines
21
with polystyrene microspheres has revealed that, in addition to intact or even empty
cryptosporidian oocysts, polybeads coated with a parasite antigen ‘cocktail’
(obtained from oocysts with sporozoites) are able to induce the same plasma
membrane modification in affected cultured cells. Contact with oocysts or parasite
antigen-coated polybeads induces actin reorganisation in cultured cells, resulting in
the formation of an F-actin network surrounding the foreign object. Encapsulation
of cryptosporidian oocysts by cultured cells is induced by parasite antigens and
occurs independently of any active invasion by motile stages (Melicherová et al.,
2018). It is likely that oocyst adherence to host cell is aided by molecules containing
N-acetyl-galactosamine on their surface. Lectin-enhanced attachment of oocysts to
host tissue/cell lines increases infection efficiency by bringing sporozoites into close
proximity with host cells (Stein et al., 2006). In addition to their adhesive function,
oocyst surface antigens may also represent a passive means of host cell manipulation
at the oocyst stage (Yao et al., 2007).
Cryptosporidia share striking similarities with epicellular gregarines, especially
in their attachment strategy, i.e. the eugregarine epimerite resembles the feeder
organelle and the entire attachment site in cryptosporidia (Fig. 7). The dense line at
the base of the feeder organelle, serving as a primary barrier between the parasite
and host cell cytoplasm, is homologous to the interface between the epimerite and
the host cell (Jirků et al., 2008; Melicherová et al., 2014, 2018; Valigurová et al., 2007,
2008, 2015). Similarly, the protococcidian E. duboscqi exhibits an extraordinary
attachment strategy, sharing features with cryptosporidia and gregarines, i.e. the
parasite itself conspicuously resembles an epicellular gregarine, while the
parasitophorous sac develops in a similar manner to that in cryptosporidia
(Tronchin and Schrével, 1977; Valigurová et al., 2007, 2008, 2015). In E. duboscqi
mature stages, a thick glycocalyx layer covers the external surface of their plasma
membrane, which probably hinders potential fusion of the parasitophorous sac with
the parasite surface. In contrast to cryptosporidian parasitophorous sacs, which
have low amounts of F-actin (Bonnin et al., 1999), actin filaments accumulate in the
E. duboscqi parasitophorous sac and are more stable than those in the surrounding
host tissue. Accumulation of host F-actin at the base of the parasitophorous sac
appears to be induced by the parasite, similarly to cryptosporidia; however, the
dense band in E. duboscqi is thinner and closely apposed to the sac’s inner
22
membrane. Restriction of polymerised α-tubulin to the parasitophorous sac wall
indicates involvement of host microcilia in the formation of the E. duboscqi
epicellular niche. Re-evaluation of epicellular development in other apicomplexans
and direct comparison of their niche with that of E. duboscqi (Fig. 7; Valigurová et al.,
2015) has revealed similarities with certain eimeriid coccidia from fish and reptiles
(Eimeria formerly known as Epieimeria, some Goussia, Acroeimeria, Choleoeimeria)
(e.g. Dyková and Lom, 1981; Lukeš, 1992; Molnar and Baska, 1986). The most
distinctive feature common to eugregarines, cryptosporidia, protococcidia and
eimeriids from poikilotherms is that they develop a specialised epicellular hostparasite
interface, reflecting analogous modes of adaptation for development in
similar environments (Melicherová et al., 2018, 2014; Valigurová et al., 2007, 2008,
2015). Extracellular but attached parasites are usually of a heteropolar nature, while
intracellular parasites are generally non-polar. Gregarines, cryptosporidia and E.
duboscqi are heteropolar cells, i.e. they exhibit a high degree of cell polarity in that
their anterior and posterior ends differ in shape, structure and function. They attach
by apical processes, i.e. epimerite in eugregarines, feeder organelle in cryptosporidia
and the apparatus formed by circularly arranged lobes, crowned by a ring of
filamentous fascicles in E. duboscqi. Epicellular eimeriids, on the other hand, appear
to be non-polar, as they have no attachment organelles (Benajiba et al., 1994). They
do, however, create projections of the parasitophorous sac equipped with pores that
enlarge the contact zone with the host cell, resembling the E. duboscqi attachment
lobes and fascicles. Presumably, the epicellular localisation leads to the occurrence
of cell polarity. All these parasites stimulate additional growth and fusion of host cell
microvilli along with modifications to the host plasma membrane, leading to
parasitophorous sac formation; in this way they develop in the cavity of a hostderived
envelope that separates them from the host internal environment. The
parasitophorous sac of cryptosporidia is not complete as they connect directly to the
host cell via a feeder organelle, while the inner membrane of the parasitophorous
sac contains the entire parasite in E. duboscqi and eimeriids. In contrast to
intracellular coccidia, evolutionary selection has presumably favoured an epicellular
niche for these parasites, allowing them to more effectively evade the host immune
response, though the parasite thereby becomes dependent upon its connection with
the host cell for nutrient acquisition (Valigurová et al., 2015).
23
Fig. 7. Schematic diagram of host-parasite interactions in Eleutheroschizon duboscqi,
eugregarines, cryptosporidia and epicellular eimeriids (taken from Valigurová et al.,
2015). Three colours are used to distinguish between the parasite (in purple); the
host cell, including parts modified due to parasitisation (in pink) and the contact
zone between the host and the parasite (in yellow), where the interrelationships of
the two organisms become more intimate. In E. duboscqi and epicellular eimeriids,
the internal space between the parasite and parasitophorous sac (PS) remained
24
colourless, even though the authors do not exclude the possibility that this region
may serve as a transitional zone for intensive interactions between the host and
parasite. A-D. Eleutheroschizon duboscqi. A. Attached zoite transforming into a
trophozoite, completely enveloped by a PS. B. Maturing trophozoite with a forming
ring of fascicles at the attachment site. C. Mature trophozoite with well-developed
attachment fascicles and lobes. D. Detail of the annular joint point (the cut-out is
marked by a red square in C). E-H. Eugregarines. E. Sporozoite freshly attached to
the host epithelial cell. F. Sporozoite transforming into a trophozoite. G. Early
trophozoite with a well-developed epimerite. H. Detail of the membrane fusion site
(the cut-out is marked by red square in G). The two cytomembranes end at the point
of membrane fusion (= osmiophilic ring). I-L. Cryptosporidia. I. Attached zoite
transforming into a trophozoite, partially enveloped by a PS. J. Young trophozoite
almost completely enveloped by a PS. The tunnel connection between the interior of
the anterior vacuole and the host cell cytoplasm develops as a result of the Y-shaped
membrane junction. K. Mature stage with a filamentous projection at the base of the
PS and fully developed feeder organelle, the lamellae of which form from the anterior
vacuole membrane. L. Detailed view of the Y-shaped membrane junction (the cut-out
is marked by a red square in K). M-P. Epicellular eimeriids. M. Invading zoite. N.
Trophozoite/meront stage enveloped by a PS with a single attachment area
(monopodial form). O. Extension of the gamont stage above the host microvillous
region leading to establishment of a new contact with the host cell apart from the
primary attachment zone by penetration of the PS membrane to the base of fused
host cell microvilli (spider-like form). P. Detailed view of the attachment area (the
cut-out is marked by a red square in O). av – anterior vacuole, b – epimeritic bud, cm
– parasite cytomembranes, cv – epimeritic cortical vesicle, db – dense band (in
cryptosporidia usually consisting of several layers), f – membrane fusion site, dl –
dense line separating the feeder organelle from the filamentous projection of the PS,
fa – attachment fascicle of filaments, fo – feeder organelle with membranous
lamellae, fom – membrane limiting the lamellae of feeder organelle, fp – filamentous
projection of the PS, fs – flask-shaped structure, hc – host cell, hm - host cell plasma
membrane, if – incomplete fusion of PS, int – interface between the host cell and
epimerite, ipm – inner membrane of the PS, is – internal space between the parasite
and PS, j – annular joint point (Y-shaped membrane junction in cryptosporidia), lo –
attachment lobe, ms – membrane-like structure limiting the cortical vesicle from the
epimerite cytoplasm, opm – outer membrane of the PS, p – pore on the PS, pm –
parasite plasma membrane, ps – parasitophorous sac, r – rhoptries, t – tail of the PS,
tu – tunnel connection.
Aside from Ditrypanocystis archigregarine, which develops within a
multimembranous envelope originating from fused enterocyte cilia, gregarines are
usually not surrounded by a host-derived sac (Butaeva et al., 2006). Host cilia,
clustering around Ditrypanocystis, lose their microtubular content and fuse to form
the membrane of a parasitophorous sac (Fig. 8), the contact area of which is
considerably enlarged. The ciliary membranes fold heavily beneath the parasite
attachment site to form a network of channels, opening in the immediate proximity
25
of the enterocyte apical surface and contacting directly with the contents of the
enterocyte transport vacuoles that cross the host cell plasma membrane. In contrast
to cryptosporidia, neither parasite membrane folds (feeder organelle) nor fusion
with the host cell membrane are formed. The trophozoite pellicle and subpellicular
microtubules remain preserved in the contact region. These host-parasite
interactions, which differ from other Selenidiidae, may result from parasitism of
polychaetes that are not primarily marine (e.g. Enchytraeus albidus).
Fig. 8. Hypothetical scheme of host-parasite interface formation in archigregarine
Ditrypanocystis sp. (taken and modified from Butaeva et al., 2006). A. Parasite
contacting cilia of the host enterocytes. B. Enterocyte cilia transform and cluster
around the parasite, with ciliary plasma membranes forming branched folds in the
host-parasite contact region. C. Plasma membrane of clustered cilia fuse to form a
parasitophorous sac (PS) with a heavily folded contact area. ci – enterocyte cilia, cm
– membrane of cilia, dmc – degraded microtubules of cilia, ftcm – folds of
transformed ciliary membrane, ipv – inner space of PS, mt – microtubules, mvi –
inner membrane of PS, mvo – outer membrane of PS, p – parasite, phc – host cell
plasma membrane, tcm – transformed ciliary membrane.
3.3 Niches of intracellular parasites
Eugregarines do not usually exhibit intracellular development. However,
intracellular stages are documented in some archigregarines, in which the
sporozoites, released into the lumen of host intestine, move through the intestinal
epithelium and reach the basal lamina to transform into trophozoites and undergo
merogony. In polychaetes highly parasitised by S. hollandei, for example,
intraepithelial cysts located near the basal lamina contain several dozen welldeveloped
merozoites (Schrével and Desportes, 2016). In S. pygospionis, small
trophozoites with cellular organisation identical to that of well-developed
26
trophozoites locate intracellularly within a parasitophorous vacuole (Paskerova et
al., 2018). The internal space of this parasitophorous vacuole contains filamentous
material and electron-dense granules, accumulation of which increases towards the
periphery. In enterocytes, a host cytoplasm rich in dense fibrillar material,
mitochondria, endoplasmic reticulum and vesicles surrounds the parasitophorous
vacuole, whereas the rest of the host cell cytoplasm is electron-lucent with rare
organelles. Similarly, reduced vegetative stages of neogregarines, which mostly
invade insect fat bodies, the haemocoel, the Malpighian ducts and intestines, are
usually intracellular or lie within tissues, with no or a reduced attachment region
(Schrével and Desportes, 2016). Some of these are reported to be enclosed within a
parasitophorous vacuole (Larsson, 1991; Vávra and McLaughlin, 1970; Žižka, 2005).
Of special interest is the occurrence of the enigmatic protist N. temporariae in
tadpoles, observed for the first time in 1920 (Nöller, 1920) and putatively assigned
to gregarines. A recent study, using both microscopy and ribosomal DNA sequencing
of N. temporariae oocysts from three different frog species (Rana temporaria, R.
dalmatina and Hyla arborea), has confirmed that this parasite belongs to the
Gregarinasina (Chambouvet et al., 2016). This is the only known case of a gregarine
found in a vertebrate. The oocysts of N. temporariae, including empty ones, are
regularly present in macrophages located in tadpole liver sinusoids and contained
within a parasitophorous vacuole (Valigurová, personal observation). All tadpoles
investigated were Nematopsis positive, while only empty oocysts were found in
organisms 4–6 weeks after metamorphosis. Oocysts are the only gregarine
developmental stage found in dissected tadpoles, making it unclear whether the
tadpoles serve as definitive or intermediate hosts (Chambouvet et al., 2016).
The poorly investigated life cycle of agamococcidian Rhytidocystis spp. in
polychaetes is expected to be streamlined and involving sporozoites that penetrate
the host intestine and then persist in the connective tissue, gonads and coelom,
developing into trophozoites (Rueckert and Leander, 2009b). Trophozoites later
form numerous sporoblasts by budding from their surface (Perkins, 2000). In a new
species of Rhytidocystis, several intracellular developmental stages covered by a
trimembrane pellicle (all of them lacking the parasitophorous vacuole), distributed
along the middle third of the host intestine (Diakin and Valigurová, 2014).
Presumably, sporozoites penetrate the enterocytes and, during trophozoite
27
maturation, move to the base of the host cell. These, unlike R. polygordiae
trophozoites reported as residing within the extracellular matrix between the
epithelial cells (Leander and Ramey, 2006), induce formation of a syncytium around
themselves from several of the surrounding epithelial cells. The apical part of the
syncytium exhibits a typical morphology, though the basal region is free of organelles
and cytoplasmic inclusions. Two or more mature stages can occur in one syncytium.
Numerous giant mitochondria with tubular cristae accumulate at the trophozoite
periphery, closely adjacent to the pellicle. The developmental stages differ in the
density of cytoplasm and morphology of some organelles (mitochondria, nucleus).
Some forms possess rhizoid-like appendages (Diakin and Valigurová, 2014). Neither
a mucron nor an apical complex is present in species investigated so far (Diakin and
Valigurová, 2014; Leander and Ramey, 2006; Rueckert and Leander, 2009b).
3.4 Feeding strategies
While the nutrition of gregarines has been the subject of extensive debate, their
mechanism of nutrition acquisition remains poorly understood. The gregarine
feeding mode depends on the long-term environmental conditions forming their
niche. Correlations between trophozoite characteristics and the environment
occupied within the host are discussed elsewhere (Leander et al., 2006).
Archigregarines, the earliest diverging apicomplexan lineage, have retained
myzocytosis as their principal feeding mode and myzocytosis is also expected to
occur in blastogregarines (Simdyanov et al., 2018). Unlike blastogregarines, where
the mucronal complex remains active (myzocytosis) during the trophozoite lifespan,
archigregarines have non-feeding gamonts, though their conoid and rhoptries
persist until at least progamic mitoses starts (Desportes and Schrével, 2013;
Simdyanov and Kuvardina, 2007; Simdyanov et al., 2018). Myzocytosis is clearly
illustrated in S. pendula, with mucronal (digestive) vacuoles inserted into the conoid
and surrounded by numerous rhoptries and micronemes (Schrével et al., 2016). The
myzocytosis process starts at the top of the conoid, with continuity of the mucronal
vacuole membrane up to its contact with the host epithelial cell indicating that
evagination through the apex of the conoid allows the parasite to suck out
nutriments from the host cell (Fig. 9). The initial mucronal vacuole then fragments
into numerous vacuoles (Schrével et al., 2016; Simdyanov and Kuvardina, 2007). An
28
axial streak of optically distinct cytoplasm reported in some species, extending from
the anterior to the posterior end and forming an expansion around the nucleus, may
represent a nutrient distribution system (Fowell, 1936; Paskerova et al., 2018). The
presumed digestive vacuoles observed in the axial row, most likely originating in the
mucron during myzocytosis, transport nutrients posteriorly along the cell axis.
Accordingly, numerous vacuoles surround the nucleus in S. pendula (Schrével, 1971).
Fig. 9. Putative feeding scheme of Selenidium archigregarines (taken and modified
from Simdyanov and Kuvardina, 2007). A. Local lysis of the host cell plasma
membrane by rhoptry (r) secretion (red arrow). B. The process of myzocytosis:
swallowing of the host cytoplasm through the temporary cytostome–cytopharyngeal
complex (red arrow) and formation of the nascent mucronal vacuole (mv). C.
Cytopharynx closure, the nascent mucronal vacuole (mv) is fully formed; the
previous vacuole is divided into smaller vacuoles that are transported via a
microtubular network (mt) into the trophozoite. The disrupted site of the host cell
plasma membrane is restored. Numbers 1–3 indicate the positions of vacuoles
formed during the preceding myzocytosis events (the current event is No. 4).
On the other hand, many eugregarines seem not to feed through the myzocytosis,
except, perhaps, at the youngest developmental stage (Hildebrand, 1976; Valigurová
et al., 2015). The apical complex of eugregarine trophozoites is reduced and instead
a complicated attachment apparatus forms. Some authors attribute the feeding
function in eugregarines to these attachment organelles (Schrével and Philippe,
1993). The concentration of host cell mitochondria and the endoplasmic reticulum
surrounding the epimerite, along with the presence of parasite mitochondria located
29
just beneath the epimeritic cortical vesicle, indicate both the existence of active
interaction between the eugregarine epimerite and the host cell (Hildebrand, 1976;
Lucarotti, 2000) and that the epimerite is a metabolically active organelle (Baudoin,
1969; Ormieres, 1977). The abundant endoplasmic reticulum observed in the
expanding epimerite of young trophozoites indicates activation of metabolic
pathways (Valigurová, 2012). As host cells affected by attached eugregarines usually
do not show any obvious pathological changes, the epimerite cortical vesicle and
vacuoles most likely absorb nutrients via a mechanism based on membrane
permeability, while numerous mitochondria underlying the cortical vesicle could
provide the energy necessary for absorption (Schrével and Philippe, 1993; Tronchin
and Schrével, 1977; Valigurová and Koudela, 2008; Valigurová et al., 2009). A
reduction in the size of cortical vesicle in some species appears to be related to the
convoluted character of their epimerites, significantly increasing the absorptive
surface (Hildebrand, 1976; Valigurová, 2012). In the monocystid eugregarine
Nematocystis, the trilaminar junction between its mucron and host cell forms
extensive folds that increase the contact zone between apposing cell membranes
(MacMillan, 1973b). Using radioisotopes, this study demonstrated that metabolites
pass directly from host cell to the trophozoite by crossing the attachment site.
However, feeding strategies differ between distant eugregarine taxa (Valigurová,
2012). An example can be found in the lecudinids, whose supposed lytic effect on
host tissue is an indicator for the nutritional function of the mucron via extracellular
secretion of enzymes and absorption of digested host tissue (Schrével and Philippe,
1993). The high concentration of actin-like proteins in the mucron of Lecudina
pellucida, corresponding to the numerous 7-nm filaments in this region, concurs with
its supposed sucker function. The protomerites, modified for attachment like those
described in G. cuneata and actinocephalid gregarines, act as feeding organelles in
eugregarines lacking an epimerite (Cook et al., 2001; Valigurová, 2012). Numerous
pores and ducts interrupting the pellicle of G. cuneata at the protomerite apex, along
with abundant dense bodies, vesicles and the Golgi apparatus in the protomerite
cytoplasm, are most likely involved in gamont nutrition and/or attachment. A
further nutrition possibility may be suggested, especially for gregarines growing in
coelomic fluid with no attachment to the host tissue, and considering the fact that
gregarines generally continue to grow after detaching from the host tissue (Schrével
30
and Philippe, 1993; Valigurová, 2012). The extensive folding of the pellicle covering
the surface of large trophozoites in marine eugregarines appears to optimise
surface-mediated nutrition (pinocytosis via micropores), and could explain the loss
of myzocytosis and apical complex in eugregarines accompanied by development of
a bulky attachment apparatus (Leander, 2008). Questions arise as to whether, and
under what circumstances, the epimerite serves as a feeding organelle, and whether
the gregarine micropores are points implicated in pinocytosis (Vivier et al., 1970) or
serve for mucus extrusion (Schrével, 1972; Valigurová et al., 2013; Warner, 1968).
The role of micropores in feeding appears to be especially true in eugregarines, the
sporozoite of which is equipped by a single micropore located in its anterior third
(Desportes, 1969; Sheffield et al., 1971), while advanced stages possess numerous
micropores located in the grooves between the epicytic folds and smaller pores
randomly distributed on the fold’s base or lateral side (Diakin et al., 2016;
Kováčiková et al., 2017b, 2018; Lucarotti, 2000; Schrével et al., 1983; Valigurová et
al., 2013; Walker et al., 1984). While no typical micropores are present in the
blastogregarines (Simdyanov et al., 2018; Valigurová et al., 2017), there is evidence
for pinocytosis in Selenidium archigregarines based on rows of micropores with
associated pinocytotic whorled vesicles interrupting the pellicle in the grooves
between the longitudinal bulges (Kováčiková et al., 2017a; Leander, 2007; Paskerova
et al., 2018; Schrével et al., 2016).
While Cryptosporidium exhibits unique features that characterise its
biochemistry and metabolism (Thompson et al., 2016), its mechanism of nutrient
acquisition remains unresolved. Presumably, the parasitophorous sac has a
protective role, while the feeder organelle, directly separating the host cell and the
parasite cytoplasm, provides an entry point for nutrients from the host (Tzipori and
Griffiths, 1998; Valigurová et al., 2007). The feeder organelle, consisting of numerous
folds that markedly enlarge the contact area between host and parasite, may be the
site of nutrient and drug transport regulation into the parasite (Perkins et al., 1999).
The feeder organelle lamellae are fringed by structures suggestive of endocytic
vesicles (Landsberg and Paperna, 1986). Freeze fracture replicas show that the
membrane folds of the feeder organelle close at the attachment site, but connect with
cytoplasmic vesicles on the opposite side (Yoshikawa and Iseki, 1992). The parasiteinduced
cytoskeletal rearrangement in the host cell probably results in the formation
31
of a network for vesicle trafficking, facilitating the movement of nutrients between
the host cell and the parasitophorous sac (Forney et al., 1999). The majority of
studies suggest that cryptosporidia rely solely on the host for nutrient acquisition
and encode for a number of transporters that likely serve this purpose (O'Hara and
Chen, 2011). The presence of ABC-cassette binding proteins at the parasite-host
interface provides further support for this hypothesis (Perkins et al., 1999). Based
on their similarity to gregarines, the question arises as to whether the feeder
organelle obtains nutrients from the host cell in an analogous manner to
myzocytosis. Considering the debatable existence of Cryptosporidium extracellular
stages reported from in vitro systems, it is unclear how this parasite, which lacks key
de novo synthesis of amino acids, fatty acids, and nucleosides, acquires nutrients
directly from an extracellular niche. As the feeder organelle has been observed in
extracellular stages from biofilms, cryptosporidia may be able to acquire nutrients
in a cell-free environment (Koh et al., 2014; Thompson et al., 2016).
It has already been emphasised that the general picture of metabolic interaction
between host and parasite in cryptosporidia resembles that documented in Eimeria
spp. (Beyer et al., 1995). However, epicellular eimeriids, though sharing features
with cryptosporidia, are likely to differ in the mode of nutrient uptake, which
probably occurs through the parasitophorous sac membrane which is considerably
enlarged by projections equipped with pores, thereby increasing the area of the
contact with the host cell (Benajiba et al., 1994; Beyer et al., 2002; Valigurová et al.,
2015). Feeding via myzocytosis would also appear to be atypical for E. duboscqi, as
its endogenous stages lack the apical complex. Further, there is no organelle similar
to the flask-shaped structure nor a mucronal vacuole in freshly attached E. duboscqi
parasites. Whether the complicated attachment apparatus of E. duboscqi is involved
in the nutrient acquisition remains unclear. Numerous pores distributed along the
entire parasite pellicle and the attachment site could be involved in feeding, with the
second option appearing more likely as some pores appear to be associated with
parasite vesicles and mitochondria (Valigurová et al., 2015).
3.5 Pathogenicity to the host
As a rule, eugregarines are considered non-pathogenic to their hosts (Henry,
1981); however, their actual impact on host fitness and viability has been poorly
32
investigated. Considering that the parasite load in eugregarines depends entirely on
the number of oocysts ingested, they are relatively benign. Theoretically, heavy
gregarine infestation in the mesenteron could affect the host’s nutritional state.
Some species could occlude the host gut, thereby preventing food passage (Lucarotti,
2000); however, this disagrees with a study on water striders Gerris buenoi, which
showed eugregarines to be rather harmless commensals as there was no reduction
in final host size or prolonged development time, despite heavy infections almost
blocking the gut passage (Klingenberg et al., 1997). Similarly, eugregarines from T.
molitor larvae kept in laboratory colonies did not appear to harm their host; instead,
they seemed to be mutualistic as their presence had a positive impact on host
development, fitness and longevity, despite heavy infection (Sumner, 1933;
Valigurová, 2012). Some substances (e.g. vitamins or enzymes) secreted by these
gregarines may be essential for larval growth. Pathogenicity is mostly attributed to
the trophozoites as they could theoretically cause some degree of damage to
parasitised tissue, depending on the dimensions and shape of their attachment
structures (Lipa, 1967). Epicellular trophozoites mostly affect the microvillous site
of epithelial cells. Microvilli are essential for efficient absorption and excretion,
hence their destruction might limit host digestion and lead to malnutrition, resulting
in weakening or even death (Valigurová, 2012). However, although the intestinal
epithelium of parasitised mealworms shows some changes that could be associated
with gregarine infection (i.e. vacuolation of individual parasitised cells),
eugregarines have no negative impact on host health (Valigurová, 2012). Likewise,
there is no evidence of direct damage to neighbouring epithelial cells caused by
trophozoites attached to intestinal tissue, even at their high densities. Presumably,
the continual regeneration of epithelial cells accounts for the apparent harmless
effect of the parasite. Even if the trophozoites destroy individual cells, therefore, the
overall damage to the epithelium is negligible and easily repaired. Even robust
epimerite with rigid hooks in Ancyrophora does not appear to induce drastic damage
to host cells (Baudoin, 1969). Species occurring in intestinal caeca appear more
pathogenic as they cause hypertrophy of parasitised cells, or even rupture of the
caecal wall, leading to secondary bacterial infection (Tanada and Kaya, 1993). The
gregarine N. temporariae from tadpoles also appears not to cause disease or fitness
impairment in the host, though livers of some tadpoles appear slightly enlarged and
33
light coloured; indeed, no mortalities of tadpoles or metamorphs have been recorded
in the field (Chambouvet et al., 2016).
In contrast, neogregarines undergo multiple division (merogony), hence the level
of parasitisation is not necessarily related to the number of oocysts ingested by the
host. Further, autoinfection occurring in some neogregarines (Naville, 1930;
Valigurová and Koudela, 2006; Weiser, 1954) appears to contribute to a rapid spread
of infection. Consequently, neogregarines cause significant pathological changes to
their insect hosts and are frequent causes of host morbidity and mortality due to cell
lysis and destruction of tissue invaded by the merogony stages (Schrével and
Desportes, 2016; Tanada and Kaya, 1993; Valigurová and Koudela, 2006). As hosts
often do not survive heavy neogregarine infection, they are considered potential
candidates for biological control of insect pests (Žižka, 1977). Neogregarine
nutritional requirements are very high and, at the end of parasite development, the
destroyed host tissue is replaced with neogregarine oocysts (Weiser, 1954). For
example, the fat body cells of Ephestia kuehniella larvae parasitised by Mattesia
dispora become vacuolated and degenerate (Valigurová and Koudela, 2006). Though
M. dispora parasitises the host cell with no obvious response, its meronts decrease
fat excretion and deposition within the parasitised cell (Weiser, 1954).
Consequently, the fat body becomes overgrown with plasma-rich cells lacking lipid
vacuoles. With progressing infection, the affected cells appear hypertrophied with
the nucleus pushed back. During the micronuclear merogony of M. dispora, a
moderate reduction in host activity and food intake can be observed, while larvae
exhibit lethargy and cease moving during the course of macronuclear merogony.
Thereafter, their bodies become pale and the larvae cease feeding, following which
the infection spreads to the entire larval body. During the final stage, the haemocoel
of the insect is packed with numerous oocysts, while the tissue is destroyed.
Massively infected larvae turn an intense pink colour before death (Valigurová and
Koudela, 2006). On the other hand, in Farinocystis triboli, which does not induce
formation of parasitophorous vacuole, presence of host mitochondria surrounding
the meronts is the only response to infection (Žižka, 1977). It is likely that the
formation of parasitophorous vacuole results from an interaction between parasite
invasion and host defence, as documented in Galleria mellonella parasitised by M.
dispora (Žižka, 2005).
34
It has been suggested that epicellular development might be considered as a
more primitive form of host-parasite association (Paperna and Landsberg, 1989).
There is evidence, however, that epicellular parasites have a lower negative effect on
the host epithelium than intracellular parasites (Eli et al., 2012). The attachment
strategy of epicellular parasites may be more progressive as, generally speaking, it is
more advantageous for the parasite to maintain acceptable fitness in their hosts. As
with the flat holes left in the epithelium (devoid of microvilli) by detached gregarine
trophozoites, detached stages of cryptosporidia and E. duboscqi leave only shallow
craters, with the parasitophorous sac remaining at the epithelial surface (Jirků et al.,
2008; Valigurová et al., 2008, 2009, 2015). In addition, cryptosporidia and E.
duboscqi regularly detach from the unmodified part of the host cell with their sacs;
while the detachment of cryptosporidia takes place in the area of the dense band, E.
duboscqi parasites expose their naked basis when tearing away from
parasitophorous sacs, leaving the intact inner sac membrane at the place of
attachment. In contrast to cryptosporidia, E. duboscqi appears to induce only
moderate alterations in the host cell. Cytoskeletal remodelling of epithelial cells
induced by cryptosporidia leads to microvillous hypertrophy, i.e. elongation and
protrusion of host cell microvilli surrounding the parasite (Forney et al., 1999). The
long microvilli clustered at the attachment site of cryptosporidia suggest an active
manipulation of the host membrane structure by the parasite. The microvilli
associated with the cryptosporidian parasitophorous sac are thick and packed with
dense bundles of F-actin. Despite having a similar attachment strategy to
cryptosporidia, E. duboscqi has a lower pathological effect on host tissue as no
pathological changes or significant extension of adjacent microvilli occurs in
parasitised tissue (Valigurová et al., 2015).
In immunocompetent hosts, cryptosporidiosis is usually an acute, self-limiting
gastrointestinal disease, characterised by watery diarrhoea, abdominal cramps,
vomiting, low-grade fever and appetite loss. The intensity of pathological alteration
depends on parasite species and load, as well as host age and immunological status.
Increased morbidity and mortality has been reported in patients with severe
immunodeficiency and in malnourished children. Other pathogens, such as
Helicobacter, Escherichia coli and rotavirus, may escalate the impacts of
cryptosporidiosis on host tissue, possibly leading to death (Santin, 2013; Tatar et al.,
35
1995). The overall effect of parasitism on host health is closely related to the
parasite’s preference for particular target tissues or cells. Presumably,
cryptosporidia prefer a specific type of cells for attachment as direct observations on
cultures have shown that, while numerous sporozoites inspect potential host cells
with their prolonged apical ends, they often leave without invasion (Melicherová et
al., 2018). Indeed, the developmental stages of C. proliferans and C. parvum
frequently occur near dividing or newly formed round cells (Melicherová et al., 2018;
Tůmová, 2019). Particularly interesting is that cryptosporidia appear to interact
with and regulate host-cell gene expression as they can inhibit (trophozoites) or
promote (sporozoites, merozoites) host cell apoptosis, depending on their
developmental stage (Chen et al., 1998; Mele et al., 2004). A loss of absorptive surface
area is frequently associated with intestinal cryptosporidiosis (Leitch and He, 2012).
For example, the major histopathological changes induced by C. parvum infecting
enterocytes of the distal small intestine, caecum and colon comprise villous atrophy,
shortening of microvilli and sloughing of enterocytes (Santin, 2013). Gastric
cryptosporidiosis in animals tends to cause mild histopathological changes with no
obvious alterations to host health status, and no, or only insignificant, inflammatory
responses in lamina propria. Cryptosporidia affect gastric tissues irregularly, in an
island-like manner (Melicherová et al., 2014; Val-Bernal et al., 2013; Valigurová et
al., 2018). For example, C. muris, a gastric species from rodents, induces less
significant pathological changes than C. proliferans (Kváč et al., 2016), where
pathological changes in mouse gastric tissues escalate during chronic phases of
cryptosporidiosis, including marked deformation of the gastric glandular epithelium
surface and a cauliflower-like appearance in gastric mucosa (Valigurová et al., 2018).
The heavily parasitised epithelium proliferates into the luminal space, resulting in
extensive folding of the stomach. Subsequently, expansion of the lamina propria
leads to an increase in the distance between gastric glands, twisting, and
deformation of the longitudinal folds, and diffuse mucosal hypertrophy occurs,
typified by the presence of giant gastric folds and intensive epithelial hyperplasia.
The gastric glands, packed with parasites and necrotic material, become markedly
dilated and hypertrophied and lose their normal architecture, while the atrophic
epithelial cells exhibit cuboidal or squamous metaplasia. A thickening of the
muscularis mucosae is typical of advanced gastric cryptosporidiosis. The tissue
36
exhibits oedema and infiltration of the lamina propria and submucosa with
neutrophils. Though stomach weight and epithelial height increase considerably,
neither clinical signs of cryptosporidiosis nor weight lost have been documented in
laboratory rodents parasitised by C. proliferans (Kváč et al., 2016; Melicherová et al.,
2014; Valigurová et al., 2018).
4 Motility in early branching Apicomplexa
4.1 Apicomplexan motility and cytoskeleton
Zoites, the highly motile stages of Apicomplexa, exhibit considerable variation in
movement, such as the progressive circular or helical gliding and non-progressive
twirling of T. gondii and Plasmodium zoites. Reported gliding rates in Apicomplexa
are usually in the range of 1–10 μm/s, with maximum rates reported from
gregarines, e.g. 22.86 μm/s in G. polymorpha and up to 60 μm/s in Porospora
gigantea (King and Sleep, 2005; King, 1988; Valigurová et al., 2013). Apicomplexans
are generally characterised by a complicated cell cortex consisting of a continuous
plasma membrane, underlined by cortical alveoli forming the IMC. The IMC connects
with cytoskeletal elements, such as the actomyosin complex, microtubules and the
network of intermediate filamentous proteins, and may be interrupted by
micropores (Morrissette and Sibley, 2002; Valigurová et al., 2013). Remarkably,
apicomplexan subpellicular microtubules are unusually stable and withstand high
pressure, cold and detergents, while actin filaments are transient. Actin is mostly
present in its globular form and microfilaments are usually detected only after
treatment with F-actin stabilising drugs such as jasplakinolide. Previous studies
(focusing predominantly on T. gondii and Plasmodium) concur with the so-called
glideosome concept applied for apicomplexan zoites, describing a substratedependent
gliding motility facilitated by a conserved form of actomyosin motor and
subpellicular microtubules. This actomyosin motor is thought to be localised
between the plasma membrane and IMC, with zoite gliding based on the locomotion
of myosin fixed to the IMC along actin filaments, together with translocation of
apically released adhesins to the parasite's posterior end, resulting in a forward
movement (Daher and Soldati-Favre, 2009; Frenal et al., 2010; Tardieux and Baum,
2016). This unique machinery (which requires a stable subpellicular network of
37
microtubules to provide structural stability and maintain polarity) is based on, and
limited by, the formation of transient actin filaments and their fixation to the IMC.
Basal apicomplexans differ from other Apicomplexa in that (i) their trophozoites
and gamonts are often motile, at least to some degree; (ii) locomotion usually differs
from substrate-dependent gliding based on glideosome, and (iii) they use several cell
motility mechanisms that correlate with various modifications of their cell cortex
(epicyte). The differing motility modes exhibited by basal Apicomplexa, comprising
bending, rolling, coiling and nematode-like motility, gliding, metaboly or peristalsis,
most likely represent specific adaptations to parasitism in different environments
within their hosts (Valigurová et al., 2013, 2017).
4.2 Motility and cytoskeleton in archigregarines and blastogregarines
The spindle-shaped trophozoites and gamonts of Selenidium archigregarines
display bending, coiling, rolling and pendular non-progressive movements, along
with cell shape contraction (Fowell, 1936; Kováčiková et al., 2017a; Leander, 2007;
Schrével et al., 2016). The pendular motility of S. pendula exhibits a regular
periodicity of about 2–3 seconds, with propagation waves generated in the parasite’s
anterior region (Desportes and Schrével, 2013). The trimembrane pellicle of most
Selenidium spp. is organised in broad longitudinal bulges separated by grooves;
though additional cortex modifications occur in species that contract their body. For
example, the motility of S. vivax trophozoites, which actively and irregularly change
their conformation by twisting, folding, shrinking and expanding cell volume, is
somewhat peristaltic (Desportes and Schrével, 2013; Leander, 2006). The folded
pellicle of contracted and semi-relaxed parasites exhibits transverse striations,
which vary in depth and number while they disappear and reappear (Rueckert and
Leander, 2009a). In general, the pellicle is underlain by longitudinally oriented
subpellicular microtubules, which tend to be arranged in a single layer, the
continuity of which is interrupted under the grooves with micropores, and deeper
located groups of irregularly arranged microtubules (Kováčiková et al., 2017a;
Paskerova et al., 2018; Schrével et al., 2016). The pellicle may act as a stiff skeletal
component, while the regular sets of subpellicular microtubules have a motility
function (Macgregor and Thomasson, 1965). It has been proposed that this
represents a unicellular analogue to the musculocuticular system of nematodes, in
38
which longitudinal muscles function antagonistically against the elastic cuticle
(Leander, 2007; Stebbings et al., 1974). Archigregarines lacking subpellicular
microtubules are non-motile, though their longitudinal bulges are supported by
arrays of fibrils reminiscent of circular myonemes (Wakeman et al., 2014). The
intracellular axial streak with laterally branching radial fibrils reported in some
Selenidium archigregarines may be involved in motility as an additional skeletal
element helping to reverse movements and maintain cell shape in bends, as
suggested by the statomotor system concept (Fowell, 1936; Paskerova et al., 2018).
The elongated, flattened trophozoites and gamonts of S. nematoides exhibit highly
active movements, with motility of both attached and detached parasites in a liquid
environment resembling a sequence of undulation, pendular, twisting and,
occasionally, spasmodic movements (Valigurová et al., 2017). In attached parasites
with typically wavy movements, the waves develop in their proximal region, just
behind the attachment area, and proceed distally, the last third of the cell being more
rigid with limited mobility. The bending movements of detached individuals also
initiate from the apical region. The blastogregarines beat at a rate of 0.51 beats per
second on average, with a beat-to-beat interval of 2.18 seconds. In S. nematoides, the
surface of the trimembrane pellicle in endogenous stages (from early trophozoites
up to gamonts) is smooth, lacking grooves or folds. A distinct glycocalyx layer,
thickening towards the apical region, covers the entire parasite and, in mature
stages, numerous pores interrupt the pedicle, mostly organised in four lateral rows
running parallel to the longitudinal cell axis. The helical arrangement of
microtubules in S. nematoides follows its serpentine body shape, similar to those of
apicomplexan zoites. The regularly arranged subpellicular microtubules exhibit a
characteristic longitudinal organisation and are nucleated from the apical polar ring,
a microtubule-organising centre unique to the Apicomplexa. In contrast to
glideosome, the majority of S. nematoides actin is present in a polymerised form and
appears to be located beneath the IMC. The subpellicular microtubules are
associated with filamentous structures (cross-linking protein complexes),
presumably of an actin-like nature (Valigurová et al., 2017). In contrast to the active
movement of S. nematoides, C. mesnili blastogregarines show only weak motility,
typified by slow and intermittent bending movements. These differences in motility
can be attributed to specific cortex modifications. The pellicle of C. mesnili, covered
39
by a trimembrane pellicle organised in longitudinal folds with flattened tops, is
underlined by numerous longitudinal and regularly distributed subpellicular
microtubules which are arranged in two layers at the tops of the folds and as a single
layer between the folds and on their lateral sides. These microtubules arise from the
fibrillar matter lying beneath the pellicle in the frontal region of the mucron. The
putative polar ring of this species is not subdivided, as observed in S. nematoides, and
does not contact with the microtubules, even though they are abundant within the
mucron (Simdyanov et al., 2018).
Due to similarities in their external morphology and movement patterns,
Siedleckia has been associated with Selenidium archigregarines. Despite bearing a
striking resemblance with overgrown apicomplexan zoites, the trophozoites and
gamonts of both genera move independently on solid substrates with no signs of
gliding motility. The motility mechanisms of S. nematoides and Selenidium
archigregarines differ from the glideosome, despite the presence of key glideosome
components such as a pellicle comprising a plasma membrane and IMC, subpellicular
microtubules, actin, myosin, micronemes and a glycocalyx layer. The subpellicular
microtubules organised in several layers appear to be the leading motor structures
in blastogregarine and Selenidium spp. motility. In both taxa, the individual
microtubules are localised within lucent areas (the so-called chambers) that most
likely act in microtubule sliding. Experimental assays have shown that polymerised
forms of actin and tubulin play an essential role in their movement (Kováčiková et
al., 2017a; Valigurová et al., 2017). Nevertheless, while treatment with membranepermeable
drugs influencing polymerisation of actin, jasplakinolide and cytochalasin
D, affects motility and actin organisation in S. pygospionis, no significant changes
occur in the position of the subpellicular microtubules, contrary to observations on
S. nematoides. The cross-linking protein complexes in S. nematoides, which
presumably anchor the subpellicular microtubules to the cytoplasmic face of IMC,
may correspond to microtubule-associated proteins (MAPs) (Valigurová et al.,
2017). MAPs are expected to control the spacing of microtubules within the cell by
interconnecting them with other cytoskeletal elements or with the plasma
membrane. It is likely that the heavy decoration of subpellicular microtubules in
Apicomplexa may account for their unusual stability. The fact that tubulin-specific
antibodies commonly do not label the full length of microtubules in apicomplexan
40
zoites may be due to occlusion of tubulin epitopes by MAPs (Tran et al., 2012). The
MAPs, such as dyneins or kinesins, are responsible for sliding between adjacent
microtubules. As with the ciliary axoneme, a mechanism with microtubules sliding
against one another could account for the undulating motility of blastogregarines
and archigregarines. A MAP-based mechanism has been proposed to explain the
undulating and bending movements in Selenidium spp. (Leander, 2006; Mellor and
Stebbings, 1980; Schrével, 1971; Stebbings et al., 1974). Numerous peripheral
mitochondria in Selenidium spp. and S. nematoides probably play an important role
in the rapid and continuous generation of ATP, which is essential for the support of
highly dynamic cell plasticity and may provide the chemical energy necessary for
MAP activity. Actively moving Selenidium spp. exhibit more ectoplasmic
mitochondria and subpellicular microtubules than less active species (Leander,
2006, 2007; Paskerova et al., 2018; Schrével, 1971; Valigurová et al., 2017). If
axoneme-like sliding of microtubules is applicable to S. nematoides, the putative actin
cytoskeleton associates with the subpellicular microtubules lengthwise in order to
position them within the cytoplasm just beneath the pellicle; otherwise, the actin
filaments could force synchronised bending of the microtubules in some cell regions,
thereby generating typical undulating motility (Valigurová et al., 2017).
4.3 Motility and cytoskeleton in eugregarines
Eugregarine sporozoites display a typical apicomplexan zoite organisation. The
sporozoites are covered by a trimembrane pellicle underlain by subpellicular
microtubules and their apical region is equipped with a conoid, polar rings, rhoptries
and numerous micronemes (Desportes, 1967; 1969; Diakin et al., 2014; Sheffield et
al., 1971). While a sinuous movement without gliding activity is reported for Pyxinia
crystalligera sporozoites (Collins, 1972), sporozoites of N. temporariae glide and
keep their banana shape, with only the apical end appearing fully flexible
(Chambouvet et al., 2016). Sporozoites of G. rigida contacting the host epithelium
form a small conical protuberance devoid of endoplasm, elongating into a slender
neck as long as the sporozoite itself (Kamm, 1920). Later stages of eugregarines
(comprising trophozoites and gamonts) exhibit diverse modes of locomotion and
appear to use several motility mechanisms, depending on their physiological and
environmental conditions. Their complicated cell cortex, the epicyte, forms a range
41
of superficial structures. The majority of eugregarines, for example, have a cortex
consisting of a dense array of longitudinal epicytic folds, while some species exhibit
further epicyte modifications such as superfolds, fusion of the plasma membrane at
the apex or a lateral site of several folds, numerous cytopilia or microvilli covering
the surface, or rows of small knobs. All these modifications lead to an increase in
gregarine surface in contact with the host environment. Bacteria found in the canals
between partially fused folds may induce their fusion (Desportes and Schrével, 2013;
Desportes et al., 1977; Diakin et al., 2017). A glycocalyx layer covering the pellicle
has been documented in a number of species (Kováčiková et al., 2017b; Philippe et
al., 1979). Most eugregarines exhibit progressive gliding motility, which is described
as an irregular, erratic process, usually corresponding to a unidirectional movement
with respect to the antero-posterior cell axis, without changes in cell shape
(Desportes and Schrével, 2013; Diakin et al., 2016, 2017; King and Sleep, 2005). As
trophozoites and gamonts in the species analysed lack subpellicular microtubules,
they must use a gliding mechanism differing from glideosome (Kováčiková et al.,
2017b, 2018; Valigurová et al., 2013). Further, eugregarines are able to free-float in
liquid, lacking any contact with the substrate, but they move at a significantly higher
rate than shown during regular gliding (Valigurová et al., 2013). Undulations of the
epicytic folds can be seen in floating individuals (Desportes and Schrével, 2013).
Eugregarine trophozoites attached to the host tissue usually show no obvious
motility, though near-surface currents of host liquids, most likely produced by the
parasite’s undulating folds, can be noticed around them (Diakin et al., 2016).
Gliding on a solid surface is accompanied by the shedding of a mucous trail
behind the gregarine. Though the material in the trail is generally designated as
mucus, its exact composition is unknown. The origin of this mucosal substance could
be related to an abundant Golgi apparatus present in the cytoplasm of eugregarines.
It has previously been suggested that the secreted mucus pushes the gregarine
forward passively (Desportes and Schrével, 2013). However, the translocation of
concanavalin-A beads posteriorly along the parasite surface at rates similar to the
forward gliding movement, and the fact that large beads comparable in mass to the
gregarine itself can be actively translocated, indicates that eugregarines generate
substantial locomotory forces (King, 1981). Hence, it is more likely that the mucus
acts as a lubricant. The increased load of mucus in eugregarine cytoplasm is
42
correlated with gliding rate, in contrast to the general architecture and
supramolecular organisation of the pellicle (Valigurová et al., 2013). More recently,
it was suggested that the eugregarine gliding mechanism involves a mechanicochemical
system related to lateral undulations of the epicytic folds, frequently
documented with undulated and straight folds alternating (Desportes and Schrével,
2013). Generally, the epicytic folds have identical ectoplasmic structures and are
built up from the plasma membrane with IMC, 12-nm filaments with properties of
intermediate filaments, rippled dense structures and basal lamina (Schrével et al.,
1983; Valigurová and Koudela, 2008; Valigurová et al., 2013; Vávra and Small, 1969;
Walker et al., 1984). The number of 12-nm filaments does not influence gliding speed
but appears to control the direction of movement. Gregarines equipped with epicytic
folds bearing a lower number of 12-nm filaments glide at a relatively high speed,
though their gliding path tends to be widely semi-circular rather than linear. The
rippled dense structures, located at the external cytomembrane, presumably serve
as supporting elements interconnecting 12-nm filaments and the plasma membrane.
The half-moon-shaped dense structure often seen to underline the 12-nm filaments
could function as a skeleton reinforcing the tips of folds directly contacting the
substrate during gliding (Kováčiková et al., 2017b; Valigurová et al., 2013).
Though involvement of actin and myosin (associated with the eugregarine
pellicle) in eugregarine motility has been reported, the actual gliding motor remains
unknown (Baines and King, 1989; Ghazali et al., 1989; Ghazali and Schrével, 1993;
Heintzelman, 2004; Heintzelman and Mateer, 2008; Kováčiková et al., 2017b;
Valigurová et al., 2013). Positive phalloidin labelling suggests that the majority of
actin in eugregarines is present in a filamentous form and could play a role in gliding
(Kováčiková et al., 2017b, 2018; Valigurová et al., 2009, 2013). The actin accumulates
in two layers of eugregarine cell cortex: the longitudinal epicytic folds and a deeper
layer of rib-like myonemes running perpendicularly to the cell axis (Heintzelman,
2004; Heintzelman and Mateer, 2008; Kováčiková et al., 2018). The myonemes,
localised at the border between the ectoplasm and endoplasm, correspond to the
concentric bundles of F-actin (Fig. 10). A further element most likely involved in
motility is the anastomosing ectoplasmic network of intermediate filamentous
proteins localised immediately beneath the epicytic folds (Beams et al., 1959;
Kováčiková et al., 2018; Valigurová et al., 2013). Of the three myosin genes
43
characterised in G. polymorpha, myosins A (93 kDa) and B (96 kDa) belong to class
XIV, restricted to the Apicomplexa, and myosin F (222 kDa) to class XII. Myosin A
localises on the longitudinal epicytic folds and subjacent rib-like myonemes, while
myosin B is restricted to the folds and myosin F to the myonemes (Heintzelman and
Mateer, 2008). If some form of actomyosin motor is present, it could be localised at
the links observed between the plasma membrane and the external cytomembrane
of the IMC, which are supposed to be related to the lateral undulations of the folds
(Desportes and Schrével, 2013; Valigurová et al., 2013). However, an experimental
study on G. garnhami, using actin-targeting drugs, contradicts the expectation that
lateral undulation of the epicytic folds provides the force behind gregarine gliding as
the wavy pattern did not change in drug-treated parasites where motility was
completely blocked (Kováčiková et al., 2018). Observations on drug-treated gamonts
demonstrate that the organisation and density of the subpellicular structures,
ectoplasmic network and myonemes, change in accordance with gliding activity.
Degradation of myonemes accompanies the blocking of gliding motility in
cytochalasin D-treated gamonts due to the depolymerisation of existing actin
filaments, while jasplakinolide-induced changes lead to further stabilisation of
already present F-actin and do not significantly affect gliding. These data suggest that
the dynamic process of actin polymerisation and rapid depolymerisation described
in glideosome is not essential for gliding in eugregarines, where polymerised actin
alone appears to be the main motor structure responsible for gliding.
Fig. 10. Schematic diagram illustrating a possible function of cortical filaments
(myonemes, ectoplasmic network) facilitating gliding motility in Gregarina
garnhami gamonts (taken from Kováčiková et al., 2018). A. Gamont in non-gliding
phase. The myonemes and the ectoplasmic network are evenly distributed around
the ectoplasm. B. Gamont during gliding on a solid substrate. Note the presence of
protruding superfolds that group together, resulting from contraction of myonemes,
along with more a compact ectoplasmic network and denser accumulation of mucus
in this region. black arrow – mucus drops, ecn – ectoplasmic network, s – superfolds,
white arrowhead – myonemes, white asterisk – cytoplasm.
44
Besides gliding, some species exhibit intense bending, curving or shortening of
the longitudinal axis, especially in their protomerite region. For example, the
trophozoites and gamonts of C. cf. communis enrich their gliding by jumping and
rotational movements, with rapid changes in gliding direction and cell flexions
(Kováčiková et al., 2017b). While it is generally believed that the eugregarine cortex
became rigid over the course of environmental adaptation, resulting in a loss of
wriggling ability (typical for archigregarines), these active movements indicate a
relatively high cellular plasticity. The rib-like myonemes girding the cell cortex and
the ectoplasmic network are thought to be involved in the bending motions of
eugregarines (Beams et al., 1959; Valigurová et al., 2013).
Coelomic eugregarines, which evolved as free-floating parasites within the host
tissue, move by pulsations of the body wall (corresponding to non-progressive
peristaltic or metabolic motility) accompanied by periodic changes in body shape
(Diakin et al., 2016; Landers, 2001; Leander et al., 2006; MacMillan, 1973a; Miles,
1966, 1968). These peristaltic waves, which occur about every two seconds, move
backward and forward, the gregarines being driven in spiral-like movements during
wave propagation (Desportes and Schrével, 2013). The annular myonemes are
considered responsible for these peristaltic movements. Abundant mitochondria
distributed uniformly in the cell cytoplasm of U. ovalis may be correlated with active
metaboly, which can be accompanied by transient modifications of the cell cortex,
such as formation of superfolds in the contracted regions (Diakin et al., 2016;
MacMillan, 1973a). The cellular deformations demonstrated by urosporid gamonts
pulsating freely within host fluids may be necessary to provoke an exchange of host
liquid around them, thereby improving the effectiveness of nutrient acquisition
(Leander, 2008). Active movements could also provide an effective protection
against adhesion of host coelomocytes to their surfaces (Diakin et al., 2016).
Urosporid eugregarines represent a perfect example of Apicomplexa exhibiting
enormous morphological and behavioural plasticity according to their localisation
within the host. Phylogenetically related species that occupy different ecological
niches in the host to decrease the intensity of species competition, often demonstrate
diverse parasitism strategies (Diakin et al., 2016; Wakeman et al., 2014). The
diversity of urosporids demonstrates an intermediate character state related to the
evolutionary transformation of trophozoites following colonisation of coelomic
45
environments. As an example, two closely related coelomic Urospora species
parasitising the polychaete T. forbesii exhibit different modes of motility. The
trophozoites of U. ovalis, located free in the host coelom, show metaboly, while the
V-shaped U. travisiae trophozoites, which are able to attach to the host tissue,
demonstrate gliding (Diakin et al., 2016). The gliding motility of U. travisiae
presumably allows the parasite to move along the inner coelomic wall and other host
tissues, as shown in other urosporids (Leander, 2008).
4.4 Motility and cytoskeleton in protococcidia and agamococcidia
In the protococcidian E. duboscqi, subpellicular microtubules, connected with
posterior polar ring, are only present during its early development (Valigurová et al.,
2015). Accordingly, in native preparations, the drop-shaped invading zoites and
earliest developmental stages, with their pointed end attached to the host
epithelium, exhibit oscillating movements, while helmet-shaped early trophozoites
show only weak movement. Despite the absence of microtubules in maturing
trophozoites and later stages of E. duboscqi, positive α-tubulin labelling of the
parasite surface and cytoplasm suggests either preservation of tubulin in a nonpolymerised
form or its presence in other tubulin-rich structures. The second option
is more likely, considering the disappearance of labelling from the parasite periphery
after treatment with oryzalin, resulting in dispersion of putatively unpolymerised αtubulin
throughout the cytoplasm. Instead of microtubules, subpellicular bands of
longitudinally oriented actin-rich filaments form beneath the IMC during
trophozoite maturation, these presumably functioning as the parasite’s
cytoskeleton. Aside from the frequent detachment of parasites along with their
parasitophorous sacs from host tissue observed in native preparations, neither
attached nor detached mature E. duboscqi stages exhibit obvious movement. In
squash preparations, some attached advanced stages exhibit slight signs of
movements and cytoplasmic streaming resembling metaboly; however, due to the
intense waving and beating motion of host enterocyte cilia it is not possible to
determine with certainty whether this is actual movement of the parasite inside the
sac (Valigurová et al., 2015; Valigurová, personal observation).
After being picked out mechanically from the host tissue, agamococcidian
Rhytidocystis spp. transform into oval or round immotile cells. The trophozoites are
46
covered by a trimembrane pellicle forming short folds, with numerous micropores
located between and on top of the folds (Diakin and Valigurová, 2014; Rueckert and
Leander, 2009b). All observed intracellular stages lack the subpellicular
microtubules, though a single microtubule located deeper within the cytoplasm can
occasionally be seen in ultrathin sections (Valigurová and Diakin, personal
observation).
4.5 Motility and cytoskeleton in cryptosporidia
While motility of apicomplexan zoites is considered the essential mechanism for
host cell invasion, motility of C. proliferans sporozoites freshly excysted from oocysts
is rather limited and featureless in a range of excystation media (Melicherová et al.,
2016). In cell cultures, however, released sporozoites can be seen probing the
plasma membrane of potential host cells with their prolonged apical end and
attempting to invade the cultured cells (Melicherová et al., 2018). This thin and
prolonged apical end is only typical for sporozoites contacting the host cell, as
sporozoites isolated from the supernatant do not exhibit apical prolongation
(Borowski et al., 2010; Melicherová et al., 2018). This behaviour appears to be
important for invasion success as cryptosporidian sporozoites are only equipped
with a single rhoptry and, therefore, have only a single attempt for successful
attachment to suitable host cell (O’Hara et al., 2005). The sporozoite of C. parvum
shows a uniform distribution of actin throughout the cell and its gliding motility
depends upon an intact actomyosin motor (Forney et al., 1998). Using the anti-actin
antibody raised against Dictyostelium, which recognises the actin in T. gondii and
Plasmodium, Western Blot analysis of C. proliferans sporozoites has confirmed the
presence of actin (42 kDa) at very low concentrations, despite negative
immunofluorescent labelling. Immunofluorescence has also detected
homogeneously distributed myosin (Mazourová, 2013, 2015; Valigurová et al.,
2014). The subpellicular microtubules of cryptosporidian sporozoites originate at
the region of the dense collar with associated apical rings and appear to spiral
around the apical region (O'Hara et al., 2005). In C. proliferans, Western Blot analysis
has revealed the presence of high consternations of α-tubulin (50 kDa), while
immunofluorescent labelling has confirmed presence of subpellicular microtubules
extending from the apical pole for up to two thirds of the sporozoite body, as shown
47
in other apicomplexan zoites (Mazourová, 2013, 2015; Morrissette and Sibley,
2002). In merozoites, the subpellicular microtubules are present in C. muris and C.
proliferans (Melicherová et al., 2014; Uni et al., 1987), while studies focusing on C.
parvum and C. wrairi failed to detected them (Current and Reese, 1986; Vetterling et
al., 1971). The trails of gliding C. parvum sporozoites are threadlike and relatively
short (typically 1-2 times the sporozoite length), with a straight to slightly curved
pattern extending from the posterior end of the parasite. The sporozoites of C.
proliferans excysted in vitro in various incubation media exhibit an oscillating
movement (short forward/backward shifts), appearing to move progressively
forward without obvious changes in cell shape (Melicherová et al., 2016). Motility of
sporozoites is lower in pure RPMI 1640 and RPMI 1640 enriched by 1 % BSA
compared to media with higher concentrations of BSA (5% and 10 %). In the medium
with 5 % BSA, sporozoites remain motile for a long period (tested over 240 minutes),
while in 10 % BSA the activity of excysted sporozoites decreases slightly. Sporozoites
in BSA free medium are most active during the first 30 minutes after excystation,
though motility rapidly declines thereafter. The motility of unexcysted sporozoites
appears to create ‘dancing’ oocysts prior to their final excystation and the liberation
of sporozoites (Melicherová et al., 2016). Frequently observed is the positioning of
unexcysted oocysts with their sutures orientated towards the cultured cell’s surface,
presumably resulting from sporozoite movement inside the oocyst. Such behaviour
may help to shorten the distance between free sporozoites and the host cells
(Melicherová et al., 2018). The enclosure of oocysts by cultured cells, as observed in
HCT-8 and HT-29 cell lines, is induced by parasite antigens (Melicherová et al., 2018).
This encapsulation occurs independently of any active invasion by motile stages and
concurs with Forney et al. (1998), who stated that, in contrast to other apicomplexan
zoites, invasion of cryptosporidian zoites is a passive process that does not require
actomyosin motility machinery as the effect of treatment with actin and myosin
inhibitors on infectivity was insignificant. In contrast, antimicrotubular drugs block
cryptosporidian infectivity for host cells (Wiest et al., 1993). Irrespective of their
structural similarities, it is evident that cryptosporidia have evolved strategies for
host cell invasion that differ significantly from those described in phylogenetically
related apicomplexan parasites.
48
5 Conclusions
Apicomplexans cause human and animal diseases that represent a major world
health problem and have a considerable impact on the global economy. Attempts to
control these pathogens are usually hampered by their localisation and strategies to
evade host immune responses and chemotherapeutics. Whilst the nature of the
diseases caused by Apicomplexa differs significantly, their common origin led them
to share specific metabolic pathways unique to this group, and these may constitute
potential targets for intervention. In contrast to well-studied vertebrate pathogens,
Apicomplexa restricted to invertebrates are considered of no economic or medical
significance and they remain poorly understood, despite their enormous diversity.
In summarising the publications produced for this thesis, as well as those closely
related to the topic, I have tried to highlight the diversity of apicomplexan survival
and parasitism strategies occurring in different environments. Two main aspects are
discussed in detail, i.e. host-parasite interactions and parasite motility, as these
represent a potential target for chemotherapeutic intervention. The study has
attempted to determine which structures and mechanisms are responsible for the
various modes of motility, attachment to host cell/tissue and nutrient acquisition in
basal lineages and how these are modified in comparison to other Apicomplexa.
Detailed observations indicate that even seemingly insignificant modifications in
apicomplexan subcellular structures may result in different parasitism strategies.
The presented data, especially those obtained on type species such as the
protococcidian E. duboscqi, blastogregarine S. nematoides and archigregarine S.
pendula, indicate the importance of further research on deep-branched
apicomplexans, which exhibit enormous diversity in subcellular organisation and
highly specialised adaptations to the parasitic life style. Such research will provide a
deeper understanding of the biology and evolutionary pathways of the Apicomplexa
in general. The results presented herein support the hypothesis of Apicomplexa
evolution progressing from myzocytotic predation to myzocytotic extracellular
parasitism, accompanied by the origination of epicellular parasitism and significant
modifications to the attachment apparatus and motility mode at the trophozoite
stage, and finally intracellular parasitism along with a rejection of trophozoite
polarity and motility.
49
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7 List of publications included in the thesis
7.1 Valigurová A. (2012). Sophisticated adaptations of Gregarina cuneata
(Apicomplexa) feeding stages for epicellular parasitism. PLoS One 7(8),
e42606.
7.2 Valigurová A., Vaškovicová V., Musilová N., Schrével J. (2013). The enigma of
eugregarine epicytic folds: where gliding motility originates? Frontiers in
Zoology 10, 57.
7.3 Melicherová J., Ilgová J., Kváč M., Sak B., Koudela B., Valigurová A. (2014). Life
cycle of Cryptosporidium muris in two rodents with different responses to
parasitization. Parasitology 141(2), 287-303.
7.4 Valigurová A., Paskerova G.G., Diakin A., Kováčiková M., Simdyanov T.G. (2015).
Protococcidian Eleutheroschizon duboscqi, an unusual apicomplexan
interconnecting gregarines and cryptosporidia. PLoS One 10(4), e0125063.
7.5 Melicherová J., Mazourová V., Valigurová A. (2016). In vitro excystation of
Cryptosporidium muris oocysts and viability of released sporozoites in different
incubation media. Parasitology Research 115(3), 1113-1121.
7.6 Diakin A., Paskerova G.G., Simdyanov T.G., Aleoshin V.V., Valigurová A. (2016).
Morphology and molecular phylogeny of coelomic gregarines (Apicomplexa)
with different types of motility: Urospora ovalis and U. travisiae from the
polychaete Travisia forbesii. Protist 167(3), 279-301.
7.7 Schrével J., Valigurová A., Prensier G., Chambouvet A., Florent I., Guillou L.
(2016). Ultrastructure of Selenidium pendula, the type species of
archigregarines, and phylogenetic relations to other marine Apicomplexa.
Protist 167(4), 339-368.
7.8 Chambouvet A., Valigurová A., Pinheiro L.M., Richards T.A., Jirků M. (2016).
Nematopsis temporariae (Gregarinasina, Apicomplexa, Alveolata) is an
intracellular infectious agent of tadpole livers. Environmental Microbiology
Reports 8(5), 675-679.
7.9 Diakin A., Wakeman K.C., Valigurová A. (2017). Description of Ganymedes yurii
sp. n. (Ganymedidae), a new gregarine species from the Antarctic amphipod
Gondogeneia sp. (Crustacea). Journal of Eukaryotic Microbiology 64(1), 56-66.
59
7.10 Kováčiková M., Simdyanov T.G., Diakin A., Valigurová A. (2017). Structures
related to attachment and motility in the marine eugregarine Cephaloidophora
cf. communis (Apicomplexa). European Journal of Protistology 59, 1-13.
7.11 Valigurová A., Vaškovicová N., Diakin A., Paskerova G.G., Simdyanov T.G.,
Kováčiková M. (2017). Motility in blastogregarines (Apicomplexa): Native and
drug-induced organisation of Siedleckia nematoides cytoskeletal elements.
PLoS One 12(6), e0179709.
7.12 Melicherová J., Hofmannová L., Valigurová A. (2018). Response of cell lines to
actual and simulated inoculation with Cryptosporidium proliferans. European
Journal of Protistology 62, 101-121.
7.13 Valigurová A., Pecková R., Doležal K., Sak B., Květoňová D., Kváč M., Nurcahyo
W., Foitová I. (2018). Limitations in the screening of potentially anticryptosporidial
agents using laboratory rodents with gastric cryptosporidiosis.
Folia Parasitologica 65, 010.
7.14 Kováčiková M., Vaškovicová N., Nebesářová J., Valigurová A. (2018). Effect of
jasplakinolide and cytochalasin D on cortical elements involved in the gliding
motility of the eugregarine Gregarina garnhami (Apicomplexa). European
Journal of Protistology 66, 97-114.
7.15 Simdyanov T.G., Paskerova G.G., Valigurová A., Diakin A., Kováčiková M.,
Schrével J., Guillou L., Dobrovolskij A.A., Aleoshin V.V. (2018). First
ultrastructural and molecular phylogenetic evidence from the
blastogregarines, an early branching lineage of plesiomorphic Apicomplexa.
Protist 169(5), 697-726.
7.16 Paskerova G.G., Miroliubova T.S., Diakin A., Kováčiková M., Valigurová A.,
Guillou L., Aleoshin V.V., Simdyanov T.G. (2018). Fine structure and molecular
phylogenetic position of two marine gregarines, Selenidium pygospionis sp. n.
and S. pherusae sp. n., with notes on the phylogeny of Archigregarinida
(Apicomplexa). Protist 169(6), 826-852.
60
8 Other publications related to the habilitation thesis
This list includes conference abstracts with results that are part of habilitation
thesis, but have not yet been published in the form of research papers.
Diakin A., Valigurová A. (2014). Development of new species of agamococcidian
Rhytidocystis sp. from Travisia forbesii. In: A variety of interactions in the marine
environment. Abstracts volume from 49th European Marine Biology Symposium,
Zoological Institute Russian Academy of Sciences, St. Petersburg, Russia, pp. 83-
84.
Diakin A., Kováčiková, M., Valigurová A. (2014). Morphology of gametocyst, oocyst
and sporozoites of extraordinary coelomic gregarine Urospora travisiae from
marine polychaete. In: A variety of interactions in the marine environment.
Abstracts volume from 49th European Marine Biology Symposium, Zoological
Institute Russian Academy of Sciences, St. Petersburg, Russia, pp. 59-60.
Valigurová A., Florent I., Mazourová V., Melicherová J. (2014). Cell motility in
sporozoites of Cryptosporidium muris. In: 5th International Giardia and
Cryptosporidium Conference, Uppsala, Sweden, p. 186.
Kováčiková M., Paskerova G.G., Diakin A., Valigurová A. (2017). Cytoskeletal elements
and motility in the archigregarine Selenidium sp.: observations on native and
drug-treated parasites. In: 15th International Congress of Protistology: Book of
Abstracts, Prague, Czech Republic, p. 113.
7.1
Valigurová A.
2012
Sophisticated adaptations of Gregarina cuneata (Apicomplexa)
feeding stages for epicellular parasitism
PLoS One 7(8), e42606
Sophisticated Adaptations of Gregarina cuneata
(Apicomplexa) Feeding Stages for Epicellular Parasitism
Andrea Valigurova´*
Department of Botany and Zoology, Faculty of Science, Masaryk University, Brno, Czech Republic
Abstract
Background: Gregarines represent a very diverse group of early emerging apicomplexans, parasitising numerous
invertebrates and urochordates, and are considered of little practical significance. Recently, they have gained more
attention since some analyses showed that cryptosporidia are more closely related to the gregarines than to coccidia.
Methodology/Principal Findings: Using a combined microscopic approach, this study points out the spectacular strategy
of Gregarina cuneata for attachment to host tissue and nutrient acquisition while parasitising the intestine of yellow
mealworm larvae, and reveals the unusual dynamics of cellular interactions between the host epithelium and parasite
feeding stages. Trophozoites of G. cuneata develop epicellularly, attached to the luminal side of the host epithelial cell by an
epimerite exhibiting a high degree of morphological variability. The presence of contractile elements in the apical region of
feeding stages indicates that trophozoite detachment from host tissue is an active process self-regulated by the parasite. A
detailed discussion is provided on the possibility of reversible retraction and protraction of the eugregarine apical end,
facilitating eventual reattachment to another host cell in better physiological conditions. The gamonts, found in contact
with host tissue via a modified protomerite top, indicate further adaptation of parasite for nutrient acquisition via epicellular
parasitism while keeping their host healthy. The presence of eugregarines in mealworm larvae even seems to increase the
host growth rate and to reduce the death rate despite often heavy parasitisation.
Conclusions/Significance: Improved knowledge about the formation of host-parasite interactions in deep-branching
apicomplexans, including gregarines, would offer significant insights into the fascinating biology and evolutionary strategy
of Apicomplexa. Gregarines exhibit an enormous diversity in cell architecture and dimensions, depending on their parasitic
strategy and the surrounding environment. They seem to be a perfect example of a coevolution between a group of
parasites and their hosts.
Citation: Valigurova´ A (2012) Sophisticated Adaptations of Gregarina cuneata (Apicomplexa) Feeding Stages for Epicellular Parasitism. PLoS ONE 7(8): e42606.
doi:10.1371/journal.pone.0042606
Editor: Ross Frederick Waller, University of Melbourne, Australia
Received April 30, 2012; Accepted July 10, 2012; Published August 10, 2012
Copyright: ß 2012 Andrea Valigurova´. This is an open-access article distributed under the terms of the Creative Commons Attribution License, which permits
unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited.
Funding: Financial support was provided by grants from the Czech Science Foundation (GPP506/10/P372). Travel expenses were partially supported by the
Ministry of Education, Youth and Sports (MEB021127). The author acknowledges support from the Department of Botany and Zoology, Faculty of Science,
Masaryk University, towards the preparation of this manuscript. The funders had no role in study design, data collection and analysis, decision to publish, or
preparation of the manuscript.
Competing Interests: The author has declared that no competing interests exist.
* E-mail: andreav@sci.muni.cz
Introduction
The alveolates (Alveolata), a major line of protists, include three
extremely diverse groups of unicellular eukaryotes: ciliates,
dinoflagellates and apicomplexans. Gregarines belong to the
phylum Apicomplexa Levine, 1970, a large group characterised
by the presence of a unique organelle called an apical complex,
and which consists entirely of parasitic genera that infect a wide
spectrum of invertebrates and vertebrates. Many of these are
intensively studied etiologic agents of globally significant human
disease, including malaria, toxoplasmosis and cryptosporidiosis. In
contrast, gregarines are restricted to the internal organs and
coelom of invertebrates and urochordates, and recently have been
classified into three orders: Archigregarinorida Grasse´, 1953;
Eugregarinorida Le´ger, 1900; and Neogregarinorida Grasse´, 1953
[1]. They are considered of no economic or medical significance
and thus, despite their enormous diversity, the general biology of
gregarines remains poorly understood. Recent phylogenetic
analyses, however, have pointed out their close affinity with
Cryptosporidium, and have drawn attention to this enigmatic group
[2,3].
Apicomplexans exhibit very specific adaptations for invading
and surviving within their hosts, which have evolved under distinct
evolutionary pressures, resulting in diverse attachment strategies
and host-parasite interactions. In general, gregarines exhibit
several known strategies for attachment to the host tissue: (i)
intracellular or intratissular localisation with or without a reduced
area of attachment in neogregarines; (ii) a mucron in archigregarines,
monocystid eugregarines and some neogregarines; (iii) a
simple epimerite in eugregarines and a few neogregarines; (iv) a
complicated epimerite equipped with various structures, e.g.
digitations, hooks or spines, hairs in eugregarines; (v) a suckerlike
protomerite or modified protomerite with rhizoids in
eugregarines. Eugregarines, similarly to cryptosporidia, are specific
with their unique epicellular localisation [4,5,6,7,8]. Their
sporozoites usually invade epithelial cells; however, some species
PLOS ONE | www.plosone.org 1 August 2012 | Volume 7 | Issue 8 | e42606
are able to invade even the intercellular space. As the majority of
eugregarines do not exhibit intracellular development, sporozoites
generally develop into large extracellular vegetative stages, called
trophozoites, exhibiting a high degree of cell polarity in that they
possess an anterior part specialised for attachment to the host cell
in general [9]. In intestinal species, the development of the
trophozoite starts after sporozoite interaction with the microvillus
border of the host epithelium, when apical organelles disappear
and an epimeritic bud derived from the conoid forms at the apical
end [4,10]. The epimeritic bud gradually transforms into a
specialised structure called the epimerite, which serves to anchor
the parasite firmly to the host cell [4,8,10]. It is already known that
epimerites of eugregarines parasitising herbivore hosts are usually
simple button-shaped; however, they are much more complicated
in carnivorous hosts, equipped with strong hooks, spines or
numerous filaments [9].
The mechanism of nutrition acquisition in gregarines, however,
is still poorly understood. Some authors attribute feeding function
to the attachment organelles, such as the epimerite or mucron [9].
The higher concentration of host cell mitochondria and
endoplasmic reticulum surrounding the epimerite, and the
presence of mitochondria under the epimeritic cortical vesicle,
indicate the existence of an active interaction between the
gregarine epimerite and the host cell [11,12]. Furthermore, the
presence of organelles associated with nutritive function suggests
that the epimerite is a metabolically active organelle [13,14,15].
Some eugregarine species are equipped with additional structures
located in the grooves between the epicytic folds covering the
gregarine body, resembling the micropores (diminished cell
mouth) reported in other apicomplexans [11,16,17]. Thus,
questions arise as to whether and under what circumstances the
epimerite serves as a feeding organelle, and whether the
micropore-like structures in gregarines are points implicated in
pinocytosis [18] or are exclusively dedicated to mucus extrusion
[19]. Similarly, the exact mechanisms responsible for trophozoite
attachment to the host cell and for abandoning the host tissue at
the end of development still remain enigmatic. There are two
contradictory hypotheses on gregarine detachment from host
tissue at the end of the trophozoite stage. One of them describes
trophozoite detachment via epimerite retraction, self-regulated by
the vegetative stage [8], while the other is based on gradual
epimerite constriction facilitated by the supposed contractility of
an osmiophilic ring surrounding the base of the epimerite and
acting as a sphincter during the separation of the epimerite from
the rest of the gregarine body [9,10,20,21,22]. All of these
questions raised by conflicting data must be satisfactorily answered
to clarify the parasitic strategies of gregarines and to better
understand the evolutionary history of the phylum Apicomplexa.
This study endeavours to address the questions set out above
and aims provide a new insight into the dynamics and architecture
of the attachment site of G. cuneata. Unique relationships with the
host epithelium, not only in trophozoites but also in more
advanced developmental stages including gamonts, are described
herein. Though there are few published works dealing with the life
cycle and host specificity of G. cuneata [23,24,25], complete data on
its early development and host-parasite interactions at the cellular
level are still lacking. Based on personal observations of four
eugregarine species (Gregarina cuneata, G. polymorpha, G. steini and G.
niphandrodes) parasitising the yellow mealworm beetle Tenebrio
molitor, in many aspects, G. cuneata appears to be the most
spectacular of them all. Conclusions are supported by identification
and detailed descriptions of structures involved in the
formation of host-parasite interactions using a combined microscopic
approach.
Materials and Methods
Larvae of the yellow mealworm, Tenebrio molitor Linnaeus, 1758
(Coleoptera, Tenebrionidae) with eugregarine infection were
obtained from colonies maintained in our laboratory. Gametocysts
of Gregarina cuneata were collected from the faeces of infected larvae
and placed in moist chambers at 25uC for maturation and
dehiscence. Larvae sterilised of eugregarines were allowed to feed
for 24 h on flour contaminated with the oocysts of G. cuneata, and
were subsequently maintained on a sterile substrate. Insects were
anesthetised with cold and dissected at different time points after
feeding with eugregarine oocysts. Squash and/or wet smear
preparations were investigated with the use of an Olympus BX51
light microscope.
For observations on living gregarines, different solutions,
including phosphate buffered saline, Insect Ringer’s solution or
Minimum Essential Medium [3% bovine foetal serum with
penicillin, streptomycin, amphotericin B and L-glutamine], were
used to prepare squash preparations.
Transmission electron microscopy
Parasitised intestines were fixed overnight at 4uC in freshly
prepared 2.78% glutaraldehyde in 0.2 M phosphate buffer for
transmission electron microscopy. The specimens were then
washed for 1 h in phosphate buffer (pH 7.0), post-fixed in 1%
osmium tetroxide in the same buffer for 3 h and dehydrated in an
alcohol series, before embedding in Epon (Polybed 812). Sections
were cut with glass knives and stained with uranyl acetate and lead
citrate. Procedures for freeze etching follow Schrevel et al. [26]
using the BAL-TEC BAF 060 freeze-etching system. Observations
were made using a JEOL 1010 TEM.
Scanning electron microscopy
Specimens were fixed overnight at 4uC in freshly prepared 3%
glutaraldehyde in 0.2 M cacodylate buffer (pH 7.4), washed
3615 min in cacodylate buffer, post-fixed in 2% osmium tetroxide
in cacodylate buffer (pH 7.4) for 2 h at room temperature and
finally washed 3615 min in the same buffer. After dehydration in
a graded series of acetone, specimens were critical point-dried
using CO2, coated with gold and examined using a JEOL JSM-
7401F field emission scanning microscope.
Fluorescence microscopy
The G. cuneata cell suspension was washed in 0.2 M phosphate
buffered saline (PBS), fixed for 15 min at room temperature in 4%
paraformaldehyde in 0.2 M PBS, washed again, and permeabilised
for 10 min in 0.1% Triton X-100 (Sigma-Aldrich). For direct
fluorescence, samples were washed for 2 h in the antibody diluent
(0.1% bovine serum albumin, 0.5% Triton X-100 and 0.1%
sodium azide in 0.1 M PBS), incubated for 2 h at room
temperature with fluorescein isothiocyanate (FITC)-phalloidin
(Sigma-Aldrich) and then washed again in antibody diluent.
Preparations were mounted in anti-fade mounting medium based
on 2.5% DABCO (Sigma-Aldrich) mixed with glycerol and 0.1 M
PBS. For indirect immunofluorescence, samples were incubated
for 2 h at room temperature in rabbit anti-myosin antibody
(smooth and skeletal, whole antiserum from Sigma-Aldrich;
dilution 1:5) or in mouse monoclonal IgG anti-actin antibody
raised against Dictyostelium actin that recognises Toxoplasma and
Plasmodium actin (provided by Prof. Dominique Soldati-Favre)
diluted in PBS with 0.1% BSA (dilution 1:500), washed three times
in PBS for 10 min and incubated with FITC-conjugated antirabbit
IgG (dilution 1:40) or anti-mouse polyvalent immunoglobulins
(1:125) in PBS with 1% BSA at 37uC for 1 h. After washing
Epicellular Parasitism in Eugregarines
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in PBS, preparations were counterstained with Evans blue (1:5000)
and mounted. Controls were labelled with FITC-conjugated
secondary antibody alone without the primary antibody. Preparations
were observed and documented using an Olympus BX60
fluorescence microscope fitted with a WB filter cube, a fully
motorized inverse epi-fluorescence microscope Olympus IX 81
equipped with Cell‘R imaging station or an Olympus IX80
microscope equipped with a laser-scanning FluoView 500 confocal
unit (Olympus FluoView 4.3 software).
Results
Feeding stages of Gregarina cuneata under light
microscopy
All vegetative stages of Gregarina cuneata exhibited epicellular
development, i.e. sporozoites and trophozoites developed attached
to microvillous sites of host epithelial cells; no developmental stage
was observed penetrating under the host cell plasma membrane.
When observed under the light microscope, it was difficult or even
impossible to detect the earliest stages, such as sporozoites
transforming into the trophozoites, and very young two-segmented
trophozoites. These stages were small and inconspicuous
(Figure 1A), and seemingly it was quite impossible to detach them
from the host tissue without any damage. The only observed
earliest trophozoite stages exhibited an irregular triangular shape,
tapering towards their apical part with a polymorphous epimerite
(Figure 1A - upper micrographs). The irregular shape of
epimerites, as shown in these micrographs, seemed to be the
consequence of mechanical damage due to the forced separation
of the gregarine from the host epithelium during the processing of
squash preparations. Three-segmented stages were irregularly
shaped, with a cylindrical deutomerite widest at its posterior
rounded end (Figure 1A - lower micrographs). Only few maturing
detached trophozoites, with an apparently non-damaged epimerite
still located on a relatively short protomerite and typical cylindrical
deutomerite, were observed in squash preparations (Figure 1B).
Their epimerites were usually conical to lance-shaped or
prolonged of irregular shape. More often, however, were
trophozoites released from the host epithelium and still bearing
the affected host cell on their epimerites (Figure 1C) or
trophozoites with ruptured epimerites (Figure 1D). The most
frequently observed stages were small-sized hyaline gamont-like
individuals (in size comparable to trophozoites) typical by a
cylindrical protomerite, which was constricted at the septum and
widely rounded at its apical top, and a prolonged cylindrical
deutomerite widest at the posterior end (Figure 1E). These stages
lacked an obvious epimerite. Gamonts of G. cuneata formed socalled
early syzygies and thus only few non-associated gamonts
could be found. This means that mature individuals transforming
into gamonts (satellites) joined to individuals still attached to the
host cell (future primites). Living (non-fixed) gamonts, either single
or associated in syzygies, exhibited a prolonged cylindrical
protomerite with a widely rounded top and cylindrical deutomerite
(Figures 1F - left micrograph, 1G). The protomerite of
paraformaldehyde fixed primites or single gamonts, however,
showed a lance-shape hyaline apical top (Figures 1F - right
micrograph, 1H). Light microscopic observations confirmed that
the number of amylopectin granules increased with the age of the
trophozoites and the cytoplasm of mature gamonts was fully
packed with amylopectin, except in the region of the lance-shaped
protomerite top in gamonts (Figures 1A–H).
Localisation of actin and myosin
The homogenous distribution of the fluorescence signal
throughout the surface of FITC-phalloidin labelled trophozoites
and gamonts (Figure 1I) corresponded to the localisation of
filamentous actin (F-actin) associated with the typical apicomplexan
cell cortex. Labelling also confirmed the presence of F-actin
in the peripheral region of growing epimerites. In all individuals,
the fluorescence signal was more evident in the area of fibrillar
septum separating the protomerite from the deutomerite. In
addition, a large circular area in the deutomerite, suggestive of a
nucleus, exhibited a more intense signal. In maturing single
gamonts, the cytoplasm of protomerite exhibited more evident
labelling of F-actin than that of deutomerite (Figure 1I), and it
corresponded to the dot-like pattern of actin labelling restricted to
the protomerite cytoplasm in individuals stained with the specific
anti-actin antibody (proved to recognize the actin in Toxoplasma
and Plasmodium) (Figure 1J). In comparison with the phalloidinstained
specimens, however, the gregarine cell cortex and septum
exhibited only slight labelling of actin when stained with this
antibody (Figure 1J).
Indirect immunofluorescence using rabbit anti-myosin antibody
revealed the presence of myosin restricted to the epimerite region
in trophozoites (Figure 1K) as well as to the gregarine cell cortex in
trophozoites and gamonts (Figures 1K–L, 1N). In contrast to Factin,
no specific labelling of myosin corresponding to the septum
was observed (Figures 1K–N). The intense fluorescence signal
observed in the area of the obviously uneven protomerite top of
some individuals lacking the epimerite most likely corresponded to
a labelling of myosin in host tissue remnants covering the
protomerite surface (Figures 1L–N). In addition, the intensely
labelled apical end of the protomerite in some individuals seemed
to be protracted or slightly retracted with an attached fragment of
the host epithelium (Figure 1M). These data suggest that, in G.
cuneata, more advanced stages than trophozoites remained in close
contact with the host epithelium and detailed electron microscopic
observations described below confirmed this assumption. In
addition, some of the primites exhibited distinct circumscribed
circular accumulation of myosin restricted to the periphery at the
base of their lance-shaped protomerite top (Figure 1N). This
structure might be related to the gamont feeding and/or
attachment, however, its exact functions remains unclear as no
comparable structure was observed under transmission electron
microscope.
Fine structure of feeding stages and their interactions
with the host epithelium
After entering the host intestine, invasive stages (sporozoites)
excysted from the oocyst and invaded the host epithelium. During
the invasion process, a slender sporozoite, tapering towards its
posterior end, attached to the host cell plasma membrane via its
apical part (Figure 2A) and, subsequently, the development of
electron-lucent epimeritic bud started (Figures 2B–E, 2G). The
apical cytoplasm of the invading parasite was packed with
numerous electron-dense micronemes (Figures 2E–F) and a more
or less translucent rhoptry-like organelle, passing through a
conoid, that seemed to empty its contents at this stage
(Figures 2D–E, 2G). In the course of transformation into a
trophozoite, the sporozoite enlarged and attained a more round
shape, and the epimeritic bud developed into an epimerite,
gradually implanting into the host epithelial cell (Figures 2H–K).
The early trophozoite was attached to the host cell via an
irregularly shaped epimerite, with its protodeutomerite hanging
free into the intestinal lumen, and developed surrounded by host
cell microvilli (Figures 2H–K). The irregularly shaped epimerite
Epicellular Parasitism in Eugregarines
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was still increasing in its size and was overlain by an indistinct
cortical vesicle. The membrane-like structure, limiting the cortical
vesicle on its cytoplasmic face, was discontinuous and often not
apparent (Figures 2H–K). A few mitochondria were observed in
the cytoplasm just beneath the epimeritic cortical vesicle
(Figure 2H). The interface between the epimerite and the host
cell was trilaminate, consisting of the epimerite and host plasma
membranes with an intercellular space in between them. In some
individuals, the epimerite plasma membrane covering the cortical
vesicle formed numerous digitations and rhizoids (Figure 2I). The
gradually expanding epimerite in the young trophozoite was very
rich in endoplasmic reticulum connected to the nuclear envelope
and often seen to be associated with the cortical vesicle covering
the epimerite (Figures 2J–K). In addition, an apically located
structure of unknown origin and function could be found close to
(or in contact with) the cortical vesicle in some maturing
trophozoites (Figure 2K). Considering its appearance and apical
localisation, this structure could correspond to the residuum of the
Figure 1. Early development of Gregarina cuneata observed using a light microscope. A. Earliest stages of trophozoites with developed
epimerites (asterisks) under transmission light (left) and in phase contrast (right). B. Early trophozoites with polymorphous epimerites (asterisks).
Transmission light. C. Detached maturing trophozoite with an epimerite surrounded by the host cell (arrowhead). Phase contrast. D. Three maturing
trophozoites exhibiting obvious injury to their epimerites (asterisks) after forced separation from the host tissue by specimen processing.
Transmission light. E. Maturing two-segmented individual exhibiting a well-developed protomerite and a cylindrical deutomerite. Note the rounded
top of the protomerite lacking an epimerite. Phase contrast. F. Detailed view of the rounded protomerite top of a living gamont (left) and of the
lance-shaped protomerite top of a chemically fixed gamont (right). Transmission light. G. Living gamonts associated in syzygy; primite (p), satellite (s).
Note the rounded top of the primite protomerite with some remnants of the host tissue (arrowhead). Phase contrast. H. Chemically fixed gamonts
associated in syzygy; lance-shaped top of the primite protomerite (asterisk), primite (p), satellite (s). I. Localisation of F-actin in early trophozoites (left)
and maturing gamont (right); epimerite (asterisk), ruptured epimerite (arrowhead), septum (arrows) separating the protomerite from the
deutomerite. Note that the septum (arrow) in the gamont is bulging into the protomerite. Direct fluorescence. J. Localisation of actin in maturing
gamont. Note the patchy accumulation of actin with a very intense signal (green) in the protomerite cytoplasm. Immunofluorescence, counterstained
with Evans blue. K. Localisation of myosin in trophozoites; epimerite (asterisk), ruptured epimerite (arrowhead). Immunofluorescence. L. Localisation
of myosin in maturing individuals. The top of the protomerite exhibits more (left) or less (right) intense labelling, suggesting the presence of host
tissue fragments. The inset shows the protomerite of more advanced stage of maturing gamont. Immunofluorescence, counterstained with Evans
blue. M. Localisation of myosin in single maturing gamonts after detachment from host epithelium. The protracted (left) and retracted (right)
protomerite tops exhibit strong labelling, suggesting the presence of host tissue fragments. Immunofluorescence; fluorescence and combination of
fluorescence with transmission light. N. Localisation of myosin in mature gamonts associated in syzygies. Note the primite (left) with a lance-shaped
top of the protomerite (asterisk) exhibiting distinct labelling in the peripheral area at its base (arrow) as well as the primite (right) with fragments of
the host tissue covering its protomerite top (arrowhead). Immunofluorescence; combination of fluorescence with transmission light.
doi:10.1371/journal.pone.0042606.g001
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rhoptry-like organelle described above in the invading stages. The
trophozoite was covered by a classical apicomplexan pellicle,
consisting of a plasma membrane and an inner membrane
complex, organised in raising longitudinal epicytic folds
(Figures 2I–J). This three-layered pellicle, however, extended only
to the protomerite top, at which point the inner membrane
complex ceased and only the plasma membrane covered the
embedded epimerite. The membrane fusion site between the
epimerite plasma membrane, host cell plasma membrane and the
membrane-like structure limiting the cortical vesicle was inconspicuous
(Figures 2H–K).
During trophozoite maturation, a thin septum developed and
separated the protomerite from the deutomerite, retaining the
large nucleus (Figure 3A). The epimerite appeared as an apical
extension of the protomerite, growing through the host cell and
interwoven with its plasma membrane, and overlain by an
indistinct cortical vesicle (Figures 3A–B). The cytoplasm of the
protomerite possessed numerous inclusions, including amylopectin
granules. The cytoplasmic area interconnecting the epimerite and
Figure 2. Early development of Gregarina cuneata observed using a transmission electron microscope. A. Invading sporozoite; host cell
microvilli (mv), sporozoite nucleus (n). B–G. Sporozoite transforming into the trophozoite stage; conoid (arrow), developing epimeritic bud (asterisks),
host cell (hc), host cell microvilli (mv), micronemes (arrowheads), microtubule (white arrow), nucleus (n), rhoptry-like organelle (r), subpellicular
microtubules (double arrowheads). H. Early trophozoite stage. Note the anterior part of the gregarine, covered by a developing cortical vesicle
(asterisks), causing an invagination of the host cell (hc) plasma membrane; host cell microvilli (mv), membrane fusion site (in circle), mitochondria
(arrowheads), nucleus (n), pellicle (double arrow). Insets show details of the membrane fusion sites. I. Early trophozoite. Note the folded plasma
membrane covering the cortical vesicle (asterisks) and forming numerous digitations; host cell (hc), host cell microvilli (mv), membrane fusion site (in
circle), nucleus (n). J. Developing trophozoite; cortical vesicle (asterisks), endoplasmic reticulum (er), host cell (hc), host cell microvilli (mv), membrane
fusion site (in circle), nucleus (n), pellicle with raising epicytic folds (double arrow). K. The apical end of another maturing trophozoite; amylopectin
granules (am), cortical vesicle (asterisks), endoplasmic reticulum (er), host cell (hc), membrane fusion site (in circle), nucleus (n), unknown structure
(st).
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protomerite contained numerous membrane cisternae and vesicles
(Figure 3B). In the course of trophozoite maturation, the epimerite
seemed to decrease and the host cell and epimerite plasma
membranes previously forming the trilaminate interface became
indistinguishable from each other (Figures 3C–E). At this stage of
gregarine epicellular development, the affected host cell exhibited
some degree of vacuolation and in some sections cellular
disorganisation (Figures 3C–E, 3I). The epimerite was irregularly
embedded into the host cell (or in close contact with it), and
formed numerous rhizoids or digitations of variable size and
shape. The cytoplasm of these epimerite digitations appeared
translucent, filled with numerous fine filamentous structures and
mitochondria-like organelles underlying the cortical vesicle. The
membrane fusion site, though not so prominent as in other
eugregarines parasitising mealworms, was still visible (Figures 3C–
E, 3I) and the freeze-etching technique revealed further details of
its typical architecture (Figures 3F–H). According to the results of
ultrathin sectioning, the border between the epimeritic cortical
vesicle and the host cell was formed by the epimerite plasma
membrane and the invaginated host plasma membrane, the
second one of which was continuous with the plasma membrane
covering surrounding microvilli. The parasite plasma membrane
covering the epimerite was continuous with plasma membrane of
the protomerite (Figure 3G). Similarly to the observations of
ultrathin sections of mature trophozoites, epimerite and host cell
plasma membranes were difficult to be distinguished from each
other. The so-called ‘membrane-like structure’ limiting the cortical
vesicle on its cytoplasmic face appeared as a membrane that was
discontinuous in some areas, but often better visible than in
ultrathin sections (Figures 3F–G). Nevertheless, it still remains
unclear whether this membranous structure beneath the cortical
vesicle was directly linked to the membrane fusion site or not
(Figures 3G–H). Longitudinally oriented epicytic folds were a
feature of both the trophozoite (Figure 3A) and gamont (e.g.
Figure 4A) stages. In the course of trophozoite development, the
decrease of epimerite proceeded and its detachment from host
epithelium initiated. The protomerite top of individuals transforming
from a trophozoite into a gamont exhibited an uneven
surface with short rhizoid-like structures irregularly attached to the
host tissue (Figure 3I) and thus resembling a retracted epimerite.
The contact with the intestinal epithelium, however, was partially
discontinuous, at least when observed in ultrathin sections.
Older stages, considered to be single maturing gamonts or
primites, exhibited protomerites with broad lance-shaped apical
ends (Figures 4A–D), similar to the light microscopic observations
on chemically fixed parasites (Figures 1F, 1H, 1N), and usually in
contact with host tissue. Less often, the protomerite top, contacting
host microvilli, appeared widely rounded (Figures 4G–H). The
apical end of the protomerite, regardless of its shape, was covered
by a trilaminate pellicle lacking the typical organisation into
longitudinal epicytic folds (Figures 4A–I). Under the scanning
electron microscope, the cylindrical protomerite reached its
maximum width at the interface between the apical part covered
by a smooth pellicle and the posterior part with a pellicle organised
into longitudinal epicytic folds (Figure 4H). The outer surface of
the widely rounded protomerite top was wrinkled, bearing
numerous non-specified globules of different size (Figures 4E–F).
The localization and size of these globules corresponded to the
myosin labelling of host tissue remnants still attached to the
protomerite surface (as shown in Figure 1L). Many of the gamonts
processed for scanning electron microscopy exhibited serious
injury on the apical part of the protomerite (Figure 4E), often
bearing scraps of host tissue (Figures 4H–I).
Detailed ultrastructural analysis of protomerite top found in
close contact with host tissue revealed its unusual organisation,
most likely dedicated to parasite food intake (Figures 5A–H). The
contact of the gamont protomerite with host tissue was uneven,
lacking any continuous intimate connection between the host cell
and parasite plasma membranes (Figures 5B–E); more often, the
protomerite top touched the host microvilli (Figure 5A). The apical
end of the protomerite was covered by a smooth trilaminate
pellicle, not organised in epicytic folds, with irregularly distributed
pore-like structures (Figures 5C–E, 5H). In some sections, the
protomerite top even showed a more undulated pattern
(Figure 5G). Using higher magnification, a dense layer with nonmembranous
character, resembling the internal lamina usually
underlining eugregarine epicytic folds, could be seen underlying
the inner membrane complex at its cytoplasmic face (Figures 5D–
E, 5H). The pore-like structures interrupting the inner membrane
complex were more concentrated in some areas. In addition,
structures similar to dense bodies, in some sections already halfemptied,
could be seen in connection with them (Figures 5C–D).
Unusual duct-like structures of unknown function could be found
in the protomerite apical cytoplasm; usually, they were linked to
the dense layer and in some sections their connection to
abovementioned pore-like structures could be seen (Figure 5C).
When observed under higher magnification, these structures
appeared as elongated dense sacs passing through the inner
membrane complex and plasma membrane and opening outwards
(Figures 5E, 5H). The protomerite cytoplasm was packed with
dense bodies, various vesicles and an abundant Golgi apparatus
(Figure 5F).
Although the ultrastructural analysis revealed localized, mild
pathological changes of the parasitised epithelium (usually limited
to the affected cell), they seemed to be of minimal or no clinical
significance. Despite often heavy infestation by G. cuneata in the
host mid-gut, the experimentally infected larvae exhibited no
obvious signs of sickness that could be considered to correlate with
the progress of parasitisation. Parasitised larvae did not show any
behavioural changes, weight loss or decreased food intake. In fact,
the presence of eugregarines (regardless of eugregarine species) in
experimentally as well as naturally infected mealworm larvae even
seemed to increase the host growth rate and to reduce the death
rate, and these larvae appeared to be more aggressive and agile in
comparison to the gregarine-free individuals.
Discussion
The observations on early development of Gregarina cuneata
generally support previously published data on another eugregarines
[4,6,7,8]. Although G. cuneata trophozoites possess epimerite
that slightly differs from those reported in other eugregarines
parasitising mealworms, they also develop epicellularly and exhibit
the same stages during their life cycle. Nevertheless, the later
developmental stages exhibit more advanced adaptations to the
epicellular parasitism and to the nutrient acquisition in intestinal
environment, and details of these will be discussed below.
Eugregarine attachment to host tissue
It has been determined that mesenteric epithelial cells are shortlived,
living only four days in T. molitor [27]. The destiny of
trophozoites with their epimerites still embedded in degenerating
epithelial cells, often observed in insect hosts, is still unknown.
Harry [28] described trophozoite detachment from the host tissue
as a random and passive process at any stage of its development,
depending on the degeneration of epithelial cells and their
extrusion under the pressure of replacement cells. He referred to
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detached trophozoites still possessing epimerites observed in
histological sections. On the contrary, recently published data
[8,11] revealed that epimerite detachment is an active process, and
thus trophozoites most likely detach and search for a new host cell
in better physiological conditions. Insofar as the vegetative phase
of the eugregarine life cycle usually lasts longer than four days
[6,29], trophozoites must be adapted either to keeping the host cell
alive during their development or for eventual reattachment to
another cell. Therefore, Lucarotti [11] speculated about hypothetical
reattachment of Leidyana trophozoites to younger cells after
abandoning senescing cells, facilitated by a retractable epimerite
and eugregarine gliding motility. The presence of contractile
elements in the area of the epimerite [20,21] and protomerite top
[30,31] serves as a more convincing argument in favour of the
structural dynamics of these cell regions and epimerite retraction
theory [8,11]. Similarly, actin-like filaments demonstrated in the
mucron of lecudinid eugregarine are considered to facilitate its
adhesion to the host cell [20,32]. As the parasite’s fixation to host
tissue might be of a temporary nature, the contact between the
host cell and parasite attachment organelle must be very loose.
Studies on attachment strategies of several eugregarines
[4,6,7,8,10,24] support this hypothesis in that they showed that,
Figure 3. Trophozoites of Gregarina cuneata observed using a transmission electron microscope. A. Trophozoite with a well-developed
epimerite (asterisk); deutomerite (d) with a nucleus, host cell microvilli (mv), protomerite (p). B. A more detailed view of the epimerite (asterisk)
shown in Fig. 3A; host cell (hc), protomerite (p) cytoplasm packed with numerous inclusions. C–E. Decreasing epimerite (asterisks) forming numerous
rhizoids and digitations (arrowheads) in more advanced stages of trophozoites as observed in different planes of sectioning; host cell (hc), host cell
microvilli (mv), membrane fusion site (in circle), protomerite (p). The inset in Fig. 3E shows the membrane fusion site in detail. F–H. Host cellepimerite
interactions visualised by a freeze-etching technique. Fig. 3G shows a more detailed view of the membrane fusion site (in circle) from
Fig. 3F. Note the border (arrowheads) between the epimerite and host cell (hc); cortical vesicle (asterisks), epicytic folds (ef) of the protomerite region
(p), host cell microvilli (mv), host cell plasma membrane (white arrow), membrane-like structure limiting the cortical vesicle on its cytoplasmic face
(white arrowheads), parasite plasma membrane (arrow). I. Trophozoite exhibiting a quite completely decreased epimerite (asterisks) with numerous
mitochondria and gradual detachment (arrowhead) from host cell (hc), membrane fusion sites (in circles), protomerite (p).
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in the course of early trophozoite development, the gradually
enlarging epimerite causes a deep invagination of the host plasma
membrane, thus allowing the parasite to anchor to the surface of
the host cell and to develop in an epicellular position. The
interface between the epimerite and the host cell consists of
epimerite and host plasma membranes, and a dense layer of
unknown nature and origin in between them [4,6,7,8,10,33]. The
contact between the epimerite and the host plasma membrane is
reported to be of the membrane-to-membrane type; lecudinid
eugregarines establish an intimate contact with the host cell
without an interspecific cell junction [34]. In fact, numerous
detached trophozoites retaining intact epimerites, commonly
observed in native or fixed squash preparations, are the evidence
that no real fusion develops between the epimerite and host
plasma membranes along the trilaminar interface. It seems that,
simultaneously with trophozoite maturation, the host and epimerite
membranes start to lose adhesion to one another and the
epimerite gradually detaches from the epithelium. I repeatedly
observed attached mature trophozoites of various species, in which
the epimerite membrane was already separated from the host
plasma membrane [7].
The feeding stages of G. cuneata exhibit an even more
spectacular adaptation to epicellular parasitism. The atypical
epimerite of G. cuneata develops from the epimeritic bud in
accordance with other eugregarines, and later in development
forms numerous digitations, deeply invaginating the plasma
Figure 4. Gamonts of Gregarina cuneata observed using an
electron microscope. A. Individual exhibiting a lance-shaped top
(arrowhead) of the protomerite (p) in contact with a host cell (hc);
deutomerite (d), microvilli (mv), septum (arrow). B. Higher magnification
of the protomerite top shown in Fig. 4A. Note the close contact of
protomerite (p) with the host cell (hc) in some areas. C. Longitudinal
section of the protomerite (p) separated from the deutomerite (d) by a
septum (arrow). Note that the tapered protomerite top, which is in
contact (arrowheads) with host cell microvilli (mv), lacks the epicytic
folds (ef) covering the rest of the gregarine body. D. A view of the
protomerite (p) top (arrowheads) in close contact with the microvillous
surface (mv) of host epithelial cells (hc); amorphous material (6),
deutomerite (d). E. Scanning electron micrograph showing the
protomerite top covered by a wrinkled plasma membrane; protomerite
epicytic folds (ef). The apical end of the protomerite is obviously
damaged (arrowheads), probably due to mechanical separation of the
gregarine from the host tissue during specimen processing. F. A more
detailed view of the protomerite top exhibiting small remnants of host
tissue still attached to its plasma membrane. G. A general view of the
protomerite (p) separated from the deutomerite (d) by a distinct
septum (arrow); epicytic folds (ef). Arrowheads indicate the rounded
protomerite top in contact with host cell microvilli. H. Scanning
electron micrograph showing the rounded protomerite top (arrowhead)
with a scrap of host tissue (t) attached; epicytic folds (ef) covering
the rest of protomerite. I. A more detailed view of the plasma
membrane covering the protomerite top shown in Fig. 4H; scrap of host
tissue (t).
doi:10.1371/journal.pone.0042606.g004
Figure 5. Host-parasite interactions in Gregarina cuneata as
observed by transmission electron microscopy. A, B. A more
detailed view of the protomerite top (p) in close contact with host
microvilli (mv) or epithelial cells (hc); amorphous material (6). C–E. A
detail of the protomerite (p) apical region covered by a three-layered
pellicle underlined by a dense layer; Golgi apparatus (g), host cells (hc),
microvilli (mv). Note the ducts (arrows) passing to the exterior,
numerous dense bodies (asterisks), semi-empty (Fig. 5C) and filled
(Fig. 5D) dense structures (in circle) directly linked to the pore-like
structures (arrowheads) interrupting the inner membrane complex. F.
Higher magnification of the Golgi apparatus frequently observed in the
protomerite cytoplasm. G. A view of the protomerite top (p) exhibiting
a more undulating pattern (arrowheads) in the area adjacent to the host
epithelium (hc) with microvilli (mv); epicytic folds (ef), numerous dense
bodies (asterisks). H. A higher magnification showing the protomerite
(p) apical region with unusual duct-like structures (arrows). This region
is obviously covered by a typical three-layered pellicle consisting of a
plasma membrane and inner membrane complex underlined by a
dense layer, but it lacks epicytic folds. Note that the inner membrane
complex is discontinuous in a periodic pattern (interrupted by pore-like
structures); amorphous material (6), dense bodies (asterisks), host cells
(hc).
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membrane on the luminal side of the affected host cell. Despite this
parasite’s firm anchoring to the brush border of the host cell,
especially in mature stages when the host and epimerite
membranes become almost indistinguishable from each other, G.
cuneata trophozoites are able to detach while retaining an intact
epimerite. Nevertheless, in contrast to other gregarines from
mealworms, detached trophozoites of G. cuneata more often were
found to have ruptured epimerites. Although the trophozoite
development is more or less identical in all eugregarines from T.
molitor, the destiny of G. cuneata mature trophozoites significantly
differs in that they form so-called early syzygies, often found to be
still attached to the host tissue. Surprisingly, the attachment site of
attached primites significantly differs from the epimerite in
younger stages, despite their resemblance at the light microscopic
level. The ‘real’ epimerite disappears (retracts) and the top of the
protomerite with a more or less undulating pattern remains in
contact with the epithelium. The attachment by means of a
modified protomerite could be facilitated by an increased
flexibility of this region, as suggested by the dot-like accumulation
of actin in the protomerite of G. cuneata as well as G. polymorpha
[30]. Comparable attachment strategy has been reported from
actinocephalid eugregarines [35] and it could be expected that
protomerites modified for attachment act as feeding organelles in
eugregarines lacking an epimerite. Similarly to G. cuneata, gamonts
of some actinocephalids lose their small globular epimerites and
subsequently attach by a modified, sucker-like protomerite [35].
The space between the epicytic folds of the attached protomerite
and the host epithelium is filled by the microvilli embedded in a
dense material interpreted to be adhesive. The authors speculate
that this material could be produced by small dense (exocytic)
vesicles in the protomerite apical cytoplasm. The space between
the host microvilli and the G. cuneata protomerite top was also filled
with an amorphous ropey material of unknown origin, probably
serving as an adhesive. These observations are supported by the
frequent presence of host tissue remnants attached to the G. cuneata
protomerite, which was confirmed by the fluorescence labelling of
myosin and not reported in other gregarines from mealworms.
Nutrient acquisition in eugregarines
Nutrition of gregarines has been the subject of extensive debate
for decades. There is evidence that feeding mode in gregarines
depends on the long-term environmental conditions forming their
niche. Correlations between trophozoite characteristics and the
environment occupied within the host are discussed elsewhere
[36]. The earliest diverging apicomplexans, archigregarines
parasitising marine invertebrates, have retained myzocytosis as
their principal mode of feeding [36]. The extensive folding of the
pellicle covering the surface of large trophozoites of marine
eugregarines seems to optimise surface-mediated nutrition (pinocytosis
via micropores), and thus could explain the loss of an apical
complex and myzocytosis in eugregarines along with the
development of a bulky attachment apparatus, such as an
epimerite or mucron [37].
The feeding strategy might even differ between distant
eugregarine taxa. For instance, the supposed lytic effect of
lecudinids on host cells indicates the nutritional function of the
mucron via extracellular secretion of enzymes and absorption of
digested host tissue [9]. In general, the epimerite cytoplasm
contains many organelles usually associated with nutritive function
[14]. Ghazali et al. [20] concluded that epimerites do not have a
direct sucker function because of the absence of actin in the G.
blaberae epimerite. On the contrary, our data confirmed the
presence of F-actin in the epimerite region of eugregarines from
mealworms [8,this study]. As host cells affected by attachment of
Gregarina spp. vegetative stages usually do not show obvious
pathological changes, the cortical vesicle and epimerite vacuoles
most likely absorb nutrients via a mechanism based on membrane
permeability [9]. Numerous mitochondria underlying the cortical
vesicle, regularly observed in various eugregarine species [7,8,10],
could provide the energy necessary for this putative absorption
mechanism. The abundant endoplasmic reticulum repeatedly
observed in the area of the expanding epimerite in young
trophozoites of G. cuneata indicates the activation of metabolic
pathways, probably involved in the synthesis and secretion of
proteins and membrane manufacturing. The significant reduction
in size of G. cuneata cortical vesicle might be related to the
convoluted character of the epimerite, significantly increasing its
absorptive surface, as reported in Didymophyes [12]. Similarly, the
trilaminar junction between the mucron of the monocystid
eugregarine Nematocystis and the host epithelial cell forms extensive
folds to increase the surface contact between their apposing cell
membranes [33]. Using radioisotopes, the study demonstrated that
metabolites pass directly from the host cell to the trophozoites by
crossing the attachment site of Nematocystis.
In gamonts of G. cuneata with their modified protomerites
contacting the host epithelium, the pore- and duct-like structures
were associated with the pellicle covering the protomerite top.
Although the function of these structures remains uncertain, they
are most likely involved in gamont nutrition and/or attachment.
The apical localisation of numerous dense bodies, various vesicles
and abundant Golgi apparatus in the protomerite cytoplasm of G.
cuneata gamonts similarly indicates the involvement of protomerite
top in the feeding.
The basic mechanisms of nutrient acquisition in gregarines,
however, are still to be resolved. Despite all these studies
attributing the major nutritional role to the attachment organelles,
another possibility must be sketched, especially when considering
the existence of gregarines growing in the coelomic fluid without
an attachment to the host tissue. In addition, eugregarines usually
continue to grow after detaching from the host tissue [9]. There
are often speculations on the functionality of the micropore-like
structures that are often observed in the spaces between epicytic
folds [11,16,17,18]; nevertheless, more elaborate analyses are
needed to determine their involvement in gregarine nutrition and/
or movement.
Pathogenicity to insect hosts
Eugregarines are probably the most frequently encountered
protists in insects and probably the most innocuous. As a rule, they
are considered to be non-pathogenic to their hosts [38]; however,
the real impact of eugregarine infection on host fitness and
viability is still poorly understood. Misinterpretation of regular
cellular processes in host tissue might significantly contribute to the
controversy surrounding the pathogenicity of eugregarines. In
addition, gregarines usually parasitise digestive epithelia that are
the first to undergo autolysis after dissection, and this could hinder
the correct determination of pathological changes induced by
gregarines and distinguishing them from the post-mortem
autolytic changes to the tissue. Some authors attributed pathogenicity
mostly to trophozoites, which theoretically might cause some
degree of damage to host tissue depending on the size and shape of
their embedded epimerites [39]. The robust epimerite of
Ancyrophora equipped with rigid hooks, however, does not appear
to induce drastic damage to the host cell [14]. In fact, although
eugregarines infecting the intestinal epithelium might cause certain
damage to affected cells, continual regeneration of these cells
accounts for the apparent harmless effect of the parasite. Usually,
even if the eugregarine trophozoites destroy individual cells, the
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overall damage to epithelial tissue is negligible and easily repaired.
Nevertheless, some species appear to reduce the host’s fitness by
occluding its gut and thus preventing the passage of food [11]. In
addition, heavy infestations of gregarines in the mesenteron can
have a significant impact on the host’s nutritional state. As
microvilli are important structures for efficient absorption and
excretion, their destruction might limit the host digestive process
and lead to its malnutrition with consequent weakening or even
death. Gregarines parasitising intestinal caeca are even much
more pathogenic, as they may cause the hypertrophy of parasitised
cells or even rupture the caecal wall, leading to a secondary
bacterial infection [40].
The eugregarines from mealworms were previously considered
to be parasitic because of their negative impact on the
development of larvae grown on a suboptimal diet [41].
Eugregarines occurring in larval T. molitor from our colonies,
however, not only do not appear to harm their host, but could
actually be considered to be mutualistic from a certain point of
view. My personal long-term observations on T. molitor confirmed
that despite heavy infection completely filling the larval mid-gut,
the presence of eugregarines seems to have a positive impact on
host development, fitness and longevity. Identical observations
were made on naturally infected mealworm larvae from our
laboratory colonies, usually parasitised by multiple eugregarine
species (G. cuneata, G. polymorpha and G. steini) simultaneously.
Similarly, Sumner [42] considered gregarines from mealworms to
be symbiotic, and necessary for the normal growth and longevity
of the host. This author even suggested that gregarines probably
secrete essential substances such as vitamins or enzymes essential
for larval growth. This study confirms that, though the affected
epithelium shows some changes, parasitisation by G. cuneata seems
to have no negative impact on host health that is essential for the
gregarine survival. Despite high densities of vegetative stages
attached to the host intestinal tissue, there is no evidence of direct
damage to neighbouring epithelial cells. Vacuolation and eventual
subsequent death of individual affected epithelial cells represented
the most marked changes that could be considered to be associated
with gregarine infection.
Morphological changes of Gregarina cuneata in different
environmental conditions
In the course of development, the epimerite of G. cuneata
undergoes dramatic changes and some of these have been shown
to be reversible depending on actual environmental conditions.
Various stimuli from the trophozoite environment, such as
changes in the chemical composition of the dissection buffer/host
tissue, pH or temperature, seem to induce significant morphological
changes of the epimerite and the protomerite top. Significant
differences in the protomerite shape, evident especially in the
primites, were noticed in this study prior to and after chemical
fixation with different cross-linking fixatives - paraformaldehyde
and glutaraldehyde. Non-fixed living gamonts exhibited a rounded
protomerite top; however, those fixed with a paraformaldehyde
solution often exhibited a lance-shaped protomerite top. These
individuals are assumed to have been mechanically detached from
the host tissue during specimen processing and simultaneously
chemically fixed, thus maintaining the real shape of the
protomerite when in close contact with the epithelium. Corresponding
stages fixed with glutaraldehyde, however, did not
exhibit such an obvious extension and tapering of their apical
ends, although the protomerite top of gamonts was often slightly
raised and covered by host tissue fragments. Only individuals
found in contact with the host tissue after fixation preserved the
lance-shaped protomerite top. Formaldehyde-based solutions fix
the tissue by cross-linking proteins; its effects are reversible by
excess water and the benefits include good tissue penetration. As
glutaraldehyde is a larger molecule, the weakness of this fixative
includes a slower rate of diffusion across membranes, resulting in
poor tissue penetration and the changes caused by fixation are
irreversible [43]. As the fixatives are known to induce remarkable
changes in cell shape, rapid fixation by paraformaldehyde is
thought to be the source of differences in the protomerite
morphology in this study. This unexpected outcome of different
fixations revealed morphological adaptations of G. cuneata to
epicellular parasitism that are not commonly observed in living
specimens. The facts discussed herein suggest that this gregarine is
able not only retract but even repeatedly protract its apical end
(epimerite or protomerite top dedicated to attachment) depending
on environmental conditions and the need to reattach to another
part of the host tissue.
Conclusions
Gregarines are important from an evolutionary perspective
because of their suspected deep-branching position within the
phylum Apicomplexa. Although some ancestral features found in
gregarines have given them a reputation of being a ‘primitive’
lineage of the Apicomplexa, the majority of them exhibit unique
and novel adaptations to their environment [37]. A wide variety of
morphological and functional adaptations that can be found in all
gregarine taxa, along with the fact that only few invertebrate
groups escaped infection with gregarines, indicates that they must
be regarded as very successful and highly specialized parasites.
The fascinating biology of these apicomplexans is derived from the
basic cellular organization of the so-called zoite, an infectious
developmental stage devoted to the invasion of host tissue. The
detachment of vegetative stage from host tissue and its eventual
reattachment, self-regulated by the parasite, might represent a
higher degree of gregarine adaptation to epicellular development
in hosts exhibiting a rapid epithelial replacement (e.g. insects). The
modified protomerite of G. cuneata gamonts, serving for attachment
to the host tissue and parasite feeding, indicates further adaptation
of eugregarines for nutrient acquisition in older developmental
stages that were previously considered to be non-vegetative. Such
modifications for epicellular parasitism do not seem to be primitive
ancestral characteristics, but rather advanced features occurring in
some eugregarines in the course of their coevolution with the host.
Acknowledgments
The author is greatly indebted to the members of the Laboratory of
Electron Microscopy (Institute of Parasitology, Biology Centre of the
Academy of Sciences of the Czech Republic, in Cˇ eske´ Budeˇjovice) for
technical assistance. Many thanks also to Prof. Dominique Soldati-Favre
(University of Geneva, Switzerland) for her kindly providing the
monoclonal anti-actin antibody.
Author Contributions
Conceived and designed the experiments: AV. Performed the experiments:
AV. Analyzed the data: AV. Contributed reagents/materials/analysis
tools: AV. Wrote the paper: AV.
Epicellular Parasitism in Eugregarines
PLOS ONE | www.plosone.org 10 August 2012 | Volume 7 | Issue 8 | e42606
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Epicellular Parasitism in Eugregarines
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7.2
Valigurová A., Vaškovicová V., Musilová N., Schrével J.
2013
The enigma of eugregarine epicytic folds:
where gliding motility originates?
Frontiers in Zoology 10, 57
The enigma of eugregarine epicytic folds: where
gliding motility originates?
Valigurová et al.
Valigurová et al. Frontiers in Zoology 2013, 10:57
http://www.frontiersinzoology.com/content/10/1/57
RESEARCH Open Access
The enigma of eugregarine epicytic folds: where
gliding motility originates?
Andrea Valigurová1*
, Naděžda Vaškovicová2
, Naďa Musilová1
and Joseph Schrével3
Abstract
Background: In the past decades, many studies focused on the cell motility of apicomplexan invasive stages as
they represent a potential target for chemotherapeutic intervention. Gregarines (Conoidasida, Gregarinasina) are a
heterogeneous group that parasitize invertebrates and urochordates, and are thought to be an early branching
lineage of Apicomplexa. As characteristic of apicomplexan zoites, gregarines are covered by a complicated pellicle,
consisting of the plasma membrane and the closely apposed inner membrane complex, which is associated with a
number of cytoskeletal elements. The cell cortex of eugregarines, the epicyte, is more complicated than that of
other apicomplexans, as it forms various superficial structures.
Results: The epicyte of the eugregarines, Gregarina cuneata, G. polymorpha and G. steini, analysed in the present
study is organised in longitudinal folds covering the entire cell. In mature trophozoites and gamonts, each epicytic
fold exhibits similar ectoplasmic structures and is built up from the plasma membrane, inner membrane complex,
12-nm filaments, rippled dense structures and basal lamina. In addition, rib-like myonemes and an ectoplasmic
network are frequently observed. Under experimental conditions, eugregarines showed varied speeds and paths of
simple linear gliding. In all three species, actin and myosin were associated with the pellicle, and this actomyosin
complex appeared to be restricted to the lateral parts of the epicytic folds. Treatment of living gamonts with
jasplakinolide and cytochalasin D confirmed that actin actively participates in gregarine gliding. Contributions to
gliding of specific subcellular components are discussed.
Conclusions: Cell motility in gregarines and other apicomplexans share features in common, i.e. a three-layered
pellicle, an actomyosin complex, and the polymerisation of actin during gliding. Although the general architecture
and supramolecular organisation of the pellicle is not correlated with gliding rates of eugregarines, an increase in
cytoplasmic mucus concentration is correlated. Furthermore, our data suggest that gregarines utilize several
mechanisms of cell motility and that this is influenced by environmental conditions.
Keywords: Actin, Cytochalasin D, Epicyte, Epicytic folds, Eugregarine, Glideosome, Gliding motility, Jasplakinolide,
Mucus, Myosin, Pellicle
Introduction
Apicomplexans are one of the most successful and diverse
groups of eukaryotic unicellular parasites that exhibit
unique adaptations to life in a wide spectrum of
vertebrate and invertebrate hosts. Many cause major human
diseases, i.e. malaria, toxoplasmosis, coccidiosis and
cryptosporidiosis. Because apicomplexan diseases are still
problematic, therapeutic research focuses either on parasitic
structures or metabolic pathways which might serve
as drug targets. The cytoskeleton of these parasites has become
a focus for drug development because it plays an
important role in various life processes, e.g., locomotion,
division, invasion and formation of parasite cell polarity
[1]. This is especially true of invasive stages of Toxoplasma
gondii and Plasmodium falciparum [2,3].
Infective stages of Apicomplexa are characterised by an
apical complex of organelles as well as a complicated cell
cortex consisting of cortical alveoli, i.e., flattened vesicles
limited by a membrane and packed into a continuous
layer (inner membrane complex), underlying the plasma
membrane. The inner membrane complex (IMC) has micropores
and connects numerous cytoskeletal elements
* Correspondence: andreav@sci.muni.cz
1
Department of Botany and Zoology, Faculty of Science, Masaryk University,
Kotlářská 2, 611 37 Brno, Czech Republic
Full list of author information is available at the end of the article
© 2013 Valigurová et al.; licensee BioMed Central Ltd. This is an Open Access article distributed under the terms of the
Creative Commons Attribution License (http://creativecommons.org/licenses/by/2.0), which permits unrestricted use,
distribution, and reproduction in any medium, provided the original work is properly cited.
Valigurová et al. Frontiers in Zoology 2013, 10:57
http://www.frontiersinzoology.com/content/10/1/57
that include an actomyosin complex, microtubules and
a network of intermediate filamentous proteins. The
invasive zoites of Apicomplexa are motile and use actinbased
gliding for host invasion and tissue traversal. This
gliding mechanism called ‘glideosome’ was first described
for Toxoplasma [4] and has been extended as a concept to
sporozoites of Plasmodium [5] and other apicomplexans
[6]. In Toxoplasma and Plasmodium, myosin A is linked
to the IMC and probably interacts with subpellicular
microtubules. The head of myosin A moves along the
actin filament, which is connected to a cell adhesion molecule
(TRAP in Plasmodium spp. or TgMIC2 in T. gondii)
via a tetrameric aldolase [6].
As deep-branching apicomplexan parasites of invertebrates
and urochordates, gregarines (Gregarinasina) are
generally thought to be of no economic importance.
Recent analyses, however, indicate a close affinity of gregarines
with species of Cryptosporidium [7,8] that parasitize
humans. Most eugregarine gamonts are covered by a
pellicle folded in numerous longitudinal epicytic folds
(e.g., Gregarina, Lecudina) [9-11] and exhibit gliding
motility [12-14]. Using a laser trap and bead translocation,
King and Sleep [15] have described gliding
as an irregular, erratic process. Some marine gregarines
(e.g. archigregarines) possess regular sets of subpellicular
microtubules under the pellicle [16,17] and typically
display a pendular or rolling movement. In contrast,
urosporidians that evolved as free-floating parasites within
the host tissue move by pulsation of their body wall corresponding
to the so-called peristaltic motility. The possible
function of epicytic folds has been often discussed, and it
is generally thought that they increase the surface area for
nutrient acquisition and facilitate actomyosin-based gliding
motility. The involvement of actin- and myosin-like
proteins in gregarine cell motility has been previously
reported [18-20]. Several electron microscopic studies
have revealed the typical organisation of epicytic folds in
eugregarines [10,11,21-23]. These suggest that there are
undulating epicytic folds located between those that do
not move [24-26], but the exact mechanism of motility
remains unclear. Therefore, it is imperative that structural
observations be integrated with biochemical and molecular
data for the actin and myosins of Gregarina species
[20,27]. Of the three myosin genes so far characterised
in Gregarina polymorpha, myosins A (93 kDa) and B
(96 kDa) belong to the class of myosin (XIV) that is
restricted to the phylum Apicomplexa [28] and myosin
F (222 kDa) to the class XII [27,29]. King and
Sleep [15] estimated that the number of myosin heads
at the site of interaction in Gregarina gamonts to be in
excess of 200 and showed that the gliding rate in a giant
eugregarine Porospora gigantea is four times higher (up to
60 μm/s) than the speed of myosin movement along the
actin filaments of a muscle sarcomere. In spite of a few
freeze-etching studies that focus on the supramolecular
cell organisation of some Gregarina species [11,22,30],
the precise location of the actomyosin complex is as
yet unknown. However, the TRAP or TgMIC2 molecules
that are in contact with the substrate suggest that the
concept of a glideosome might help shed light on the
role of the mucus [12] in the gliding mechanism of
gregarines [31-33].
Laboratory-reared colony of the mealworm Tenebrio
molitor parasitized by three species of Gregarina has
permitted the comparison of gliding by G. cuneata, G.
polymorpha and G. steini under identical environmental
conditions. This study was performed so that the respective
roles of the apical and lateral parts of the
epicytic folds in apicomplexan zoite gliding could be
discerned. Our intent was to evaluate the presumptive
involvement of specific subcellular components such as
the 12-nm filaments, rippled dense structures [11], and
mucus in eugregarine gliding motility [32,33] using both
the experimental and morphological approaches.
Results
Light microscopic observations on gregarine movement
behaviour and gliding motility
The pellicle (epicyte) appeared as a thick but transparent
layer of even width covering the entire gregarine (Figures 1A,
2A and 3A). Longitudinal striations that were easily recognisable
in the pellicle corresponded to the epicytic folds.
During gliding, the shape of the cell varied by species. In
G. cuneata and G. polymorpha, the changes of direction
during gliding seemed to be controlled by protomerite activity.
In G. polymorpha the protomerite and deutomerite
were very flexible. The bending of the protomerite may
take place in any plane, but in G. polymorpha it was sometimes
so extensive that the axis of the protomerite (or
even with the one-third of the deutomerite) came to form
a right or even acute angle with that of the deutomerite.
This behaviour was especially noted when gamonts encountered
barriers in their gliding path. In G. polymorpha,
a partial pulling of the protomerite into the deutomerite,
so that the corresponding pellicle became pleated, could
be observed. A slight bending of the protomerite could be
detected also in G. cuneata, but the angle between the
planes of protomerite and deutomerite was only obtuse.
In contrast, the protomerite of G. steini did not show any
changes during gliding; only a slight bending of the
gamont deutomerite, usually in its posterior half, was
observable when turning to the side. In syzygies of all
species, the satellite seemed to be passive and just
followed the path given by the obviously active primite,
and this path corresponded to a forward unidirectional
gliding. In all three species, the gliding locomotion of
single and associated gamonts was usually discontinuous
with multiple stops and frequent changes of direction, and
Valigurová et al. Frontiers in Zoology 2013, 10:57 Page 2 of 27
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often occurred with discernible changes in speed. The
gregarines glided in an almost linear pattern. The gliding
movement, however, was not constant and varied
among species as well as every gliding individual. The
maximum speed of gliding gamonts achieved during our
observations was 5.66 μm/s in single gamonts and
8.49 μm/s in syzygies of G. cuneata, 22.86 μm/s in single
gamonts and 16.18 μm/s in syzygies of G. polymorpha,
and 9.42 μm/s in single gamonts and 9.25 μm/s in syzygies
of G. steini (Table 1). Based on contact stimuli, gregarines
were able to quite rapidly change the direction of their
otherwise straightforward gliding to avoid a barrier in their
gliding path. When a gliding gamont encountered a barrier,
it usually endeavoured to bore or wriggle its way through.
Obviously, gamonts of G. steini exhibited the most continuous
and constant gliding with a linear or widely semicircular
track. The gliding of G. cuneata gamonts was
characterised by multiple and prolonged stops, and thus
many individuals did not exhibit gliding during the recording.
In contrast, gamonts of G. polymorpha covered the
greatest distance per unit of time in one continuous track.
Treatment of gregarine gamonts with jasplakinolide and
cytochalasin D
Various concentrations of both drugs were used to treat
gregarine gamonts. Concentrations of jasplakinolide
(JAS), a strong actin stabilizer, lower than 5 μM had no
effect. Thus, in order to obtain reliable results on active
and vital gregarines, they were treated with 5, 10, 20 and
30 μM concentrations of JAS in Ringer’s solution. Gregarines
not only survived in these very high doses of JAS,
but even actively moved for the next 90–150 min, depending
on the drug concentration and gregarine species
(Table 2). Experiments revealed that JAS treatment led
to an increased speed of gliding movement beyond
5 minutes after drug application, followed by subsequent
decrease to normal in all three species. Afterwards, individual
reactions rates to JAS differed by species with the
most rapid occurring in G. steini whose gamonts moved
up to 90 minutes. The most delayed reaction to the drug
(inhibition of gliding motility) exhibited syzygies of
G. cuneata, which moved in large numbers up to
150 minutes after JAS application. After a uniform
period of 1 hour, independently on above mentioned
concentrations of JAS, all three species exhibited obvious
cellular changes including shrivelling and some degree of
cytoplasmic disorganisation. In all species, cell shape restoration
took place immediately after returning gamonts
to the Ringer’s solution. Nevertheless, the time needed for
full recovery of gregarines along with the restoration of
their motility varied by species (or even individuals) and
applied JAS concentrations. The most rapid recovery has
been observed in G. cuneata. In contrast, gamonts of
G. steini needed much longer time, and on top of that,
some of them did not survive the experiment. In all control
preparations, the gamonts continued to move until the end
of the experiment. Interestingly, during the first 30 minutes,
in contrast to G. polymorpha and G. steini, gamonts of
G. cuneata moved more rapidly in a drop of Ringer’s solution
than was observed on microbiological agar only
slightly moistened with Ringer’s solution. Although their
movement in this period resembled regular gliding in contact
with the substrate, detailed observations revealed that
they were rather free-floating in a liquid. After 30 minutes,
gamonts of all three species sank to the surface of the
microscopic slide and started to glide in a regular way,
exhibiting the same speed of movement as observed during
the motility experiments performed on moistened agar or
in the Bürker counting chamber described above (Table 1).
Treatments with cytochalasin D (10, 20 and 30 μM in
Ringer’s solution), an inhibitor of actin polymerisation,
completely inhibited gregarine motility in a species- and
concentration dependent manner; i.e. at 10-30 minutes
in G. steini, 30-75 minutes in G. polymorpha and 75-120
minutes in G. cuneata. The cellular changes, observed
in all assays after a uniform period of 30 minutes, were
less obvious than those induced by JAS. In all species,
cell shape restoration took place immediately after
returning to the Ringer’s solution. The time needed for
full recovery and restoration of gregarine motility varied
by species and drug concentrations (Table 3).
Confocal microscopic analysis of actomyosin motor
In all three species, the homogenous distribution of the
fluorescence signal throughout the surface of phalloidinand
antibody-labelled gamonts corresponded to the localisation
of an actomyosin motor associated with the
apicomplexan cell cortex. Phalloidin labelling confirmed the
Table 1 The speed in observed gliding gregarines
Species Gliding speed in single gamonts (μm/s) Gliding speed in syzygies (μm/s)
Minimum -
maximum
Mean Standard
deviation
N. of
gamonts
N. of
records
Minimum -
maximum
Mean Standard
deviation
N. of
syzygies
N. of
records
G. cuneata 0.38 - 5.66 2.20 1.51 18 24 1.15 - 8.49 3.96 2.47 8 8
G.
polymorpha
3.46 - 22.86 9.91 5.15 12 16 1.05 - 16.18 6.73 4.08 14 17
G. steini 1.32 - 9.42 5.02 2.19 33 39 2.51 - 9.25 5.33 1.62 23 23
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Table 2 The treatment of living gregarines with jasplakinolide
Species/JAS concentration Gregarina cuneata Gregarina polymorpha Gregarina steini
5 μM JAS 10 μM JAS 20 μM JAS 30 μM JAS 5 μM JAS 10 μM JAS 20 μM JAS 30 μM JAS 5 μM JAS 10 μM JAS 20 μM JAS 30 μM JAS
Changes/time left after
drug application
(in minutes)
Initial increase of gliding
speed
≥ 5 min+ ≥ 5 min+ ≥ 5 min++ ≥ 5 min++ ≥ 5 min+ ≥ 5 min+ ≥ 5 min++ ≥ 5 min++ ≥ 5 min+ ≥ 5 min+ ≥ 5 min++ ≥ 5 min++
Decrease of gliding speed
to the normal rate
≥ 90 min+ ≥ 20 min++ ≥ 20 min++ ≥ 20 min++ ≥ 90 min+ ≥ 20 min++ ≥ 20 min++ ≥ 20 min++ ≥ 90 min+ ≥ 20 min++ ≥ 20 min++ ≥ 20 min++
Further decrease of gliding
speed
≥ 100 min+ ≥ 30 min++ ≥ 30 min++ ≥ 30 min++ ≥ 100 min+ ≥ 30 min++ ≥ 30 min++ ≥ 30 min++ ≥ 100 min+ ≥ 30 min+ ≥ 30 min++ ≥ 30 min++
Cellular changes
(shrivelling)
≥ 60 min ≥ 60 min ≥ 60 min ≥ 60 min ≥ 60 min ≥ 60 min ≥ 60 min ≥ 60 min ≥ 60 min ≥ 60 min ≥ 60 min ≥ 60 min
Complete stoppage of
gliding motility
≤ 150 min ≤ 140 min ≤ 100 min ≤ 100 min ≤ 150 min ≤ 140 min ≤ 100 min ≤ 100 min ≤ 130 min ≤ 120 min ≤ 90 min ≤ 90 min
Recovery of cell shape
after washing in Ringer’s
solution
≤ 1 min ≤ 1 min ≤ 1 min ≤ 1 min ≤ 1 min ≤ 1 min ≤ 1 min ≤ 1 min ≤ 1 min ≤ 1 min ≤ 1 min ≤ 1 min
Full recovery of motility
after washing in Ringer’s
solution
≥ 5 min ≥ 5 min ≥ 5 min ≥ 5 min ≤ 10 min ≥ 10 min ≥ 10 min ≥ 10 min ≤ 30 min ≥ 30 min ≥ 30 min ≥ 30 min
The symbol + indicates an intensity of observed change ranging from + to +++ when compared among gregarine species and JAS concentrations. Applied only if the listed phenomenon showed significant differences
in its intensity.
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Table 3 The treatment of living gregarines with cytochalasin D
Species/Cytochalasin D concentration Gregarina cuneata Gregarina polymorpha Gregarina steini
10 μM CytD 20 μM CytD 30 μM CytD 10 μM CytD 20 μM CytD 30 μM CytD 10 μM CytD 20 μM CytD 30 μM CytD
Changes/time left after drug application (in minutes)
Initial decrease of gliding speed ≥ 5 min+ ≥ 5 min++ ≥ 5 min+++ ≥ 5 min+ ≥ 5 min++ ≥ 5 min+++ ≥ 5 min+ ≥ 5 min++ ≥ 5 min+++
Further decrease of gliding speed ≥ 20 min+ ≥ 10 min++ ≥ 10 min+++ ≥ 15 min+ ≥ 10 min++ ≥ 10 min+++ ≥ 10 min+ ≥ 10 min++ ≥ 10 min+++
Cellular changes (shrivelling) ≥ 30 min ≥ 30 min ≥ 30 min ≥ 30 min ≥ 30 min ≥ 30 min ≥ 30 min ≥ 30 min ≥ 30 min
Complete stoppage of gliding motility ≤ 120 min ≤ 90 min ≤ 75 min ≤ 75 min ≤ 60 min ≤ 30 min ≤ 30 min ≤ 20 min ≤ 10 min
Recovery after washing in Ringer’s solution ≤ 1 min ≤ 1 min ≤ 1 min ≤ 1 min ≤ 1 min ≤ 1 min ≤ 1 min ≤ 1 min ≤ 1 min
Full recovery after washing in Ringer’s solution ≥ 5 min ≤ 10 min ≤ 10 min ≥ 5 min ≤ 10 min ≤ 10 min ≥ 5 min ≥ 10 min ≥ 10 min
The symbol + indicates an intensity of observed change ranging from + to +++ when compared between gregarine species and concentrations of cytochalasin D. Applied only if the listed phenomenon showed
significant differences in its intensity.
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presence of filamentous actin (F-actin) in the gregarine cell
cortex, the fibrillar septum separating the protomerite from
the deutomerite and the area of the nucleus (Figures 1B, 2B
and 3B). After treatment with 10 μM JAS for 10 minutes,
when gregarines glided with increased speed, the F-actin
staining became more bright and confined to the cell
cortex, the septum and the perinuclear space (Figures 1C-E,
2C-E and 3C-D). Higher magnification revealed numerous
transverse and oblique actin filaments in the area of epicytic
folds (Figures 1E, 2E and 3D). In addition, several aggregations
of F-actin were observed in the cytoplasm of G.
cuneata protomerite and deutomerite (Figure 1C-D). This
Figure 1 Actin and myosin in Gregarina cuneata gamonts. A. Gamonts in syzygy; primite (p), satellite (s). LM, transmitted light. B. Localisation
of F-actin in a gamont; nucleus (asterisk), septum (arrowhead) between protomerite and deutomerite. CLSM, phalloidin-TRITC. C-D. Localisation of
F-actin in gamonts (previously associated in syzygy) treated for 10 minutes with 10 μM JAS. Intense labelling is restricted to the cortex and
cytoplasmic F-actin aggregations; septum (arrowhead), nucleus (asterisk). Figure C shows a primite. CLSM (left) and merged CLSM/transmitted
light (right), phalloidin-TRITC. Figure D shows a satellite. CLSM, phalloidin-TRITC. E. The deutomerite of a gamont treated for 10 minutes with
10 μM JAS. F-actin localisation corresponds to the cortex and nucleus (asterisk). Upper two figures show the gamont middle plane; lower figure shows the
cortex in the area of epicytic folds. Merged CLSM/transmitted light (upper) and CLSM, phalloidin-TRITC. F. F-actin localisation in a gamont treated for 150
minutes with 10 μM JAS. Note the decreased labelling of cell cortex and septum (arrowhead), and formation of numerous cytoplasmic aggregations of
F-actin; nucleus (asterisk). CLSM, phalloidin-TRITC. G. The deutomerite of a gamont treated for 150 minutes with 10 μM JAS. F-actin labelling is restricted to
the cortex in the area of epicytic folds (lower); cytoplasmic F-actin aggregations (upper). CLSM, phalloidin-TRITC. H. Actin localisation in previously
associated gamonts; septum (arrowheads). CLSM, IFA. I. Actin localisation in a gamont ghost; nucleus (asterisk), septum (arrowhead). CLSM, IFA. J. Myosin
labelling in a maturing gamont. CLSM, IFA. K. Myosin labelling in a mature gamont is restricted to the cortex, but not to the septum (arrowhead). Merged
CLSM/transmitted light, IFA. L. Labelling of myosin in a gamont cortex shows a pattern of longitudinal rows; septum (arrowhead). CLSM, IFA. Figures H, I
and J show merged FITC (antibody) and rhodamine (counterstaining with Evans blue) fluorescence channels.
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was accompanied by an obvious decrease of diffuse F-actin
in all analysed gregarines. The localisation of F-actin did not
significantly change even after 150 minutes incubation in
10 μM JAS, when gregarines completely stopped their
movement and showed obvious cellular changes, but the intensity
of cell cortex labelling significantly decreased and
numerous small aggregations of F-actin appeared in the cell
cytoplasm (Figures 1F-G, 2F-G and 3E). The transverse
Figure 2 Actin and myosin in Gregarina polymorpha gamonts. A. Gamonts in syzygy; primite (p), satellite (s). LM, transmitted light. B. F-actin
localisation in a gamont; nucleus (asterisk), septum (arrowhead). CLSM, phalloidin-TRITC. C-D. F-actin localisation in gamonts (previously associated in
syzygy) treated for 10 minutes with 10 μM JAS. The more intense labelling is restricted to the cortex; septum (arrowhead), nucleus (asterisk). Figure C
shows a primite. CLSM (left) and merged CLSM/transmitted light (right), phalloidin-TRITC. Figure D shows a satellite. CLSM, phalloidin-TRITC. E. The
deutomerite of a gamont treated for 10 minutes with 10 μM JAS. F-actin localisation corresponds to the cortex; nucleus (asterisk). Upper two figures
show the gamont middle plane; lower figure shows the cortex in the area of epicytic folds. Merged CLSM/transmitted light (upper) and CLSM,
phalloidin-TRITC. F. F-actin localisation in a gamont treated for 150 minutes with 10 μM JAS, showing decreased labelling of cortex and septum
(arrowhead). Numerous cytoplasmic F-actin aggregations give the labelling homogenous appearance. CLSM, phalloidin-TRITC. G. The deutomerite
of a gamont treated for 150 minutes with 10 μM JAS; rosette-like aggregations of F-actin (arrows), nucleus (asterisk). Merged CLSM/transmitted light
(upper) and CLSM (lower), phalloidin-TRITC. H. Actin localisation in a gamont; nucleus (asterisk), septum (arrowhead). The indistinct labelling (green)
is more evident in the cortex covering the anterior part of cell. CLSM, IFA. I–J. Myosin localisation in a gamont. The labelling is restricted to the cortex,
with a pattern of longitudinal rows; septum (arrowhead). CLSM, IFA. K. The anterior part of the gamont shown in J. Myosin localisation is restricted to
the cortex, but not to the septum (arrowhead). Merged CLSM/transmitted light, IFA. L. Myosin localisation in a gamont ghost; septum (arrowhead).
CLSM, IFA. Figures H, J and L show merged FITC (antibody) and rhodamine (counterstaining with Evans blue) fluorescence channels.
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actin filaments in epicytic folds appeared to fuse into a homogeneous
layer. In G. polymorpha, rosette-like aggregations of
F-actin were situated in the cell periphery (Figure 2G).
Gamonts stained with the specific anti-actin antibody
(known to recognise the actin in Toxoplasma and
Plasmodium) exhibited a similar actin localisation, however,
in comparison with the phalloidin-stained specimens,
only slight labelling of actin was associated with the
cell cortex and the septum in G. cuneata (Figure 1H–I)
and G. polymorpha (Figure 2H). Gamonts of G. steini
Figure 3 Actin and myosin in Gregarina steini gamonts. A. Gamonts associated in syzygy; primite (p), satellite (s). LM, transmitted light.
B. Localisation of F-actin in three single gamonts; nucleus (asterisk), septum (arrowheads). CLSM, phalloidin-TRITC. C. Localisation of F-actin in
gamonts associated in syzygy treated for 10 minutes with 10 μM JAS. The intense labelling is restricted to the cortex, septum (arrowheads) and
nucleus (asterisks). CLSM (left) and merged CLSM/transmitted light (right), phalloidin-TRITC. D. The deutomerite of a gamont treated for 10
minutes with 10 μM JAS. The localisation of F-actin corresponds to the cortex. Upper two figures show the gamont middle plane; lower figure is
a view of the cortex in the area of epicytic folds. Merged CLSM/transmitted light (upper) and CLSM, phalloidin-TRITC. E. Localisation of F-actin in
gamonts associated in syzygy treated for 150 minutes with 10 μM JAS. Note the decreased labelling of cell cortex and septum (arrowheads);
nucleus (asterisks). Numerous indistinct, small cytoplasmic aggregations of F-actin give the labelling a more homogenous appearance. CLSM, phalloidinTRITC.
F. Actin localisation in gamonts associated in syzygy; nucleus (asterisks). The septum (arrowhead) exhibits an intense labelling. CLSM, IFA. G.
Myosin labelling in gamonts associated in syzygy; septum (arrowhead). CLSM, IFA. H. An optical plane of the gamont cortex showing the distribution of
myosin corresponding to the epicytic folds; septum area (arrowhead). CLSM, IFA. I. Myosin localisation in a single gamont. The labelling is restricted to
the cortex and exhibits a pattern of longitudinal rows. A slight labelling in the septum area is shown (arrowhead). Merged CLSM/transmitted light (left)
and CLSM (right), IFA. Figures F, G and H show merged FITC (antibody) and rhodamine (counterstaining with Evans blue) fluorescence channels.
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exhibited a higher intensity of actin labelling with this
antibody, and gamonts associated in syzygy labelled with a
different intensity in that a higher concentration of actin
was observed in the primite (Figure 3F).
Similarly to actin, the myosin was restricted to the cell
cortex (Figures 1J–L, 2I–L and 3G–I), but no specific labelling
corresponding to the septum was observed. When
focussing on the gregarine surface, the organisation of myosin
exhibited a pattern of longitudinal rows corresponding
to the epicytic folds (Figures 1L, 2I–J and 3H-I). The
primites and satellites in syzygies (Figure 3G) exhibited
more or less identical intensity of labelling.
The results of immunofluorescent labelling using an antiα-tubulin
antibody were negative (data not shown) in
agreement with the absence of subpellicular microtubules.
Mucus shedding
Only phase contrast microscopy was able to show
mucus shedding and trail formation of gamonts gliding
on agar (Figure 4A–F). The most evident mucous trails
were that of associated (Figure 4C) or single (Figure 4D)
gamonts of G. polymorpha due to the distance travelled
by them, which left long and regular mucous paths. In
contrast, G. cuneata (Figure 4A) and G. steini (Figure 4E)
gliding gamonts exhibited irregular and short paths
containing greater accumulations of mucus, especially
when these species were avoiding barriers or altering
direction (Figure 4B and F).
The mucous substances were visualized by Alcian blue
staining within the gamont cytoplasm (Figure 4G–I).
The most intense labelling occurred in the deutomerite
cytoplasm of G. polymorpha (Figure 4H). In contrast to
G. cuneata (Figure 4G) and G. steini (Figure 4I), this
gregarine showed an increased volume of mucus in the
protomerite cytoplasm, which might be correlated with
the increased motility of the G. polymorpha protomerite.
Ultrastructural features of the pellicle
The pellicle in all the species was organised in numerous
longitudinal narrow folds, which were more raised in the
deutomerite region than in the protomerite. The epicytic
folds of the deutomerite were more or less undulated,
depending on the species. Similarly, the number of
epicytic folds per square micrometer varied among species,
and their number did not significantly changed in
the course of gamont growth (Table 4). The gamonts
G. cuneata were covered by almost linear folds and numerous
mucus-like drops were often present in the grooves
separating them (Figure 5A–F). Occasionally, rings of undulated
folds could be observed, especially in the posterior
half of the gamont deutomerite (Figure 5E). The pellicle
appeared to be slightly helically coiled along the gregarine
longitudinal axis, resulting in a helical course of epicytic
folds (Figure 5A, the syzygy on the right). The pellicle
of G. polymorpha gamonts exhibited zones of almost
linear folds alternating with zones of much undulated
folds; however, surprisingly when considering the results of
the Alcian blue staining, only a few mucus drops could be
detected (Figure 6A–E). The epicytic folds of G. steini were
undulated in a more regular pattern than in G. polymorpha,
and numerous mucus-like drops covered the entire surface
of the gamonts (Figure 7A–G). Depending on the species,
more or less evident constriction could be found at the
interface between the protomerite and deutomerite; however,
no interruption of folds, running from the gamont
apical to its posterior end, was present in this area
(Figures 5A, 6A and 7A). Syzygies were caudo-frontal,
i.e. the posterior end of the primite deutomerite was
joined with the apical part of the satellite protomerite
(Figures 5A, 6A and 7A). The connection area appeared
as a collar-like junction composed of modified epicytic
folds of the primite deutomerite meshing in a gear-like
manner with the folds of the sucker-like apical region
of the satellite (Figures 5B–C, 6B–C and 7B–D). In
G. cuneata, two or more satellites were often found to be
associated with one primite (Figure 5A and E). In some
cases, a large primite was associated with several tiny satellites
(up to four satellites associated with one primite were
observed). Occasionally, three individuals of G. cuneata
were seen to be associated in a row, the last one of which
was the smallest.
The folded pellicle was three-layered, composed of the
superficial plasma membrane covering the entire gregarine
and a middle lucent region, underlined by two distinct
and tightly apposed membranes, i.e., external and
internal cytomembrane, forming the inner membrane
complex (IMC). Epicytic folds emerged from the peripheral
ectoplasm bounding the endoplasm. Three types of
associated structures were constantly present in each
fold: an internal lamina, 12-nm filaments and rippled
dense structures. The internal lamina, running under the
IMC, not only linked the bases of epicytic folds but also
bifurcated just beneath each fold, and its thinner part extended
to the individual folds (Figures 5G, 6G and 7I).
The localisation along with the organisation of internal
lamina suggest its function in the stabilisation of individual
folds as well their interconnection. The thickness of
the internal lamina varied by species; i.e., 50–75 nm in
G. polymorpha (Figure 6F–G), 17–30 nm in G. cuneata
(Figure 5G–H) and 8–11 nm in G. steini (Figure 7H–I).
The compact organisation of the internal lamina usually
disappeared when reaching the region of 12-nm filaments.
In fact, the 12-nm filaments seemed to be embedded in
the widened area of the internal lamina (Figures 5G, 6G
and 7I). The 12-nm filaments, exhibiting the properties
of intermediate filaments, ran under the IMC along
the longitudinal axis of each fold. Their numbers varied
by species and developmental stage, i.e., in gamonts of
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Figure 4 Gliding motility and mucus. A–F. Mucous trail (arrowheads) left behind gliding gregarines; syzygy of Gregarina cuneata (A, B),
syzygy (C) and single gamont (D) of Gregarina polymorpha, syzygy of Gregarina steini (E, F). Note the increased mucus shedding by the syzygy of
G. cuneata exhibiting rotary movement (B). G–I. Light micrographs showing the volume of mucus in gamonts of G. cuneata (G), G. polymorpha
(H) and G. steini (I) revealed with Alcian blue staining at low pH.
Table 4 The number of epicytic folds in deutomerite of gamonts
Species G. cuneata G. polymorpha G. steini
Gamont N. Number of
folds/μm2
Deutomerite
perimeter (μm)
Number of
folds/μm2
Deutomerite
perimeter (μm)
Number of
folds/μm2
Deutomerite
perimeter (μm)
1 4.5 53.4 4.3 94.3 5.6 31.4
2 4.4 98.1 4.2 103.7 6.3 34.6
3 3.9 113.1 4.9 106.8 6.3 37.7
4 3.6 114.0 4.3 110.0 5.9 58.0
5 3.2 150.9 4.5 132.0 5.0 66.0
6 3.3 163.4 3.6 144.5 4.0 113.1
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G. cuneata up to 7 (Figure 5G) and in G. polymorpha
up to 10 filaments were observed (Figure 6G), while in
G. steini only 2–4 filaments could be seen (Figure 7I).
Their number increased with fold maturation, i.e., new
rising folds contained fewer filaments than the older
and evidently higher epicytic folds. Rippled dense structures,
located between the plasma membrane and IMC,
appeared as electron-dense, triangle-shaped structures
Figure 5 Pellicle architecture in Gregarina cuneata gamonts. A. Gamonts associated in syzygy; primite (p), satellite (s). SEM. The syzygy on the
right is composed of one primite (p) and two satellites (s1, s2). B. Higher magnification of the junction between the posterior end of the primite
deutomerite (d) and the apical part of the satellite protomerite (p). SEM. C. A detail of the junction (arrow) between folded pellicles covering the
primite deutomerite (def) and satellite protomerite (pef). SEM. D. Organisation of linear epicytic folds covering the deutomerite. SEM. E. Higher
magnification of the junction between the primite (p) and two satellites (s1, s2) shown in panel A. SEM. F. A higher magnification of deutomerite
epicytic folds (ef); grooves (g) between folds, mucus drops (*). SEM. G. Cross section of deutomerite epicytic folds; grooves (g) with mucus (*)
between folds (ef), 12-nm filaments (arrowhead), inner membrane complex (imc), internal lamina (double arrowhead), plasma membrane (pm),
rippled dense structures (white arrowhead), unknown dense structure (arrow). TEM. H. Cross section of deutomerite epicytic folds (ef); internal
lamina (double arrowhead), rib-like myonemes (arrowheads). TEM. I. Detailed view of an epicytic fold in cross section revealing filamentous
connections (arrowheads) localised between the plasma membrane (pm) and inner membrane complex (imc). TEM. J. Cross section showing the
organisation of the deutomerite pellicle; ectoplasmic network (arrows), epicytic folds (ef), duct (*), rib-like myonemes (arrowheads). The insect
shows the micropore located in the groove between two epicytic folds. TEM. K. Organisation of the pellicle and ectoplasmic network (n) during
gregarine movement; epicytic folds (ef), deutomerite (d), protomerite (p). TEM. L. Higher magnification of a septum (arrow) separating the
protomerite (p) from the deutomerite (d); epicytic folds (ef). Note the ectoplasmic network (n) connected to the septum. TEM.
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with base lying on the external cytomembrane and median
running in between two adjacent 12-nm filaments. Their
number varied by species and developmental stages,
in correlation with the number of 12-nm filaments
(Figures 5G, 6G and 7I). Unknown dense and usually
half-moon-shaped structures underlined the 12-nm
filaments at their cytoplasmic face in all gregarines. This
structure achieved its maximum length in G. polymorpha,
in which its ends were in obvious contact with the internal
lamina extending to the top of the fold (Figure 6G). In
G. cuneata, this structure was evidently shorter and
thicker (Figure 5G), and in G. steini it was shortest and in
Figure 6 Pellicle architecture in Gregarina polymorpha gamonts. A. Gamonts associated in syzygy; primite (p), satellite (s). SEM. B. Higher
magnification of the junction between the posterior end of the primite deutomerite (d) and apical part of the satellite protomerite (p). SEM.
C. Detailed view of the junction (arrows) between folded pellicles covering the primite deutomerite (def) and the satellite protomerite (pef). SEM.
D. Organisation of undulated epicytic folds covering the deutomerite. SEM. E. Higher magnification of deutomerite epicytic folds (ef); grooves (g)
between folds. SEM. F. Cross section of the deutomerite pellicle; epicytic folds (ef), grooves (g), internal lamina (double arrowhead), rib-like
myonemes (arrowhead). TEM. G. Cross section of deutomerite epicytic folds; 12-nm filaments (arrowhead), inner membrane complex (imc),
internal lamina (double arrowhead), plasma membrane (pm), rippled dense structures (white arrowhead), unknown dense structure (arrow). TEM.
H. Detailed view of epicytic folds in cross section revealing filamentous connections (arrowheads) localised between the plasma membrane (pm)
and inner membrane complex (imc). TEM. I. Pellicle organisation revealed in a mechanically ruptured gamont; cytoplasm (c) with ectoplasmic
network, epicytic folds (ef). SEM. J. The view of an ectoplasmic face (*) of a pellicle separated from the gamont cytoplasm; epicytic folds (ef). SEM.
K. The septum (arrow) separating the protomerite (p) from the deutomerite (d); epicytic folds (ef). TEM.
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some sections it was even reversed with its ends facing
the apical top of the fold (Figure 7I). As the internal
lamina lacked its typical compact look in this area, it
is possible that the mentioned half-mooned structures
represent its component. Careful analysis of the pellicle
covering the lateral part of epicytic folds revealed novel
thin filamentous connections interconnecting the IMC
and the plasma membrane (Figures 5I, 6H and 7I).
In addition to structures restricted to the epicytic
folds, an ectoplasmic network and rib-like myonemes,
present to some degree in the deutomerite ectoplasm of
all studied species, could be observed in some ultrathin
Figure 7 Pellicle architecture in Gregarina steini gamonts. A. Gamonts associated in syzygy; primite (p), satellite (s). SEM. B. Higher
magnification of the junction between the posterior end of the primite deutomerite (d) and apical part of the satellite protomerite (p). SEM.
C, D. Detailed views of the junction (arrow) between folded pellicles covering the primite deutomerite (def) and satellite protomerite (pef). SEM.
E, F. Organisation of more and less undulated epicytic folds covering the deutomerite. SEM. G. Higher magnification of deutomerite epicytic folds
(ef); grooves (g) between folds, pore-like structure (arrow). SEM. H. Organisation of the deutomerite pellicle in cross section; epicytic folds (ef),
grooves with mucus (*), internal lamina (double arrowhead). TEM. I. Cross section of deutomerite epicytic folds. Note the filamentous connections
(white arrows) localised between the plasma membrane (pm) and inner membrane complex (imc); 12-nm filaments (arrowhead), rippled dense
structures (white arrowhead), unknown dense structure (arrow). TEM. J. The septum (arrow) and ectoplasmic network (n); deutomerite (d),
epicytic folds (ef), protomerite (p). TEM. K. Golgi apparatus in deutomerite cytoplasm. TEM. L. The duct (arrow) opening outwards to the groove
between epicytic folds. TEM.
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sections. The ectoplasmic network contacting the bases
of the epicytic folds was most prominent in fully matured
gamonts of G. cuneata (Figure 5J–L), especially at
the septum periphery or in areas with an obviously
pleated pellicle (Figure 5K). This local pleating of the
pellicle seemed to be the result of gregarine movement.
The rib-like myonemes, running perpendicularly to the
longitudinal axis of the gregarine and located beneath the
deutomerite ectoplasm, were very distinct in gamonts
of G. cuneata (Figure 5H and J) and G. polymorpha
(Figure 6F), but were hard to detect and often absent
in G. steini. The septum separating the protomerite from
the deutomerite was well developed in all three species
(Figures 5L, 6K and 7J). The micropores, interrupting
the IMC, were sometimes seen in the grooves between
the folds of G. cuneata (Figure 5J). In addition, ducts,
appearing as elongated dense sacs passing through the
pellicle and opening outwards, were often present in the
ectoplasm of G. cuneata and G. steini gamonts (Figures 5J
and 7L).
Membranes as exposed by freeze etching
The freeze-etching technique confirmed the presence of
three fracture planes in the pellicle of all analysed gregarines,
corresponding to the three membranes, i.e., two
cytomembranes (IMC) underlying the plasma membrane.
The general aspects of the epicytic folds in freezefracture
are shown in Figures 8, 9 and 10, and their
architecture corresponded to the TEM observations. At
the tip of each fold, double linear rows of tightly aligned
intramembranous particles (IMP) were observed in both
fractures of the external and internal cytomembranes.
These rows seemed to correspond to the apical site of
the 12-nm filaments and to the base of the triangleshaped,
rippled dense structures. The number of these
lines roughly coincided with the number of 12-nm filaments.
The freeze-etching approach proved to be a
strong tool for visualisation of structures that are hardly
documented on TEM micrographs including round
micropores (110–130 nm in diameter) at the base of
the grooves between the folds (Figures 8A, 8G, 9E, 9G
and 10E) or smaller pores (35–50 nm in diameter) randomly
distributed on the base or on the lateral side of the
folds (Figures 8A–D, 8F-G, 9A–E, 9G, 10A, 10C and
10E–F), ducts and cisternae often connected these pores
or to the pellicle (Figures 8B, 8E, 8H, 9F–G and
10A), as well as mucus drops often present in the grooves
between the folds of G. cuneata (Figure 8B) and G. steini
(Figure 10A–D and F).
The densities of the IMP for the fracture faces of the
plasma membrane and both cytomembranes are
summarised in Table 5. The density of the IMP in membranes
differed in analysed gregarines; in general, the
values in G. polymorpha were significantly lower in
comparison to G. cuneata and G. steini. In G. cuneata
and G. polymorpha, the IMP density in the IMC was
lower than in the plasma membrane. In G. steini, however,
the IMP density in the plasma membrane was
lower than those in the IMC. The IMP in all three membranes
showed a high variability in their size distribution
(see histograms in Figures 11, 12 and 13); nevertheless,
only particles in a range of 6–14 nm were included in
statistical calculations to obtain data comparable with
those previously published on other apicomplexans.
Membranes forming the pellicle in G. cuneata, especially
the plasma membrane, were the most extraordinary
given the wide range of the IMP size along with its distribution
(Figure 11).
Discussion
Gliding movement is a feature observed in a wide range
of unicellular organisms including diatoms, flagellates,
apicomplexan zoites, and gregarines. The speed of these
organisms varies along with the special locomotive
structures and is affected mostly by their physiological
status and surrounding environmental conditions. The
reported motility rates in Apicomplexa are usually in the
range of 1–10 μm/s, and the maximal rate was observed
in gregarines [34]. To minimise a potential effect of different
environmental conditions on gregarine motility in this
study, we took advantage of a naturally mixed infection
with three Gregarina species occurring in the intestine of
larval mealworms kept under laboratory conditions. In
addition, the experimental part of this work, including the
light microscopic observations on gliding and treatment
of living gamonts with JAS and cytochalasin D, has been
performed on suspensions consisting of all three species
(often from a single host). Göhre [35] states that gregarines
parasitising the intestine of larval T. molitor are distributed
based on intestinal pH, i.e., G. cuneata inhabits a
part of the intestine with pH 4.5–5.5, and G. steini
(pH 5.5–8.2) inhabits a part of the intestine together with
G. polymorpha (pH 6.4–7.5). Therefore, it could be
expected that the mixed suspensions of gamonts are not
preferable for motility assays, nevertheless, pH-restricted
localisation of gregarines in mealworms applies only to attached
trophozoites [36] not to gamonts that are usually
found in luminal part of the mesenteron. During our observations,
eugregarines isolated from one host were gliding
at speeds ranging from 0.38 to 22.86 μm/s, while the
highest speed was reached by gamonts of G. polymorpha
and the lowest by G. cuneata. To explain these evident
differences study focussed on structures that were generally
considered to be responsible for gregarine gliding epicytic
folds and mucus. Longitudinal folds formed by
the pellicle represent the most conspicuous feature differentiating
eugregarine trophozoites and gamonts from the
other apicomplexans. The presence of the swellings along
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the longitudinal epicytic folds of gliding gregarines suggests
that these structures, pushed against the substrate,
may provide the force for gliding [33], and the mucus
seems to enhance the efficiency of their interaction with
the substrate to produce gregarine forward movement. It
is important, however, to mention that gregarine gamonts
in performed experiments were able to move without any
contact with the substrate when put in a drop of Ringer’s
Figure 8 Pellicle organisation in Gregarina cuneata gamonts as revealed by the freeze-etching. A. The general view of fractured epicytic
folds; cytoplasm (c), micropore (encircled), cytoplasm of folds (*), groove (g), EF face of external cytomembrane (Ee), EF face of internal
cytomembrane (Ei), IMP alignments (arrowhead), PF face of external cytomembrane (Pe), PF face of internal cytomembrane (Pi), plasma
membrane (arrow), pores (white arrows). B. The base of the epicytic folds; cytoplasm of folds (*), deutomerite cytoplasm (c), duct (encircled),
grooves (g), mucus (x), PF face of the external cytomembrane (Pe), pores (white arrows). C, D. The fracture of the epicytic fold; cytoplasm (c),
EF of the external cytomembrane (Ee), EF face of the internal cytomembrane (Ei), PF face of the external cytomembrane (Pe), PF face of the
internal cytomembrane (Pi), pores (white arrows), IMP alignments (arrowheads). E. The base of the fold with a duct opening outwards (arrow) to
the groove; deutomerite cytoplasm (c), EF of the external cytomembrane (Ee), PF face of the external cytomembrane (Pe), PF face of the internal
cytomembrane (Pi). F. The longitudinal fracture of the fold; cytoplasm (c), cytoplasm of fold (*), PF face of the external cytomembrane (Pe),
PF face of the internal cytomembrane (Pi), pores (white arrows). G. The grooves (g) between the folds (ef); cytoplasm (c), mucus (x), numerous
pores (some of them are shown by white arrows), PF face of the external cytomembrane (Pi). The inset shows a micropore and a pore of smaller
size. H. A detail of the grove between folds (ef) showing the part of micropore (arrow) with vesicle (v); deutomerite cytoplasm (c). I. The top of
the fold; EF face of the plasma membrane (Ep), IPM alignments (arrowheads), PF face of the external cytomembrane (Pe). The arrowhead in the
circle shows the direction of shadowing.
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solution, i.e., for approximately 30 min, they were floating
in a liquid until sinking to the surface of microscopic slide.
These observations conflict with the supposed need of
their contact with some solid matter [37]. Although some
correlations can be found between speed and the number
of epicytic folds, i.e., gamonts of G. cuneata are quipped
by fewer folds (3.2–4.5) per square micrometer than small
gamonts of G. steini (4.0–6.3) or similarly-sized gamonts
Figure 9 Pellicle organisation in Gregarina polymorpha gamonts as revealed by the freeze-etching. A. The general view of fractured
epicytic folds; cytoplasm of folds (*), deutomerite cytoplasm (c), EF face of external cytomembrane (Ee), IMP alignments (arrowhead), groove (g),
PF face of external cytomembrane (Pe), PF face of internal cytomembrane (Pi), pores (white arrows). B. The fractured epicytic fold; deutomerite
cytoplasm (c), EF face of the internal cytomembrane (Ei), groove (g), PF face of the external cytomembrane (Pe), pores (white arrows). C. The
fracture of the lower epicytic fold; cytoplasm of folds (*), deutomerite cytoplasm (c), EF face of the internal cytomembrane (Ei), EF face of the
plasma membrane (Ep), IMP alignments (arrowhead), PF face of the internal cytomembrane (Pi), pore (white arrow). D. The fracture of higher
epicytic folds; cytoplasm of folds (*), deutomerite cytoplasm (c), EF of the internal cytomembrane (Ei), EF of the plasma membrane (Ep), groove
(g), IMP alignments (arrowhead), PF face of the internal cytomembrane (Pi), pore (white arrow). The inset shows the IMP alignments located on
the EF face of the external cytomembrane (white arrowhead) and on the PF of the internal cytomembrane (arrowhead) at the top of the epicytic
fold. E. The grooves (g) between epicytic folds with numerous pores (some of them shown by white arrows); cytoplasm of folds (*), deutomerite
cytoplasm (c), EF face of the internal cytomembrane (Ei), PF face of the external cytomembrane (Pe). Note the large opened micropore (arrow).
The inset shows a micropore and three pores of different sizes. F. The base of the epicytic fold (ef) with a duct (encircled); cytoplasm (c). G. The
base of the epicytic folds showing the micropore connected to a vesicle (encircled); cytoplasm (c), pores (white arrows). The arrowhead in the
circle shows the direction of shadowing.
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of G. polymorpha (3.6–4.9), these differences were only
slight and we do not consider them to significantly influence
the gliding rate. More important, however, seem to be
undulations of folds documented by SEM. In accordance
with Vávra and Small [24], the folds of glutaraldehyde-fixed
G. cuneata were almost linear, forming occasional rings of
undulated folds, while the pellicle of G. polymorpha formed
zones of undulated as well as almost linear folds, and the
folds in G. steini were undulated in quite a regular pattern.
Microscopic observations indicate that the lateral undulations
of folds arise during gliding. Nevertheless, the question
of whether epicytic folds might be responsible for
Figure 10 Pellicle organisation in Gregarina steini gamonts as revealed by the freeze-etching. A–C. Fractured epicytic folds; cytoplasm
of epicytic folds (*), deutomerite cytoplasm (c), ducts (encircled), EF face of the external cytomembrane (Ee), EF face of the internal
cytomembrane (Ei), EF face of the plasma membrane (Ep), groove (g), mucus (x), PF face of the external cytomembrane (Pe), PF face of the
internal cytomembrane (Pi), PF face of the plasma membrane (Pp), pores (white arrows). D. General view of the epicytic folds showing the
grooves (g) with mucus drops (x); cytoplasm of epicytic folds (*), deutomerite cytoplasm (c). E. The cytoplasmic face of the grooves between
epicytic folds with micropores and numerous pores (some of them shown by white arrows); cytoplasm of epicytic folds (*), EF face of the internal
cytomembrane (Ei), PF face of the external cytomembrane (Pe). The inset shows the detailed view of micropore and four pores of different sizes.
F. Fractured epicytic folds showing their IMP alignments (arrowheads) located on the PF of the internal cytomembrane (Pi) and on the EF face
of the external cytomembrane (Ee); cytoplasm of epicytic folds (*), EF face of internal cytomembrane (Ei), grooves (g), PF face of external
cytomembrane (Pe), pores (white arrows). The inset shows EF of the plasma membrane (Ep) with mucus drops (x). The arrowhead in the circle
shows the direction of shadowing.
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gliding in gregarines can be satisfactorily answered only
after careful analysis of all subcellular components forming
the complicated pellicle in gregarines.
The ‘glideosome’ concept and eugregarine gliding
First, it must be highlighted that no similar structures
resembling epicytic folds can be found on the surface
of the zoite stage (e.g., sporozoites in gregarines or
Plasmodium, tachyzoites of Toxoplasma), which are
used to illustrate the mechanism of gliding motility in
apicomplexans. The concept of the ‘glideosome’ [4] describes
apicomplexan zoites as actively entering host cells
and moving by a substrate-dependent gliding motility,
which requires coordinated interactions between parasite
surface adhesins and its cytoskeleton. This machinery is
considered an unusual form of eukaryotic locomotion.
The so-called actomyosin motor, which is generally assumed
to be embedded between the plasma membrane
and the IMC, consists of immobilised unconventional
myosins, short actin stubs, and TRAP-family invasins.
This motor is expected to be oriented by subpellicular microtubules
[38]. Micronemal proteins, inserted into the
plasma membrane, are carried along the IMC by the
motor and interact with the parasite substrate, or associate
with a GPI-anchored protein interacting with the substrate,
resulting in gliding [38].
Considering the possible application of this concept
for eugregarine gliding, the first striking inconsistency is
the fact that in the eugregarines analysed here, there are
no subpellicular microtubules in the epicytic folds (confirmed
also by negative results of immunolabelling).
Thus, a question arises concerning the real motor in their
motility. It could be expected that enigmatic 12-nm filaments,
running under the IMC and exhibiting the properties
of intermediate filaments [34,39], could support the
actomyosin motor in a similar way. Longitudinal arrays
of IMP found in the area of the 12-nm filaments and
the rippled dense structures [11,22,30,40] are comparable
to the lines of higher particle density overlaying the
subpellicular microtubules in Eimeria or Plasmodium
sporozoites [41,42]. Our data show, however, that the
number of 12-nm filaments does not influence the speed
of gregarine gliding (up to 7 filaments in G. cuneata vs. 10
filaments in G. polymorpha and maximally 4 filaments in
G. steini), but rather seem to control the direction of
movement. Indeed, despite folds equipped by a low number
of 12-nm filaments, gamonts of G. steini glided with
relatively high speed, but their gliding path was rather
widely semi-circular than linear. The question remains
whether apical rippled dense structures, with their base located
at the external cytomembrane, serve as supporting
elements interconnecting 12-nm filaments and the plasma
membrane. Such speculation is supported by the existence
of filamentous interconnections occasionally observed between
their tips and the plasma membrane in ultrathin
sections [39]. The half-moon-shaped dense structure
underlining the 12-nm filaments could play the role of a
‘skeleton’ reinforcing the tips of folds, which contact the
substrate during gliding.
The apicomplexan gliding motility relies on the dynamic
turnover of actin, the polymerisation of which is
controlled by a number of regulators. A general model
for the organisation of the apicomplexan actomyosin
motor depicts actin filaments lying in the space between
the parasite IMC and plasma membrane, parallel to the
plasma membrane [43]. In gregarines, actin is generally
expected to be localised in the epicytic folds [20,27].
Using a specific anti-actin antibody known to recognise
the actin in Toxoplasma and Plasmodium, actin in all three
gregarine species was localised. The unusual nature of
apicomplexan actin, where its unpolymerised form seems
to have an increased potential to form filaments relative to
vertebrate actin, is discussed elsewhere [44]. In contrast to
other eukaryotic cells, which maintain comparable amounts
of globular and F-actin, in Toxoplasma more than 97% of
actin is found in its globular form and in Plasmodium,
where more F-actin may be recovered, the filamentous
fraction appears to represent a collection of short polymers
[27]. The apparent lack of visible, stable filaments in
apicomplexans, however, does not fit eugregarines, in
Table 5 A protein particle density in the fracture faces of the membranes of studied gregarines
Species Gregarina cuneata Gregarina polymorpha Gregarina steini
Membrane Face Density = number of
particles/μm2
(± SE)
Kp Density = number of
particles/μm2
(± SE)
Kp Density = number of
particles/μm2
(± SE)
Kp
Plasma membrane PF 2244 ± 283 0.81 1446 ± 158 0.59 2265 ± 154 1.27
EF 2770 ± 96 2473 ± 147 1783 ± 233
External
cytomembrane
PF 1420 ± 190 1.13 602 ± 265 0.70 2588 ± 189 0.68
EF 1260 ± 211 863 ± 202 3820 ± 211
Internal
cytomembrane
PF 1993 ± 253 1.33 1276 ± 200 1.57 2339 ± 132 1.24
EF 1502 ± 273 814 ± 246 1886 ± 274
The size of IMP is in range 6-14 nm.
Kp partition coefficient defined as the ratio of number of particles per μm2
in the PF face/number of particles per μm2
in the EF face.
SE standard error.
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which the phalloidin labelling revealed that the presence
of F-actin does not require filament-stabilising drugs
[45,46]. Still, the results of JAS (a toxin that stabilizes actin
filaments and induces actin polymerisation) and cytochalasin
D (disrupts actin filaments and inhibits actin polymerisation)
treatments are the clear evidence of an essential
role of actin in eugregarine gliding. Both probes are
membrane-permeable probes and thus suitable for examining
actin dynamics in living cells, and are known to
disrupt Toxoplasma motility and invasion [1]. The treatment
of Toxoplasma tachyzoites with 1–2 μM JAS
inhibited their gliding and cell invasion, but once the drug
Figure 11 Histograms illustrating the IMP size distribution in Gregarina cuneata; protoplasmic (PF) and exoplasmic fracture (EF) faces
of the plasma membrane (PM), external cytomembrane (EC), internal cytomembrane (IC). The mean diameter (nm) of IMP, with its
standard error, is given for each. Ordinate - number of particles; abscissa - particle diameter in nm.
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was removed the parasites were able to invade host
cells [43]. Furthermore, JAS treatment increased the rate
of Toxoplasma gliding [43], indicating that filaments are
rate-limiting for motility and also caused frequent reversals
of direction [3]. In Eimeria sporozoites, the inhibitor
of actin polymerisation, cytochalasin B, reversibly
inhibited the gliding; nevertheless, the bending was only
slightly less [47]. In agreement with these studies, the
treatment of living gregarines with JAS and cytochalasin
D suspended their gliding motility, and they were able to
recover after returning to normal physiological conditions
in insect saline. In spite of high doses of both probes used
Figure 12 Histograms illustrating the IMP size distribution in Gregarina polymorpha; protoplasmic (PF) and exoplasmic fracture (EF)
faces of the plasma membrane (PM), external cytomembrane (EC), internal cytomembrane (IC). The mean diameter (nm) of IMP, with its
standard error, is given for each. Ordinate - number of particles; abscissa - particle diameter in nm.
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in this study, prolonged incubations were necessary to inhibit
gregarine gliding completely. Similar to Toxoplasma
tachyzoites [43], JAS application led to an initial increased
gliding activity, which gradually decreased until
complete blocking. In contrast, reversals or changes of
gliding direction and apical protrusion were not observed
in gregarine gamonts. Interestingly, an enhanced deposition
of actin, resembling an apical protrusion, occurred
on the apical end of the migrating trophozoites of
eugregarine Ascogregarina [48].
High concentrations of JAS can increase the density of
actin filaments adjacent the plasma membrane [49].
Figure 13 Histograms illustrating the IMP size distribution in Gregarina steini; protoplasmic (PF) and exoplasmic fracture (EF) faces of
the plasma membrane (PM), external cytomembrane (EC), internal cytomembrane (IC). The mean diameter (nm) of IMP, with the standard
error, is given for each. Ordinate - number of particles; abscissa - particle diameter in nm.
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Despite studies reporting competitive binding of JAS
and phalloidin with F-actin [50], this work also revealed
an increase in F-actin labelling in the cell cortex after a
short treatment of living gamonts with JAS (corresponding
to the period of their increased motility). These
observations are supported by another study reporting
that washing of cells before fixation and staining with
phalloidin-TRITC to remove JAS revealed brighter staining
of F-actin [51]. Prolonged treatment with JAS (until
complete inhibition of gliding), however, resulted in an
obvious decrease in F-actin labelling. These data concur
with studies reporting that treatment of living cells with
JAS causes a redistribution of their actin cytoskeleton,
formation of F-actin aggregates and cell shape change,
and can result in a patchy appearance of cortical actin
[52]. Observations on gregarines during cytochalasin D
treatment are supported by another study in which
gliding was inhibited by cytochalasin B [53]. With regards
to the Ca++
activation of the actomyosin system, it is
interesting that the anti-psychotic drug trifluoperazine
can inhibit gregarine gliding. This observation suggests
that the calcium-binding protein calmodulin might
affect motility even though extracellular Ca++
is not
required [53].
Although the localisation of actin seems to be more
diffuse in gregarines, myosin seems to be organised in
longitudinal rows corresponding to epicytic folds similar
to observations in G. blaberae [54]. The micrographs
obtained with the anti-myosin antibody (smooth and
skeletal, the whole antiserum from Sigma-Aldrich, Czech
Republic) showed similar localization of myosin as obtained
by Heintzelman with specific antibodies directed against
the myosins A, B and F [20,27]. Nevertheless, the commercial
antibody used in this study is developed for immunofluorescence
and in the absence of conclusive results from
Western Blotting this point needs further investigation.
The presence of previously unreported tiny filamentous
connections observed in some ultrathin sections between
the plasma membrane and the IMC suggests that the actomyosin
complex could be restricted to the lateral parts of
the epicytic folds and, thus, this could be the source of the
lateral undulations described above.
Subcortical filamentous structures
In the course of environmental adaptations, the cortex
of eugregarines became very rigid and, hence, they lost the
wriggling ability of archigregarines. Nonetheless, when
considering active movements of the protomerite, the cellular
plasticity must be relatively high. Indeed, additional
but very prominent forms of motility, such as bending,
curving or shortening of the longitudinal axis and intense
movements of the protomerite, commonly observed in
eugregarines, were attributed to the presence of contractile
elements entitled ‘myocyte’ as early as one hundred
years ago [37]. A network of intermediate filamentous
proteins can be often found associated with the gregarine
cortex. Early studies reporting the filamentous character
of eugregarine ‘myonemes’ suggest that they may consist
of actin microfilaments [55], and recent studies confirmed
them to be actin-rich [20]. An additional robust population
of actin filaments, forming a series of annular
(rib-like) myonemes girding the cell cortex, was reported
from G. polymorpha but not seen in other apicomplexans
[20,27]. Actin and myosin A were detected in both the
epicytic folds and rib-like myonemes, while myosin B was
exclusively restricted to folds [20]. Later on, a WD40
repeat-containing myosin designed myosin F, unique to
the Apicomplexa and associated with rib-like myonemes,
was reported in G. polymorpha [27]. Interestingly, the
bending of the protomerite that is expected to be related
to the ectoplasmic network and rib-like myonemes occurred
in G. polymorpha and G. cuneata gamonts, which
are indeed proven to possess these structures, while the
stiff gamonts of G. steini missing the rib-like myonemes
on ultrathin sections showed no shape changes during
their rapid gliding. Despite the work reporting myosinand
actin-like proteins restricted to the vegetative stages
of G. blaberae [54], the presence of these proteins was
documented in both the gamonts as well as the trophozoites
of Gregarina representatives [45,46]. Nevertheless, we
do not exclude that there may exist some correlation
between the abundance or form of actin and gregarine
developmental stage.
Shedding of mucous material
The next point that is worth noting with regard to the
glideosome is the lack of micronemes in gregarine
gamonts. Eugregarine gliding resembles the gliding motility
in sporozoites of Plasmodium [56]. The trail left behind
gliding ookinetes of Plasmodium was shown to correspond
to the release of the Pbs25 and the circumsporozoite
thrombospondin-related protein (CTRP) [57]. The material
observed in the trail left after gliding eugregarines is generally
designated as mucus, but more detailed biochemical
analyses are needed to determine the exact composition.
It could be expected the origin of this mucous material is
related to numerous Golgi apparatuses present in the
cytoplasm of gregarines [46,58,59]. The longest mucous
trail was left behind gamonts of G. polymorpha. Similarly,
Alcian blue staining at low pH, which proved to be very
helpful to visualise mucous substances (i.e., glycosaminoglycans)
[59,60], showed the highest amount of mucous
material in the cytoplasm of G. polymorpha and lowest in
G. cuneata. The results of this staining, however, conflict
with the seemingly mucus-free surface of G. polymorpha
and the abundant mucus-like drops covering the surface
of G. cuneata, supported by the observations on the secretion
of a mucus-like material in the grooves between
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G. cuneata folds [24]. Accepting the possibility that the
increased production of mucus allows gregarines to glide
with higher speeds, the differences in its viscosity could
be the reason for mentioned observations on mucus
secretion. Nevertheless, the origin of these drops remains
unclear as no reliable conclusions about the chemical
composition could be drawn from electron
microscopic observations. Furthermore, the way of mucus
secretion is not clear. It could be expected that openings or
pore-like structures observed in the grooves between the
folds might be related to mucus. Freeze-etching data further
supported these speculations by the demonstration
of numerous micropores in the pellicle lined by a collar
and often in connections with some cisternae, vesicles
or ducts, similar to those observed in G. garnhami
[22]. Generally, the micropore is defined as an organelle
formed by the apicomplexan pellicle, which is
composed of two concentric rings (in transverse section),
the inner of which corresponds with an invagination
of the outer pellicle membrane. Micropores
are assumed to have a feeding function but their real
function is still poorly understood. A typical micropore
was documented in ultrathin sections of the pellicle of
G. cuneata (Figure 5J), and we assume that this structure
corresponds to the pores revealed by the freezeetching.
Similar structures were observed in the pellicle
of trophozoites of another Gregarina species [22,61] or
in the pellicle of Plasmodium ookinete [62]. In addition,
numerous tiny pore-like structures, located at the base
and lateral side of epicytic folds, were found on the
fractured faces of membranes, especially on the protoplasmic
faces of IMC. The typical organisation of the
proteins forming these structures proved them as pores.
Intramembranous particles
We did not find any correlation between IMP density in
membranes forming the pellicle and the gregarine gliding
rate. Although the density of IMP, as well as the Kp,
in studied gregarines differs, the overall values are
highest in G. steni and lowest in G. polymorpha, both of
which glide at high speed. Only IMP with their size ranging
from 6 to 14 nm were used for statistical evaluation
of their densities in order to get data comparable with
those already published on other apicomplexans (Table 6).
Most conspicuous differences in IMP densities can be
seen between the values reported for IMC of G. garnhami
[11] and our data, especially when considering that both
studies focused on representatives of the same genus. It
must be highlighted, however, that the statistical values differ
considerably when including all visible IMP. A magnification
of 56,000X used for statistics in this study allows
visualisation of tiny IMP of 1 nm in diameter. That is why
we included histograms illustrating the IMP size distribution
in all analysed membranes to show their size variability
among species as well as the frequency of particles with
their sizes beyond this range. Considering the frequency of
particles beyond the size range of 6–14 nm, especially in G.
cuneata, it is questionable if this range set in previous studies
really offers reliable data on IMP densities. An example
is the differences in Kp for membranes when considering
all detectable IMP in contrast to statistical values calculated
for a size range 6–14 nm shown in Table 5; i.e., the Kp for
the G. cuneata plasma membrane is 0.91 (in contrast to
0.81 for a set range of 6–14 nm), for the external cytomembrane
is 0.93 (1.13) and for the internal cytomembrane is
1.49 (1.33); Kp for the G. polymorpha plasma membrane is
1.69 (0.59), for the external cytomembrane is 0.38 (0.70)
and for the internal cytomembrane is 1.41 (1.57); and the
Kp for the G. steini plasma membrane is 1.02 (1.27), for the
external cytomembrane is 0.93 (0.68) and for the internal
cytomembrane is 0.90 (1.24).
Conclusions
Neither the general architectural features of the pellicle, including
the number of epicytic folds or its subcellular components,
nor the supramolecular organisation of the
plasma membrane and IMC (density of IMP and their Kp)
correlate with a gliding rate in eugregarines. Phalloidin and
Table 6 Densities of IMP (particles/μm2
) in different apicomplexans
Species Plasma membrane External cytomembrane Internal cytomembrane
EF PF PF EF EF PF
Gregarina cuneata 2770 ± 96 2244 ± 283 1420 ± 190 1260 ± 211 1502 ± 273 1993 ± 253
Gregarina polymorpha 2473 ± 147 1446 ± 158 602 ± 265 863 ± 202 814 ± 246 1276 ± 200
Gregarina steini 1783 ± 233 2265 ± 154 2588 ± 189 3820 ± 211 1886 ± 274 2339 ± 132
Gregarina blaberae1
977 ± 235 1469 ± 233 285 ± 39 133 ± 34 158 ± 72 297 ± 33
Eimeria nieschulzi2
218 ± 21 648 ± 73 2360 ± 133 29 ± 7 146 ± 31 1780 ± 97
Plasmodium knowlesi3
185 ± 25 2198 ± 528 17511 ± 228 38 ± 15 48 ± 28 574 ± 200
The size of IMP is in range 6-14 nm.
1
Values taken from Schrével et al. [11].
2
Values taken from Dubremetz and Topier [41].
3
Values taken from McLaren et al. [63].
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antibody labelling repeatedly confirmed the presence of
actin and myosin restricted to the cell cortex. Moreover,
the reaction of gregarines to the application of JAS and cytochalasin
D serves as indirect proof of the importance of
actin dynamic polymerisation during gregarine gliding. The
location of the actomyosin complex seems to be restricted
to the lateral parts of the epicytic folds rather than to their
tips, as the number of 12-nm filaments and rippled dense
structures running along their length does not influence
the speed of gliding. The results of Alcian blue staining
along with the mucous trail left behind gliding gamonts are
the proof that the increased load of mucus in the cytoplasm
correlates with gliding rate, as shown in G.
polymorpha vs. G. cuneata, however, the viscosity of the
mucus of the seemingly mucus-free surface of G.
polymorpha needs further investigation. It is also worthy
to highlight that despite the basic concept describing a
substrate-dependent gliding in gregarines [31], for some
period subsequent to drugs application to the cell suspension,
gamonts were free-floating in a liquid lacking any
contact with the substrate but with a significantly higher
rate than exhibited during regular gliding.
Gregarines retained some ancestral features and are considered
to be deep-branching apicomplexans. They evolved
an enormous morphological and ecological diversity, and
exhibit unique and novel adaptations to surrounding environment.
Various gregarines parasitizing terrestrial and
marine invertebrates not only exhibit diverse modes of locomotion
(e.g., gliding in eugregarines with well-developed
epicytic folds vs. bending, rolling or coiling known from
marine archigregarines that probably evolved from hypertrophic
zoite and retained subpellicular microtubules [9,17],
and finally peristalsis-like movements observed in
urosporidians), but even might use several mechanisms of
cell motility depending on their actual physiological and
environmental conditions. An understanding of the
mechanism of gregarine motility and host cell invasion
would offer significant insights into the parasitic strategy
of apicomplexan parasites and evolution of obligate intracellular
parasitism from free-living photosynthetic ancestors.
Materials and methods
Material collection
Larvae of the yellow mealworm, Tenebrio molitor Linnaeus,
1758 (Coleoptera, Tenebrionidae) infected with gregarines
were obtained from colonies maintained in our laboratory.
Gamonts of Gregarina cuneata, G. polymorpha and G. steini
were collected from the intestinal lumen of naturally infected
larvae. As all three eugregarine species can be simultaneously
present in the larvae of T. molitor, experimental infections of
larvae previously sterilised of gregarines were performed
using infective oocysts in order to obtain a model infected
with a single species for electron microscopic analyses.
Protocols concerning experimental infection and following
dissection of infected larvae are described elsewhere [8,45].
Light microscopic observations on gliding motility
Single gamonts and gamonts associated in syzygy were
removed from the host and placed on glass slides in
Minimum Essential Medium [enriched with 3% bovine
foetal serum with penicillin, streptomycin, amphotericin
B and L-glutamine] (Sigma-Aldrich, Czech Republic). Incubation
in this medium increased the viability of gregarines
after isolation from host intestine. Light microscopic
(LM) observations of gliding movement, orientation, and
conformational changes were made. Results were confirmed
by observations on gregarines incubated in Ringer’s
insect physiological solution (pH 7.2) [64]. For speed calculations,
short video records were taken using Bürker
counting chamber. Individual cell speeds (in micrometers
per second) were calculated from individual gregarine
tracks by measurements of distances between initial and
final positions covered over the time interval. The interval
between selected recorded positions was normalised to
1 second using the ImageJ2x software developed at the
National Institutes of Health.
For observations on mucus shedding, living gamonts
were put on microscopic slides covered by a thin
layer of microbiological agar, slightly moistened with
Ringer’s solution and observed using phase contrast
microscopy.
For treatment of gregarines with jasplakinolide (JAS;
Invitrogen, Czech Republic) and cytochalasin D (Invitrogen,
Czech Republic), living gamonts of G. cuneata, G. steini
and G. polymorpha (a mixture of suspensions obtained
from the guts of several hosts) were put on single cavity
microscope glass slides with a drop of JAS or cytochalasin
D in Ringer’s insect physiological solution (pH 7.2). The
JAS was reconstituted in dimethyl sulfoxide (DMSO) to
prepare a 1 mM stock solution and diluted in Ringer’s solution
to prepare final working concentrations (5, 10, 20 and
30 μM JAS). Similarly, the 1 mM stock solution of cytochalasin
D in DMSO was diluted in Ringer’s solution to obtain
working concentrations (10, 20 and 30 μM cytochalasin D).
Control assays of living gregarines were performed in pure
Ringer’s solution as well as corresponding concentrations of
DMSO in Ringer’s solution.
Cell suspensions, squash and/or wet smear preparations
were investigated with the use of a motorised
Olympus microscope BX61 equipped with Olympus
DP71 digital camera and software (Olympus Stream Motion
version 1.5.1).
Mucus staining with alcian blue
Living gamonts were centrifuged at 10 000 × g for
30 minutes and subsequently fixed in freshly prepared
Valigurová et al. Frontiers in Zoology 2013, 10:57 Page 24 of 27
http://www.frontiersinzoology.com/content/10/1/57
4% paraformaldehyde in phosphate buffered saline (PBS).
After washing in 0.2 M PBS, cell suspensions were stained
with Alcian blue (pH 1.3) for 1 hour, rinsed with 0.1 M
hydrochloric acid and washed again in PBS [59]. For light
microscopic analyses, the drops of stained cell suspension
in PBS were dropped onto microscopic slides and covered
by a cover slip.
Electron microscopy
Cell suspensions were fixed overnight at 4°C in freshly
prepared 2.5–3% glutaraldehyde in 0.2 M cacodylate
buffer (pH 7.4). Procedures for sample processing for
transmission (TEM) and scanning electron microscopy
(SEM) follow Valigurová et al. [8,45]. Observations were
made using a JEOL 1010 TEM and JEOL JSM-7401 F Field
emission scanning microscope.
Freeze etching
Cell suspensions were fixed overnight at 4°C in freshly
prepared 3% glutaraldehyde in 0.2 M cacodylate buffer
(pH 7.4) followed by cryoprotection with 20% glycerol
(w) treatment, concentrated on glass slides by using
tweezers and put on the gold carrier. Specimens were
then frozen in melting liquid nitrogen (-210°C), and for
one cycle three carriers were mounted on a gold stand
(subcooled in liquid nitrogen). Using the manipulator,
the gold stand was transported into the working chamber
(Freeze-etching system device, BAF 060 BAL-TEC) cooled
to a temperature of -100°C at a pressure of 10-5
Pa.
Subsequently, the specimens were cut and fractured
with a microtome knife, etched (ice sublimation) for
2 minutes, and replicas were prepared by vacuumdeposition
of platinum (the angle of evaporation was
45°, the thickness of layer 2.4 nm) and carbon (90°
angle of evaporation, 22.2-nm thick layer) onto the
frozen, fractured surface. The replicas were cleaned
with 7% sodium hypochlorite and chromo-sulphuric
acid to remove all the biological material, and washed
in distilled water. The pieces of replica were mounted
on copper grids and examined under a transmission
electron microscope (Morgagni 268 D, FEI). Statistical
evaluation of intramembranous particles (IMP) per a
unit area (1 μm2
) and histograms illustrating the IMP
size distribution were done in ACC (Adaptive Contrast
Control) developed by the Institute of Mathematics, Faculty
of Mechanical Engineering, University of Technology, Brno.
The nomenclature follows that proposed in Branton et al.
[65] and used in Schrével et al. [11].
Confocal laser scanning microscopy
Cell suspensions were washed in 0.2 M PBS, fixed
for 15 minutes at room temperature in freshly prepared
4% paraformaldehyde in 0.2 M PBS, washed,
and permeabilised for 15 minutes in 0.1-0.5% Triton
X-100 (Sigma-Aldrich, Czech Republic). Protocols for
direct staining of filamentous actin with phalloidin–
tetramethylrhodamine B isothiocyanate (phalloidin-TRITC;
Sigma-Aldrich, Czech Republic), as well as indirect immunofluorescent
antibody (IFA) staining using the rabbit
anti-myosin antibody (smooth and skeletal, the whole
antiserum, Sigma-Aldrich, Czech Republic), the mouse
monoclonal IgG anti-actin antibody that was raised
against Dictyostelium actin (provided by Prof. Dominique
Soldati-Favre) and mouse monoclonal anti-α-tubulin antibody
(Clone B-5-1-2, Sigma-Aldrich, Czech Republic)
follow Valigurová et al. [45,46]. Similarly, living gamonts
treated for 10 and 150 minutes with 10 μM JAS were
briefly washed in 0.2 M PBS and fixed for subsequent phalloidin
labelling. Preparations were observed and documented
using an Olympus IX80 microscope equipped with
a laser-scanning FluoView 500 confocal unit (Olympus
FluoView 4.3 software).
Abbreviations
Act: Actin; CLSM: Confocal laser scanning microscopy; DMSO: Dimethyl
sulfoxide; EF: Exoplasmic fracture; F-act: Filamentous actin; FITC: Fluorescein
isothiocyanate; IFA: Indirect immunofluorescent assay; IMC: Inner membrane
complex; IMP: Intramembranous particles; JAS: Jasplakinolide; Kp: Partition
coefficient; LM: Light microscopy; Myo: Myosin; PF: Protoplasmic fracture;
PBS: Phosphate buffered saline; SE: Standard error; SEM: Scanning electron
microscopy; TEM: Transmission electron microscopy;
TRITC: Tetramethylrhodamine B isothiocyanate.
Competing interests
The authors declare that they have no competing interests.
Authors’ contributions
AV conceived and designed the study, carried out the research, performed
the experiments and microscopic analyses, and wrote the manuscript.
JS contributed substantially to the conception and design of the study,
interpretation of experimental data and to the writing of the manuscript.
NV implemented freeze-etching techniques, performed related statistical
analyses and data interpretation. NM assisted in the laboratory work, material
collection and acquisition of data. All authors read and approved the final
manuscript.
Acknowledgements
The authors are greatly indebted to the members of the Laboratory of
Electron Microscopy (Institute of Parasitology, Academy of Sciences of the
Czech Republic, in České Budějovice) for technical assistance. Many thanks to
MSc. Veronika Toporcerová for help with preparation of some of the
scanning electron micrographs. Financial support was provided by the
project GPP506/10/P372 from the Czech Science Foundation. Travel
expenses were partially supported by a Czech-French bilateral project
MEB021127 (Barrande 2011-2012) from the Ministry of Education, Youth and
Sports of the Czech Republic. The first author acknowledges support from
the Department of Botany and Zoology, Faculty of Science, Masaryk
University, towards the preparation of this manuscript.
Author details
1
Department of Botany and Zoology, Faculty of Science, Masaryk University,
Kotlářská 2, 611 37 Brno, Czech Republic. 2
Department of Biophysics, Faculty
of Medicine, Masaryk University, Kamenice 3, 625 00 Brno, Czech Republic.
3
Muséum National d’Histoire Naturelle, UMR 7245 CNRS/MNHN, CP 52, 61
rue Buffon, 75231 Paris Cedex 05, France.
Received: 19 January 2013 Accepted: 24 August 2013
Published: 22 September 2013
Valigurová et al. Frontiers in Zoology 2013, 10:57 Page 25 of 27
http://www.frontiersinzoology.com/content/10/1/57
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doi:10.1186/1742-9994-10-57
Cite this article as: Valigurová et al.: The enigma of eugregarine epicytic
folds: where gliding motility originates? Frontiers in Zoology 2013 10:57.
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7.3
Melicherová J., Ilgová J., Kváč M., Sak B., Koudela B., Valigurová A.
2014
Life cycle of Cryptosporidium muris in two rodents
with different responses to parasitization
Parasitology 141(2), 287-303
Life cycle of Cryptosporidium muris in two rodents with
different responses to parasitization
JANKA MELICHEROVÁ1
†, JANA ILGOVÁ1
†, MARTIN KVÁČ2
, BOHUMIL SAK2
,
BŘETISLAV KOUDELA3
and ANDREA VALIGUROVÁ1
*
1
Department of Botany and Zoology, Faculty of Science, Masaryk University, Kotlářská 2, 611 37 Brno, Czech Republic
2
Institute of Parasitology, Biology Centre of the Czech Academy of Sciences, Branišovská 31, 370 05 České Budějovice,
Czech Republic
3
Department of Pathological Morphology and Parasitology, University of Veterinary and Pharmaceutical Sciences,
Palackého 1/3, 612 42 Brno, Czech Republic
(Received 8 April 2013; revised 25 July and 20 August 2013; accepted 21 August 2013; first published online 11 October 2013)
SUMMARY
This study focuses on mapping the life cycle of Cryptosporidium muris in two laboratory rodents; BALB/c mice and the
southern multimammate rat Mastomys coucha, differing in their prepatent and patent periods. Both rodents were
simultaneously experimentally inoculated with viable oocysts of C. muris (strain TS03). Animals were dissected and
screened for the presence of the parasite using a combined morphological approach and nested PCR (SSU rRNA) at
different times after inoculation. The occurrence of first developmental stages of C. muris in stomach was detected at 2·5 days
post-infection (dpi). The presence of Type II merogony, appearing 36 h later than Type I merogony, was confirmed in both
rodents. Oocysts exhibiting different size and thickness of their wall were observed from 5 dpi onwards in stomachs of both
host models. The early phase of parasitization in BALB/c mice progressed rapidly, with a prepatent period of 7·5–10 days;
whereas in M. coucha, the developmental stages of C. muris were first observed 12 h later in comparison with BALB/c mice
and prepatent period was longer (18–21 days). Similarly, the patent periods of BALB/c mice and M. coucha differed
considerably, i.e. 10–15 days vs chronic infection throughout the life of the host, respectively.
Key words: cryptosporidia, development, gastric, oocyst, pathology, Type II merogony.
INTRODUCTION
The phylum Apicomplexa includes significant and
widespread unicellular pathogens of humans and
animals and one of these is the genus Cryptosporidium
that is the causative agent of zoonotic disease of the
gastrointestinal and respiratory tract, called cryptosporidiosis.
In general, the progress of infection
in immunocompetent individuals is mild, without
fatal consequences, and usually self-curing or selflimiting
(Chappell et al. 1999). The gastric species
Cryptosporidium muris parasitizes epithelial cells in
the glandular part of the gastric mucosa and was
first described in mice by Tyzzer (1907) as a
Cryptosporidium species. Uni et al. (1987) extended
his work and provided new morphometrical data
of its developmental stages. Although C. muris is
considered to be a predominantly rodent species,
occasionally it can be transmitted from its natural
host to other animals (Aydin and Ozkul, 1996;
Hůrková et al. 2003; Pavlásek and Ryan, 2007;
Lupo et al. 2008) and possibly humans (Katsumata
et al. 2000; Guyot et al. 2001; Gatei et al. 2002;
Tiangtip and Jongwutiwes, 2002; Palmer et al. 2003).
The general life cycle, which is well documented
for Cryptosporidium parvum parasitizing the intestine
(Current and Reese, 1986), includes four phases of
development: excystation, merogony, gametogony
and sporogony. Each phase occurs in a particular
chronological period and depends on both the species
of Cryptosporidium as well as the host. Diversity in
patent and prepatent periods has been reported
from various animals (Tarazona et al. 1998; Hijjawi
et al. 2001; Fayer and Santin, 2009). The infective
oocysts are transmitted by the fecal–oral route. The
invasive sporozoites, being released from oocysts
inside the host, evolve to trophozoites and multiply in
the asexual phase involving two types of merogony.
Type II merogony produces four merozoites and has
been recorded in C. parvum and Cryptosporidium
wrairi but not confirmed in C. muris. The sexual part
of the life cycle, gametogony, is typified by the
occurrence of numerous macrogamonts and microgamonts
that produce 16 non-flagellated microgametes.
After macrogamont fertilization, the zygote
is enveloped by a wall and matures into the oocyst,
which sporulates inside the host. A few studies report
the existence of two types of oocysts, i.e. thick- and
thin-walled (Current and Reese, 1986; Uni et al.
1987).
* Corresponding author: Department of Botany and
Zoology, Faculty of Science, Masaryk University,
Kotlářská 2, 611 37 Brno, Czech Republic. E-mail:
andreav@sci.muni.cz
† These authors contributed equally to this work.
287
Parasitology (2014), 141, 287–303. © Cambridge University Press 2013
doi:10.1017/S0031182013001637
Due to lack of efficient treatment of cryptosporidiosis,
further knowledge on cryptosporidia biology
and host–parasite interactions, depending on the host
immunological status, is still required. Our aim was
to map the complete life cycle of C. muris in two
laboratory rodents, BALB/c mice and the southern
multimammate rat Mastomys coucha, and to verify
the presence of Type II merogony and thin-walled
oocysts during the life cycle of C. muris, using a
combined microscopic approach, i.e. electron microscopy
as well as observations on native and stained
preparations using light microscopy (LM), and
supplemented with molecular detection. We also
attempted to record and re-evaluate the stagedependent
characteristics depicted in the detailed
drawings of Tyzzer (1910) and for this reason, we
have followed the techniques used in his study. Since
the endogenous developmental stages of C. muris
attack the stomach tissue, we expected to find some
differences in its life cycle when compared with the
findings for C. parvum.
MATERIALS AND METHODS
Laboratory animal models
The southern multimammate rat (M. coucha; the
Institute of Parasitology Biology Centre, ASCR,
v.v.i. in České Budějovice) and BALB/c mice (Anlab
s.r.o, Czech Republic) were used for the purposes
of this study. Animals were housed in plastic cages
(5 animals per cage) with sterile wood-chip bedding
and supplied with sterilized food and water
ad libitum. The rearing of animals is regulated by
Czech legislation (Act No 246/1992 Coll., on protection
of animals against cruelty). These documents
are consistent with the legislation of the European
Commission. All housing, feeding and experimental
procedures were conducted under protocols approved
by the Institute of Parasitology, Biology
Centre of the Academy of Sciences of the Czech
Republic and Central Commission for Animal
Welfare, Czech Republic (Protocol # 066/2010).
The minimum number of animals has been involved
to produce statistically reproducible results.
Parasite
Cryptosporidium muris strain TS03 originating from
an East African mole rat (Tachyoryctes splendens)
previously characterized by Kváč et al. (2008, 2011)
was used for all experiments in this study. Over
10 years the C. muris TS03 has been passaged through
susceptible laboratory hosts, M. coucha and SCID
mice, in the Institute of Parasitology Biology Centre,
ASCR, v.v.i. in České Budějovice. Fresh rat feces
were collected regularly every morning. The presence
of oocysts in feces was verified using the staining
method according to Miláček and Vítovec (1985).
Oocysts were purified using Sheather’s sugar flotation
method (Arrowood and Sterling, 1987) and modified
caesium chloride gradient centrifugation (Kilani and
Sekla, 1987). Obtained oocysts of C. muris were either
immediately used for experimental inoculations of
laboratory animals or mixed with aqueous potassium
dichromate (2·5% w/v, final concentration) and stored
at 4 °C. Oocysts less than 1 week old were exclusively
used for the experiment in this study. Before their
use, the residue of aqueous potassium dichromate
was removed by centrifugation at 1000 g for 10 min,
the supernatant was aspirated, and the pelleted
oocysts were resuspended in distilled water. This
procedure was repeated three times.
Experimental inoculations and parasitological
dissection of laboratory rodents
Rodents, 8–10 weeks old, were inoculated per os
using the oesophagus gavage with a dose of 106
viable
oocysts of C. muris that had previously been checked
for their viability via propidium iodide staining
(Sauch et al. 1991). Inoculations of both rodent
models were performed simultaneously. Animals
(one per host model), were euthanased by cervical
dislocation at 0·5, 1, 6 and 12 h post infection (hpi)
and then consecutively every 12 h until 11 dpi, and
onwards at 12, 13, 14, 18, 21 and 28 dpi. In
M. coucha, additional parasitological examination
wasperformed, ofanimals keptfor4,8and18months,
and 2 years after inoculation. The negative controls
of both models, BALB/c mice and M. coucha, were
prepared to evaluate pathological alterations in gastric
tissue induced by cryptosporidiosis. A total of 33
individuals of BALB/c mice and 37 individuals of
M. coucha were examined. In subsequent repetition of
the experiment, performed to detect cryptosporidia
during the early phase of parasitization, the infective
doses were increased to 108
oocysts of C. muris per
animal. Animals were dissected at selected time
points after inoculation (a total of 8 individuals of
BALB/c mice and 17 individuals of M. coucha).
Excision of the stomach, consisting of both the
glandular and non-glandular part was performed. As
endogenous developmental stages of C. muris are
known to be restricted to the glandular part, this area
of the gastric tissue was divided lengthways through
the curvatura major and then transversely. The resulting
four sections were processed using the various
approaches described in the following subchapters.
Differences between the course of parasitization in
groups of BALB/c mice and M. coucha were tested by
the non-parametric Mann–Whitney U test. Testing
was performed using the software Statistica®
10.0
(StatSoft CR, Prague, Czech Republic).
Native and stained preparations
As the first step, the blood serum of both BALB/c
and M. coucha was obtained from non-infected
288Janka Melicherová and others
controls. A total of 10 μL (BALB/c) or 30 μL
(M. coucha) of adequate sera were subsequently
added to the processed gastric tissue in order to
release the C. muris developmental stages. Native
preparations, prepared from suspensions obtained
from scrapings of the gastric luminal epithelium,
were evaluated immediately via LM and checked
for the presence of cryptosporidia. Preparations
with gastric scrapings were also stained with Lugol’s
iodine. The protocols according to Giemsa and
Wright staining procedures (Tyzzer, 1910) were
used to visualize the characteristic morphological
features of individual C. muris developmental stages.
For this purpose, specimens were fixed with a vapour
of 2% osmium tetroxide (v/v). Slightly modified
staining procedures followed Tyzzer (1910). Preparations
were viewed and documented using the light
motorized microscope Olympus BX61 equipped
with phase contrast optics and Olympus BX51 with
Nomarski interference-contrast microscopy (DIC).
Electron microscopy
Samples of gastric tissue were fixed in freshly
prepared 3% glutaraldehyde (v/v) in cacodylate
buffer and further processed for transmission (transmission
electron microscopy (TEM)) and scanning
electron microscopy (SEM) as described elsewhere
(Valigurová et al. 2007, 2008). The grids for TEM
were coated with Formvar film at a thickness of
15–25 nm. Ultrathin sections transferred to the
Formvar coated copper grids were evaluated using
a JEOL 1010 transmission electron microscope.
Micrographs from SEM were performed by JEOL
JSM-7401F. In the case of negative results obtained
during the early phase of infection, samples of gastric
tissue already viewed under SEM were broken
into smaller pieces to expose the interior of the closed
gastric pits, were re-coated with gold and re-
evaluated.
Histology
Gastric tissue was fixed in AFA (alcohol – formalin –
acetic acid) solution and processed as described
previously (Valigurová et al. 2008). The blocks in
Histoplast II were cut using a Zeiss Hyrax M 300
rotary microtome. The 7 μm thick sections were
stained with haematoxylin–eosin (H&E). Preparations
were viewed and documented using the
light motorized microscope Olympus BX61.
Molecular method; nested PCR
This method was chosen to verify the presence (or
absence) of specific DNA of developmental stages
of C. muris in the stomachs of both animal models
during the early phase of parasitization (up to 3 dpi).
In addition, this approach was essential to disprove
the localization of C. muris developmental stages in
other organs such as the gall bladder, pancreas,
oesophagus, spleen, liver, Peyer’s patches, duodenum,
jejunum, ileum, colon, lung and kidney.
Total DNA was extracted from tissue by bead
disruption for 60 s at 5·5 m s−1
using 0·5 mm glass
beads in a FastPrep®
–24 Instrument (MP Biomedicals,
CA, USA), followed by isolation/purification
using a commercially available kit according to the
manufacturer’s instructions (DNeasy Blood and
Tissue Kit, Qiagen, Hilden, Germany). Purified
DNA was stored at −20 °C prior to being used for
PCR. The presence of C. muris was confirmed by
sequence analysis of the SSU gene using previously
described methods (Jiang et al. 2005). The positive
(sample containing DNA of C. hominis) and negative
control were included in each PCR. Secondary PCR
products were detected by agarose gel electrophoresis
(2·0%) and visualized by ethidium bromide staining
(0·2 μg mL−1
).
RESULTS
The effect of parasitization by C. muris on
laboratory rodents
Laboratory rodents exhibited significantly different
responses to parasitization and chronology of pathological
changes of gastric tissue induced by the
parasite, in contrast to the negative control of gastric
tissue (Figs 1A–C and 2A, B); nevertheless, the
sequence of individual changes during the acute
phase of parasitization corresponded in both models.
At the beginning of parasitization, the gastric tissue
of both host models was irregularly affected by
endogenous developmental stages of C. muris in an
island-like manner and individual parasitized gastric
pits were surrounded by areas of healthy tissue
(Figs 1D, F and 2D, F). This specific feature of
C. muris hampered the localization of parasites in
host stomachs and made the detection of its earliest
developmental stages almost impossible in both
rodent models. Therefore, a variety of different
microscopic approaches along with nested PCR
were necessary to confirm the presence or absence
of C. muris developmental stages during the early
phase of infection.
The BALB/c mice showed a significantly shorter
prepatent period (P<0·05), in contrast to that of
M. coucha, and the first oocysts in their feces
were detected at 7·5 dpi with range of 7·5–10 dpi.
The patent period in BALB/c mice lasted 10–15 dpi.
The first obvious alterations of its gastric surface
were noticeable after 3 dpi (Figs 1D–F). Some gastric
pits were slightly open and thus enlarged (Fig. 1E).
At 8 dpi, the majority of the gastric glandular part
was obviously affected (Fig. 1G–I), whereas the
289Life cycle of Cryptosporidium muris in two rodent models
Fig. 1. Pathology of gastric cryptosporidiosis in BALB/c mice. (A) Cross-section of gastric glands; negative control.
LM (Histology, H&E). (B) The surface of the gastric epithelium showing empty gastric pits (arrow). Inset shows a
detailed view inside an empty gastric pit; negative control. SEM. (C) Cross-section of a gastric gland; negative control.
TEM. (D) Longitudinal section of gastric epithelium; gastric pits (arrow) parasitized by C. muris (arrowhead) at 5·5 dpi.
LM (Histology, H&E). (E) Cross-section of a gastric gland with C. muris individuals at 3 dpi. TEM. (F) Surface of the
gastric epithelium showing gastric pits filled with C. muris (arrowhead) at 7 dpi; an apparently empty gastric pit (arrow).
Inset shows a detailed view inside the gastric pit filled by C. muris developmental stages. SEM. (G) Cross-section of
enlarged gastric glands after 8·5 dpi (arrow). LM (Histology, H&E). (H) Gastric pits completely filled by C. muris (arrow)
at 8·5 dpi; the surface is covered by a mucus layer (asterisk). SEM. (I) Cross-section of a gastric gland completely filled
with various developmental stages of C. muris at 8 dpi. TEM. (J) Longitudinal section of a stomach showing parasitized
gastric pits (arrow) and glands (arrowhead) at 13 dpi. LM (Histology, H&E). (K) The folds of the enlarged gastric
epithelium (arrowhead); C. muris developing at the surface of epithelium (arrows) at 13 dpi. SEM. (L) Hypertrophy and
hyperplasia of the gastric tissue; numerous developmental stages of C. muris (arrow) at 13 dpi. SEM.
290Janka Melicherová and others
Fig. 2. Pathology of gastric cryptosporidiosis in Mastomys coucha. (A) Cross-section of gastric glands; negative control.
LM (Histology, toluidine blue). (B) Cross-section of the gastric gland; negative control. TEM. (C) Surface of the gastric
epithelium at 1 dpi; gastric pits (arrowhead). SEM. (D) Cross-section of the parasitized gastric gland (arrow) surrounded
by non-parasitized glands (arrowhead) at 5 dpi. LM (Histology, toluidine blue). (E) Cross-section of the gastric gland
filled with C. muris (arrow) at 5 dpi. TEM. (F) Surface of the gastric epithelium exhibiting seemingly empty and closed
gastric pits (arrowhead) and parasitized gastric pits (arrow) at 5 dpi. SEM. (G) A detailed view inside the gastric pit with
stages of C. muris (arrow) at 5·5 dpi. SEM. (H) Cross-section showing an acute phase of parasitization; obvious
dilatation of the gastric glands filled with various stages of C. muris (arrow) at 10·5 dpi. LM (Histology, toluidine blue).
(I) Surface of the gastric epithelium exhibiting enlarged gastric folds (arrow) covered by mucus (asterisk) at 21 dpi.
SEM; (J) Oblique section showing hyperplasia of the gastric gland filled with C. muris at 8 months PI; a massive
291Life cycle of Cryptosporidium muris in two rodent models
non-glandular part exhibited no changes. The most
obvious changes were observed at the peak of
parasitization around 13 dpi. Pathological modifications
of the host tissue induced by the parasite
included an intensive epithelial hyperplasia (Fig. 1K)
and a mucosal hypertrophy without inflammatory
exudates. The gastric surface of the glandular part
was markedly deformed due to intense pathological
changes, including extension of gastric longitudinal
folds, and numerous cryptosporidia were seen to be
attached to superficial gastric epithelium outside
pits (Fig. 1J–L). The most significant feature that
distinguished the progress of parasitization in both
host models was the host self-recovery of all BALB/c
mice from cryptosporidiosis that occurred after
21 dpi. Similarly, the pathological modifications
of gastric tissue in BALB/c mice gradually retreated
from 21 dpi onwards. At 28 dpi, the necrotic
cells were almost completely replaced by new ones
and the gastric tissues appeared to be completely
regenerated.
In M. coucha, the prepatent period was significantly
longer (P<0·05) and the first oocysts in the
feces were detected at 18 dpi. In the early phase of
parasitization, the gastric tissue showed no observable
changes and no parasites were detected in closed
pits evaluated under SEM (Fig. 2C). In the course of
parasitization, individual gastric pits were progressively
invaded by parasites and thus became slightly
dilated at 5 dpi (Fig. 2D–G). Subsequently, the
dilatation of gastric glands and pits became increasingly
noticeable. At the peak of parasitization at
10–14 dpi, almost all pits and glands were filled by
C. muris (Fig. 2H). The further progress of parasitization
caused a distinctive diffuse mucosal hypertrophy
characterized by the formation of enlarged
or giant gastric folds (Fig. 2I, L and N), without
inflammatory exudates, and subsequently an intensive
epithelial hyperplasia occurred (Fig. 2M).
The massive increase in the volume of the lamina
propria, located beneath the epithelium, caused an
enlarged distance between individual affected gastric
glands (Fig. 2J–K) and longitudinal folds were
twisted and deformed. When the infection entered
a chronic phase, all the above-described pathological
alterations of parasitized gastric tissue became
much more obvious (Fig. 2N–P) and the feces of
M. coucha remained positive on the presence of
C. muris oocysts.
The life cycle of C. muris
The life cycle of C. muris was studied and documented
in detail at set time intervals after inoculation
of both host models; BALB/c mice and M. coucha.
Oocysts released active sporozoites (Figs 3A and
7O, P), which rapidly penetrated deep into the
bottom of the pits of the gastric glands to avoid the
adverse conditions in the host stomach. The first
documented attached developmental stages of
C. muris in both hosts (for chronology and dimensions
see Table 1) were oval to ovoid young
trophozoites observed at 2·5 dpi in BALB/c mice
and at 3 dpi in M. coucha (Figs 3B and 4A–F). In
preparations stained with Giemsa and Wright, the
apical part of the attachment site of the parasitophorous
sac (PS) enveloping the trophozoite stained
intensively pink, thereby confirming its host cell
origin, and the cytoplasmic region packed with
lamellae of the feeder organelle appeared transparent
(Fig. 3B–D). The alteration of the gastric microvillous
surface and the gradual formation of a PS
from the host cell plasma membrane were observed
(Fig. 4A–C). The formation of lamellae of the feeder
organelle from the membrane of an anterior vacuole
was observed exclusively in the earliest trophozoites
(Fig. 4B and C). The apical part of the mature
trophozoites enveloped by the PS, which were
mechanically detached from the host tissue, exhibited
strong adhesive properties, causing the clustering of
trophozoites and subsequent formation of rosettes
(Fig. 3C). A slight elongation of host cell microvilli
surrounding some of the attached parasites was
observed. Further observations on early development
of C. muris were described in detail previously
(Valigurová et al. 2007, 2008) and hence not reported
again.
The progress of the asexual phase into Type I
merogony was characterized by clusters of chromatin
localized in the peripheral cytoplasm of the parasite
(Fig. 3E). Daughter nuclei formed inside a Type I
meront undergoing a nuclear division (Fig. 5B–D).
The asynchrony of nuclear division often resulted
in an odd number of chromatin clusters. By the
process of multiple budding from the residuum still
attached to the host cell (ectomerogony), motile and
invasive merozoites differentiated (Figs 3F–G and
5A, E–O). Almost the entire cytoplasm of the mother
meront appeared to be used up for merozoite
increase in the volume of the lamina propria (lp). TEM. (K) Cross-section showing hyperplasia of the parasitized gastric
glands filled with numerous C. muris individuals (arrow) at 8 months PI. LM (Histology, toluidine blue). (L)
Hypertrophy of the gastric glandular part at 8 months PI; enlarged gastric pits (arrowhead), parasite expanding from the
pits to the luminal gastric epithelium (arrow). SEM. (M) View inside the dilated gastric pit filled by C. muris (arrow) at
8 months PI; hyperplasia of gastric cells (arrowhead). SEM. (N–P) Heavily modified gastric epithelium covered by
numerous C. muris parasites (arrow) during the course of chronic infection after 2 years PI. SEM.
292Janka Melicherová and others
Fig. 3. The life cycle of Cryptosporidium muris as observed in smear preparations stained with Wright. (A) Free, infective
sporozoites. (B–D) Trophozoite stage; gradually maturing trophozoites (B), clustering of trophozoites (C), trophozoite
attached to a host cell (D). (E–G) Type I merogony; clusters of chromatin in the cytoplasm at the periphery of parasite
(E), budding of merozoites from the meront cytoplasm (F, G). (H–I) Free, motile Type I merozoites. (J–K) Type II
merogony; first mitotic division (J), four daughter nuclei formed following the second mitotic division (K). (L) Type II
merozoites inside the parasitophorous sac. (M–N) Macrogamont stage; maturing macrogamonts (M) transforming into
mature ones (N) typified by an increased number of amylopectin granules, giving their cytoplasm (purple) a foam-like
appearance; (O) Mature macrogamont and microgamont exhibiting multiple divisions of nuclei. (P) Microgamont stage;
small compact nuclei localized at the cytoplasm periphery or at the distal end of the microgamont. (Q–R) Thick-walled
oocysts; zygote (left) transforming into an oocyst (right) still enveloped by the parasitophorous sac (Q), more advanced
stages of oocyst sporulation following release from the sac (R). (S–T) Thin-walled oocysts enveloped by parasitophorous
sacs, one of them releasing four sporozoites (T).
293Life cycle of Cryptosporidium muris in two rodent models
formation. Type I merogony usually resulted in the
formation of 6–8 merozoites, most likely depending
on the number of nuclear divisions (Figs 3H, I and
5K, L). Occasionally, five merozoites still attached to
the residual body were observed (Fig. 5O). Type I
merozoites (9·64×1·52 μm), which were released
from the PS, either recycled to form another
generation of Type I meronts, or transformed into
the Type II meront stage and produced four
merozoites. Observation on Type II merogony is
well documented by scanning electron micrographs
showing the ruptured PS releasing four merozoites
and further confirmed by native/stained smear
preparations (Figs 3J–L and 5P–S). The Type I
meronts (9·51×7·25 μm) were observed in both hosts
at the same time as the trophozoites; however,
Type II meronts (9·36×7·39 μm) were first observed
at 4 dpi in BALB/c mice and at 4·5 dpi in M. coucha.
Merozoites exhibited a typical subcellular organization
(Fig. 5M and T), including presence of
subpellicular microtubules, however no mitochondria
were observed. In both host models, the Type I
merozoites were usually present simultaneously with
Type II merozoites, the latter being slightly shorter
and broader. Merozoites seemed to possess several
rhoptries (Fig. 5J and T).
Released merozoites (it is generally assumed that
Type II merozoites) initiated sexual multiplication,
called gametogony, via differentiating into either
female (Fig. 3M–O) or male microgamonts (Fig. 3O
and P). Macrogamonts (8·04×6·49 μm) were detected
in both hosts at 5 dpi and microgamonts
(6·18×5·54 μm) were successfully documented
slightly later, i.e. at 5·5 dpi in BALB/c mice and at
6 dpi in M. coucha. The mature macrogamonts were
easily recognized, as their cytoplasm was packed with
numerous oval-shaped amylopectin granules, the
quantity of which was directly proportional to the
maturation stage (Figs 3M–O and 6A–F, I). Their
cytoplasm possessed typical wall-forming bodies
localized just beneath the pellicle (Fig. 6D, E).
Non-mature microgamonts in stained smears
resembled the meront stage but contained smaller
and more compact nuclei (Fig. 3O). The nucleus of
the microgamonts divided four times and gave rise
to the 16 clusters of chromatin usually localized at the
distal end or periphery of microgamonts (Fig. 3P).
After nuclear division, daughter nuclei migrated
out of the microgamont cytoplasm into the space of
the PS and subsequently developing microgametes
budding from the surface of the microgamont were
observed (Figs 3P and 6G–K). The beginning of
microgamete budding was accompanied by an
obvious evagination of the microgamont pellicle
(Fig. 6J and K). The formation of microgametes
appeared to be asynchronous (Fig. 6J and K) and
resulted in the formation of up to 16 microgametes.
After separation of the mature microgametes, a
large residual body remained attached to the host
cell via feeder organelles (Fig. 6K). The mature
non-flagellated, bullet-shaped microgametes exhibited
a slightly flattened, expanded anterior region
and possessed an elongated, condensed nucleus
(Fig. 6L–P). Microgametes were covered by a
relatively thick plasma membrane and an additional
single membrane was observed beneath this in the
anterior half of the body. This single membrane
formed an invagination beneath the apical cap and
appeared to join the anterior conical structure
(Fig. 6O). Originating at the anterior conical structure
localized just beneath the apical cap, numerous
(at least 10) parallel-arranged microtubules extended
posteriorly surrounding the nucleus (Fig. 6M–P).
The microtubule forming a loop was observed
(Fig. 6P) behind the posterior end of nucleus and it
might play a role in pushing out the nucleus from the
microgamete towards the macrogamont nuclear
envelope during the fertilization process.
After macrogamonts became fertilized by microgametes
(Fig. 6Q–R) their wall appeared thicker.
Through a transient stage of the diploid zygote
(Figs 3Q and 7A, C), they rapidly developed into
oocysts (Figs 3R and 7A–M). The oocyst stage
occurred within the same time period as macrogamonts,
i.e. after 5 dpi in both hosts. In the course of
sporulation, oocysts formed four sporozoites that
were coiled around a large residual body (Fig. 7F–I).
Based on TEM observations, oocysts appeared to
sporulate in situ whilst being enveloped by the intact
PS, as they exhibited already well-differentiated
sporozoites (Fig. 7F, H and I). Both types of oocysts
were observed: the typical thick-walled ones and a
second type resembling the so-called ‘thin-walled’
oocysts (Figs 3R–T, 7H and I). Nevertheless,
variability in oocyst wall thickness was very high.
Table 1. The occurrence of specific developmental
stages of C. muris
Developmental
stages of
C. muris Size (μm)
First
detection
in the
stomach of
BALB/c
mice
First
detection
in the
stomach of
M. coucha
Trophozoites 7·85×4·89 2·5 dpi 3·0 dpi
Type I Meront 9·51×7·25 2·5 dpi 3·0 dpi
Type II Meront 9·36×7·39 4·0 dpi 4·5 dpi
Free merozoites 9·64×1·52 4·0 dpi 4·5 dpi
Macrogamont 8·04×6·49 5·0 dpi 5·0 dpi
Microgamont 6·18×5·54 5·5 dpi 6·0 dpi
Thick-walled
oocysts
7·48×5·86 5·0 dpi 5·0 dpi
Thin-walled
oocysts
8·03×6·69 5·0 dpi 14·0 dpi
Sporozoites in
stomach
11·72×0·98 6·0 dpi 14·0 dpi
Oocysts in feces 7·48×5·86 7·5 dpi 18·0 dpi
dpi = days post-infection; hpi = hours post-infection.
294Janka Melicherová and others
The suture present on the surface of a well-developed
oocyst marks the place where sporozoites were
released (Fig. 7D, J–L and N). As the ‘thin-walled’
oocysts found in BALB/c mice at 5 dpi appeared to
open inside the host stomach, they are presumed to be
responsible for the presence of free sporozoites
(11·72×0·98 μm) observed in scrapings of gastric
mucosa at 6 dpi (Fig. 7P) onwards and they appeared
to result in autoinfection (Fig. 7O). In M. coucha, no
‘thin-walled’ oocysts were detected in the initial
phase of parasitization, but they were observed from
14 dpi onwards, and the free sporozoites occurred
simultaneously with them. In BALB/c mice, thickwalled
oocysts (7·48×5·86 μm) that were shed into
the environment in host feces were detected at
7·5 dpi, whereas this did not occur in feces of
M. coucha until 18 dpi.
To detect early infection by C. muris, the experiment
was repeated using a higher infective dose of
108
oocysts per host. Subsequently, additional
microscopic and molecular analyses were performed.
Based on PCR results, C. muris DNA was detected in
the stomach of BALB/c mice at 1, 6, 12, 36, 48 and
72 hpi. Using the combined approach of SEM and
PCR, other organs from BALB/c mice including
pancreas, oesophagus, spleen, liver, Payer’s patches,
kidneys, lungs, duodenum, jejunum, ileum, gall
bladder and feces were examined for the presence
of C. muris developmental stages and specific
Cryptosporidium DNA but none was found. The
parasitization of the stomach in M. coucha was
confirmed at 6, 48 and 72 hpi. The same organs as
in BALB/c mice (except for the gall bladder) were
analysed and showed positive results in the duodenum
at 48 hpi and in the jejunum at 72 hpi. Using
molecular tools, C. muris developmental stages were
also found in the feces of M. coucha at 36 and 72 hpi.
DISCUSSION
Understanding the life cycle of parasitic organisms is
fundamental to explain their pathogenicity. Parasitic
strategies, such as the ability to enter into the host and
avoid the host defence mechanisms, must be closely
analysed. The essential factors influencing infectivity
are morphological structure and the physiological
processes occurring inside the parasite and host. The
sensitivity of hosts to C. muris depends on its specific
isolate or strain (Xiao et al. 1999; Kváč et al. 2008;
Kodádková et al. 2010). Our observations on the life
cycle of C. muris (strain TS03) in different rodent
models showed a dissimilar chronology with respect
to the occurrence of individual developmental stages.
As the physiology and immune response to parasitization
of both host models varies in many aspects,
the host anatomy and tissue morphology together
with enzymatic secretion appear to be important in
the understanding and evaluation of differences in
parasitization of both hosts with C. muris.
Localization
The gastric pathogen C. muris clearly prefers the acid
environment of the stomach and the common use of
hydrochloric acid to accelerate the in vitro excystation
of its oocysts also serves as proof of this. The pH
values in the gastrointestinal tract of a rat or mouse
are about 5·5–6 in the mouth and intestinal parts,
whereas those in the stomach rapidly decrease to 2–4
(Ward and Coates, 1987; McConnell et al. 2008). The
stomach is comprised of a non-glandular and a
glandular part. As the mucosa of the non-glandular
stomach is lined by keratinized, stratified squamous
epithelium without secretory activity, its surface is
unsuitable for parasite attachment and development.
Fig. 4. Trophozoite stage. (A) Longitudinal section of an attached zoite gradually becoming enveloped by the
parasitophorous sac (ps); endoplasmic reticulum (er), microvilli (mv), parasite nucleus (n). TEM. (B) Section of an early
trophozoite enveloped by the parasitophorous sac (ps); anterior vacuole (av), endoplasmic reticulum (er), microvilli
(mv), parasite nucleus (n). TEM. (C) Detailed view of an anterior part of a young trophozoite. Note the folded
membrane of the anterior vacuole and its gradual transformation into the lamellae of the feeder organelle (arrows);
parasitophorous sac (ps). TEM. (D) Young trophozoite. LM (native preparation). (E) Trophozoite enveloped by the
parasitophorous sac. LM (native preparation); (F) View of a maturing trophozoite completely enveloped by the
parasitophorous sac; elongated host cell microvilli (mv). SEM.
295Life cycle of Cryptosporidium muris in two rodent models
Fig. 5. Merogony. (A) Meront with six merozoites. LM (native preparation). (B–D) Asynchronous mitotic division of
nuclei in a meront; cytoplasm with abundant endoplasmic reticulum (er), filamentous cytoplasm (asterisk), lamellae of
feeder organelle (fo), nucleoli of future merozoites (n), parasitophorous sac (ps), pellicle of meront (arrow). TEM.
Fig. D shows a meront with five daughter nuclei in cross section. TEM. (E) Meront with prominent endoplasmic
reticulum (er), feeder organelle (fo) and lipid inclusion (asterisk). Note the formation of merozoites from the cytoplasm
of the mother meront (arrowhead) still enveloped by an intact parasitophorous sac (ps). TEM. (F) Merozoite still
attached to residual body with lipid inclusion (arrow); released merozoite (arrowhead). LM (native preparation).
(G) Type I meront with budding merozoites (m). SEM. (H) Ruptured parasitophorous sac (ps) revealing more
advanced stages of budding merozoites (m). SEM. (I) Daughter merozoites budding from a Type I meront still
enveloped by a parasitophorous sac (ps); apical ends of merozoites (arrows), endoplasmic reticulum (er), nuclei (n).
TEM. (J) Detailed view of budding merozoite from Fig. 5I; residual body of meront, nucleus (n), rhoptries (arrows).
TEM. (K) Two parasitophorous sacs filled with Type I merozoites (arrow) inside and one free merozoite. LM (native
preparation). (L) Merozoites being released from the parasitophorous sac. LM (native preparation). (M–N) Formation
of merozoites (m) from the cytoplasm of a meront; ducts of merozoite rhoptries (arrows), large residual body (rb) with
endoplasmic reticulum (er) and lipid inclusion (asterisk), parasitophorous sac (ps). TEM. (O) Odd numbers (5) of
mature Type I merozoites attached to the residual body. SEM. (P–Q) Type II merogony producing four shorter
merozoites. LM (native preparation). (R) Cross-section of a Type II meront; four merozoites with nucleus (n) and
rhoptries (arrows). TEM. (S) Ruptured parasitophorous sac (ps) releasing four Type II merozoites (m). SEM.
(T) Mature merozoites in a parasitophorous sac in longitudinal section; dense granules (×), micronemes (arrowhead),
rhoptries (arrows).
296Janka Melicherová and others
Fig. 6. Sexual multiplication. (A–B) Macrogamonts packed with numerous amylopectin granules giving them a
foam-like appearance, feeder organelle (white arrowhead). LM (native preparation). (C) Maturing macrogamont with a
few amylopectin granules (ag), prominent endoplasmic reticulum (er) situated in the centre, feeder organelle (fo),
filamentous projection (asterisk), lipid inclusion (x), nucleus (n), pellicle of macrogamont (arrowhead). TEM.
(D) Macrogamont with wall-forming bodies (arrow) localized just beneath its pellicle (arrowhead); the lamellae of the
feeder organelle (fo), cytoplasm with prominent amylopectin granules (ag), endoplasmic reticulum (er), parasitophorous
sac (ps). TEM. (E) Macrogamont with numerous wall-forming bodies (arrow), feeder organelle (white arrowhead).
LM (native preparation). (F) Numerous macrogamonts at different stages of development. LM (native preparation).
(G,H) Microgamonts with budding microgametes localized at their distal ends or periphery (arrowhead), feeder
organelle (white arrowhead). LM (native preparation). (I) Various stages of mature macrogamonts (ma) and one
microgamont (mi) with microgametes (arrowhead); host cell (hc). TEM. (J) Microgamont showing an asynchronous
budding (arrows) of microgametes (mi); feeder organelle (fo), parasitophorous sac (ps), residual body (rb).TEM.
(K) Microgamont exhibiting a formation of microgametes (mi) inside a parasitophorous sac (ps); dense band
(arrowhead), feeder organelle (fo), residual body (rb). TEM. (L) Two microgamonts (mi) at different developmental
stages stained with Lugol’s iodine; free bullet-shaped microgametes (right). LM. (M) Higher magnification of three
microgametes in longitudinal section. Note the apical cap (asterisks) and the elongated nucleus (n) surrounded by
microtubules (arrow). TEM. (N) Higher magnification of a microgamete in cross-section; microtubules (arrow)
surrounding the nucleus (n). TEM. (O) Microgamete in tangential section; apical cap (asterisk) showing the
invagination of the inner single membrane (arrowhead) attached to an anterior conical structure; microtubules (arrow),
nucleus (n). TEM. (P) Further microgamete in longitudinal section; concentric lamellae located posteriorly to the apical
cap (white arrow), microtubules (arrows), microtubule forming a loop (arrowhead) around posterior end of nucleus (n).
TEM. (Q) The microgamete (arrowhead) penetrating the parasitophorous sac most likely enveloping a macrogamont
(ma). SEM. (R) Free microgametes (mi) close to the macrogamont (ma), apical caps of microgametes (asterisks). TEM.
297Life cycle of Cryptosporidium muris in two rodent models
In contrast, the glandular part forms gastric pits lined
by a simple columnar epithelium. Simple tubular
gastric glands, found on the bottom of the gastric pit,
are dedicated to the production of acids to maintain
the low pH level of the stomach juice and proteases, as
well as mucous substances. The wall of the glandular
part of the stomach is thicker and creates longitudinal
folds (Ward and Coates, 1987; Ghoshal and Bal,
1989). Considering all these morphological features,
despite its extremely acid environment providing less
hospitable conditions for parasites than the intestine,
the gastric glandular part appears to be more suitable
for Cryptosporidium development when compared
with the non-glandular part. Hence, the question
arises how the invasive stages of these minute
parasites avoid the effects of gastric juices containing
strong acids. Matsubayashi et al. (2011) showed that
the viability of sporozoites from oocysts excysted in
medium at pH 7·0 is approximately 90% in the gastric
species, Cryptosporidium andersoni, in contrast to 56%
viable sporozoites for the intestinal species
C. parvum. At a lower pH, however, the viability of
Fig. 7. Oocyst sporulation and excystation. (A) Various developmental stages from zygotes to oocysts. LM (native
preparation). (B) Non-sporulated oocysts possessing a large residual body (arrow). LM (native preparation). (C) Zygote
with numerous amylopectin granules (ag) after fertilization; developing oocyst wall (arrow), parasitophorous sac (ps).
TEM. (D) Mature oocyst inside parasitophorous sac. SEM. (E) Oocyst in retreating parasitophorous sac (ps). SEM.
(F) Longitudinal section of an oocyst in parasitophorous sac (ps); feeder organelle (fo), large residual body (rb), formed
sporozoites (s), oocyst wall (arrow). TEM. (G) Excystation of sporozoites from the oocyst. LM (native preparation).
(H) Thin-walled oocyst with sporozoites (s) inside a complete parasitophorous sac (ps); residual body (asterisk) with
amylopectin granules (ag), wall of oocyst (arrowhead). TEM. (I) Thick-walled oocyst inside an intact parasitophorous
sac (ps), amylopectin granules (ag), sporozoites (s). TEM. (J) Oocyst equipped with a typical suture (arrow). SEM.
(K) The excystation of sporozoites from the oocyst. LM (native preparation). (L) Empty oocyst and free sporozoites.
LM (native preparation). (M) Oocyst with an even surface considered to be thin-walled. SEM. (N) The ruptured empty
oocyst. SEM. (O) Free sporozoite invading the gastric epithelium. SEM. (P) Invading sporozoite. LM (native
preparation).
298Janka Melicherová and others
gastric sporozoites gradually declined to 61% after 3 h
incubation at pH 2. Consequently, in comparison
with intestinal cryptosporidia, the better adaptation
of gastric species to an acidic environment enables
them to survive for longer in the stomach
(Matsubayashi et al. 2010).
In general, cryptosporidia have evolved efficient
strategies to avoid the potential harmful effects of the
gastrointestinal tract, e.g. direct exposure to digestive
enzymes, possible mechanical damage or their peeling
off from the host tissue. Probably one of the most
important of these strategies is enveloping by the PS
of host cell origin (Valigurová et al. 2007, 2008).
Another is invading a proper shelter (e.g. gland or
pits) and becoming covered by a mucus layer,
especially in gastric species. To prevent too-rapid
erosion of the gastric mucosa due to a hostile stomach
environment, the surface cells and cells of the gland
necks produce mucous secretions that form a
relatively thick protective layer, which often hampers
the ability to view inside the gastric gland using
SEM. Even very careful washing of gastric tissue is
often not sufficient to clear the entire surface from
mucus, which forms a compact and non-transparent
layer after chemical fixation. As even this protection
is not perfect, the shedding of dead cells from the
gastric pits into the lumen is often observed and
might also be one of the reasons for the occasional
detachment of PS filled with cryptosporidia and their
subsequent presence in the host intestine or feces.
Another curiosity is the progress and mode of
parasitization by C. muris as well as its dispersal
throughout the gastric tissue in an irregular, islandlike
pattern. Obviously, invasive stages preferred
regions that were already occupied by C. muris. This
evidently non-competitive behaviour could be understood
as exploiting the colonization of epithelial
tissue (cell) already modified by the parasite. Stained
smear preparations, showing parasites oriented by
their apical ends to each other, confirmed this
speculation.
Pathological changes
Both host models displayed the same pathological
manifestations, but significantly differed in the
chronology of parasitization. Taylor et al. (1999)
observed the correlation between increased infective
dose and subsequent shortening of the prepatent
period of mice as well as the increased shedding of
oocysts. In this study, increased infective doses
used for inoculation did not influence the success of
parasite detection in the host stomach in the early
course of parasitization, even if all mentioned
detection methods including molecular tools were
used.
The pathological alterations induced by the multiplication
of cryptosporidia in BALB/c mice, were
compared with other studies (Ozkul and Aydin,
1994; Aydin and Ozkul, 1996; Taylor et al. 1999;
Kváč et al. 2008). Our observations correspond
with previous reports on the self-curing process in
BALB/c mice infected with different strains of
C. muris linked with migration and proliferation of
T-cells in the site of infection (Jalovecká et al. 2010;
Kváč et al. 2011).
In M. coucha, the immune system obviously failed
and modifications of gastric tissue persisted during
the entire chronic phase of infection. The parasite
continued to expand from affected gastric pits to the
surrounding healthy gastric epithelium and these
observations are in concordance with previous study
(Valigurová et al. 2008). Kváč et al. (2008) and Kváč
and Vítovec (2003) observed the dispersal of parasitization
in the abomasum of domestic ruminants
from the region of the curvature major. Pathological
alterations in this study concur with their histological
observations. In contrast to Taylor et al. (1999), the
infiltration of the lamina propria was not observed
and these findings correspond with the study of Kváč
et al. (2008).
Life cycle
In general, excystation is a very rapid process (Lumb
et al. 1988). As it was almost impossible to record
invading stages and the first observed stages were
usually trophozoites, more sensitive molecular tools
were used to detect C. muris DNA. Considering the
irregular manner of parasitization, negative results
of PCR for the presence of C. muris in stomachs of
M. coucha could be the consequence of tissue excision
from the non-parasitized part. Although molecular
methods specifically proved the presence of C. muris
DNA in the feces or stomach, they did not provide
information concerning the presence of its concrete
developmental stages, i.e. residual oocysts passing
through the body vs real infection. For example, after
unsuccessful attachment of zoites to the epithelium
or due to the mechanical detachment of the PS in
the area of the dense band reported previously
(Valigurová et al. 2008), various stages apart from
oocysts might be present in host feces. This could
also explain positive testing of M. coucha duodenum
and jejunum. Miller and Schaefer (2007) claim
that intact or excysted oocysts in the first hours
following inoculation were neither detected in the
host (stomach, caecum and colon) nor in its feces.
They suggested that oocysts were destroyed during
normal transit through the gut to the feces.
Considering the persistence of sporulating oocysts
released from the stomach and passing through the
intestine to the feces, it is questionable whether
well-developed and resistant oocysts used for inoculation
could be destroyed by digestion if not excysted.
Acknowledging such a possibility, sporulating
oocysts leaving the host should also be destroyed.
299Life cycle of Cryptosporidium muris in two rodent models
Several studies highlight extraordinary epicellular
development of cryptosporidia and their
phylogenetic affinity with gregarines (e.g. Barta and
Thompson, 2006; Valigurová et al. 2007; Valigurová
et al. 2008). Despite few studies reporting a complete
development of cryptosporidia in cell-free cultures
(Hijjawi et al. 2001, 2010; Borowski et al. 2010), we
did not noticed any extracellular stages resembling
those of gregarines and developing without host cells.
This study did not focus on the formation of the
attachment site and the epicellular localization of
C. muris as newly obtained data confirmed our
previous published findings (Valigurová et al. 2007,
2008). Because of the unique niche of C. muris
developing within the PS, this parasite requires a
more sophisticated defence strategy against the
unfriendly host environment when compared with
typical intracellular coccidians. The PS is a product
of the host cell extensions enveloped by the plasma
membrane, modified by products of the parasite
(Robert et al. 1994; Bonnin et al. 1995; McDonald
et al. 1995; O’Hara et al. 2004; Valigurová et al. 2007,
2008) and thus perfectly masks the parasite from host
immune mechanisms (McDonald et al. 1992, 1996).
Although we noticed the formation of several layers
of PS in early stage of trophozoites, we did not
observe the pre-parasitophorous vacuole described
by Huang et al. (2004).
Generally, the occurrence of two generations of
merogony is described for cryptosporidia (Vetterling
et al. 1971; Current and Reese, 1986) and even
merogony including three generations has been
observed for C. baileyi (Current et al. 1986). Only
one published report on Type II merogony in
C. muris exists to date (Aydin, 1999), but because
this is exclusively based on TEM, it cannot be
considered reliable. This observation might be
influenced by the plane of sectioning and only serial
sectioning or a complete view of the entire parasite (as
obtained by LM or SEM) would avoid any possible
misinterpretations. This study, however, succeeds to
reliably document the presence of both the Type I
and Type II merogony. Considering three cycles of
mitotic divisions in Type I meronts, it might be
possible that Type II merogony comprises only two
generations of nuclei, resulting in the formation of
four merozoites. In addition, an anomaly in the final
number of Type I merozoites often occurred. In
accordance with a previous TEM study on C. muris
(Uni et al. 1987), we observed subpellicular microtubules
in merozoites. The failure of studies on
C. parvum (Current and Reese, 1986) and C. wrairi
(Veterling et al. 1971) to document subpellicular
microtubules, might be due to the known instability
of these structures and to their frequent damage due
to improper fixation. In addition, as the basic
mechanism of apicomplexan motility is expected to
be based on the orientation of the actomyosin motor
by subpellicular microtubules (Dubremetz et al.
1998), it is not likely that motile merozoites lack
these structures.
Concerning the sexual phase of the life cycle, the
most easily recognized stages were macrogamonts.
Microgamonts are often found in close proximity to
macrogamonts, probably to facilitate macrogamont
fertilization, considering that microgametes are nonflagellated.
The ratio of their occurrence in comparison
to that of macrogamonts, however, was much
lower. This might be the consequence of a large
number of microgametes (up to 16) produced by a
single microgamont, that are sufficient to fertilize the
surrounding macrogamonts. The presence of minute
microgamonts in low numbers might be the reason
for our failure to detect them in the early phase of
parasitization (they were detected later than oocysts).
Fertilization as described previously (Current and
Reese, 1986; Aydin and Ozkul, 1996), was not
observed using TEM, but we observed migrating
microgametes towards the macrogamont surface and
found similar coupled individuals in smears stained
with Lugol’s iodine as well as in SEM preparations.
Endogenous stages of thick-walled oocysts were
detected in the stomachs of both hosts. In the same
time period, oocysts enveloped by an obviously
thinner wall were noticed in BALB/c mice, whereas
similar ‘thin-walled’ oocysts in M. coucha were
absent. We speculate that sporulated oocysts might
also have been present during earlier phases of
infection in M. coucha, but stomach dimensions and
anatomical features probably did not allow their
detection. Since the first microscopic detection of
sporulated oocysts of C. muris in host feces was
possible only after several hours or even days, the
destiny of oocysts found inside the host stomach and
identified as thick-walled oocysts remains enigmatic.
In the course of early parasitization, we repeatedly
observed structures resembling empty oocysts in
mucosal scrapings or fecal smears, and often, the wall
of these structures appeared to be very thin. We
consider them to be oocysts, but since the identification
of developmental stages based exclusively on
native (unstained) preparations is problematic and
usually not reliable, we cannot conclude this with
absolute certainty. Current and Reese (1986) claimed
to show endogenous thick-walled oocysts releasing
sporozoites (Fig. 23 in their study), but this stage
more resembles an earlier stage of merogony than an
oocyst. Numerous released sporozoites found in
mucosal scrapings 6 dpi in BALB/c mice and 14 dpi
in M. coucha support the existence of oocysts that
release sporozoites in an early phase of infection.
Considering the variability of wall thickness in
mature oocysts observed in ultrathin sections during
this study, several conflicts arose. Firstly, it appears
almost impossible to strictly identify oocysts as thickor
thin-walled, even via electron microscopy.
Furthermore, the size of the oocyst wall in cryptosporidia
is not uniform and varies significantly
300Janka Melicherová and others
during oocyst maturation. The study that reported
the existence of so-called ‘thin-walled’ oocysts
(Current and Reese, 1986; Aydin, 1999), described
them as structures comprising a single unit membrane,
in contrast to that of thick-walled oocysts
that contain both the inner and outer membrane.
Despite the high-quality micrographs showing the
wall in maturing oocysts in detail, the magnification
in micrographs showing sporulated oocysts in the
mentioned study is not sufficient to evaluate the true
number of oocysts in membranes. Moreover, we
often observed detached PS containing oocysts in
various stages of sporulation and the outer layer
reported to be the outer membrane of the oocyst wall
in Fig. 45 in Current and Reese (1986) actually more
resembles the PS shown in micrographs showing
attached stages. Whether the variability in the
thickness of the oocyst wall found in the stomach is
the consequence of oocyst structural disorder, their
malfunction or simply misinterpretation of observations
caused by the oblique plane of ultrathin
sectioning, remains open. The ‘thin-walled’ oocysts
are believed to be responsible for autoinfection.
Nevertheless, oocysts release only four sporozoites
equipped with a single rhoptry and thus having only
a single attempt for successful attachment to the
suitable host cell (Tetley et al. 1998; Petry and Harris,
1999; Blackman and Bannister, 2001; Petry, 2004;
O’Hara et al. 2005). In contrast, merogony produces
six to eight merozoites, which seem to possess several
rhoptries and numerous micronemes (Figs 5J and
5T in this study), providing them more chances for
successful attachment (Current and Bick, 1989;
Tetley et al. 1998; Jirků et al. 2008). Since the cyclic
multiplication of parasites during Type I merogony
(asexual division resulting in identical daughter cells)
is undoubtedly more than sufficient for the infestation
of the host tissue by C. muris and the zygote is
the only heterozygous diploid stage in the life cycle of
cryptosporidia (McLauchlin and Nichols, 2002), the
multiplication via autoinfective oocysts (sexual stage)
appears to be of benefit to increase the genetic
variability of the parasite and thereby its fitness and
probably infectivity.
CONCLUSIONS
A combined approach used in this study was crucial
to achieve reliable and complex data on the life cycle
of C. muris in two host models. The most problematic
issue proved to be the detection of parasite developmental
stages in gastric pits during the early phase of
parasitization. Light micrographs helped to record
the presence of individual developmental stages of
C. muris in host stomachs and to visualize their
typical morphological features in staining preparations,
and these are in accordance with Tyzzer’s
detailed depictions shown in his work (Tyzzer, 1910).
The Wright staining procedure has been evaluated to
be the most suitable method, since it allows a reliable
and relatively rapid identification of all developmental
stages found in scrapings from host stomach.
ACKNOWLEDGEMENTS
The authors are greatly indebted to Ing. Lenka Hlásková
and RNDr. Dana Květoňová from the Laboratory of
Veterinary and Medical Protistology, and to the staff of the
Laboratory of Electron Microscopy (both groups from the
Institute of Parasitology, Academy of Sciences of the Czech
Republic, in České Budějovice), for their valuable advice
and help. Many thanks to Monika Nechvílová from the
Department of Pathological Morphology and Parasitology,
University of Veterinary and Pharmaceutical Sciences, for
her help with laboratory animals.
FINANCIAL SUPPORT
Financial support was provided by a postdoctoral grant
GPP506/10/P372 from the Czech Science Foundation
(Investigator: A.V.). J.M., J.I. and B.S. were partially
supported by the project GAP505/11/1163 from the Czech
Science Foundation and M.K. by a grant from the Ministry
of Education, Youth and Sports of the Czech Republic
(LH11061). The authors acknowledge support from the
Department of Botany and Zoology, Faculty of Science,
Masaryk University, in the preparation of this manuscript.
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303Life cycle of Cryptosporidium muris in two rodent models
7.4
Valigurová A., Paskerova G.G., Diakin A., Kováčiková M.,
Simdyanov T.G.
2015
Protococcidian Eleutheroschizon duboscqi, an unusual
apicomplexan interconnecting gregarines and cryptosporidia
PLoS One 10(4), e0125063
RESEARCH ARTICLE
Protococcidian Eleutheroschizon duboscqi, an
Unusual Apicomplexan Interconnecting
Gregarines and Cryptosporidia
Andrea Valigurová1
*, Gita G. Paskerova2
, Andrei Diakin1
, Magdaléna Kováčiková1
,
Timur G. Simdyanov3
1 Department of Botany and Zoology, Faculty of Science, Masaryk University, Kotlářská 2, 611 37, Brno,
Czech Republic, 2 Department of Invertebrate Zoology, Faculty of Biology, Saint-Petersburg State
University, Universitetskaya emb. 7/9, St. Petersburg, 199034, Russian Federation, 3 Department of
Invertebrate Zoology, Faculty of Biology, Lomonosov Moscow State University, Leninskiye Gory 1–12,
Moscow, 119234, Russian Federation
* andreav@sci.muni.cz
Abstract
This study focused on the attachment strategy, cell structure and the host-parasite interactions
of the protococcidian Eleutheroschizon duboscqi, parasitising the polychaete Scoloplos
armiger. The attached trophozoites and gamonts of E. duboscqi were detected at
different development stages. The parasite develops epicellularly, covered by a host cellderived,
two-membrane parasitophorous sac forming a caudal tipped appendage. Staining
with Evans blue suggests that this tail is protein-rich, supported by the presence of a fibrous
substance in this area. Despite the ultrastructural evidence for long filaments in the tail, it
stained only weakly for F-actin, while spectrin seemed to accumulate in this area. The attachment
apparatus consists of lobes arranged in one (trophozoites) or two (gamonts) circles,
crowned by a ring of filamentous fascicles. During trophozoite maturation, the internal
space between the parasitophorous sac and parasite turns translucent, the parasite trilaminar
pellicle seems to reorganise and is covered by a dense fibrous glycocalyx. The parasite
surface is organised in broad folds with grooves in between. Micropores are situated at the
bottom of the grooves. A layer of filaments organised in bands, underlying the folds and
ending above the attachment fascicles, was detected just beneath the pellicle. Confocal microscopy,
along with the application of cytoskeletal drugs (jasplakinolide, cytochalasin D,
oryzalin) confirmed the presence of actin and tubulin polymerised forms in both the parasitophorous
sac and the parasite, while myosin labelling was restricted to the sac. Despite positive
tubulin labelling, no microtubules were detected in mature stages. The attachment
strategy of E. duboscqi shares features with that of cryptosporidia and gregarines, i.e. the
parasite itself conspicuously resembles an epicellularly located gregarine, while the parasitophorous
sac develops in a similar manner to that in cryptosporidia. This study provides a
re-evaluation of epicellular development in other apicomplexans and directly compares
their niche with that of E. duboscqi.
PLOS ONE | DOI:10.1371/journal.pone.0125063 April 27, 2015 1 / 27
a11111
OPEN ACCESS
Citation: Valigurová A, Paskerova GG, Diakin A,
Kováčiková M, Simdyanov TG (2015) Protococcidian
Eleutheroschizon duboscqi, an Unusual
Apicomplexan Interconnecting Gregarines and
Cryptosporidia. PLoS ONE 10(4): e0125063.
doi:10.1371/journal.pone.0125063
Academic Editor: Tobias Spielmann, Bernhard
Nocht Institute for Tropical Medicine, GERMANY
Received: December 23, 2014
Accepted: March 20, 2015
Published: April 27, 2015
Copyright: © 2015 Valigurová et al. This is an open
access article distributed under the terms of the
Creative Commons Attribution License, which permits
unrestricted use, distribution, and reproduction in any
medium, provided the original author and source are
credited.
Data Availability Statement: All relevant data are
within the paper.
Funding: AV, AD and MK were supported by the
Czech Science Foundation, Project no. P505/12/
G112 (ECIP). GGP was supported with grants from
St. Petersburg State University (1.42.514.2013,
1.42.1277.2014). TGS was supported by the grant of
the Council of President of the Russian Federation
NSh-1801.2014.4. A part of the electron microscopic
analysis was performed at the User Facilities Center
of Lomonosov Moscow State University under
financial support of Ministry of Education and Science
Introduction
Phylum Apicomplexa Levine 1980, emend. Adl et al. 2012 [1] represents one of the most successful
groups of eukaryotic unicellular organisms, consisting entirely of parasitic genera that
infect a broad range of vertebrates and invertebrates. In contrast to the apicomplexan etiologic
agents of globally significant human and animal diseases (e.g. malaria, toxoplasmosis, cryptosporidiosis),
the enormously diversified basal groups of Apicomplexa, that are restricted to invertebrate
hosts, remain poorly understood. Nevertheless, they appear to be very important in
the comprehension of evolutionary pathways and phylogenetic relations within the phylum
Apicomplexa.
Apicomplexans evolved various adaptations for invading and surviving within their hosts. It
is assumed that ancestral apicomplexans parasitised marine annelids, and their radiation and
adaptation to the parasitic life style took place before the era of vertebrates. First, they spread to
other marine invertebrates (turbellaria, crustaceans, echinoderms, etc.), then to freshwater and
terrestrial invertebrates, and finally to vertebrates [2]. The apicomplexan zoite is characterised
by a high degree of cell polarity in that it has an apical pole equipped with a so-called apical
complex, usually comprising specialised secretory organelles (rhoptries, micronemes), polar
rings, and a conoid. This unique invasion apparatus, traditionally used as the best defining feature
for the phylum Apicomplexa, is initially linked with a myzocytosis-based mode of feeding
as it is in colpodellids and archigregarines [3,4]. Most likely, apicomplexan evolution progressed
from myzocytotic predation to myzocytotic extracellular parasitism, a characteristic of
lower gregarines and cryptosporidia, and finally to intracellular parasitism which is typical for
coccidia. This means apicomplexans demonstrate two main determinative evolutionary trends:
i) the origination of intracellular parasitism in typical coccidia and Aconoidasida, accompanied
by a rejection of trophozoite polarity and motility; and ii) the origination of epicellular parasitism,
observed mostly in gregarines, with subsequent modifications of attachment apparatus
along with the motility mode/mechanism in the trophozoite stage. Recent studies have pointed
out the unique epicellular localisation of cryptosporidia within the host cell-derived parasitophorous
sac (PS), and the similarities in their attachment and feeding strategy with gregarines.
Thus, these parasites reflect analogous modes of adaptation to a similar environment within
the host [5,6]. Based on phylogenetic analyses reporting the close affinity of gregarines and
cryptosporidia [7,8], speculation that cryptosporidia represent a ‘missing link’ between the
gregarines and coccidia is frequently discussed.
One of the poorly explored basal apicomplexan lineages is the order Protococcidiorida
Kheisin, 1956 (subclass Coccidiasina Leuckart, 1879; class Conoidasida Levine, 1988) comprising
four families: Eleutheroschizonidae Chatton & Villeneuve, 1936; Myriosporidae Grassé,
1953; Angeiocystidae Léger, 1911 and Grelliidae Levine, 1973 [9]. Protococcidia are expected
to lack merogony, and their gamogony and sporogony occurs extracellularly [9]. Genus
Eleutheroschizon Brasil, 1906 was placed in the family Eleutheroschizonidae, representatives of
which are characterised by epicellular development [9,10,11]. Their gamonts detach from the
host tissue and disperse into the environment where gametogenesis and sporogenesis take
place. Oocysts contain fan-shaped clusters of sporozoites, with one end of each sporozoite attached
to the residuum [9]. There are reported only two species of the genus Eleutheroschizon,
the type species E. duboscqi Brasil, 1906 from Scoloplos armiger and E. murmanicum Awerinzew,
1908 from Ophelia limacina (Rathke) [11,12]. Apart from the original description [11]
and one study focusing on the life cycle of E. duboscqi [10], no further studies dealing with this
parasite have been published. The aim of this study was to provide a morphological analysis of
the attachment strategy, cell cortex and cytoskeleton of trophozoites and gamonts of
E. duboscqi Interconnecting Gregarines and Cryptosporidia
PLOS ONE | DOI:10.1371/journal.pone.0125063 April 27, 2015 2 / 27
of Russian Federation and at the Core Facility
Centres ‘Culturing of microorganisms’ and
‘Development of molecular and cell technologies’ of
St. Petersburg State University. AV, AD and MK
acknowledge support from the Department of Botany
and Zoology, Faculty of Science, Masaryk University,
towards the preparation of this manuscript. The
funders had no role in study design, data collection
and analysis, decision to publish, or preparation of
the manuscript.
Competing Interests: The authors have declared
that no competing interests exist.
Eleutheroschizon duboscqi, a representative of marine apicomplexans, which shares features of
both the gregarines and coccidia.
Materials and Methods
Material collection
The polychaetes Scoloplos armiger (Müller, 1776) were collected from 2006 to 2014 at the sandsilt
littoral zone close to the White Sea Biological Station of M. V. Lomonosov Moscow State
University (66°33.200' N, 33°6.283' E) and the Marine Biological Station of St. Petersburg State
University (66°18.770' N; 33°37.715' E). Both stations are situated in the Kandalaksha Bay of
the White Sea. The dissection of polychaetes and extraction of parasites were performed using
a MBS-10 stereomicroscope. Squash preparations with living parasites and semi-thin sections
stained with toluidine blue were investigated with the use of a Leica DM 2000 microscope connected
to a DFC 420 digital camera, a Zeiss Axio Imager.A1 connected to an AxioCam MRc5
digital camera or with an Olympus microscope BX61 equipped with DP71 digital camera.
Electron microscopy
Specimens were fixed in an ice bath in 2.5–5% (v/v) glutaraldehyde, in different concentrations
of cacodylate buffer (0.05–0.15 M; pH 7.4; osmolarity was reached up to 720 mOsm by adding
NaCl), for over two hours. For transmission electron microscopy (TEM), the specimens were
then washed in buffer used for fixation or in filtered (0.22 μm Millipore) sea water and postfixed
in 1–2% (w/v) OsO4 in the same buffer for 1–3 h at room temperature. Some specimens
were fixed with 3% glutaraldehyde-ruthenium red [0.15% (w/v) stock water solution] in 0.2 M
cacodylate buffer (pH 7.4) and postfixed with 1% OsO4-ruthenium red in the same buffer. The
subsequent procedure follows published protocols [6,13,14]. Observations were made using
microscopes JEM-1010 (JEOL) and LEO 910 (Zeiss). For scanning electron microscopy (SEM),
the specimens were washed in buffer used for fixation or in filtered (0.22 μm Millipore) sea
water, processed according to Valigurová et al. [13,15] and examined using microscopes JSM-
7401F —FE SEM (JEOL), LEO 420 (Zeiss) or GEMINI Supra 40V (Zeiss).
Confocal laser scanning microscopy
Fragments of parasitised intestines were washed in 0.1 M phosphate buffered saline (PBS),
fixed for 30 minutes at room temperature in freshly prepared 4% paraformaldehyde in 0.1 M
PBS (PFA) or in ice-cold methanol, washed, and permeabilised for 15–30 minutes in 0.5% Triton
X-100. Some of the PFA fixed samples were stained with Evans blue. Protocols for direct
staining of filamentous actin (F-actin) with phalloidin—tetramethylrhodamine B isothiocyanate
(phalloidin-TRITC; Sigma-Aldrich, Czech Republic), as well as indirect immunofluorescent
antibody (IFA) staining using a rabbit anti-myosin antibody (smooth and skeletal, the
whole antiserum), a rabbit anti-chicken spectrin antibody (the whole antiserum), a mouse
monoclonal anti-α-tubulin antibody (Sigma-Aldrich, Czech Republic) and a mouse monoclonal
IgG anti-actin antibody that was raised against Dictyostelium actin (provided by Prof.
Dominique Soldati-Favre), follow Valigurová et al. [13,15]. For double labelling, specimens
were incubated in phalloidin-TRITC after washing off the secondary antibody. Some preparations
were counterstained with DAPI. Confocal laser scanning microscopic (CLSM) observations
were made with an inverted Olympus IX81 microscope equipped with a laser-scanning
FluoView 500 confocal unit (FluoView 4.3 software); using the rhodamine (Evans blue), tetramethylrhodamine
isothiocyanate (TRITC—phalloidin, anti-myosin), fluorescein isothiocyanate
(FITC—anti-actin, anti-α-tubulin, anti-spectrin) and/or UV (DAPI) filter sets. Some
E. duboscqi Interconnecting Gregarines and Cryptosporidia
PLOS ONE | DOI:10.1371/journal.pone.0125063 April 27, 2015 3 / 27
micrographs were processed using the software Fiji (an image processing package based on
ImageJ developed at the National Institutes of Health). For better interpretation of fluorescent
data, a set of 6 linguistic variables, such as No fluorescence (-), Very weak (-/+), Weak (+), Medium
(++), Strong (+++) and Very strong (++++), were used (symbols are only illustrative and
not used in the text). Quantification of fluorescence intensity was made using the visual assessment
of CLSM micrographs (using the raw images), comparison was made to those obtained
from two sets of control samples: negative (omitting the primary antibody or phalloidin) and
positive controls (labelled, but not treated with cytoskeletal drugs).
Experimental part
As a control for potential false positive results from the fluorescent labelling of F-actin and microtubules,
specimens were treated with probes that influence the polymerisation of actin and
tubulin: jasplakinolide (JAS, Invitrogen; a toxin that stabilises actin filaments and induces actin
polymerisation), cytochalasin D (Invitrogen, Czech Republic; a drug disrupting actin filaments
and inhibiting actin polymerisation), and oryzalin (Sigma-Aldrich, Czech Republic; a dinitroaniline
herbicide acting through the disruption/depolymerisation of microtubules). Drugs were
reconstituted in dimethyl sulfoxide to prepare a 1 mM stock solution. The final concentration
of these membrane-permeable probes lower than 5 μM had no obvious effect. To obtain reliable
results on vital cells, final solutions of 10 and 30 μM JAS, cytochalasin D and oryzalin prepared
in filtered (0.22 μm Millipore) sea water were applied. Controls were performed in pure
filtered sea water as well as corresponding concentrations of dimethyl sulfoxide in filtered
sea water.
Results
Light microscopic observations
We observed several development stages of E. duboscqi in squash preparations with living parasites.
The earliest developmental stages, presumably zoites during the invasion process, were
drop-shaped cells, 1 μm in length, with their pointed end attached to the host tissue (Fig 1A).
At the light microscope (LM) level, it was not possible to identify the presence or absence of
any membrane structure enclosing these parasites. The next stage was represented by helmetshaped
trophozoites, 2–10 μm in length, with a wide basal part attached to the host cell, and a
large nucleus located near the base. They were enveloped by a parasitophorous sac (PS) of host
membrane origin (Fig 1B–1E). The caudal part of the sac, in all attached parasites, was usually
prolonged in a prominent translucent, tail-like appendage (Fig 1B–1I). The length of this tail
varied. Gamonts corresponded to the helmet-shaped cells, about 20 μm in length, with a granular
cytoplasm containing numerous light-refracting amylopectin granules (Fig 1F–1N). Two
morphs of E. duboscqi gamonts were observed, i.e. macro- and microgamonts, agreeing with
the life cycle description by Chatton and Villeneuve [10]. Macrogamonts possessed one large
nucleus with a dense nucleolus (Fig 1K), while microgamonts had numerous small nuclei with
fragmented nucleoli (Fig 1M and 1N). During light microscopic observations, parasites along
with their PS frequently detached from host tissue (Fig 1J).
Ultrastructural analysis
With the exception of putative zoite stages (Fig 1A), trophozoites and gamonts of various developmental
stages, corresponding to our light microscopy data (Fig 1B–1N), were observed
under the electron microscope. All these attached stages were covered by a PS.
E. duboscqi Interconnecting Gregarines and Cryptosporidia
PLOS ONE | DOI:10.1371/journal.pone.0125063 April 27, 2015 4 / 27
The earliest trophozoites, about 2 μm in length, that obviously transformed from attached
zoites a short time before fixation, were already enveloped by a loose but complete PS (Fig 2A).
Early trophozoites were barrel-shaped (Fig 2B). The space between the parasite and PS was
filled by numerous vesicles of various sizes. The fine structure of the pellicle was not discernible
in all observed early stages (Fig 2A and 2B). The attachment site at the base of PS was mostly
indistinct (Fig 2B). Ultrathin sections revealed a dense, equally thick and continuous, double
layer which separated the unmodified part of the host cell from its apical region bearing an attached
parasite. Early trophozoites were equipped with subpellicular microtubules (21–23 nm
in outer diameter) connecting to a ring-like structure, presumably a posterior ring (Fig 2B). No
organelles of the apical complex were detected (Fig 2A and 2B).
During progressive maturation, the trophozoites underwent changes in their shape and size
(Figs 2B–2L and 3A–3J). Gradually they took the shape of a helmet and reached up to 10 μm in
Fig 1. Light microscopic observations on attached stages of Eleutheroschizon duboscqi. A. Putative zoite of E. duboscqi invading the host intestinal
epithelium. LM, bright field. B-C. Early trophozoites within an already formed PS. Note the caudal prolongation of the sac into a tail. LM, bright field. D.
Maturing trophozoite. LM, bright field. E. Mature trophozoite. LM, phase contrast. F. A gamont stage. LM, phase contrast. G. Gamont exhibiting a prolonged
tail at the PS. LM, bright field. H. Various stages of trophozoites and gamonts attached to the host intestinal epithelium. LM, differential interference contrast.
I. A macrogamont after fixation in PFA. Note the separation of PS from the parasite cortex. LM, bright field. J. Detached macrogamont still enveloped by a PS.
LM, bright field. K-N. Macrogamonts (K-L) and microgamonts (M-N) in semi-thin sections. LM, bright field, Toluidine blue. arrow—parasite, arrowhead—tail of
the PS, asterisk—parasite attachment site, h—host tissue, n—parasite nucleus/nuclei.
doi:10.1371/journal.pone.0125063.g001
E. duboscqi Interconnecting Gregarines and Cryptosporidia
PLOS ONE | DOI:10.1371/journal.pone.0125063 April 27, 2015 5 / 27
Fig 2. Ultrastructural features of Eleutheroschizon duboscqi early development. A. Early trophozoite in the process of transformation from an attached
zoite, already enveloped by a host-derived PS. TEM. B. Early trophozoite. Note the numerous vesicles in the space between parasite and PS, especially in
caudal region. TEM. C-E. Young trophozoite. D shows the space between the parasite caudal region and PS; E shows the host parasite interface at the
attachment site. TEM. F-H. Maturing trophozoite. G shows an annular joint point of two host membranes; H focuses on the parasite caudal region and PS.
TEM. I. Early trophozoite. SEM. J. Young trophozoite. SEM. K. Mature trophozoite. TEM. L. Detailed view of the attachment site of the trophozoite shown in
K, focusing on the developing fascicles of filaments and the annular joint point. TEM. a—parasite amylopectin, asterisk—space between the parasite and PS,
black arrow—PS, black arrowhead—parasite plasma membrane, black double/paired arrowheads—parasite cytomembranes, c—parasite cytoplasm, cr—
crystalloid body, er—parasite endoplasmic reticulum, fa—attachment fascicles, fi—short attachment filaments, g—glycocalyx, h—host cell, mc—host
microcilia, mt—parasite subpellicular microtubules, mv—host microvilli, n—parasite nucleus, r—parasite posterior ring, sf—parasite subpellicular filaments,
t—tail of the PS, v—vesicles, white arrow—host cell plasma membrane, white arrowhead—dense band, white double arrowhead—base of the PS
(membrane of host cell origin), x—forming attachment fascicles.
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Fig 3. Fine structure of Eleutheroschizon duboscqi mature trophozoites. A. Mature trophozoite transforming into a gamont stage. TEM. B. Detailed view
of the annular joint point and a well-developed fascicle of filaments. TEM. C. The view of mitochondria and a micropore (white circle) at the attachment site.
The inset shows the micropore in detail. TEM. D. Higher magnification of the caudal region. TEM E. Two partially detached, mature trophozoites. SEM. F.
The attachment site of a partially detached, mature trophozoite with well-developed fascicles and short filaments. SEM. G. Diagonal section of the apical part
of a mature trophozoite. TEM. H-I. Craters left after detachment of mature trophozoites with well-developed attachment fascicles. Flat holes organised in one
circle correspond to the developing lobes. SEM. J. A crater left after a trophozoite of more advanced stage as indicated by the presence of one circle of deep
holes corresponding to well-developed lobes and one extra lobe starting the formation of a second circle. SEM. a—parasite amylopectin, black arrow—PS,
black arrowhead—parasite plasma membrane, black asterisk—space between the parasite and PS, black double/paired arrowheads—parasite
cytomembranes, c—parasite cytoplasm, er—parasite endoplasmic reticulum, fa—attachment fascicles, fh—holes in the host tissue left after the fascicles of
the detached parasite, fi—short attachment filaments, g—glycocalyx, h—host cell, l—attachment lobe, lh—holes in the host tissue left after the lobes of the
detached parasite, m—parasite mitochondria, mv—host microvilli, n—parasite nucleus, p—parasite, sf—parasite subpellicular filaments, white arrow—host
cell plasma membrane, white arrowhead—dense band, white asterisk—empty attachment site, white double arrowhead—base of the PS.
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length. The developmental stage, defined as a young trophozoite (3–5 μm in length), was identifiable
due to a large nucleus in the apical position, a crystalloid body of unknown nature, subpellicular
microtubules, and a cytoplasm rich in rough endoplasmic reticulum (Fig 2C–2E).
The pellicle membranes appeared to be folded, loose and still discontinuous in some regions
(Fig 2C–2E). In contrast to the earliest stages, the interface between the unmodified and modified
part of the host cell at the base of the PS appeared clearly delineated, comprising of a membrane
of host cell origin (about 7–9 nm in thickness, corresponding to the plasma membrane
covering adjacent microvilli) underlain by a 4–7 nm thick dense band (Fig 2E). In maturing
trophozoites, the next developmental stage, the pellicle seemed to be more distinct and compact
along the entire surface, and its membranes became more evident than in parasites of previous
developmental stages (Fig 2F). The internal space between the parasite and surrounding
PS continued to clarify so that it appeared translucent with several vesicles. The border between
the parasite PS and the neighbouring microvilli area formed a so called annular joint point.
That is where the host cell plasma membrane, forming the base of PS, comes closer to the plasma
membrane covering the neighbouring microvilli. Both membranes continuously proceed
into the rising PS, forming its inner and outer membranes, respectively (Fig 2G). The 4–7 nm
thick dense band underlying the inner PS membrane in the attachment site ended in this area
and did not continue into the PS. No structures obviously belonging to the parasite and connected
directly to the host-derived membranes of the PS were seen. In the attachment area of
parasites, one ring of filaments began to form just below the annular joint point (Fig 2F). Maturing
trophozoites still exhibited subpellicular microtubules (Fig 2H). In the course of trophozoite
development, the caudal part of the PS gradually formed a prolongation being
characteristic for E. duboscqi, the tail (Fig 2C and 2H). This is easily seen when comparing an
early trophozoite without a tail (Fig 2I) with a young trophozoite with a developing tail on the
PS (Fig 2J). In mature trophozoites, the next developmental stage, a cell coat (glycocalyx) appeared
as a thick layer of fibrous material (Figs 2K, 2L and 3A–3D). The pellicle was distinct; it
was comprised of a plasma membrane, two adjacent cytomembranes (i.e. the inner membrane
complex, IMC) and a thin dense layer underlining the inner cytomembrane (Fig 3C and 3D).
Under the pellicle, a thick layer of subpellicular filaments emerged (Figs 2L and 3B). The cytoplasm
was filled with a large nucleus in the apical position, and an increasing rough endoplasmic
reticulum. The crystalloid body observed in young trophozoites disappeared, while
peripheral amylopectin granules appeared and increased in number (Figs 2K, 3A and 3D).
Large mitochondria underlying the parasite pellicle appeared, especially at the attachment site
(Fig 3C). With increasing age, typical apicomplexan micropores (154 nm in outer diameter
and 132 nm deep when measured from the plane of cytomembranes to the bottom of the micropore)
formed on the parasite surface (Fig 3C). The internal space of the PS enveloping the
mature trophozoites became completely translucent (Fig 3A–3D). At the attachment site of
parasite, the above-mentioned ring of filaments continued to develop: fascicles of long filaments
alternating with short filaments appeared (Figs 2K, 2L, 3A, 3B and 3E–3G). The parasites
formed thick outgrowths, i.e. lobes, which were organised in a single circle at the
attachment site, just below the ring of filaments (Fig 3A and 3E). Correspondingly, craters in
the host tissue, left after detached trophozoites, confirmed the circular organisation of the fascicles
of filaments and lobes at the attachment site (Fig 3H–3J). These were seen as a peripheral
circle of narrow but very deep holes, left after the well-developed fascicles of filaments, and
larger flat holes, organised in one central circle and corresponding to the developing lobes (Fig
3H and 3I). In trophozoites transforming into gamonts, the attachment lobes became more
and more prominent, as also documented by the deeper holes left after detached individuals
(Fig 3J). In addition, the second circle of lobes started to develop in the centre of the attachment
site (Fig 3A). Accordingly, detached parasites left one peripheral circle of holes corresponding
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to the well-developed lobes and one central, extra hole indicating the start of the formation of a
second circle of lobes (Fig 3J).
The oldest developmental stages observed during our study were gamonts reaching about
20 μm in length (Figs 4A–4K, 5A–5E, 6A–6K, and 7A–7K). All gamonts were contained within
a well-developed PS, with a prominent tail (Fig 4A and 4D–4G). A few individuals bearing two
or three tails were observed (Fig 4K). The surface of gamonts exhibited about 12 (10–13,
n = 20) shallow grooves showing through the PS under the SEM (Figs 4A, 4H, 4I, and 7A).
Macrogamonts were characterised by a centrically located, large, roundish nucleus with one
nucleolus, abundant amylopectin granules, large lipid droplets, and prominent dense bodies
varying in shape and size (Fig 4B and 4D). In contrast, microgamonts possessed several nuclei
(up to 20 in a longitudinal section) with fragmented nucleoli and cell inclusions (amylopectin
granules, lipid droplets) of a smaller size (Fig 4C). Pores were often observed at the caudal region
of the PS (Fig 4E and 4F). Dense substances staining intensively with ruthenium red (RR),
most likely mucosubstances secreted by the parasite, were detected on the PS surface close to
these pores (Fig 4E). The internal space between the PS and parasite was mostly translucent,
while the internal space of the PS tail was packed with thin filaments running longitudinally
(Fig 4E and 4G). The parasite surface (Fig 4G and 4J) bore a dense layer of fibrous glycocalyx
(80–85 nm in thickness). Gamonts were often observed with a ruptured PS revealing their surface
(Fig 4H and 4I). In such cases, the parasite pellicle in the caudal region was covered by a fibrous
material (Fig 4H) that might correspond to the filaments observed in the PS tail under
TEM. Small piles of unknown material organised in circles were observed to be present on the
caudal part of the parasite surface (Fig 4I). The dense glycocalyx was visible under SEM as a
woolly coat covering the surface (Fig 4I). Completely detached parasites, but still enveloped by
a PS and located far away from the host tissue, were rarely detected in ultrathin sections.
Observations on mechanically detached gamonts under SEM revealed their complicated attachment
sites, comprising of two circles of massive lobes crowned by a ring of fascicles of long
filaments, alternating with short filaments (Fig 5A–5D). Detached individuals were covered by
a PS, except for their attachment sites that seemed to be covered by a parasite pellicle only. The
naked attachment lobes appeared as smooth protuberant, hemispheric structures. Detached
gamonts left characteristic craters on the host intestinal tissue, indicating that they were fully
matured, with well-developed attachment fascicles and two circles of lobes (Fig 5E–5G). The
basal part of the PS was comprised of the host membrane (i.e. the inner membrane of PS) underlined
by a dense band and was still present after parasite detachment (Fig 5H), corresponding
to the observations of detached parasites with naked attachment sites.
Ultrathin sections of attached gamonts confirmed the SEM observations on the attachment
site, and showed that lobes represent structures belonging to the parasite, as they are covered
by a parasite pellicle and are filled with its cytoplasm packed with large mitochondria, various
vesicles and the endoplasmic reticulum (Fig 6A, 6B, 6G and 6H). The pellicle membranes covering
the lobes were usually well-preserved (Fig 6G). In contrast to lobes, attachment fascicles
were located in the translucent space between the parasite and PS, and were not covered by the
parasite pellicle (Fig 6A, 6B, 6D, 6E, 6J and 6K). Although the membranes of the parasite pellicle
were not clearly distinguishable in this area, the short filaments (15–35 nm thick) and fascicles
of longer filaments (about 60 nm thick) that seemed to arise from the pellicle and evidently
extended through the glycocalyx, were deeply anchored into the IMC (Fig 6C–6E, 6J and 6K).
Some sections clearly showed the hook-shaped short filaments that were anchored into the
outer cytomembrane of IMC (Fig 6C). The subpellicular layer of filaments, localised just beneath
the IMC, ended above the ring of the filaments and fascicles (Fig 6D and 6E). The organisation
of structures at the annular joint point corresponded with the observations on younger
stages (Fig 6F). The dense band (4–8.5 nm thick) underlying the inner PS membrane at the
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Fig 4. Morphology of Eleutheroschizon duboscqi gamonts. A. Attached gamont. SEM. B. Macrogamont with a large central nucleus. TEM. C.
Microgamont with several nuclei. TEM. D. Macrogamont enclosed by host tissue. TEM, RR. E. The PS tail of the macrogamont shown in D. Note the pores
and the mucosubstances present in their surroundings. TEM, RR. F. High magnification of the caudal PS part with the tail showing numerous pores. SEM. G.
Detailed view of the tail and gamont caudal part. TEM, RR. H. Upper view of an individual with a ruptured PS. SEM. I. The caudal region of a naked individual.
SEM. J. High magnification of the interface between the parasite and PS in the area of the tail. TEM, RR. K. Gamont with two tails at the PS. SEM. a—
parasite amylopectin, arrow—PS, asterisk—space between the parasite and the PS, black arrowhead—parasite plasma membrane, black double/paired
arrowheads—parasite cytomembranes, c—parasite cytoplasm, db—parasite dense bodies, fa—attachment fascicles, g—glycocalyx, h—host cell, l—
attachment lobe, ld—parasite lipid droplets, m—parasite mitochondria, mc—host microcilia, n—parasite nucleus, p—parasite, po—pore, s—
mucosubstances, t—tail of the PS, v—vesicles, white arrowhead—base of the PS.
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attachment site ended in this area. Numerous typical micropores (130–155 nm in outer diameter,
40–50 in inner diameter, the distance between the lumen of the duct and collar periphery is
about 50–58 nm) were distributed at the attachment site of the parasite, especially in between
individual lobes (Fig 6G–6I). Vesicles were rarely seen to be connected with micropores located
at the attachment site of the parasite (Fig 6H). At the basal part of PS, an accumulation of
Fig 5. Architecture of attachment site of Eleutheroschizon duboscqi gamonts. A. Host intestinal tissue with a detached gamont revealing its attachment
site at the base of PS. SEM. B. Detail of the gamont attachment site. SEM. C. Detailed view of the fascicles of long filaments alternating with short filaments,
organised in ring. SEM. D. A detail of attachment fascicles. SEM. E. Host intestinal tissue with an attached parasite and a crater left after detached ones.
SEM. F. A detail of crater left after gamont with well-developed attachment fascicles and two circles of lobes. SEM. G. Host epithelium showing the crater left
after the parasite detached. TEM. H. A detail of the PS membrane remains covering the crater. TEM. arrow—PS, fa—attachment fascicle of filaments, fh—
holes in the host tissue left after the fascicles of the detached parasite, fi—short attachment filaments, l—attachment lobe, lh—holes in the host tissue left
after the lobes of the detached parasite, mv—microvilli and cilia of the host enterocyte, p—parasite, white arrowhead—dense band, white asterisk—empty
attachment site, white double arrowhead—base of the PS.
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Fig 6. Fine structure of the attachment site of Eleutheroschizon duboscqi gamonts. A. Macrogamont with a ruptured PS. TEM. B. Oblique section of the
attachment site. TEM. C. A detail showing the hook-shaped short filaments anchored into the parasite outer cytomembrane. TEM. D-E. The attachment
fascicles in a longitudinal section. The subpellicular layer of filaments is localised just beneath the parasite IMC and ends above the fascicles. TEM; D is stained
with RR. F. The annular joint point of two host membranes. TEM. G. A detail of the attachment lobe packed with endoplasmic reticulum and mitochondria. Note
the cross-sectioned micropore. TEM. H. Detailed view of vesicles connected with the micropores located in the area of attachment lobes. TEM. I. Longitudinal
section of a micropore localised at the parasite attachment site. TEM. J. Detailed view of the attachment fascicles of long filaments alternating with short
filaments. TEM. K. The basal part of PS showing an accumulation of fine filaments in the host cell cytoplasm surrounding the PS invaginations with attachment
fascicles. TEM, RR. a—parasite amylopectin, asterisk—space between the parasite and PS, black arrow—PS, black arrowhead—parasite plasma membrane,
black double/paired arrowheads—parasite cytomembranes, c—parasite cytoplasm, db—parasite dense bodies, er—parasite endoplasmic reticulum, fa—
attachment fascicle of filaments, fi—short attachment filaments, g—glycocalyx, h—host cell, hf—filaments in host cell cytoplasm, l—attachment lobe, ld—
parasite lipid droplets, m—parasite mitochondria, mv—microvilli and cilia of the host enterocyte, n—parasite nucleus, sf—parasite subpellicular filaments, v—
parasite vesicle, white arrow—host cell plasma membrane, white arrowhead—dense band, white double arrowhead—base of the PS. Micropores are
indicated by white circles.
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filaments 7–10 nm thick (most likely of F-actin nature) was documented in the host cell cytoplasm
surrounding the invaginations of PS with attachment fascicles (Fig 6K).
The cross-sections of mature gamonts confirmed that the surface was organised in weakly
expressed broad folds with shallow grooves between them (Fig 7B). Each fold was underlain by
a broad band of subpellicular filaments (apparently, it corresponds to the thick layer of subpellicular
filaments observed in maturing trophozoites). Various planes of sectioning confirmed
that 5–9 nm thick subpellicular filaments were oriented longitudinally (Fig 7C–7H) and
formed 22–59 nm thick bands. Micropores were located at the bottom of the grooves, i.e. between
these bands. Sectioning also revealed a regular arrangement of interruptions of subpellicular
filaments, corresponding to the distribution of micropores (Fig 7B–7H). A lot of
mitochondria could be observed in the cortical zone of the gamonts, and some were located
so closely to micropores that they seemed to be connected to them (Fig 7F). In contrast to the
individuals with the plasma membrane underlined by two well-preserved cytomembranes
(Fig 7I), several gamonts clearly possessed discontinuous cytomembranes or a disorganised
pellicle. Vesicles present in the area of subpellicular filaments can be evidence that cytomembranes
underwent re-building due to pellicle reorganisation (completion or renewal) in a growing
parasite (Fig 7J and 7K).
Fluorescent visualisation of cytoskeletal elements
For the unspecific visualisation of proteins in E. duboscqi, Evans blue staining viewed with a
rhodamine filter set was used (Fig 8A–8E). Besides typical staining of cytoplasmic contents, it
revealed a relatively high concentration of unspecified proteins in the PS tail and in the area of
the parasite attachment site.
The strong phalloidin labelling revealed a high accumulation of filamentous actin (F-actin)
in the layer corresponding to the host-derived PS as well as in the brush border of the host epithelium
(Fig 8F–8P). The parasite surface and cytoplasm exhibited a fluorescent signal of medium
intensity, similar to the cytoplasm of surrounding host cells. The PS tail exhibited Factin
staining of weak to medium intensity (Fig 8H and 8I). In agreement with the electron
microscopic observations, the basal part of the PS enveloping the trophozoites and young
gamonts showed numerous lobes organised in one circle (Fig 8F–8I), while lobes in mature
gamonts formed two circles (Fig 8J–8P). The attachment site often exhibited brighter fluorescence
than the rest of the parasite enclosed within the PS. After incubation in 10 μM JAS for 9
hours, the F-actin staining became very strong in the area of the PS but only slightly increased
on the parasite surface (Fig 8Q). Treatment for 7 hours, with the concentration of JAS increased
to 30 μM, induced even more advanced stabilisation of actin filaments, resulting in
further amplification of the fluorescent signal (Fig 8R). Interestingly, the treatment with
10 μM cytochalasin D for 9 hours first caused depolymerisation of F-actin in the host tissue
and parasites, while the F-actin restricted to the PS remained intact (Fig 8S) as it stained with
the same intensity as non-treated controls. After incubation in 30 μM cytochalasin D for 7
hours, the fluorescence signal was very weak in all the host tissue, host-derived PS and parasites
(Fig 8T).
Parasites labelled with the specific anti-actin antibody (known to recognise the actin in
Toxoplasma and Plasmodium) exhibited fluorescence signal of medium intensity (Fig 9A and
9B). The immunolocalisation of actin differed from F-actin labelling in that the antibody did
not label the PS, but labelled the host tissue with the same intensity as the parasite enclosed
within the sac (Fig 9B). A slightly increased labelling was noticed at the base of the PS (Fig 9A),
especially when viewing individual optical sections. A weak staining of the PS tail was observed
in all individuals. The treatment with 30 μM JAS for 7 hours resulted in increased labelling of
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Fig 7. Fine structure of a parasitophorous sac and pellicle in Eleutheroschizon duboscqi gamonts. A. Attached gamonts. SEM. B. Cross-sectioned
gamont showing its surface organised in 12 broad folds and shallow grooves corresponding to the regularly arranged interruptions of subpellicular filaments.
TEM. C. Longitudinal section showing the organisation of PS, gamont pellicle and the subpellicular layer of filaments that is repeatedly interrupted in areas
corresponding to the localisation of micropores. TEM, RR. D. Superficial section of the PS and the gamont pellicle. The channel-like structures located in the
space between the PS and parasite correspond to the folding of the PS observed under SEM. TEM, RR. E. Tangential section of the gamont surface
underlined with subpellicular layer of filaments. TEM, RR. F. Diagonal section of the gamont surface revealing mitochondria connected with micropores.
TEM, RR. G. Cross-section of pellicle showing the subpellicular filaments interrupted in the micropore area. TEM, RR. H. Almost longitudinal section of
pellicle with interrupted subpellicular filaments. TEM, RR. I. Pellicle with continuous cytomembranes. TEM. J-K. Re-building of the parasite IMC indicated by
the discontinuous cytomembranes and numerous vesicles located between the parasite plasma membrane and the subpellicular layer of the filaments. TEM,
RR. a—parasite amylopectin, arrow—PS, asterisk—space between the parasite and PS, black arrowhead—parasite plasma membrane, black double/paired
arrowheads—parasite cytomembranes, db—parasite dense bodies, er—parasite endoplasmic reticulum, g—glycocalyx, h—host tissue, ld—parasite lipid
droplet, m—parasite mitochondria, p—parasite, sf—parasite subpellicular filaments, v—vesicles, white arrowheads—channel-like structures. Micropores are
indicated by white circles, interruptions of subpellicular filaments—by white asterisks.
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Fig 8. Fluorescent visualisation of an Eleutheroschizon duboscqi parasitophorous sac. A. Macrogamont stained with Evans blue. CLSM (lower) and
CLSM in a combination with transmission LM (upper two). B-E. Trophozoites (B-D) and a gamont (E) stained with Evans blue. CLSM, output image not
coloured. F-H. Localisation of F-actin in trophozoites. One circle of lobes is visible in the attachment site of the trophozoite shown in G. CLSM in a
combination with transmission LM (F) and CLSM (G, H), phalloidin-TRITC/DAPI. I. F-actin labelling of a putative young microgamont with two primary nuclei.
CLSM, phalloidin-TRITC/DAPI. J. F-actin in a microgamont with numerous nuclei. CLSM, phalloidin-TRITC/DAPI. K-M. F-actin in a putative macrogamont.
CLSM in a combination with transmission LM (K) and CLSM (L, M), phalloidin-TRITC/DAPI. N-P. F-actin in a macrogamont equipped with attachment lobes
organised in two circles. CLSM, phalloidin-TRITC. The intensity of signal for F-actin shown in F-P was strong for PS and medium for parasites. Q. Labelling of
F-actin in an individual treated for 9 hours with 10 μM JAS showing the very strong labelling of PS. Individual optical sections also revealed a slightly
increased F-actin labelling of the parasite. CLSM, phalloidin-TRITC/DAPI. R. Treatment with 30 μM JAS for 7 hours resulted in further increase of F-actin
labelling in the PS, parasite and host tissue. The individual with several nuclei corresponds to the microgamont stage. CLSM, phalloidin-TRITC/DAPI. S.
Visualisation of F-actin in an individual (putative young microgamont with two primary nuclei) treated for 9 hours with 10 μM cytochalasin D. Note the strong
labelling of PS in contrast to the parasite and host tissue exhibiting only very weak signal. CLSM, phalloidin-TRITC/DAPI. T. Very weak F-actin labelling in a
specimen treated for 7 hours with 30 μM cytochalasin D. CLSM, phalloidin-TRITC/DAPI. A-L, N-O, Q-T are composite views created by flattening a series of
optical sections, while M and P represent single median optical sections. All samples were fixed in PFA. arrow—tail of the PS, asterisk—parasite attachment
site, black arrowhead—PS, h—host tissue, n—parasite nucleus, white arrowhead—parasite pellicle.
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Fig 9. Immunolocalisation of Eleutheroschizon duboscqi cytoskeletal proteins. A-B. Actin labelling with a medium intensity in a trophozoite (PFA
fixation). CLSM, IFA (A) and CLSM in a combination with transmission LM, IFA/DAPI (B). B represents a single median optical section. C. Actin labelling in a
macrogamont treated with 30 μM JAS for 7 hours (PFA fixation). Note the increased accumulation of parasite actin (FITC) organised in longitudinal bands
exhibiting strong fluorescence and strong F-actin (TRITC) labelling with a diffuse character. CLSM, IFA/phalloidin-TRITC. D. A gamont exhibiting a more
diffuse actin (FITC) labelling of medium intensity after treatment with 10 μM cytochalasin D for 9 hours (PFA fixation). The F-actin (TRITC) labelling of the
parasite did not change significantly. CLSM, IFA/phalloidin-TRITC. E. Very strong myosin (TRITC) labelling restricted to the PS and host tissue (PFA
fixation). CLSM, IFA/DAPI. F. Strong spectrin (FITC) labelling of the PS in a macrogamont (PFA fixation). CLSM, IFA/DAPI. Single median optical section. G.
Labelling of α-tubulin (FITC) of strong intensity in a young microgamont (PFA fixation). CLSM, IFA/DAPI. H-I. A trophozoite (fixed in ice-cold methanol)
exhibiting a labelling of medium intensity for α-tubulin (FITC) and very strong intensity for myosin (TRITC). CLSM, IFA/DAPI. J. Labelling of α-tubulin (FITC)
and myosin (TRITC) in an early trophozoite treated for 7 hours with 10 μM oryzalin (fixed in ice-cold methanol). The fluorescence signals for both antibodies
did not change significantly. CLSM, IFA. K-L. Localisation of α-tubulin (FITC) and myosin (TRITC) in an individual (probably a young microgamont) treated
with 30 μM oryzalin for 3 hours (fixed in ice-cold methanol). The fluorescence signal for tubulin became very weak, while it remained very strong for myosin.
CLSM, IFA/DAPI. M. Co-localisation of α-tubulin (FITC) and F-actin (TRITC) in a macrogamont treated for 7 hours with 10 μM oryzalin (PFA fixation). CLSM,
IFA/phalloidin-TRITC. N-O. Labelling of α-tubulin (FITC) and F-actin (TRITC) in a maturing trophozoite treated for 3 hours with 30 μM oryzalin (PFA fixation).
CLSM, IFA/phalloidin-TRITC/DAPI. In both the preparations (M-O), there was almost no fluorescence signal for α-tubulin, while the F-actin labelled with a
strong intensity. arrow—tail of the PS, asterisk—parasite attachment site, black arrowhead—PS, h—host tissue, n—parasite nucleus, white arrowhead—
parasite pellicle.
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actin with a strong fluorescence signal, revealing its higher accumulation in longitudinal bands
corresponding to the localisation of subpellicular filaments (Fig 9C). Nevertheless, the counterstaining
with phalloidin did not show increased actin polymerisation in this area; the strong Factin
labelling of the parasite had diffuse character. The treatment with 10 μM cytochalasin D
for 9 hours caused more homogenous labelling of actin with a medium intensity dispersed
within the parasite cytoplasm, but only slightly decreased staining of F-actin (Fig 9D). The very
strong labelling of myosin was restricted to the PS and host tissue (Fig 9E and 9H). Spectrin appeared
to be dispersed in low concentrations in the parasite cytoplasm and surrounding host
tissue (medium signal), while the PS, especially in the tail region, stained with a strong intensity
(Fig 9F). Immunolabelling with an anti-α-tubulin antibody, used for visualisation of subpellicular
microtubules and related structures, was repeatedly very strongly positive for the brush
border of the host intestinal epithelium densely covered by microcilia. Both the parasite and
the PS unexpectedly (formaldehyde is known to not satisfactory preserve microtubules) exhibited
medium to strong labelling in PFA-fixed samples that were processed immediately after
fixation (Fig 9G), while only weak to medium labelling was observed in those fixed in methanol
(Fig 9H and 9I). Though more diffuse, the labelling of the same intensity was still noticeable at
the parasite periphery after treatment with 10 μM oryzalin for 7 hours (Fig 9J). After incubation
in 30 μM oryzalin for 3 hours, the peripheral labelling became weak to very weak in methanol-fixed
samples, and putatively unpolymerised α-tubulin seemed to be more dispersed
throughout the cytoplasm (Fig 9K and 9L). The very strong labelling of myosin in methanolfixed
samples was restricted to the periphery of PS and independent of oryzalin treatment (Fig
9H, 9J and 9K). High doses of oryzalin induced more frequent detachment of E. duboscqi,
along with its sac, from the host tissue. To confirm that the modified microtubules were the
reason for parasite detachment but not the redistribution of F-actin, the correct fixation for
phalloidin staining was essential to retain the quaternary protein structure of F-actin (methanol
destroys its native conformation and is not suitable for F-actin staining with phalloidin).
Hence, control PFA fixation was performed for the co-localisation F-actin and α-tubulin. Parasites
treated either with 10 μM or 30 μM oryzalin and subsequently fixed in PFA exhibited no
changes in the F-actin distribution (Fig 9M and 9N), except for, when compared to the rest of
PS, stronger labelling of the tail (Fig 9N). In contrast to methanol-fixed samples, the labelling
with anti-α-tubulin antibody was almost undetectable in parasites and less conspicuous in the
host brush border (Fig 9M–9O).
Discussion
This study confirmed the epicellular localisation of the protococcidian Eleutheroschizon
duboscqi on the gut epithelium of the polychaete Scoloplos armiger, as described in the original
studies [10,11]. We use the term ‘parasitophorous sac’ to underline the peculiar localisation
of the developmental stages of E. duboscqi in the host-derived two-membrane structure.
Parasitophorous sac is the preferable term introduced for the first time by Paperna and Vilenkin
[16] for the host-derived structure enveloping cryptosporidia. We believe that the term
‘parasitophorous vacuole’ to describe the location of epicellular organisms like Cryptosporidium
and Eleutheroschizon is misleading, because it refers solely to a vacuolar space bordered
by a membrane [5,17]. The parasitophorous sac (PS) is an epicellular structure (niche) enveloping
the entire parasite composed of two continuous host plasma membranes on the
outer and inner sides, enclosing a thin layer of host cell cytoplasm. In addition, a dense band
of microfilaments separates the unmodified and modified parts of the host cell ([5], this
manuscript).
E. duboscqi Interconnecting Gregarines and Cryptosporidia
PLOS ONE | DOI:10.1371/journal.pone.0125063 April 27, 2015 17 / 27
Fig 10. Schematic diagram of host-parasite interactions in Eleutheroschizon duboscqi, eugregarines, cryptosporidia, and epicellular eimeriids.
The diagrams of E. duboscqi, gregarines and cryptosporidia are based on our personal observations enriched by published data. The diagram of eimeriids
represents our interpretation and summary of published micrographs, where only maturing or mature stages were clearly shown [36–38,40,42– 44,64,65]. In
this scheme, we refer to the host-derived envelope (described as a parasitophorous vacuole throughout literature) of eimeriids in epicellular location as a
parasitophorous sac (PS) due to its organisation similar to that in cryptosporidia and E. duboscqi. Three colours are used to distinguish between the parasite
(in purple), the host cell including its parts modified due to parasitisation (in pink) and the contact zone between the host and the parasite (in yellow) where the
interrelationships of the two organisms become more intimate. In the case of host-parasite cellular interactions in E. duboscqi and epicellular eimeriids, the
internal space between the parasite and PS remained colourless, even though we do not exclude the possibility that this region may serve as a transitional
zone for intensive interactions between the host and its parasite. A-D. Eleutheroschizon duboscqi. A. Attached zoite transforming into a trophozoite stage,
already completely enveloped by a PS. B. Maturing trophozoite with a forming ring of fascicles at the attachment site. The tail forms at the caudal part of the PS.
C. Mature trophozoite with a prominent tail. Note the presence of attachment fascicles and lobes. D. Detailed view of the annular joint point (the cut-out is
marked by a red square in C). E-H. Eugregarines. E. Sporozoite immediately after attachment to the host epithelial cell. F. Transformation of the sporozoite
into a trophozoite stage. G. Early trophozoite with a well-developed epimerite. H. Detailed view of the membrane fusion site (the cut-out is marked by red
E. duboscqi Interconnecting Gregarines and Cryptosporidia
PLOS ONE | DOI:10.1371/journal.pone.0125063 April 27, 2015 18 / 27
Attachment strategy and host-parasite interactions in E. duboscqi
compared to other epicellular apicomplexans
According to our analysis, E. duboscqi develops within the host-derived PS, resembling cryptosporidia
[5,6,18,19,20]. Moreover, the parasite attaches to the host cell with the help of the complicated
attachment site, by analogy with an invading gregarine [6,13,15,21–24] (Fig 10A–10L).
Gregarines are mostly intestinal epicellular parasites, equipped with a specialised attachment
apparatus that might also serve as a feeding organelle (epimerite, mucron, modified protomerite)
[13,15,21,22,25–29]. Archigregarines suck out the host cell cytoplasm using
organelles of the apical complex [3,30,31]. Such a mechanism of feeding is called myzocytosis.
On the contrary, many eugregarines seem to not feed through myzocytosis, except, probably,
at their youngest developmental stages [32]. Their apical complex of organelles is reduced, and
a new, more complicated, attachment apparatus forms (Fig 10G). This apparatus does not penetrate
the host cell, but simply causes the invagination of the host plasmalemma (Fig 10E–10H)
[6,15,22]. Cryptosporidia have a typical feeder organelle that attaches to the host cell and remains
separated from the host cytoplasm by a dense line (Fig 10K and 10L) [5,6]. Similar to
gregarines and cryptosporidia, E. duboscqi has a complicated attachment apparatus (fascicles
and lobes in circles) (Fig 10C); and it remains unclear whether this apparatus is involved in the
parasite feeding. Numerous pores are distributed along the entire pellicle of E. duboscqi, including
the attachment site and some of them seem to be connected with vesicles and mitochondria.
These pores may participate in parasite feeding. The apical complex of organelles is
absent in E. duboscqi during the endogenous phase of its life cycle; and, apparently, feeding
through myzocytosis is not typical for this parasite. Neither organelle similar to the flaskshaped
structure described in the early stages of the eugregarines [6,13,22,23,32] nor the
mucronal vacuole, characteristic of the archigregarine Selenidium [3,30], was detected in freshly
attached E. duboscqi individuals. The flask-shaped structure observed in gregarines with an
opening towards the apical pole (Fig 10E) initially appears electron-dense, but, with the formation
of the cortical vesicle, it turns electron-lucent [6], suggesting that it might be a rhoptry
emptying its enzymes. As rhoptry proteins are generally expected to be involved in the transformation
of the host cell membrane into a parasitophorous vacuole (PV) [33], we cannot exclude
the possibility that a similar structure may be present in E. duboscqi at a stage younger
than the early trophozoite already enveloped by a PS.
Epicellular gregarines are usually not surrounded by any sac of host cell origin (Fig 10E–
10H), except for the archigregarine Ditrypanocystis sp., which is enveloped by a multimembranous
structure, originated from fused cilia of the host cell [24]. Cryptosporidia are enveloped
square in G). The two cytomembranes end at the point of membrane fusion, where the osmiophilic ring is formed. I-L. Cryptosporidia. I. Attached zoite
transforming into a trophozoite stage, partially enveloped by an incomplete PS. J. Young trophozoite almost completely enveloped by a PS. Note the tunnel
connection between the interior of the anterior vacuole and the host cell cytoplasm that developed as the result of the Y-shaped membrane junction. K. Mature
stage with a prominent filamentous projection at the base of the PS and with a fully developed feeder organelle, the lamellae of which formed from the anterior
vacuole membrane. L. Detailed view of the Y-shaped membrane junction (the cut-out is marked by a red square in K). M-P. Epicellular eimeriids. M. Invading
zoite. N. Trophozoite/meront stage enveloped by a PS with a single attachment area (monopodial form). O. Extension of the gamont stage above the
microvillous region leading to an establishment of a new contact with the host cell apart from the primary attachment zone by penetration of the PS membrane
to the base of fused microvilli (spider-like form). P. Detailed view of the attachment area (the cut-out is marked by a red square in O). av—anterior vacuole, b—
epimeritic bud, cm—parasite cytomembranes, cv—epimeritic cortical vesicle, db—dense band (in cryptosporidia usually consisting of several layers), f—
membrane fusion site, dl—dense line separating the feeder organelle from the filamentous projection of the PS, fa—attachment fascicle of filaments, fo—
feeder organelle with membranous lamellae, fom—membrane limiting the lamellae of feeder organelle, fp—filamentous projection of the PS, fs—flask-shaped
structure, hc—host cell, hm—host cell plasma membrane, if—incomplete fusion of PS, int—interface between the host cell and eugregarine epimerite,
consisting of host cell plasma membrane, epimerite plasma membrane and a dense layer in between, ipm—inner membrane of the PS, is—internal space
between the parasite and PS, j—annular joint point (Y-shaped membrane junction in cryptosporidia), lo—attachment lobe, ms—membrane-like structure
limiting the cortical vesicle from the epimerite cytoplasm, opm—outer membrane of the PS, p—pore on the PS, pm—parasite plasma membrane, ps—
parasitophorous sac, r—rhoptries, t—tail of the PS, tu—tunnel connection.
doi:10.1371/journal.pone.0125063.g010
E. duboscqi Interconnecting Gregarines and Cryptosporidia
PLOS ONE | DOI:10.1371/journal.pone.0125063 April 27, 2015 19 / 27
by the PS, the inner membrane of which came from the plasmalemma presented on the host
cell microvilli (Fig 10I–10L). In gregarines, as well as in cryptosporidia, the membrane fusion
site is formed at the contact area between the parasite and host cell. In contrast, there is no connection
between the host and E. duboscqi plasma membranes at the annular joint point. The internal
space between the E. duboscqi plasmalemma and the inner membrane of the PS does not
seem to be a part of the parasite. It rather resembles the space between the PV and intracellular
coccidia located inside the host cell. As all early stages of E. duboscqi were seen to be completely
contained within a PS, this host cell-derived envelope must develop much more rapidly than
that documented in cryptosporidia, in which, during the invasion process, a tight-fitting membrane
fold of the invaded host cell gradually rises up along the zoite, resulting in the formation
of the PS [5,20]. The presence of a crystalloid body, a typical feature of sporozoites [26,34], in
the stages attached and completely enveloped by the PS, confirms that the earliest observed
stages were only slightly modified sporozoites, after their attachment to the host cell. Interestingly,
the cortical vesicle in eugregarine Didymophyes gigantea was interpreted as a periparasitic
space between the host and parasite, functioning as a PV [35]. At first glance, the gregarine
cortical vesicle indeed resembles the internal space of the PV due to its translucent appearance
with traces of an opaque or filamentous material [6,13,22,23]. In Gregarina garnhami, fine tubular
structures pass through the cortical vesicle and attach to the epimerite-host cell interface
[21]. We could speculate that the cortical vesicle is in fact an incomplete PV, restricted to the
embedded apical region (epimerite) of the gregarine. It most likely develops from fused flat vesicles
distributed in the epimerite periphery, originating from the parasite endoplasmic reticulum,
and turns into a single large vesicle filled with microfilaments [23]. This vesicle is limited
on its cytoplasmic face by a membranous structure, often discontinuous or multi-layered
[6,15,21]. It retracts along with the epimerite during detachment of mature trophozoite from
the host cell [15].
Epicellular localisation within the host tissue has also been described for certain eimeriid
coccidia from fish (some Eimeria or former Epieimeria, some Goussia) [36–41] and reptiles
(Choleoeimeria, Acroeimeria) [42,43]. According to multiple studies, they are localised at the
enterocyte apical site among microvilli and covered by a double membrane envelope (the
enterocyte and PV membranes), but a single PV membrane in contact areas (Fig 10M–10P).
The arrangement of the PV membrane in contact zones is species specific based on various
projections and undulations. No direct contact between the parasite and parasitised cell was
observed [37,38,40,42–44]. During intracellular development of Eimeria anguillae merozoites,
the PV with a parasite inside is expelled into the apical region of the parasitised cell, hereby taking
an epicellular position [37]. In Goussia pannonica and G. janae, the parasites either attach
to the host cell at a single contact area resulting from multiple fusions of microvilli (‘monopodial’
form) or are located above the microvillar zone, connected with the epithelium (occasionally
to more than one enterocyte) through multiple thin projections in a spider-like
arrangement [40,44]. The PV membrane of the spider-like projections have been described to
be closely apposed to the enterocyte plasmalemma and to penetrate to the base of the villus
where it contacts the host cytoplasm. The referenced micrographs, though not provided in satisfactory
magnification, however, do not show any PV membrane penetrating into the host cytoplasm,
which would indicate an intracellular (epicytoplasmic) position of the parasite. It
rather suggests a deep invagination of the enterocyte membrane, at the connection with PV
projections, i.e. epicellular localisation. Representatives of Acroeimeria are also considered to
be epicellular parasites developing within a PV bulging above the epithelium surface [43].
From there, despite intracellular or epicellular initial stages of the development, some eimeriid
coccidia of poikilothermic animals localise at the host cell apical part and are surrounded by
the host-derived two-membrane PV.
E. duboscqi Interconnecting Gregarines and Cryptosporidia
PLOS ONE | DOI:10.1371/journal.pone.0125063 April 27, 2015 20 / 27
Cryptosporidia, E. duboscqi and gregarines represent heteropolar cells; i.e. they exhibit a
high degree of cell polarity in that their anterior and posterior ends differ in shape, structure
and function. Extracellular but attached parasites are usually of a heteropolar nature, while the
intracellular ones are generally non-polar. Epicellular eimeriids seem to be non-polar as they
do not have any attachment organelles and seem to take up nutrients exclusively via the PV
[37]. Nevertheless, they create projections of the PV [37,38] equipped with pores; these projections
enlarge the contact area with the host cell and resemble the E. duboscqi attachment lobes
and fascicles. Similarly, the complicated attachment organelles in many eugregarines seem to
significantly increase the absorptive surface [13,29,35]. Apparently, the epicellular localisation,
independent of its origin (whether initial stages of the parasite development are extracellular or
intracellular), leads to the occurrence of cell polarity. All the above-mentioned apicomplexans
form a specialised host-parasite interface, reflecting analogous modes of adaptation for survival
and development in a similar host environment (i.e. the gastrointestinal epithelial brush border)
[45] (Fig 10A–10P). It seems that, once attached to a proper host cell, all these parasites
stimulate additional growth and subsequent fusion of host microvilli as well as further modifications
to the host plasmalemma, leading to PS/PV formation. They tend to create a host-derived
envelope around themselves and develop in its cavity so as to be separated from the host
cytoplasm/environment. In cryptosporidia, the PS is not complete as they are directly connected
to the host cell via a feeder organelle (i.e. they form the so-called Y-shaped junction between
the host and parasite membranes), while in eimeriids and E. duboscqi, the inner
membrane of this envelope contains the entire parasite. The mode of connection with the host
cell remains inconclusive in the earliest developmental stages of E. duboscqi. Most likely it
never invades the host cytoplasm, but attaches to the apical site of the host cell for a short time
(until the formation of the PS).
It has been suggested that eimeriid epicellular development might be considered as a more
primitive form of host-parasite association [42]. Some studies showed, however, that parasites
developing epicellularly seem to have a less negative effect on the host epithelium than intracellular
ones [46]. In general, it is more advantageous for the parasite to maintain its host in acceptable
fitness; from this perspective, the evolution of this attachment strategy could be more
progressive. While the most serious pathological changes in E. anguillae are induced by intracellular
stages and vary depending on the intensity of parasitisation, the epicellular development
causes alterations to the host epithelium, such as local swellings or a reduction in the number of
microvilli [37]. Similar changes are reported in cryptosporidiosis, where the destiny of the host
depends on its health status and immunocompetence [47,48]. In E. anguillae, the host epithelium,
discharged from parasites, has lesions on the intestinal surface resembling circular holes
[37]. Cryptosporidia and E. duboscqi leave only shallow craters with PS remains at the epithelial
surface after their detachment [5]. Similarly, detached gregarine trophozoites leave flat holes in
the epithelium lacking microvilli [15]. Microvilli are essential for digestion and nutrient absorption
and their destruction might lead to host malnutrition with consequent weakening or even
death. Although epicellular parasites destroy individual cells, the overall damage to epithelium
in mild infections is negligible and often easily repaired thanks to its continual regeneration.
Host actin distribution and architecture of E. duboscqi parasitophorous
sac in comparison to cryptosporidia
Phalloidin staining, along with the application of drugs that influence the polymerisation of
actin, confirmed the presence of actin filaments in host tissue and their increased accumulation
in the PS surrounding E. duboscqi. Moreover, the higher doses of cytochalasin D, required for
destroying actin filaments in the PS wall, suggest that the polymerised form of actin is more
E. duboscqi Interconnecting Gregarines and Cryptosporidia
PLOS ONE | DOI:10.1371/journal.pone.0125063 April 27, 2015 21 / 27
stable in the host-derived PS than in surrounding host tissue. This accumulation of host actin
filaments might be induced by invading E. duboscqi, similarly to cryptosporidia, which induce
the rearrangement and accumulation of actin and actin-binding proteins to their attachment
site during invasion [49,50–52]. Cryptosporidia, however, show a low amount of F-actin in
their PS [53]. Furthermore, the wall of the E. duboscqi PS exhibited a high accumulation of myosin.
In cryptosporidia, the motor activity of host myosin seems to play an important role, putatively
in association with microvilli extensions, in the formation of a parasite niche [49,52].
This theory is supported by a model for the protrusion of membranous structures, illustrating
the significant involvement of myosin in the movement of actin filaments toward the apical
surface of membrane extensions [54]. The involvement of host microcilia in the formation of
the E. duboscqi PS is indicated by the presence of α-tubulin restricted to the PS wall, especially
in the caudal region with the tail. The tubulin seemed to be in polymerised form, as incubation
in oryzalin resulted in the vanishing of fluorescence as well as frequent detachment of parasites
with their sacs from epithelium observed in vitro.
In contrast to intracellular coccidia, evolutionary selection presumably favoured the unique
epicellular niche for cryptosporidia to more effectively evade the host immune response,
though as a consequence, the attached parasite became dependent upon its connection with
the host cell for nutrient acquisition. In E. duboscqi, the strategy could be similar. Host actin
polymerisation and subsequent membrane protrusion are considered to be important for the
establishment of a productive infection site in cryptosporidia [52], in which induced membrane
extensions encapsulate the parasite and form the PS, with a dense band in the host cytoplasm
located just beneath the attachment zone [5,6,20]. This band consists of electron-dense
microfibrils interwoven perpendicularly [55], with an adjacent filamentous network of polymerised
actin [52]. The dense band underlining the base of E. duboscqi is much thinner and
closely apposed to the PS inner membrane. CLSM confirmed an increased accumulation of Factin
at the attachment site. In cryptosporidia, the actin reorganisation and formation of dense
bands supported by the actin plaque were shown to be intimately involved in parasite anchoring
and retention at the host cell apical surface [5,49,56]. An explanation for such a cytoskeletal
rearrangement in parasitised cells is that this process results in the formation of a network for
vesicle trafficking, facilitating the movement of nutrients between the host cell and the PS [49].
This hypothesis is in agreement with our observations on the accumulation of tiny filaments located
within host cytoplasm surrounding the invaginations of PS membrane in the area of
E. duboscqi attachment fascicles. These fascicles develop during E. duboscqi trophozoite maturation
and seem to anchor the growing PS with the parasite to the host cell, while host filaments
overlapping them could strengthen this fixation and prevent mechanical detachment.
Both parasites, cryptosporidia and E. duboscqi, regularly detach (most likely due to damage)
along with their sacs from the unmodified part of the host cell. While the detachment of cryptosporidia
takes place in the area of a dense band [5], individuals of E. duboscqi tear away from
their sacs at the base, thereby exposing their naked basal region (i.e. covered by the parasite pellicle
only) and leaving the intact inner membrane of the PS at the place of previous attachment.
It seems that E. duboscqi induces only moderate alterations of the host cell in contrast to cryptosporidia,
in which, along with actin remodelling, remarkable alteration of host membrane organisation
has been documented [57]. Cytoskeletal remodelling of the host cell induced by
cryptosporidia can be noted as microvillous hypertrophy, i.e. elongation and protrusion of host
cell microvilli surrounding the parasite [49]. The persistence of long microvilli clustered at the
attachment site suggests an active manipulation of the host membrane structure by the parasite.
The microvilli associated with the cryptosporidian PS were particularly thick and contained
dense bundles of F-actin. In E. duboscqi, we did not observe any significant extension of
the adjacent host cell microvilli; only a few microcilia were occasionally attached to the PS
E. duboscqi Interconnecting Gregarines and Cryptosporidia
PLOS ONE | DOI:10.1371/journal.pone.0125063 April 27, 2015 22 / 27
surface (Fig 4A and 4K) and no obvious pathological changes were seen in the parasitised area.
Despite a similar strategy of PS formation and architecture with cryptosporidia, epicellular
stages of E. duboscqi seem to have less of a pathological effect on host tissue.
The function of the tail of the E. duboscqi PS remains unknown. Staining with Evans blue
showed that the tail is protein-rich, confirmed by the SEM observations of the presence of a fibrous
substance at the caudal region of parasites with a ruptured PS. Numerous pores along
with a dense matrix, intensively stained by RR, in their vicinity indicate the role of the tail in
the transportation of mucus and/or other parasite metabolites outside. Despite the presence of
long filaments, the F-actin labelling in the tail turned out to be less distinct than the rest of the
PS, but surprisingly, the incubation with oryzalin resulted in much stronger F-actin labelling in
the tail in contrast to the proximal part of the PS. Tails of different sizes in maturing trophozoites
may be a result of extra growth of the PS. Spectrin accumulation seemed to increase towards
the PS caudal area with a tail, suggesting that plasma membrane proteins are
accumulated in this zone. So, hypothetically, the fibrous substance could be a stock of membrane
material needed for the PS development during the parasite maturation and growth. It
remains unclear as to whether the ruptured PS observed in our preparations is a consequence
of damage due to material processing or it represents a natural process in the life cycle allowing
E. duboscqi gamonts to leave the host cell. It could be also a result of incomplete fusion of the
PS during early parasite development.
Cell cortex and cytoskeleton of E. duboscqi
In the course of transformation of E. duboscqi trophozoites into gamonts, a thick glycocalyx
layer appears on the external surface of the parasite plasmalemma. Similar fibrillar coat, most
likely of glycocalyx nature, has been documented covering the entire parasite in Acroeimeria
pintoi [43]. In E. duboscqi, it forms earlier than the subpellicular filaments and seems to be essential
for the attached stages. The most important role of glycocalyx might be a mechanical
defence against potential fusion of the PS with the parasite surface. Importantly, the short and
long attachment filaments that arise from the pellicle are anchored in the IMC and extend
through the glycocalyx. They evidently represent a modified form of the glycocalyx.
The pellicle of E. duboscqi also appears unique in that it seems to re-build or reorganise during
the parasite development. Vesicles observed under the plasma membrane in the area of discontinuous
or absent cytomembranes of some gamonts indicate a process of membrane
insertion into the cortex, most likely needed for pellicle completion during parasite growth.
The barely distinguishable pellicle of early trophozoites provides further support for the hypothesis
of its repetitive reorganisation. It is not clear how the parasite undergoes fertilisation
and if its life cycle comprises additional free stages, but in vitro we often observed detachment
of E. duboscqi along with PS, and documented detached parasites enveloped by the PS under
electron microscope. All this suggests the participation of the PS in parasite protection from
the surrounding environment even after detachment from the host tissue, probably as a consequence
of pellicle reorganisation during E. duboscqi development.
The cytoskeleton of this parasite comprises subpellicular microtubules, but only during
early development. While the posterior ring of Toxoplasma gondii does not connect with subpellicular
microtubules [58], in E. duboscqi it does, similar to the ‘posterior polar ring’ in tachyzoites
of Besnoitia besnoiti [59] or the so-called ‘proximal polar ring’ in Plasmodium
sporozoites [60]. Usually, these structures are only recognised as pellicular (IMC) thickening.
Although the microtubules disappeared during E. duboscqi trophozoite maturation and were
not present in gamonts, we obtained positive α-tubulin labelling of the parasite surface and cytoplasm,
suggesting the preservation of tubulin either in its non-polymerised form or its
E. duboscqi Interconnecting Gregarines and Cryptosporidia
PLOS ONE | DOI:10.1371/journal.pone.0125063 April 27, 2015 23 / 27
presence in other tubulin-rich structures. The second speculation seems to be more likely, because
incubation in oryzalin resulted in the disappearance of labelling from the parasite periphery,
and the putatively unpolymerised α-tubulin seemed to be more dispersed throughout the
cytoplasm. Apicomplexan microtubules are selectively susceptible to drug-induced disruption;
after prolonged treatment in 2.5 μM oryzalin all tubulin is usually unpolymerised and dispersed
[61]. Oryzalin prevents the formation of microtubules in T. gondii daughter cells, but
has a more moderate effect on existing microtubules in the mother cell [62,63]. This is in agreement
with our results on E. duboscqi, where high drug doses must be applied for prolonged period
to observe a dispersed character of tubulin staining. Of special interest are also the
subpellicular bands of longitudinally oriented filaments, located beneath the parasite IMC, that
form during the trophozoite maturation. Positive labelling of the parasite surface for actin/Factin
and the effect of cytoskeletal drugs on its staining suggest that these filaments are actinrich.
Considering the lack of subpellicular microtubules in mature stages, these bands could
play the role of the parasite cytoskeleton.
Conclusions
The endogenous stages of Eleutheroschizon duboscqi life cycle exclusively comprise trophozoites
and gamonts. They develop in the intestine of Scoloplos armiger being attached to the host
cell in an epicellular position and covered by a host-derived parasitophorous sac. Attached parasites
share features of cryptosporidia and gregarines, i.e. they conspicuously resemble a maturing
trophozoite of epicellular eugregarines with morphologically pronounced attachment
apparatus, but are contained within a PS similar to that in cryptosporidia. However, E. duboscqi
parasites have no intimate contact with the enterocyte membrane. The parasite pellicle seems
to reorganise repeatedly during development. Detached parasites are enveloped by a PS covering
their distal area above the attachment site, hereby suggesting that after detachment from
the host tissue they preserve this envelope of host origin, providing them ongoing protection in
a potentially unfriendly surrounding environment.
Acknowledgments
Authors would like to thank the staff of the White Sea Biological Station of Lomonosov Moscow
State University (WSBS MSU) for providing them with facilities for field sampling and
material processing, as well as their kind and friendly approach. GP is also grateful to the staff
of the Marine Biological Station of Saint-Petersburg State University at the White Sea (MBS
SPbSU) for providing facilities for staying and sampling. The authors are greatly indebted to
the members of the Laboratory of Electron Microscopy (Institute of Parasitology, BC ASCR in
České Budějovice) for technical assistance and to Dr. Peter Stuart (Masaryk University) for English
editing of the manuscript. Many thanks to Prof. Dominique Soldati-Favre (University of
Geneva) for kindly providing the monoclonal anti-actin antibody. GP deeply thanks Dr. Prof.
Rudolf Entzeroth who invited her to the Technical University of Dresden (Michail Lomonosov
DAAD stipendium, 2006) and organised all the things needed for the preliminary investigation
of Eleutheroschizon duboscqi, and Markus Günther (Technical University of Dresden) who assisted
her with electron microscopy.
Author Contributions
Conceived and designed the experiments: AV. Performed the experiments: AV. Analyzed the
data: AV GGP AD MK TGS. Contributed reagents/materials/analysis tools: AV GGP AD MK
TGS. Wrote the paper: AV. Discovered Eleutheroschizon duboscqi in Scoloplos armiger from
the White Sea and shared her findings with other authors for further collaboration, and
E. duboscqi Interconnecting Gregarines and Cryptosporidia
PLOS ONE | DOI:10.1371/journal.pone.0125063 April 27, 2015 24 / 27
contributed considerably to manuscript writing: GGP. Performed field sampling: AV GGP AD
TGS. Contributed to the study design: GGP AD TGS. Performed light and electron microscopic
analyses: AV GGP AD MK TGS. Performed CLSM analyses: AV. Assisted in sample processing
for CLSM: MK. Reviewed and edited the manuscript, and approved its final version: AV
GGP AD MK TGS.
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E. duboscqi Interconnecting Gregarines and Cryptosporidia
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7.5
Melicherová J., Mazourová V., Valigurová A.
2016
In vitro excystation of Cryptosporidium muris oocysts and viability
of released sporozoites in different incubation media
Parasitology Research 115(3), 1113-1121
ORIGINAL PAPER
In vitro excystation of Cryptosporidium muris oocysts and viability
of released sporozoites in different incubation media
Janka Melicherová1
& Veronika Mazourová1
& Andrea Valigurová1
Received: 2 September 2015 /Accepted: 19 November 2015 /Published online: 18 December 2015
# Springer-Verlag Berlin Heidelberg 2015
Abstract This study aimed to evaluate and document the
excystation process of Cryptosporidium muris oocysts in various
incubation media, and to monitor the behaviour of
excysting and freshly excysted sporozoites. A test of oocyst
viability, using fluorescent double staining with fluorescein
diacetate and propidium iodide, was performed prior to each
experimental assay. Light microscope observations confirmed
that relatively often only three sporozoites were released; the
fourth one either left the oocyst later together with a residual
body or remained trapped within the oocyst wall. These results
suggest that successful oocyst excystation is not limited
by the viability of all four sporozoites. Darkening of oocysts to
opaque and their specific movement (the so-called Boocyst
dancing^) preceded the final excystation and liberation of
sporozoites, while the dormant oocysts appeared refractive.
The process of excystation in C. muris is not gradual as generally
described in cryptosporidia but very rapid in an eruptive
manner. Experiments were performed using oocysts stored at
4 °C for various time periods, as well as oocysts freshly shed
from host rodents (Mastomys coucha) of different ages. The
most suitable medium supporting high excystation rate (76 %)
and prolonged motility of sporozoites was RPMI 1640,
enriched with 5 % bovine serum albumin (BSA). Our results
emphasize that to reliably evaluate the success of in vitro
excystation of cryptosporidia, not only the number of released
sporozoites in a set time period should be taken into consideration
but also their subsequent activity (motility), as it is
expected to be essential for the invasion of host cells.
Keywords Cryptosporidium muris . Excystation rate .
Motility . Oocyst . Sporozoite . Viability test
Abbreviations
BSA bovine serum albumin
FDA fluorescein diacetate
LM light microscopy
PBS phosphate-buffered saline
PI propidium iodide
Introduction
The genus Cryptosporidium, belonging to the phylum
Apicomplexa, was previously placed to the subclass
Coccidiasina, Leuckart, 1879. Based on more recent phylogenetic
and comparative morphologic analyses, Cryptosporidium
spp. are considered to be close relatives of gregarines (Barta and
Thompson 2006; Carreno et al. 1999; Kváč et al. 2008;
Valigurová et al. 2007, 2008). Cryptosporidia differ significantly
from representatives of order Eucoccidiorida due to (i) the
putative lack of sporocysts; (ii) the presence of microgamonts
producing 16 non-flagellated, bullet-shaped microgametes; and
finally (iii) their unique epicellular localization within a hostderived
parasitophorous sac (Current and Reese 1986;
Valigurová et al. 2008). The host specificity of cryptosporidia
varies according to species and genotype (Feng et al. 2011; Kar
et al. 2011).
Two main branches are recognized within gastrointestinal
cryptosporidia, i.e. species/genotypes infecting either the intestine
or the stomach (Pavlasek and Ryan 2007). The process
of excystation initiates when oocysts reach the gastrointestinal
tract. In a proper host environment, oocysts excyst and release
four infectious sporozoites, capable of actively invading the
* Janka Melicherová
jankam@sci.muni.cz
1
Department of Botany and Zoology, Faculty of Science, Masaryk
University, Kotlářská 2, 611 37 Brno, Czech Republic
Parasitol Res (2016) 115:1113–1121
DOI 10.1007/s00436-015-4841-0
host epithelial cells. Excystation of oocysts under in vitro conditions
has been studied predominately with intestinal species,
where various protocols have been tested (Kato et al. 2001;
Robertson et al. 1993; Smith et al. 2005). Whereas excystation
of cryptosporidian intestinal species can be stimulated with
acidic components or bile salts, gastric species seem to require
raised temperature only (Gold et al. 2001; Widmer et al. 2007).
Hence, different protocols for in vitro experiments must be
applied for successful excystation of Cryptosporidium muris
oocysts resulting in the release of viable sporozoites.
In this study, we focused on the gastric parasite C. muris
strain TS03. Although C. muris is specific for rodent hosts, it
can be occasionally found in other mammals (FitzGerald et al.
2011; Kodádková et al. 2010), including humans (Gatei et al.
2002; Palmer et al. 2003). The main aim of this study was to
compare the viability of C. muris oocysts collected from host
rodents at different time points of the patent period and stored
for different periods of time. Moreover, a detailed investigation
of the entire excystation process could reveal the behaviour
of freshly excysted sporozoites prior to the active invasion
of appropriate host cells. This study underlines that the
success rate of in vitro excystation should consider not only
the number of released sporozoites in a set time period but also
the activity (motility) of freshly excysted sporozoites, which is
essential for the invasion process of host cells.
Materials and methods
C. muris
Faeces from multimammate rats, Mastomys coucha, experimentally
inoculated with C. muris (strain TS03) oocysts (dose
of 1×106
oocysts suspended in 200 μl of distilled water),
were collected daily and stored in an aqueous solution of
2.5 % potassium dichromate (K2Cr2O7) at 4 °C. The TS03
strain of C. muris was characterized in detail by Kváč et al.
(2008), with descriptions of differences in its genetic characteristics,
infectivity and pathogenicity from other isolates of
gastric Cryptosporidium spp. Collected oocysts were purified
using the Sheather’s sugar flotation method (Arrowood and
Sterling 1987) and a modified caesium chloride gradient centrifugation
(Kilani and Sekla 1987). The purified oocyst suspension
was stored in 0.1 M phosphate-buffered saline (PBS),
pH 7.2, at 4 °C.
Animals were housed in plastic cages (five animals per
cage) with sterile wood chip bedding and supplied with sterilized
food and water ad libitum. The rearing of animals is
regulated by Czech legislation (Act No 246/1992 Coll., on
protection of animals against cruelty). These documents are
consistent with the legislation of the European Commission.
All housing, feeding and experimental procedures were conducted
under protocols approved by University of Veterinary
and Pharmaceutical Sciences Brno and Central Commission
for Animal Welfare, Czech Republic (protocol # 066/2010).
The minimum number of animals has been involved to produce
statistically reproducible results.
Oocyst viability test
Double staining with fluorescein diacetate (FDA) and
propidium iodide (PI) was performed to verify the viability
of oocysts prior to all experimental assays. Some samples
were stained only with PI. Since PI, a red fluorescent nuclear
and chromosome counterstain is not live cell permeable, it is
commonly used to detect the dead cells in a population, using
an excitation maximum at 535 nm and fluorescence emission
maximum at 617 nm (Jones and Senft 1985). The principle of
the FDA test is the ability of living cells to metabolize the
esterase substrate to a derivative that can be subsequently
detected or quantified by measuring the fluorescence, using
an excitation maximum at 450–490 nm, and fluorescence
emission maximum at 520 nm. Non-fluorescent FDA is a
non-polar compound that easily penetrates the plasma membrane
of viable cells, where it converts into a fluorescent metabolite
(fluorescein). Fluorescein cannot penetrate cell membranes
and can be detected by a fluorescence microscope at an
appropriate pH, when accumulated in some concentrations.
Hence, a green fluorescent signal as well as no or a weak
fluorescence signal (due to a low concentration of accumulated
metabolite) indicates a viable oocyst with living sporozoites.
Stock solutions were prepared by dissolving FDA in acetone
(5 mg/1 ml) and PI in deionized water (1 mg/1 ml).
Oocysts were suspended in 300 μl of RPMI in an Eppendorf
microtube, and subsequently 100 μl of FDA (4 μl/ml in PBS,
pH 7.2) and 30 μl of PI (1 mg/ml in distilled water) working
solutions were added. Specimens were incubated for 10 min in
the dark at room temperature (RT). Microscopic slides with
stained oocyst suspensions were viewed with a fluorescence
microscope, Olympus BX60.
In vitro excystation of oocysts
In order to investigate the effect of various factors on the
excystation rate, in vitro incubations of oocysts in various
media were performed and the process of oocyst excystation
was evaluated. Simultaneously, the effect of a specific medium
on the motility rate of sporozoites was monitored. Oocysts
of C. muris were not chemically pre-treated (e.g. with sodium
hypochlorite routinely used for oocysts disinfection) to avoid
any alternations of the oocyst wall, potentially causing the
death of enclosed sporozoites. To obtain the maximum possible
excystation rate, various excystation media were applied,
including distilled water with 0.6 % HCl (0.1 N HCl, SigmaAldrich,
Czech Republic), 0.1 M phosphate-buffered saline
(PBS) enriched by 10 % trypsin or 5 % bovine serum albumin
1114 Parasitol Res (2016) 115:1113–1121
(BSA) (Sigma-Aldrich, Czech Republic), Ringer’s solution
(containing 154.00 mM NaCl, 5.64 mM KCl, 2.16 mM
CaCl2, 11.10 mM dextrose and 2.38 mM NaHCO3 in 1 L
H2 O) and finally RPMI 1640 (Sigma-Aldrich,
Czech Republic) with 5 % BSA or without BSA. Volumes
of 4×106
C. muris oocysts were diluted in 1000 μl of a selected
medium and divided into 200-μl aliquots in Eppendorf
microtubes to achieve a final concentration of 8×105
oocysts.
The incubation temperature was set at 37 °C.
The second set of experimental assays was performed
using the most effective medium, RPMI 1640, enriched with
BSA at the following concentrations: 1, 5, 10, 20 and 50 %.
Oocyst excystation counts were performed in a Bürker chamber
(with at least 100 oocysts counted) where ten squares were
used for counting released sporozoites and excysted and
unexcysted oocysts. Excystation percentage was calculated
as [number of excysted oocysts/ a total number of counted
oocyst]×100. Excystation rate was monitored at set time intervals
of 5, 15, 30, 60, 90, 120, 180 and 240 min. Statistical
evaluation of the excystation rate in all tested media was performed
using the non-parametric methods of the Kruskal–
Wallis test. Furthermore, motility of released sporozoites
was recorded. The result of excystation was evaluated and
documented (imaging, video) using a motorized light microscope
Olympus BX61 at phase contrast, equipped with a digital
camera Olympus DP71 and the software Olympus Stream
Motion v. 1.5.1.
Results
Excystation of oocysts in various media
Oocysts used in the first set of experiments were collected
from the faeces of M. coucha at the beginning of the last part
of the chronic infection period (8 months post-inoculation).
These relatively old oocysts (4 months old) were incubated for
1–3 hours at 37 °C in different incubation media (mammalian
Ringer’s solution, PBS with 10 % trypsin, PBS with 5 % BSA,
PBS with 0.6 % HCl and finally RPMI 1640 with or without
5 % BSA), to check their viability after prolonged storage
(data not shown). The highest ratio (50 %) of excystation, a
process where viable sporozoites are released from the oocyst,
was observed in RPMI 1640 containing 5 % BSA, where the
motility of excysted sporozoites was the most evident. A pure
RPMI 1640 without any supplements also increased the rate
of oocyst excystation, but the difference in motility of sporozoites
was negligible when compared to a medium enriched
with BSA. Excystation of these oocysts in Ringer’s solution
reached 45 % after 2 hours at 37 °C, but the motility of freshly
excysted sporozoites was very low. Reduction of medium pH,
achieved by adding 0.6 % HCl, was shown not to significantly
increase the excystation rate. PBS that contained 10 % trypsin
or 5 % BSA supported the motility of sporozoites, but its
positive effect on the excystation process was not confirmed;
i.e. less than 30 % oocysts excysted successfully. As the obtained
results seemed to correlate with the length of oocyst
storage (at 4 °C), and most likely also with the age of the host
rodent, further experiments with younger oocysts were designed
to test the influence of various concentrations of BSA
diluted in RPMI 1640 on the excystation rate and sporozoite
motility.
Viability of C. muris oocysts
Viability of C. muris oocysts stored for different time periods
was compared to freshly collected oocysts, 2-week- and 4month-old
oocysts stored in PBS at 4 °C. The percentage of
viable oocysts was determined for each age group individually
by subtracting the percentage of dead oocysts from the percentage
of total oocysts. The viability of oocysts was evaluated
using double PI/FDA staining or simple PI staining for
fluorescence analysis. If the oocysts with sporozoites stained
red, they were considered non-viable because PI had penetrated
through their damaged membrane regions. FDA causes live
cells to fluoresce green under blue light excitation. The viability
of oocysts correlated negatively with the length of their
storage after collection from host faeces. The highest percentage
of viability (99 %) was documented in fresh oocyst samples
(Fig. 1a, b), whereas after 2 weeks, the viability of oocysts
decreased to 95 %. The oocysts were often stained with both
dyes; i.e. a more or less intense PI staining of sporozoites was
observed in an oocyst emitting green fluorescence (Fig. 1c).
The 4-month-old oocysts showed 20 % viability (Fig. 1d).
The viability of oocysts also depended on the phase of the
host patent period. At the end of the patent period, M. coucha
excreted C. muris oocysts that were about 50 % less viable
than oocysts excreted at the beginning of the patent period.
The fluorescent test of oocyst viability was used also prior to
each experimental assay.
Excystation rate and sporozoite motility in various
concentrations of BSA
After primary evaluation of the viability of C. muris oocysts
stored for different time periods, an experimental group of 2week-old
oocysts from a rodent host of reproductive age was
chosen as the most appropriate material for further research.
Evaluation of the oocyst excystation ratio and the motility of
released sporozoites was performed in various concentrations
of BSA (1, 5, 10 and 50 %) diluted in RPMI 1640, as well as
in RPMI 1640 without BSA (Table 1). In view of the fact that
the condition of dispersion was not achieved in our data (tested
by statistical Bartlett’s test), statistical comparison of the
excystation rate in different media at selected time periods was
tested using the non-parametric Kruskal–Wallis test (Fig. 3).
Parasitol Res (2016) 115:1113–1121 1115
Detailed observations on oocyst excystation revealed that a
typical darkening of oocysts predicted that activated sporozoites
are ready to be released from the oocyst (Fig. 2a). A
vibrating movement of these dark and opaque oocysts, resembling
a dance (oocyst dancing), was observed preceding the
excystation and indicated the activation of oocysts. Repeated
observations showed that the final release of sporozoites from
the oocyst is not gradual, but unexpectedly very rapid and
ejection-like. The dormant oocysts appeared refractive when
observed in the medium by light microscopy.
Excystation with the highest concentration of BSA (50 %)
was affected negatively and seemed to block the oocyst suture
opening, as the oocysts remained refractive with light microscopy
and did not excyst till the end of experiment (Fig. 3
(yellow line) and Fig. 2d). The process of excystation was
successfully initiated in pure RPMI 1640 as well as RPMI
1640 with 1–10 % BSA, almost at the same rate. The first
obvious differences in the excystation rate appeared after the
first 15 min of oocyst incubation (p<0.001, for RPMI 1640
with 0–5, 1–5 and 5–10 % BSA). In the course of the entire
experiment, numerous Bdancing oocysts^ were observed. The
significant peak period of excystation was noticed after
30 min (p < 0.001, for RPMI 1640 with 0–5, 1–5 and 5–
10 % BSA), when an obvious variation in the excystation rate
occurred in individual media. The highest excystation rate was
observed in a medium with 5 % BSA. The second peak period
of excystation was documented after 1 hours (p<0.018, for
RPMI 1640 with 0–5 % BSA), when the percentage of
excystation reached over 50 % in all tested media (except
media with 50 % BSA). The third peak period of excystation
occurred after 3 hours (p<0.001, for RPMI 1640 with 0–5 %
and 0–10 % BSA), when the percentage of excystation in
Fig. 1 Viability test of C. muris
oocysts using PI or double PI/
FDA staining. a, b Fresh oocysts
stained with PI. Non-viable, redstained
oocysts are shown by
arrows. Fluorescence combined
with transmission LM (a) and
fluorescence (b). c Fluorescent
visualization of 2-week-old
oocysts double stained with PI/
FDA. Note the viable, greenstained
oocyst and the green
oocysts with red spots (white
arrowheads). d Fluorescent
detection of dead oocysts and
oocyst with dying sporozoites
(white arrowhead) from a 4month-old
sample. Stained with
PI/FDA. Inset shows the released
dead sporozoites and two
sporozoites still attached to the
residual body (white arrowhead).
Stained with PI (Colour figure
online)
Table 1 The percentage of oocyst excystation in RPMI with or without BSA
Percentage of excystation
0 min 5 min 15 min 30 min 60 min 90 min 120 min 180 min 240 min
RPMI 0 18,9 ± 4.2 25 ± 1.7 28,7 ± 2.7 48,5 ± 0.8 56,1 ± 0.9 60,5 ± 0.7 65 ± 0.9 67 ± 1.8
1 % BSA 0 23,3 ± 2.2 27 ± 0.9 32,5 ± 1.7 50,6 ± 0.6 57,2 ± 0.5 61,3 ± 0.5 67,2 ± 1.4 68,4 ± 0.9
5 % BSA 0 29,6 ± 1.6 46,5 ± 1.5 50 ± 1.4 54,5 ± 1.0 59,7 ± 1.7 68,4 ± 0.3 72,9 ± 2.2 76 ± 1.8
10 % BSA 0 25 ± 1.1 32 ± 0.8 36,7 ± 0.9 51,7 ± 1.7 58,5 ± 1.8 63,5 ± 0.7 68,9 ± 1.6 74 ± 2.5
50 % BSA 0 0 0 0 0 0 0 0 0
Mean % ± SD, n = 3
BSA bovine serum albumin
1116 Parasitol Res (2016) 115:1113–1121
media without BSA or with 1 % of BSA reached almost maximum.
The blue and black lines (RPMI 1640 without BSA and
RPMI 1640 with 1 % BSA) in the graph started to be constant
or only increased slightly, while the red and green lines demonstrating
the excystation in media with 5 and 10 % BSA
continued to gradually increase (Fig. 3). Furthermore, in pure
medium and 1 % BSA, more dormant oocysts were observed.
All experimental assays were stopped after 4 hours (p<0.001,
for RPMI 1640 with 0–5 % and 1–5 % BSA) because of a
natural decrease in the viability of excysted sporozoites, correlating
with the length of their in vitro incubation.
Although the percentage of excystation in a pure RPMI
1640 medium was comparable with the percentage of
excystation in this medium supplemented with BSA, the motility
of excysted sporozoites was significantly lower in the
former one. Focusing exclusively on the excystation process,
the best results (76 % excystation at 37 °C) were obtained in
RPMI 1640 enriched by 5 % BSA (Fig. 3, red line). In this
medium, the activity of sporozoites was obviously higher and
their motility did not significantly change (decrease) with time
(the majority were motile till the end of the experiment), in
contrast to those incubated in medium without BSA. In the
Fig. 2 Excystation of C. muris
oocysts in different incubation
media. Phase contrast LM. a
RPMI 1640 medium. b RPMI
1640 enriched by 5 % BSA. c
RPMI 1640 with 10 % BSA. Note
the contact of sporozoites with
their apical ends (black asterisk).
d RPMI 1640 with 50 % BSA.
Darkened oocysts ready for
excystation (black arrowhead),
released viable sporozoites (black
arrows), non-viable sporozoites
(white arrows) and sporozoites
remaining inside the oocyst
(black double arrowheads),
unexcysted oocysts (white
arrowheads) and empty oocysts
(white asterisk) (Colour figure
online)
0
10
20
30
40
50
60
70
80
RPMI 1% BSA 5%BSA 10% BSA 50% BSA
*
0-5%
1-5%
5-10%
*
0-5%
1-5%
5-10%
*
0-5%
1-5%
*
0-5%
*
0-1%
*
0-5%
1-5%
*
0-5%
0-10%
*
0-5%
1-5%
Fig. 3 Curves showing the
percentage of oocysts’ excystation,
depending on tested medium and
time of incubation. Black line
represents the pure RPMI medium,
blue line the RPMI medium
enriched with 1 % BSA, red line
the medium with 5 % BSA and
green line the medium with 10 %
BSA. Statistical Kruskal–Wallis
test assessed significant differences
for each time period of incubation
between tested media. The p value
was smaller than 0.01 and
statistically significant differences
between all tested media are
marked with asterisk (colour figure
online)
Parasitol Res (2016) 115:1113–1121 1117
RPMI 1640 medium with 10 % BSA, the excystation ratio
achieved almost 74 % (Fig. 3, green line). The motility of
released sporozoites was comparable with their motility in
the medium with 5 % BSA. Nevertheless, in the medium with
10 % BSA, the sporozoites seemed to be active for a shorter
time, i.e. their motility decreased more rapidly, and by the end
of experiment, more of them appeared less active.
The elongated, slender sporozoites exhibited an oscillating
movement (short forwards/backwards shifts) appearing to
move progressively forward and without obvious changes in
cell shape (Fig. 2c). This movement was observed, at least to
some degree, in all tested media (except for 50 % BSA), but
the activity of excysted sporozoites decreased with lower concentrations
of BSA. In the medium enriched with 1 % BSA,
oocysts reached an excystation rate of 68.5 % after 4 hours at
37 °C (Fig. 3, blue line). The lowest motility of sporozoites
was recorded in the medium without BSA, where a 67 %
excystation ratio was observed (Fig. 3, black line). These sporozoites
were most active during the first 30 min after
excystation, but thereafter, motility rapidly declined.
Discussion
Viability of C. muris oocysts
Viability testing of C. muris oocysts using fluorescent double
PI/FDA staining is generally considered standard procedure
prior to experimental work (Jones and Senft 1985). However,
it is important to mention that Boyd et al. (2008) investigated
the membrane integrity and cell viability using the FDA/PI
staining and claimed that the staining of viability is dependent
upon a number of other factors (the effect of FDA diluent or
the time elapsed from the staining procedure). Likewise,
Smith and Smith (1989) observed that some cysts could not
be stained with either FDA or PI. While some studies claim
that the viability of Cryptosporidium parvum oocysts can be
maintained in appropriate conditions for 6 months (Fayer et al.
1998a, b), others recommend working with oocysts younger
than 3 months (Kar et al. 2011). Likewise, Jenkins et al.
(2003) recorded a 50 % decrease in excystation success after
3 months of oocyst storage, while 9-month-old oocysts were
non-viable. Furthermore, other studies concerning the viability
of cryptosporidian oocysts found them to be affected by
biotic and abiotic factors, such as age of oocysts, temperature,
purification procedure, storage period or chemical composition
of media (Fayer et al. 1998a, b; Jenkins et al. 2003;
Reduker et al. 1985; Reinoso et al. 2008; Robertson et al.
1992). Based on our empirical knowledge, prolonged storage
is not acceptable for successful in vitro experiments with C.
muris (strain TS03). Oocysts of C. muris used in our studies
preserve their infectivity only for a short period of storage; i.e.
after 4 months, the viability of oocysts stored in PBS
decreased to 20 %. Of special interest is a study claiming that
it is almost impossible to reduce the viability of all oocysts in a
sample to zero (Robertson et al. 1992). This hypothesis was
supported by an experiment showing that even putatively nonviable
oocysts of C. parvum (not excysted during in vitro
excystation experiments) could infect host laboratory models
(Neumann et al. 2000). In a study reporting the lowest infective
dose of nine oocysts in total for C. parvum for humans
(Okhuysen et al. 1999), a possible explanation could be that
the viability test is not sensitive enough to detect sporadic
viable oocysts. On the other hand, our in vitro experiments
also provided support for such a controversial hypothesis, as
despite almost negative viability tests (PI/FDA), an
excystation rate of up to 50 % was achieved in 4-month-old
oocysts incubated in RPMI 1640 with 5 % BSA.
The influence of incubation medium on excystation rate
In the current study, the excystation process of C. muris oocysts
collected from faeces of M. coucha was evaluated in
various excystation media. The majority of published studies
focus on the intestinal pathogen C. parvum, which is more
widespread. Our research deals with a gastric parasite, originally
obtained from Tachyoryctes splendens and described as
C. muris by Kváč et al. (2008), and subsequently maintained
in laboratory rodent hosts. During its monoxenous life cycle,
the oocysts of C. muris are ingested into the stomach, a suture
present on the oocyst surface opens provided the environmental
chemistry and physical factors are favourable. For study
purposes, the entire process of C. muris oocyst excystation,
normally taking place in vivo within the host stomach, was
performed in vitro in a series of experiments. These ran under
modified conditions in order to examine the possible effect of
individual media components on the excystation process. The
composition of these media as well as the incubation times
(usually from 15 min up to 4 hours) varied in individual experimental
sets. Basic physiological solution and a saline solution,
though applicable in some experimental studies and also
tested in this study, do not represent media suitable for longterm
in vitro cultivation. Based on personal long-term experience,
RPMI 1640 was chosen as a proper medium for our
study because of its wide applicability for in vitro cultivation
of various cell cultures and unicellular parasites. This medium
was first supplemented with trypsin and HCl because these
have been recorded as having a reversible stimulatory effect
on sporozoite motility (Smith et al. 2005; Widmer et al. 2007).
However, no similar impact was observed in C. muris during
this study. Hence, BSA was used to enrich the excystation
medium because of its known positive effect on the motility
of some organisms (Galvani et al. 2007; Harrison et al. 1982).
Although the excystation rate was not significantly affected
by BSA supplementation, the motility of sporozoites was lower
in pure RPMI 1640, and that enriched by 1 % BSA, when
1118 Parasitol Res (2016) 115:1113–1121
compared to the media with higher concentrations of BSA (5
% and 10 %). The most suitable medium for further studies is
the one supplemented with 5 % BSA, in which sporozoites
remained motile during the entire experiment (the activity of
excysted sporozoites slightly decreased in 10 % BSA).
Excystation in medium with 50 % BSA failed. We assume
that a medium with such a high concentration of BSA represents
a very dense and viscous environment that could seal the
oocyst, thereby preventing it from excysting.
Previous studies indicate that cryptosporidian oocysts generally
only require the temperature to reach 37 °C for successful
excystation, but the excystation rate can accelerate as pH
decreases, e.g. by adding HCl or NaOCl (Fayer and Leek
1984; Forney et al. 1996; Smith et al. 2005; Gold et al.
2001; Reduker et al. 1985) . Pre-treatment of oocysts with
bleaching agents seems unnecessary (Arrowood 2002;
Robertson et al. 1993), but various chemical factors such as
trypsin, taurocholic acid or hypochloric acid can be added to
the incubation medium as stimuli for excystation. These acidic
chemical substances are not essential for gastric cryptosporidia
excystation, but they slightly accelerate the excystation process
(Widmer et al. 2007).
Our data showing that the age of C. muris oocysts (i.e. length
of their storage) and the age of the host organism represent
further important factors are inconsistent with a study of
Widmer et al. (2007), in which no significant influence of these
factors was recorded. The patent period of M. coucha progresses
continuously into a chronic infection and results in the death of
the host (usually about 1–1.5 years after experimental inoculation)
or, in rare cases, the infection with C. muris spontaneously
disappears (personal unpublished data). The amount of oocysts
in faeces decreases with increasing age of the host. Hence, the
best period for parasite collection from host faeces seems to be
up to 4 months from the beginning of the patent period.
Behavioural activity of sporozoites during the excystation
process
Surprisingly, our observations showed that the release of sporozoites
from oocysts is an extremely rapid process, resembling
the shooting out of sporozoites through the suture in the
oocyst wall. It is known that the most significant excystation
factors, appropriate pH and temperature, modify permeability
of the oocyst wall (Matsubayashi et al. 2011; Jenkins et al.
2003; Robertson et al. 1993). Hence, we expect that some
factors might increase the pressure inside the oocyst by absorbing
liquids through the permeable oocyst wall (most likely
through the suture) into the enlarging residual body.
Subsequently, the hypothetical excessive pressure within an
oocyst results in shot-like releasing of sporozoites to the surrounding
environment.
Other chemicals usually added to the incubation media can
also influence the behaviour of oocysts or sporozoites.
Although trypsin or BSA do not increase the excystation success
rate, they can cause translucency in oocysts and increase
the motility of excysted, as well as still unexcysted sporozoites
(Smith et al. 2005; Galvani et al. 2007; Harrison et al. 1982).
The motility of unexcysted sporozoites seems to be a source of
dancing oocysts, observed at the start of the excystation process
in this study. Enlarging the load of amylopectin in the
residual body just before excystation can serve as an energy
source for sporozoite activation (Fayer et al. 1998b; Harris et
al. 2004). Similar to gregarines (Steele et al. 2012), the activation
of C. muris sporozoites was obvious due to the darkening
of individual sporozoites within the enclosed oocyst
until both became opaque. Furthermore, our data concur with
that of Kar et al. (2011), where usually only three sporozoites
were released from oocysts while the fourth one remained
within the oocyst walls, eventually being liberated along with
the residual body. This could be a strategy to preserve one
viable sporozoite by protecting it within the oocyst wall in
case of potential failure of those already excysted. However,
it is more likely that the remaining sporozoite is already dead.
This corresponds to the red spots observed in some green
oocysts stained with FDA/PI. If the second variant applies,
the dying of sporozoites within the oocyst must be gradual,
and the oocyst excystation is induced by the other three, still
viable sporozoites.
It still remains unknown whether the percentage of successfully
excysted oocysts correlates with the infectivity of released
sporozoites. Previous studies, as well as results from
experiments using BSA herein, indicate that excysted sporozoites
require binding of specific lectins to their surface to
become invasive.
Conclusions
This study demonstrated that at the beginning of excystation
assays, the excystation process was only slightly accelerated
by adding the suitable concentration (5–10 %) of BSA to the
incubation medium, while the motility of sporozoites increased
significantly. Hence, although not for oocyst
excystation as such, the choice of an appropriate excystation
medium is undoubtedly essential for activation and prolonged
motility of excysted sporozoites, as well as for further successful
in vitro cultivations. Data presented herein also revealed
that requirements for successful oocyst excystation in gastric
species C. muris differ from those in intestinal species
C. parvum.
Acknowledgments The authors are greatly indebted to Monika
Nechvílová from the Department of Pathological Morphology and Parasitology,
University of Veterinary and Pharmaceutical Sciences in Brno,
for her kind help with laboratory animals. Many thanks go to Prof. Joseph
Schrével from Muséum national d'Histoire naturelle (MNHN) in Paris for
his invaluable advice on experiments and RNDr. Danka Haruštiaková,
Parasitol Res (2016) 115:1113–1121 1119
Ph.D. (Institute of Biostatistics and Analyses, Faculty of Medicine and
Faculty of Science, Masaryk University in Brno) for her help with statistical
evaluation. Great thanks are to Dr. Peter Daniel Stuart for English
correction of this text.
Compliance with ethical standards
Financial support Financial support was provided by the project
GAP505/11/1163 from the Czech Science Foundation. Travel expenses
were partially covered by the Mobility project from Ministry of
Education, Youth and Sports (7AMB14FR013). Authors acknowledge
support from the Department of Botany and Zoology, Faculty of
Science, Masaryk University, towards the preparation of this manuscript.
Conflict of interest The authors declare that they have no conflict of
interest.
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Parasitol Res (2016) 115:1113–1121 1121
7.6
Diakin A., Paskerova G.G., Simdyanov T.G., Aleoshin V.V., Valigurová A.
2016
Morphology and molecular phylogeny of coelomic gregarines
(Apicomplexa) with different types of motility: Urospora ovalis
and U. travisiae from the polychaete Travisia forbesii
Protist 167(3), 279-301
Protist, Vol. 167, 279–301, June 2016
http://www.elsevier.de/protis
Published online date 17 May 2016
ORIGINAL PAPER
Morphology and Molecular Phylogeny of
Coelomic Gregarines (Apicomplexa) with
Different Types of Motility: Urospora
ovalis and U. travisiae from the
Polychaete Travisia forbesii
Andrei Diakina,1
, Gita G. Paskerovab,1
, Timur G. Simdyanovc
,
Vladimir V. Aleoshind
, and Andrea Valigurováa
aDepartment of Botany and Zoology, Faculty of Science, Masaryk University, Kotláˇrská 2,
611 37, Brno, Czech Republic
bDepartment of Invertebrate Zoology, Faculty of Biology, St. Petersburg State University,
Universitetskaya emb. 7/9, Saint-Petersburg, 199 034, Russian Federation
cDepartment of Invertebrate Zoology, Faculty of Biology, Lomonosov Moscow State
University, Leninskie Gory, Moscow 119 234, Russian Federation
dBelozersky Institute for Physico-Chemical Biology, Lomonosov Moscow State University,
Leninskie Gory, Moscow 119 234, Russian Federation
Submitted November 18, 2015; Accepted May 5, 2016
Monitoring Editor: C. Graham Clark
Urosporids (Apicomplexa: Urosporidae) are eugregarines that parasitise marine invertebrates, such as
annelids, molluscs, nemerteans and echinoderms, inhabiting their coelom and intestine. Urosporids
exhibit considerable morphological plasticity, which correlates with their different modes of motility
and variations in structure of their cortical zone, according to the localisation within the host.
The gregarines Urospora ovalis and U. travisiae from the marine polychaete Travisia forbesii were
investigated with an emphasis on their general morphology and phylogenetic position. Solitary ovoid
trophozoites and syzygies of U. ovalis were located free in the host coelom and showed metabolic
activity, a non-progressive movement with periodic changes of the cell shape. Solitary trophozoites of
U. travisiae, attached to the host tissue or free floating in the coelom, were V-shaped. Detached trophozoites
demonstrated gliding motility, a progressive movement without observable cell body changes.
In both gregarines, the cortex formed numerous epicytic folds, but superfolds appeared exclusively on
1
Corresponding authors;
e-mail diakin@sci.muni.cz (A. Diakin), gitapasker@yahoo.com (G.G. Paskerova).
http://dx.doi.org/10.1016/j.protis.2016.05.001
1434-4610/© 2016 Elsevier GmbH. All rights reserved.
280 A. Diakin et al.
the surface of U. ovalis during metabolic activity. SSU rDNA sequences obtained from U. ovalis and U.
travisiae revealed that they belong to the Lecudinoidea clade; however, they are not affiliated with other
coelomic urosporids (Pterospora spp. and Lithocystis spp.), but surprisingly with intestinal lecudinids
(Difficilina spp.) parasitising nemerteans.
© 2016 Elsevier GmbH. All rights reserved.
Key words: Urosporidae; marine eugregarines; ultrastructure; gliding and metaboly; superfolds; 18S rDNA
phylogeny.
Introduction
Apicomplexa Levine 1980, emend. Adl et al. 2012
(Adl et al. 2012) consist entirely of unicellular
organisms parasitising various animals. Some of
them cause important human and domestic animal
diseases (e.g. malaria, toxoplasmosis, and
cryptosporidiosis); therefore, these species have
been intensively studied in different aspects of
biology, medicine and phylogeny. However, basal
apicomplexans (e.g. gregarines, agamococcidia,
blastogregarines, and protococcidia), inhabiting
exclusively invertebrate hosts, are crucial for our
understanding of the evolutionary pathways of Apicomplexa,
yet they still remain poorly investigated.
Gregarines parasitise a broad range of terrestrial
and aquatic invertebrates (annelids, turbellarians,
arthropods, echinoderms, and urochordates), and
inhabit different sites within the host organism, e.g.
the gut and its derivatives (the Malpighian tubules,
respiratory trees), the body cavity, and the reproductive
system.
According to their morphological features, life
cycles, and host range, gregarines are usually
subdivided into three groups: Archigregarinorida
Grassé, 1953, Eugregarinorida Léger 1900, and
Neogregarinorida Grassé, 1953 (Adl et al. 2012;
Desportes and Schrével 2013; Grassé 1953;
Perkins et al. 2000). In contrast to coccidia, developmental
stages of gregarines are predominantly
extracellular and of large dimensions. The feeding
stages (trophozoites) of gregarines are usually
motile and heteropolar, with opposite ends differing
in their structure and function. Usually gregarines
undergo their vegetative phase of development
when attached to the host tissue; the majority
of them lack the form of asexual reproduction
called merogony (=schizogony). Another important
characteristic of the gregarine life cycle is the presence
of a pre-sexual association, the so-called
syzygy, usually consisting of two partners. There
are several types of syzygies: caudo-frontal (headto-tail),
frontal (head-to-head), caudal (tail-to-tail),
and lateral (partners are in contact at their lateral
surfaces). In the majority of eugregarines and archigregarines,
the partners in the syzygies retain the
motility characteristic of solitary trophozoites. In the
course of time, the partners become hemispherical
in shape, form a common envelope (a gametocyst
wall), and undergo gametogenesis, followed
by sporogenesis (Desportes and Schrével 2013;
Grassé 1953; Perkins et al. 2000).
Gregarines show a great variety of cell shapes
and different modes of motility that seem to correlate
with trophozoite localisation within the host.
Gregarines from the intestine are generally vermiform
(archigregarines) and demonstrate pendular
(rolling) motility, or elongated (most eugregarines)
and show gliding motility. Parasites from the body
cavity, tissues, or the reproductive system are
usually oval or roundish (some of the urosporids
and monocystids), or dendritic (Pterospora spp.).
As a rule, such gregarines possess metabolic or
peristaltic motility (Nematopsis magna, Lithocystis
schneideri, Urospora neapolitana); some of them
are non-motile (Gonospora varia, Lythocystis foliaceae,
Urospora chiridotae) (Coulon and Jangoux
1987; Desportes and Schrével 2013; Dyakin and
Paskerova 2004; Dyakin and Simdyanov 2005;
Frolov 1991; Landers and Gunderson 1986; Levine
1977; MacMillan 1973; Miles 1968; Perkins et al.
2000; Schrével 1964, 1971a, b, and others). While
gliding is a progressive movement, both pendular
(rolling) and metaboly are non-progressive. In
addition, metaboly is accompanied with periodic
changes of the cell body shape.
The exact mechanism of gregarine motility
remains unknown. Gliding motility seems to be
facilitated by the complex organisation of the parasite’s
cortical zone. The pellicle forms longitudinal
epicytic folds with special sets of filamentous structures
in their apex (the so-called rippled dense
structures [RDS and 12-nm apical filaments) and
an internal lamina, which underlays the inner membrane
complex (IMC) (Schrével et al. 1983; Vivier
1968). The polymerised form of actin and cytoplasmic
mucus, excreted outside the cell, both actively
participate in gregarine gliding (Valigurová et al.
Morphology and Molecular Phylogeny of Coelomic Gregarines 281
2013). Peristaltic or metabolic motility is accompanied
by the forming of one or several contracted
regions running from one end to the other along
the longitudinal axis of the cell. It was assumed
that this type of motility can be facilitated by the
presence of a subpellicular cytoskeletal network;
however, its nature, whether it is composed of fibrils
or microtubules, remains unknown (MacMillan
1973; Warner 1968).
The family Urosporidae, established by L.
Léger (Léger 1892), combining the monocystid
gregarines with heteropolar oocysts, nowadays
comprises several genera of parasites inhabiting
various marine and freshwater invertebrates
(mainly echinoderms and polychaetes, as well as
oligochaetes, sipunculids, molluscs, nemerteans)
(Desportes and Schrével 2013; Dogiel 1906, 1909,
1910; Grassé 1953, Levine 1977; Perkins et al.
2000; Pixell-Goodrich 1915, 1950). To date, most
investigations concerning urosporids are either
faunistic studies or morphological descriptions of
various developmental stages (mostly trophozoites
and oocysts), performed at light microscopic level.
Only a few members have been studied from an
ultrastructural viewpoint (Corbel et al. 1979; Dyakin
and Simdyanov 2005; Landers and Leander 2005;
Pomory and Lares 1998), and even fewer have
been investigated at the molecular biological level
(Leander et al. 2006).
The type genus Urospora Schneider, 1895 unites
monocystid gregarines from the body cavity or
tissues of hosts, with lateral or frontal syzygies,
and with heteropolar oocysts possessing a thin
appendage at one end and a conical transparent
funnel at the other (Desportes and Schrével
2013; Grassé 1953; Levine 1977). In the present
study, we investigated the morphology and molecular
phylogeny of gregarines of two closely related
urosporid species Urospora ovalis Dogiel, 1910 and
U. travisiae Dogiel, 1910, parasites of the body cavity
of the marine polychaete Travisia forbesii Johnston,
1840, noting, in addition, the biodiversity and
adaptations of gregarines from coelomic habitats.
Results
All dissected hosts (approx. 400 individuals) were
infected with Urospora ovalis and Urospora travisiae
(Fig. 1A). The parasites inhabited the host
body cavity. In addition, spherical gametocysts with
typical urosporid oocysts were found. The ovalshaped
oocysts were heteropolar, with a funnel and
a tail at opposite ends, and about 20 ␮m (n = 40) in
length, 7 ␮m (n = 40) in width (Fig. 1A, inset).
The intensity of parasitisation by both gregarine
species varied during the summer season. In the
case of U. ovalis, it reached up to 50 parasites
per host in June/July, while no more than 5 in
August/early September. In the case of U. travisiae,
it also reached up to 50 parasites per host (in rare
cases, no more than 5) during the entire summer
season. For U. ovalis, both solitary trophozoites
(June/August) and syzygies (August/early September)
were found, while for U. travisiae, mostly
solitary trophozoites were observed, and syzygies
were found in a few cases only.
The main species characteristics of U. ovalis and
U. travisiae are summarised in Table 1.
General Morphology and
Ultrastructure of Urospora ovalis
The solitary trophozoites of U. ovalis, occurring
freely in the host body cavity, were ovoid with
rounded ends and showed no signs of heteropolarity
under the light microscope (LM) (Fig. 1A-D).
The cell size varied widely: 19.6 - 294.0 ␮m (av.
179 ␮m, mode 252 ␮m, n = 36) in length and 12.6
- 187.6 ␮m (av. 114 ␮m, mode 134 ␮m, n = 36) in
width. We did not observe any young stages of U.
ovalis.
Some parasites were glued to the host peritoneal
epithelium by means of a mucous substance surrounding
them. (Fig. 1B-C). Living parasites which
had fallen out of the host during dissection showed
characteristic metabolic (peristaltic) activity, during
which cells alternately contracted at their ends
causing the migration of the cytoplasm from one
end to the other (Fig. 1D, Supplementary Material
Video 1). Several superficial, longitudinal ridges
formed in the contracted regions during gregarine
movement. These ridges were visible even in histological
sections (Fig. 1B-C).
Under SEM, these ridges corresponded to the
so-called superfolds that run along the surface
of the contracted region (Fig. 2A, C). However,
during processing for electron microscopy (EM),
solitary motile cells of U. ovalis completely contracted
in most cases, and superfolds identical to
those observed in contracted regions appeared
on the entire gregarine surface (Fig. 2B, E). In
transversal sections, both narrow (0.4-0.5 ␮m wide
at the base) and wide (2.5-3.5 ␮m wide at the
base) superfolds of uniform height (1 ␮m on average)
were present at the surface of contracted cells
(Fig. 3A-C).
The cells of U. ovalis were covered with a typical
three-layered pellicle consisting of the plasma
282 A. Diakin et al.
Figure 1. Localisation of Urospora ovalis and U. travisiae in the polychaete Travisia forbesii and light microscopic
observations of U. ovalis A. Sagittal section of the host infected with U. ovalis and U. travisiae (white
arrowheads) located in the body cavity (bc). The black rectangles mark the parasites presented in Figure 1B (U.
ovalis) and Figure 5A (U. travisiae) at high magnification. Ant – anterior end of the host, D – dorsal side of the
host; il – intestinal lumen of the host, ph – pharynx; oo – oocytes of the host, Post – posterior end of the host, V
– ventral side of the host. LM, H&E. The inset shows an oocyst with a funnel (f) and a tail (t) at opposite ends.
LM. B – C. Histological sections of trophozoites of U. ovalis located in the host body cavity (bc), in different
planes. Note mucous substance (ms) surrounded parasites and ridges (black arrows) at the parasite surface.
oo – oocytes of the host. LM, H&E. D. Micrograph of the solitary U. ovalis trophozoite during cell metaboly. Note
contracted region (black arrowheads) of the cell with ridges at the surface (black arrows). n – nucleus. PC. E.
Syzygy of U. ovalis. Note ridges (black arrows) at free ends of gamonts. vac – vacuoles. BF.
membrane and the inner membrane complex
(IMC). The pellicle formed numerous thin and
waved epicytic folds (Fig. 2D). Their width was usually
about 90 nm, while their height cyclically varied
within the range from 0.5 to 1.4 ␮m. (Fig. 3B-C, E).
The number of epicytic folds in 1 ␮m of the surface
varied from 3 to 7, in dependence of the degree
of cell contraction. The rippled dense structures
and 12-nm apical filaments were not distinguishable;
however, there was a single electron-dense
rod located just beneath the IMC in the apical
part of each epicytic fold (Fig. 3D-E). The 19-
24 nm thick internal lamina underlay the IMC and
formed curved bridges in the basal part of each
fold, thereby separating the cytoplasm of folds from
the rest of the gregarine cytoplasm (Fig. 3D-E).
Under SEM the folds in non-contracted, as well
as contracted regions were undulating or waving
(Fig. 2D-E). Each superfold bore 10-20 epicytic
folds (Fig. 3B-C).
Numerous cortical microtubules were arranged
in transversal bundles located just beneath the pellicle.
They did not form a continuous ring in the cell
periphery, and some of the microtubules extended
into the superfolds (Fig. 3B-D).
Typical apicomplexan micropores, appearing
as short cylindrical invaginations of the plasma
membrane terminated by a vesicle (about 55 nm
in diameter), could occasionally be observed
(Fig. 3F). The cylindrical part was enforced by
an electron-dense collar (ca. 130 nm in diameter),
formed by the IMC and the internal lamina. In addition,
numerous structures resembling micropores
(micropore-like structures) were located in the gregarine
cortex between the epicytic folds (Fig. 3E,
G-H). In these structures, the IMC and internal
lamina formed an electron-dense cone-shaped collar
situated beneath the intact plasma membrane
(Fig. 3E, G). Numerous cytoplasmic vesicles (about
0.2 ␮m in diameter) filled with an electron-dense,
Morphology and Molecular Phylogeny of Coelomic Gregarines 283
Table 1. Main species characters of Urospora ovalis and U. travisiae.
Urospora ovalis Urospora travisiae
Host Travisia forbesii
Localisation in the host coelom
Attachment to the host tissues non-attached attached
Cell shape ovoid V-like
Number of cell axis 1 2
Cell dimension (average, ␮m) of mature
trophozoites
179 × 114 395 × 380 per
branch
Cell motility metaboly or
peristalsis
(non-progressive
movement)
gliding
(progressive
movement)
Cell polarity - +
Attachment site - +
Cortex
Epicytic folds typical for gliding eugregarines
Superfolds + Cytoplasm
differentiation into ectoplasm
and endoplasm
- +
Oocysts spindle-shaped, heteropolar, with a funnel and the
hairy-tail at opposite ends
Distribution Barents Sea (Murmansk coast), White Sea (Karelian coast)
References Dogiel 1910; present study
homogeneous material were usually situated in
the vicinity of micropore-like structures as well as
deeper within the parasite cytoplasm (Fig. 3B).
Some of them were in contact with the collar. Except
for a few cases, there was no fusion between these
vesicles and the plasma membrane (Fig. 3G-H).
The electron-dense droplets occasionally observed
between the epicytic folds could be secreted by
these vesicles (Fig. 3E).
There was no obvious division of cell cytoplasm
into two zones, ectoplasm and endoplasm, as is
typical for eugregarines. Many amylopectin granules
were irregularly distributed in the cytoplasm,
and several mitochondria could be observed at
the cell periphery (Fig. 3A-C). The dictyosomes of
Golgi apparatus, surrounded by numerous small
and round vesicles, were mainly localised in the
central zone of the cell (Fig. 3A).
The cytoplasm was packed with numerous and
different structures, exhibiting a lower quantity at
the cell periphery in comparison to the central cell
region (Fig. 3A). Electron-dense, homogeneous
inclusions of uncertain shape (di) accumulated predominantly
around the nucleus. Some of them were
in group of 2-3 and were usually accompanied
by small transparent vesicles (Fig. 3A, I). Another
type of inclusions was represented by large and
round electron-transparent vacuoles (ov) with a
loose filamentous content (Fig. 3A, J). In addition,
electron-dense, roundish vacuoles (dv) with a
heterogeneous granular content were found in the
cytoplasm (Fig. 3A, K).
The eccentrically located nucleus possessed one
large and several small nucleoli. The large nucleolus
lay close to the nuclear envelope; the rest
of the nucleoplasm was homogenous with a finegrained
content (Fig. 3A). This nucleolus consisted
of two parts: the larger one was dense, homogeneous
and directed towards the centre of the
nucleus, while the smaller one was heterogeneous
and faced the rough and two-layered nuclear envelope.
There were several inclusions of nucleoplasm
in both parts of the nucleolus (Fig. 3A).
Mature trophozoites (gamonts) in syzygies were
connected to each other by their ends. The free
ends of partners in syzygies were rounded (in rare
cases) or bulb-shaped (in most cases). In the first
case, syzygies showed active metabolic motility
comparable with that of solitary parasites. In the
second case, they moved much more slowly. In all
cases, both partners demonstrated ridges on their
surface in contracted regions (Fig. 1E), similar to
those observed in moving solitary trophozoites
(Fig. 1D). The gamonts in all of the observed
284 A. Diakin et al.
Figure 2. General morphology and surface ultrastructure of solitary Urospora ovalis trophozoites. A. General
view of a trophozoite fixed during its movement. Contracted (cr) and non-contracted (ncr) regions are visible
in the cell. White double arrowheads indicate superfolds. SEM. B. General view of a completely contracted
trophozoite with superfolds (white double arrowheads) that ran over the entire cell surface. SEM. C. Detailed
view of Figure 2A showing a transition between the contracted (cr) and non-contracted (ncr) regions of the cell.
White double arrowheads mark superfolds in the contracted region. SEM. D. Details of epicytic folds (white
arrow) covering the non-contracted region. Note electron-dense droplets (black arrow) located between the
epicytic folds. SEM. E. Details of superfolds (white double arrowheads) and epicytic folds (white arrow) of a
contracted cell. Black arrow points to an electron-dense droplet between epicytic folds. SEM.
syzygies were larger than solitary trophozoites,
and their cytoplasm was completely filled with
large transparent vacuoles (Fig. 1E).
During processing for EM, gamonts in syzygies
changed their shape by rounding their free ends
(Figs 1E vs. 4A). In contrast to solitary trophozoites,
fixed gamonts had no superfolds on their surface
(Figs 2A-C vs. 4A-C, Figs 3A-C vs. 4E). The height
of epicytic folds in syzygies cyclically varied within
the range from 0.7 to 2 ␮m, so that longitudinal
sets of high epicytic folds alternating with lower
ones were good visible on the syzygies surfaces
(Fig. 4A-E). The number of epicytic folds in 1 ␮m of
the surface varied from 3 to 5. There were droplets
of mucus between these folds (Fig. 4C). Neither
micropores, nor similar structures interrupting the
cortex were found in all examined ultrathin sections
(Fig. 4E).
The contact zone between two syzygy partners
appeared simple, lacking an additional collar or
other pellicle modifications (Fig. 4D). Usually, the
sets of high epicytic folds of both partners coincided
with each other (Fig. 4A, D). The free ends
of gamonts in syzygy exhibited almost identical
superficial morphology: they were slightly bulged-in
(Fig. 4F-G).
The cytoplasm of syzygy partners was filled with
a huge amount of electron- transparent vacuoles
with loose filamentous content (Fig. 4E), similar to
those found in solitary trophozoites (Fig. 3J, ov),
but extremely enlarged in volume. Among them,
there were many other inclusions such as lipid
droplets and amylopectin granules. Electron-dense
vacuoles and inclusions resembling the “di” and
“dv” found in solitary trophozoites (Fig. 3I, K) were
also observed (Fig. 4E).
Morphology and Molecular Phylogeny of Coelomic Gregarines 285
Figure 3. Fine structure of solitary Urospora ovalis trophozoites. A. Transversal section of a solitary trophozoite
contracted during fixation. The nucleus (n) with the nucleolus (nuc) inside is located eccentrically in
the cell. The cytoplasm is enriched by numerous and different inclusions: ag – amylopectin granules, di electron-dense
homogeneous inclusions of uncertain shape, dv – electron-dense vacuole with granular content,
286 A. Diakin et al.
General Morphology and
Ultrastructure of Urospora travisiae
Solitary trophozoites were found in the host body
cavity (Fig. 1A). Some gregarines were attached
to the intestine wall (Fig. 5A) and occasionally to
the blood vessels, but they easily detached during
host dissection or sample manipulation. The
young trophozoites and syzygies of U. travisiae
were observed rarely during this study (Fig. 5B-C).
Young trophozoites were elongated, and drop-like
in shape, with up to three transverse constrictions
at the tapering end. The length of them varied from
100 to 130 ␮m (n = 2) (Fig. 5B). The nucleus was
located in the widest part of the cell. Detached
young trophozoites exhibited a gliding motility, with
a wide, rounded leading end.
Mature trophozoites of U. travisiae were Vshaped.
They possessed two narrowing branches
and attached to the host tissue with the tip where
the branches converged, the so-called attachment
tip. In syzygies, the V-shaped partners were in contact
with each other by means of this attachment tip
(Fig. 5C); however, the contact was not strong, and
partners easily disassociated.
Each branch had 5-15 transverse permanent
constrictions; thus, branches appeared as a string
of pearls with differently sized beads (Fig. 5DG).
The angle between the branches in a cell
varied in the range from 10◦
to 180◦
, generally
from 90◦
to 130◦
(n = 25) (Figs 5D-G, 6A). Commonly,
one of the branches was longer than the
other: 195-660 ␮m (av. 392 ␮m; standard deviation
(SD) = 99.4; n = 25) vs. 165-580 ␮m (av. 352 ␮m;
SD = 96.8; n = 25). A single oval nucleus with 2-3
nucleoli was usually situated in the longest branch
close to the attachment tip (Fig. 5D-G).
Detached V-shaped trophozoites demonstrated
the typical gliding motility. Cells with an angle of
about 10◦
- 100◦
between their branches usually
moved with the attachment tip forward. The gliding
path was straight or curved, as if the cell branches
possessed equal or different motion forces, respectively
(Supplementary Material Video 2). In cases
where the angle was about 180◦
, cells glided with
one of the branches forward.
The attachment tip appeared like a plateau, usually
surrounded by a circular, wide furrow (Fig. 6B).
Some traces of the epicytic folds were visible on
the surface of the plateau, while the well-developed
longitudinal folds extended radially from the circular
furrow. Most of these aforementioned folds ran parallel
till the distal end of each branch (Fig. 6A-E).
On the lateral surface of the cell, the folds, passing
from opposite branches, converged and merged
with each other, while on the inner side of the Vshaped
cell, the epicytic folds passed continuously
from one branch to another (Fig. 6F). There were
no evident changes in the form or structure of these
folds in the region close to the attachment tip and at
the constrictions between individual beads (Fig. 6EG).
Some of the folds ended at constriction regions,
while others continued to the next beads. Additional
epicytic folds that passed only on the surface of the
beads also appeared (Fig. 6D-E).
The parasites were covered by a typical threelayered
pellicle consisting of plasma membrane
underlain by the closely adjacent membranes of
IMC. The cortex of U. travisiae did not exhibit
any secondary superfolds. Epicytic folds reached
0.5 ␮m in height and 0.1 ␮m in width, and were regularly
distributed with a distance of about of 0.1 ␮m
between them, 3-4 folds per 1 ␮m (Figs 6C, G,
7A-D). As in the epicytic folds of U. ovalis, the rippled
dense structures and 12-nm apical filaments
were indistinguishable in the apex of the epicytic
fold; however, a dense fibrillar rod was observed
just beneath the IMC. The pellicle was underlain
ov – electron-transparent vacuole with loose filamentous content. Ga – dictyosome of the Golgi apparatus. TEM.
B – C. Details of the narrow (B) and wide (C) superfolds bearing the epicytic folds (ef). Note a micropore-like
structure (black double arrowhead) and electron-dense vesicles (white arrowhead). ag – amylopectin granules,
mit – mitochondria, mt – microtubules. TEM. D. Details of the trophozoite cortex showing peripheral transversal
microtubules (mt). A single electron-dense rod (r) located just beneath the IMC is visible at the apex of one
completely visible epicytic fold. inl – internal lamina, ov – electron-transparent vacuole with loose filamentous
content. TEM. E. Transversal section of the gregarine cortex showing micropore-like structures (black double
arrowhead) accompanied by electron-dense vesicles (white arrowhead). Electron-dense rods (r) are visible in
the epicytic folds. Note electron-dense droplets (black arrow) between the epicytic folds. inl – internal lamina.
TEM. F. Details of a typical micropore (mp). TEM. G. Details of a micropore-like structure (black double arrowhead)
in contact with an electron-dense vesicle (white arrowhead). TEM. H. Details of an electron-dense vesicle
beneath the trophozoite cortex; note the membrane (white arrow) limiting the vesicle. TEM. I. Higher magnification
of a dense inclusion (di) of uncertain shape. TEM. J. Higher magnification of an electron-transparent
vacuole (ov) with loose filamentous material. TEM. K. Higher magnification of an electron-dense vacuole (dv)
with granular content. TEM.
Morphology and Molecular Phylogeny of Coelomic Gregarines 287
288 A. Diakin et al.
by an internal lamina, which did not form links at
the base of individual epicytic folds (Fig. 7A). Typical
micropores were situated between the folds
(Fig. 7A). Several micropore-like structures, similar
to those in U. ovalis, were seen between the epicytic
folds. They appeared as a cone-shaped collar,
formed by an IMC and internal lamina, and electrondense
vesicles were associated with them (Fig. 7B).
The same vesicles were also found deeper within
the parasite cytoplasm (Fig. 7D). Electron-dense
inclusions, oval in profile and apparently located
between the pellicle membranes, were seen at the
base or at the lateral sides of most folds (Fig. 7B,
D). Transversal subpellicular microtubules underlay
the bases of epicytic folds, as observed in U. ovalis
(Figs 6G, 7A).
In cross-sections of a branch near the attachment
tip, the cell was almost round, and the cytoplasm
was subdivided into an ectoplasm and endoplasm
(Fig. 7C-E). The endoplasm was packed
with electron-transparent inclusions of irregular
shape, vacuoles with homogeneous translucent
content, electron-dense vesicles, numerous lipid
droplets, and dictyosomes of the Golgi apparatus.
The ectoplasm mostly comprised mitochondria and
electron-dense vesicles (Fig. 7C-E).
Molecular Phylogenetic Analysis
The new SSU sequences of U. ovalis (1623 bp)
and U. travisiae (1604 bp) were obtained by direct
sequencing of PCR products. There are 3.5% differences
between them (46 substitutions and 3 indels)
across the distance of their pairwise alignment,
where they completely overlapped (1604 sites), and
distinctive nucleotides did not show any polymorphism
(Supplementary Material Fig. S1). Like other
lecudinid and urosporid SSU rDNA sequences, the
novel sequences were considerably shorter in comparison
with other eukaryotes (U. ovalis: 1604 bp
vs 1731 bp in Bigelowiella natans across the same
interval of the alignment; U. travisiae: 1623 bp vs
1742 bp in B. natans; the full length of SSU rDNA
of B. natans and the large majority of other eukaryotes
is about 1800 bp) and contained motifs specific
to other lecudinid and urosporid gregarines.
The constructed Bayesian (Fig. 8) and maximumlikelihood
(ML) trees of SSU rDNA of 99 OTUs
showed similar topology with two exceptions: (i)
the flipped positions of two eugregarine clades:
Gregarinoidea + Cephaloidophoroidea and Stylocephalids;
and (ii) the branching order of
archigregarines (data not shown). However, both
of these variable branching patterns were weakly
supported by both methods (Fig. 8).
The Bayesian tree of SSU rDNA sequences
(Fig. 8) fitted recent opinions on alveolate phylogeny:
the main robust clades are ciliates,
dinoflagellates and apicomplexans. The backbone
of the apicomplexans was weakly supported; nevertheless,
the sequences clustered into several
major well-supported clades: (1) coccidia and
hematozoa (not supported by BP in ML analyses);
(2) cryptosporidia; (3) Actinocephaloidea (CavalierSmith
2014) consisting of neogregarines and some
terrestrial eugregarines, e.g., Monocystis spp. and
representatives of family Actinocephalidae (not
supported by BP (91%); (4) Gregarinoidea (Clopton
2009); (5) Cephaloidophoroidea (Rueckert et al.
2011a); (6) the clade of Polyplicarium spp.
and related environmental sequences (not supported
by BP (74%)); and (7) Lecudinoidea (=
Urosporoidea in Cavalier-Smith 2014), a clade consisting
of the marine aseptate gregarine families
Lecudinidae and Urosporidae, and the unusual gregarine
Veloxidium leptosynaptae. The SSU rDNA
sequences from archigregarines did not form a
common clade. All gregarine and cryptosporidia
subclades formed a monophyletic clade – however,
with low supports (PP = 0.81, BP = 43%), and these
subclades were shuffled inside this common clade,
i.e. their branching order was unresolved because
of low support both in Bayesian and ML analyses.
The new SSU rDNA sequences of U. ovalis and
U. travisiae belonged to the Lecudinoidea clade
➛
Figure 4. General morphology and fine structure of Urospora ovalis syzygies. A. General view of a syzygy.
Black arrowheads mark sets of high epicytic folds. SEM. B. Detailed view of the surface showing the sets of
high epicytic folds (black arrowheads). SEM. C. Higher magnification of the gregarine surface with low and
high epicytic folds (ef). Note the droplets of mucous substance (black arrows) between folds. SEM. D. Detailed
view of the contact between two syzygy partners. Black arrowheads indicate sets of high epicytic folds at the
surface of both gamonts. SEM. E. Transversal section of a gamont showing the alternating sets of high (black
arrowheads) and low epicytic folds (white arrowheads). Inclusions of the cytoplasm are similar to that in the
cytoplasm of solitary trophozoites: ag – amylopectin granules, di – electron-dense homogeneous inclusions of
uncertain shape, dv – electron-dense vacuole with granular content, ld – lipid droplets, ov – electron-opaque
vacuole with loose filamentous material. TEM. F-G. Semi-axial view (F) and higher magnification (G) of the free
ends of gamonts. Note that the apex of the free ends is slightly bulged-in. SEM.
Morphology and Molecular Phylogeny of Coelomic Gregarines 289
Figure 5. Light microscopic observations of Urospora
travisiae gregarines. A. Histological section of the
trophozoite clamped between the folds of the host
intestine (in). Asterisk indicates the attachment tip, n –
nucleus, b – bead of the branch in the cell. LM, H&E. B.
Young trophozoite with a single tapering branch bearing
two transverse constriction near the end. Asterisk
marks the attachment tip, b – beads of the branch
in the cell. BF. C. Syzygy of two V-shaped partners
attached to each other by their attachment tips (asterisk).
b – beads of the branches. BF. D – G. Micrographs
of trophozoites with different angles between their
branches: D – approx. 140◦; E – 180◦; F – 70◦; G
– 10◦. Note the different number of the beads (b) in
their branches and the oval nucleus (n) with 2-3 nucleoli
(nu) situated in the longest branch close to the
attachment tip (asterisk) of each individual.
(Fig. 8). Within this clade, however, they closely
affiliated with lecudinids (Difficilina species), but not
with other urosporids (Pterospora spp. and Lithocystis
spp.).
Despite the fact that the new sequences were
fully supported by both the BI and ML analyses in
the 99 OTUs tree, topological tests were performed
using the resulting phylogeny of 23 selected lecudinid
and urosporid sequences (Fig. 9A-I, Table 2).
The topology of the main clades in this tree (Fig. 9A)
remained the same as the topology of those in
the 99 OTUs tree. Further, 8 trees with alternative
topologies were constructed (Fig. 9B-I), in
which the clade U. ovalis + U. travisiae sequentially
changed their position. We tested all of these
topologies using a set of the most common tests.
The majority of topologies were discarded with the
exception of three permissible ones: the Bayesian
tree, the alternative topology F (but not the BP test),
and the alternative topology I (Fig. 9A, F, I, Table 2).
All these trees contained the clade Difficilina spp. +
Urospora spp. in contrast to the discarded topologies,
where this combination was absent.
Discussion
The morphology of the gregarines investigated in
this study completely corresponds to the original
description of Urospora ovalis and U. travisiae
trophozoites presented by V. A. Dogiel (1910).
According to the original descriptions, oocysts of
U. ovalis and U. travisiae were heteropolar (with
a funnel and a tail at opposite ends), very similar,
but differ in size; oocysts of U. travisiae were
larger than those of U. ovalis (Dogiel 1910). All
oocysts observed in the present study were of the
same size and typical of representatives of the
genus Urospora. Although, species affiliation of the
observed oocysts was not identified in this study,
they obviously belong to one of the investigated
species.
The investigated gregarines differ in cell shape
and morphology (Table 1). In U. ovalis, the solitary
trophozoites do not demonstrate any signs
of cell heteropolarity. This phenomenon was also
described for gamonts of Gonospora ormieri
(Porchet 1978). Young and mature trophozoites
of U. travisiae are heteropolar. The young individuals
of U. travisiae can attach to the substrate
with the wider end, which is most likely the anterior
end. We assume that during gregarine growth,
a second branch of the cell starts to develop, so
that the cell gradually transforms from a monoaxial
to a V-like biaxial form. The anterior end of the
290 A. Diakin et al.
Table 2. Results of alternative topology tree tests (alignment of 23 OTUs, 1676 bp).
Tree topology – ln L BPa ELWb KHc SHd WSHe AUf
Bayesian
consensus tree
(Fig. 9, A)
13536.74 0.84736 0.84736 1.0 1.0 1.0 0.9023058
Alternative topologies
B 13686.58 0 0 0 0 0 0
C 13747.51 0 0 0 0 0 0
D 13813.34 0 0 0 0 0 0
E 13813.32 0 0 0 0 0 0
F 13541.49 0.04601 0.07659232 0.11657 0.64057 0.37045 0.1236613
G 13698.65 0 0 0 0 0 0
H 13747.51 0 0 0 0 0 0
I 13541.15 0.10663 0.1215358 0.13904 0.65436 0.41608 0.1578981
aBootstrap Probability (Felsenstein 1985);
bExpected-Likelihood Weights (Strimmer and Rambaut 2002);
cP-value of the Kishino-Hasegawa test (Kishino and Hasegawa 1989);
dP-value of the Shimodaira-Hasegawa test (Shimodaira and Hasegawa 1999);
eP-value of the Weighted Shimodaira-Hasegawa Test (Shimodaira and Hasegawa 1999);
fP-value of the Approximately Unbiased test (Shimodaira 2002). P-values <0.05 discard the suggested topology.
Permissible topologies are bolded.
young cell remains as the attachment site of V-like
shaped cell. The older (or primary) branch is usually
longer and contains the nucleus. It is important
to note that trophozoites of Pterospora spp. can
be characterised as biaxial as well (Dogiel 1910;
Landers 1991, 2001; Landers and Gunderson
1986; Leander 2008).
Most eugregarines possess a cortex with a complicated
structure (Schrével et al. 1983; Vivier 1968;
Vivier et al. 1970). Investigated gregarines from
Travisia forbesii also possessed a well-developed
cortex with numerous longitudinal epicytic folds.
In both species, rippled dense structures (RDS)
and 12-nm apical filaments were not well preserved
and were poorly detectable only in a few
folds, despite the application of different fixation
protocols. Electron-dense rods, observed in both
species, were located in the apex of the longitudinal
folds, under the IMC, and appear to be
similar to those described in Gonospora beloneides,
Lankesteria spp., Gregarina spp., Difficilina
cerebratuli, Thiriotia pisae, Ganymedes vibiliae,
and Porospora portunidarum (Corbel et al. 1979;
Desportes et al. 1977; Simdyanov 1995a, 2009;
Valigurová et al. 2013). It was assumed that it has
the role of reinforcing the fold tips (Valigurová et al.
2013).
Micropores, appearing as invaginations of the
plasma membrane encircled by a collar formed
from the IMC, are typical for apicomplexans.
The exact function of these structures, although
often discussed, remains unclear. Micropores could
function as organelles for the acquisition of nutrients
(Chobotar and Scholtyseck 1982; Scholtyseck
1973; Scholtyseck and Mehlhorn 1970; Vivier
et al. 1970) or as extrusomes for mucus secretion
(Desportes and Schrével, 2013; Philippe and
Schrével 1982; Valigurová et al. 2013; Vegni Talluri
and Dallai 1983). We observed typical micropores
and micropore-like structures (MLS) in U. ovalis
and U. travisiae. We assume that typical micropores
play a role in the gregarine’s acquisition
of nutrients, while the second structures serve to
secrete mucus onto the parasite surface in between
the folds. Under SEM and TEM, the excreted
mucus appears as droplets (Fig. 3E, 4B), as was
also demonstrated in other gregarines (Simdyanov
1995a, 2009; Valigurová et al. 2013; Walker et al.
1984). In addition, the pore-like structures interrupting
the IMC (and the plasma membrane, in some
cases), but lacking the collar, were documented in
the attachment site at the top of the protomerite
of Gregarina cuneata gamonts (Valigurová 2012).
These structures could also play a role in the secretion
of adhesive material, which is often present
between the attached gregarine and adjacent host
tissue. Mucous material was also observed on
the sucker-like protomerite of some actinocephalid
eugregarines (Cook et al. 2001).
In a series of transverse sections of the studied
gregarines, we observed the non-uniform accumulation
of presumably mucus-secretory vesicles
under the pellicle. It allows us to speculate that
mucus excretion occurs with different intensities
Morphology and Molecular Phylogeny of Coelomic Gregarines 291
in various zones of the parasite surface at the
same moment. Therefore, an eruptive (or impulsive)
nature of the mucus excretion could take place
in the investigated gregarines. We can assume
that the release of the mucus by gregarine into
the environment may serve as a protective mechanism
against attack by host coelomocytes. It is
expected that the gliding motility of eugregarines is
facilitated by the specific structure of the epicytic
folds; therefore, mucus secretion may also help
to decrease frictional forces during forward movement.
This was previously proposed for Gregarina
spp., in which the mucus load in the gregarine cytoplasm
was positively correlated with gliding speed
(King 1981, 1988; Mackenzie and Walker 1983;
Valigurová et al., 2013; Vávra and Small 1969).
Gliding motility is characteristic of the majority of
eugregarines. This motility can easily be observed
in free (non-attached) eugregarines contacting with
a substrate, when they move forward, usually without
any obvious changes in their cell shape (unlike
metaboly or rolling). However, when attached to the
host tissue, eugregarines do not usually demonstrate
any signs of motility, although near-surface
currents in the internal environment of the host
can be noticed around parasites. Obviously, parasites
produce these currents by themselves. It is
important to note that contact between a gregarine
and solid matter might not be necessary for gregarine
motion, as Gregarina spp. gamonts are able
to free-float in a liquid lacking any contact with the
substrate and with a significantly higher rate than
exhibited during regular gliding (Valigurová et al.
2013).
Metaboly is another type of motility, which is
accompanied by significant changes in cell shape.
Gregarines demonstrating metabolic (or peristaltic)
motility, or even immobility, as a rule, possess an
epicyte of unusual organisation. Various modifications
of the epicyte typical for eugregarines along
with the loss of gliding motility have been reported
in representatives of the families Monocystidae and
Urosporidae, parasitising the coelom, respiratory
trees, and seminal vesicles of various invertebrates
(Dyakin and Simdyanov 2005; Frolov 1991;
Landers 1991, 2001; Landers and Gunderson
1986; Landers and Leander 2005; MacMillan
1973; Miles 1968; Vinckier 1969; Vinckier and
Vivier 1968). Urospora ovalis and U. travisiae both
possess a typical epicyte; nevertheless, the trophozoites
and young syzygies of U. ovalis demonstrate
metabolic motility, during which the cell cortex in
the contracted regions generates several superfolds.
Similar superfolds have been documented in
trophozoites of Nematocystis magna, a monocystid
eugregarine demonstrating peristaltic-like motility
(MacMillan 1973; Miles 1968). Mitochondria
observed in U. ovalis are similar to those reported in
Pterospora floridensis (Landers 2002). We assume
that the relatively great number of mitochondria distributed
uniformly in the cell cytoplasm appears to
be correlated with the active metaboly of U. ovalis.
The motility of gregarines seems to be an adaptation
to their localisation in a certain niche in the
host body. Gregarines inhabiting the host digestive
tract usually possess pendular or gliding motility
types (e.g. Selenidium spp., Lecudina spp.). They
develop clamped in narrow spaces between the
intestinal folds or between the intestine and food.
We assume that motility in intestinal gregarines
might be necessary to provoke an exchange of host
internal environmental liquids around them in order
to improve the effectiveness of their nutrient acquisition
and/or reduce frictional forces to retain the
attachment to host tissues, as also suggested by
Leander (2008). This also applies to U. travisiae,
which attaches to the outer wall of the intestine.
Metaboly seems to be characteristic of detached
parasites (numerous representatives of the families
Urosporidae and Monocystidae) inhabiting the
host body cavities. Presumably, monocystids have
acquired this type of motility as an adaptation to
their life style, being detached and motile within the
gametes agglomeration of oligochaete hosts. Moreover,
some urosporids could have evolved their
motility as an adaptation for living in the liquid environments
of host cavities without any attachment to
host tissue. In addition, motility in gregarines could
provide an effective protection against the adhesion
of host coelomocytes to their surfaces, as shown for
other coelomic parasites (De Ridder and Jangoux
1984; Coulon and Jangoux 1988, 1991; Siedlecki
1903).
Both gregarines investigated in this study
showed signs of active metabolism, represented by
the presence of various vacuoles and inclusions.
We assume that, along with an increase in amylopectin
load, the maturation of trophozoites of both
species is accompanied by an increase in the quantity
and size of these inclusions and vacuoles, which
occupy almost the entire cell volume. Their function
remains unclear; however, we can suggest that
their contents may be used for further gametocyst
and oocyst wall formation, similar to wall-forming
bodies in coccidia (Long 1982).
The frontal and lateral syzygies are characteristic
of urosporids (Levine 1977). In the present study,
we documented end-to-end syzygy in U. ovalis and
frontal syzygy in U. travisiae. We cannot identify the
exact type of syzygy present in U. ovalis gregarines:
292 A. Diakin et al.
Figure 6. General morphology and fine structure of Urospora travisiae trophozoites. A. General view of a
trophozoite. White double arrowhead points to the attachment site placement, b - beads of the branches.
SEM. B. Detailed view of the attachment tip; traces of the epicytic folds on the surface of the plateau (tef) and
well-developed epicytic folds (ef) extended from the furrow are well visible. SEM. C. Higher magnification of the
gregarine surface demonstrating epicytic folds. SEM. D. Higher magnification of the distal part of the branch
with one bead. White dots mark the beginning/end of additional epicytic folds on the bead surface. SEM.
E. Higher magnification of the constricted region between two individual beads. Asterisks mark the epicytic
folds terminating near the region of constriction. Dis – distal end of the branch, Pro – proximal end of the branch.
Morphology and Molecular Phylogeny of Coelomic Gregarines 293
caudo-frontal, frontal, or caudal. The syzygy of U.
travisiae is obviously frontal, as partners are in
contact by their attachment tips. Syzygies of U.
travisiae are comparable with those of Pterospora
spp., in which partners are of V-like shape, with
two piriform branches possessing posterior dendritic
trunks (Landers 1991, 2001; Landers and
Gunderson 1986; Landers and Leander 2005).
Molecular phylogenetic analyses confirmed the
morphological data: both species belong to the
clade Lecudinoidea comprising representatives of
the families Lecudinidae and Urosporidae. Previously,
it was shown that, within the Lecudinoidea
clade, some species have small interspecific differences
(Mita et al. 2012; Rueckert et al. 2015).
Alternatively, some of them have high intraspecific
differences in SSU rDNA (Rueckert et al. 2011b),
even in the species with negligible morphological
interspecific differences (Rueckert et al. 2010). In
a pair of studied species U. ovalis and U. travisiae
a set of distinctive sites in the sequences was
identified by direct sequencing of PCR products
from genomic DNA obtained from 10-20 individuals
of each species, and distinctive nucleotides
were neither polymorphic nor ambiguous (Supplementary
Material Fig. S1). Consequently, there is
a genetic hiatus conforming to considerable morphological
differences between gregarines of these
species. Further investigations including single-cell
sequencing and analysing of other genetic markers,
such as ITS2, LSU rDNA and the whole ribosomal
operon sequences, are desirable for revealing of
distinctions, undiscovered by cell-pool sequencing,
between U. ovalis and U. travisiae.
In our study, the representatives of the families
Lecudinidae and Urosporidae were partially mixed
with each other (clades corresponding to these families
were not well-supported). At the same time,
representatives of the family Urosporidae exhibited
a clear diagnostic feature, probably a synapomorphy:
the characteristic morphology of oocysts, with
a funnel at one of the poles and one or more projections
of the oocyst wall, sometimes quite long,
at the opposite pole (Dogiel 1906, 1909, 1910;
Léger 1892). Therefore, we assume that SSU rDNA
phylogeny cannot resolve the real branching order
in the clade Lecudinoidea, either because of the
insufficient sensitivity of the method or because of
the limited number of taxon samples. Furthermore,
removing the Veloxidium leptosynaptae sequence
(see below) from the analysis led to the uniform
shuffling of all lecudinids and urosporids within the
clade (data not shown). The high sensibility of gregarine
SSU rDNA phylogeny to taxonomic sample
size was noted earlier (Simdyanov et al. 2015).
On the other hand, SSU rDNA phylogeny confirms
the close relations between lecudinids and
urosporids, which were included in the ‘aseptate’
eugregarines, parasites of marine invertebrates,
according to Grassé’s concept of host-parasite
coevolution (Grassé 1953).
In this study, Veloxidium leptosynaptae, an
unusual gregarine from the intestine of the sea
cucumber Leptosynapta clarcki, was closely affiliated
with the clade Lecudinoidea (Figs 8, 9). In
addition, the SSU rDNA sequence of V. leptosynaptae
possessed motifs that were unique to this
clade. However, on the basis of the bending motility
and surface morphological characteristics of the
gregarine, the authors of the original description
of V. leptosynaptae (Wakeman and Leander 2012)
suggested that this species belongs to the order
Archigregarinorida. Nevertheless, they noticed that
the ‘Veloxidium clade’ branched as the nearest
sister lineage to the clade of marine lecudinids
and urosporids. We have several objections to
such a classification of V. leptosynaptae: 1) the
syzygy of V. leptosynaptae appears frontal (typical
for lecudinids) rather than caudal (typical for
archigregarines); and 2) some eugregarines are
also capable of bending their body, sometimes quite
dramatically (Hildebrand 1981; Simdyanov, 1995b;
Valigurová et al. 2013). Therefore, further TEM
studies of the cortex are necessary to establish
the taxonomic position of V. leptosynaptae more
reliably.
It was suggested that coelomic gregarines
evolved more than once from different marine
intestinal eugregarines (Leander et al. 2006). One
of the possible ways for the transition from intestinal
to coelomic parasitism was demonstrated in
some marine eugregarines from polychaetes when,
during host reproductive metamorphosis, intestinal
gregarines located in the host body cavity for a
short time (Durchon and Vivier 1961). Such temporary
location of gregarines in the host coelom
➛
SEM. F. Higher magnification showing converging and merging epicytic folds (black arrows) that cover the
lateral side, black arrowheads mark folds passing from one branch (br1) to another (br2) on the inner surface
of the cell near the attachment tip, and folds (white arrowheads) arising from the attachment tip (white double
arrowheads). SEM. G. Tangential section of the constricted region between adjoining beads. ef – epicytic folds;
mt – microtubules. TEM.
294 A. Diakin et al.
Figure 7. Fine structure of the cortex and cytoplasm of Urospora travisiae trophozoites. A. Transversal section
of a trophozoite demonstrating the cell cortex in detail. Note a typical micropore (mp) between epicytic folds
(ef) and an electron-dense rod (r) in the apex of each fold. inl – internal lamina, mit – mitochondrion, mt –
microtubules. TEM. B. Transversal section of the gregarine cortex showing a micropore-like structure (black
double arrowhead) with a closely located electron-dense vesicle (white arrowhead), and an electron-dense
inclusion between cortex cytomembranes (inc). TEM. C. Transversal section of one of the branches near the
attachment tip. The cytoplasm is subdivided into ectoplasm (ect) and endoplasm (end). TEM. D. Transversal
section of a trophozoite demonstrating the cell cortex and ectoplasm (ect) in detail. Ga – Golgi apparatus, inc –
electron-dense inclusion; mit – mitochondrion, vac – electron-transparent vacuoles. TEM. E. Detailed view of
the cell ecto- (ect) and endoplasm (end) in transversal section; ld – lipid droplets; mit – mitochondrion. TEM.
Morphology and Molecular Phylogeny of Coelomic Gregarines 295
Figure 8. SSU rDNA Bayesian tree of alveolates (99 OTUs, alignment of 1557 bp) constructed using the
GTR+ +I model. Numbers at the nodes denote Bayesian posterior probabilities (numerator) and ML bootstrap
percentage (denominator). Black disks on the branches indicate the Bayesian posterior probabilities and the
bootstrap percentages equal to or more than 0.95 and 95%, respectively. A black box highlights the sequences
from Urospora ovalis and U. travisiae.
296 A. Diakin et al.
Figure 9. Results of topology tests of 23 selected lecudinid and urosporid sequences. A. SSU rDNA Bayesian
tree of selected OTUs (alignment of 1676 bp) constructed using the GTR+ +I model (8 categories). Numbers
at the nodes denote Bayesian posterior probabilities (numerator) and ML bootstrap percentage (denominator).
A black box highlights the sequences from Urospora ovalis and U. travisiae; black disks on the branches
indicate the Bayesian posterior probabilities and the bootstrap percentages equal to or more than 0.95 and
95%, respectively. B – I. alternative topologies which were tested together with tree A; there are just three
permissible topologies: A, F, and I, marked by circles. Abbreviations: D = Difficilina spp., Le = Lecudina spp. +
Lankesteria spp., Li = Lithocystis sp., Pa = Paralecudina polymorpha, Pt = Pterospora spp., U = Urospora spp.,
V = Veloxidium leptosynaptae.
Morphology and Molecular Phylogeny of Coelomic Gregarines 297
could be fixed in their life cycle. It is an argument
that supports the hypothesis that haemocoelic gregarines
originated from intestinal ones in insects
(Léger 1892). The explanation for such a transition
is that phylogenetically related parasites, occupying
different ecological niches in the host, decrease the
intensity of species competition, and demonstrate
diverse adaptations to parasitism (Wakeman et al.
2014).
Conclusions
This study revealed that the closely related eugregarines
Urospora ovalis and U. travisiae, inhabiting
the coelom of the polychaete Travisia forbesii,
demonstrate different strategies of parasitism. The
V-shaped cells of U. travisiae attach to the host and
retain gliding motility when detached. In contrast,
the cells of U. ovalis are oval-shaped, non-attached,
and exhibit peristaltic activity. Both gregarines
possess a typical organised cortex with epicytic
folds of similar structure. In metabolic U. ovalis, the
cell cortex generates superfolds in the contracted
regions.
Methods
The cells of both gregarine species (Urospora ovalis and
Urospora travisiae) were isolated from the coelom of the marine
polychaete Travisia forbesii Johnston, 1840. The hosts were
collected from June to July each year from 2004 to 2006
and in the second half of August each year from 2011 to
2013 at upper sublittoral of two sites: in the vicinity of the
Marine Biological Station of Saint-Petersburg State University
(inlet Yakovleva, Chupa Inlet, Kandalaksha Bay, White Sea,
66◦
18 99 N, 33◦
49 95 E) and the White Sea Biological Station
of Moscow State University (Rugozerskaya Inlet, Kandalaksha
Bay, White Sea, 66◦
33 12 N, 33◦
06 17 E).
The dissection of hosts and subsequent manipulation with
parasites was performed under MBS-10 stereomicroscopes
(LOMO, Russia). Light micrographs were provided using an
MBR-1 microscope (LOMO, Russia) equipped with phase contrast
and connected to a Canon EOS 300D digital camera.
The gregarines of both species were isolated separately with
thin glass pipettes, washed in Millipore filtered sea water (SW)
(Millex-GC 0.22 ␮m) and subsequently prepared for light, electron
microscopy and DNA extraction.
Histological procedure: Several entire worms were anesthetised
and fixed in AFA (Alcohol–Formalin–Acetic Acid)
fixative solution. The material was dehydrated through a graded
alcohol series, cleared in xylene, infiltrated in a graded series
of xylene/Histoplast II (3:1, 1:1, 1:3) and finally embedded in
Histoplast II (Sigma-Aldrich, Czech Republic). Serial sections
(transversal, sagittal, and coronal) of the fixed worms were
prepared on a Microm HM 360 rotary microtome and stained
with haematoxylin-eosin. Micrographs were obtained using an
Olympus BX61 microscope equipped with an Olympus DP 71
digital camera.
Electron microscopy: The hosts were dissected and the
parasites were collected separately from the host body cavity
using thin glass pipettes. The trophozoites were fixed in 2%
or 2.5% glutaraldehyde in 0.1 M cacodylate buffer (CB), 0.1 M
PBS, or SW. For transmission electron microscopy the gregarines
were then post-fixed with 1% OsO4 (Os) in 0.2 M CB,
0.1 M PBS or SW, dehydrated in an ethanol series, and embedded
into Epon blocks. The ultra-thin sections were stained
according to standard protocols (Reynolds 1963) and observed
with LEO-910 and JEOL-1010 transmission electron microscopes.
For scanning electron microscopy, fixed trophozoites
were critical point dried in liquid CO2 and then coated with gold.
The samples were observed with a JEOL JSM-7401F scanning
electron microscope.
DNA isolation, PCR and sequencing: Individual trophozoites
of each species, about 10 and 20 trophozoites of
Urospora ovalis and Urospora travisiae, respectively, were isolated
from dissected hosts, washed three times in Millipore
filtered SW, and deposited into 0.5 ml microcentrifuge tubes.
All samples were fixed and stored in RNA-later reagent (Life
Technologies, USA). DNA extraction was performed with the
Diatom DNA Prep 200 kit (Isogen, Russia).
The new partial SSU rDNA sequences (1623 bp for U. travisiae
and 1603 bp for U. ovalis) were amplified with Encyclo PCR
kit (Evrogen, Russia) using a T3000 Thermocycler (Biometra,
Germany) according to the following protocol: initial denaturation
at 95 ◦
C for 3 min; 40 cycles of 95 ◦
C for 30 sec, 45 ◦
C for
30 sec, and 72 ◦
C for 1.5 min; and a final extension at 72 ◦
C for
10 min with primers 5 -GTAGTCATAYGCTTGTCTYGC-3 (forward)
and 5 - GATCCTTCTGCAGGTTCACCTAC -3 (reverse).
Only weak bands of an expected size were obtained by electrophoresis
in agarose gel; therefore, small pieces of the gel
were sampled from those bands (using pipette tips under a transilluminator)
and re-amplification PCR with ColoredTaq DNA
polymerase kit (Silex, Russia) using the DNA Engine Dyad
thermocycler (Bio-Rad) and the same primers was performed.
PCR products of the expected size were gel isolated using a
Cytokine DNA isolation kit (Cytokine, Russia) and sequenced
using an ABI PRISM BigDye Terminator v. 3.1 reagent kit
on an Applied Biosystems 3730 DNA Analyzer automatic
sequencer. The newly obtained SSU rDNA sequences (GenBank
Accession numbers: Urospora ovalis KR868712 and
Urospora travisiae KR868713) were preliminarily identified by
BLAST analysis including the built-in NJ-tree tool.
Molecular phylogenetic analysis: The two novel SSU
rDNA sequences were aligned with 97 other SSU rDNA
sequences, representing the major lineages of apicomplexans,
as well as dinoflagellates, ciliates, heterokonts and rhizarians
as outgroups, using the MUSCLE 3.6 programme (Edgar 2004)
and manual tuning with the BioEdit 7.0.9.0 programme (Hall
1999). After removing hypervariable regions, the length of the
alignment of the final 99 operational taxonomic units (OTUs)
was 1557 sites. Bayesian analysis of this alignment was conducted
using the MrBayes 3.2.1 programme (Ronquist and
Huelsenbeck 2003). The programme was set to operate using
the following parameters: nst = 6, ngammacat = 8, rates =
invgamma, covarion = yes; parameters of Metropolis Coupling
Markov Chains Monte Carlo (mcmc): nchains = 4, nruns = 4,
temp=0.2, ngen = 7 000 000, samplefreq = 1 000, burninfrac =
0.5 (the first 50% of 7 000 sampled trees, i.e. the first 3500,
were discarded in each run). An average standard deviation of
split frequencies of 0.013232 was achieved at the end of calculations.
Maximum-likelihood analysis of the 99 OTU alignment
and calculations of Bayesian tree bootstrap support were performed
with the RAxML 7.2.8 programme (Stamatakis 2006)
under the GTR+ +I model with 4 categories of discrete gamma
298 A. Diakin et al.
distribution. The procedure included bootstrap analysis with
1000 replicates and 100 independent runs of ML analysis. All
of these computations were performed using the University of
Oslo Bioportal free service (www.bioportal.uio.no).
For the testing of alternative topologies, another alignment of
23 selected OTUs including lecudinids and urosporids (all available
sequences from GenBank) was created. The sequences of
Veloxidium leptosynaptae and Paralecudina polymorpha were
used as outgroups for this analysis. This gave us the opportunity
to include 119 additional nucleotides from hypervariable
regions in the analyses, so that the final length of the alignment
increased to 1676 bp. Topology tests for the 23 OTUs
Bayesian tree were performed using the TREEFINDER programme
under the same model as in the Bayesian analyses
(GTR+ +I, 8 categories) (Jobb 2011; Jobb et al. 2004).
List of Abbreviations
BF – Bright Field light microscopy; CB – Cacodylate
Buffer; DIC – Differential Interference Contrast
microscopy; IMC – Inner Membrane Complex;
H&E – Haematoxylin & Eosin staining; LM – Light
Microscopy; ML – Maximum-likelihood analyses;
MLS – Micropore-like structures; OTU – Operational
Taxonomic Unit; PBS – Phosphate Buffered
Saline; PC – Phase Contrast light microscopy;
RDS – Rippled Dense Structure; SEM – Scanning
Electron Microscopy; SW – Sea Water; TEM
– Transmission Electron Microscopy
Author Contributions
AD and GGP conceived and designed the study,
performed field sampling, carried out the research,
performed the light and electron microscopic
analyses, and wrote the manuscript. AD and
TGS designed and performed the molecularbiology
experiments and phylogenetic analyses.
VVA provided the laboratory equipment, necessary
reagents, and materials, and gave consultations.
AV contributed to material collection and
processing (2011-2013), to the light microscopic
observations, and to the interpretation of microscopic
data. All authors contributed to the writing
of the manuscript, and read and approved the final
manuscript.
Acknowledgements
The authors are grateful to the staff of the Marine
Biological Station of Saint-Petersburg State University
(MBS SPbSU) and the Nikolai Pertsov White
Sea Biological Station of Lomonosov Moscow State
University (WSBS MSU) for providing them with
on-site facilities for field sampling and material
processing. The authors are greatly indebted to
members of the Laboratory of Electron Microscopy
(Institute of Parasitology, BC ASCR in ˇCeské
Budˇejovice) for their technical assistance. AD and
GGP would like to thank Andrej Dobrovolskij (Dept.
of Invertebrate Zoology, St. Petersburg State University)
for initiating this study, and also Vladimir
Semenov and Tatiana Kharkievitch (Sechenov
Institute of Evolutionary Physiology and Biochemistry
of the Russian Academy of Sciences) for
their assistance with electron microscopy. A part of
the present study was performed at the Resource
Centres (Culture Collections of Microorganisms,
Molecular and Cell Technologies, Observatory of
Environmental Safety) of St. Petersburg State
University. GGP is grateful to Professor Rudolf
Entzeroth for providing facilities for her research
at Dresden Technical University (TUD) and to
Markus Gunther (TUD) for his assistance with SEM.
DNA sequencing was performed at the “Genome”
DNA sequencing centre (Engelhardt Institute of
Molecular Biology, Russian Academy of Sciences,
www.genome-centre.ru). AV and AD were funded
by project No. GBP505/12/G112 from the Czech
Science Foundation (ECIP - Centre of Excellence)
and acknowledge support from the Department
of Botany and Zoology, Faculty of Science,
Masaryk University towards the preparation of this
manuscript. GGP was supported by St. Petersburg
State University grants (1.42.1493.2015,
1.42.1099.2016), a Michail Lomonosov stipendium
(DAAD), and TUD grants. TGS was supported by
grants from the Council of the President of the
Russian Federation (NSh-7770.2016.4) and from
the Russian Foundation of Basic Research (15-
29-02601). The phylogenetic analysis in this study
was supported by Russian Scientific Foundation
grant No. 14-50-00029. The authors also grateful
to Matthew Nicholls for English proof-reading.
Appendix A. Supplementary data
Supplementary data associated with this article
can be found, in the online version, at
http://dx.doi.org/10.1016/j.protis.2016.05.001.
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Available online at www.sciencedirect.com
ScienceDirect
7.7
Schrével J., Valigurová A., Prensier G., Chambouvet A.,
Florent I., Guillou L.
2016
Ultrastructure of Selenidium pendula,
the type species of archigregarines, and phylogenetic relations
to other marine Apicomplexa
Protist 167(4), 339-368
Protist, Vol. 167, 339–368, August 2016
http://www.elsevier.de/protis
Published online date 18 June 2016
ORIGINAL PAPER
Ultrastructure of Selenidium pendula, the
Type Species of Archigregarines, and
Phylogenetic Relations to Other Marine
Apicomplexa
Joseph Schrévela,1
, Andrea Valigurováb
, Gérard Prensierc,2
, Aurélie Chambouvetd
,
Isabelle Florenta
, and Laure Guilloue
aUnité Molécules de Communication et Adaptation des Microorganismes, (MCAM, UMR
7245), Muséum National Histoire Naturelle, Sorbonne Universités, CNRS, CP 52, 57 Rue
Cuvier, 75005 Paris, France
bDepartment of Botany and Zoology, Faculty of Science, Masaryk University, Kotláˇrská 2,
611 37 Brno, Czech Republic
cCell Biology and Electron Microscopy Laboratory, Franc¸ois Rabelais University, 10
Boulevard Tonnellé, BP 3223, 37032 Tours Cedex, France
dLaboratoire des Sciences de l’Environnement Marin (LEMAR), UMR6539
UBO/CNRS/IRD/IFREMER, Institut Universitaire Européen de la Mer (IUEM),
Technopole Brest Iroise, 29280 Plouzané, France
eSorbonne Universités, Université Pierre et Marie Curie - Paris 6, CNRS, UMR 7144,
Station Biologique de Roscoff, Place Georges Teissier, CS90074, 29688 Roscoff cedex,
France
Submitted February 3, 2016; Accepted June 12, 2016
Monitoring Editor: Frank Seeber
Archigregarines, an early branching lineage within Apicomplexa, are a poorly-known group of invertebrate
parasites. By their phylogenetic position, archigregarines are an important lineage to understand
the functional transition that occurred between free-living flagellated predators to obligatory parasites
in Apicomplexa. In this study, we provide new ultrastructural data and phylogenies based on
SSU rDNA sequences using the type species of archigregarines, the Selenidiidae Selenidium pendula
Giard, 1884. We describe for the first time the syzygy and early gamogony at the ultrastructural level,
revealing a characteristic nuclear multiplication with centrocones, cryptomitosis, filamentous network
of chromatin, a cyst wall secretion and a 9+0 flagellar axoneme of the male gamete. S. pendula belongs
to a monophyletic lineage that includes several other related species, all infecting Sedentaria Polychaeta
(Spionidae, Sabellaridae, Sabellidae and Cirratulidae). All of these Selenidium species exhibit
similar biological characters: a cell cortex with the plasma membrane - inner membrane complex subpellicular
microtubule sets, an apical complex with the conoid, numerous rhoptries and
1
Corresponding author; fax +33 1 40 79 34 99
2
This work is dedicated to the memory of Brigitte Arbeille-Brassart.
e-mail schrevel@mnhn.fr (J. Schrével).
http://dx.doi.org/10.1016/j.protis.2016.06.001
1434-4610/© 2016 Elsevier GmbH. All rights reserved.
340 J. Schrével et al.
micronemes, a myzocytosis with large food vacuoles, a nuclear multiplication during syzygy and young
gamonts. Two other distantly related Selenidium-like lineages infect Terebellidae and Sipunculida,
underlying the ability of archigregarines to parasite a wide range of marine hosts.
© 2016 Elsevier GmbH. All rights reserved.
Key words: Archigregarines; Apicomplexa; Selenidium pendula; ultrastructure; phylogeny; sporozoite.
Introduction
Apicomplexa, a large subgroup of the Alveolata,
are unicellular parasites infecting a wide range of
invertebrate and vertebrate hosts. Most known Apicomplexa
belong to Coccidia and Haemosporidia
and are involved in human and veterinary diseases
(malaria, toxoplasmosis, coccidiosis, babesiosis,
piroplasmosis). However, a very large group of
the early branching Apicomplexa, the gregarines,
is comparatively poorly known. Most of the gregarines
infect invertebrate hosts and usually do not
have deleterious effects on their hosts (Desportes
and Schrével 2013). The number of Apicomplexa
species is estimated to be ∼2,000-6,000, however
the ability of gregarines to infect a wide range of
insects could significally enhance this estimation to
several thousand or more than one million (e.g., the
Coleoptera (beetles) class corresponds to about
40% of the insect biodiversity with an expected
species number around 1 to 2 million) (Schrével
and Desportes 2013).
All apicomplexan species are characterized
by an infective life stage, the so-called zoite,
a polarized cell with an original apical complex.
This apical complex is an assembly of
specific organelles including club-shaped rhoptries,
filament-like micronemes, dense granules
and apical polar rings. In Apicomplexa, the presence
of a conoid in the apex of the zoite,
observed in coccidia and gregarines, defines the
Conoidasida Levine, 1988. In contrast, no conoid is
observed in Haemosporidia and Piroplasmida designated
Aconoidasida Mehlhorn et al., 1980. Except
gregarines and some other taxa developing in epicellular
localization, such as cryptosporidia, most
Apicomplexa have an intracellular development in
their host cells and there, the apical organelles
as well as the conoid, play an essential role in
cell invasion processes through sophisticated cascades
of molecular interactions (Boothroyd and
Dubremetz 2008; Bradley et al. 2005; Santos et al.
2009). In Apicomplexa displaying an intracellular
life style, the cycle usually occurs in two hosts,
the sexual phase being performed in the definitive
host while asexual phases occur in one or several
intermediate hosts. Gametogenesis, as observed
in Coccidia or Haemosporidia, exhibits a clear
anisogamy with production of small flagellated male
gametes (microgametes) and large non-flagellated
female gametes (macrogametes). After fertilization,
the sporogony produces sporozoites in the definitive
hosts while asexual schizogny or merogony,
producing merozoites, is realized in intermediate
hosts. In contrast, most gregarines exhibit
an extracellular development and their entire life
cycle usually occurs within a single host. Their
zoites transform into large vegetative cells, the
trophozoites, with an extraordinary diversity in
their morphologies and behaviours. In addition to
this extracellular development, gregarines share
a unique sexual phase. The sexual association
between two gamonts, named syzygy, produces
a cyst where the gametogenesis differentiates
a large and equal number of male and female
gametes; at this stage, this cyst is called a gametocyst.
Then, fertilization and sporogenesis take
place within the cyst yielding the final stages with
the sporocysts usually containing each 8 sporozoites.
These sporocysts can survive for a long
period generally waiting for their ingestion by their
specific hosts. Gregarine biochemistry and physiology
are still poorly documented. Studies of their
zoite apical apparatus as well as of the variation
of their cytoskeleton and microtubule organizing
centers (MTOCs), with unique organization
as the 6+0 or 3+0 flagellar axonemes described
for some male gametes (Prensier et al. 1980;
Schrével and Besse 1975), contributed, however,
to a more general understanding of many biological
aspects of Apicomplexa including pathogenic
species.
Among Apicomplexa, there is a consensus on the
stem group of archigregarines commonly found in
Polychaeta, Sipunculida and some Hemichordata.
These marine gregarines represent the earliest
diverging lineage of Apicomplexa (Leander 2007a;
Schrével 1971b). The type species of archigregarines
is Selenidium pendula Giard 1884 and
its life cycle was established during the second
part of the 20th century (Schrével 1966,
1970). Beside this type species, a long series
Selenidium pendula: Ultrastructure and Phylogeny 341
of contributions have been performed on other
Selenidium and related species at the cytological
(Brasil 1907, Caullery and Mesnil 1899,
1900, Ray 1930, Reed 1933) and ultrastructural
(Leander 2006, 2007b; Macgregor and Thomasson
1965; Schrével 1968, 1970, 1971a; Simdyanov
and Kuvardina 2007, Vivier and Schrével 1964,
1966) levels and more recently also at the molecular
level through the analysis of SSU rDNA
sequences (Leander et al. 2003a; Leander 2006,
2007b; Rueckert and Leander 2009; Wakeman
and Leander 2012, 2013; Wakeman et al. 2014).
Most of these studies focused on the trophozoite
stages with few descriptions on nutrition modalities
(Schrével 1968; Simdyanov and Kuvardina 2007).
Additionally, these studies highlighted several
incongruities among Selenidiidae at the molecular
level that could not be elucidated in absence
of the type species of the family. Here, we report
on the cell organization of the Selenidium pendula
trophozoite with a special attention to the conoid,
the abundance of rhoptries and micronemes, and
we provide the first ultrastructural description of the
syzygy (pairing stage), the early gamogony with the
cryptomitosis and the secretion of the cyst walls.
We also provide the first phylogenetic analysis of
the SSU rDNA gene sequences encompassing
the type species of archigregarines S. pendula.
Molecular phylogenetic analyses revealed three
lineages within archigregarines, S. pendula belonging
to the Selenidiidae that includes parasites of
Spionidae, Sabellidae, and Sabellariidae, all polychaete
annelids, as well as two Selenidiidae–like
lineages, parasites of hosts belonging to Terebellidae
and Sipunculida, respectively.
Results
The Trophozoite of Selenidium pendula
The mature S. pendula trophozoite is a crescentshaped
cell of about 150 ␮m in length with a
circular cross section of about 35 ␮m in diameter.
The cell surface exhibits about 30 striations
in phase contrast light microscopy as well as in
scanning electron microscopy (SEM), appearing
as a series of longitudinal bulges of about 2.5-
3 ␮m in width separated by grooves (Fig. 1A).
The trophozoite is inserted into the intestinal
epithelium of the Scolelepis squamata polychaete
worm by a special apical apparatus called the
mucron (Figs 1B, 2A-C). In transmission electron
microscopy (TEM), a tropism for host cells rich in
granules can be observed (Fig. 1B). The mucron of
S. pendula corresponds to the attachment apparatus
anchoring the parasite to the host epithelial cell.
In SEM, the mucron appears as a regular mammiliform
area without bulges and grooves (Fig. 2A).
After detachment of the trophozoite, the trace of
the mucron in the host cell is very regular with
sometimes a small hole in a subcentral position
(Fig. 2B).
A series of short microvilli is seen at the periphery
of the epithelial cells (Fig. 2B). All around
the trophozoite attachment, numerous long ciliary
structures of the host epithelium are observed
(Figs 1A, 2B).
Asexual schizogony in S. pendula could be an
explanation to the exceptional clotting of trophozoites,
with thousands and thousands of cells that
obstruct the intestinal lumen of some Scolelepis
squamata hosts. In vivo, trophozoites are dispersed
along the host intestine, except for the first
thirty segments. The distinction between these two
intestinal regions is facilitated by the yellow color of
the first segments versus the green color of the posterior
region. Motility of the S. pendula trophozoites
is clearly of pendular type, as proposed by Giard
(1884) for the species diagnosis, and the stroboscopic
records show regular pendular beats with a
period of about 0.2 second (Golstein and Schrével
1982).
In TEM cross sections, the bulges of S. pendula
exhibit a characteristic ultrastructure described for
the first time in Selenidium hollandei (Vivier and
Schrével 1964). The plasma membrane is underlain
by a regular flat vesicle designated as the inner
membrane complex (imc) while a very slight cell
coat covers the cell surface. Under these three
cortical membranes, a regular set of longitudinal
subpellicular microtubules and some other dispersed
microtubules within the cortical cytoplasm
are seen below the bulges but not in the area of
the grooves (Fig. 3B-C). In TEM cross sections,
each subpellicular microtubule of S. pendula is surrounded
by an electron-lucent sheath (Fig. 3C) as
observed in S. hollandei (Vivier and Schrével 1964),
Platyproteum (Selenidium) vivax (Leander 2006),
Selenidium serpulae (Leander 2007b) and Selenidium
terebellae (Wakeman et al. 2013). Abundant
mitochondria are present under the subpellicular
network of the bulges.
Different ectoplasmic structures along the
grooves are observed with lamellar elements,
dense material structures that crossed the imc and
are in contact with the plasma membrane (Fig. 3B,
D-F). Under SEM, series of holes are observed
in the grooves with an irregular distribution and
distances ranging from 0.3-0.4 ␮m to 0.8-0.9 ␮m
342 J. Schrével et al.
Figure 1. Scanning and transmission electron microscopy of Selenidium pendula trophozoites fixed to the
intestine of the polychaete worm Scolelepis squamata (A-B). Abbreviations: bulge (B), dense granule (DG),
food vacuole (FV), groove (G), intestinal epithelium (IE), mucron (MU), rhoptry (R). A. SEM micrograph of
trophozoites with their apical region inserted into the intestinal epithelium, exhibiting on this face about 18
longitudinal bulges separated by grooves. The long filamentous structures covering the intestinal epithelium
correspond to ciliary structures (arrows). B. Longitudinal TEM section of a trophozoite with the apical end
designated as mucron containing a food vacuole and numerous rhoptries. In the intestinal epithelium, the
trophozoite preferentially anchors to the host cells enriched in dense granules having mucous secretions.
(Fig. 3A). Such a distribution seems to correspond
to the opening sites of the above-mentioned
ectoplasmic structures and their density might
indicate a role that was previously underestimated.
Interestingly, the longitudinal microtubular bundles,
abundantly distributed beneath the cortex in
the trophozoite apical part corresponding to the
mucron, could represent the biogenesis site of the
Selenidium pendula: Ultrastructure and Phylogeny 343
Figure 2. Apex of the Selenidium pendula trophozoite (A-D). Abbreviations: bulge (B), conoid (Co), food vacuole
(FV), groove (G), intestinal epithelium (IE), microneme (mn), microtubules (mt), microvilli (mv), mucron
(MU), rhoptry (R). A. SEM micrograph of the apex surface showing that bulges and grooves of the epicyte start
from a regular mammiliform area corresponding to the external surface of the mucron. Small folds (arrows) are
observed on the bulges located on the internal curvature of the cell B. SEM micrograph of intestinal epithelium
after the detachment of a mucron, with small microvilli on the periphery, a small hole in the subcentral position
(white arrow) and the long ciliary structures (black arrow). C. TEM micrograph of a median longitudinal section
of the apex with several food vacuoles that enter via the conoid and are surrounded by an accumulation of
rhoptries and micronemes. D. TEM longitudinal section of the apical region (= trophozoite apex with numerous
micronemes and rhoptries) revealing that the subpellicular microtubule bundles start before the differentiation
of the epicytic bulges of the cell surface.
344 J. Schrével et al.
Figure 3. Cell surface and cortex of Selenidium pendula trophozoite (A-F). Abbreviations: bulge (B), groove (G),
inner membrane complex (imc), microneme (mn), microtubules (mt), mitochondrion (M), myelin-like structure
(st myel), plasma membrane (pm), pore (p), rhoptry (R), vesicle (ves). A. SEM view of the cell surface with
the apertures of pores along the grooves (arrows). B-F. TEM cross sections of the cortex with the plasma
membrane, the dilated inner membrane complex and the subpellicular microtubules under the epicytic bulges
(C). In cross section, each microtubule is surrounded by a white hexagonal area. Ectoplasmic organelles in the
grooves, connected to the cortical membranes via the imc, contain lamellar structures (arrow in B) or dense
material (white arrow in D). These organelles form an annular ring in cross section parallel to the cell surface
(white arrow in E) corresponding to the cross section of a micropore or myelin-like structures (F).
longitudinal networks of the subpellicular microtubules
(Fig. 2D).
Conoid and Myzocytosis
The conoid of S. pendula is a truncated cone
of about 225 nm height, with apical and distal
diameters about 260 nm and 1 ␮m respectively
(Fig. 4A-C). In TEM cross sections, the diameter
of filaments is about 23-32 nm; 9 sections are well
identified in one side of the conoid, while only an
opaque layer can be observed on the other side due
to their spiral organization (Fig. 4B-C). This structure
is quite similar to the well-described conoid of
Toxoplasma gondii (Hu et al. 2002, 2006) but the
apical polar ring is not present in the distal part of
S. pendula mucron and the preconoidal rings are
not clearly identified in its apical part but a dilatation
of the imc and the ends of the subpellicular microtubules
are unambiguously demonstrated (Fig. 4A,
Selenidium pendula: Ultrastructure and Phylogeny 345
Figure 4. Conoid in the Selenidium pendula mucron (A-C). Abbreviations: dense granule (DG), conoid (Co),
food vacuole (FV), food vacuole membrane (fvm), inner membrane complex (imc), host intestinal epithelium
(IE), parasite plasma membrane (pm). A-B. Two longitudinal sections of the S. pendula mucron, showing the
conoid structure and the opening, allowing a contact between the fvm and the host cell, visible in A. C. High
magnification showing the 9 cross sections of the microtubular network forming the conoid.
346 J. Schrével et al.
C, white arrows). This imc dilatation could correspond
to a site of a Microtubule Organizing Center
(MTOC) able to generate the subpellicular microtubules
since abundant bundles are found in the
anterior area of the trophozoite (Fig. 2D). In few
TEM cross sections, dense structures corresponding
to the neck of the rhoptries are observed inside
the conoid (Fig. 2C).
Myzocytosis, the predatory mode of nutrition
characteristic of archigregarines, is clearly illustrated
in S. pendula with food vacuoles inserted
inside the conoid (Figs 2C, 4A-B). In the axis of the
mucron, one or several clear food vacuoles, likely
formed via the conoid, are present (Fig. 2C). These
food vacuoles are surrounded by many rhoptries
and micronemes, two apical organelles characteristic
of zoites (Figs 1B, 2C). As shown by the
continuity of the food vacuole membrane up to its
contact with the host epithelial cell, an evagination
process through the apex of the conoid has
occurred, allowing the parasite to suck out the nutriments
from the host. This myzocytosis process
starts at the top of the conoid (Fig. 4A). The food
vacuoles are large, reaching sometimes up to 7 ␮m,
and several additional food vacuoles of about 2 or
3 ␮m are observed in the axis of the trophozoite
(Figs 2C, 5A-B). The lumen of the food vacuoles
has a low electron-dense aspect with some vesicles
and the membrane of the food vacuole exhibits
a very irregular border with numerous digitations.
Vital staining with low concentrations of neutral
red (1 0
/00) allowed to visualize large vacuoles of
about 4x2 ␮m located in the apex of the S. pendula
trophozoite with several small vesicles (data
not shown). This observation is in agreement with a
fragmentation of the initial food vacuole into numerous
vacuoles present in the anterior part of the
trophozoite (Fig. 5A).
Rhoptries, Micronemes, and Intrareticular
Granules in Trophozoites
In addition to the conoid, the apical end of S.
pendula trophozoites exhibits about 8-10 rhoptries
corresponding to the long, electron-dense
club-shaped, tubular or saccular organelles. They
appear in the trophozoite as cylindrical organelles
reaching up to 6 ␮m in length, with a diameter of
0.3-0.4 ␮m in the basal bulbous. At the apex, a
rhoptry neck could be observed. The rhoptry orientation
usually follows the direction of the conoid. In
some cases, the rhoptry neck penetrates the conoid
(Fig. 2C).
The rough endoplasmic reticulum (RER) and the
Golgi apparatus of S. pendula show an original
association between the swollen cisternae containing
numerous intrareticular granules of about
0.5-1 ␮m and the first saccule of the cis-region
of the Golgi apparatus (Fig. 6D). Similar associations
are observed in S. hollandei (Vivier and
Schrével 1966) but not in Selenidiidae species
parasitizing Cirratulidae (Schrével 1971a), Serpulidae
(Leander 2006), Terebellidae (Wakeman et al.
2014) or Sipunculida (Simdyanov and Kuvardina
2007, Leander 2006). Some micrographs show an
accumulation of numerous micronemes close to the
nuclear envelope (Fig. 7D) or to the Golgi apparatus
(Fig. 6C) with annular sections likely corresponding
to the neck of micronemes (Fig. 6B). The relation
of these RER-Golgi apparatus to the biogenesis of
the rhoptries and/or the micronemes is not clear,
since numerous micronemes are mixed with large
rhoptries (Fig. 6A).
Nucleus and the Perinuclear Cytoplasm
The ovoid nucleus of the S. pendula trophozoite is
characterized by the presence of a large spherical
nucleolus of about 4-5 ␮m in diameter (Fig. 7A).
No accumulation of chromatin is observed in
the nucleoplasm and the nuclear envelope lacks
the nuclear lamina as observed in S. hollandei
(Schrével 1971a; Vivier 1967). The nuclear envelope,
typically comprising two membranes, is rich in
nuclear pores (about 5 per ␮m) regularly distributed
all over the entire nuclear surface (Fig. 7C). In tangential
sections, the pores appear as rings of about
100-110 nm in their largest diameter with the presence
of a central particle of about 10 nm in diameter
(Fig. 7C).
The periphery of the nucleus exhibits a special
cytoplasmic area comprising a regular, 0.5 ␮m thick
fibrillar zone, lacking any organelle, and surrounding
the nucleus in a distance of 1.5-2 ␮m from the
nuclear envelope (Fig. 7A-B, E). This fibrillar zone
corresponds to the axial ducts described in living
cells (Schrével 1970).
Apicoplast-like Organelles
In the trophozoite of S. pendula, organelles with
four membranes are frequently observed (Fig. 5C)
and they appear morphologically similar to the
apicoplast of Toxoplasma and Plasmodium (Lim
and McFadden 2010) with some dense structures
(Fig. 5D) or in contact with multilamellar organelle
(Fig. 5E).
Selenidium pendula: Ultrastructure and Phylogeny 347
Figure 5. Food vacuoles and rhoptries in the apex of the S. pendula trophozoite (A-B), and apicoplast-like
organelles (C, D, E). Abbreviations: conoid (Co), food vacuole (FV), host intestinal epithelium (IE), microneme
(mn), mitochondrion (M), fragmented food vacuoles similar to pinocytotic vesicles (pv), rhoptry (R). A. TEM
cross section with the initial food vacuole passing through the conoid and the fragmented food vacuoles similar
to the pinocytotic vesicles (pv) observed in S. hollandei (Schrével 1968). Numerous rhoptries are accumulated
around these food vacuoles. B. Another cross section showing the irregular shapes of the initial food vacuole and
the intravacuolar vesicles. (C-E). Apicoplast-like organelles, characterized by the presence of four membranes
morphologically similar to the apicoplast of Toxoplasma and Plasmodium.
Nuclear Multiplication During the Syzygy
and Young Gamonts
The sexual phase of the S. pendula life cycle starts
with the syzygy, characterized by the pairing of
two haploid trophozoites, now called gamonts: one
male and one female. During the young syzygy
stage of S. pendula, the two gamonts are linked
by their posterior parts, while their pendular motility
continues with waves starting from the apex to the
posterior end (Schrével 1970).
In TEM, each gamont exhibits a similar intracellular
organization with a nucleus of about 20 ␮m in
diameter containing a spherical nucleolus of about
5 ␮m in diameter (Supplementary Material 1). In
each nucleolus, several clear areas are observed
with sizes varying from 0.3 to 1 ␮m in diameter
(Supplementary Material 1, arrows). The cell
348 J. Schrével et al.
Figure 6. Rhoptries, micronemes and Golgi apparatus (A-D). Abbreviations: amylopectin granule (am), Golgi
apparatus (Go), intrareticular granule (ig), microneme (mn), mitochondrion (M), rhoptry (R). A-B. TEM cross
sections of an accumulation of rhoptries and micronemes within the cytoplasm. The micronemes appear as
long-necked bottles, the necks appear as dense rings in cross sections (white thick arrow in B). C-D. Golgi
apparatus and mitochondrion occur close to the micronemes; the cis-region of the Golgi apparatus usually
contains numerous intrareticular granules (D).
Selenidium pendula: Ultrastructure and Phylogeny 349
Figure 7. Nuclear area of Selenidium pendula trophozoite (A-E). Abbreviations: amylopectin granule (am),
intrareticular granule (ig), microneme (mn), nuclear pores (np), nucleus (N), nucleolus (nu). A-B. TEM cross
sections of the nucleus containing a spherical nucleolus (A) and surrounded by the regular fibrillar zone without
organelles (white arrows in B). This fibrillar area is delimited by large vesicles of the rough endoplasmic reticular
containing numerous granules and amylopectin granules. C. Tangential section of the nuclear envelope exhibits
numerous pores. D. Occasional accumulation of micronemes can be observed near the nucleus. E. Higher
magnification of the micronemes and intrareticular granules.
350 J. Schrével et al.
surface and the cytoplasm of the two gamonts
also exhibit a similar organization (Supplementary
Material 1).
A clear characteristic of archigregarines belonging
to the family Selenidiidae is the early nuclear
multiplication within the two gamonts at the site
corresponding to the initial trophozoite nucleus that
occurs before the encystment of the gamonts. The
localization of the nuclei at the initial site of the
trophozoite nucleus is clearly shown by the DAPI
staining highlighting the DNA-containing structures
(Fig. 8A-B). Bright spots are observed inside spherical
structures, each of them corresponding to a
nucleus. In about two hours, the pendular motility
of each gamont is progressively reduced and
cyst formation occurs with a widening of the nuclear
zone in the gamont’s median plane (Fig. 8A-B). In
TEM, the concentration of the nuclei at this stage
is not easy to observe due to the relatively high
rate of this process. In favourable cross sections,
the nuclei are observed in the central area of the
gamont and before the secretion of the cyst wall.
Each spherical nucleus is about 5 ␮m in diameter
(Fig. 8C). From this central site, the nuclei migrate
to the periphery of each gamont while the cyst wall
is forming (Fig. 8D). In many nuclei of the gamonts,
centrocones and other stages of cryptomitosis were
detected.
Centrocones and Cryptomitosis in
Gamonts
The mitosis in S. pendula gamonts is a closedmitosis,
also called cryptomitosis, with the persistence
of the nuclear envelope as observed in all
Apicomplexa (Francia and Striepen 2014). All the
nuclei of the S. pendula gamonts are spherical with
a diameter of about 5 ␮m and many are associated
to a cupule with microtubules radiating from
the Microtubule Organizing Center (MTOC) in order
to form half-spindles (Fig. 9A-B). The chromatin is
localized all around the internal face of the nuclear
envelope as shown in TEM images (Fig. 9A-B). This
chromatin forms an electron dense filamentous network
with spotty dark nodes and in some cases, an
important dense accumulation is observed inside
the nucleoplasm (Fig. 9A). This dense accumulation
of at least 1 ␮m could correspond to the bright
spots visualized by the DAPI-staining (Fig. 8B).
The distribution of chromatin in S. pendula nucleus
appears as a continuous filamentous network quite
similar to the model of Apicomplexa cryptomitosis
proposed by Francia and Striepen (2014).
In S. pendula gamonts, many nuclei exhibit a
centrocone resulting probably from the high rate
of nuclear divisions since the chronology from
the syzygy to the encystment of the gamonts
represents only 2-3 hours (Schrével 1970). The
centrocone depends upon the MTOC that appears
as an electron dense annular structure of about
200 nm in diameter. From the MTOC, microtubules
radiate and form a half-spindle that pushes the
nuclear envelope without penetration in the nucleus
(Fig. 9A). The resulting cupule exhibits an outer
diameter of about 1.6-1.9 ␮m and the distance
from the MTOC to the inner border of the cupule
is about 1.4-1.6 ␮m. This typical centrocone can
duplicate and the second centrocone migrates to
the opposite direction of the initial cupule (Fig. 9B).
Micrographs with two centrocones are rather rare
and an intranuclear spindle was not observed most
likely due to the high rate of the progamic nuclear
division in S. pendula.
As the progamic nuclei migrate from the central
part of the gamont to the periphery, the
cryptomitosis continues after this migration, since
the duplication of the centrocones is observed
in the border of the cyst where the wall is
secreted.
Modifications of the Gamont Cell Surface
and Secretion of the Gametocyst Wall
When the gametocyst wall is forming, the cortical
membranes of each gamont are strongly modified
(Fig. 10A-C). The plasma membrane is always
present but the imc is disorganized with a series of
folds and clear dissociation from the plasma membrane
(Fig. 10B-C). In TEM, the gametocyst wall
exhibits two major layers, a homogeneous internal
layer of about 500-700 nm in thickness and a fuzzy
external layer with long filaments reaching about
300 nm. The total thickness of the gametocyst wall
at the beginning of gamogony is about 1 ␮m. The
secretion of this wall is the result of two types of
vesicles, one with rather electron dense components
(vesicle 1) and the second with a network
of very spotty filaments (Fig. 10A). The mechanism
of discharge of these two types of vesicles was not
clearly observed. As the gametocyst wall formation
occurs only two hours after the early syzygy step,
the secretion is probably the result of accumulations
of numerous intrareticular granules in the cisterns of
the rough reticulum endoplasm that represent storage
material for this process (Fig. 6D). However, a
potential dual function of the RER-Golgi apparatus
for both the formation of rhoptries and micronemes
and the storage of material for gametocyst formation
needs further investigations (Fig. 6A).
Selenidium pendula: Ultrastructure and Phylogeny 351
Figure 8. Nuclear development during the syzygy of Selenidium pendula (A-D). Abbreviations: nucleus (N),
cyst wall (CW). A-B. Fluorescence staining with DAPI showing nuclei in the median plane of each gamont
corresponding to the initial position of the nucleus at the beginning of the syzygy stage. Their numbers are
quite similar in the two gamonts and some bright spots are observed in some nuclei (B). C-D. TEM cross
sections in an early cyst where the nuclei are accumulating in the central position of the gamont while the cell
wall is not secreted (C) and later after the secretion of the cyst wall where the nuclei migrate to the gamont
periphery (D).
The Gametocyst and the Sporocyst Walls
The gametogenesis is a fast process in S. pendula,
lasting about one hour (Schrével 1970). After
the series of progamic nuclear divisions yielding
syncytium nuclei in the same gametocyst, cellularization
occurs, producing flagellated male gametes
and female gametes without flagellum. In TEM, the
gametocyst wall is more compact with dense layers
(Fig. 11A-B). The fuzzy coat observed at the beginning
of the gamogony is now very irregular in width
and the internal homogenous layers are more electron
dense (Fig. 11A). In some cases the internal
layers show a regular opaque layer of 0.3 ␮m and
an irregular homogenous layer with a lower electron
density (Fig. 11A).
In cross sections, the flagellar axoneme of the
male gamete of S. pendula exhibits a 9+0 pattern
(Fig. 11B). After fecundation, the life cycle
moves into the sporogony phase with the formation
of sporocysts corresponding to the evolution of
the zygotes toward the sporozoite formation inside
each sporocyst. A new secretion process occurs
around this sporocyst (Fig. 11C). The thickness of
the sporocyst wall is about 0.1 ␮m with small thin
spine-like digitations of about 0.2 ␮m (Fig. 11C).
352 J. Schrével et al.
Figure 9. Centrocones and mitosis stages during the gametogenesis of Selenidium pendula (A-B). Abbreviations:
centrocone (CC), chromatin (Ch), cyst wall (CW), dense layer (dl), filamentous layer (fl), nuclear envelope
(en), mitochondrion (M), microtubule-organizing center (MTOC), microtubule (mt), nucleus (N), vesicle type 1
(V1) and type 2 (V2). A. Early gametogenesis stage before the secretion of the gametocyst wall exhibiting
centrocone where the microtubules radiate from the MTOC to the cupule of the nuclear envelope forming a
truncated cone. The chromatin covers the inner face of the nuclear envelope and a dense accumulation is
observed in the nucleus. B. A second centrocone migrating on the other side of the nucleus. No intranuclear
spindle is observed, the chromatin is attached to the persistent nuclear envelope. Two types of vesicles are
observed one with dense granules (V1) and a second one with filamentous material (V2). The gametocyst wall
exhibits an external filamentous layer and an internal dense layer.
Molecular Phylogenetic Analyses of the
SSU rDNA Sequence
Type species are important to build solid bridges
between molecular phylogenies and taxonomy.
A phylogenetic tree was constructed using 115
sequences including nine novel small subunit
(SSU) rDNA sequences (two sequences from S.
pendula, the type species for Selenidiidae, one
from S. hollandei, one from Lecudina pellucida,
the type species for Lecudinidae, and 5 from L.
tuzetae, all specimen isolated from host organisms
collected in the Roscoff area, France) and 106
previously published ones available from public
databases, taking into account all available data
for archigregarine species (Table 1). Sequences
known to produce extreme long branches in SSU
rDNA-based phylogenies, such as those of the
gregarines Trichotokara spp. and Pyxinia robusta,
were excluded from this analysis. Globally, the Maximum
Likelihood and the Bayesian tree topologies
were congruent (Fig. 12) and in good agreement
with recently published phylogenies (Wakeman and
Leander 2013; Wakeman et al. 2014). The two
early lineages emerging among Apicomplexa were
from marine gregarines with archigregarines and
eugregarines. Interestingly the phylogenetic position
of the type species Lecudina pellucida (Fig. 12)
fell within the Lecudinidae, with a good support
with lecudinids of tunicates represented by the
Lankesteria genus. In the terrestrial gregarines, the
Gregarina lineage belongs to rather old insects
such as Coleoptera, Blattaria, and Orthoptera, in
contrast to the Ascogregarina lineage that infects
more recent insects according to the most recent
knowledge on insect evolution (Misof et al. 2014).
An analysis of the SSU rDNA sequences
clearly demonstrated the paraphyly of Selenidiidae,
which are split into three major groups
(Fig. 13, Supplementary Material 2-4). The type
species Selenidium pendula is closely related to
Selenidium boccardiella (Wakeman and Leander
2013). These two gregarines infect members
of the Spionidae family of Polychaeta. Similarly,
Selenidium pendula: Ultrastructure and Phylogeny 353
Figure 10. Reorganisation of the cortical membranes in Selenidium pendula gamonts and ultrastructure of
the gametocyst wall during the gametogenesis stage (A-D). Abbreviations: gametocyst wall (CW), dense layer
(dl), epicytic folds (ef), filamentous layer (fl), inner membrane complex (imc), plasma membrane (pm), vesicle
type 1 (V1) and type 2 (V2). A-C. TEM cross sections of the gamont’s periphery where the epicytic folds are
disorganized with dissociation of the inner member complex under the plasma membrane. The wall of the
gametocyst exhibits a filamentous external layer and a more homogenous internal layer; the vesicles of type 2
are probably involved in the cyst wall construction. D. Higher magnification of the cyst wall with large amount
of filaments attached to the surface of the internal homogenous layer.
S. hollandei is closely related to S. neosabellariae
and S. identhyrsae (Wakeman and Leander
2013), these three species being parasites of hosts
belonging to Sabellariidae. Parasites infecting Spionidae
and Sabellariidae diverged from 3.4 to
13.7% from each other (sequence identity, Supplementary
Material 2, 3).
Selenidium parasites of Terebellidae group form
a second divergent lineage with a wider global
divergence with true Selenidiidae of 25.8-28.2%,
354 J. Schrével et al.
Figure 11. Gamonts with gametes and young sporocysts after the fertilization (A-C). Abbreviations: amylopectin
granule (am), axoneme (Ax), gametocyst wall (CW), dense granule (DG), dense layer (dl), filamentous
layer (fl), mitochondrion (M), nucleus (N), sporocyst wall (SW). A. Gametocyst wall after the formation of the
gametes exhibits a third layer with more dense material under the two layers observed in more early gamont
stages (Figure 12). Residual amylopectin and dense granules are observed between the gametes and the
gametocyst wall. B. Cross sections of flagellar axonemes indicate a male gamete (B) and two serial sections
are of a 9+0 pattern. C. After the fertilization process the young sporocysts are surrounded by a thin wall
covered by very small spines.
Selenidium pendula: Ultrastructure and Phylogeny 355
(Supplementary Material 2, 3). Finally, all Selenidiidae
described in Phascolosoma formed a third
group which is the most divergent (26.3 - 28.8% of
divergence with the two precedent groups, Supplementary
Material 2, 3).
Discussion
Selenidium spp. and archigregarines
Since 2003, the morphology of some trophozoites
of Selenidiidae and related archigregarines
was investigated using SEM and more than 25
SSU rDNA sequences were deposited in the
GenBank/EMBL/DDBJ databases (Leander 2006,
2007b; Leander et al. 2003a; Rueckert and
Leander 2009; Wakeman and Leander 2012, 2013;
Wakeman et al. 2014). However, data on sexual
stages (gamogony and sporogony) were missing.
By combining electron microscopic descriptions
with phylogenies using SSU rDNA sequence data,
new Selenidium species have been proposed
such as S. pisinnus Rueckert and Leander, 2009,
S. boccardiella Wakeman and Leander, 2012,
S. idanthyrsae Wakeman and Leander, 2012, S.
neosabellariae Wakeman and Leander, 2013, S.
sensimae Wakeman and Leander, 2013 and S.
melongena Wakeman et al., 2014. A new genus
Platyproteum (Rueckert and Leander 2009) was
erected to replace the former species Selenidium
vivax Gunderson and Small 1986. A new enigmatic
genus related to archigregarines was also
proposed as Veloxidium (Wakeman and Leander
2013). In their discussion, Leander and co-workers
produced a comparative table with morphological
data of all Selenidiidae described in the
last century (Ray 1930; Schrével 1970, 1971b).
Few mistakes within Selenidiidae were reported
in this table (Table 2 in Wakeman and Leander
2012) as for example the mention of S. spionis,
presented as a parasite of Polyrabdina spionis.
Polyrhabdina spionis is in fact a lecudinid
gregarine and the host of S. spionis is the polychaete
Scolelepis fuliginosa Claparède, 1870 now
called Malacoceros fuliginosus (Claparède, 1870)
Schrével and Desportes, 2013. However, the SSU
rDNA sequences analysis of S. spionis revealed
lineages inside archigregarines and Leander and
co-workers underlined the importance of future
work on additional Selenidium-like gregarines
especially the type species S. pendula (Wakeman
et al. 2014). This current work on the type species
S. pendula Giard, 1884 and on S. hollandei Vivier
and Schrével, 1966 therefore enlighten with less
ambiguities parts of the evolutionary history of
archigregarines.
The SSU rDNA sequence phylogeny trees with
the different SSU rDNA sequences of archigregarines
(Table 1), show three clearly delimited
lineages among Selenidiidae (Figs 12-13, Supplementary
Material 3, 4). A major group corresponds
to the true-Selenidium lineage, for which sexual
stages (syzygy to sporocyst) have been
described. Its members are parasites of Sedentaria
polychaetes, such as S. pendula that
infects the Spionidae family, S. hollandei infecting
Sabellariidae and Selenidium cf. meslini infecting
Sabellidae. These true-Selenidium share common
important features, such as a nuclear multiplication
during the syzygy, the gamogony and the
sporocysts with usually four sporozoites. Many
archigregarines have developed atypical variations
in their cell morphology and their motility from pendular
to rolling type, with subpellicular microtubule
sets under the inner membrane complex (imc), but
without the gliding type observed in eugregarines.
Trophozoites of true-Selenidium exhibit a threemembrane
cortex where the imc forms a complete
envelope underlying the plasma membrane, with
sets of longitudinal subpellicular microtubules running
under the large folds designated as bulges
(Schrével 1970a, 1971a; Schrével et al. 2013;
Vivier and Schrével 1964). The grooves correspond
to the striations well described this last century by
light microscopic (Brasil 1907; Ray 1930; Schrével
1970). The cytoplasm beneath the grooves is
devoid of microtubules but exhibits micropores and
residual membranous organelles in connection with
the imc (Schrével et al. 2013; Vivier and Schrével
1964). These parasites feed by myzocytosis using
the conoid located at the apex of the trophozoite
(Schrével 1968; Simdyanov and Kuvardina 2007;
this work).
In this true-Selenidium lineage, the sexual stage
starts by the syzygy where the formation of progametic
nuclei is observed inside the gamont nucleus
before encystment. This observation in the type
species S. pendula (Fig. 8) is the confirmation of
histological previous descriptions by Caullery and
Mesnil (1900), Ray (1930), Reed (1933), Tuzet
and Ormières (1958) and in vivo observations by
Schrével (1970). This gamogony is quite different
from all other eugregarines where the first
gamogony division starts inside the cyst and is
followed by successive series of nuclear divisions
called progamic mitoses without cytokinesis such
as in Lecudina tuzetae (Kuriyama et al. 2005).
So, the Lecudina type gamogony produces a syncytium
until the cellularization process yielding the
356 J. Schrével et al.
Coleoptera
Diptera
Lepidoptera
Hymenoptera
Odonata, Hemiptera
Coleoptera, Lepismatids
Capitellid Polychaeta
Blattaria, Orthoptera
Lecudinids
from Polychaeta
Lecudinids
from Tunicates
Urosporids
Incertae sedis
True Selenidiidae (Lineage I)
Selenidiidae from Sipunculida (III)
Selenidiidae from Terebellids (II)
Eugregarines
Marine
gregarines
Hexapod
gregarines
Heterokonts
Colponemids
Ciliates
Dinoflagellates
Perkinsids
Colpodellids
Chromerids
Hematozoans
Coccidians
Cryptosporidians
Archigregarines
Crustacean
gregarines
Marine
gregarines
Marine
gregarines
Rhizaria
Hexapod
gregarines
Gregarina niphandrodes FJ459747
Gregarina polymorpha FJ459748
Selenidium hollandei LN901445
Ascogregarina sp ex Ochlerotatus japonicus DQ462458
Ascogregarina armigerei DQ462459
Ascogregarina culicis from Vietnam DQ462457
Ascogregarina taiwanensis from Japan DQ462454
Paraschneideria metamorphosa FJ459755
Mattesia geminata AY334568
Ophryocystis elektroscirrha AF129883
Apicystis bombi FN546182
Psychodiella chagasi FJ865354
Geneiorhynchus manifestus FJ459739
Hoplorhynchus acanthatholius FJ459750
Syncystis mirabilis DQ176427
Prismatospora evansi FJ459756
Stylocephalus giganteus FJ459761
Colepismatophila watsonae FJ459738
Polyplicarum citrusae JX535336
Uncultured eukaryote CCA38 AY179975
Polyplicarum curvarae JX535340
Polyplicarum translucidae JX535348
Polyplicarum lacrimae JX535344
Uncultured eukaryote clone DSGM13 AB275013
Uncultured alveolate clone AT416 AF530523
Ammonia beccarii U07937
Gregarina kingi FJ459746
Blabericola haasi FJ459753
Blabericola migratory FJ459754
Gregarina blattarum FJ459741
Gregarina tropica FJ459749
Amoebogregarina nigra FJ459737
Gregarina coronata FJ459743
Gregarina diabrotica FJ459745
Gregarina sp Phaedon JF412715
Gregarina cloptoni FJ459742
Ganymedes sp SR2010 FJ976721
Thiriotia pugettiae HQ876006
Heliospora cf longissima HQ891115.2
Lecudina tuzetae Roscoff 2014b LN901450
Lecudina tuzetae Roscoff 2013a LN901447
Lecudina tuzetae Roscoff 2014a LN901449
Lecudina tuzetae Roscoff 2013b LN901448
Lecudina tuzetae Roscoff 2012 LN901446
Lecudina tuzetae AF457128
Lecudina pellucida LN901442
Lankesteria abbotti DQ093796
Lankesteria ascidiae JX187607
Lankesteria chelyosomae EU670240
Lankesteria cystodytae JF264840
Pterospora floridiensis DQ093794
Difficilina paranemertes FJ832159
Difficilina tubulani FJ832160
Uncultured eukaryote DSGM6 AB275006
Pterospora schizosoma DQ093793
Lithocystis sp DQ093795
Veloxidium leptosynaptae JN857966
Selenidium sp1 KCW2013 KC110863
Selenidium serpulae DQ683562
Selenidium sensimae KC110869
Selenidium sp2 KCW2013 KC110864
Selenidium cf echinatum KC110874
Selenidium pendula LN901444
Selenidium cf mesnili JN857968
Selenidium boccardiellae JN857969
Selenidium pendula LN901443
Selenidium identhyrsae JN857967
Selenidium neosabellariae KC110871
Uncultured alveolate clone BAQA40 AF372776
Selenidium orientale FJ832161
Selenidium pisinnus FJ832162
Selenidium melongena 2 KC890800
Selenidium terebellae 2 KC890804
Cryptosporidium serpentis AF151376
Cryptosporidium baileyi AJ276096
Cryptosporidium parvum AB089290
Goussia desseri GU479641
Eimeria tenella AF026388
Toxoplasma gondii M97703
Besnoitia besnoiti AY833646
Babesia microti strain RI gi 399217317
Babesia bigemina JQ437264
Theileria parva L02366
Theileria orientalis AB520957
Colpodella pontica AY078092
Colpodella tetrahymenae AF330214
Chromera velia strain CMS22 DQ174731
Colpodella edax AY234843
Alveolata sp CCMP3155 HM245049
Parvilucifera infectans KF359485
Parvilucifera rostrata KF359483
Parvilucifera prorocentri FJ424512
Perkinsus atlanticus KF359485
Duboscquella sp AB295041
Ichthyodinium chabelardi FJ440623
Uncultured marine dinoflagellate clone Ma131 1A49 FJ0326782
Amoebophrya RCC1626 HQ658161
Syndinium turbo DQ146404
Hematodinium perezi EF065717
Gonyaulax polyedra AF377944
Alexandrium catenella AY347308
Pfiesteria piscicida DQ991382
Akashiwo sanguinea AY831412
Prorocentrum micans AY803739
Stentor coeruleus AF357145
Paramecium tetraurelia AF149979
Sterkiella histriomuscorum FJ545743
Colponema edaphicum KF651068
Colponema sp Vietnam KF651083
Chrysolepidomonas dendrolepidota AF123297
Cylindrotheca closterium DQ082742
Hyphochytrium catenoides X80344
Phytophthora megasperma M54938
Bigelowiella sp U02075
//
0.2
x3
97/0.9
91/0.9
94/0.9
82/1
97/0.9
77/0.6
75/0.6
60/0.9
82/0.9
100/0.8
93/0.8
72/0.8
79/0.9
98/1
67/1
97/1
98/1
98/0.9
92/1
98/1
80/0.5
96/0.7
83/1
42/0.9
54/0.7
89/1
39/0.6
92/0.9
90/1
100/0.8
59/0.9
48/1
43/0.8
98/1
44/1
44/1
72/1
89/1
97/1
47/0.7
49/1
98/1
58/1
57/1
97/1
Cephaloidophora cf communis HQ891113
Selenidium pendula: Ultrastructure and Phylogeny 357
60
79
51
100
Selenidium sp1 KCW2013 Isolate 1 KC110863
Selenidium sp2 KCW2013 Isolate 2 KC110868
Selenidium sp2 KCW2013 Isolate 3 KC110864
Selenidium sp2 KCW2013 Isolate 1 KC110865
Selenidium sp1 KCW2013 Isolate 2 KC110866
Selenidium sp1 KCW2013 Isolate 3 KC110867
Selenidium serpulae DQ683562
Selenidium sensimae KC110869
Selenidium sensimae KC110870
Selenidium cf mesnili JN857968
Selenidium neosabellariae KC110873
Selenidium neosabellariae KC110872
Selenidium neosabellariae KC110871
Selenidium identhryrsae JN857967
Selenidium pendula LN901443
Selenidium cf echinatum KC110874
Selenidium hollandei LN901445
Selenidium cf echinatum KC110875
Selenidium pendula LN901444
Selenidium boccardiella JN857969
0.02
100
100
78
78
100
100
100
100
82
100
65
Spionidae
Serpulidae
Sabellidae
Sabellaridae
Cirratulidae
100
Selenidium pendula LN901443
Selenidium pendula LN901444
Selenidium hollandei LN901445
100
Figure 13. Molecular phylogenetic analysis by Maximum Likelihood method of Selenidiidae lineage retrieved
from polychaete annelids (host families in bold black).
The evolutionary history was inferred by using the Maximum Likelihood method based on the General Time
Reversible model (Nei and Kumar 2000). The tree with the highest log likelihood (-5446.5092) is shown. A
discrete Gamma distribution was used to model evolutionary rate differences among sites (5 categories with
gamma parameter = 0.2711). The rate variation model allowed for some sites to be evolutionarily invariable
(+I), 39.8364% sites). The tree is drawn to scale, with branch lengths measured in the number of substitutions
per site. Novel sequences are highlighted in grey boxes.
gametes. Another clear difference concerns the
degree of condensation of the chromatin with a
continuous filamentous network attached to the
nuclear envelope all around the nucleus in S.
pendula cryptomitosis. Chromosome condensation
does not seem to occur in S. pendula in contrast to
cryptomitosis of L. tuzetae (Kuriyama et al. 2005)
and Grebnickiella gracilis (Moblon-Noblot 1980).
Sporogony then leads to spherical sporocysts that
differentiate usually into four sporozoites per sporocyst
(Ray 1930; Schrével 1970).
Other Selenidium-like species infecting sipunculids
and Terebellidae are only known through
their trophozoites and their localization within
hosts (Leander 2006; Wakeman et al. 2014).
The intestinal trophozoite of S. terebellae Ray
1930 exhibits large bulges but differences with the
true-Selenidium have been observed. As an example,
a regular layer of about 30-33 nm in thickness
(Supplementary Material 5 and Wakeman et al.
2014) similar to the internal lamina of eugregarines
(Schrével et al. 1983) or to some euglenoid cortex
(Mignot 1966) is attached to the imc. Numerous
sets of longitudinal subpellicular microtubules are
immediately under this regular dense layer and
many residual membranous organelles are highly
concentrated under the imc of the grooves (Supplementary
Material 5). S. melongena trophozoites
were described in the same host as S. terebellae,
but inside the coelom, an unusual localization
for archigregarines (Wakeman et al. 2014). The
cortex of S. melongena Exhibits 30-40 epicytic
folds helically arranged along the axis of the cell.
Surprisingly, although the subpellicular sets of
microtubules were not observed in TEM, a strong
fluorescent labelling of alpha-tubulin was detected
below the helical folds. S. melongena are nonmotile
without pendular or rolling motility nor gliding.
➛
Figure 12. Maximum Likelihood (ML) tree inferred on an alignment of 115 small subunit (SSU) rDNA sequences
corresponding to 9 Selenidium and Lecudina species from this current study (highlighted in grey boxes) and
106 sequences from diverse eukaryotes corresponding mostly to representatives of Alveolata with one Rhizaria
as outgroup. The Maximum Likelihood method is based on the General Time Reversible +G +I model (Nei and
Kumar 2000). The tree is drawn to scale, with branch lengths measured in the number of substitutions per site.
A branch was shortened by a multiple (3) of the length of substitutions/site scale bar. There were a total of
1153 positions in the final dataset. ML evolutionary analyses were conducted in MEGA6 (Tamura et al. 2013).
Numbers at the branches denote ML bootstrap percentage (first value). Bayesian posterior probabilities are
also indicated (second value). Black dots on branches denote bootstrap percentages above 99% and Bayesian
posterior probabilities superior to 0.97.
358J.Schréveletal.
Table 1. List of the SSU rDNA sequence numbers of archigregarines and Veloxidium initially included in this group and the references: 1. This
work; 2. Leander et al. 2003; 3. Leander et al. 2007; 4. Rueckert and Leander 2009; 5. Wakeman and Leander 2012; 6. Wakeman and Leander
2013; 7. Wakeman et al. 2014.
Archigregarines SSU rDNA
sequences
Ref. Host Infraclass Order Family
Selenidium pendula LG LN901443 1 Scolelepis squamata Canalipalpata Spionida Spionidae
Selenidium pendula IF LN901444 1 Scolelepis squamata Canalipalpata Spionida Spionidae
Selenidium boccardiella JN857969 5 Boccardiella ligerica Canalipalpata Spionida Spionidae
Selenidium mesnili JN857968 5 Myxicola infundibulum Canalipalpata Sabellida Sabellidae
Selenidium hollandei LN901445 1 Sabellaria alveolata Canalipalpata Sabellida Sabellariidae
Selenidium neosabellariae KC110871
KC110872
KC110873
6 Neosabellaria cementarium Canalipalpata Sabellida Sabellariidae
Selenidium identhyrsae JN857967 6 Idanthyrsus saxicavus Canalipalpata Sabellida Sabellariidae
Selenidium serpulae DQ683562 3 Serpula vermicularis Canalipalpata Sabellida Serpulidae
Selenidium sensimae KC110869
KC110870
6 Spirobranchus giganteus Canalipalpata Sabellida Serpulidae
Selenidium Sp1 KC110863
KC110866
KC110867
6 Spirobranchus giganteus Canalipalpata Sabellida Serpulidae
Selenidium Sp2 KC110864
KC110865
KC110868
6 Spirobranchus giganteus Canalipalpata Sabellida Serpulidae
Selenidium terebellae AY196709 2 Thelepus sp, Canalipalpata Terebellida Theleponidae
Selenidium terebellae KC890803
KC890804
KC890805
KC890806
7 Thelepus japonica Canalipalpata Terebellida Theleponidae
Selenidium melongena KC890799
KC890800
KC890801
KC890802
7 Thelepus japonica Canalipalpata Terebellida Terebellinae
Selenidium cf echinatum KC110874
KC110875
6 Dodecaceria concarum Canalipalpata Terebellida Cirratulidae
Selenidium vivax AF236097 2 Phascolosoma agassizii Sipunculida Phascolosimida Phascolosomatidae
Platyproteum vivax AY196708 4 Phascolosoma agassizii Sipunculida Phascolosimida Phascolosomatidae
Filipodium phascolosoma FJ832163 4 Phascolosoma agassizii Sipunculida Phascolosimida Phascolosomatidae
Selenidium pisinnus FJ832162 4 Phascolosoma agassizii Sipunculida Phascolosimida Phascolosomatidae
Selenidium orientale FJ832131 4 Themiste pyroidea Sipunculida Golfingiida Veloxidium leptosynaptae
Veloxidium leptosynaptae JN857966 5 Leptosynapta clarki Echinodermata Apodida Synaptidae
Selenidium pendula: Ultrastructure and Phylogeny 359
According to Wakeman et al. (2014), such atypical
cell organization of S. melongena trophozoites
seems to be closer to lecudinids than to Selenidiidae.
These original observations as well as the
lack of description of syzygy and sporocysts require
future work, especially to explain the way by which
S. melongena can infect its host.
The third Selenidium-like lineage described here
corresponds to a group of intestinal parasites of
Sipunculida. These parasites are mainly known
from SEM and TEM observations on Selenidium
vivax trophozoites (Leander 2006). Renaming of
S. vivax as Platyproteum vivax was supported by
archigregarine flat shape when observed under
TEM with important sets of longitudinal subpellicular
microtubules and numerous mitochondria
probably in relation to the very active plasticity of S.
vivax (Rueckert and Leander 2009). This cellular
organization appears similar to that of S. hollandei
(Schrével 1970). Here also, descriptions of the
syzygy with their characteristic progametic nuclei
as well as the sporocysts require clarifications. This
point is also important for Filipodium trophozoites
where numerous microvilli from 1.6-10 ␮m long and
about 0.15 ␮m in diameter were clearly described
in TEM (Hoshide and Todd 1996).
All archigregarines are intestinal parasites of
Annelida belonging to the clade Sedentaria except
one Selenidium metchnikovi reported in Hemichordata
(Léger and Duboscq 1917). In contrast, many
lecudinids are intestinal parasites of the clade
Errantia from Annelida (Schrével and Desportes
2013). This separation between archigregarines
and marine lecudinid eugregarines is probably
related to the different modes of living of their hosts.
For instance, lecudinids are adapted to the errant
and predatory life of Errantia while archigregarines
are adapted to sedentary life of Sedentaria with
microphage species living below stones, or as tube
builders or ingesting sediment as the representatives
of the family Spionidae or surface deposit
feeders with head appendages (Sabellidae, Sabellariidae).
Evolutionary history of Annelida is still
poorly understood as the classic morphological
cladistic analysis with a monophyletic Polychaeta
(Rouse and Fauchald 1997) was challenged in
the light of the recent molecular evidences. Today,
Polychaeta are inferred to be paraphyletic with the
inclusion of the Clitella (earthworms) and the nonsegmented
taxa Echiura and Sipunculida (Struck
et al. 2011). Complexity of the phylogeny of Selenidium
species may reflect the one of their hosts.
The true-Selenidium lineage within the Selenidiidae
family likely forms the core of archigregarines while
the two other distant lineages infecting respectively
the Sipunculids and Terebellids orders, could be
considered as related Selenidium-like lineages.
These results, deduced from molecular phylogeny
analyses need to be confirmed at the biological and
cellular levels but are crucial since they open new
trends in evolutionary history among Apicomplexa.
The enigmatic Veloxidium leptosynaptae, initially
placed within archigregarines after phylogenetic
analyses (Wakeman and Leander 2012) was
later included within lecudinids and urosporids
(Wakeman et al. 2014). In our phylogenetic studies,
it also groups with lecudinids and urosporids
with strong supports (Fig. 12).
Apicoplasts, Conoid, MTOC and
Rhoptries are Major Cell Structures in the
Evolution of Apicomplexa
Gregarines represent interesting models to investigate
the evolution from free-living flagellated
alveolates status, likely photosynthetic, to obligatory
parasites among Apicomplexa.
In archigregarines, the presence of an apicoplast
remains an open question. Presence of
a functional plastid is reported in Chromera, a
free-living photosynthetic relative to Apicomplexa
(Lim and McFadden 2010). The apicoplast, a nonphotosynthetic
plastid of red algae origin, is well
documented in some Apicomplexa species such
as Plasmodium, Toxoplasma, Eimeria, Babesia,
Theileria. This relict plastid is limited by four
membranes indicating its secondary endosymbiont
origin. In the eugregarine Gregarina niphandrodes,
the apicoplast seems to be absent (Toso and
Omoto 2007). Here in S. pendula, apicoplast-like
organelles are regularly observed in trophozoites
at the ultrastructural level. Interestingly Ray (1930)
reported the visualization of a dark spot stained with
Heidenhain’s haematoxylin, associated to each
merozoite nucleus in S. mesnili parasitizing the
polychaete Myxicola infundibulum. Such an observation
at the light microscopy level was also
observed by TEM, revealing the presence of an
organelle with four membranes close to the anterior
part of each S. hollandei merozoite (Schrével
1971b).
The apical phagotrophy in the free-living predators
of alveolates, with open conoid and rhoptries,
may be at the origin of the anchoring device of
archigregarines like Selenidium, characterized by
their mucron and the myzocytosis function. The
conoid of S. pendula, similar to that of S. hollandei
(Schrével 1968) and S. orientale (Simdyanov and
Kuvardina 2007), is conserved in large trophozoites
and appears similar to the conoid of sporozoites
360 J. Schrével et al.
from eugregarines Stylocephalus africanus
(Desportes 1969) and Ascogregarina (Lankesteria)
culicis (Sheffield et al. 1971). Among Conoidasida,
the conoid of T. gondii is the most investigated
at the structural and molecular levels, with the
construction of unique coma-shaped tubulin sheets
to form a spiral cone-shaped structure (Hu et al.
2002, 2006). As S. pendula is the archigregarine
type species and an early branching Apicomplexa,
its conoid appears a good model to study the transition
between Apicomplexa with closed conoid and
free-living alveolate ancestors with open conoid,
as found in the early branching dinoflagellates as
Colpodella (Brugerolle 2002; Leander et al. 2003b)
or Psammosa pacifica (Okamoto and Keeling
2014).
Recently, the hypothesis of molecular links
between Apicomplexa and algal ancestors was
suggested with the demonstration of similar components
in the apical complex of Myzozoa and the
flagellar apparatus of protists. This hypothesis was
mainly supported by the localization of striated fiber
assemblies (Francia et al. 2012) and SAS-6 proteins
(de Leon et al. 2013). T. gondii striated fiber
assemblins (TgSFA2 and TgSFA3) proteins whose
orthologs are found in the rootlet associated with
the basal bodies from green algae, polymerize into
a dynamic fiber that emerges from the centrosomes
immediately after their duplication (Francia et al.
2012). Genetic experiments showed that the two
proteins TgSFA2 and 3 play an essential role in
the cell division of the T. gondii since cytokinesis
is blocked in their absence. This Tg SFA fiber thus
provides a robust spatial and temporal organizer
for the parasite cell division. Also, Francia et al.
(2012) indicated that other comparable SFA fibers
were observed in previous ultrastructural studies on
Eimeria (Dubremetz 1973, 1975) and Plasmodium
(Schrével et al. 2008).
The SAS-6 protein is well known in the centriolar
biogenesis of eukaryotes from protists to
vertebrates (Leidel et al. 2005; van Breugel et al.
2011). This protein was described in the centrocone
during T. gondii cryptomitosis (de Leon et al.
2013). In addition a novel SAS-6 like (SAS-6L) protein
family that shares an N-terminal domain with
SAS-6 but without the coiled-coil tails was localized
above the T. gondii conoid (de Leon et al.
2013). Genomic analyses showed that SAS-6L
is an ancient protein found in diverse eukaryotic
lineages: Trypanosoma, Leishmania, ciliates and
Apicomplexa (Hodges et al. 2010; de Leon et al.
2013). In Trypanosoma brucei trypomastigotes, the
Tb SAS-6L was observed near the basis of the
flagellum, consistent with the basal body location.
In T. gondii, the Tb SAS-6L antibody labelled the
apex of tachyzoites, and after conoid extrusion triggered
by ionomycin treatment, it labelled the tip of
the “true” conoid. The SAS-6L and SAS-6 antibodies
did not colocalize in T. gondii, the former one
labelling the centriole and the latter one labelling
the conoid tip (de Leon et al. 2013).
Complex connections between the “pseudoconoid”
or “incomplete conoid” and the flagellar
apparatus were also shown, by conventional TEM
and 3D reconstruction, in the apical complex of
Psammosa pacifica, a predator relative of apicomplexans
and early dinoflagellates (Okamoto and
Keeling 2014).
The MTOC of the centrocones of S. pendula
appears as a disc similar to that observed in other
eugregarines such as L. tuzetae where 9 singlets
could be detected in favourable TEM sections
(Kuriyama et al. 2005). From these MTOC discs,
microtubules radiated to form a cone involved in
the cup-shaped invaginations of the nuclear envelope.
The continuity of these MTOC during the life
cycle could be in agreement with a centriolar-like
structure since a 9+0 axonemal pattern is observed
in S. pendula male gamete (Fig. 11B). The question
of the subpellicular microtubule biogenesis is
not clear. The conoid is not, by itself, the MTOC
since it is absent in the zoites of Hematosporida
and Piroplasmida. The two polar rings, observed at
the apex of the Eimeria or Plasmodium zoites were
proposed as the MTOC sites generating the subpellicular
microtubules (Russel and Burns 1984),
but these two polar rings were not observed in S.
pendula. The imc dilatation at the border of the proximal
opening of the conoid could fulfil this function
(Fig. 4). The exceptional accumulation of microtubule
bundles in the anterior part of the mucron,
before the regular subpellicular microtubule sets of
the epicytic bulges (Fig. 2D), is in agreement with
the strong labelling of S. melongena apex with fluorescent
anti-alpha tubulin (Wakeman et al. 2014).
Biogenesis of these abundant microtubule bundles
needs further analysis.
Rhoptries are characteristic of the apicomplexan
zoites and also of the Selenidiidae trophozoites
(Schrével et al. 2013 for a review). Interestingly,
presence of numerous intracytoplasmic thread-like
bodies described by Ray (1930) in the apex of different
Selenidium trophozoites was visualized after
iron haematoxylin staining (Heidenhain’s haematoxylin).
By their sizes reaching 8-12 ␮m depending
on the Selenidium species and their localization,
these thread-like structures could correspond to
the rhoptries described from TEM such as in S.
pendula (Fig. 5A), S. hollandei (Schrével 1968)
Selenidium pendula: Ultrastructure and Phylogeny 361
and S. orientale (Symdyanov and Kurvidina 2007).
Ray (1930) considered these thread-like structures
as one of the morphological characters of each
Selenidium species, however the abundance of
rhoptries detected in TEM is in fact a general character
for archigregarines (Schrével et al. 2013,
for review). Biological functions of many apicomplexan
rhoptry proteins remain largely unknown.
In Plasmodium and Toxoplasma, the most investigated
apicomplexans at the molecular level, there
is growing evidence to suggest that the rhoptry neck
proteins are predominantly involved in host-cell
adhesion with some sharing evolutionary origins
among apicomplexans. In contrast, the rhoptry
bulb proteins appear mainly genus specific, suggesting
that they evolved secondarily to become
highly specific to their host cells (Counihan et al.
2013). In S. pendula, food vacuole membranes
may have arised from numerous rhoptries localized
within the apex. A strong membrane trafficking is
expected to produce the large and abundant food
vacuoles observed during myzocytosis (Fig. 4A).
Therefore Selenidium rhoptry proteins could play
a role in producing intracellular food vacuole in
contrast to Apicomplexa with an intracellular development,
where the rhoptry proteins seem involved
in the parasitophorous vacuole elaboration such as
in Plasmodium and Toxoplasma.
Archigregarines and Eugregarines: Two
Early Branching Lineages Among
Apicomplexa
The transition from the free-living alveolates
to apicomplexan parasites was supported by
comparative ultrastructural studies and molecular
phylogeny analyses of basal lineages, such
as dinoflagellates (together with perkinsids) and
apicomplexans (including colpodellids) (Leander
and Keeling 2003). The myzocytosis is the most
plesiomorphic features of apicomplexans with
archigregarines having a closed conoid (Schrével
1968, 1971b), and colpodellids the sister lineage
of Apicomplexa with an open conoid (Kuvardina
et al. 2002). In perkinsids, representing the earliest
diverging sister lineage of dinoflagellates
(Saldarriaga et al. 2003), an open conoid is also
observed (Perkins 1996). These three types of
parasites also share rhoptry-like organelles and,
together with their phylogenetic positions, they
confidently infer that a common ancestor of apicomplexans
and dinoflagellates had an apical complex
involved in the acquisition of nutrients from the cytoplasm
of prey cells (Leander and Keeling 2003).
Among the high diversity of gregarines in
invertebrates, Polychaeta, an animal class known
to be present at the Cambrian biodiversity explosion
and to represent one of the earliest Bilateria
organisms (De Rosa et al. 2005; Schrével and
Desportes 2013), is well infected by gregarines.
This situation supports the evolutionary prelude of
marine gregarines to the apicomplexan radiation
(Leander 2007). The initial archigregarine radiation
is supported by the “hypersporozoite” cell organization
of the trophozoite, the myzocytosis and
the pendular or rolling motility (Schrével 1971b;
Schrével and Desportes 2015). The subsequent
eugregarine radiation, with an adaptation to the
intestinal biome and an extracellular development,
could have emerged from intestinal lecudinid gregarines.
Here, their cell cortex is quite different
from archigregarines by the presence of numerous
epicytic folds, without the regular sets of subpellicular
microtubules but with a sophisticated
distribution of 12-nm filaments, apical rippled dense
structures at the top of the folds (Schrével et al.
1983;Vivier 1968). Their gliding motility depends
upon an actin-myosin system but the molecular
mechanochemical properties are far from being
understood (Heintzelman 2004; Valigurová et al.
2013). The myzocytosis, similar to the archigregarine
model, is not observed in these marine
eugregarines: their nutrition process is realized
through a bulbous attachment apparatus usually
designated by mucron.
The gregarine colonization of the coelom in
invertebrate hosts by transmigration of the sporozoites
through the intestinal epithelium and a
coelomic development reveal additional adaptations
of eugregarines to their host environment.
These adaptations are a significant evolutionary
step of marine gregarines as suggested by Leander
(2007a), and represent an antithesis to any notion
of “primitiveness”. One of the best evidence is
the unique adaptation of the coelomic eugregarine
Diplauxis hatti to its host Perinereis cultrifera where
a strict synchronization is observed between the
maturation of the polychaete gametes and the
sexual phases (gamogony and sporogony) of the
parasite (Prensier et al. 2008). This example illustrates
how gregarines are well adapted to their host
environment. For instance, D. hatti is adapted to
P. cultrifera but cannot invade other Nereidae host
as Hedistes (Nereis) diversicolor nor Nereis pelagica.
The extreme adaptation of some gregarines to
their host environments could explain some unexpected
situations such as the reduction observed
from the canonical 9+2 flagellar pattern, in the male
gametes, with a 9+0 pattern in S. pendula (this
362 J. Schrével et al.
study), 6+0 in L. tuzetae (Schrével and Besse 1975)
and 3+0 in D. hatti (Prensier et al. 1980). The 9+0
pattern of Selenidium, close to the 9+2 normality,
may be correlated to a fertilization phase lasting
about 1 hour in a 1-day sexual phase (gamogony
and sporogony), the 6+0 pattern of L. tuzetae, may
result from a fertilization realized in few hours within
a cyst, during a 3 days sexual phase of the Lecudina
life cycle (Schrével 1969). More impressively,
the 3+0 pattern in D. hatti could have been selected
over evolution because of the fertilization step lasting
only few hours in a highly extended complete
life cycle, lasting 2.5 years (Prensier et al. 2008).
Such evolutionary proposal, suggesting that each
gregarine develops its own programme according
to its environment is in agreement with the notion
of regressive evolution in microorganisms proposed
by Lwoff (1944). This type of regressive evolution
could probably continue with other coelomic gregarines
with the disappearance of the flagellum in
male gametes of Gonospora species as suggested
from histology studies (Schrével 1963; Trégouboff
1918). Expression of the own program of each
coelomic eugregarines is also observed with the
variations in their epicytic cell surface transformations
with digits, surface swelling in Pterospora,
microvillosities in Diplauxis or the development of
peristaltic motility instead of gliding, sometimes
a pendular motility is observed in young trophozoite
and peristaltic motility during the fast growing
period of the same trophozoite as observed in D.
hatti (see Schrével et al. 2013 for a review).
Conclusion
Molecular phylogenetic analyses of archigregarines
demonstrate that S. pendula, the type
species of archigregarines, belongs to a lineage
with a large number of Selenidium parasites
of Spionidae, Sabellaridae, Sabellidae, Cirratulidae
families of the Sedentaria Polychaeta. All
these Selenidium exhibit similar biological characters
such as the cell cortex with a plasma
membrane, imc (inner-membrane-complex) and
subpellicular microtubules, the apical complex with
a conoid, the myzocytosis with large food vacuoles
and abundance of large rhoptry organelles, the
nuclear multiplication during the syzygy and the
early gamonts. Two other Selenidium-like lineages
are observed in the Terebellidae and Sipunculida
where the sexual characters are not available at
this time. Such a status underlines an adaptation
of the family Selenidiidae to their host families and
this first early evolutive lineage could correspond
to the transition step between the free-living flagellated
alveolates and the Apicomplexa, before the
diversification of the marine eugregarines without
the typical myzocytosis realized through the conoid
but with a gliding motility.
Methods
Preparation of annelids and gregarines: Isolates of the gregarine
Selenidium pendula Giard, 1884 type species, were
collected from the intestine of the polychaete worm Scolelepis
squamata (O. F. Müller, 1806) (previously named Nerine cirratulus,
Delle Chiaje, 1831) on the French coast of the English
Channel at the “Station Biologique de Roscoff”, in 2007 then
again in 2012. Isolates of the gregarines Selenidium hollandei
Vivier and Schrével, 1966, Lecudina pellucida (Mingazzini,
1891) type species and different isolates of L. tuzetae Schrével,
1963 were also collected from the intestines of polychaete
worms from the same area, in 2007, 2012, 2013 and 2014
(Table 2).
After washing in seawater, each collected worm was kept,
at the laboratory temperature, in a separate Petri dish. The
medium (seawater) was changed daily. For long-term conservation,
the collected worms were rinsed with 0.22 ␮m filtered
seawater and stored at 4 ◦
C. In order to collect Selenidium pendula
Giard, 1884, the anterior part of the Scololelepis squamata
worms, with a yellow color, was discarded since the parasites
were always absent, then the worms were cut transversally
in series of segments of about 1 to 1.5 cm of length. To collect
S. hollandei, L. pellucida and L. tuzetae, a similar type of
microdissection was performed from their corresponding hosts,
under a classic binocular microscope, in order to expose the
intestinal epithelial surface to the seawater. In addition, and
only in the case of L. tuzetae, cysts excreted with feces were
collected from the Petri dishes of individually kept Neanthes
(Nereis) diversicolor (O. F. Müller, 1776). Trophozoites of S. pendula,
attached to the intestine, were easily detected, in spite of
their rather small sizes (usually 150 -180 ␮m x 30-35 ␮m), by
their white color - contrasting to the characteristic green color
of the intestinal epithelium of the worm - and by their active
pendular movements. In highly infected Scolelepis squamata,
trophozoites and sexual stages of S. pendula (syzygies and
young cysts) were also collected in Petri dishes, among the
gametes released from hosts during the dissection. S. hollandei
trophozoites were easily observed in host epithelium by their
very active rolling movements, immediately after sectioning the
post abdominal segment of their hosts, Sabellaria alveolata
Linnaeus, 1767.
Electron microscopy: For transmission electron
microscopy (TEM), intestinal epithelial tissues of Scolelepis
squamata highly infected with trophozoites of S. pendula were
collected and fixed in 5% (v/v) glutaraldehyde in 100-150 mM
phosphate or 0.2 M cacodylate buffer (pH 7.3), at 4 ◦
C, for
6 to 12 hours. The syzygy and gametocytes of S. pendula,
not attached to the epithelium, were collected directly in
the seawater from the Petri dishes and fixed in the same
conditions. After washing either in the same buffer or in buffer
containing 0.3 M sucrose, the samples were post-fixed with 1%
(w/v) OsO4 in the same buffer for 1 hr, then processed through
standard dehydration, infiltration, and embedding procedures,
in Epon or Araldite mixtures, with the corresponding solvents
(i.e. propylene oxide or acetone respectively), at room temperature.
The blocks were thin sectioned, collected on grids
Selenidiumpendula:UltrastructureandPhylogeny363
Table 2. Summary of biological, geographical and molecular data, for original isolates in this study. The number of corresponding stages used for
DNA preparations is indicated; T, trophozoite; C, cyst. Gene Accession numbers of the new sequences are available from the EMBL database.
Gregarine Host Location Isolate names Stage Gene Access
number (18S)
Selenidium pendula Giard 1884 Scolelepis squamata
(O. F. Müller 1806)
English Channel,
Roscoff, Aber,
Lat:48◦43 35.25 N,
Long:3◦59 22.54 W.
Selenidium pendula
LG
50-70 T LN901443
Selenidium pendula Giard 1884 Scolelepis squamata
(O. F. Müller 1806)
English Channel,
Roscoff-Aber 2012,
Lat:48◦43 35.25 N,
Long:3◦59 22.54 W.
Selenidium pendula
IF
50-70 T LN901444
Selenidium hollandei Vivier &
Schrével 1966
Sabellaria alveolata
(Linnaeus 1767)
English Channel,
Saint-Efflam-Ile Rouge
Lat:48◦40 57.96 N,
Long:3◦35 32.52 W.
Selenidium hollandei
LG
50-70 T LN901445
Lecudina pellucida (Mingazzini
1891)
Perinereis cultrifera
(Grübe 1840)
English Channel,
Roscoff-Ile de la Souris,
Lat:48◦43 41.73 N,
Long:3◦59 22.10 W.
Lecudina pellucida
LG
50-70 T LN901442
Lecudina tuzetae Schrével 1963 Neanthes (Nereis)
diversicolor (O. F.
Müller 1776)
English Channel,
Roscoff-Penzé 2012,
Lat:48◦37 40.07 N,
Long:3◦57 13.40 W.
Lecudina tuzetae
Roscoff 2012 IF132
7 C LN901446
Lecudina tuzetae Schrével 1963 Neanthes (Nereis)
diversicolor (O. F.
Müller 1776)
English Channel,
Roscoff-Penzé 2013,
Lat:48◦37 40,07 N,
Long:3◦57 13.40 W.
Lecudina tuzetae
Roscoff 2013a IF181
30 C LN901447
Lecudina tuzetae Schrével 1963 Neanthes (Nereis)
diversicolor (O. F.
Müller 1776)
English Channel,
Roscoff-Penzé 2013,
Lat:48◦37 40.07 N,
Long:3◦57 13.40 W.
Lecudina tuzetae
Roscoff 2013b IF462
2 C LN901448
Lecudina tuzetae Schrével 1963 Neanthes (Nereis)
diversicolor (O. F.
Müller 1776)
English Channel,
Roscoff-Penzé 2014,
Lat:48◦37 40,07 N,
Long:3◦57 13.40 W.
Lecudina tuzetae
Roscoff 2014a IF171
50 C LN901449
Lecudina tuzetae Schrével 1963 Neanthes (Nereis)
diversicolor (O. F.
Müller 1776)
English Channel,
Roscoff-Penzé 2014,
Lat:48◦37 40,07 N,
Long:3◦57 13.40 W.
Lecudina tuzetae
Roscoff 2014b IF172
50 C LN901450
364 J. Schrével et al.
and stained with saturated uranyl acetate in 50% (v/v) ethanol
for 1-3 min then in lead citrate. Sections were observed with a
Hitachi HU 11 E electron microscopy (Hitachi Ltd, Japan) or a
JEOL 1010 TEM.
For SEM, the intestines were open along the axis of the polychaete,
and the body parts highly infected by S. pendula were
carefully washed in 0.22 ␮m-filtered seawater before fixation
in glutaraldehyde as done above for TEM. After the post fixation
in 1% OsO4 in 0.2 M cacodylate buffer, specimens were
dehydrated in a graded series of acetone, critical point-dried in
liquid CO2 and coated with gold. The samples were examined
in a JEOL JSM-7401F FE SEM.
DNA isolation and sequencing: For the LG isolates (S.
pendula LG, S. hollandei LG, L. pellucida LG Table 1), groups
of ∼50-70 isolated trophozoites were washed at least three
times in 0.22 ␮m-filtered seawater and DNA was extracted
from individual parasites using a modified GITC (Guanidinium
isothiocyanate) protocol (Chomczynski and Sacchi 2006). Individuals
were placed in 50 ␮l of the GITC extraction buffer and
crushed using an adjusted micro-pilon (Kimble Chase®). Tubes
were incubated at 72 ◦
C for 20 min. Next, one volume of cold
isopropanol was added at −20 ◦
C overnight for DNA precipitation.
The following day, samples were centrifuged (20,000 g,
15 min at 4 ◦
C) and supernatants removed. The DNA pellet
was cleaned using 70% ethanol (100 ␮l), followed by a last
centrifugation (20,000 g, 10 min). Supernatant was removed
and the DNA pellet was hydrated into 20 ␮l of sterile distilled
water and stored at −20 ◦
C. For S. pendula IF, a group of ∼50-
70 isolated trophozoites were washed at least three times in
0.22 ␮m filtered seawater and genomic DNA was isolated by
using a phenol-chloroform extraction procedure as previously
described for Plasmodium falciparum (Florent et al. 2000), and
the purified DNA pellet was rehydrated into 20 ␮l of sterile distilled
water and stored at -20 ◦
C.
For L. tuzetae Roscoff 2012 IF462, DNA was isolated by
using the phenol-chloroform extraction procedure described
above, from 2 cysts, collected from the feces of a single Neanthes
(Nereis) diversicolor (O. F. Müller, 1776) host individually
kept in a Petri dish. The purified DNA pellet was rehydrated
into 20 ␮l of sterile distilled water and was stored at -20 ◦
C.
Finally, for the 4 remaining L. tuzetae Roscoff, DNA extractions
were performed using MasterPureTM
Complete DNA and RNA
Purification kit (Epicentre, Illumina Inc. USA) following supplier’s
recommendations for Cell Samples manipulations, with minor
modifications, from respectively 7 cysts (IF131), 50 cysts (IF171
and IF172) and 30 cysts (IF181). Briefly, each group of cysts
was isolated from the feces of a single N. diversicolor host individually
kept in a Petri dish, from which each cyst was then
extensively washed, one by one, in three successive drops of
0.22 ␮m filtered seawater supplemented with antibiotics penicillin
(100 U/mL), streptomycin (100 ␮g/mL) and gentamycin
(50 ␮g/mL) (Gibco, Life Technologies, USA) then pooled again.
Then, isolated and washed cysts were submerged in 300 ␮L
Tissue-and-Cell lysis solution, submitted to five series of freezing
(liquid nitrogen) and thawing (37 ◦
C) before addition of
Proteinase K then RNAse A and, after sample processing as
recommended, isolated DNA pellets were rehydrated in 35 ␮l
TE (10 mM Tris-pH 7.5 and 1 mM EDTA) prior to subsequent
storage at -20 ◦
C.
These DNA extraction products were then used as templates
in various series of PCR amplifications, in order to amplify the
SSU rRNA gene of these gregarines, then sequenced using the
Sanger sequencing methodology.
LG samples. The PCR mix (15 ␮l final volume) contained
1–6 ␮l of the DNA extract, 330 ␮M of each deoxynucleoside
triphosphate (dNTP), 2.5 mM of MgCl2, 1.25 U of GoTaq® DNA
polymerase (Promega Corporation), 0.17 ␮M of both primers,
1× of buffer (Promega Corporation). The PCR cycle, run in
an automated thermocycler (GeneAmp®PCR System 9700,
Applied Biosystem, USA), was programmed to give an initial
denaturating step at 95 ◦
C for 5 min, 35 cycles of denaturating
at 95 ◦
C for 1 min, annealing at 55 ◦
C for 45 s and extension
at 72 ◦
C for 1 min 15 s, and a final extension step at
72 ◦
C for 7 min. PCR products were cloned into a TOPO TA
cloning kit (Invitrogen®), following manufacturer’s recommendations.
Inserts inside white colonies were screened by PCR
(same procedure as before). Positive PCR products were purified
(ExoSAP-IT® For PCR Product Clean-Up, USB®) and
sequenced using the Big Dye Terminator Cycle Sequencing
Kit version 3.0 (PE Biosystems®) and an ABI PRISM model
377 (version 3.3) automated sequencer with specific internal
primers.
The list of primers used for both PCR amplifications and
Sanger sequencing is provided in the table of the Supplementary
data 6.
IF samples. PCR amplifications were done using Hot firepol
DNA polymerase as recommended (Solis BioDyne, Estonia),
in a 50 ␮l final volume supplemented with 2 mM MgCl2, 200 ␮M
each dNTPs and 200 nM forward (P4+T or WL1) and reverse
(EukP3) primers (Supplementary Material 6) and 1 ␮l of isolated
gregarine DNAs. PCR cycles, run in an automated thermocycler
(GeneAmp®PCR System 9700, Applied Biosystem, USA),
were programmed to give an initial denaturation step at 95 ◦
C
for 4 min, 30 cycles of denaturation at 95 ◦
C for 30 s, annealing
at 51 ◦
Cfor 30 s and extension at 72 ◦
C for 2 min, and a final
extension step at 72 ◦
C for 7 min. PCR products were purified
using IllustraTM
GFXTM
PCR DNA and Gel Band Purification
kit (GE Healthcare, France) and were cloned into pGEM®
-T
Easy vector (Promega, Madison WI, USA) using supplier’s recommendations.
DNA sequences were obtained from positive
clones selected by PCR using T7 and Sp6 universal primers
flanking the pGEM®
-T Easy vector cloning site, using T7, Sp6
and internal primers such as LWA1, LWA3, PIF3F and PIF3R
(Table 6), by the Sanger method (Beckman Coulter Genomics,
Takeley, UK). Raw were edited using the BioEdit 7.1.3.0 program
(Hall 1999) and assembled by using MEGA6 (Tamura
et al. 2013).
Phylogenetic analyses: SSU rDNA sequences from nine
Selenium and Lecudina species were aligned to 106 rDNA
sequences from diverse eukaryotes, mostly corresponding to
representatives of Alveolata with one Rhizaria as outgroup.
Sequences were aligned using the online version of MAFFT
version 7 (http://mafft.cbrc.jp/alignment/server/ Katosh and Toh
2010), using the secondary structure of RNA (Q-INS-I option)
and further refined manually taking as a reference the secondary
structure of T. gondii small subunit rRNA (Gagnon et al.
1996). Ambiguously aligned positions were manually removed
which yielded a confident alignment of 1350 positions. A GTR
substitution model with gamma-distributed rate variation across
sites was suggested as the best-fit model by JModeltest V2.1.3
(Darriba et al. 2012). Accordingly, a Bayesian phylogenetic tree
was constructed with MrBayes v.3.2.3 (Ronquist et al. 2012)
using lset nst=6 rates=Invgamma Ngammacat=4 parameters.
Four simultaneous Monte Carlo Markov chains were run from
random trees for a total of 13,000,000 generations in two parallel
runs. A tree was sampled every 1000 generation and 25%
of the trees were discarded as “burn-in”. A consensus tree was
constructed from the post-burn-in trees and posterior probabilities
were calculated in MrBayes. Maximum Likelihood analyses
were performed with MEGA 6.06 (Tamura et al. 2013) using
the GTR+G+I model. Bootstraps were estimated from 1,000
replicates.
Selenidium pendula: Ultrastructure and Phylogeny 365
The phylogenetic tree for the Selenidiidae lineage from
polychate annelids (Fig. 13) was contructed using the same
alignment but for a subset of 20 sequences; all position containing
gaps and missing data were eliminated; there were a
total of 1,416 positions in the final dataset. Maximum Likelihood
analyses were performed with MEGA 6.06 (Tamura et al.
2013) using the GTR+G+I model. Bootstraps were estimated
from 1,000 replicates.
Estimate of evolutionary divergence between
sequences: Evolutionary divergence between sequences
was computed by using the MEGA 6.06 (Tamura et al.
2013) using a subset of sequences extracted from the main
phylogenetic alignment. For the analysis of the Selenidiidae
lineage (Supplementary Material 4) the analysis involved 33
nucleotide sequences for 16 distinct species, there were a
total of 2088 positions in the final dataset and all positions
containing gaps and missing data were eliminated.
Acknowledgements
Many thanks to Dr Isabelle Desportes (MNHN) for
her fruitful comments, Dr Marc Dellinger (MNHN)
for help in primer design and Dr Marc Gèze (MNHN)
for imaging on the MNHN-CEMIM Platform. The
careful illustrations of the figures were realized
by Doanh Baccam (MNHN). This study was supported
by ECO-NET Project 2131QM (Égide,
France), PHC-Barrende projects 24663ND (2011-
2012) and 31266SL (2014-2015), Interdisciplinary
Programs of the MNHN (ATM-Microorganismes,
ATM-Emergence des clades, des biotes et des
cultures) and by the French governmental ANR
under ANR-10-LABX-0003 BCDiv, ANR-11-IDEX-
0004-0 and the ANR-14-CE02-0007-01 (HAPAR).
AV was funded by the Czech Science Foundation
project No. GBP505/12/G112 (ECIP), and her travel
expenses were supported by projects MEB021127
and 7AMB14FR013 from the Ministry of Education,
Youth and Sports of the Czech Republic.
Appendix A. Supplementary Data
Supplementary data associated with this article
can be found, in the online version, at
http://dx.doi.org/10.1016/j.protis.2016.06.001.
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Supplementary data 1. General view of a young syzygy of Selenidium pendula in TEM.
Since the size of each gamont was too large for a single TEM micrograph, in this composite
view we indicated the missing area between the two paired gamonts by a white band.
Abbreviations: nucleus (N), nucleolus (nu). The sexual stages of gregarines start with the
pairing of two gamonts. In one syzygy the gamont’s cytoplasm as well the nuclei exhibit a
similar organization. The spherical nucleoli of paired gamonts are of similar sizes and both
contain several clear areas (Arrows).
Supplementary data 2. Estimates of evolutionary divergence between sequences. The
numbers of base differences per site from between sequences are shown. The analysis
involved 33 nucleotide sequences. All ambiguous positions were removed for each sequence
pair. There were a total of 2088 positions in the final dataset. Evolutionary analyses were
conducted in MEGA6 (Tamura et al. 2013).
Supplementary data 3. Estimates of evolutionary divergence between sequences of
Selenidiidae family.
The number of base differences per site between sequences of two given species is indicated
as %. The analysis involved 33 nucleotide sequences for 16 distinct species. Indeed, for some
species, up to 5 distinct sequences were taken into account in the calculations (this sequence
number is indicated for each species in the first column, no indication corresponding to a
unique sequence). Evolutionary analyses were conducted in MEGA6 (Tamura et al. 2013).
There were a total of 2088 positions in the final dataset. For the pairwise analysis, all
ambiguous positions were removed for each sequence pair.
S. pendula
(Lineage I)
S. hollandei
(Lineage I)
S. terebellae
(Lineage II)
S. orientale
(Lineage III)
S. pendula (2seq) 1.2% 8.5 - 9.5 % 25.8 - 26.9 % 28.5 % Lineage I
S. boccardiella 3.4 - 4.2 % 8.9 % 25.9 - 26.3 % 27.2 % Lineage I
S. neosabellariae
(3seq)
8.1 - 10.2 % 3.0 - 3.7 % 26.8 - 27.6 % 27.3 - 28.4 % Lineage I
S. hollandei 8.5 - 9.5 % - 27.5 - 27.7 % 28.8 % Lineage I
S. identhyrsae 8.5 - 10 % 3.4% 27.0 - 27.3 % 27.7% Lineage I
S. cf mesnilli 8.7 - 9.5 % 9.8 % 26.9 - 27.3 % 28.1 % Lineage I
S. echinatum
(2seq)
8.8 - 10.7% 11.4 % 26.1 - 26.6 % 28.5% Lineage I
S. sensimae
(2seq)
10 - 11.3 % 12.1 % 26.4 - 26.8 % 28.1 - 28.2 % Lineage I
S. serpulae 10.2 - 10.9 % 11 % 26.6 - 26.9 % 27.6 % Lineage I
S. sp1 (3seq) 10.4 - 11.4 % 11.4 - 11.6 % 26.2 - 26.7 % 27.2 - 27.3 % Lineage I
S. sp2 (3seq) 11.7 - 13.4 % 13.3 - 13.7 % 26.8 - 27.5 % 28.5 - 29.1 % Lineage I
S. terebellae
(5seq)
25.8 - 26.9 % 27.5 - 27.7 % 0.1 - 0.4 % 20.3 - 20.6 % Lineage II
S. melongena
(4seq)
25.9 - 27.2 % 28 - 28.2 % 12.5 - 13 % 19.5 - 19.7 % Lineage II
S. pisinnus 26.3 - 26.8 % 28.4 % 17.6 - 17.8 % 15.2 % Lineage III
S. orientale 28.5 % 28.8 % 20.3 - 20.6 % - Lineage III
Plathyproteum
vivax
30 - 30.5 % 31 % 25.6 - 25.8 % 28.6 Lineage III
Filipodium
phascolosomae
30.9 % 32.1 % 25.8 - 26.1 % 27.7 % Lineage III
Supplementary data 4. Molecular phylogenetic analysis by Maximum Likelihood method of
the Selenidiidae lineage parasites of polychaete annelids and the Selenidiidae-like lineage
parasites of the Terebellidae family.
The evolutionary history was inferred by using the Maximum Likelihood method based on the
General Time Reversible model (Nei and Kumar 2000). The tree with the highest log
likelihood (-9626.0015) is shown. A discrete Gamma distribution was used to model
evolutionary rate differences among sites (5 categories (+G, parameter = 0.5500). The rate
variation model allowed for some sites to be evolutionarily invariable (+I), 16.4799% sites).
The tree is drawn to scale, with branch lengths measured in the number of substitutions per
site. The analysis involved 33 nucleotide sequences. All positions containing gaps and
missing data were eliminated. There were a total of 1374 positions in the final dataset.
Evolutionary analyses were conducted in MEGA6 (Tamura et al. 2013).
Supplementary data 5. Cross sections of Selenidium terebellae Ray 1930 intestinal parasite
of the polychaete Terebella lapidaria Linnaeus 1776 from Roscoff (A-D). Abbreviations:
amylopectin granule (am), bulge (B), dense layer (dl), groove (G), inner membrane complex
(imc), mitochondria (M), microtubule (mt), nucleus (N). A. General view of a trophozoite
with large bulges separated by grooves. B. Cross section of the cortex with a dense layer
between the imc and the subpellicular microtubules. C. Cross section along a groove with
accumulation of membranous residual organelles in the cytoplasm under the groove. D.
Detail of the dense layer and the cross section of the microtubules (original data).
7.8
Chambouvet A., Valigurová A., Pinheiro L.M.,
Richards T.A., Jirků M.
2016
Nematopsis temporariae (Gregarinasina, Apicomplexa, Alveolata)
is an intracellular infectious agent of tadpole livers
Environmental Microbiology Reports 8(5), 675-679
Nematopsis temporariae (Gregarinasina, Apicomplexa,
Alveolata) is an intracellular infectious agent of
tadpole livers
Aurelie Chambouvet,1
* Andrea Valigurova,2
Lara M. Pinheiro,3
Thomas A. Richards1,4
and
Miloslav Jirk˚u5
*
1
Biosciences, University of Exeter, Geoffrey Pope
Building, Exeter, EX4 4QD, UK.
2
Department of Botany and Zoology, Faculty of Science,
Masaryk University, Kotlarska 2, Brno, 611 37, Czech
Republic.
3
National Council for Technological and Scientific
Development (CNPq), Brazil.
4
Integrated Microbial Biodiversity Program, Canadian
Institute for Advanced Research (CIFAR), Toronto,
Canada.
5
Biology Centre, Czech Academy of Sciences, Institute
of Parasitology, Branisovska 31, Ceske Budejovice, 370
05, Czech Republic.
Summary
Amphibians are in decline as a result of habitat
destruction, climate change and infectious diseases.
Tadpoles are thought susceptible to infections
because they are dependent on only an innate
immune system (e.g. macrophages). This is because
the frog adaptive immune system does not function
until later stages of their life cycle. In 1920, N€oller
described a putative infectious agent of tadpoles
named Nematopsis temporariae, which he putatively
assigned to gregarine protists (Apicomplexa). Here,
we identify a gregarine infection of tadpoles using
both microscopy and ribosomal DNA sequencing of
three different frog species (Rana temporaria, R. dalmatina,
and Hyla arborea). We show that this protist
lineage belongs to the subclass Gregarinasina
Dufour 1828 and is regularly present in macrophages
located in liver sinusoids of tadpoles, confirming the
only known case of a gregarine infection of a
vertebrate.
Introduction
Amphibian populations are in crisis with 48% of populations
reported as declining (Stuart et al., 2004). The emergence
of infectious diseases is thought to be a major factor (Daszak
et al., 2003; Martel et al., 2013). Amphibian physiology
varies considerably during the life cycle. Tadpoles have a
weak adaptive immunity with fewer antibody classes, poorer
B and T lymphocyte function, no consistent expression of
the major histocompability complex (MHC) class I protein
and a poor switch from IgM to IgY (Du Pasquier et al.,
1989). Tadpoles, therefore, rely on an innate immune system
that provides rapid and nonspecific protection. As such
tadpoles host a diversity of different microbial organisms,
acting as either definitive or intermediate hosts. Specifically,
investigation of tadpole livers have identified a diversity of
alveolate protists (Jirk˚u et al., 2002; 2009; Davis et al.,
2007; Chambouvet et al., 2015) for which their role as putative
parasites is unclear.
One enigmatic group of alveolates are the gregarines.
Phylogenetic analyses show gregarines branch within
the subphylum Apicomplexa Levine, 1980, emend. Adl
et al. 2012 (Leander et al., 2003; Adl et al., 2012), which
also includes parasites of mammals, e.g. Plasmodium
spp. All described gregarines are parasites (Leander
et al., 2003) and are known to infect many groups of
invertebrates, particularly annelids and insects (Leander,
2008). In 1920, N€oller described a gregarine named
Nematopsis temporariae infecting the liver tissue of the
frog Rana temporaria (N€oller, 1920). Here, we report the
identification of an infectious microbe fitting this description
from three species of frog tadpoles sampled in the
Czech Republic using molecular and microscopy data.
Results and discussion
During an amphibian population survey in the Czech
Republic we identified a gregarine-like intracellular infection
of liver cells from tadpoles of R. temporaria, R. dalmatina
and H. arborea. These tadpoles showed no signs of disease
or impairment of fitness/function, although livers of
some tadpoles appeared slightly enlarged and light coloured,
they were not yellowish as previously reported for
Perkinsea (Alveolata) infections (Davis et al., 2007). No
mortalities of tadpoles or metamorphs were recorded in the
Received 23 December, 2015; accepted 21 April, 2016. For correspondence:
*E-mail a.chambouvet@exeter.ac.uk; Tel. 144 (0) 1392
726527; Fax 144 (0)1392 263434. **E-mail miloslav.jirku@seznam.cz;
Tel. 142 (0) 739014817; Fax 142 (0)-38-5310388.
VC 2016 Society for Applied Microbiology and John Wiley & Sons Ltd
Environmental Microbiology Reports (2016) 8(5), 675–679 doi:10.1111/1758-2229.12421
field. Dissections of the tadpoles were carried out using
standard procedures identifying the protist infection in multiple
samples (n5 20 R. damatina, 20 R. temporaria and 15
H. arborea) from Zajecˇı potok, Brno, Czech Republic
(49.23765N, 16.60637E) and Radun, Czech Republic
(49.88997N, 17.94375E). All specimens were in Gosner
stage 26 or higher (Supporting Information Table S1 – and
see below for discussion of sampling for N. temporariae
beyond metamorphosis). The observed morphological characteristics
are consistent with the original description of N.
temporariae, specifically the protists observed possess
monozoic oocysts and are morphologically and morphometrically
consistent with the original description of N. temporariae
(see description below), we therefore assign the
gregarine-like oocysts to this species.
Standard light microscopy squash examination of liver,
gall bladder, skin, heart, intestine and tail muscle of all
examined tadpoles from the two localities revealed the
presence of N. temporariae oocysts exclusively in host livers,
demonstrating the intracellular microbial infection was
not present in other host tissues examined. Samples of all
examined tissues from each tadpole were fixed in 10%
buffered formalin and glutaraldehyde, processed routinely,
stained either with haematoxylin and eosin or ToluidineBlue
and examined by light or transmission electron
microscopy. Each oocyst is ovoid, asymmetrical with one
side usually flattened measuring 15.5 (14.0–17.0) 3 6.5
(5.0–7.5) lm (Fig. 1A and B). Using light microscopy, sporozoites
appeared transversely striated that corresponds to
micronemes organized in parallel layers (Fig. 1C). On a
few occasions, we observed a free sporozoite, keeping its’
overall banana shape during gliding movement, with only
apical end appearing fully flexible (Fig. 1D). Oocysts were
the only developmental stage of N. temporariae consistently
sampled, making unclear if the tadpoles serve as
definitive or intermediate host of N. temporariae.
Fig. 1. Oocysts and free sporozoite of Nematopsis temporariae of Rana dalmatina tadpoles using microscopic analysis; fresh mount NIC (A–
D), histological section stained with Toluidine-Blue (E), transmission electron microscopy (TEM) (F).
A. Intracellular oocyst (arrow) with single sporozoite (s) in a macrophage (white arrowheads).
B. Macrophage containing oocysts of both N. temporariae and G. noelleri (arrowhead); the macrophage as well as G. noelleri oocyst are ruptured
by pressure during the squash preparation; N. temporariae oocysts are mechanically flattened, making the sporozoites more dispersed
than normal; see the pigment granules upper right.
C. Composite micrograph of oocysts containing sporozoites showing distinct transverse striation.
D. Composite micrograph of a free sporozoite in gliding motion.
E. Macrophage containing two oocysts of N. temporariae (arrows) in lumen (L) of liver sinusoid.
F. Macrophage (white arrowheads) containing oocyst of N. temporariae (arrow); n – macrophage nucleus, s – sporozoite. A, B, C, D in the same scale.
676 A. Chambouvet et al.
VC 2016 Society for Applied Microbiology and John Wiley & Sons Ltd, Environmental Microbiology Reports, 8, 675–679
In most preparations (n 5 40), both N. temporariae
oocysts and Goussia oocysts (i.e. protists cell with a
fine elastic oocyst wall and four dizoic sporocysts measuring
7.5 (7.0–8.0) 3 4.7 (4.0–5.0) (n 5 50) – Eimeriorina
Leger, 1911, Apicomplexa) were observed to occupy
the same cells (Fig. 1B) (Jirk˚u et al., 2009). However, in
the H. arborea samples inspected (n 5 15), this coinfection
was not identified. In tadpole liver histological
sections stained with Toluidine-Blue, oocysts were readily
identified due to their characteristic morphology (Fig.
1E). Similarly as in fresh preparations, some oocysts
were empty, sometimes containing residual granules.
Interestingly, histological and transmission electron
microscopy (TEM) examinations revealed presence of
oocysts exclusively in phagocytic cells in liver sinusoids
(Fig. 1E). Both non-pigmented (c.f. Kupffer cells) and
pigmented (containing melanosomes) cell types were
identified (Fig. 1A, B, E and F). The oocysts-containing
cells belong to a macrophage lineage as reflected by
their amoeboid nature with a notable variability in size
and shape, typical filopodia, irregularly shaped nucleus,
the presence of various quantities of lysosomes and
phagosomes, poorly developed rough endoplasmic reticulum,
Golgi bodies, a well-developed cortical microvacuolar
system, small mitochondria and eventually
melanosomes (e.g. Guida et al., 1998) (Fig. 1F).
To investigate progression of the Nematopsis infection,
an additional 45 tadpoles of Rana dalmatina (Gosner
stages 33–42) were collected at Zajecˇı potok on the
first of July 2004. Twenty-five tadpoles were euthanized
by pithing and examined as described above for a presence
of Nematopsis demonstrating presence of the
infection in liver tissue. Additionally, tadpoles of R. dalmatina
were kept in captivity beyond metamorphosis to
assess the fate of Nematopsis oocysts in metamorphosed
animals. A subset of 20 juvenile (and later subadult)
frogs in total were dissected at intervals of 2
weeks for the first 2 months, then every one month for
the third and fourth months, and every 3 months for
the rest of the experiment up to the 15th month postmetamorphosis.
In both fresh and histological preparations
of livers from hosts examined, all tadpoles
investigated were Nematopsis positive, while for organisms
4–6 weeks after metamorphosis, only empty
oocysts were found.
In parallel to the histology analysis, we selected two
liver samples from two different species: R. temporaria
and H. arborea (four in total) and isolated 10–15 cells by
mouth pipetting for DNA extraction. Using the eukaryotic
forward primer (Euk1F) with the general -non-metazoanreverse
primer (Supporting Information Table S2), we
PCR amplified and double strand sequenced (1000 bp
of SSU gene) 10 clones per liver sample. All sequences
recovered showed  97% identity. A conserved portion
of the alignment was selected to design a ‘Nematopsis’
specific forward primer. This primer NEM-1F was used
in association with the primer 28S-R1 targeting the 50
of
the LSU rRNA gene from R. temporaria (three samples),
H. arborea (three samples) and R. dalmatina (two samples
– Supporting Information Table S2). For each liver
sample, three independent PCR amplifications were
mixed and cloned. Three clones per sample were double
strand sequenced (see SMM and Supporting Information
Table S2).
Currently, there is only one sequence of the complete
ribosomal RNA encoding gene belonging to the Gregarinasina
Dufour, 1928 available in the Genbank nr database
(Gregarina sp. JF412715, March 2016). To allow for
comprehensive taxon sampling, phylogenetic analysis
was, therefore, based on an alignment of the SSU gene
that encompassed the V4 and V9 loops. The sequence
alignment included 65 publically available sequences previously
used for phylogenetic analysis (Rueckert et al.,
2011; Wakeman et al., 2014) and 24 clone sequences
recovered here. The ML and Bayesian phylogenies recovered
a weakly supported backbone as previously
described in phylogenies of the gregarines (Rueckert
et al., 2011; Wakeman et al., 2014) (Fig. 2A). However,
the SSU rDNA gene sequences recovered from the tadpole
tissue form a highly supported clade (1/100/100)
and branch with moderate bootstrap values (1/77/100)
with the terrestrial gregarine clade 1 sequences (Rueckert
et al., 2011; Wakeman et al., 2014) (Fig. 2A). Many
alveolate genomes are highly AT rich (Gardner et al.,
2002; Kopecna et al., 2006). We conducted Log-Det distance
bootstrap analysis to account for differential base
composition as a source of artifact (Foster and Hickey,
1999). This phylogenetic method provides strong support
for phylogenetic association of Nematopsis with the terrestrial
gregarines. This clade encompasses gregarine
pathogens of a wide range of invertebrates, e.g. damselflies,
earthworms, dragonflies, green darners, mosquitoes
and sandflies (Fig. 2A). The phylogenetic results show
that N. temporariae belongs to gregarines and confirms
that this is the first example of a member of the subclass
Gregarinasina, Dufour 1828, infecting a vertebrate.
Eukaryotic ribosomal RNA gene clusters (rRNA
genes) are typically present in multiple copies within a
nuclear genome (Long and Dawid, 1980). The internal
transcribed spacers (ITS1 and ITS2) that separate the
SSU, 5.8S and LSU genes have a high rate of sequence
variation. We generated 24 independent clone sequences
from eight liver samples (three clones per sample).
These sequences showed between 96% and 99%
sequence identities across the SSU-ITS1-5S-ITS2 ribosomal
sequences (Fig. 2B and C, and Supporting Information
Table S3). Considering only single nucleotide
polymorphisms that occurred in at least two independent
Gregarines in tadpole cells 677
VC 2016 Society for Applied Microbiology and John Wiley & Sons Ltd, Environmental Microbiology Reports, 8, 675–679
Fig. 2. A. RAxML tree investigating the phylogenetic placement of N. temporariae. The phylogeny is calculated from 89 sequences and 1276
alignment positions. Bayesian posterior probability, ML and Log-Det bootstrap values were notated using the following convention: support values
are summarized by black circles when  0.9/80%/80% and white circles when this is not the case but all values are  0.6/50%/50%, actual
values are shown for key branching relationships. The double-slashed line represents branches shortened by 1=2. The identification of the different
clades was reported as described in (Rueckert et al., 2011; Wakeman et al., 2014). B. Unrooted maximum likelihood phylogenetic tree of
the ribosomal RNA gene cluster sequences. The colours of the clone’s names identified the tadpole liver tissue samples and the host taxonomy
(see key). C. Representation of the ribosomal gene cluster and the relative position of the different primer used in this study (not to scale).
For each region of the rRNA gene cluster the number of SNPs were indicated in brackets if the mutation is retrieved in at least two independent
clones. The ITS1 region where at least two separate nucleotide motifs have been detected is represented using http://weblogo.berkeley.
edu.
678 A. Chambouvet et al.
VC 2016 Society for Applied Microbiology and John Wiley & Sons Ltd, Environmental Microbiology Reports, 8, 675–679
clone sequences, we identified SNPs that identify variation
specific for distinct rDNA-types. The main region of
polymorphism was located within the ITS1 region identifying
a minimum of two major rDNA-types (Fig. 2C), representing
either inter or intraindividual genetic diversity.
This study represents the first molecular and microscopic
description of the association between a gregarine
and a vertebrate, and importantly shows that the N.
temporariae oocysts form intracellular infections of tadpole
cells. It is unclear whether tadpoles serve as definitive
or intermediate hosts. These results provide the
molecular tools for studying this infectious agent with
regard to wider environmental ecology and specifically
distribution in amphibian populations.
Acknowledgements
A.C. is supported by a Marie Curie Intra-European and
EMBO Long-Term Fellowships (FP7-PEOPLE-2011-IEF-
299815-PARAFROGS and ATL-1069-2011). A.V. was
funded from Czech Science Foundation (GBP505/12/
G112-ECIP) and acknowledges support from the Department
of Botany and Zoology of Masaryk University. L.M.P.
was supported by the National Council for Technological
and Scientific Development (CNPq) - Brazil (process no.
237499/2013-4). M.J. was supported by institutional support
(RVO: 60077344, Institute of Parasitology, BC CAS)
and acknowledges David Modry (VFU Brno, Czech Rep.)
for support and introduction to amphibian parasites.
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Supporting information
Additional Supporting Information may be found in the
online version of this article at the publisher’s web-site:
Table S1. Detail of the tadpoles sampled, providence and
the corresponding Nematopsis clone library sequences
recovered.
Table S2. Primers used in this study.
Table S3. Identity percentage between the ribosomal operons
sequences of Nematopsis temporariae from the different
clone sequences. Data were calculated using the SIAS
website (Available at http://imed.med.ucm.es/Tools/sias.
html).
Gregarines in tadpole cells 679
VC 2016 Society for Applied Microbiology and John Wiley & Sons Ltd, Environmental Microbiology Reports, 8, 675–679
7.9
Diakin A., Wakeman K.C., Valigurová A.
2017
Description of Ganymedes yurii sp. n. (Ganymedidae), a new
gregarine species from the Antarctic amphipod
Gondogeneia sp. (Crustacea)
Journal of Eukaryotic Microbiology 64(1), 56-66
ORIGINAL ARTICLE
Description of Ganymedes yurii sp. n. (Ganymedidae), a New
Gregarine Species from the Antarctic Amphipod
Gondogeneia sp. (Crustacea)
Andrei Diakina
, Kevin C. Wakemanb
& Andrea Valigurovaa
a Department of Botany and Zoology, Faculty of Science, Masaryk University, Kotlarska 2, Brno 611 37, Czech Republic
b Center for Global Communication Strategies, The University of Tokyo Meguro-ku, Komaba Campus, Tokyo 153-8902, Japan
Keywords
Apicomplexa; Cephaloidophoroidea; eugregarine;
marine benthic amphipod; molecular
phylogeny; ultrastructure; Weddell Sea.
Correspondence
A. Diakin, Department of Botany and Zoology,
Faculty of Science, Masaryk University,
Kotlarska 2, Brno 611 37, Czech Republic
Telephone number: +420-549-49-8844;
FAX number: +420-549-49-8331;
e-mail: ondredyakin@gmail.com
Received: 23 March 2016; revised 7 June
2016; accepted June 7, 2016.
doi:10.1111/jeu.12336
ABSTRACT
A novel species of aseptate eugregarine, Ganymedes yurii sp. n., is described
using microscopic and molecular approaches. It inhabits the intestine of Gondogeneia
sp., a benthic amphipod found along the shore of James Ross Island,
Weddell Sea, Antarctica. The prevalence of the infection was very low and
only a few caudo-frontal syzygies were found. Morphologically, the new species
is close to a previously described amphipod gregarine, Ganymedes
themistos, albeit with several dissimilarities in the structure of the contact
zone between syzygy partners, as well as other characteristics. Phylogenetic
analysis of the 18S rDNA from G. yurii supported a close relationship between
these species. These two species were grouped with other gregarines isolated
from crustaceans hosts (Cephaloidophoroidea); however, statistical support
throughout the clade of Cephaloidophoroidea gregarines was minimal using
the available dataset.
THE Antarctic is an exceptional continent, with extreme
climates and environmental conditions above and below
the water surface. While not many species are thought to
survive in these high latitudes, the diversity of marine
fauna in this part of the world is enormously rich, especially
in the plankton, nekton, and benthos. A number of
reviews are dedicated to the climatology, geology,
palaeontology, and fauna of vertebrates (fish, birds, and
mammals) from the Antarctic region (Barbosa and Palacios
2009; Eastman 2005). Similarly, many investigations were
devoted to apicomplexans parasitising fish, birds, and krill
(Avdeev 1985, 1987; Barber and Mills Westermann 1988;
Golemansky 2011; Takahashi et al. 2003, 2004, 2008,
2009). To date, only one study has been devoted to crustacean
gregarines occurring in littoral/sublittoral invertebrates
(Lipa and Rakusa-Suszczewski 1980).
Apicomplexans are a diverse group of unicellular parasites,
inhabiting almost all known phyla of multicellular
organisms. Some Apicomplexa such as Toxoplasma, Plasmodium,
Eimeria, and Cryptosporidium are well known, as
these parasites cause harmful diseases in humans and
domestic animals. The invasive stages (zoites) in this
group are characterised by the presence of an ‘apical
complex’, comprising a set of subcellular organelles (conoid,
rhopries, and miconemes), specialised for invading
and subsequently modifying the host cells (Chobotar and
Scholtyseck 1982; Scholtyseck 1973; Scholtyseck and
Mehlhorn 1970).
Gregarines, in contrast to coccidia, largely exist as extracellular
parasites of a broad range of invertebrate groups,
for example, terrestrial insects, aquatic annelids, and crustaceans.
Most gregarines inhabit the host intestinal lumen,
are elongated and heteropolar, cylindrical, or vermiform in
shape. Feeding stages (=trophozoites) generally develop
attached to the host cell via their modified anterior end.
Usually, the trophozoites are subdivided into three parts:
epimerite (attachment function), and protomerite followed
by a deutomerite containing a large nucleus. The last two
regions are separated by a fibrillar septum. Gregarines
with such organisation are classified as being “tricystid”
or “septate” gregarines. On the other hand, gregarines
that are not subdivided represent the “monocystid” form,
and are known as aseptate gregarines. It should also be
mentioned that, generally, trophozoites, as well as subsequent
sexual stages (=gamonts), exhibit gliding motility
and possess a unique organisation of the cell cortex. The
© 2016 The Author(s) Journal of Eukaryotic Microbiology © 2016 International Society of Protistologists
Journal of Eukaryotic Microbiology 2017, 64, 56–6656
Journal of Eukaryotic Microbiology ISSN 1066-5234
pellicle of eugregarines forms longitudinal epicytic folds
equipped with special sets filamentous structures in their
apex, represented by rippled-dense structure and 12-nm
filaments. Before gametogenesis, the gamonts join into a
sexual association called syzygy. Later on, the paired gregarines
form a common envelope (gametocyst), under
which further processes, including gametogenesis, fertilisation
and formation of invasive stages (sporogenesis),
take place (Desportes and Schrevel 2013; Frolov 1991;
Grasse 1953; Long 1982; Perkins et al. 2000; Simdyanov
2007).
Many septate and aseptate eugregarines have been
described from different crustacean hosts from different
marine and terrestrial aquatic localities, and have traditionally
been distinguished based on their general morphology
using light microscopy. These gregarines have been separated
into different families including the Cephaloidophoridae,
Porosporidae, Uradiophoridae, Ganymedidae, and
others. However, this system of families and nomenclature
at the level of genus and species remains unsettled
(Desportes and Schrevel 2013; Grasse 1953; Levine
1977a,b; Perkins et al. 2000; Simdyanov 2007). Furthermore,
the data collected from each group are not uniform,
for example, only some of these groups have been investigated
using electron microscopic approaches and/or
molecular techniques (Desportes and Theodorides 1969,
1985; Rueckert et al. 2011; Simdyanov et al. 2015; Takahashi
et al. 2009; Theodorides and Desportes 1975).
Recent phylogenetic analyses of SSU rDNA sequences
showed that gregarines from different crustacean hosts
clustered in a single clade, together with a number of
environmental sequences (Rueckert et al. 2011). This finding
was confirmed by phylogenetic analyses of LSU rDNA
and whole ribosomal operon (SSU + 5.8S + LSU) (accession
numbers HQ891113.2 – HQ891115.2) (Simdyanov
et al. 2015).
The family Ganymedidae was established by Huxley
(1910) and comprises intestinal aseptate gregarines possessing
ball-like and cup-like structures at the anterior and
posterior ends of the cell, respectively. He described the
type species, Ganymedes anaspidis, from a mountain
shrimp, Anaspides tasmaniae Thomson, 1892 (Huxley
1910). Later many aseptate gregarines were described
from different freshwater and marine crustacean hosts
(Cirripedia, Amphipoda, Decapoda etc.) (Jones 1968, 1969;
Jones et al. 1994; Prokopowicz et al. 2010; Theodorides
and Desportes 1972, 1975). Subsequently, Levine (1977a,
b) made a taxonomical revision of this genus; he placed all
species lacking the above-mentioned ball-like and cup-like
structures into a new genus, Paraophioidina, and only one
species was retained in the genus Ganymedes, namely
G. anaspidis. Perkins et al. (2000) and Simdyanov (2007)
followed this opinion. In the latest revision of gregarine
species, this point of view was rejected and many species
were returned and assigned to the genus Ganymedes
(Desportes and Schrevel 2013).
In this study, we describe the general morphology and
molecular phylogeny, based on SSU sequence data, of a
new Antarctic gregarine, Ganymedes yurii sp. n. For this,
we used a combined approach of transmission and scanning
electron microscopy, and molecular phylogenetic
analyses.
MATERIALS AND METHODS
Gondogeneia sp. Barnard, 1972, an amphipod, was collected
in January and February 2013 in the littoral and
upper sublittoral zone of Cape Lachman (63°470
32″S,
57°460
86″W), James Ross Island, Weddell Sea, Antarctica.
The amphipods were transported to the laboratory and
kept in cold conditions. About 400 crustaceans were dissected
under a stereomicroscope (MST 131, Poland). Parasites
released from the host intestine were collected
using a thin glass pipette. Light microscopic observations
of living parasites were performed using an Olympus
CX41 Microscope (Olympus Corp., Tokyo, Japan)
equipped with phase contrast and connected to an Olympus
Camedia C-7070 Digital Camera (Olympus Corp.).
For electron microscopy, parasites were fixed in 2.5%
glutaraldehyde in Millipore-filtered sea water (SW) (MillexGC
0.22 lm). For transmission electron microscopy
(TEM), gregarines were then postfixed with 1% OsO4
(Os) in 0.2 M cacodylate buffer, dehydrated in an ethanol
series and embedded in Epon blocks. Ultra-thin sections
were made using Reichert Ultracut E and Leica UTC ultramicrotomes,
stained according to a standard protocol
(Reynolds 1963), and observed under a JEOL-1010 transmission
electron microscope (JEOL Ltd., Peabody, MA,
USA). For scanning electron microscopy (SEM), fixed
trophozoites were postfixed for 2 h in 2% osmium tetroxide
in 0.2 M cacodylate buffer, dehydrated, dried with
CO2 using Emitech K850, and then coated with gold using
Emitech K550 sputter coaters. The samples were
observed under a JEOL JSM-7401F field emission scanning
microscope (JEOL Ltd.).
For molecular analysis, gamonts in syzygy were fixed in
100% ethanol and stored at room temperature. Genomic
DNA was later extracted with the standard protocol provided
in the MasterPure Complete DNA & RNA Purification
Kit (Epicentre Biotechnologies, Madison, WI).
However, the final elution step was lowered to 4 ll. Outside
primers, PF1 50
–GCGCTACCTGGTTGATCCTGCC–30
and SSUR4 50
–GATCCTTCTGCAGGTTCACCTAC–30
(Leander
et al. 2003), were used in a 25 ll PCR with EconoTaq
2X Master Mix (Lucigen Corp., Middleton, WI). The following
programme was used on a thermocycler for the initial
amplification: Initial denaturation at 94 °C for 2 min; 35
cycles of denature at 94 °C for 30 s, anneal at 52 °C for
30 s, extension at 72 °C for 1 min 50 s, final extension
72 °C 5 min. Subsequently, internal primers F2 50
–GCYTG
AAAAGGTGACDATCTG–30
and R2 50
–CATATCTGCTAAG
GTTCTG–30
were paired with outside primers in a nested
PCR using the following programme on a thermocycler:
Initial denaturation for 94 °C for 2 min; 25 cycles of denature
at 94 °C for 30 s, anneal at 52 °C for 30 s, extension
at 72 °C for 1 min 30 s; final extension at 72 °C for 7 min.
The newly obtained DNA sequence from Ganymedes
yurii was initially identified by BLAST (http://
© 2016 The Author(s) Journal of Eukaryotic Microbiology © 2016 International Society of Protistologists
Journal of Eukaryotic Microbiology 2017, 64, 56–66 57
Diakin et al. Morphology and Phylogeny of Ganymedes yurii sp. n.
blast.ncbi.nlm.nih.gov). This sequence was then aligned
with 89 additional alveolate sequences selected from
NCBI/GenBank (to cover the diversity of apicomplexans,
including some dinoflagellates as an outgroup), using
MUSCLE 3.8.31 (Edgar 2004). The alignment of 90 OTUs
was subsequently edited and fine-tuned using MacClade
4.08 (Maddison and Maddison 2005). Garli0.951-GUI
(www.bio.utexas.edu/faculty/antisense/garli/Garli.html) was
used to analyse the 90-sequence alignment (1,170
unambiguously aligned positions; gaps excluded) with
maximum-likelihood (ML). Jmodeltest 0.1.1 selected a
general-time reversible (GTR) model of nucleotide substitutions
(Posada and Crandall 1998) that incorporated invariable
sites and a discrete gamma distribution (eight
categories) (GTR + Γ + I model: a = 0.6430 and fraction of
invariable sites = 0.2160) under Akaike Information Criterion
(AIC) and AIC with correction (AICc.). ML bootstrap
analyses were performed on 500 pseudo-replicates, with
one heuristic search per pseudo-replicate (Zwickl 2006),
using the same programme set to the GTR
model + Γ + I. Bayesian analysis of the 90 OTU-dataset
was performed using the programme MrBayes 3.1.2
(Huelsenbeck and Ronquist 2001). The programme was
set to operate with GTR, a gamma-distribution, and four
Monte Carlo Markov Chains (MCMC; default temperature
= 0.2). A total of 10,000,000 generations were calculated
with trees sampled every 100 generations and with
a prior burn-in of 1,000,000 generations (10,000 sampled
trees were discarded; burn-in was checked manually).
When the average split fell below 0.01, the programme
would terminate. All other parameters were left at the
default. A majority rule consensus tree was constructed
from 90,000 postburn-in trees. Posterior probabilities correspond
to the frequency at which a given node was
found in the postburn-in trees.
RESULTS
The prevalence and abundance of gregarines within the
host were low; approximately 10% of crustacean hosts
were parasitised with no more than 2–5 syzygies per
host. Mostly, head-to-tail syzygies with two partners
were found, while no solitary trophozoites, gametocysts
or other stages were observed. Both partners in syzygy
were of monocystid form, with the nucleus situated in
the middle of the cell. Primites (anterior syzygy partner)
were usually slightly curved, longer and more slender
than the satellite (posterior syzygy partner) (190–160 lm
vs. 140–150 lm, respectively) (Fig. 1A, B, 2A). The cytoplasm
of both partners was packed with a granular content,
corresponding to the grains of amylopectin. The
thin transparent cortical layer was seen on the lateral
sides of the cells; however, on the side of anterior ends
of the primite and satellite, and posterior end of the
satellite this zone was thicker (Fig. 1A, B). In contrast to
the satellite, the anterior end of the primite possessed a
transparent vacuolar zone (Fig. 1A–C). The nuclei of both
partners were ovoid in shape and each contained one
round nucleolus (Fig. 1D). The contact between the
primite and satellite was simple, without any visible
interdigitations at LM level (Fig. 1E). Only once we
observed a multiple association, when three differently
sized satellites attached to the posterior end of the
primite (Fig. 1F). All syzygies exhibited unidirectional gliding
motility.
Under transmission electron microscopy, the cells were
round in cross-section. The bulk of cytoplasm (=endoplasm)
was packed with amylopectin granules and the
zone of ectoplasm was very thin (Fig. 2B, C). The nucleus
possessed granular karyoplasm and an uneven nuclear
envelope (Fig. 2D, E). The parasites were covered with
numerous epicytic folds (about 1 lm in length and 0.1 lm
in width), which ran along the surface of the both partners
(Fig. 2A, C, F, 3A, D, E), and most of these folds were
wavy (Fig. 2F, 3H). The gregarine surface exhibited specific
areas where some folds formed a loop; i.e. these folds
went towards one of the ends, but reversed in the
Figure 1 Bright field light microscopic observations of Ganymedes
yurii. A. A general view of syzygy in the coronal plane, showing the
attachment point (at) with transparent vacuoles (v), nuclei (n), primite
(Pr) and satellite (Sa). B. A general view of syzygy with nuclei (n) in
sagittal plane. C. Anterior end of the primite. (D.) Ovoid nucleus with
one nucleolus. E. Contact between the primite (Pr) and satellite (Sa).
F. Multiple association of a primite (Pr) with three attached satellites
(Sa1-3). Scale bars: 50 lm.
© 2016 The Author(s) Journal of Eukaryotic Microbiology © 2016 International Society of Protistologists
Journal of Eukaryotic Microbiology 2017, 64, 56–6658
Morphology and Phylogeny of Ganymedes yurii sp. n. Diakin et al.
opposite direction (Fig. 2F). Such patterns were found in
all observed cells.
The three-layered pellicle was of typical apicomplexan
organisation, having a plasma membrane, underlain by
two closely adjacent membranes of the inner membrane
complex (imc) (Fig. 3A, B). The base of the folds was
underlain with a thick internal lamina, which formed
bridges with dense triangular rods (Fig. 3A, I). The internal
lamina, located underneath the pellicle, continued into the
folds, where it became thinner and more dense (Fig. 3A,
B). At the top of each epicytic fold, four to five rippleddense
structures could be observed, situated between the
plasma membrane and imc. An electron-dense rod was
also seen in the apex of epicytic folds, underlying the imc
(Fig. 3A, B). Typical micropores were rarely found
between the epicytic folds, they appeared as cylindrical
invaginations of the plasma membrane (approximately
40 nm in diameter, and 50 nm in length) ending with a
vesicle (approximately 50 nm in diameter). The cylindrical
part of micropore was enforced by the internal lamina;
however, no typical collar formed by imc was observed
(Fig. 3C).
In addition to the aforementioned typical micropores,
two different structures were observed in contact with
the pellicle of the gregarine (Fig. 3D–G). The first was
membranous vesicles. These were round. These were
round in shape, with a central, and electron-dense inclusions.
At the point of contact with the pellicle, the plasma
membrane did not form an invagination, but the imc
formed a cone-shaped, electron-dense collar, instead of
Figure 2 General morphology of Ganymedes yurii. A. General SEM
view of syzygy with a primite (Pr) and satellite (Sa). B. Transverse section
of a gamont, the bulk of the cell corresponds to endoplasm (end);
TEM. C. Higher magnification view showing the gamont’s cell cortex
with epicytic folds (ef), ectoplasm (ect) and endoplasm (end) filled
with amylopectin granules (ag); TEM. D. Tangential section of the
nucleus; TEM. E. Higher magnification view of the nuclear envelope;
TEM. F. Higher magnification view of the gamont surface covered by
longitudinal epicytic folds (ef) and looped epicytic folds (lf); SEM.
Scale bars: A = 50 lm; B = 20 lm; C, F = 2 lm; D = 5 lm;
E = 500 nm.
Figure 3 Organisation of cell cortex in Ganymedes yurii. A. General
view of epicytic folds formed by plasma membrane (pm), inner membrane
complex (imc) and internal lamina (il). Rippled-dense structures
(black arrows) were situated between pm and imc. Electron-dense
rod (ro) was found under imc in the fold apex. The internal lamina
formed triangular rod (tr) at the base of the folds; TEM. B. Higher
magnification of the top of epicytic fold, showing rippled-dense structures
(black arrows) and electron-dense rod (ro); TEM. C. Typical
micropore, showing the vacuole of micropore (vm) and cylindrical
enforcement (en). D. and E. General view of the cortex with nonfused
(ef) and fused (ff) epicytic folds, membranous vesicles (mv) with
cone-shaped collar (cc), and teardrop granular inclusions (gi); TEM. F.
Details of membranous vesicles with cone-shaped collar (cc); TEM. G.
High magnification of teardrop granular inclusions (gi); TEM. H. Scanning
electron micrograph showing fussed epicytic folds (ff) and starts/
ends of the fusion (white arrows); SEM. I. Cross-section of fussed
folds, showing the plasma membrane (pm) with inner membrane
complex (imc) and triangle rod (tr) at their base. Note the canals (ca)
formed between the fussed folds; TEM. J. Transversally sectioned
fussed folds with mucous (mu) material inside the canal; TEM. Scale
bars: A, I = 200 nm; B and C, F = 100 nm; D and E, G, J = 500 nm;
H = 2 lm.
© 2016 The Author(s) Journal of Eukaryotic Microbiology © 2016 International Society of Protistologists
Journal of Eukaryotic Microbiology 2017, 64, 56–66 59
Diakin et al. Morphology and Phylogeny of Ganymedes yurii sp. n.
cylindrical collar present in typical micropores (Fig. 3D, F).
The same membranous structures were also found in the
ectoplasm (Fig. 3E). The second structures (gi) that came
in a contact with the pellicle contained granular content
and were round or teardrop in shape; no membrane surrounded
these structures. No invaginations of the plasma
membrane or the imc collar were observed in this region
(Fig. 3E, G).
Fused epicytic folds were observed in cross-sections of
the cells. Commonly two or three folds fused along their
lateral surfaces (Fig. 3D, E, G–J). The fusion of the fold
was discontinuous throughout the length of the cell; some
folds were fused across a longer distance, while others
were only fused across a short distance (Fig. 2F, 3H). At
the point of fusion, only the imc membranes of two adjacent
folds were present. Usually, a triangular canal was
seen at the base of fused folds, while one or two narrow
canals in the middle were observed (Fig. 3D, E, G, I).
Occasionally, a mucus-like substance could be observed in
the space between fused folds (Fig. 3J).
During preparation of samples for SEM, some cells previously
associated in syzygy disassociated, so it was possible
to compare the surface morphology of anterior and
posterior ends of both the primite and satellite (Fig. 4A, B).
The anterior end of primite was convex and had an ovalshaped
attachment area with a wrinkled surface (Fig. 4A,
C). The epicytic folds near this attachment tip were straight
(Fig. 4C). Many small pores were visible on the surface of
attachment area (Fig. 4D). In longitudinal sections through
the anterior part of primite, the cytoplasm was subdivided
into several zones along the longitudinal cell axis (Fig. 4E).
The most anterior area (zone 1) was packed with heterogeneous
electron-dense roundish inclusions; this was presumed
to be condensed mitochondria. It was followed by
a concaved zone (zone 2) filled with various opaque and
transparent round inclusions (Fig. 4F). These two zones
were generally located eccentrically to the longitudinal axis
of the cell. The next zone was thicker and packed with various
vacuolar and granular inclusions, while the rest of the
cell was filled with numerous amylopectin granules
(Fig. 4E–G). The apical part of the primite was covered
with a typical three-membrane pellicle, underlain by a
dense, thick internal lamina. The internal lamina and imc
were interrupted at some points (Fig. 4F, inset), which corresponded
to the pores observed under SEM (Fig. 4D). At
the free posterior end of the satellite, wavy epicytic folds
converged in the centre, where a light depression was
usually observed (Fig. 4H–I). At the posterior end of the
satellite, a lentil-shaped granular zone was situated without
amylopectin granules (Fig. 4J).
While under light microscopy the contact site between
the primite and satellite appeared simple (Fig. 1E), electron
microscopic observations revealed its complex organisation
(Fig. 5, 6). Two collars were observed at the
posterior end of the primite. The innermost collar had an
uneven edge, and tightly adhered to the anterior most part
of the satellite. The outermost collar did not adhere to the
innermost one, and the epicytic folds running along the
surface of the primite merged to this collar (Fig. 5A, B,
6A, B). The junction site of the partners from the side of
the primite was crateriform, and was surrounded by the
aforementioned collars. The radial ridges were observed
on the surface of this area (Fig. 6A). The anterior end of
the satellite, which contacted the primite, was cap-like
and not surrounded by any collar (Fig. 6C, D). Usually, in
the centre, it had a light depression from which many narrow
grooves radiated. Small round and shallow caverns
were found on the surfaces between these grooves
(Fig. 6D).
Figure 4 Morphology of the primite’s anterior and satellite’s posterior
ends. A. General view of the primite with attachment tip (asterisk) at
the anterior end (Pra), and free posterior end (Prp); SEM. B. General
view of satellite disassociated from the primite during sample processing.
Anterior end (Saa) has a cap-like appearance, while the posterior
(Sap) is rounded; SEM. C. Details of the transition between the
surface of attachment tip (asterisk) and the rest surface covered with
epicytic folds (ef); SEM. D. High magnification view of the attachment
tip with pores (white arrowheads) on the surface; SEM. E. Cross-section
of the anterior part of the primite showing three zones (z1, z2,
z3) containing vacuoles; the bulk of the cytoplasm (bc) is packed with
amylopectin granules (ag); TEM. F. High magnification view of the
transition between the zones 1 (z1) and 2 (z2), white arrowheads
mark the pores; (insertion) – high magnification of pellicle showing
one pore; TEM. G. High magnification of the transition between the
zone 3 (z3) and the main bulk of the cytoplasm; TEM. Axial H. and lateral
I. views of the posterior end of the satellite; SEM. J. Longitudinal
section of the satellite’s posterior end, showing a transparent zone
(tz); the bulk of cytoplasm (bc) is filled with amylopectin granules (ag);
TEM. Scale bars: A, B = 20 lm; C, G = 2 lm; D = 1 lm; E, I,
J = 10 lm; F = 1 lm (insertion = 100 nm); H = 5 lm.
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Morphology and Phylogeny of Ganymedes yurii sp. n. Diakin et al.
The junction site of both syzygy partners was covered
with three-membrane pellicle, underlain with a thick internal
lamina. Occasionally, membranous structures resembling
“mv” (aforementioned) were found in the contact
with the pellicle of the junction area (Fig. 5E). The profile
of the ridges of the primite crateriform posterior end and
the grooves at the satellite anterior end corresponded to
each other (Fig. 5D, E). The cytoplasm of both ends, in
general, had the same organisation as what was described
previously. The posterior end of the primite exhibited the
same granular zone without any amylopectin granules, as
seen in the free posterior end of the satellite. However, in
the satellite, there was only a zone of dense granules
(zone 2), immediately followed by a cytoplasm filled with
amylopectin granules (Fig. 5C, D). As described previously,
the innermost collar tightly adjoined to the satellite
and possessed the rod consisting of fibrillar-like material.
In contrast, the outermost collar did not possess a rod,
but some epicytic folds merged with it (Fig. 5B, F, 6D).
The new SSU rDNA sequence generated from Ganymedes
yurii sp. n. had a 77% similarity to a closely related
species, Ganymedes themistos; 354 sites were mismatched
across their pairwise alignment of 1,553 bp.
Maximum-likelihood and Bayesian analysis (Fig. 7) generally
agreed with previous work in this field. Clades comprising
Cephaloidophoroidea (Rueckert et al. 2011),
Lecudinoidea (Simdyanov and Diakin 2013) (=Urosporoidea
in Cavalier-Smith 2014), Gregarinoidea (Clopton 2009) and
Actinocephaloidea (Cavalier-Smith 2014) were recovered.
Other selected gregarines including archigregarines, as
well as eugregarines from sipunculids and polychaetes,
were dispersed between these main clades. Additional
alveolate sequences were also grouped into the main
clades, namely cryptosporidia, rhytidocystids, coccidia, and
piroplasmids. The relationships between these clades
were unresolved with this current dataset.
The novel SSU rDNA sequence obtained from G. yurii
sp. n. was strongly affiliate with the SSU rDNA
sequence from a previously investigated species, namely
Ganymedes themistos. Both of these sequences were
grouped with robust support in the family Ganymedidae,
which is incorporated into the clade Cephaloidophoroidea,
comprising gregarines from crustaceans, as
well as phylogenetically related environmental
sequences. The sequences of two Ganymedes spp.
branched early within the entire clade Cephaloidophoroidea,
albeit, with low posterior probability and bootstrap
support (Fig. 7).
TAXONOMIC SUMMARY
Family Ganymedidae Huxley 1910
Genus Ganymedes Huxley 1910
Ganymedes yurii sp. n. (Fig. 1A, B, 2A)
Figure 5 Morphology of junction between the primite and satellite.
A. General view of the contact between the primite (Pr) and satellite
(Sa); SEM. B. High magnification view of the junction area, showing
the outer (oc) and inner (ic) collar; SEM. C. Cross-sectioned contact
zone showing the transparent zone (tz) of the primite’s posterior end
(Pr) and zone 2 (z2) of the satellite’s anterior end (Sa); TEM. D. Detail
of the junction; white arrows mark ridges of the primite’s posterior
end (Pr), black arrows mark grooves in the satellite’s anterior end
(Sa); TEM. E. High magnification view of the primite’s (Pr) ridges and
satellite’s (Sa) grooves coinciding to each other. Membranous vesicles
(mv) is present in the primite. Thick internal lamina (il) underlies the
pellicle of both the primite and satellite; TEM. F. Cross-section of the
periphery of the junction site between the primite (Pr) and satellite
(Sa), showing the outer collar (oc) bearing epicytic folds (ef) and the
inner collar (ic) with rod of fibrillar material. Scale bars: A, C = 10 lm;
B, D = 2 lm; E, F = 500 nm.
Figure 6 Surface morphology of the primite and satellite junction
ends after disassociation; SEM. A. General view of the crateriform
posterior end of the primite disassociated from satellite, showing
prominent outer collar (oc), inner collar (ic) and radial ridges (rr). B.
High magnification view of inner collar (ic), outer collar (oc) with epicytic
folds (ef). C. General view of the anterior end of a satellite
detached from the primite, showing a central depression (asterisk)
and grooves (gr) radiating from this depression. D. High magnification
view of the peripheral region of the anterior end of the satellite, showing
caverns (cav) and radial grooves (gr). Scale bars: A, C = 10 lm; B,
D = 2 lm.
© 2016 The Author(s) Journal of Eukaryotic Microbiology © 2016 International Society of Protistologists
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Diakin et al. Morphology and Phylogeny of Ganymedes yurii sp. n.
Figure 7 Combined maximum-likelihood (ML) and Bayesian interference tree derived from phylogenetic analyses of the 90 OTUs dataset. 1.170
unambiguously aligned sites) of small subunit (SSU) rDNA sequences. This tree was inferred using the GTR + Γ + I substitution model (Àln
L = 16,7382.83829, gamma shape = 0.62738, proportion of invariable sites = 0.2381). Numbers at the nodes denote the ML bootstrap percentage
(numerator) and Bayesian posterior probabilities (denominator). Bootstrap support values are listed above Bayesian posterior probabilities. Black dots
on branches denote the bootstrap support values and Bayesian posterior probabilities of 95/0.95 or higher, respectively. Bootstrap and Bayesian values
less than 55 and 0.95, respectively, were not added to this tree, the Bayesian values lower than 0.9 marked /“–”. The novel sequence generated
in this study is highlighted in a black box. Some branches were shortened by multiples of the length of the substitutions/site scale bar.
© 2016 The Author(s) Journal of Eukaryotic Microbiology © 2016 International Society of Protistologists
Journal of Eukaryotic Microbiology 2017, 64, 56–6662
Morphology and Phylogeny of Ganymedes yurii sp. n. Diakin et al.
Diagnosis. Syzygy caudo-frontal, the primite being slightly
curved and longer than satellite. Vacuolar region present
at the anterior end of the primite. Nucleus situated in the
centre of the cell. Dimensions of the primite varied
between 190–160 lm and satellite between 140–150 lm
(Fig. 1, 2A). The posterior end of the primite is crateriform
(cup-like), with two peripheral collars at the point of satellite
attachment (Fig. 4A, 5A, B, 6A). Satellites possess
cap-like anterior end (Fig. 4B, 6C). GenBank accession
number KU726617 of small subunit rDNA distinguishes G.
yurii from other investigated species.
Type host. Gondogeneia sp. Barnard 1972 (Arthropoda,
Crustacea, Malacostraca, Amphipoda, Caliopioidea, Ponto-
geneiidae).
Type host habitat. Cape Lachman, James Ross Island,
Weddell Sea, Antarctica (63°470
32″S, 57°460
86″W). Littoral
and upper sublittoral zone.
Type material deposition. Parasites on gold sputtercoated
SEM stubs have been deposited at the Dept. of
Botany and Zoology, Faculty of Science, Masaryk University,
Brno, Czech Republic (Fig. 2A, 4A, B, 5A, B, 6A, C).
Etymology. The name of this species refers to a Russian
name Yury (in the honour of first author’s father name).
Localisation in host. Intestinal lumen.
Gene sequence. Sequence of SSU RNA GenBank accession
number KU726617.
DISCUSSION
The genus Ganymedes contains several species of aseptate
eugregarines infecting various groups of crustacea.
All known gregarine species within the genus Ganymedes
are aseptate and form caudo-frontal syzygy. On the anterior
end of the primite, they have a transparent zone (balllike
structure), and most have a cup-like structure at the
posterior end, as it is described in this study. However,
some possess cup-like anterior ends of the satellite, as it
was described for G. themistos. Under light microscopy,
G. yurii differs from previous described species by its
form and size of the cells/partners in syzygy. Most of the
described species possess elongated and flexible cells
(Huxley 1910; Jones 1968, 1969; Prokopowicz et al. 2010;
Theodorides and Desportes 1972, 1975), in case of species
studied in this work, the syzygy partners are shortened
and are more or less rigid.
Surface morphology of G. themistos and G. yurii is similar;
both gregarines are covered by longitudinal epicytic
folds that run from one end of the cell to the other (as is
the case with most of eugregarines). However, G. yurii
possesses unique “looped folds” on the surface, the function
of which remains unclear. The ultrastructure of epicytic
folds was studied in several eugregarines from
crustacean hosts, namely Porospora portunidarum, Thiriotia
pisae, G. vibiliae, G. eucopiae, Uradiophora maetzi,
Cephaloidophora cf. communis, and Heliospora cf. longissima
(Desportes and Theodorides 1985; Desportes et al.
1977; Simdyanov et al. 2015). All of them show similar
morphology among folds, which appear club-shaped in
cross-section. At the top of each fold, there are several
rippled-dense structures (3–6), and in some species, the
12-nm filaments were observed. Many of the aforementioned
species, except C. cf. communis and H. cf. longissima,
have electron-dense rods in the apical part of the
fold; this has also been found in G. yurii. Similar structures
were also found in other eugregarines, for example, in
Gregarina steini, G. polymorpha and G. cuneata, parasites
of mealworm larvae (Valigurova et al. 2013), urosporids
Gonospora beloneides (Corbel et al. 1979), Urospora ovalis
and U. travisiae (Diakin et al. 2016). Therefore, it can be
assumed that this structure appeared (or was lost) in different
lineages of eugregarines independently. The function
of this dense rod is unknown; however, Valigurova
et al. (2013) suggested that in mentioned Gregarina spp.
the half-moon-shaped rod underlying the 12-nm filaments
could serve as a ‘skeleton’ reinforcing the apex of the
folds, which is in contact with the substrate while the gregarine
is gliding.
In G. yurii, we observed micropores typical for apicomplexans,
and in addition, two types of structures that differed
in their morphology making contact with the cortex
of the cell. However, there is no consensus regarding the
function of these structures (typical micropores and two
additional structures described in this study). Some
authors assume that micropores could take part in the process
of nutrient acquisition (Chobotar and Scholtyseck
1982; Scholtyseck 1973; Scholtyseck and Mehlhorn 1970;
Vivier et al. 1970), while others suppose that they are in
fact extrusomes, and function more in the process of
secreting mucus (Desportes and Schrevel 2013; Philippe
and Schrevel 1982; Valigurova et al. 2013; Vegni Talluri
and Dallai 1983). In the light of these studies, and our
recent observations, the following could be taken into consideration:
(i) typical micropores could serve in the process
of nutrient acquisition, and (ii) different structures
observed to be in contact with the pellicle correspond to
phases of mucous excretion. Previously, structures similar
to teardrop granular inclusions (gi) were described in several
species (Gregarina spp. and Urospora spp.). It is possible
that at different stages of the life cycle or during the
excretion process, these structures could look like vesicles
or ducts in transverse sections of the cells (Diakin et al.
2016; Valigurova et al. 2013). However, it is important to
mention that we did not observe any droplets of mucus
on the surface of the cell; nonetheless, mucus-like substances
between the laterally fused folds were detected.
Lateral fusions of epicytic folds are not unique to
G. yurii. In Porospora portunidarum, for example, some
folds are fused as well; however, it could be caused by
the presence of bacteria disposed between the folds (Desportes
et al. 1977). This phenomenon was also reported in
the septate gregarine Leidyana tinei (Leidyaniidae) (Vivier
et al. 1970); in which 2–3 folds were fussed (similar to
G. yurii). Similar fusion has been reported in Monocystis
agilis and M. herculea, (Monocystidae), aseptate eugregarines
found in the seminal vesicles of Oligochaetes. In
the case of M. herculea, 2–7 folds are fused at their tips;
in M. agilis folds are fused at their tips, and 2–3 lateral
points (Vinckier 1969; Vinckier and Vivier 1968). In all
© 2016 The Author(s) Journal of Eukaryotic Microbiology © 2016 International Society of Protistologists
Journal of Eukaryotic Microbiology 2017, 64, 56–66 63
Diakin et al. Morphology and Phylogeny of Ganymedes yurii sp. n.
mentioned cases, the canals between the folds were
formed. It was proposed that for monocystids the fusion
of the folds serves to reduce the friction force during
metabolic movement of the parasite between seminal
cells (Frolov 1991). In the case of intestinal gregarines, the
functionality of the fused folds remains unknown.
Pores localised on the surface of the gregarine attachment
site were described in several species: C. cf. communis,
Gregarina cuneata and G. yurii (Rueckert et al.
2011; Simdyanov et al. 2015; Valigurova 2012; present
study). In all cases, these pores exhibit similar morphology;
e.g. imc and the internal lamina are interrupted in this
region, while no obvious interruption or invagination of
plasma membrane is observable. Interestingly, only C. cf.
communis and G. yurii has heterogeneous inclusions situated
under the pellicle. Simdyanov et al. (2015) considered
these inclusions to be microneme-like structures, due to
their elongated shape. In contrast, the inclusions from
G. yurii were round. We assume that this could be condensed
mitochondria. Another possible explanation is that
these structures could represent vesicles varying in shape
and containing mucus-like substances or adhesive material,
which can be extruded outside the cell, facilitating the
parasites adhesion to the host, as it was shown in C. cf.
communis, G. cuneata, and some actinocephalid eugregarines
(Cook et al. 2001; Simdyanov et al. 2015; Valigurova
2012).
The ball-like structure described in all representatives
of the genus Ganymedes obviously corresponds to the
vacuolar zone found in G. yurii (most likely zone 3)
(Fig. 4E). The fine structure of anterior end of primite of
G. yurii appears simple in comparison to the previously
described gregarines from crustaceans, despite all of
them being septate. As it was described in this study,
the cytoplasm in the anterior end is subdivided into three
zones differing in structure, which are not separated from
each other by any fibrillar material (septum). The apical
end in septate species is also subdivided into three
zones; however, these zones are separated by a septum.
The number of septae varies in different species: e.g.
one septum separates the protomerite and deutomerite
in Heliospora cf. longissima, Callynthrochlamys phronimae;
while two septae divide the cell of Cephaloidophora
cf. communis into the epimerite, protomerite, and deutomerite
(Desportes and Theodorides 1969; Simdyanov
et al. 2015).
The fine structure of the contact site between the primite
and satellite of G. yurii is comparable to the previously
described syzygy contact zone in Callynthrochlamys phronima
(Desportes and Theodorides 1969). Both of these species
possess collars formed by the posterior end of
primite at the periphery of the contact site: two collars are
present in G. yurii, whereas C. phronima has a single collar.
The surface of the junction zone in the primite and
satellite are covered with a three-membrane pellicle lacking
epicytic folds; however, in G. yurii, radial ridges (on
the primite) and grooves (on the satellite) were documented.
In contrast to this, the anterior end of satellite in
G. themistos forms the collar, and only a small area lacks
epicytic folds, whereas the rest of the pellicle covering
the junction site is folded (Prokopowicz et al. 2010). In
Cephaloidophora phrosinae, only one of the partners (primite)
possesses modified epicytic folds in the contact zone
(with bifurcated or flattened tops), while the anterior end
of satellite has a smooth surface. It is assumed that the
role of these modifications is to increase the surface of
the contact area (Desportes et al. 1977).
Phylogenetic analyses of the novel sequence generated
from G. yurii supported a close relationship with
G. themistos, a previously described gregarine that was
also isolated from an amphipod. Both of these sequences
formed a basal clade within the large clade (Cephaloidophoroidea),
comprising other eugregarines from crustaceans
and some environmental sequences. As in
previous studies, this clade was long-branching, relative to
other clades of gregarines and apicomplexans (Prokopowicz
et al. 2010; Rueckert et al. 2011). It is generally
assumed that all gregarines so far described from crustaceans
are phylogenetically closely related to each other,
despite the wide range and global distribution of the
hosts. Nonetheless, the phylogenetic analyses of 18s
rDNA were unable to resolve the relationships along the
backbone and among many of gregarines isolated from
crustaceans hosts (Simdyanov et al. 2015).
In conclusion, the new species described in this study,
G. yurii, shares common features with G. themistos and
with other investigated gregarine species from crustaceans.
But, there exists specific regions throughout the
sequence that clearly distinguish each species among
members of the genus Ganymedes, and gregarines from
other crustaceans, in general. While all crustacean gregarines
share two common features: (i) all of them are intestinal
parasites of crustaceans, and (ii) phylogenetic analyses
group these isolates in a single clade – Cephaloidophoroidea.
Despite this, it is difficult to resolve the phylogenetic
backbone and morphological trends within this group. To
solve this problem, further investigations focusing on
molecular phylogeny in connection with ultrastructural
studies are needed.
ACKNOWLEDGMENTS
Authors are grateful to all participants of the expedition to
The Johann Gregor Mendel Station that took place in
2013. Special thanks to Doc. RNDr. Martin Vacha, Ph.D.
(Dept. of Experimental Biology, Faculty of Science,
Masaryk University) for help in material collection and host
manipulation. Many thanks to Stanislav Malavin (Zoological
Institute of Russian Academy of Science, Saint-Petersburg)
for determination of the host. AD thanks Timur G.
Simdyanov and Kirill V. Mikhailov for helpful consultations.
AD and AV are greatly indebted to the members of the
Laboratory of Electron Microscopy (Institute of Parasitology,
BC ASCR in Ceske Budejovice) for technical assistance.
The authors acknowledge the financial support
from a project ECIP – Centre of Excellence from Czech
Science Foundation No. GBP505/12/G112, and Dr. Aika
Yamaguchi for assistance with genetic sequencing.
© 2016 The Author(s) Journal of Eukaryotic Microbiology © 2016 International Society of Protistologists
Journal of Eukaryotic Microbiology 2017, 64, 56–6664
Morphology and Phylogeny of Ganymedes yurii sp. n. Diakin et al.
CONFLICT OF INTEREST
The authors declare no conflict of interest.
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Morphology and Phylogeny of Ganymedes yurii sp. n. Diakin et al.
7.10
Kováčiková M., Simdyanov T.G., Diakin A., Valigurová A.
2017
Structures related to attachment and motility in the marine
eugregarine Cephaloidophora cf. communis (Apicomplexa)
European Journal of Protistology 59, 1-13
Available online at www.sciencedirect.com
ScienceDirect
European Journal of Protistology 59 (2017) 1–13
Structures related to attachment and motility in the marine eugregarine
Cephaloidophora cf. communis (Apicomplexa)
Magdaléna Kováˇcikováa,∗, Timur G. Simdyanovb, Andrei Diakina, Andrea Valigurováa
a
Department of Botany and Zoology, Faculty of Science, Masaryk University, Kotláˇrská 2, 611 37 Brno, Czech Republic
b
Department of Invertebrate Zoology, Faculty of Biology, Lomonosov Moscow State University, Leninskiye Gory 1-12, Moscow 119234,
Russian Federation
Received 2 December 2016; received in revised form 7 February 2017; accepted 28 February 2017
Available online 7 March 2017
Abstract
Gregarines represent a highly diversified group of ancestral apicomplexans, with various modes of locomotion and hostparasite
interactions. The eugregarine parasite of the barnacle Balanus balanus, Cephaloidophora cf. communis, exhibits
interesting organisation of its attachment apparatus along with unique motility modes. The pellicle covered gregarine is arranged
into longitudinal epicytic folds. The epimerite is separated from the protomerite by a septum consisting of tubulin-rich filamentous
structures and both are packed with microneme-like structures suggestive of their function in the production of adhesives
important for attachment and secreted through the abundant epimerite pores. Detached trophozoites and gamonts are capable of
gliding motility, enriched by jumping and rotational movements with rapid changes in gliding direction and cell flexions. Actin in
its polymerised form (F-actin) is distributed throughout the entire gregarine, while myosin, detected in the cortical region of the
cell, follows the pattern of the epicytic folds. Various motility modes exhibited by individuals of C. cf. communis, together with
significant changes in their cell shape during locomotion, are not concordant with the gliding mechanisms generally described
in apicomplexan zoites and indicate that additional structures must be involved (e.g. two 12-nm filaments; the specific dentate
appearance of internal lamina inside the epicytic folds).
© 2017 Elsevier GmbH. All rights reserved.
Keywords: Actin; ␣-Tubulin; Apicomplexa; Cell motility; Eugregarine; Myosin
Abbreviations: AB, alcian blue; CLSM, confocal laser scanning
microscopy; FITC, fluorescein isothiocyanate; IFA, indirect immunofluorescent
assay; IMC, inner membrane complex; LM, light microscopy;
PBS, phosphate buffered saline; PFA, paraformaldehyde; RR, ruthenium
red; SEM, scanning electron microscopy; TEM, transmission electron
microscopy; TRITC, tetramethylrhodamine isothiocyanate.
∗Corresponding author. Fax: +420 549 49 8331.
E-mail address: kovacikova@sci.muni.cz (M. Kováˇciková).
Introduction
Apicomplexans (Apicomplexa Levine 1980, emend. Adl
et al. 2012) are one of the most investigated group of protists,
comprising exclusively parasites of vertebrates as well
as invertebrates. Besides important pathogens of human and
agricultural animals (Toxoplasma gondii, Plasmodium spp.,
Cryptosporidium spp., and Eimeria spp.), this group comprises
highly diversified basal lineages, including gregarines.
Gregarines are obligate parasites that inhabit a wide range of
http://dx.doi.org/10.1016/j.ejop.2017.02.006
0932-4739/© 2017 Elsevier GmbH. All rights reserved.
2 M. Kováˇciková et al. / European Journal of Protistology 59 (2017) 1–13
terrestrial, freshwater, and marine invertebrates and urochordates
(Desportes and Schrével 2013). In particular, gregarines
occurringinmarineenvironmentshaveretainedspecificcharacteristics
inferred to be ancestral and are considered to be
deep-branching apicomplexans. Molecular phylogenetic evidence
is concordant with this interpretation (Leander 2008).
In general, the development of gregarines takes places in
the digestive tract, reproductive organs and body cavities of
their hosts, hereby explaining their varied shapes and motility
modes. Gregarine trophozoites are attached to host tissue by
a specialised apical part forming a mucron or an epimerite
(Schrével and Desportes 2015). After the vegetative phase of
their life cycle, trophozoites detach and transform into sexual
stages, called gamonts, which are usually motile (Schrével
and Desportes 2015).
The invasive stages (zoites) of apicomplexans are characterised
by a unique set of organelles called the apical
complex. Their pellicle consists of a plasma membrane which
is underlain by a closely apposed inner membrane complex
(IMC). The pellicle is associated with numerous cytoskeletal
elements such as microtubules, a network of intermediate
filament-like proteins, actin, and myosin (Morrissette and
Sibley 2002). While motile apicomplexan zoites are reported
to employ a unique mechanism of substrate-dependent gliding
facilitated by a conserved form of the actomyosin motor
(the so called glideosome concept) first proposed for Toxoplasma
gondii and Plasmodium spp. (Kappe et al. 2004;
Keeley and Soldati 2004; Opitz and Soldati 2002), the precise
machinery involved in the motility of apicomplexan ancestral
groups remains unclear. Indeed, in gregarine trophozoites
and gamonts, several types of motility have been described.
For example, the pendular or rolling movement in archigregarines
of the family Selenidiidae Brasil, 1907 seems to
be facilitated by regular sets of subpellicular microtubules
(Schrével et al. 1974; Stebbings et al. 1974). Meanwhile, the
motility mode mostly displayed by intestinal eugregarines is
a form of progressive linear gliding usually without obvious
changes in cell shape (King 1981, 1988) and accompanied by
the secretion of mucus leaving a trail behind (Mackenzie and
Walker 1983; Valigurová et al. 2013; Walker et al. 1979). The
pellicle of most eugregarines forms numerous longitudinal
epicytic folds. Electron microscopic analyses showed these
folds to form lateral undulations, which were suggested to
provide the force behind the gregarine gliding (Schrével and
Philippe 1993; Vávra and Small 1969; Vivier 1968). Actin
and myosin restricted to the cell cortex were identified in
eugregarines of the genus Gregarina Dufour, 1828, and their
involvement in gliding motility was proposed (Ghazali et al.
1989; Heintzelman 2004; Valigurová et al. 2013). As subpellicular
microtubules were not observed in the investigated
species of eugregarines, the mechanism of their gliding motility
must differ from that described by the glideosome concept
(Valigurová et al. 2013). Another type of motility, exhibited
by coelomic (Urosporidae Léger, 1892 and Monocystidae
Bütschli, 1882) and some intestinal eugregarines (e.g. Didymophyes
gigantea), is the so-called peristaltic or metabolic
movement (Desportes and Schrével 2013; Diakin et al. 2016;
Hildebrand and Vinckier 1975; Landers and Leander 2005;
Leander et al. 2006; MacMillan 1973). These diverse modes
of gregarine motility represent specific adaptations to parasitism
in different environments (Valigurová et al. 2013).
The present study focuses on the eugregarine
Cephaloidophora cf. communis (Cephaloidophoridae
Kamm, 1922) parasitising the barnacle Balanus balanus
Linnaeus, 1758 (Crustacea: Cirripedia). We performed
ultrastructural and immunological analyses of structures,
which are expected to be related to the attachment to host
tissue and to the unique motility mode displayed by the
trophozoites and gamonts of this eugregarine.
Material and Methods
The gregarine Cephaloidophora cf. communis was isolated
from the intestine of its marine crustacean host Balanus balanus.
Hosts were collected between 2013 and 2015 from the
White Sea environment close to the White Sea Biological Station
of Lomonosov Moscow State University (66◦33.190 N,
33◦06.550 E). Parasites were separated from host’s intestine
in filtered seawater using entomological needles and
then transferred to embryo dishes with a 30 mm cavity for
careful washing in filtered seawater and subsequent fixation
procedures. The dissection and manipulation of parasites
were performed using a stereomicroscope MBS-1 (LOMO,
Russia). For native preparations, individual gregarines were
put on microscope glass slides with filtered seawater and
their motility was monitored using a Leica DM 2000 light
microscope connected to a DFC 420 digital camera (Leica
Microsystems, Germany).
For transmission electron microscopy (TEM), specimens
were fixed in an ice bath in 2.5–3% (v/v) glutaraldehyde in filtered
seawater or in 0.15 M cacodylate buffer (pH 7.4). Some
specimens were fixed with 3% glutaraldehyde–ruthenium red
[0.15% (w/v) stock solution in Milli-Q water] in cacodylate
buffer or with 2.5% glutaraldehyde–alcian blue [1%
(w/v) stock solution in Milli-Q water] in filtered seawater.
Fixed samples were rinsed 3× for 20 min and post-fixed in
2% (w/v) OsO4 for 2 h in the same buffer as used for fixation.
After rinsing 3× for 20 min in the same fixation buffer
and after dehydration in an acetone series, specimens were
embedded in Epon (Polybed 812). Ultrathin sections were
obtained with diamond knives using a Leica EM UC6 ultramicrotome
(Leica Microsystems, Germany) and stained with
uranyl acetate and lead citrate. Sections were examined under
a JEM-1010 (Jeol, Japan).
For scanning electron microscopy (SEM), specimens were
fixed in 2.5–5% (v/v) glutaraldehyde in 0.2 M cacodylate
buffer (pH 7.4), washed 3× for 15 min in cacodylate buffer,
post-fixed in 2% (w/v) OsO4 for 2 h in the same buffer,
and finally washed 3× for 15 min in cacodylate buffer.
After dehydration in an acetone series, parasites were critical
M. Kováˇciková et al. / European Journal of Protistology 59 (2017) 1–13 3
point-dried with CO2, coated with gold, and observed using
a JEOL JSM-7401F (Jeol, Japan).
For confocal laser scanning microscopy (CLSM), specimens
were fixed in 4% paraformaldehyde (PFA) in 0.1 M
phosphate buffered saline (PBS) at room temperature for
45 minandthenwashed3×for15 minbeforefurtherprocessing.
Afterwards, the parasites were permeabilised in 0.3%
Triton X-100 (Sigma-Aldrich, Czech Republic) for 20 min.
For the direct fluorescent staining of F-actin, samples were
incubated overnight at room temperature with phalloidintetramethylrhodamine
B isothiocyanate (phalloidin-TRITC;
Sigma-Aldrich, Czech Republic) and then washed 3× for
10 min in 0.1 M PBS. For indirect immunofluorescence, the
samples were incubated overnight at 4 ◦C with the following
primary antibodies: mouse monoclonal IgG anti-actin raised
against Dictyostelium actin (provided by Prof. Dominique
Soldati-Favre), rabbit anti-myosin (smooth and skeletal, the
whole antiserum, dilution 1:5; Sigma-Aldrich, Czech Republic)
and mouse monoclonal anti-␣-tubulin (Clone B-5-1-2,
dilution 1:1000; Sigma-Aldrich, Czech Republic), all in
PBS with 0.1% BSA. After washing 3× for 10 min in
0.1 M PBS, the specimens were incubated at 37 ◦C for 4 h
with the following secondary antibodies: FITC-conjugated
anti-mouse (polyvalent immunoglobulins, dilution 1:125;
Sigma-Aldrich, Czech Republic) and TRITC-conjugated
anti-rabbit IgG (whole molecule, dilution 1:200; SigmaAldrich,
Czech Republic), both in PBS with 1% BSA. They
were then washed again. Controls were incubated with a
mixture of secondary antibodies alone, i.e. without primary
antibodies. Preparations were mounted in VECTASHIELD
Hard Set Mounting Medium (Vector laboratories, USA).
Samples were examined under an Olympus IX80 microscope
equipped with a laser-scanning Fluo View 500 confocal unit
(Fluo View 3.4 software; Olympus, Japan). Fluorescence was
visualisedusingtheTRITC(phalloidin,anti-myosin/544 nm)
and FITC (anti-actin, anti-␣-tubulin/457–515 nm) lasers sets.
Some micrographs obtained under CLSM were processed
using Fiji software (an image processing package based on
ImageJ developed at the National Institutes of Health).
Results
Light microscopic observations of gregarine motility.
The detached trophozoites and gamonts isolated from host
intestine exhibited a very active gliding movement (Video
S1, Supplementary material). When compared to gliding
motility in other eugregarines, the behaviour of C. cf. communis
during gliding differed in several aspects. In addition
to progressive and rapid gliding, these gregarines displayed
jumping or jerky movements, during which gliding was discontinuous
and combined with abrupt stopping (Video S2,
Supplementarymaterial).Moreover,rotationsaroundthelongitudinal
cell axis, rapid changes in direction (Video S3,
Supplementary material), and flexion in the area of the septum
separating the protomerite from the deutomerite and in
the first third of the deutomerite were observed during gliding
(Video S4, Supplementary material). In some cases, the
protomerite partially retracted into the deutomerite (Video
S4). In addition, the parasites were capable of gliding motility
even when contacting the microscopic slide via their
apical or posterior regions only (Video S5, Supplementary
material). Reverse movement was also observed (Video S6,
Supplementary material).
Electron microscopic analyses. Trophozoites of C. cf.
communis exhibited tricystid morphology; i.e. the cell
was dived into three morphologically distinct regions: the
epimerite, protomerite, and deutomerite (Fig. 1A–C). The
lenticular epimerite, situated in the middle of the protomerite
apical end, appeared as a thickened hyaline region when
observed under the light microscope (Fig. 1A). Detailed SEM
observations of the epimerite surface revealed a wrinkled
plasma membrane with numerous pores, lacking epicytic
folds. The epimerite of one specimen bore a long tiny protrusion
in its central part (Fig. 2A–C). The epimerite was
separated from the protomerite by a septum (Figs 1 A, 2
D–G).Thetubularstructures(20 ± 1 nmindiameter)forming
the septum were difficult to detect (Fig. 2G). Another distinct
septum separated the protomerite from the deutomerite,
inside which a prominent nucleus was situated (Figs 1 A, C,
2 D, 3 G). A superficial constriction was evident at the interface
between the protomerite and deutomerite in the area of
the septum (Figs 1 B, 2 A, 3 A, F). The cytoplasm of the
protomerite and deutomerite was divided into an inner granular
endoplasm packed with various cytoplasmic organelles
and inclusions, and an outer thinner hyaline ectoplasm lying
under the pellicle and free of amylopectin grains (Figs 1 A,
C, 4 A, B).
Both the epimerite and protomerite comprised abundant
electron lucent vesicles of unknown origin and function surrounding
the septum, giving this area a foam-like appearance
(Fig. 2D–F), and microneme-like organelles (Figs 2 E, F,
3 D, E). One SEM specimen with a mechanically ruptured
cortex revealed obvious differences between epimerite and
protomerite organisation. While the epimerite contained only
cylindrical organelles corresponding to the microneme-like
structures shown by TEM, numerous filamentous structures
were visible under the pellicle of the protomerite (Fig. 2H).
The typical gregarine pellicle of C. cf. communis, consisting
of a plasma membrane and an inner membrane complex
(IMC) (Fig. 3C), formed numerous longitudinally arranged
epicytic folds (Figs 1 B, 2 A). The folds of the protomerite
were lower in the gregarine apical end but their height
increased posteriorly (Fig. 3A, B, D). In contrast, the folds of
the deutomerite part were higher and their height was equal
along the entire deutomerite (Fig. 4A–C, E, F). The epicytic
folds exhibited an undulating appearance and new emerging
folds between the already existed ones were observed (Figs
1 B, 2 B, C, 3 A, B, F, 4 E–G, 5 A–F). A thick internal lamina
situated under the IMC linked the base of the folds, while a
thin layer of the lamina separated and extended to each fold
(Fig. 4C, D, F, G). A sparse network of filamentous structures
4 M. Kováˇciková et al. / European Journal of Protistology 59 (2017) 1–13
Fig. 1. General morphology of Cephaloidophora cf. communis trophozoites. A. A trophozoite exhibiting tricystid morphology. LM, bright
field. B. General view of a trophozoite. SEM. C. A longitudinal section of a trophozoite. TEM.
black arrow—septum separating the epimerite from the protomerite, black arrowhead—septum separating the protomerite from the deutomerite,
dm—deutomerite, double black arrowhead—constriction of the cell in the area of septum separating the protomerite from
the deutomerite, double white arrowhead—ectoplasm, ep—epimerite, n—nucleus, pm—protomerite, white arrow—epicytic folds, white
arrowhead—endoplasm.
(8 ± 1 nm thick) was detected inside the ectoplasm underlying
the internal lamina (Fig. 4D). The nature of this network
remains unknown, as, in some sections, these putative filaments
seemed to demarcate vesicle-like structures. Each
epicytic fold possessed in the apical part three rippled dense
structures with their bases located at the external cytomembrane,
and two poorly visible 12-nm filaments located under
the IMC (Fig. 4F, inset). The thin layer of internal lamina
extending into the folds and underlying the IMC had a specific
dentate appearance (5 ± 1 nm in diameter) (Fig. 4G,
H). In tangential sections, these dentations had a filamentous
appearance and were revealed to be organised angle-wise to
the epicytic fold longitudinal axis (Fig. 4H).
Typical apicomplexan micropores were present in the
grooves between the epicytic folds, distributed at irregular
distances from each other. Their duct (35–40 nm in diameter),
interrupting the IMC, was lined by a cone-shaped, electrondense
collar (120–130 nm in diameter). These micropores
were ending with a vesicle (Figs 3 B, 5 E–H). Staining with
alcian blue and ruthenium red confirmed the presence of a
glycocalyx layer forming a network along with an intense
secretion of mucus accumulating between the epicytic folds
(Fig. 5B, C). Further, amylopectin granules were clearly
visible in the cytoplasm of stained gregarines (Figs 4 B, 5 A,
C). Similarly, SEM observations showed mucus-like drops
to be situated at the folds’ apex and in the grooves between
them (Figs 4 E, 5 D, E). In the ectoplasm, just beneath the
pellicle, numerous cisternae containing mucous substances
were observed (Fig. 5C).
Confocallaserscanningmicroscopicanalysis.Thestaining
of ␣-tubulin showed a distinct ring structure located in the
area of the epimerite-protomerite septum (Fig. 6B–D, compare
with Fig. 6A). In addition, after ␣-tubulin labelling, a
strong fluorescent signal was visible in some parasites in a
form of a funnel-like structure extending from the epimerite
centre and ending at the septum (Fig. 6C–F). In other specimens,
the labelling of ␣-tubulin within the epimerite had the
appearance of concentric rings (Fig. 6H, I). Also, clusters
of putatively unpolymerised ␣-tubulin were dispersed in the
cytoplasm of some individuals (Fig. 6C, D, H, I).
Filamentous actin occurred in the entire cell, with a slightly
increasing intensity around both septa and in the area of the
nucleus (Figs 6 F–I, 7 A, B). In addition, specific antibody
labelling confirmed the localisation of actin in the gregarine
cytoplasm in the form of abundant clusters occurring espe-
ciallyintheepimerite-protomeriteregion(Fig.7B,C,E,F).A
few actin clusters were visible in the deutomerite cytoplasm
of some individuals, but their occurrence was mostly occasional
(Fig. 7A–C, E). Myosin was restricted exclusively to
the gregarine cortex (Figs 6 B, C, 7 C–F). Superficial optical
sections showed that the organisation of myosin followed the
pattern of longitudinally-organised epicytic folds (Fig. 7C,
D).
M. Kováˇciková et al. / European Journal of Protistology 59 (2017) 1–13 5
Fig. 2. Attachment region of Cephaloidophora cf. communis trophozoites. A. A front view of the cell showing the protomerite with
centrally located epimerite. SEM. B, C. Frontal (B) and lateral (C) views showing the epimerite covered by a wrinkled plasma membrane
with numerous pores and a central protrusion. SEM. D. Longitudinal section of the protomerite and anterior part of the deutomerite. TEM. E,
F. Higher magnification of the epimerite separated from the protomerite by a septum comprising electron lucent vesicles and microneme-like
structures. TEM. G. The area of the septum separating the epimerite from the protomerite. The inset shows a different view of the filaments
in the septum. TEM. H. A mechanically ruptured cell revealing differences in organisation between the epimerite and the protomerite. The
inset shows a detail of filamentous structures under the pellicle. SEM.
black arrow—septum separating the epimerite from the protomerite, black arrowhead—septum separating the protomerite from the deutomerite,
black asterisk—epimerite protrusion, dm— deutomerite, double black arrowhead—filamentous structures in the protomerite,
double white arrowhead—filaments in septum, elv—electron lucent vesicles, ep—epimerite, mns—microneme-like structures, n—nucleus,
pm—protomerite, white arrow—epicytic folds, white arrowhead—epimerite pores.
Discussion
In our study on C. cf. communis, only trophozoites and
gamonts were collected after dissection of the host intestine,
but previous studies reported also intracellular stages occurring
in the hyaline vacuoles of the enterocytes (Lacombe
et al. 2002). The parasites exhibited tricystid morphology,
with the attachment apparatus – the epimerite – in the apical
part of the cell. The epimerite was defined as a rudimental
and rounded or lenticular structure with evenly distributed
superficial pores (Rueckert et al. 2011; this study). In addition,
a long thin protrusion rising from the central part of the
epimerite was observed in one individual, probably facilitating
more stable attachment to the host gut tissue. So far, it
6 M. Kováˇciková et al. / European Journal of Protistology 59 (2017) 1–13
Fig. 3. Protomerite architecture in Cephaloidophora cf. communis. A. Protomerite and the apical part of deutomerite covered by dense
arrays of longitudinal epicytic folds. SEM. B. Higher magnification of epicytic folds covering the protomerite. SEM. C. The three-layered
pellicle. TEM. D. Higher magnification of protomerite epicytic folds in cross section. TEM. E. Detail of protomerite apical region showing
microneme-like structures. The inset shows microneme-like structures in detail. TEM. F. The constriction at the protomerite-deutomerite
septum. SEM. G. Septum separating the protomerite from the deutomerite. TEM.
black arrow—micropores, black arrowhead—septum separating the protomerite from the deutomerite, dm—deutomerite, double black
arrowhead—constriction of the cell in the area of septum separating the protomerite from the deutomerite, double white arrowhead—IMC,
elv—electron lucent vesicles, mns—microneme-like structures, pm—protomerite, white arrow—epicytic folds, white arrowhead—plasma
membrane.
cannot be disproven that the protrusion is only an artefact.
On the other hand, with regard to its thickness, this structure
seems extremely fragile; thus, it could be broken off during
the fixation procedure. Another possibility is that the protrusion
is retracted before fixation. Inside the apical parts of
the epimerite and the protomerite, numbers of electron dense
vesicles similar to micronemes were detected. These structures
are supposed to release a glutinous secretion through the
pores situated on the epimerite and therefore to be responsible
for parasite attachment (Simdyanov et al. 2015). The
phenomenon of gluing was noted during our observation of
C. cf. communis, where gregarines were capable of attaching
to the surface of cover slides. This adhesive ability is
also known in other eugregarines, where a dense adhesive
material likely produced by exocytic vesicles (Cook et al.
2001) is released through pore-like structures located at the
top of the protomerite or the attachment site and which is
thought to have the function of sticking to the host tissue
(Diakin et al. 2017; Valigurová 2012). Alternatively, these
poresmightplayaroleingamontnutrition(Valigurová2012).
The space inside the epimerite and protomerite is packed with
translucent vesicular structures of unknown origin and function.
These electron-lucent vesicular structures of foam-like
appearance are supposed to be formed by a highly developed
endoplasmic reticulum (Simdyanov et al. 2015).
In comparison with other eugregarines (Devauchelle 1968;
Devauchelle and Oger 1968; Tronchin and Schrével 1977;
Valigurová and Koudela 2005; Valigurová et al. 2009;
M. Kováˇciková et al. / European Journal of Protistology 59 (2017) 1–13 7
Fig. 4. Deutomerite architecture in Cephaloidophora cf. communis. A. Deutomerite and a part of protomerite in longitudinal section. TEM.
B. Deutomerite in transversal section. AB, TEM. C, D. Detail of cortex and ectoplasm organisation. TEM. E. Detail of deutomerite epicytic
folds. Note new folds appearing between the already existing ones. SEM. F. Higher magnification of deutomerite epicytic folds. Inset shows a
detail of the folds’ tip comprising three rippled dense structures and two 12-nm apical filaments. TEM. G, H. The dentation of internal lamina
inside the epicytic folds. White rectangle demarcates the superficial section of epicytic folds. TEM.
af—12-nm apical filaments, black arrow—dentation of internal lamina, black arrowhead—mucus drop, dm—deutomerite, double black
arrowhead—filamentous structures in ectoplasm , double white arrowhead—ectoplasm, il—internal lamina, n—nucleus, rds—ripple dense
structures, white arrow—epicytic folds, white arrowhead—endoplasm.
8 M. Kováˇciková et al. / European Journal of Protistology 59 (2017) 1–13
Fig. 5. Micropores, mucus secretion and glycocalyx in Cephaloidophora cf. communis. A. General view of posterior region showing
cytoplasm with inclusions and amylopectin granules and epicytic folds. RR, TEM. B. Glycocalyx layer forming a network between the
epicytic folds. RR, TEM. C. A view of the glycocalyx network and mucus accumulation between the folds and cisternae with mucous material
located under the pellicle. The inset shows cisternae demarcated by a white rectangle in more detail. AB, TEM. D, E. A view of mucus-like
drops situated at the folds’ apex and in the grooves between them along with the micropores. SEM. F. Superficial cross section showing the
cytoplasm and micropores. Inset shows the duct and electron-dense collar of micropores. TEM. G, H. Longitudinal sections of micropores.
TEM.
am—amylopectin , black arrow—micropores, black arrowhead—mucus, c—cisternae, d—duct, double white arrowhead—IMC, e—electrondense
collar, g—glycocalyx, vm—vesicle of micropore, white arrow—epicytic folds, white arrowhead—plasma membrane.
Valigurová 2012), the epimerite of C. cf. communis is
not retracted/discarded after trophozoite detachment. In
Cephaloidophora spp. the epimerite is also present during
the intracellular phase of trophozoites development
(Poisson 1924) and is covered by a pellicle (Simdyanov
et al. 2015), not only by a plasma membrane as in other
species. Hence, the question arises of whether this structure
represents an epimerite or is the modified apical end of the
protomerite dedicated to attachment or eventually to feeding,
as described in eugregarines from mealworms (Devauchelle
1968; Valigurová 2012). The remaining epimerite could
serve for the hypothetical reattachment of trophozoites to
younger host cells after the abandonment of senescing cells,
as suggested in some other eugregarines (Lucarotti 2000;
Valigurová et al. 2009; Valigurová 2012).
In addition, labelling with ␣-tubulin antibody detected a
ring structure located in the area of the septum separating
the epimerite from the protomerite. In TEM sections, this
M. Kováˇciková et al. / European Journal of Protistology 59 (2017) 1–13 9
Fig. 6. Distribution of ␣-tubulin, myosin and F-actin in Cephaloidophora cf. communis trophozoites. A. General view of a PFA-fixed
trophozoite. LM. B, C. Distribution of ␣-tubulin (FITC) forming a ring structure in the region of the epimerite-protomerite septum and the
localisation of myosin (TRITC). CLSM, IFA. D, E. Localisation of ␣-tubulin (FITC). CLSM, IFA. F. Double labelling of ␣-tubulin (FITC)
and F-actin (TRITC). CLSM, IFA/phalloidin-TRITC. G. Localisation of F-actin (TRITC). CLSM, phalloidin-TRITC. H, I. Double labelling
of ␣-tubulin (FITC) and F-actin (TRITC). CLSM, IFA/phalloidin-TRITC. F: CLSM in combination with transmission LM.
black arrow—septum separating the epimerite from the protomerite.
structure was composed of tubular elements (20 ± 1 nm in
diameter). The above mentioned ␣-tubulin-rich filamentous
structures could be responsible for the epimerite controlling
the attachment to and detachment from host tissue in accordance
with the theory of epimerite retraction (Devauchelle
1968; Valigurová and Koudela 2008; Valigurová et al. 2009;
Valigurová 2012). The presence of ␣-tubulin in clusters
localisedingregarinecytoplasm,indicatestheunpolymerised
form of tubulin. In contrast, neither positive ␣-tubulin
labelling nor tubular structures on ultrathin sections were
detected in the septum separating the protomerite from the
deutomerite.
Micropores are suggested to have a role in cell surface
nutrition after the detachment of trophozoites from the host
tissue (Warner 1968). In addition, their connection with cisternae
or ducts was demonstrated by freeze-etching analysis,
indicating that they might be related to mucus secretion
(Valigurová et al. 2013). Superficial observations under SEM
confirm the presence of mucus drops in between and on the
top of epicytic folds. In addition, a strongly stained network
was detected between the epicytic folds after the application
of fixatives binding to mucopolysaccharides, thereby indicating
the presence of a glycocalyx layer covering the gregarine
surface. The presence of glycocalyx layer (glycoconjugates)
10 M. Kováˇciková et al. / European Journal of Protistology 59 (2017) 1–13
Fig. 7. Distribution of actin and myosin in Cephaloidophora cf. communis trophozoites. A, B. Co-localisation of actin (FITC) and F-actin
(TRITC) in a single optical section (A) and composite picture (B). CLSM, IFA/phalloidin-TRITC. C. Composite picture of actin (FITC)
clusters accumulated within epimerite and protomerite cytoplasm and myosin (TRITC) restricted to the cell cortex. CLSM, IFA. D. Higher
magnification showing myosin (TRITC). CLSM, IFA. E, F. Localisation of actin (FITC) clusters and myosin (TRITC) in single optical
sections. CLSM, IFA. A, E: CLSM in combination with transmission LM.
was documented from the surface of different gregarine
species (Philippe et al. 1979; Simdyanov and Kuvardina
2007). Glycocalyx allows the parasite to interact with and
respond to its external environment and represents an important
protective barrier against hostile forces (Guha-Niyogi
et al. 2001).
The epicytic folds covering the C. cf. communis surface
exhibit an undulating arrangement and typical architecture
as described in other eugregarine species (Desportes and
Schrével 2013; Reger 1967; Schrével et al. 1983; Simdyanov
et al. 2015; Vávra and Small 1969; Valigurová et al. 2013;
Walkeretal.1984).Itwasproposedthatpolymerisationofthe
12-nm filaments located in the tip of the epicytic fold under
the IMC plays a significant role in the gliding motility and cell
morphogenesis of eugregarine trophozoites with longitudinal
folds, whose lateral undulation could be mediated by an acto-
myosinsystem(SchrévelandPhilippe1993;VávraandSmall
1969; Vivier 1968). A more recent study on trophozoites
of Gregarina spp. showed that the number of 12-nm filaments
does not influence the speed of gregarine gliding, but
seems to control the direction of movement (Valigurová et al.
2013). According to observations on C. cf. communis possessing
only two 12-nm apical filaments in each epicytic fold
(Simdyanov et al. 2015; this study), it was also demonstrated
that the gliding path of species equipped with a low number of
12-nm filaments was rather semi-circular than linear and the
individuals glided with a relatively high speed. Interestingly,
each epicytic fold exhibited structures oriented angle-wise
to the longitudinal axis of the fold, which appeared in cross
section as dentations of the internal lamina, and have not so
far been described in other eugregarines. These structures
formed by internal lamina might possibly participate as a
scaffold strengthening the epicytic folds involved in gliding
motility.
In general, the glideosome, described in apicomplexan
invasive stages (zoites) (first announced for Toxoplasma
gondii and later for Plasmodium spp.), powers the gliding
motility essential for their migration to the appropriate location
in the host organism and for host cell invasion (Opitz
and Soldati 2002). In this model, an actomyosin motor is
expected to be embedded between the parasite plasma membrane
and the IMC, and to require a stable subpellicular
network of microtubules (Dubremetz et al. 1998; Kappe et al.
2004; Keeley and Soldati 2004; Matuschewski and Schüler
2007). Nowadays, the glideosome concept continues to be
redefined as new components are discovered, but the principal
mechanism still appears largely to be valid (Heintzelman
2015). Despite this, it has emerged that this mechanism cannot
be applied to basal apicomplexans such as gregarines
(Valigurová et al. 2013).
M. Kováˇciková et al. / European Journal of Protistology 59 (2017) 1–13 11
Eugregarines are usually capable of gliding motility, but
the exact mechanism behind their movement must differ sig-
nificantlyfromthegenerallyacceptedglideosomeconcept,as
the presence of subpellicular microtubules and micronemes
was, in the investigated species, disproven (Valigurová et al.
2013). In contrast to the substrate-dependent gliding of apicomplexan
zoites, eugregarines from the genus Gregarina
were able to move for a certain time without any contact with
the substrate (Valigurová et al. 2013). In our study, the gliding
movement of C. cf . communis also exhibited unusual characteristics
(e.g. additional jumping and rotational movements,
rapid changes in direction, and cell flexions) and, similarly
to other eugregarines, subpellicular microtubules were not
observed in the cortex.
The localisation of actin and myosin, which are considered
to constitute the main motility motor in apicomplexan
zoites, were studied within eugregarines mostly from the
genus Gregarina from terrestrial insect hosts and the genus
Lecudina Mingazzini, 1891 from marine polychaete hosts
(Baines and King 1989; Ghazali et al. 1989; Ghazali and
Schrével 1993; Heintzelman 2004; Valigurová et al. 2013).
The presence of filamentous actin fluorescently tagged with
phalloidin was studied only in gregarines parasitising mealworms.
It localised in their cortex, in the septum separating
the protomerite and deutomerite, and in the area of the
nucleus (Valigurová et al. 2009; Valigurová 2012; Valigurová
et al. 2013). In C. cf. communis, F-actin was distributed
homogenously throughout the entire gregarine, but particularly
in the area of both septa and in the position surrounding
the nucleus; however, the intensity of the fluorescence signal
was low. A different situation was described for other
apicomplexans, where F-actin was considered to be highly
unstable, forming short filaments (Matuschewski and Schüler
2007). For example, in T. gondii, actin filaments were not
detected without treatment by F-actin stabilising drugs (e.g.
jasplakinolide)andactinexistedprimarilyinitsglobularform
(Dobrowolski et al. 1997). In Gregarina spp., positive phalloidin
labelling gave an indication that the majority of actin
was present in filamentous form (Valigurová et al. 2013). On
the other hand, actin fluorescently localised with a monoclonal
antibody known specifically to recognise actin in T.
gondii and Plasmodium spp. showed only clusters of actin
predominantly distributed in the area of epimerite and protomerite
cytoplasm. This situation is similar to that described
in Gregarina cuneata, where a dot-like pattern of actin staining
was demonstrated in the protomerite region of mature
gamonts, suggesting that this actin could be responsible for
the increased flexibility of this region (Valigurová 2012). In
the present study, some individuals showed similar dot-like
actin staining with a less intensive signal in the deutomerite
region. The occurrence of actin clusters in the deutomerite,
however, was irregular and occurred mostly in smaller individuals.
This difference could be the result of features being
specific to particular developmental stages (younger parasites)
or eventually to particular gamont types (primite vs.
satellite), and could indicate the diverse intensity of their
motility. In previous studies (Heintzelman 2004; Valigurová
2012; Valigurová et al. 2013), the localisation of actin in
the eugregarine cortex was described with two distinct orientations
(parallel and perpendicular) within the parasite.
In spite of this, however, in our study the antibody used
for actin labelling in apicomplexans failed to detect cortical
actin. On the other hand, positive phalloidin labelling suggested
that the polymerised form of actin could play a role
in C. cf. communis gliding, as proposed for Gregarina spp.
(Valigurová et al. 2013). The presence of the XIV subclass of
unconventional myosins is characteristic of all so far investigated
Apicomplexa groups (Foth et al. 2006; Frénal et al.
2008), and structural homologies exist between the myosins
of gregarines and other apicomplexans (Heintzelman 2004).
Similarly to previous studies (Ghazali and Schrével 1993;
Heintzelman 2004; Valigurová et al. 2013), the myosin in C.
cf. communis was localised in the cell cortex, following a pattern
of longitudinal epicytic folds. The actomyosin complex
in eugregarines was assumed to be localised in the lateral
parts of epicytic folds (Valigurová et al. 2013).
Conclusion
The structural organisation of the attachment apparatus,
the epimerite, in the marine eugregarine C. cf. communis
exhibited a very specific pattern including numerous irregularly
distributed pores in the plasma membrane. These pores
are thought to release adhesives, most likely produced by
the microneme-like structures localised in the parasite apical
end and facilitating its adhesion to the host tissue. More stable
anchoring to the host is likely to be secured by the tiny
long protrusion rising from the epimerite centre, although
its presence and function in the eugregarine is ambiguous.
Accordingly, it may be the case that the attachment strategy
combines both mechanical and chemical means.
Despite the fact that the presence of the basic motor proteins,
actin and myosin, was demonstrated in this study, the
motility mode in C. cf. communis obviously differs from
the substrate-dependent gliding described for apicomplexan
zoites. This outcome is also supported by the absence of subpellicular
microtubules, which are essential components in
apicomplexan motor machinery. Actin, predominantly occurring
in polymerised form, was dispersed throughout the entire
cell, while myosin was localised to the gregarine cortex. Nevertheless,
the exact position of actin microfilaments as well
as the role of the putative actomyosin motor in eugregarines
remain to be elucidated. The unusually active and variable
mode of gliding motility in this gregarine is facilitated by the
architecture of its epicytic folds. Each fold comprises two 12nm
filaments responsible for the gregarine’s variable gliding
path, as previously proposed for Gregarina spp. (Valigurová
et al. 2013), and internal lamina which exhibits an extraordinary
dentate appearance. Furthermore, gliding is supported
by the intensive secretion of mucopolysaccharides densely
coating the entire gregarine surface.
12 M. Kováˇciková et al. / European Journal of Protistology 59 (2017) 1–13
Acknowledgements
Financial support was provided by the Czech Science
Foundation, project No. GBP505/12/G112 (ECIP). Financial
support was also provided by the Russian Foundation
of Basic Research Nos. 15-29-02601 and 17-04-02098. The
authors would like to thank the staff of the White Sea Biological
Station of Lomonosov Moscow State University (WSBS
MSU) for providing facilities for field sampling and material
processing. We are grateful to the Laboratory of Electron
Microscopy,BiologyCentreCAS,aninstitutionsupportedby
the Czech-BioImaging large RI project (LM2015062 funded
by MEYS CR), for their help with obtaining the EM data presented
in this paper. The authors are also extremely grateful
to Prof. Dominique Soldati-Favre (University of Geneva) for
providing the monoclonal anti-actin antibody.
Appendix A. Supplementary data
Supplementary data associated with this article can be
found, in the online version, at http://dx.doi.org/10.1016/
j.ejop.2017.02.006.
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7.11
Valigurová A., Vaškovicová N., Diakin A., Paskerova G.G., Simdyanov
T.G., Kováčiková M.
2017
Motility in blastogregarines (Apicomplexa):
Native and drug-induced organisation of Siedleckia nematoides
cytoskeletal elements
PLoS One 12(6), e0179709
RESEARCH ARTICLE
Motility in blastogregarines (Apicomplexa):
Native and drug-induced organisation of
Siedleckia nematoides cytoskeletal elements
Andrea Valigurova´1
*, Nadězˇda Vasˇkovicova´2
, Andrei Diakin1
, Gita G. Paskerova3
, Timur
G. Simdyanov4
, Magdale´na Kova´čikova´1
1 Department of Botany and Zoology, Faculty of Science, Masaryk University, Kotla´řska´ 2, Brno,
Czech Republic, 2 Institute of Scientific Instruments of the CAS, v. v. i., Kra´lovopolska´ 147, Brno, Czech
Republic, 3 Department of Invertebrate Zoology, Faculty of Biology, Saint-Petersburg State University,
Universitetskaya emb. 7/9, St. Petersburg, Russian Federation, 4 Department of Invertebrate Zoology,
Faculty of Biology, Lomonosov Moscow State University, Leninskiye Gory 1–12, Moscow, Russian
Federation
* andreav@sci.muni.cz
Abstract
Recent studies on motility of Apicomplexa concur with the so-called glideosome concept
applied for apicomplexan zoites, describing a unique mechanism of substrate-dependent
gliding motility facilitated by a conserved form of actomyosin motor and subpellicular microtubules.
In contrast, the gregarines and blastogregarines exhibit different modes and mechanisms
of motility, correlating with diverse modifications of their cortex. This study focuses
on the motility and cytoskeleton of the blastogregarine Siedleckia nematoides Caullery et
Mesnil, 1898 parasitising the polychaete Scoloplos cf. armiger (Mu¨ller, 1776). The blastogregarine
moves independently on a solid substrate without any signs of gliding motility;
the motility in a liquid environment (in both the attached and detached forms) rather resembles
a sequence of pendular, twisting, undulation, and sometimes spasmodic movements.
Despite the presence of key glideosome components such as pellicle consisting of the
plasma membrane and the inner membrane complex, actin, myosin, subpellicular microtubules,
micronemes and glycocalyx layer, the motility mechanism of S. nematoides differs
from the glideosome machinery. Nevertheless, experimental assays using cytoskeletal
probes proved that the polymerised forms of actin and tubulin play an essential role in the
S. nematoides movement. Similar to Selenidium archigregarines, the subpellicular microtubules
organised in several layers seem to be the leading motor structures in blastogregarine
motility. The majority of the detected actin was stabilised in a polymerised form and appeared
to be located beneath the inner membrane complex. The experimental data suggest
the subpellicular microtubules to be associated with filamentous structures (= cross-linking
protein complexes), presumably of actin nature.
PLOS ONE | https://doi.org/10.1371/journal.pone.0179709 June 22, 2017 1 / 29
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OPEN ACCESS
Citation: Valigurova´ A, Vasˇkovicova´ N, Diakin A,
Paskerova GG, Simdyanov TG, Kova´čikova´ M
(2017) Motility in blastogregarines (Apicomplexa):
Native and drug-induced organisation of Siedleckia
nematoides cytoskeletal elements. PLoS ONE 12
(6): e0179709. https://doi.org/10.1371/journal.
pone.0179709
Editor: Gordon Langsley, Institut national de la
sante´ et de la recherche me´dicale—Institut Cochin,
FRANCE
Received: September 13, 2016
Accepted: June 2, 2017
Published: June 22, 2017
Copyright: © 2017 Valigurova´ et al. This is an open
access article distributed under the terms of the
Creative Commons Attribution License, which
permits unrestricted use, distribution, and
reproduction in any medium, provided the original
author and source are credited.
Data Availability Statement: All relevant data are
within the paper and its Supporting Information
files.
Funding: AV, AD and MK were funded from the
project from Czech Science Foundation No.
GBP505/12/G112 (ECIP - Centre of excellence) and
acknowledge support from the Department of
Botany and Zoology, Faculty of Science, Masaryk
University, towards the preparation of this
Introduction
Apicomplexans (Apicomplexa Levine 1980, emend. Adl et al. 2012 [1]) belong to the most
monitored group of unicellular parasites. As many of them cause major human and animal
diseases, recent research has focused on the motility of apicomplexan invasive stages (zoites)
representing a potential target for chemotherapeutic intervention. Apicomplexan zoites are
characterised by a typical apical complex of organelles and a complicated cell cortex consisting
of a continuous plasma membrane underlined by cortical alveoli (inner membrane complex =
IMC). The IMC can be interrupted by micropores and connected with numerous cytoskeletal
elements such as actomyosin complex, microtubules and a network of intermediate filamentous
proteins [2, 3].
Although apicomplexans share a number of cytoskeletal structures with other eukaryotic
organisms, a number of remarkable differences makes them unique. First of all, apicomplexan
subpellicular microtubules are unusually stable and withstand high pressure, cold, and detergents
that are often used for their isolation, while actin filaments (F-actin) are extraordinarily
transient [2] and actin is present mostly in its globular form [4]. Except for the study demonstrating
the presence of long actin filaments in Theileria [5], apicomplexan microfilaments can
be usually observed only after treatment with F-actin stabilising drugs such as jasplakinolide
[2]. So far published studies, focusing mostly on Toxoplasma gondii and Plasmodium spp., concur
with the so-called glideosome concept applied for motile zoites, describing their unique
mechanism of substrate-dependent gliding motility facilitated by a conserved form of actomyosin
motor [6–10]. This motor is expected to be localised between the parasite plasma membrane
and IMC, and the gliding is based on the locomotion of myosin along actin filaments
together with the translocation of apically released adhesins to the parasite’s posterior end
[11]. The above-mentioned differences in the apicomplexan cytoskeleton correspond to this
machinery, which is based on and limited by the formation of transient actin filaments and
their fixation to the IMC, and requires a stabile subpellicular network of microtubules.
In contrast to vertebrate pathogens, the motility mechanism in early emerging groups of
Apicomplexa, such as lower coccidia and gregarines parasitising invertebrates and urochordates,
still remains unclear. The basal apicomplexans studied to date are covered by a typical
three-layered pellicle and use several mechanisms of motility, correlating with diverse modifications
of their cortex. Among these organisms, only a few model archigregarines and eugregarines
were investigated for specific aspects in their motility behaviour and related structures
[3, 4, 12–43]. These studies showed that gregarine locomotion differs from the substratedependent
gliding observed in apicomplexan zoites.
The present study focuses on the blastogregarine, Siedleckia nematoides Caullery et Mesnil,
1898 (Apicomplexa: Siedleckiidae), parasitising the polychaete Scoloplos cf. armiger from the
family Orbiniidae. Blastogregarines are characterised by a permanent multinuclearity and
complicated life cycle: gametogenesis goes through a budding of mononuclear or multinuclear
spherical bodies at the posterior end of parasites and their further transformation into macrogametes
and microgametes correspondingly [44]. Here, using a combined microscopic
approach, for the first time we present an experimental study on the motility of the apicomplexan
restricted to the marine invertebrate host.
Materials and methods
Material collection
The polychaetes Scoloplos cf. armiger (Mu¨ller, 1776), parasitised with Siedleckia nematoides,
were collected at the sand-silt littoral zone at the White Sea Biological Station of M. V.
Motility in blastogregarines (Apicomplexa)
PLOS ONE | https://doi.org/10.1371/journal.pone.0179709 June 22, 2017 2 / 29
manuscript. NV was supported by MEYS CR
(LO1212) and its infrastructure by MEYS CR and
EC (CZ.1.05/2.1.00/01.0017). TGS was supported
by the grants from the Council of President of the
Russian Federation NSh-7770.2016.4 and from the
Russian Foundation of Basic Researches 15-29-
02601. The funders had no role in study design,
data collection and analysis, decision to publish, or
preparation of the manuscript.
Competing interests: The authors have declared
that no competing interests exist.
Abbreviations: CLSM, confocal laser scanning
microscopy; EF, exoplasmic fracture face; FE,
freeze-etching; FITC, fluorescein isothiocyanate;
IFA, indirect immunofluorescent assay; IMC, inner
membrane complex; IMP(s), intramembranous
particle(s); JAS, jasplakinolide; Kp, partition
coefficient; LM, light microscopy; LP, large pore;
MP, medium pore; PF, protoplasmic fracture face;
PFA, 4% paraformaldehyde in 0.1 M phosphate
buffered saline; RR, ruthenium red; SD, standard
deviation; SE, standard error; SP, small pore; SEM,
scanning electron microscopy; TEM, transmission
electron microscopy; TRITC,
tetramethylrhodamineisothiocyanate.
Lomonosov Moscow State University (66˚33.1900
N, 33˚06.5500
E) and the Marine Biological
Station of Saint-Petersburg State University (66˚18.770’ N, 33˚37.715’ E), both situated
in the Kandalaksha Bay of the White Sea. The polychaetes were collected within the framework
of regular scientific work at White Sea Biological Station of M. V. Lomonosov Moscow
State University (WSBS), which is situated in the buffer zone of Kandalaksha State nature
reserve. According agreement between WSBS and the reserve, the biological station can collect
animals for the scientific work on its own territory and other sites situated in the buffer
zone of the reserve. The field sampling locality at the Marine Biological Station of SaintPetersburg
State University is not part of any national park or private territory, so no special
permission for their collection was required. The polychaetes S. armiger are not an endangered
or protected species in those regions. Animal capturing, handling and dissecting was
designed to avoid distress and unnecessary suffering. Parasitological dissection of polychaetes
and manipulation with parasites were performed using MBS-1 stereomicroscope
(LOMO, Russia).
Experimental motility assays and light microscopy
Parasites were treated with commercial membrane-permeable probes influencing the polymerisation
of actin—jasplakinolide (JAS, Invitrogen, Czech Republic) and cytochalasin D
(Invitrogen, Czech Republic), and microtubule-disrupting/antimitotic agents such as oryzalin
(Sigma-Aldrich, Czech Republic) and colchicine (Sigma-Aldrich, Czech Republic). As a
concentration of these probes lower than 5 μM has no obvious effect on gregarines [3], final
concentrations of 10 and 30 μM for JAS, cytochalasin D and oryzalin, and 10 and 100 mM
for colchicine in filtered (0.22 μm Millipore) seawater were applied to obtain reliable results
on vital parasites. Cytochalasin D, JAS and oryzalin were reconstituted in dimethyl sulfoxide
(DMSO) to prepare a 1 mM stock solution and diluted in filtered seawater to prepare
final working concentrations, while colchicine was reconstituted directly in filtered seawater.
Experimental assays that were processed for further microscopic analyses were performed
in embryo dishes with a 30 mm diameter cavity. For continuous light microscopic
observations of changes occurring during each assay, small pieces of host intestine with
attached blastogregarines were put on single cavity microscope glass slides with a drop of
drug diluted in filtered seawater. Controls were performed in filtered natural seawater and
corresponding concentrations of DMSO in filtered seawater. Embryo dishes with parasites
were kept in refrigerator with a temperature set point of 10˚C. Behavioural and morphological
changes of parasites induced by drugs’ application were monitored using a light microscope
Leica DM 2000 connected to a DFC 420 digital camera. To assess the parasites’ beat
frequency, the motility of S. nematoides trophozoites and gamonts attached to the host epithelium
was monitored at set time intervals. The number of beats performed over period of
30 seconds was counted by taking a sample of six (at least) randomly selected individuals.
Average time for each beat was calculated for the anterior-most region of the cell, where
waves develop.
Three repetitions of each experiment were performed in the course of three years. Each
year, for controls and every drug treatment, twelve fragments of intestines, with not less
than twenty parasites attached at the experiment beginning, were investigated. During
each experiment a fragment with attached parasites was periodically selected for video
recording of the parasite motility under the light microscope. At the end of each experiment,
these fragments were equally divided into three portions and fixed for microscopic
analyses (scanning and transmission electron microscopy, confocal laser scanning
microscopy).
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Electron microscopy
Specimens were fixed in an ice bath in freshly prepared 2.5–5% (v/v) glutaraldehyde either in
cacodylate buffer (0.05–0.15 M; pH 7.4) or in filtered seawater. For transmission electron
microscopy (TEM), the specimens were then washed 3×20 min in the same buffer as used for
fixation, and post-fixed in 1–2% osmium tetroxide (OsO4) in the same buffer for 1–3 h. Alternatively,
specimens were fixed with 3% glutaraldehyde-ruthenium red [0.15% (w/v) stock
water solution] in 0.2 M cacodylate buffer (pH 7.4) and post-fixed with 1% OsO4-ruthenium
red in the same buffer [45]. The following procedure was based on previously published protocols
[46]. Observations were made using a JEM-1010 (JEOL). For scanning electron microscopy
(SEM), the specimens were washed 3×15 min in the same buffer as used for fixation,
processed according to Valigurova´ et al. [46, 47] and examined using a JSM-7401F –FE SEM
(JEOL), GEMINI Zeiss Supra 40VP and REM LEO 420 (Zeiss).
Freeze-etching
Parasitised pieces of the intestine of freshly collected polychaetes, fixed at 4˚C in freshly prepared
5% (v/v) glutaraldehyde in 0.15 M cacodylate buffer (pH 7.4) or 2.5% (v/v) glutaraldehyde
in 0.1 M phosphate buffered saline, were washed in the same buffer and processed
according to Valigurova´ et al. [3] using the freeze-etching system device BAF 060 (BALTEC).
The replicas were cleaned with 5% sodium hypochlorite, 70% sulphuric acid and 50%
chromo-sulphuric acid, washed in distilled water and mounted on copper grids for examinations
using a transmission electron microscope Morgagni 268 D (FEI). Evaluation of
intramembranous particles (IMP) per a unit area (1 μm2
) and the size of IMP were performed
in ImageJ software. The nomenclature follows that proposed in Branton et al. [48]
and used in Schre´vel et al. [17].
Confocal laser scanning microscopy
Fragments of parasitised intestines were fixed for 45 min at room temperature in freshly prepared
4% paraformaldehyde in 0.1 M phosphate buffered saline (PFA) or in ice-cold methanol.
Samples were carefully washed before further processing and permeabilised for 15–40 min in
0.3–0.5% Triton X-100 (Sigma-Aldrich, Czech Republic). Protocols used for the direct staining
of filamentous actin with phalloidin–tetramethylrhodamine B isothiocyanate (phalloidinTRITC;
Sigma-Aldrich, Czech Republic) and indirect immunofluorescent antibody (IFA)
staining using the mouse monoclonal IgG anti-actin antibody that was raised against Dictyostelium
actin and recognises the actin in Toxoplasma and Plasmodium (provided by Prof. Dominique
Soldati-Favre), mouse monoclonal anti-α-tubulin antibody (Cat. No. T5168, SigmaAldrich,
Czech Republic), and rabbit anti-myosin antibody (Cat. No. M7648, Sigma-Aldrich,
Czech Republic) follow Valigurova´ et al. [46, 47]. All preparations were counterstained for
localisation of the cell nuclei with Hoechst 33342 (Molecular Probes, Czech Republic) and analysed
using an Olympus IX80 microscope equipped with a laser-scanning FluoView 500 confocal
unit (FluoView 4.3 software). Fluorescence was visualised using the TRITC (phalloidin,
anti-myosin), FITC (anti-actin, anti-α-tubulin) and/or UV (Hoechst) filter sets. All specimen
from one experimental assay (= particular staining and controls) were processed using the
same protocol, and micrographs from confocal laser scanning microscopy (CLSM) were
obtained under identical image capture conditions (filters, the laser intensity). Some micrographs
were processed using the Fiji software (an image processing package based on ImageJ
developed at the National Institutes of Health).
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Results
Observations of the motility of Siedleckia nematoides trophozoites and
gamonts
Parasites developed being attached between microvilli of the host intestinal epithelium. The
earliest observed stages of S. nematoides were the lancet-shaped early trophozoites (Fig 1A and
1B) exhibiting only barely visible pendular movement. The pendular movement of more
advanced stage, young trophozoites (Fig 1C), was more evident. The elongated and flattened
maturing trophozoites and gamonts (Fig 1D–1G) of S. nematoides exhibited very active types
of movement. In addition to the pendular and twisting motility of attached parasites, their
movement was typically wavy, with waves developing in the proximal region of the cell (just
behind the attachment area) and proceeding to its distal end, while the last third of the cell
appeared more rigid with limited mobility (S1 Video). Detached individuals either showed the
same kind of movement, or simply bent from side to side with movement initiated by the
proximal region (Fig 1H and 1I; S2 Video). A spasmodic movement was often observed in
physiologically stressed parasites (during experiments or prolonged observations under the
light microscope).
Ultrastructural analysis of the cortex organisation
Parasites attached to the enterocyte via the mucron; the cytoplasm in their anterior region contained
numerous rhoptries and micronemes (Fig 2A–2C). The surface of all observed S. nematoides
individuals, comprising developmental stages from early trophozoites up to gamonts,
appeared smooth, lacking any grooves or folds (Figs 1B–1F, 1I, 2C and 2D).
Ruthenium red staining revealed the presence of distinct glycocalyx layer covering the
entire parasite. This cell coat was evidently thicker in the parasite apical region (Fig 2A and
2C) than in its middle to distal part (Fig 2G) (apical part 85 ± 4 nm vs. distal part 26 ± 2 nm).
The parasite was covered by a typical apicomplexan pellicle consisting of a plasma membrane
and IMC (Fig 2E and 2F). The IMC consisted of external and internal cortical cytomembranes,
from which the former was usually poorly preserved. Only a few ultrathin sections showed
well preserved and closely apposed membranes of IMC (Fig 2E). More often, these two cortical
cytomembranes were separated from each other, thereby forming a translucent and optically
empty space between them (Fig 2F). Freeze-etching revealed that the cytomembranes were
unusually undulated, while the plasma membrane was almost smooth (Fig 2H and 2I).
Analysis of the supramolecular organisation of the plasma membrane showed that intramembranous
particles (IMP) are evenly distributed and arranged in a regular pattern (Fig
2H). The size of IMPs in S. nematoides pellicle membranes varied in the range from 0.5 to 21.3
nm, dependent on the membrane and its fractured face (Table 1). To compare our statistical
data with known data on other apicomplexans [17, 49, 50], we additionally analysed density of
IMP ranging from 6 to 14 nm (Table 2). The partition coefficient (Kp) was used as a tabulated
factor for specimen comparison.
Numerous pores, mostly organised in four lateral rows (two per each flattened side) running
parallel to the longitudinal cell axis, were observed (Fig 1D–1E). The distance between
two lateral rows was 2.77 ± 0.05 μm (= width of the cell per flattened side) and the distance
between individual pores in a row was in the range from 0.5 to 1.9 μm (0.94 ± 0.06 μm). While
the rows of pores were conspicuous in some specimens (Figs 1D and 3A), in others they were
less distinct (Fig 1E), or even not detected (Fig 1F and 1I). Young trophozoites did not exhibit
any pores at their surface under SEM (Fig 1B and 1C), but because of their sporadic presence
in our samples, it remains unclear whether the presence of pores is exclusively restricted to the
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Fig 1. General view of Siedleckia nematoides trophozoites and gamonts. A-B. An early trophozoite. C. A young
trophozoite. D. Composite micrograph of trophozoites attached to the host intestinal epithelium between microvilli and cilia.
Note the pores organised in longitudinal rows, two per each flattened side. E. Attached trophozoite with a smooth surface
showing the pores organised in rows. F. Attached gamont lacking the pores. G. Two parasites attached to the brush border of
the host intestinal epithelium. H. Composite micrograph showing the sequence of movement of a single detached parasite. I.
Detached parasite. A, G-H: LM, bright field; B-F, I: SEM. black asterisk–parasite apical end, h–host tissue, n–nucleus, white/
grey/black arrows–row of pores.
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older stages. At least three types of different sized pores were documented: small (16.6 ± 0.9
nm), medium (48.7 ± 3.2 nm), and large (123.0 ± 4.6 nm). The small and medium pores
occurred only in the fracture plane of both the cortical cytomembranes, but were never
Fig 2. Cortex organisation in Siedleckia nematoides. A. Apical end of a parasite attached to the host enterocyte. Note the well-developed layer
of glycocalyx. B. A detail of parasite apical end focusing on organisation of subpellicular microtubules. C. General view of parasite cross-sectioned
in the anterior region. D. General view of a parasite cross-sectioned in the middle region. E. The cross-sectioned pellicle with well-preserved and
adjacent cortical cytomembranes. F. Longitudinally-sectioned pellicle with obviously separated cortical cytomembranes. G. Cortex of parasite
cross-sectioned in the middle region. H. Protoplasmic fracture face of the plasma membrane with pores. I. Fractured plasma membrane and
cortical cytomembranes. A, C, G: RR TEM; B, D-F: TEM; H-I: FE TEM. black arrowhead–plasma membrane, black asterisk–rhoptry, black circle–
pore, double/paired black arrowhead–IMC, ei–EF of the internal cytomembrane, ep–EF of the plasma membrane, g–glycocalyx, h–host tissue, mi–
mitochondria, mv–mucronal vacuole, n–nucleus, pe–PF of the external cytomembrane, pp–PF of the plasma membrane, white arrowhead–
subpellicular microtubule, white arrows–pores, x—micronemes.
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detected in the plasma membrane. All three types of pores were present in each lateral row
(Fig 3C–3H). Their arrangement within rows was not regular, but the large pores usually alternated
with several medium- and small-sized pores (Fig 3D–3H). In the anterior parasite
region, the pores within a lateral row were organised in a single line (Fig 3F-top), while posteriorly,
the small- and medium-sized pores started to form double lines (Fig 3E, 3F-bottom and
3G and 3H). The large pores were usually connected to a vesicle, containing a coiled lamellar
structure or dense material and with a duct opening towards the IMC (Fig 3B and 3C). These
vesicles were seen below the subpellicular microtubules, while the duct was situated in the
plane of microtubule outermost layer (Fig 3B). The simple dense circle, visible in superficial
ultrathin sections showing the outermost layer of subpellicular microtubules, seems to correspond
to the vesicle duct, while the rosette-like organisation of dense particles could be a protein
bridge connecting the vesicle duct with the pore at the IMC (Fig 3D). In replicas revealing
the fractured IMC, the large pores appeared widely opened (Fig 3E). Interestingly, even in
areas of lateral rows, the signs of pores were only rarely observed in fracture faces of the plasma
membrane (Figs 2H and 3E). Besides the pores organised in four lateral rows, additional rows
and randomly distributed pores were observed (Fig 3E–3H).
The subpellicular microtubules arose from the apical pole (Fig 2A and 2B) and run to the
parasite posterior end. Their arrangement appeared to be slightly helically twisted along the
longitudinal cell axis (Fig 4A). Cross-sections showed the organisation of microtubules in
Table 1. The sizes and density of IMP in individual fracture faces of pellicle membranes in Siedleckia nematoides.
Membrane Face Size of IMP (nm) Density of IMP (particles/μm2
)
Mean Median SD SE Min Max Mean ± SE
Plasma membrane PF 7.3 7.0 2.9 0.1 0.5 21.3 3109 ± 90
EF 4.9 4.3 2.7 0.1 0.5 19.1 314 ± 56
External cytomembrane PF 6.8 6.6 3.1 0.1 0.5 19.1 2877 ± 213
EF 5.7 5.2 2.4 0.1 1.0 14.1 5758 ± 357
Internal cytomembrane PF 6.1 6.1 2.4 0.1 0.5 13.3 4352 ± 279
EF 4.9 4.6 1.9 0.1 0.5 12.1 3411 ± 260
A total number of all sizes of IMP in membrane fracture was used for density calculation. SD–standard deviation; SE–standard error.
https://doi.org/10.1371/journal.pone.0179709.t001
Table 2. Density of IMP (particles/μm2
) in different apicomplexans.
Species Plasma membrane External cytomembrane Internal cytomembrane
EF PF Kp PF EF Kp EF PF Kp
Gregarina blaberae1
977 ± 235 1469 ± 233 1.5 285 ± 39 133 ± 34 2.1 158 ± 72 297 ± 33 1.9
Gregarina cuneata 2770 ± 96 2244 ± 283 0.8 1420 ± 190 1260 ± 211 1.1 1502 ± 273 1993 ± 253 1.3
Gregarina polymorpha 2473 ± 147 1446 ± 158 0.6 602 ± 265 863 ± 202 0.7 814 ± 246 1276 ± 200 1.6
Gregarina steini 1783 ± 233 2265 ± 154 1.3 2588 ± 189 3820 ± 211 0.7 1886 ± 274 2339 ± 132 1.2
Eimeria nieschulzi2
218 ± 21 648 ± 73 3.0 2360 ± 133 29 ± 7 81.4 146 ± 31 1780 ± 97 12.2
Plasmodium knowlesi3
185 ± 25 2198 ± 528 11.9 1751 ± 228 38 ± 15 46.1 48 ± 28 574 ± 200 12.0
Siedleckia nematoides 183 ± 8 2926 ± 135 16.0 2745 ± 220 458 ± 15 6.0 797 ± 60 3342 ± 128 4.2
The size of IMP is in range 6–14 nm. Kp—partition coefficient defined as the ratio of number of particles per μm2
in the PF face/number of particles per μm2
in the EF face.
1
Values taken from [17]
2
Values taken from [49]
3
Values taken from [50].
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Fig 3. Distribution of the pores on the Siedleckia nematoides surface. A. Detail of pellicle surface with a well visible row of pores. B. Different
longitudinally-sectioned vesicular structures connected to the pellicle and corresponding to the pores observed by SEM. C. An almost superficial section
of a parasite revealing the pores and vesicles organised in row. D. Superficially-sectioned cortex showing the layer of subpellicular microtubules and a
row of pores of various size. E. Fractured pellicle revealing the row of differently sized pores located on the PF of the internal cytomembrane, but not
visible at the plasma membrane. F. A general view of the longitudinally fractured pellicle revealing the external cytomembrane with a lateral row of pores
and few randomly distributed pores. The large empty arrowheads with labels show the direction towards anterior (an) and posterior (po) parasite ends.
The inset shows the fractured pellicle and pores demarcated by black rectangle in more detail. G. Fractured pellicle showing pores organised in rows;
few pores are distributed randomly. H. A fragment of fractured pellicle where several rows of variously sized pores are visible. Inset shows a more
detailed view of area demarcated by black rectangle, with alternating small and large pores organised in row. A: SEM; B, D: TEM; C: RR TEM; E-H: FE
TEM. black arrowhead–plasma membrane, black arrows–additional row of pores, double/paired black arrowhead–IMC, ee–EF of the external
cytomembrane, ei–EF of the internal cytomembrane, pe–PF of the external cytomembrane, pi–PF of the internal cytomembrane, pp–PF of the plasma
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several layers; one of them was continual and located just beneath the IMC, while the other
intermittent layers were to be found deeper in the cytoplasm (Figs 2G and 4E–4H). The number
of microtubule layers significantly increased towards the parasite anterior region (Fig 2C).
In ultrathin sections, the outer diameter of the microtubules was 22.9 ± 0.3 nm and the inner
diameter was 9.8 ± 0.5 nm, while in replicas, the outer diameter was 31.8 ± 0.5 nm and inner
14 ± 1 nm. In cross-fractured replicas, the microtubules appeared as rosettes (Fig 4F and 4H),
and the diameter of the putative microtubule subunits corresponded to 5.8 ± 0.2 nm. Ultrathin
sections showed individual microtubules to be localised within more lucent areas (in contrast
to the surrounding cytoplasm), the so-called ‘chambers’, with a diameter of 35 ± 2 nm (Figs
2E, 2G, 4E and 4G). The distance between the IMC and microtubules forming the outer layer
was about 23.5 ± 0.7 nm.
In longitudinal sections of the microtubule layer, tiny filamentous structures were detected
between individual microtubules (Fig 4B). These structures, running parallel to the microtubules,
seemed to interact periodically with them via short oblique filamentous connections
(Fig 4B). More superficial sections revealed these connections as filamentous structures being
wound around each microtubule (Fig 4C). Ultrathin cross-sections as well as freeze-etching
data revealed a complex of two large and one small particles around each microtubule (Fig
4D–4H and 4J). The small particle (8.9 ± 0.6 nm in ultrathin sections, 8.8 ± 0.5 nm in replicas)
was embedded in the IMC and most likely serves as an anchor for an underlying microtubule
(Fig 4D–4G and 4J). One of the large particles (19.4 ± 1.2 nm in ultrathin sections, 15±1 nm in
replicas) interconnected the small particle and microtubules, and was localised within the electron-lucent
microtubule chamber. The second large particle was located diagonally to the first
one, below the microtubule (Fig 4D–4H and 4J). All these observations suggest that these electron-dense
particles may play role of cross-linking protein complexes that anchor the subpellicular
microtubules to the cytoplasmic face of the internal cortical cytomembrane, thereby
forming a series of microtubule-membrane bridges along entire length of each microtubule.
Filamentous structures, 3.2 ± 0.2 nm (max = 9.65 nm) thick in ultrathin sections and
5.6 ± 0.2 nm (max = 9.7 nm) thick in replicas, were seen connecting the plasma membrane
with the IMC (Fig 4G and 4I). Analysis of the cross-fractured pellicle confirmed the presence
of large particles between the plasma membrane and the external cortical cytomembrane (Fig
4L). In oblique fractured specimens, these appeared as short filaments situated in the supraalveolar
space (Fig 4K). In replicas, the glycocalyx was seen as an agglomeration of large particles
located on the external surface of the plasma membrane (Fig 4K and 4L).
Parasites’ motility and cortex organisation after treatment with
cytoskeletal drugs
To monitor the role of individual elements of the putative motility motor in S. nematoides, the
living parasites were treated with commercial probes influencing the de-/polymerisation of
cytoskeletal proteins. To investigate the involvement of subpellicular microtubules in parasite
motility, incubation of living parasites with oryzalin or colchicine (toxins causing the disruption
of the microtubules) was performed. To verify the essential role of actin microfilaments,
drugs with a contradictory effect, i.e. jasplakinolide (stabilises actin filaments and induces
actin polymerisation) and cytochalasin D (disrupts actin filaments and inhibits actin polymerisation)
were applied to living parasites. All experimental assays were performed on parasites
membrane, t–subpellicular microtubules, white arrows–lateral row of pores, white arrowheads–randomly distributed pores, white circles indicate some
of the large pores, white rectangle demarcates the doubled row of pores.
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Fig 4. Organisation of the subpellicular microtubules in Siedleckia nematoides. A. A superficial section of a cortex
revealing the pores and subpellicular microtubules being helically twisted along the longitudinal cell axis. B-C. Higher
magnification of the longitudinally-sectioned subpellicular microtubules. Note the rows of filamentous structures running
parallel to the adjacent microtubules (grey arrows) and filamentous connections with the microtubules (black arrows). D.
Cytoplasmic face of the internal cytomembrane with IMP alignments (white arrows) that correspond to the localisation of
subpellicular microtubules. E. The pellicle covering the anterior part of parasite, underlain by one continuous and several
intermittent layers of subpellicular microtubules sectioned in cross (left) and tangential (right) plane. F. The view (similar to E)
of fractured pellicle underlain with several layers of subpellicular microtubules. G. The cross-sectioned cortex in the middle
region of parasite, showing the organisation of subpellicular microtubules with cross-linking protein complexes. H. Fractured
subpellicular microtubules with cross-linking protein complexes. I. The detail of pellicle covered by a thick glycocalyx layer. J.
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attached to the host tissue and those that detached spontaneously in the course of each experiment.
Treated individuals survived in extremely high doses of all of the used cytoskeletal
probes and showed signs of motility for the next couple of hours (2 h in 10 mM colchicine, up
to 1 h in 100 mM colchicine, 8 h in 10 μM oryzalin, 5–7 h in 30 μM oryzalin, 8 h in 10 μM JAS,
6 h in 30 μM JAS, 8 h in 30 μM cytochalasin D and more than 9 h in 10 μM cytochalasin D)
(Table 3).
Despite extremely high concentrations of all of the applied cytoskeletal drugs, the surface of
parasites appeared undamaged (Figs 5A–5I and 6A–6H) as also seen in control parasites incubated
in seawater or proportionate concentrations (10 and 30 μM) of DMSO (data not
shown). Furthermore, after careful rinsing and returning of the treated parasites to pure seawater,
the majority recovered to normal motility in the period from 10 to 120 min (Table 3).
The treatment of parasites with antimicrotubule agents (colchicine and oryzalin) confirmed
the gradual disruption of microtubules, correlating with the increasing drug concentrations
and prolongation of the incubation period (Fig 5A–5I), and resulted in the complete blocking
of parasite motility. During the initial acceleration in parasite movement occurring in the first
10–20 min after drugs’ application, some of the parasites showed an oscillating movement (S3
Video). Afterwards, parasites gradually decreased their movement until they completely
stopped (Table 3). Drug-treated parasites exhibited irregular, spasmodic movement (mostly
turning over from side to side) and the majority of them laid on the surface of the host tissue.
In contrast to the very wavy movement of controls, attached parasites treated with antimicrotubule
probes exhibited less pendular and twisting movements; their bodies appeared to be
more rigid with obvious limitations in motility. The drug-induced cell rigidity first appeared
in the posterior half of the parasite which bended in the anterior-most region (S4 Video), and
afterwards the motility gradually ceased (S5 Video). During experiments with high doses of
oryzalin, the frequent detachment of parasites from host tissue has been observed. The drugtreated
parasites, especially those with larger dimensions (i.e. more mature), were more frequently
found to be spirally curled (Fig 5C), but no signs of cell damage or collapse were seen.
Ultrathin sections revealed that lower concentrations, i.e. 10 mM colchicine and 10 μM oryzalin,
induced a gradual depolymerisation of subpellicular microtubules: in oryzalin treated
parasites they appeared less distinct (Fig 5G), while in colchicine a few microtubules were
completely lacking, as easily seen in the otherwise continuous outermost microtubule layer
(Fig 5B). The majority of parasites treated with 10 μM oryzalin stopped moving after 8 h, while
this effect was seen after 2 h in 10 mM colchicine. Higher doses, 100 mM colchicine and
30 μM oryzalin, obviously disrupted microtubules more rapidly, as proved by a rapid decrease
of parasite motility within a considerably shorter time (20–60 min in 100 mM colchicine, 5–7
h in 30 μM oryzalin) and empty regions interrupting their outermost layer of subpellicular
microtubules (Fig 5D, 5E and 5I). In comparison to oryzalin, colchicine seemed to be more
efficient as it unambiguously caused the disruption of more than half of the microtubules
(viewed in cross-section) in a considerably shorter time period (1 h). At the end of experiment,
the number of subpellicular microtubules was 28.3 ± 1.9 per 1 μm of pellicle length in nonThe
high magnification of cross-sectioned microtubules partially revealing the organisation of tubulin protofilaments. K-L.
Various views of fractured pellicle revealing the cross-linking protein complexes. A, E, G, J: TEM; B-C, I: RR TEM; D, F, H,
K-L: FE TEM. black arrow–filamentous structures around subpellicular microtubules, black arrowhead–plasma membrane, c–
cytoplasm, double/paired black arrowhead–IMC, g–glycocalyx, grey arrow–filamentous structures located between individual
microtubules, grey arrowhead–protein complexes localised between the plasma membrane and IMC, iti–inner surface of the
true (= not fractured) internal cytomembrane, white arrow–protein complex embedded in the IMC, white arrowhead–
subpellicular microtubule. Black circles mark some of the large pores. White ellipse encircles the cross-linking protein
complexes anchoring the subpellicular microtubules to the internal cytomembrane.
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treated control parasites, 22.02 ± 1.9 in parasites treated in 10 mM colchicine, 11.6 ± 1.4 in 100
mM colchicine, 25.8 ± 1.3 in 10 μM oryzalin, and 20.6 ± 0.9 in 30 μM oryzalin.
When incubating with probes influencing actin polymerisation, the speed of parasite movement
increased during the first 5–40 min in 10 μM JAS and 5–15 min in 30 μM JAS (S6
Video), followed by a gradual decrease of the motility intensity to zero (in 10 μM JAS after 8 h
and in 30 μM JAS after 6 h) (Table 3). The movement of individuals treated with JAS appeared
spasmodic and irregular, with parasites bending from side to side (S7 Video). Blocked parasites
were twisted and completely non-motile. In cytochalasin D, after an initial increase of speed
(30–40 min in 10 μM cytochalasin D and 20–60 min in 30 μM cytochalasin D) (S8 Video), the
intensity of parasite movement gradually ceased (S9 Video) until it completely stopped in
Table 3. The treatment of living individuals of Siedleckia nematoides with cytoskeletal drugs.
Changes / Time left
after drug
application
Drug / Concentration
Colchicine Oryzalin Jasplakinolide Cytochalasin D
10 mM 100 mM 10 μM 30 μM 10 μM 30 μM 10 μM 30 μM
Initial increase of
movement speed
(0
compared to
control)
10 min 10 min 20 min 20 min ! 5 min ! 5 min ! 30 min ! 20 min
*0.55 ± 0.03
beats/s
**1.86 ± 0.11
s
*0.56 ± 0.01
beats/s
**1.78 ± 0.04
s
*0.57 ± 0.03
beats/s
**1.77 ± 0.10
s
*0.59 ± 0.09
beats/s
**1.85 ± 0.33
s
*0.59 ± 0.04
beats/s
**1.70 ± 0.09
s
*0.54 ± 0.04
beats/s
**1.99 ± 0.12
s
*0.61 ± 0.05
beats/s
**1.68 ± 0.11
s
*0.59 ± 0.05
beats/s
**1.79 ± 0.13
s
Oscillating
movement
+ ! 10 min + ! 10 min + ! 20 min + ! 20 min - - - First
documented
decrease of
movement speed
20 min 15 min ! 60 min ! 45 min ! 60 min ! 30 min ! 60 min ! 120 min
Δ Δ *0.40 ± 0.05
beats/s
**2.53 ± 0.32
s
*0.39 ± 0.04
beats/s
**2.64 ± 0.28
s
*0.41 ± 0.02
beats/s
**2.51± 0.12
s
*0.49 ± 0.06
beats/s
**2.42 ± 0.24
s
*0.43 ± 0.03
beats/s
**2.42 ± 0.20
s
*0.47 ± 0.04
beats/s
**2.33 ± 0.23
s
Progressive
decrease of
movement speed
! 30 min ! 20 min ! 120 min ! 60 min ! 120 min ! 60 min ! 300 min ! 240 min
*0.23 ± 0.03
beats/s
**4.88 ± 0.65
s
*0.21 ± 0.02
beats/s
**5.15 ± 0.46
s
*0.18 ± 0.02
beats/s
**5.92 ± 0.60
s
*0.22 ± 0.04
beats/s
**5.29 ± 0.69
s
*0.35 ± 0.04
beats/s
**3.51 ± 0.72
s
*0.35 ± 0.04
beats/s
**3.38 ± 0.61
s
*0.27 ± 0.05
beats/s
**5.19 ± 0.82
s
*0.22 ± 0.01
beats/s
**4.75 ± 0.27
s
Spasmodic
movement
(bending)
- - + + Restricted to
anterior region
+ + Prevailing
from side to
side
+ + Prevailing to
one side
Obvious cell
rigidity (especially
in posterior half)
+ + + + - - - -
Complete
stoppage of
motility
120 min 60 min 480 min 420 min 480 min 360 min ! 540 min 480 min
Recovery of
motility in majority
of
blastogregarines
after washing in
seawater
120 min 90 min ! 60 min ! 60 min ! 60 min ! 60 min ! 60 min ! 60 min
The symbol + indicates some observed change in the character of motility;—no obvious changes; ! changes appeared after the noted time period;
changes appeared only during the noted time period
* beat frequency (beats/s = in Hz equivalent to 1 beat cycle per second, average ± standard error of the mean)
** beat to beat interval (an average time between the two beats ± standard error of the mean)
Δ beat frequency not measurable due to high sample variability.
0
Control individuals continued beating for more than 9 h (entire experiment duration); they beat at a rate of 0.51 ± 0.02 beats/s with beat to beat interval of
2.18 ± 0.13 s.
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Fig 5. Organisation of the subpellicular microtubules in Siedleckia nematoides after treatment with cytoskeletal drugs. A-B. Treatment
with 10 mM colchicine for 2 h: A. Attached gamont. B. Cross-sectioned cortex with subpellicular microtubules. C-E. Treatment with 100 mM
colchicine for 1 h: C. Attached gamont. D-E. General view (D) and higher magnification (E) of the cross-sectioned cortex with subpellicular
microtubules. F-G. Treatment with 10 μM oryzalin for 8 h: F. Attached gamont. G. Cross-sectioned cortex with subpellicular microtubules. H-I.
Treatment with 30 μM oryzalin for 7 h: H. Attached trophozoite and gamont. I. Cross-sectioned cortex with subpellicular microtubules. A, C, F, H:
SEM; B, D-E, G, I: TEM. black asterisk–parasite apical end, black arrowhead–plasma membrane, double/paired black arrowhead–IMC, h–host
tissue, white arrowhead–subpellicular microtubule. White ellipses demarcate the regions with disrupted microtubules.
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Fig 6. Organisation of the cortex in Siedleckia nematoides after treatment with cytoskeletal drugs. A-B. Treatment with 10 μM JAS for 8
h: A. Attached gamont. B. Cross-sectioned cortex with subpellicular microtubules. C-D. Treatment with 30 μM JAS for 6 h: C. Attached gamont.
D. Cross-sectioned cortex with subpellicular microtubules. E-F. Treatment with 10 μM cytochalasin D for 9 h: E. Attached gamont. F. Crosssectioned
cortex with subpellicular microtubules. G-H. Treatment with 30 μM cytochalasin D for 8 h: G. Attached gamont. H. Cross-sectioned
cortex with subpellicular microtubules. A, C, E, G: SEM; B, D, F, H: TEM. black asterisk–parasite apical end, black arrowhead–plasma membrane,
double/paired black arrowhead–IMC, h–host tissue, white arrowhead–subpellicular microtubule. White rectangles highlight the reduced spacing
between microtubule layers.
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most individuals after 8 h when incubated with 30 μM cytochalasin D. Nevertheless, in 10 μM
cytochalasin D, although significantly suppressed and spasmodic, the majority of parasites
continued to move until the end of each experiment (more than 9 h). The ultrastructural
observations in parasites after the application of actin de-/polymerising agents did not allow
us to provide unequivocal interpretations (Fig 6A–6H). Ultrastructural analysis of parasites
treated with JAS indicated only moderate changes in contraction and condensation of filamentous
structures around the subpellicular microtubules (Fig 6B and 6D). Localisation of these
structures corresponds to the cross-linking protein complexes, consisting of proteins embedded
in the IMC and the network around subpellicular microtubules, which apparently anchor
the microtubules to the internal cytomembrane. With an increasing concentration of JAS (10
and 30 μM), these complexes seemed to contract, thereby changing the usually regular distribution
of subpellicular microtubules (Fig 6B and 6D). The most notable was the reducing or
vanishing of spacing between the outer continuous and inner discontinuous microtubule layers
(Fig 6B and 6D) (35.4 ± 0.4 nm in 10 μM JAS and 32.5 ± 0.2 nm in 30 μM JAS when compared
to 44.8 ± 0.8 nm in controls; calculated from two adjacent microtubule layers). In
individuals treated either with 10 or 30 μM cytochalasin D, density of all above-mentioned
structures did not significantly differ from the normal status (Fig 6F and 6H). However, similarly
to JAS, the spacing between the individual microtubule layers was reduced to 38.1 ± 1.3
nm in 10 μM cytochalasin D and 34.3 ± 1.1 nm in 30 μM cytochalasin D (Fig 6F and 6H). No
conspicuous changes have been documented in the density of protein complexes localised
between the plasma membrane and IMC in JAS or cytochalasin D-treated individuals (Fig 6B,
6D, 6F and 6H). The distance between the internal cytomembrane of the IMC and the outer
continuous microtubule layer, however, was significantly reduced (36.3 ± 0.2 nm in controls
vs. 22.5 ± 0.3 nm in 10 μM JAS, 23.7 ± 0.2 nm in 30 μM JAS, 23.0 ± 0.5 nm in 10 μM cytochalasin
D, and 25.3 ± 0.3 nm in 30 μM cytochalasin D).
Confocal laser scanning microscopic analysis of cytoskeletal structures
before and after treatment with cytoskeletal drugs
Fluorescence labelling was used to visualise the arrangement of cytoskeletal structures before and
after treatment of S. nematoides with cytoskeletal drugs. The myosin accumulated at the parasite
periphery (Fig 7A and 7C). Similar localisation was documented in α-tubulin immunolabelling
used for visualisation of the subpellicular microtubules (Fig 7B–7G). The labelling with an antiα-tubulin
antibody was also strongly positive for brush border of host intestinal epithelium. Colocalisation
of myosin and α-tubulin showed both proteins continuously distributed in the cell
periphery of young and mature parasites, with increasing labelling intensity towards the parasite
posterior region, and overlapped to some degree (Fig 7C). The superficial optical section in the
area of the parasite cortex showed patchy organisation of α-tubulin, just beneath the parasite pellicle,
organised in barely visible, tiny longitudinal lines corresponding to the subpellicular microtubules
(Fig 7D and 7G). Though, methanol-fixed individuals showed more intense labelling of
myosin and α-tubulin (Fig 7A–7D), the less intense fluorescence signal in PFA-fixed samples
allowed the pattern of cytoskeletal elements staining to be more precisely identified (Fig 7E–7G).
The labelling also revealed patchy distribution of α-tubulin in the posterior half of parasites (Fig
7E). Younger parasites exhibited obviously stronger labelling of α-tubulin.
Incubation with microtubule destroying agents resulted in an overall decrease of the fluorescent
signal for α-tubulin labelling (Fig 7H–7M). PFA-fixed samples were investigated for
the distribution of F-actin and α-tubulin to verify that the influenced microtubules were the
reason for the changes in parasite motility after the application of high doses of oryzalin or colchicine,
but not potential F-actin redistribution (= treated parasites exhibited no changes in FMotility
in blastogregarines (Apicomplexa)
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Fig 7. Distribution of myosin (TRITC), α-tubulin (FITC) and F-actin (TRITC) in Siedleckia nematoides before and after application of
cytoskeletal drugs. A–G. Non-treated parasites. A-B. Single optical section revealing the localisation of myosin (A) and α-tubulin (B). The inset
in B shows a localisation of α-tubulin in a caudal part of an individual from another optical section. C. Composite view showing the co-localisation of
myosin and α-tubulin in a single optical section of parasites shown in A-B. The inset shows a co-localisation of myosin and α-tubulin in a caudal part
Motility in blastogregarines (Apicomplexa)
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actin distribution; Fig 7E–7G vs. 7H–7M). Incubation with 10 mM colchicine for 2 h resulted in
a more diffuse pattern of α-tubulin labelling, which was especially apparent in the caudal region
of attached parasites (Fig 7H). Individuals treated for 1 h with 100 mM colchicine showed a further
decrease in tubulin labelling, the localisation of which remained restricted to the caudal
region only (Fig 7I and 7J). Similarly, parasites treated with 10 μM oryzalin for 8 h (Fig 7K)
showed less intensive staining of α-tubulin in contrast to non-treated ones. The intensity of αtubulin
labelling in parasites treated with 30 μM oryzalin for 7 or 5 h respectively was further
decreased (Fig 7L and 7M). The patchy distribution of α-tubulin organised in a line underlying
the pellicle corresponded to the localisation of subpellicular microtubules (Fig 7M-inset).
The phalloidin labelling confirmed the presence of F-actin in the blastogregarine cortex
and cytoplasm (Fig 8A–8D), with a slightly increased staining in the anterior half of the cell.
The pattern of staining in the caudal end exhibited a more spotted and fibrous pattern. The
brush border of the host epithelium also stained intensively with phalloidin. Parasites labelled
with the specific anti-actin antibody exhibited a patchy accumulation of actin (Fig 8C and 8D).
Actin was distributed almost homogenously in younger stages (Fig 8C), while in mature
gamonts (Fig 8D), it mostly accumulated in their middle part. The immunolocalisation of
actin differed from direct F-actin labelling with phalloidin in that the antibody did not bind to
the cell periphery corresponding to the cortex, and labelled the host tissue with intensity comparable
to the labelling of parasites. The treatment with 10 μM JAS for 8 h resulted in the overall
stronger immunolabelling of actin, with a spotted character being obvious, especially in the
caudal regions of parasites (Fig 8E). The phalloidin staining of F-actin in parasites showed no
or only a slight increase in fluorescence signal when compared to the control samples. The
incubation with 30 μM JAS for 6 h induced a more advanced stabilisation of actin filaments,
resulting in amplification of the fluorescence signal for phalloidin labelling (Fig 8F). These parasites
also exhibited a strong immunolabelling of actin, distributed within entire cell and often
more accumulated in parasite apical region. Parasites treated with 10 μM (9 h) or 30 μM (8 h)
cytochalasin D exhibited low or almost no F-actin labelling (Fig 8G and 8H), nevertheless the
signal for antibody labelling of actin did not change significantly.
Discussion
Blastogregarines, comprising a single genus Siedleckia, represent rather a problematic group of
unclear taxonomic position within Apicomplexa. Besides a unique life cycle [44, 51, 52], these
enigmatic organisms also exhibit unusually active motility, composed of pendular and twisting
movements.
The motility in gregarines vs. the concept of glideosome in apicomplexan
zoites
Typical movement generally described for apicomplexan zoites is a substrate-dependent gliding,
relying on dynamic turnover of actin, the unpolymerised form of which seems to have an
of an individual from another optical section. D. The localisation of α-tubulin in the superficial region of a gamont. E. Co-localisation of α-tubulin and
F-actin in parasites of various developmental stages. F. Composite view revealing the co-localisation of α-tubulin and F-actin in a macrogamont and
microgamont in a single optical section. G. More superficial optical section revealing the localisation of α-tubulin in macrogamont shown in F. H-J.
Co-localisation of α-tubulin and F-actin in parasites treated with colchicine: H. 10 mM colchicine (2 h). I-J. 100 mM colchicine (1 h). K-M. Colocalisation
of α-tubulin and F-actin in parasites treated with oryzalin: K. 10 μM oryzalin (8 h). L. 30 μM oryzalin (7 h). M. Labelling of α-tubulin
in a trophozoite treated with 30 μM oryzalin (5 h). The inset shows a co-localisation of α-tubulin and myosin in the trophozoite caudal region. Note
the patchy distribution of α-tubulin underlying the pellicle, corresponding to the localisation of subpellicular microtubules. A-C, M: CLSM, IFA/
Hoechst, methanol fixation; D: CLSM, IFA, methanol fixation; E-F: CLSM, IFA/phalloidin-TRITC, PFA fixation; G: CLSM, IFA, PFA fixation; H-L:
CLSM, IFA/phalloidin-TRITC/Hoechst, PFA fixation. black arrowhead–parasite caudal end, black asterisk–parasite apical end, h–host tissue, white
arrowheads–tiny longitudinal lines corresponding to the subpellicular microtubules.
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increased potential to form filaments relative to vertebrate actin [53]. This so-called glideosome
concept, so far described for zoites of Toxoplasma and few other apicomplexans [6–10], requires
coordinated interactions between surface adhesins and the cytoskeleton of the parasite. The
actomyosin motor is described as being embedded between the plasma membrane and the
IMC, and oriented by subpellicular microtubules [11]. The myosin A is linked to the IMC
against the subpellicular microtubules and its head moves along the actin filament connected to
a cell adhesion molecule (TgMIC2, TRAP, MIC2) [9, 10]. The complex of adhesins and actin filaments
is transported towards the posterior end of the cell. As a result, the actomyosin motor
generates cell gliding by transposing transmembrane adhesins through the plasma membrane
by bearing against the IMC [54]. The structure and function of the glideosome require re-evaluation,
as recent results have shown that numerous glideosome components (including actin and
myosin) can be knocked out without complete blocking of motility [55–57].
Fig 8. Phalloidin (TRITC) and antibody (FITC) staining of actin in Siedleckia nematoides before and after application of cytoskeletal
drugs. A-D. Non-treated parasites: A-B. Localisation of F-actin with phalloidin in parasites attached to the host tissue. C. Double labelling with
phalloidin and specific anti-actin antibody. D. Double labelling with phalloidin (left) and anti-actin antibody (right), image split into two separate
channels. E-F. Double labelling with phalloidin (left) and anti-actin antibody (right) in parasites treated with JAS, images split into two
separate channels: E. 10 μM JAS (8 h). F. 30 μM JAS (6 h). G-H. Double labelling with phalloidin (left) and anti-actin antibody (right) in
parasites treated with cytochalasin D, images split into two separate channels: G. 10 μM cytochalasin D (9 h). H. 30 μM cytochalasin D (8 h).
A-B, D-H left: CLSM, phalloidin-TRITC/Hoechst; C: CLSM, IFA/phalloidin-TRITC/Hoechst; D-H right: CLSM, IFA/Hoechst; A-H: PFA fixation. black
arrowhead–parasite caudal end, black asterisk–parasite apical end, h–host tissue.
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Moreover glideosome apparently cannot be applied to all apicomplexans; e.g., gregarines
exhibit diverse modes of locomotion and seem to use several mechanisms of cell motility. Different
modes of gregarine motility could represent specific adaptations to a parasitism in different
environments within hosts. Most eugregarines, covered by a cortex consisting of a
dense array of longitudinal epicytic folds and exhibiting progressive linear gliding, were shown
to use a modified machinery to move forwards on a solid surface, as they lack both the subpellicular
microtubules and micronemes [3]. In contrast, coelomic eugregarines move by pulsation
of their body corresponding to the peristaltic or metabolic motility accompanied with
periodic changes of the body shape [24, 26–33]. The spindle shaped trophozoites and gamonts
of archigregarines (Selenidium spp.), possessing regular sets of subpellicular microtubules, display
a bending, coiling, and rolling or pendular movements along with contracting their cell
shape [34–43, 58]. These movements could be also described as nematode-like [43, 58]. Both
the pendular and peristaltic movement are non-progressive.
Due to similarities in external morphology and movement patterns, genus Siedleckia has
been associated with Selenidium archigregarines [59]. Besides the pendular movement similar
to that in Selenidium [34, 36, 37], additional motility modes, such as twisting, undulation,
spasmodic and thrashing movements, could be observed depending on the stage and physiological
status of S. nematoides. Apicomplexan zoites also show variability in their movement;
e.g. circular/helical gliding (progressive) and twirling (non-progressive) of Toxoplasma and
Plasmodium zoites, but their motility relies on contact with a substrate [60]. Trophozoites and
gamonts of S. nematoides, however, despite bearing a striking resemblance with overgrown
apicomplexan zoites, showed no signs of gliding motility. Though the pendular/twisting movements
of S. nematoides might be considered to be reminiscent of twirling in Toxoplasma tachyzoites
(occurring when the parasite rights itself vertically, remaining attached to the substrate
by its posterior end and spinning clockwise) [60], the individuals of S. nematoides continued
to bend/twist also after detachment from the host tissue. In this view, the movements of S.
nematoides are most comparable to the attached waving of Plasmodium sporozoites that attach
at their apical ends and then exhibit waving or flexing, or may swivel or rotate [61].
Actin filaments
The apparent lack of visible, stable filaments does not fit for all apicomplexans, as in S. nematoides,
some eugregarines and protococcidian Eleutheroschizon duboscqi, the phalloidin labelling
revealed the presence of F-actin, even without the application of filament-stabilising
probes [3, 46, 47, 62]. The research on involvement of actin- and myosin-like proteins in gregarine
cell motility has been restricted to representatives of the genus Gregarina [3, 4, 12, 15, 16,
21]. The gregarine movement is often attributed to the F-actin cytoskeleton that is assumed to
exist in the form of a myocyte (= outer layer of longitudinal and inner layer of circular myonemes)
underlying the pellicle [12, 24, 25, 30, 32, 34] and to the ectoplasmic network [63], e.g.
the peristalsis is accompanied by the contraction of circular myonemes [24, 25].
In S. nematoides, the F-actin is distributed throughout the parasite cytoplasm and cell cortex.
Using a specific antibody recognising the Toxoplasma and Plasmodium actin, previous
studies successfully localised the actin within both the pellicle and cytoplasm of several gregarine
species [3, 46]. In S. nematoides, however, the actin immunolabelling was of patchy character
and restricted to the cytoplasm only. Under TEM, only short filamentous structures with
a maximal thickness of 9.7 nm were seen oriented perpendicularly between the plasma membrane
and IMC. These structures did not react to the JAS or cytochalasin D treatment indicating
that they are not of actin origin, and fluorescently-localised F-actin might be located
deeper, i.e. beneath the IMC. Irregular distribution of microtubules observed in JAS-treated
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parasites (reduced spacing of individual microtubule layers occurred also in cytochalasin Dtreated
individuals) along with a shortened distance between the internal cytomembrane and
the outer continuous microtubule layer, was probably caused by the condensation and contraction
of the cross-linking protein complexes that anchor the microtubules to the internal
cytomembrane. Localisation of these complexes corresponded to the filamentous structures
associated with microtubules in longitudinal sections. To our best knowledge, there is no data
indicating that JAS and cytochalasin D influence the polymerisation of other proteins than
actin. Hence we conclude that protein complexes observed in our study could be of actin
nature. Although this observation requires further analysis, it could support the idea that the
actin filaments might be localised in this area. Recent studies documented the microtubuleassociated
F-actin in Plasmodium gametocytes [64]. As this actin cytoskeleton was found in
non-motile gametocytes [64], it is likely that F-actin plays a rather structural role, thereby providing
a template for microtubule positioning. In such cases, it must be stable rather than
dynamic. Similarly to our results in S. nematoides, the treatment with cytochalasin D did not
disassemble the microtubule-associated F-actin in Plasmodium gametocytes [64]. Another
study performed on Arabidopsis also suggested the possibility of crosstalk between the F-actin
and cortical microtubules, as JAS treatment affected the orientation and parallel ordering of
microtubules and stabilised actin filaments were found to align with and move along microtubules
[65]. Similarly, disruption of the actin filaments using cytochalasin B affected microtubule
organisation in developing Zinnia elegans [66]. Altogether, these apparently highlystabilised
filamentous structures in S. nematoides might represent novel F-actin cytoskeleton
supporting the layers of numerous subpellicular microtubules.
Subpellicular microtubules
The regularly arranged subpellicular microtubules in S. nematoides exhibit a characteristic longitudinal
organisation and are nucleated from the apical polar ring, a microtubule-organising
center (MTOC) unique to the Apicomplexa [2]. The helical arrangement of microtubules in
apicomplexan zoites [2] and S. nematoides follow their serpentine body shape. The arrangement
of subpellicular microtubules varies among Apicomplexa, but their number, length, and
organisation are particular for the developmental stage of a species. The high number of S.
nematoides subpellicular microtubules concurs with a positive correlation between the species
dimensions and the number of subpellicular microtubules. In contrast to apicomplexan zoites
with subpellicular microtubules usually ending in the region below the nucleus (2/3 of the cell
length) [2], in S. nematoides, the microtubules extend along the entire length of the cell. Higher
accumulation of α-tubulin in blastogregarine caudal region, accompanied by the lack of an
increased number of microtubules in this area, is indicative of unpolymerised form of accumulated
tubulin.
Drug-treated apicomplexans lacking subpellicular microtubules are generally non-motile
and nonpolar [2]. Probes disrupting microtubules are usually effective only in certain taxonomic
groups. Oryzalin is known to inhibit growth of protists but not to disrupt the vertebrate
microtubules. For example, in T. gondii it binds to α-tubulin and prevents the formation of
new microtubules in daughter cells, while the effect on existing stable microtubules in the
mother cell is rather moderate [67, 68]; after prolonged treatment (40 h) in 2.5 μM oryzalin, all
tubulin is unpolymerised and dispersed [69]. In contrast, colchicine binds to β-tubulin and
effectively blocks microtubule assembly in animal cells [70]. For Chromista, corresponding to
S. nematoides, the minimal effective concentration of colchicine is 10 mM [70]. Drugs disrupting
dynamic microtubules are expected to be completely ineffective against the subpellicular
microtubules of extracellular apicomplexans, thus indicating that these microtubules are not
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dynamic [68]. In S. nematoides, however, prolonged incubation in high doses of oryzalin and
colchicine led to a gradual vanishing of subpellicular microtubules. The motility of drugtreated
blastogregarines attached to the host tissue was often limited to their apical region,
equipped with numerous layers of subpellicular microtubules, and persisted for the longest
incubation period. The effect of colchicine in extremely high doses (100 mM) applied for a
shorter time was considerably more effective than oryzalin. While in colchicine-treated parasites
the microtubules completely disappeared in some regions, in those treated with oryzalin,
they rather gradually faded away. Experiments performed on archigregarine S. fallax, using
0.1–2% colchicine diluted in seawater, showed similar results; i.e. the movement ceased in 19–
360 min depending on drug concentration, while the motility of the cilia of the host’s epithelium
was not affected [41]. Interestingly, the pellicular folding of treated archigregarines was
considerably less pronounced or non-existent, suggesting that subpellicular microtubules are
important in the formation and the maintenance of the longitudinal bulges. On the other
hand, the presence of the second and third row of more precisely arranged microtubules on
the inner curvature of bent parasites indicates the role of microtubules in archigregarine motility
[41]. Accordingly, archigregarines lacking subpellicular microtubules were shown to be
non-motile, although their longitudinal bulges were supported by arrays of fibrils reminiscent
of circular myonemes [39]. Our ultrastructural data on drug-treated individuals of S. nematoides
suggest that continuity of the outermost layer of subpellicular microtubules is essential
for normal movement. According to the doses, the motility of drug-treated individuals
stopped at a specific time on a regular basis, despite obvious differences in the number of preserved
microtubules in ultrathin sections. This indicates that the changes must first occur
along the length of microtubules; in longitudinal sections they appeared wavy (data not
shown) and, despite their presence, non-functional. Further support can be found in the observations
on a differential susceptibility of T. gondii subpellicular and spindle microtubules to
drugs, which could be either influenced by associated proteins specifically interacting with
some of the microtubule population, or, more likely, by the length that is required to become
functional [68].
The cross-linking protein complexes in S. nematoides that likely anchor the subpellicular
microtubules to the cytoplasmic face of IMC, might correspond to proteins, such as microtubule-associated
proteins (MAPs), which are important in controlling the local interactions of
microtubules with other structures. In general, MAPs are thought to control the spacing of
microtubules within the cell via microtubules interconnecting with other parts of the cytoskeleton
or the plasma membrane. It is likely that heavy decoration of subpellicular microtubules
in Apicomplexa may account for their unusual stability. Observations, when tubulin-specific
antibodies do not label the full length of microtubules, suggest that tubulin epitopes were
occluded by MAPs [71]. The MAPs, such as dyneins or kinesins, are known to be responsible
for sliding between adjacent microtubules [72]. A mechanism similar to that shown in ciliary
axoneme, with the microtubules sliding against one another, could account for the undulating
motility of archigregarines [42, 72] and blastogregarines. MAP-based mechanism has been
already proposed to explain the undulating and bending movements in Selenidium representatives
[40–42, 72]. The sliding might be localised during the bending movements and occurred
uniformly during overall contractions of a trophozoite [41]. Numerous peripheral mitochondria
detected in Selenidium archigregarines and S. nematoides seem to play an important role
in the rapid and continuous generation of ATP needed to support the highly dynamic cell plasticity
([40, 58, 72], this study), and might provide the chemical energy necessary for MAP
activity [72]. Accordingly, actively moving Selenidium species possess more subpellicular
microtubules and ectoplasmic mitochondria than less active ones. In Selenidium, the microtubules
in deeper layers appear to be orientated obliquely to the longitudinal cell axis [58].
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Although numerous ultrathin sections of S. nematoides did not allow us to unequivocally assess
the orientation of deeper microtubules, in longitudinal sections they appeared to be more
undulated and arranged loosely when compared to those organised in the outermost layer. In
addition, single microtubules or their small clusters oriented obliquely or perpendicularly to
the longitudinal cell axis were detected in few sections.
Finally, the organisation of subpellicular microtubules along with their placement within
the electron-lucent hexagonal chambers (= sheaths) in S. nematoides corresponds to the cortex
organisation in archigregarines of the family Selenidiidae [35, 37, 39–41, 58, 72]. The function
of these chambers remains to be elucidated, but they could play an important role in microtubule
sliding.
Pellicle
In Selenidium trophozoites, the multi-layered pellicle, usually with broad longitudinal bulges separated
by grooves [37, 58], might act as a stiff skeletal component, while the subpellicular microtubules
function in cell motility [73]. It has been proposed that together they represent a
unicellular analogue to the musculocuticular system of nematodes, in which longitudinal muscles
function antagonistically against an elastic cuticle [41, 58]. Despite the striking similarity in
motility mode and mechanism with Selenidium, however, the surface of S. nematoides is smooth.
The Kp of the plasma membrane in S. nematoides is significantly higher than in other apicomplexans
analysed to date (Table 2), indicating that more proteins must be anchored to its
protoplasmic face (the cortical supra-alveolar space). It is necessary to highlight that the widely
varied sizes of IMPs in S. nematoides pellicle membranes (Table 1) had a significant impact on
statistics, compared to studies, where only particles in the range from 6 to 14 nm were included
in the statistical calculations ([17, 49, 50], Table 2 in this study). The IMC refers to singlemembrane
flattened, cortical alveoli, which underlie the plasma membrane and are coupled to
a supporting cytoskeletal network of intermediate filaments [54]. Suture lines can be observed
at the edges where the alveoli contact each other [74]. In the IMC of S. nematoides, no sutures
were observed, so we assume that the IMC is formed from a single fused alveolus as also
described in Plasmodium life stages (except for gametocytes) [75]. The values of Kp in cortical
membranes of S. nematoides are higher than in eugregarines, but lower than in Eimeria or
Plasmodium (Table 2). In apicomplexans with gliding motility, the IMC outer leaflet anchors
the actomyosin motor; whereas the cytoplasmic face is intimately associated with the subpellicular
microtubules and alveolins generating cell rigidity [54]. The PF of the internal cytomembrane
in highly motile apicomplexan zoites shows longitudinal rows of 9 nm IMPs that likely
anchor the cytoskeleton to the IMC, while the non-motile stages apparently lack them [74].
Similar subpellicular network intimately associated with the pellicle cytoplasmic face extends
along the cell in T. gondii [76] and forms a resilient membrane skeleton that stabilises the alveoli
[75]. Furthermore, a double linear array of IMPs might overlay the microtubules. As the
helical path of some zoites during gliding corresponds to their helically coiled microtubules
and linear IMPs arrays, these IMPs may function as anchorage points for an axial motor system
[77]. In contrast, no regularly organised, linear IMP arrays were detected in the fractured
planes of S. nematoides cortical cytomembranes. Interestingly, despite a lack of longitudinal
IMP arrays in the IMC, the cytoplasmic face of the IMC exhibited imprints rather than IMP
alignments matching the localisation of subpellicular microtubules (Fig 4D).
Micropores and pores
Apicomplexan micropores are defined as organelles formed by the pellicle and composed of
two concentric rings (in transverse section), the inner of which corresponds to invagination of
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the plasma membrane. They are generally assumed to possess a feeding function, e.g. endocytosis
of the host cell cytoplasm during the vegetative phase of parasite development. In contrast
to other pores, micropores are usually less numerous and widely scattered [74]. Although the
presence of typical micropores (= cytostomes) has been confirmed in various apicomplexans,
including extracellular gregarines [3, 18, 49, 74], no identical structures were seen in S. nematoides.
Nevertheless, the pellicle of S. nematoides bears numerous pores of three different sizes;
the majority of them are organised in four laterally located, longitudinal rows. Although pores
were easily detected in cortical membranes by freeze-etching method, they do not seem to
interrupt the plasma membrane. In S. nematoides ultrathin sections, the plasma membrane in
the region of pores appears straight and not invaginated. Thus, these pores, observed only in
some SEM preparations, were most likely visualised due to a specific fixative osmolarity. In
eugregarines, the micropores are often located at the base of the grooves between the epicytic
folds, while the smaller pores are randomly distributed on the base or on the lateral side of
the folds [3, 17, 18, 78]. The diameter sizes of pores in S. nematoides correspond to those in
Gregarina spp. [3]. In S. nematoides, the function of pores arranged in lateral rows remains to
be elucidated. It is possible that they represent an alternate route to transport motor proteins
between the parasite cytoplasm and the cortical supra-alveolar space, independent of the route
through the apical complex, as it was described in Plasmodium ookinetes [74]. The largest
pores in S. nematoides, connected to a vesicle containing lamellar structure, resemble the ectoplasmic
structures observed in the bottom of the grooves in Selenidium representatives [37, 40,
58], which are assumed to serve for pinocytosis [58]. Based on comparable sizes and the putative
absence of typical micropores in S. nematoides, the largest pores detected in this blastogregarine
could possess similar function to the apicomplexan micropores.
Glycocalyx
A thick cell coat, the glycocalyx, covers the cell surface of S. nematoides. The thickness of the
glycocalyx layer significantly increased towards parasites’ apical end, hereby, suggesting that it
is produced by some of the apical organelles; e.g. micronemes. Similar reinforcing of glycocalyx
in the parasite’s apical region was documented in Selenidium archigregarines ([79], personal
unpublished data). The often highly decorated glycocalyx of unicellular parasites allows
them to interact with and respond to their environment, and is often essential for their virulence
[80].
Conclusions
Similar to archigregarines of the genus Selenidium, investigated blastogregarine S. nematoides
infects the intestines of marine invertebrates and exhibits ecological, morphological, and
motility traits inferred to reflect the early evolutionary history of apicomplexans. Despite the
presence of key glideosome components such as three-layered apicomplexan pellicle, actin
(including its filamentous form), myosin restricted to the cell cortex, subpellicular microtubules,
numerous micronemes and prominent glycocalyx layer (where adhesins might be
located), the motility mechanism of S. nematoides most likely differs from the glideosome
machinery. Parasites move independently on a solid substrate and show no signs of gliding
motility. We pointed to a possible role of the polymerised form of actin and tubulin in S. nematoides
motility, which could be described as a combination of pendular, twisting, undulation,
and sometimes spasmodic movements. Similar movements were described in Selenidium
archigregarines. As already proposed for motility mechanism in Selenidium spp., our observations
suggest that the subpellicular microtubules organised in several layers are the real leading
motor structures. The majority of S. nematoides actin is stabilised in a polymerised form and
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appears to be located beneath the IMC. The filamentous structures (i.e. cross-linking protein
complexes) associated with subpellicular microtubules reacted to the JAS and cytochalasin D
treatment, resulting in changes in spacing of microtubules, hereby indicating that they could
be of actin origin. If the axoneme-like sliding mechanism of microtubules is applicable for S.
nematoides motility, it is possible that this putative actin cytoskeleton associates lengthwise
with subpellicular microtubules to position them within the cytoplasm just beneath the pellicle.
Otherwise, the actin filaments may force the synchronised bending of microtubules in
some cell regions and this way generate the typical undulating motility of S. nematoides.
Supporting information
S1 Video. The motility of Siedleckia nematoides individuals attached to the host intestine
during incubation in seawater.
(MP4)
S2 Video. The motility of detached Siedleckia nematoides individuals during incubation in
seawater.
(MP4)
S3 Video. The modified motility of detached Siedleckia nematoides individuals incubated
with 30 μM oryzalin for 20 minutes. Note: one cell demonstrates oscillating movement, while
another cell–decreased bending motility.
(MP4)
S4 Video. The slightly limited motility of attached and detached Siedleckia nematoides
individuals incubated with 10 μM oryzalin for 30 minutes.
(MP4)
S5 Video. The limited motility of attached Siedleckia nematoides individuals incubated
with 100 mM colchicine for 15 minutes.
(MP4)
S6 Video. The increased motility of attached Siedleckia nematoides individuals incubated
with 30 μM jasplakinolide for 5 minutes.
(MP4)
S7 Video. The decreased motility of attached Siedleckia nematoides individuals incubated
with 10 μM jasplakinolide for 3 hours.
(MP4)
S8 Video. The slightly increased and modified motility of attached and detached Siedleckia
nematoides individuals incubated with 30 μM cytochalasin D for 25 minutes.
(MP4)
S9 Video. The decreased and limited motility of detached Siedleckia nematoides individuals
incubated with 30 μM cytochalasin D for 6 hours.
(MP4)
Acknowledgments
The authors would like to thank the staff of the White Sea Biological Station of Lomonosov
Moscow State University (WSBS MSU) and the Marine Biological Station of St. Petersburg
State University (MBS SPbSU) for providing facilities for field sampling and material processing,
as well as their kind and friendly approach. The authors are greatly indebted to the
Motility in blastogregarines (Apicomplexa)
PLOS ONE | https://doi.org/10.1371/journal.pone.0179709 June 22, 2017 25 / 29
Laboratory of Electron Microscopy, Biology Centre CAS, supported by the Czech-BioImaging
large RI project (LM2015062 funded by MEYS CR) for their support and technical assistance
with obtaining of EM data presented in this paper. The present study also utilised equipment
of core facility centres ‘Culturing of microorganisms’, ‘Observatory of Environmental Safety’
and ‘Development of molecular and cell technologies’ of St. Petersburg State University. Many
thanks to Prof. Dominique Soldati-Favre (University of Geneva) for kindly providing the
monoclonal anti-actin antibody.
Author Contributions
Conceptualization: AV.
Formal analysis: AV NV AD.
Investigation: AV MK NV AD GGP TGS.
Methodology: AV MK NV AD GGP TGS.
Resources: AV NV GGP AD MK TGS.
Validation: AV MK NV.
Writing – original draft: AV.
Writing – review & editing: AV NV GGP MK AD TGS.
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7.12
Melicherová J., Hofmannová L., Valigurová A.
2018
Response of cell lines to actual and simulated inoculation
with Cryptosporidium proliferans
European Journal of Protistology 62, 101-121
Available online at www.sciencedirect.com
ScienceDirect
European Journal of Protistology 62 (2018) 101–121
Response of cell lines to actual and simulated inoculation with
Cryptosporidium proliferans
Janka Melicherováa, Lada Hofmannováb, Andrea Valigurováa,∗
a
Department of Botany and Zoology, Faculty of Science, Masaryk University, Kotláˇrská 2, 611 37 Brno, Czech Republic
b
Department of Pathological Morphology and Parasitology, University of Veterinary and Pharmaceutical Sciences, Palackého tˇr. 1946/1, 612
42 Brno, Czech Republic
Received 16 May 2017; received in revised form 8 December 2017; accepted 13 December 2017
Available online 15 December 2017
Abstract
The need for an effective treatment against cryptosporidiosis has triggered studies in the search for a working in vitro model.
The peculiar niche of cryptosporidia at the brush border of host epithelial cells has been the subject of extensive debates. Despite
extensive research on the invasion process, it remains enigmatic whether cryptosporidian host-parasite interactions result from
an active invasion process or through encapsulation. We used HCT-8 and HT-29 cell lines for in vitro cultivation of the gastric
parasite Cryptosporidium proliferans strain TS03. Using electron and confocal laser scanning microscopy, observations were
carried out 24, 48 and 72 h after inoculation with a mixture of C. proliferans oocysts and sporozoites. Free sporozoites and
putative merozoites were observed apparently searching for an appropriate infection site. Advanced stages, corresponding to
trophozoites and meronts/gamonts enveloped by parasitophorous sac, and emptied sacs were detected. As our observations
showed that even unexcysted oocysts became enveloped by cultured cell projections, using polystyrene microspheres, we
evaluated the response of cell lines to simulated inoculation with cryptosporidian oocysts to verify innate and parasite-induced
behaviour. We found that cultured cell encapsulation of oocysts is induced by parasite antigens, independent of any active
invasion/motility.
© 2017 Published by Elsevier GmbH.
Keywords: Antigen; Apicomplexa; Cryptosporidium muris; Cryptosporidium proliferans; Encapsulation; Invasion
Introduction
The genus Cryptosporidium (Tyzzer, 1907) is an economically
significant and zoonotic pathogen of the gastrointestinal
Abbreviations: BSA, bovine serum albumin; CLSM, confocal laser
scanning microscopy; F-actin, filamentous actin; FBS, foetal bovine
serum; FITC, fluorescein isothiocyanate; HPI, hours post inoculation; PBS,
phosphate buffered saline; SEM, scanning electron microscopy; TRITC,
tetramethylrhodamine isothiocyanate.
∗Corresponding author.
E-mail address: andreav@sci.muni.cz (A. Valigurová).
tract, mainly threatening cattle/livestock and humans. In
healthy hosts, cryptosporidiosis is usually a self-limiting
disease; however, it produces a chronic and debilitating
condition in immunocompromised hosts, with no effective
treatment (Evering and Weiss 2006). One factor limiting the
development of a functioning anti-cryptosporidial drug is the
lack of any long-term in vitro cultivation system, although
significant progress in long-term cultivation of C. parvum
was recently reported (Morada et al. 2016). Furthermore,
gaps remain in our understanding of cryptosporidian invasion
mechanisms and formations of host-parasite interactions.
https://doi.org/10.1016/j.ejop.2017.12.003
0932-4739/© 2017 Published by Elsevier GmbH.
102 J. Melicherová et al. / European Journal of Protistology 62 (2018) 101–121
Using molecular markers (SSU rDNA, actin, HSP70 and
COWP1 genes), two sister/related cryptosporidian groups
have been distinguished (Xiao et al. 1999a,b) that differ in
their localisation within the host’s body, i.e. a gastric group,
of which the most studied species are C. andersoni, C. muris,
C. galli, C. serpentis, C. proliferans, and C. fragile, and an
intestinal group, of which the best known species are C.
parvum, C. baileyi, C. pestis, C. hominis, C. canis and C. felis.
The cryptosporidian life cycle, both within the host organism
and in vitro culture systems, shows distinct variation between
these two groups as well as species. While the life cycle of the
intestinal species C. parvum has been investigated in numerous
in vivo and in vitro studies (Arrowood 2002; Fayer and
Ungar 1986; Hijjawi et al. 2001; King et al. 2011; Upton et al.
1994, 1995), the life cycle of gastric species has only been
studied occasionally (Aydin and Ozkul 1996; Melicherová
et al. 2014; Taylor et al. 1999; Uni et al. 1987; Valigurová
et al. 2007, 2008). Cryptosporidian species infecting intestinal
sites are distinguished considerably by their response to
the host environment or other biological factors when compared
to gastric species. For example, C. parvum, an intestinal
species, exhibits a shorter prepatent period and a more rapid
occurrence of mature oocysts in host faeces than C. proliferans
(Melicherová et al. 2014; Tzipori 1983). Many studies
havedescribedthesuccessfulinvitrocultivationofC.parvum
and complete development of its asexual and sexual stages
for a range of cell lines (Kato et al. 2001; King et al. 2011;
Rosales et al. 1993; Yu et al. 2000). Gastric species, however,
which develop in the more inhospitable environment of the
host’s stomach, appear to depend on more specific factors
influencing parasite survival (Choi et al. 2004). Hence, longterm
maintenance of gastric species has yet to be achieved in
cell culture and the yield of newly formed oocysts still tends
to be less than the initial inoculum (Hijjawi et al. 2002).
While some refer to cryptosporidia as intracellular
though extracytoplasmic parasites, the others prefer the
term “epicellular” to describe the peculiar localisation of
cryptosporidian developmental stages (Barta and Thompson
2006; Bartoˇsová-Sojková et al. 2015; Borowski et al. 2010;
Cavalier-Smith 2014; Clode et al. 2015; Dumenil 2011; Ryan
et al. 2016; Valigurová et al. 2008). Following attachment of
the sporozoites to the surface of the host cell, the parasite apical
plasma membrane interacts with the host’s microvilli and
the parasite gradually gets enveloped by a host-derived parasitophorous
sac. The sac serves as a protective coat against
the hostile conditions found in the host’s gastrointestinal tract
and reinforces attachment of the parasite to the host’s epithelium
(Valigurová et al. 2008). Interestingly, cryptosporidian
extracellular stages have been reported in cell-free cultures
(Aldeyarbi and Karanis 2015; Boxell et al. 2008; Hijjawi et al.
2004, 2010; Koh et al. 2013, 2014; Rosales et al. 2005; Ryan
et al. 2016), though other studies have failed to repeat these
results (Girouard et al. 2006).
This study focuses on the formation of host-parasite
interactions in gastric pathogen Cryptosporidium proliferans
cultivated in vitro. Our original intent was to establish a working
in vitro system for gastric cryptosporidia using protocols
from previously published studies (Upton et al. 1994, 1995;
Woods et al. 1995); however, repeated observations of an
unusual cell line reaction to the parasite oocysts prompted
us to design further experiments to investigate the reaction
of cell lines to presence of foreign objects. Cell lines were
inoculated with either cryptosporidia or polystyrene microspheres
in order to compare their innate and parasite-induced
behaviour.Acombinedmicroscopicapproachusingscanning
electron and confocal laser scanning microscopy was used to
evaluate modifications to the inoculated cultures.
Material and Methods
Specification of cell lines and cultivation
conditions
Human ileocecal adenocarcinoma cell lines (HCT-8;
ATCC
®
CCL244) and human colorectal carcinoma cell lines
(HT-29; ATCC
®
HTB-38 TM) were cultured in RPMI 1640
medium with l-glutamine (Sigma–Aldrich, Prague, Czech
Republic), supplemented with 10% foetal bovine serum
(FBS), sodium bicarbonate (2 g/l), penicillin G (100 U/ml),
streptomycin (100 ␮g/ml) and amphotericin B (250 ␮g/ml)
(BioTech, Prague, Czech Republic). Our choice of HCT-8
and HT-29 cell lines was based on published data reporting
high achievement in cultivating the intestinal species C.
parvumandC.hominis(Arrowood2002;Flaniganetal.1991;
Hashim et al. 2006) and the complete development of gastric
C. andersoni in the HCT-8 cell line (Hijjawi et al. 2002).
The cells lines were maintained in 25-cm2 tissue culture
flasks in a 5% CO2 atmosphere at 37 ◦C and 100% humidity
and passaged every 2–3 days. A confluent monolayer of
cells was lifted from the flask’s surface by using a solution
consisting of 0.25% (wt/vol) trypsin and 0.53 mM ethylenediaminetetraacetic
acid (EDTA) in phosphate buffered saline
(PBS). The cell suspension was centrifuged for 3 min at
216.3g in 15 ml falcon tubes. The resultant pellet was then
re-suspended in maintenance medium and split between new
flasks or well plates.
Preparation of cell culture for inoculation
For the experimental inoculations, confluent monolayers
were washed with PBS, trypsinised and then centrifuged.
The resultant pellets from two flasks (2 × 25 cm2) were resuspended
in fresh medium and split between 12-well test
plates (3.2 cm2 per well), each containing a 15 mm diameter
cover glass on the bottom. The concentrated cell suspension
was applied to each well and topped-up with RMPI 1640
(100 ml) with l-glutamine enriched with sodium bicarbonate
(0.3 g/ml), bovine bile (0.02 g/ml), glucose (0.1 g/ml), folic
acid (25 mg/ml), 4-aminobenzoic acid (100 mg/ml), calcium
pantothenate (50 mg/ml), ascorbic acid (875 mg/ml), antibi-
J. Melicherová et al. / European Journal of Protistology 62 (2018) 101–121 103
otics (100 U/ml penicillin and 100 ␮g/ml streptomycin),
antimycotics (250 ␮g/ml amphotericin B) and 5% FBS
(BioTech, Prague, Czech Republic). This ‘supplemented
medium’ was used for all experimental inoculations in our
study as it produced better physiological conditions for keeping
the culture and appeared to support the parasite invasion
process. After two days, cells adhering to the cover glasses
had created a 70–80% confluent monolayer.
Characterisation of parasites
The gastric species used in this and some of our published
studies (Melicherová et al., 2014, 2016) was previously characterised
as Cryptosporidium muris strain TS03, originating
from an East African mole rat (Tachyoryctes splendens)
(Feng et al. 2011; Kváˇc et al. 2008). A recent study of Kváˇc
et al. (2016), however, concluded that this strain differs genetically
from C. muris and other known cryptosporidia, and
proposed a new species name for it, C. proliferans.
Microscopic identification of C. proliferans developmental
stages was based on their description and measurements
given in previous studies (Melicherová et al. 2014; Kváˇc et al.
2016).
Preparation of parasites
Oocysts of C. proliferans (strain TS03), originating from
the Institute of Parasitology Biology Centre at the Academy
of Sciences of the Czech Republic ( ˇCeské Budˇejovice), were
passaged in experimentally inoculated southern multimammate
mice (Mastomys coucha) kept at the Department of
Pathological Morphology and Parasitology at the University
of Veterinary and Pharmaceutical Sciences in Brno (UPVS)
in agreement with Czech legislation (Act No 246/1992 Coll.,
on the Protection of Animals Against Cruelty). All housing,
feeding and experimental procedures were conducted under
Protocol 31-2014 approved by the UPVS and the Central
Commission for Animal Welfare, Czech Republic.
Faeces were collected in morning each day and stored with
aqueous potassium dichromate (2.5% w/v, final concentration)
at 4 ◦C. Oocysts were purified using Sheather’s sugar
flotation method (Arrowood and Sterling 1987) and modified
caesium chloride gradient centrifugation (Kilani and Sekla
1987) then stored in PBS at 4 ◦C for a maximum of four
weeks. Viability of the C. proliferans oocysts was assessed
using the fluorescein diacetate/propidium iodide staining
protocol (Jones and Senft 1985). Based on the protocol standardized
in our previous study (Melicherová et al. 2016),
oocysts of C. proliferans were excysted in a 37 ◦C water bath
using RPMI 1640 medium with 5% bovine serum albumin
(BSA) for 30–60 min, until the percentage of oocyst excystation
reached 40–50%. Then the suspension was centrifuged
for 3 min at 100g and resultant pellet was re-suspended in
supplemented medium. Prior to inoculation of cell cultures,
activity/motilityofexcystedsporozoiteswasmonitoredusing
the Olympus IX70 research inverted tissue culture micro-
scope.
Inoculation of cell cultures with cryptosporidian
oocysts or with foreign non-living objects
The 70–80% confluent cell line monolayer was washed
with 0.1 M PBS and a mixture of 1 × 104 unexcysted/excysted
oocysts and sporozoites of C. proliferans
was placed into each of 12-well plates topped-up with 1 ml
of fresh supplemented medium. Using the light microscope
Olympus IX70, behaviour of parasites (such as estimation
of the ratio between the excysted and intact oocysts, monitoring
of the sporozoites’ viability) inoculated into the cell
culture was continuously controlled during the entire experiment
and prior to each fixation for subsequent procedures.
Cover glasses with inoculated culture were analysed 24-, 48and
72-h post inoculation (HPI) using confocal laser scanning
microscopy (CLSM) and scanning electron microscopy
(SEM).
Simulated inoculation of HCT-8 and HT-29 cell lines with
non-living foreign objects (polystyrene microspheres; specified
more precisely in following sections) was undertaken
in order to verify whether the embracing of C. proliferans
oocysts by cell lines was provoked by the parasite itself or
whether it represents an innate reaction of the culture to foreign
objects in general. The same supplemented medium was
used for cultures inoculated with either the parasite or the
foreign objects.
Preparation of parasite antigen-coated polybead
microspheres
Suspensions of C. proliferans oocysts were partially
excysted (30 min) in sterile PBS, sonicated by placing the
material on ice, centrifuged at 10,314g to remove undisrupted
remnants and resulting supernatant was used as C. proliferans
antigen. A model 150 V/T ultrasonic homogeniser
(230 V/50 Hz, power 0–150 W, timer 0–15 min; Biologics
Inc., Manassas, Virginia, USA) fitted with a 5/32 diameter
stepped titanium micro tip (300 ␮l–15 ml processing
volume, very high intensity) was used to obtain C. proliferans
protein homogenate. The C. proliferans antigen-coated
polystyrene microspheres were prepared according to protocols
of Valigurová et al. (2014). The coated polystyrene
microspheres were re-suspended in RPMI 1640 medium supplemented
with vitamins and glucose, and split between
each well with a total volume of approximately 1000 microspheres/200
␮l.
Scanning electron microscopy
HCT-8 and HT-29 cell lines were inoculated with either
excysted oocysts of C. proliferans or polystyrene microspheres,
i.e. 6 ␮m Polybead
®
Polystyrene Red Dyed Micro-
104 J. Melicherová et al. / European Journal of Protistology 62 (2018) 101–121
Fig. 1. Scanning electron micrographs of HCT-8 cell lines inoculated with Cryptosporidium proliferans at 24 HPI. (A) Excysting C. proliferans
oocyst releasing a sporozoite (black arrowhead); suture (black arrow). (B) Free sporozoite with typically prolonged apical end (black
arrowhead). (C) A general view of the HCT-8 monolayer inoculated with C. proliferans; newly formed round cultured cells (c), oocysts (white
arrowheads) and invading sporozoites (black arrowheads).
spheres (CAT#15714) and 10 ␮m Polybead
®
Microspheres
(CAT#17136; Polysciences Europe GmbH, Hirschberg an
der Bergstrasse, Germany) coated with protein homogenate
obtained from C. proliferans oocysts. Samples on cover
glasses were fixed in 2.5% glutaraldehyde in pH 7.2 phosphate
buffer at 24-, 48- and 72-HPI. Negative controls were
evaluated in a similar manner. The specimens were then
examined under a JEOL JSM-6300 scanning electron microscope
(JEOL, Peabody, MA, USA).
Confocal laser scanning microscopy
Cell lines were grown in RPMI 1640 supplemented with
vitamins and glucose on cover slides placed in 12-well plates
(15 mm). Following inoculation with either C. proliferans, or
pure/C. proliferans antigen-coated polystyrene microbeads
(10 ␮m Fluoresbrite
®
YG Microspheres, CAT#18140 or
10 ␮m Polybead
®
Microspheres, CAT#17136; Polysciences
Europe GmbH, Hirschberg an der Bergstrasse, Germany),
samples were fixed in freshly prepared 4% paraformaldehyde
(v/v) in 0.2 M PBS, washed in 0.1 M PBS and permeabilised
in0.3%TritonX-100.Thereafter,thespecimenswerewashed
with antibody diluent (containing 0.1 M PBS, 0.1% Triton X-
100, 0.1% bovine serum albumin and 0.1% sodium azide at
pH 7.4), incubated with TRITC-phalloidin (Sigma–Aldrich,
Prague, Czech Republic), washed with antibody diluent
and counterstained with Hoechst 33342 (Life Technologies,
Prague, Czech Republic). Preparations were mounted in
anti-fade based on 2.5% DABCO (Sigma–Aldrich, Prague,
Czech Republic) mixed with glycerol and 0.1 M PBS or
VECTASHIELD
®
(Vector Laboratories, Burlingame, CA,
USA) and viewed under an Olympus IX80 microscope
equipped with a laser scanning FluoView 500 confocal unit
(Olympus FluoView 4.3 software), using the tetramethylrhodamine
isothiocyanate (TRITC-phalloidin), UV (Hoechst)
and fluorescein isothiocyanate (Fluoresbrite
®
YG Microspheres
with excitation and emission spectra similar to FITC)
filter sets (Olympus Czech Group, Prague, Czech Republic).
Results
Experimental inoculation of HCT-8 and HT-29
cell lines with Cryptosporidium proliferans strain
TS03
In both cell lines (HCT-8 and HT-29), a 70–80% confluence
provided an appropriate environment for culture
inoculation and parasite development. Furthermore, the conditions
used (see Section ‘Material and methods’) clearly
inhibited the formation of cell multilayers, thus preventing
unwanted overgrowth of parasites with cultured cells.
Oocyst excysted continuously within the well plates with supplemented
medium, while free motile sporozoites occurred
repeatedly throughout the experiment.
The first interaction between the parasite developmental
stages (unexcysted/excysted oocysts) and the cell line monolayer
was noted at 24 HPI. Of the two cell lines examined, the
reaction of the HCT-8 cell line to the parasite invasion process
was more intense, as shown by formation of clearly elongated
microvilli around the excysting oocysts and sporozoites. As
expected,excystationofunexcystedoocystscontinuedduring
cell line incubation (Fig. 1A). Released sporozoites probed
the cultured cell membrane with their prolonged apical end
and attempted to invade the cultured cells (Fig. 1B). Developmental
stages of C. proliferans were frequently found near
dividing or newly formed round cells, with oocysts even
observed stuck to the surface of the young cells (Fig. 1C).
Slightly elongated cultured cell microvilli were observed
surrounding some of the attached sporozoites. At 48 HPI,
numerous viable sporozoites were observed searching for
an appropriate invasion/attachment site within the HCT-8
cell culture (Fig. 2A–F). Invading sporozoites were partially
embedded in cultured cells with obviously altered microvilli
(Fig.2A–D).Remarkably,somesporozoiteshadalreadybeen
embraced by the cultured cell membrane (Fig. 2E, F). Early
developmental stages corresponding with their size and shape
to trophozoites (Fig. 2G–I), as well as more advanced stages
resembling meronts or gamonts (Fig. 2J), were found. The
J. Melicherová et al. / European Journal of Protistology 62 (2018) 101–121 105
Fig. 2. Scanning electron micrographs of HCT-8 cell lines inoculated with Cryptosporidium proliferans at 48 HPI. (A, B) Invading sporozoite
(black arrowhead); elongated microvilli of cultured cell (white arrow). (C) Excysting oocyst with opened suture (black arrow) releasing a
sporozoite (black arrowhead). Note that the sporozoite immediately invades the cultured cell. (D) Oocyst with suture (black arrow) embedded
into a newly formed, round cell and an invading sporozoite (black arrowhead). (E) Sporozoite (black arrowhead) partially covered by membrane
fold (white arrow) formed by the cultured cell. (F) Free sporozoites (black arrowheads), one of which is already covered by a cultured cell
membrane fold (white arrow). (G) Oocyst with a free sporozoite (black arrowhead) and an early trophozoite (asterisk). (H) Detail of the
early trophozoite (asterisk) shown in G. (I) A more mature trophozoite stage of C. proliferans. (J) A newly-formed round cell with attached
meront-/gamont-like stage within the parasitophorous sac. Note the elongated microvilli and membrane fold fragments (white arrows) of
cultured cell. (K) Oocyst (white arrowhead) partially embraced by plasma membrane fold of a single cultured cell (white arrows). (L) Oocyst
(white arrowhead) enveloped by cultured cell projection and a single free sporozoite (black arrowhead).
microscopic identification was based on description and measurements
of particular developmental stages reported in our
previous study (Melicherová et al. 2014). Though the stages
enveloped by parasitophorous sacs were hardly distinguishable
from each other under SEM, there were some differences
that allowed us to distinguish the trophozoites from more
106 J. Melicherová et al. / European Journal of Protistology 62 (2018) 101–121
Fig. 3. Scanning electron micrographs of HCT-8 cell lines inoculated with Cryptosporidium proliferans at 72 HPI. (A) Free sporozoites (black
arrowheads) attracted by the newly formed round cells (c). (B) Two sporozoites (black arrowhead) apparently checking the target site on the
cultured cell’s surface. (C) Three free sporozoites (black arrowhead), one of which is already covered by the cultured plasma membrane fold
(white arrow). (D) Newly formed cultured cell (c) invaded by a sporozoite (black arrowhead) and one free sporozoite (black arrowhead on
the left). (E) Detail of the invading sporozoite (black arrowhead) shown in D, partially embraced by a projection of the cultured cell (white
arrow). (F) Putative merozoite partially covered by the fold of the cultured cell’s plasma membrane (white arrow). (G) Sporozoite torn from
the monolayer. Note the apical end with remnants of the cultured cell plasma membrane (white arrow). (H) Detached zoite transforming into a
trophozoite stage, completely enveloped by a parasitophorous sac (white arrow). (I) More advanced developmental stage (meront or gamont)
located within the parasitophorous sac and surrounded by elongated microvilli.
mature stages; i.e. trophozoites were more elongated in shape
and significantly smaller in size. Nevertheless, it was almost
impossible to distinguish the meronts stages from gamonts
or zygotes. Excysting (Fig. 2C, D, G) or unexcysted C. proliferans
oocysts were still observable on the surface of the
cell lines, often completely or partially enveloped by cultured
cell projections/plasma membrane fold (Fig. 2K, L).
At 72 HPI, despite the increasingly obvious depleting effect
of parasites, 40–50% of the previous cell culture monolayer
was still preserved and new young round cells were observed
(Fig. 3A, D). By 72 HPI, the quantity of sporozoites trying to
invade culture cells had increased (Fig. 3A–E), with invading
sporozoites inducing a cell culture response resulting in the
gradual formation of a parasitophorous sac from the cultured
cell’s plasma membrane (Fig. 3C–E). There was evidence
that some sporozoites had been torn from the monolayer, with
the remains of the invaded cell at their apical end (Fig. 3G).
Zoites, with their size and shape corresponding to merozoites,
lying on the surface of cell culture establishing another
layer, were partially covered by cultured cell membrane folds
(Fig. 3F), while others were completely enveloped by parasitophorous
sacs (Fig. 3H). Few stages, completely enveloped
by parasitophorous sac and corresponding to meronts or
gamonts were detected (Fig. 3I).
HT-29 cell lines, incubated under the same conditions
as HCT-8 and inoculated with an equal number of
J. Melicherová et al. / European Journal of Protistology 62 (2018) 101–121 107
Fig. 4. Scanning electron micrographs of HT-29 cell lines inoculated with Cryptosporidium proliferans at 24 HPI. (A) Unexcysted oocyst
with closed suture (black arrow) at the cell culture surface. Note the elongated microvilli (white arrow) adhering to the oocyst’s surface. (B,
C) Oocyst with closed suture (black arrow) embedded to the cultured cell, partially (C) or almost completely (B) encapsulated by its plasma
membrane (white arrow). (D) Excysted oocyst with opened suture (black arrow) lying on the monolayer surface. The response of the cultured
cell is demonstrated by the elongated microvilli (white arrow). (E) Excysting oocyst releasing three sporozoites (black arrowheads); suture
(black arrow). (F) Two sporozoites (black arrowheads) apparently checking the cultured cell’s surface. (G) Free sporozoite (black arrowhead)
partially covered by the cultured cell’s membrane fold (white arrow). (H) Zoite completely overlapped by a cultured cell (white arrow).
(I) Modified area (a) following previous interaction with parasite, characterised by the remnants of raised and altered plasma membrane of
cultured cell (white arrow).
excysted/unexcysted oocysts, showed a different reaction to
inoculation with C. proliferans. In contrast to HCT-8, far
more unexcysted oocysts with obviously closed sutures were
observed on the HT-29 cell line surface at 24 HPI (Fig. 4A).
Both unexcysted and empty oocysts were partially or completely
encapsulated by the plasma membrane of cultured
cells (Fig. 4B, C). Empty oocysts with opened sutures were
observed in contact with elongated HT-29 cell microvilli
(Fig. 4D). Oocysts continued to excyst and viable sporozoites
were released (Fig. 4E, F). Free sporozoites were often
noticed partially covered by cultured cell folds (Fig. 4G).
Some zoites were completely overlapped by a new layer of
HT-29 cells (Fig. 4H). Previous sites of parasites’ interaction
with the cell monolayer surface were clearly observable
due to preservation of raised cultured cell plasma membrane
folds (Fig. 4I). At 48 HPI, free sporozoites continued to
invade cultured cells (Fig. 5A). Numerous empty or unexcysted
oocysts were observed on the HT-29 cell line surface,
with some being embraced by cultured cells (Fig. 5A–C).
Developmental stages, corresponding to meronts or gamonts
(Fig. 5D, E) and sporadic individuals with their shape resembling
oocysts (Fig. 5F), were detected developing within
parasitophorous sacs on the cell line surface. At 72 HPI,
a number of empty and unexcysted oocysts were observed
being encapsulated by the cultured cells to a greater or lesser
degree (Fig. 6A–C). The imprints from previous parasites’
occurrence and modified sites of cultured cells with raised
plasma membrane folds were often noticed (Fig. 6D–F).
No free sporozoites or merozoites were observed at 72
HPI.
108 J. Melicherová et al. / European Journal of Protistology 62 (2018) 101–121
Fig. 5. Scanning electron micrographs of HT-29 cell lines inoculated with Cryptosporidium proliferans at 48 HPI. (A) Excysted oocyst (white
arrowhead) and a sporozoite (black arrowhead) invading young HT-29 cell. (B) Two oocysts stuck into cultured cells, one of which is partially
covered by a cultured cell’s membrane fold (white arrow). (C) Three oocysts adhering to the surface of the cultured cells; biological debris
adhering to the oocyst surface (white arrow), suture (black arrow). (D, E) More advanced developmental stage (meront or gamont) within
parasitophorous sac; elongated microvilli (white arrow). (F) Mature developmental stage resembling oocyst within parasitophorous sac.
Fig. 6. Scanning electron micrographs of HT-29 cell lines inoculated with Cryptosporidium proliferans at 72 HPI. (A) Unexcysted oocyst
gradually being enveloped by the cultured cell’s plasma membrane (white arrow). (B) Detailed view of the plasma membrane fold (white
arrow) raising along the oocyst shown in A; suture (black arrow). (C) Empty oocyst immersed into and partially embraced by the cultured
cell (white arrow). (D) Circular imprint (white arrows) left after oocyst (white arrowhead) torn away from the cultured cell. (E) Remnants of
raised plasma membrane (white arrow) of cultured cell left after previous parasitisation by C. proliferans. (F) Deep imprint of the parasite
visible on the surface of the cultured cell (a).
J. Melicherová et al. / European Journal of Protistology 62 (2018) 101–121 109
Fig. 7. Fluorescence visualisation of F-actin in HCT-8 cell lines inoculated with Cryptosporidium proliferans at 24, 48 and 72 HPI. (A–D)
HCT-8 cells with C. proliferans oocysts at 24 HPI. One oocyst is partially immersed (white arrow), while the second is completely enveloped by
the cultured cell (white arrowhead). CLSM with transmitted light (A, C) and CLSM (B, D). C–D represent different optical planes confirming
that oocysts is completely embedded in the cultured cell. (E, F) Cultured cells at 48 HPI with one C. proliferans oocyst stuck to their surface
(white arrow) and one embedded (white arrowhead). CLSM with transmitted light (E) and CLSM (F). (G, H) Two oocysts embedded in
cultured cells (white arrowheads) at 72 HPI. CLSM with transmitted light (G) and CLSM (H). All micrographs are composite views created
by flattening a series of optical sections using the TRITC (F-actin stained with phalloidin) and UV (nuclei stained with Hoechst) filter sets.
The same experiments, run under identical conditions,
were also evaluated using phalloidin staining of filamentous
actin (F-actin) for confocal laser scanning microscopy
(CLSM). Different optical sections of the culture revealed
the actual localisation of oocysts as well as their interactions
with cultured cells. At 24 HPI, C. proliferans oocysts in both
HCT-8 and HT-29 cell lines were predominantly located in
gaps in the discontinuous monolayer and usually stuck to the
lateral side of the cultured cells (Figs. 7A–D and 8A, B). At
48 HPI, when the monolayer was more continuous, oocysts
occurred mainly on the free surface of cell culture, more or
less embedded in individual cells. While fluorescently visualised
cultured cell actin filaments started to envelope the
oocysts in HT-29, a similar reaction was not observed in HCT-
8 (Figs. 7E, F and 8C, D). At 72 HPI, oocysts were deeply
embedded within the monolayer, with increased accumulation
of F-actin surrounding the oocysts in the HT-29 cell lines
(Fig. 8E, F), but negligible reorganisation of F-actin in the
HCT-8 cell lines (Fig. 7G, H).
Experimental inoculation of HCT-8 and HT-29
cell lines with polystyrene microspheres
At 24 HPI, polystyrene microspheres were recorded lying
on the surface of both the HCT-8 and HT-29 cell lines, with
no significant response from individual cultured cells (Figs.
9A, B and 10A, B) and only biological debris adhering to the
microspheres (Fig. 9C). Nevertheless, tiny elongated cultured
cell microvilli were attached to the microspheres (Fig. 9D).
At 48 HPI, the microspheres appeared to be clustering in
gaps in the HCT-8 monolayer (Fig. 9E), while others were
adhered to the culture’s surface (Figs. 9F, G and 10C). At 72
HPI, similar results were observed, with no real interaction
between microspheres and cells recorded (Figs. 9H–J and
10D–F). It is necessary to mention that daily washing and
changing of the medium considerably reduced the total number
of polystyrene microspheres adhering to the cell culture
over time.
Fluorescence observations on HCT-8 and HT-29 cell lines
inoculated with polystyrene microspheres were performed
at the same time as those above. At 24 HPI, distribution of
actin filaments within the cultured cells of both cell lines indicated
no response to the polystyrene microspheres (Fig. 11A,
B) or only shallow imprints of the cultured cell cytoskeleton
(Fig. 12A–C). Evaluation of cultures at 48 HPI showed
similar results, i.e. cultured cell actin filaments had not
enveloped the microspheres (Fig. 12D–F); at most, microsphere
imprints were seen on cell surfaces (Fig. 11C, D).
Even at 72 HPI, no change in F-actin organisation was noticed
(Fig. 11E, F).
110 J. Melicherová et al. / European Journal of Protistology 62 (2018) 101–121
Fig. 8. Fluorescence visualisation of F-actin in HT-29 cell lines inoculated with Cryptosporidium proliferans at 24, 48 and 72 HPI. (A, B)
HT-29 cells with C. proliferans oocysts (white arrowheads) at 24 HPI. CLSM with transmitted light (A) and CLSM (B). (C, D) Deeply
immersed oocysts (white arrowheads), obviously enveloped by cultured cell F-actin at 48 HPI. CLSM with transmitted light (C) and CLSM
(D). (E, F) Increased accumulation of F-actin surrounding the C. proliferans oocysts (white arrowheads) at 72 HPI. CLSM with transmitted
light (E) and CLSM (F). All micrographs are composite views created by flattening a series of optical sections using the TRITC (F-actin
stained with phalloidin) and UV (nuclei stained with Hoechst) filter sets.
Simulated parasitisation of HCT-8 and HT-29
cell lines with C. proliferans antigen-coated
polystyrene microspheres
On the basis of data obtained during the previous two
experiments, further trials were designed in which HCT-8
and HT-29 cell lines were inoculated with polystyrene microspheres
coated with C. proliferans antigens (obtained from
oocyst/sporozoite homogenate).
At 24 HPI, HCT-8 cell lines displayed elongated filamentous
microvilli of cultured cells embracing the foreign
object (Fig. 13A–C). At 48 HPI, the cell lines were showing
an increased response to the antigen-coated microspheres
(Fig. 13D, E), with microspheres now covered with cultured
cell plasma membrane folds (Fig. 13F). At 72 HPI, we documented
the gradual formation of cultured cell ‘shelters’,
covering the microspheres in a manner similar to parasitophorous
sacs (Fig. 13G–I). Formation of these shelters
J. Melicherová et al. / European Journal of Protistology 62 (2018) 101–121 111
Fig. 9. Scanning electron micrographs of HCT-8 cell lines inoculated with polystyrene microspheres at 24, 48 and 72 HPI. (A) HCT-8
monolayer with polystyrene microspheres (white arrowheads) on its surface at 24 HPI. (B) Lateral view at 24 HPI of three polystyrene
microspheres laying on the surface of cultured cells that show no reaction to the foreign objects. (C) A polystyrene microsphere at 24 HPI
covered with biological debris (black arrow). (D) Polystyrene microspheres at 24 HPI with adhering microvilli of surrounding cells (white
arrow). (E) Polystyrene microspheres (white arrowhead) located mostly within monolayer gaps at 48 HPI. (F) Polystyrene microspheres
(white arrowheads) laying on the surface of the HCT-8 monolayer at 48 HPI. (G) A polystyrene microsphere with debris (black arrow) on its
surface at 48 HPI. (H) The surface of HCT-8 multilayer with a polystyrene microsphere (white arrowhead) at 72 HPI. (I, J) Detailed view of
the top (I) and lateral side (J) of the same polystyrene microsphere at 72 HPI. Cultured cells show no reaction to the foreign objects.
often included both mechanisms; i.e. encapsulation as well
as overgrowth by cultured cells.
The inoculation of HT-29 cell lines with C. proliferans
antigen-coated microspheres showed similar results. Compared
to HCT-8 cells, HT-29 cells typically had shorter
filamentous microvilli; hence, interactions between cultured
cells and the microspheres were more obvious. At 24 HPI,
the microspheres were apparently immersed in the cells and
accumulating into small groups (Fig. 14A–C). At 48 HPI, the
antigen-coated microspheres had been partially enveloped
by cultured cell plasma membranes (Fig. 14D–F), and had
been completely covered by the cell culture by 72 HPI
(Fig. 14G–I).
Simulated parasitisation with C. proliferans antigencoated
fluoresbrite polystyrene microspheres also showed
both cell lines reacting to the presence of foreign objects.
Distribution of F-actin in cell lines at 24 HPI indicated that
many microspheres were either deeply imprinted into individual
cultured cells or were already overgrown by a newly
formed layer of cells, and that microspheres had apparently
been surrounded by elongated cultured cell microvilli (Figs.
15A, B and 16A–C). Observations at 48 HPI revealed further
changes in the distribution of F-actin, with numerous holes
in the cell lines apparent. The microspheres had been partially
enveloped by cultured cell actin filaments (Fig. 16D–F),
indicating the raising of plasma membrane folds (Fig. 15C).
Prolonged incubation to 72 HPI resulted in encapsulation
of the microspheres by cultured cells (Figs. 15D, E and
16G–K). Similar to cultures inoculated with parasites, the
HT-29 showed more obvious changes in F-actin organisation
in cultures inoculated with parasite antigen-coated microspheres
when compared to the HCT-8 cell lines.
112 J. Melicherová et al. / European Journal of Protistology 62 (2018) 101–121
Fig. 10. Scanning electron micrographs of HT-29 cell lines inoculated with polystyrene microspheres at 24, 48 and 72 HPI. (A) HT-29
monolayer with polystyrene microspheres (white arrowheads) on its surface at 24 HPI. (B) Newly formed, round cells with polystyrene
microspheres on their surface at 24 HPI. (C) HCT-8 multilayer surface with a polystyrene microsphere at 48 HPI. (D) Polystyrene microspheres
covered with biological debris (black arrow) at 72 HPI. (E, F) Detailed view of the top (E) and lateral side (F) of the same polystyrene
microsphere at 72 HPI. Cultured cells show no reaction to the foreign objects.
Discussion
In vitro systems have the benefit of allowing investigations
into host-parasite interactions that cannot be observed within
the host’s own body, and thus the establishment of a successful
in vitro culture system of Cryptosporidium spp. has been a
challenge for many researchers. Since achievement of the first
in vitro cultivation of cryptosporidian asexual developmental
stages in 1983 (Woodmansee and Pohlenz 1983), only little
progress has been made. While a further study reported the
successful cultivation of human Cryptosporidium isolates,
including both asexual and sexual stages in several cell lines
(Current and Haynes 1984), anecdotal reports suggest that
attempts to replicate these results have been disappointing
(Arrowood 2002). Although later protocols appear suitable
for developing cryptosporidian stages, long-term maintenance
in culture systems has proven unsuccessful as the
infection tends to peak and decline at around 48–72 HPI
(Borowski et al. 2010; Gut et al. 1991; McDonald et al. 1990;
Rosales et al. 1993; Upton et al. 1994; Wu et al. 2009).
In this study, C. proliferans strain TS03 (previously C.
muris) was cultivated on the HCT-8 and HT-29 cell lines used
in previous studies reporting completion of Cryptosporidium
sp. life cycle in vitro (Alcantara Warren et al. 2008; Flanigan
et al. 1991; Hijjawi et al. 2001, 2002; Upton et al. 1995).
Based on suggestions in the studies mentioned above, the
RPMI 1640 cultivation medium used in our study was supplemented
with antibiotics/antimycotics, FBS, l-glutamine
and sodium bicarbonate and enriched with vitamins and glucose.
In contrast to protocols using 10% FBS (Choi et al.
2004; Flanigan et al. 1991; McDonald et al. 1990) or 1%
FBS (Borowski et al. 2010; Hijjawi et al. 2001, 2002), however,
the concentration of FBS in our study was set at 5%
based on observations on sporozoite motility using C. proliferans
strain TS03 (Melicherová et al. 2016). Although there
is some indication that components of FBS could inhibit parasite
growth (Woods and Upton 2007), another study has
shown that FBS can enhance the motility of sporozoites
(Upton et al. 1995), consistent with our study. In general,
appropriate concentrations of serum proteins (FBS, FCS, and
BSA) appear to have a positive influence on parasite motility
(Hijjawi et al. 2002; Melicherová et al. 2016; Upton et al.
1995). In addition, four vitamins (folic acid, 4-aminobenzoic
acid, calcium pantothenate and ascorbic acid) were added
to the medium as they have been shown to support interaction
between the parasite (C. parvum) and host cells (Hijjawi
et al. 2001; Upton et al. 1995). While investigating the optimal
cultivation conditions for C. muris in human stomach
adenocarcinoma cell lines, further study failed to notice any
positive or negative effect of vitamins on parasite growth;
though they do appear beneficial for the maintenance of cell
cultures inoculated with pathogens (Choi et al. 2004). The
positive impact of glucose on parasite growth has previously
been described (Upton et al. 1995). Based on the results of
previous studies, standardised in vitro cultivation conditions
(i.e. 37 ◦C, 5% CO2, 95% O2) were adjusted as a reduc-
J. Melicherová et al. / European Journal of Protistology 62 (2018) 101–121 113
Fig. 11. Fluorescence visualisation of F-actin in HCT-8 cell lines inoculated with polystyrene microspheres at 24, 48 and 72 HPI. (A, B)
HCT-8 cells inoculated with fluorescent microspheres (white arrowheads), showing no response to foreign objects at 24 HPI. CLSM. (C, D)
HCT-8 cell lines at 48 HPI, showing a deep imprint only below the fluorescent microsphere (white arrowhead). CLSM. (E, F) HCT-8 cells
with fluorescent microsphere (white arrowhead), showing no response to inoculation with foreign objects at 72 HPI. CLSM. A, C and E
are composite views created by flattening a series of optical sections using the TRITC (F-actin stained with phalloidin), FITC (fluorescent
microspheres) and UV (nuclei stained with Hoechst, fluorescent microspheres) filter sets. B, D and F are views created by flattening a series
of optical sections using only the TRITC (F-actin stained with phalloidin) filter set.
tion in the level of O2 in the atmosphere has been seen to
have a stimulating effect on host cell penetration and subsequent
parasite development (Borowski et al. 2010; Hijjawi
et al. 2001; Upton et al. 1995). In our experiments, while
the effect of O2 reduction on parasite development was not
significant it did have a positive effect on the cell lines, i.e.
cultured cell mortality was reduced and culture viability was
prolonged. Our preliminary (non-published) data indicated
that the multilayers formed by older cultures hindered detection
of attached cryptosporidia. Hence, we used 48-h 70–80%
confluent monolayers exclusively for experimental inoculation
with C. proliferans. While evaluating freshly confluent
(48-h) and aged (up to six-days) HCT-8 cell monolayers for
their ability to support C. parvum infection, another study
showed that it was possible to use cell monolayers up to
three weeks old as they developed the same number of parasite
infection clusters as freshly confluent samples (Sifuentes
and Di Giovanni 2007). In this study, majority of C. proliferans
developmental stages were found near dividing or newly
formed round cells, hereby supporting the hypothesis that
cryptosporidia prefer dividing cultured cells (Widmer et al.
2006).
In contrast to study documenting early trophozoites of
C. parvum after six hours (Borowski et al. 2010), the first
early trophozoites of C. proliferans were detected at 48 h.
Here we should highlight that, while numerous cryptosporidian
sporozoites were observed inspecting potential host cells
with their prolonged apical ends, they often left without invasion.
In accordance with Borowski et al. (2010), the thin
and prolonged apical end is a typical feature for sporozoites
that appear to be making host cell contact, while the sporozoites
isolated from supernatant do not exhibit this apical
prolongation. Moreover, our data (only few attached and successfully
developing parasites observed throughout the entire
experiment) suggest that the HCT-8 and HT-29 cell lines are
appropriate for cultivation of both intestinal (e.g. C. parvum
and C. hominis) and gastric (e.g. C. andersoni) pathogens
that infect humans (Arrowood 2002; Hashim et al. 2006;
114 J. Melicherová et al. / European Journal of Protistology 62 (2018) 101–121
Fig. 12. Fluorescence visualisation of F-actin in HT-29 cell lines inoculated with polystyrene microspheres at 24 and 48 HPI. (A–C) HT-29
cells with fluorescent microsphere (white arrowhead), showing no reaction to foreign objects at 24 HPI. CLSM. (D–F) HT-29 cells with deeply
immersed fluorescent microsphere (white arrowhead), showing no reaction to foreign objects at 48 HPI. CLSM. A and D are composite views
created by flattening a series of optical sections using the TRITC (F-actin stained with phalloidin) and FITC (fluorescent microspheres) filter
sets. B and E are views created by flattening a series of optical sections using the TRITC (F-actin stained with phalloidin) filter set only. C and
F are composite views created by flattening a series of optical sections using the TRITC (F-actin stained with phalloidin), FITC (microspheres)
and UV (nuclei stained with Hoechst, fluorescent microspheres) filter sets.
Jiang et al. 2014; Morada et al. 2016; Perez Cordon et al.
2007; Upton et al. 1994, 1995), but not the gastric pathogen
C. proliferans, which mostly occurs in rodents.
Detailed electron microscopic studies showed that invading
zoites of Cryptosporidium spp. became partially
enveloped by the host cell membrane, resulting in formation
of parasitophorous sac enclosing the entire parasite (e.g.
Lumb et al. 1988; Valigurová et al. 2007, 2008). Unexpectedly,
our in vitro system showed that embracing of parasites
by the cultured cell membrane was not only induced by sporozoites
but also unexcysted or empty oocysts, which were
routinely observed encapsulated by the membrane. Analysis
of oocyst adherence to hosts cells has revealed that
thepresenceofmoleculescontainingN-acetyl-galactosamine
on oocysts may help them attach (Stein et al. 2006). The
lectin-enhanced attachment to host cells could increase the
efficiency of the infection process by bringing sporozoites
into close proximity with host cells. Moreover, the excystation
does not change the lectin binding sites on the oocyst
(Stein et al. 2006). We speculate, the strong adhesion of
oocysts to the cell line surface in our in vitro study could
be explained by an excess of such molecules, the levels of
which could be reduced in vivo while passing through the
host’s digestive system. Numerous unexcysted oocysts with
sutures obviously closed were still present in the cell lines,
even at 48 and 72 HPI. Hence, the question arises as to
whether the oocysts described in other studies (e.g. Choi et al.
2004; Hijjawi et al. 2001; Rosales et al. 1993) were actually
newly-formed oocysts or were simply oocysts from the original
inoculation that had not been washed away during culture
rinsing. Curiously, sutures visible on the unexcysted oocysts
in our study were usually orientated toward the cultured
cell’s surface. Furthermore, sutures were often embedded in
the cell culture and thus were difficult to distinguish from
attached cryptosporidia. This positioning could be the result
of sporozoites’ movement inside the oocyst (in addition to
oocyst balance and the pull of gravity), with such behaviour
helping to shorten the distance between released sporozoites
and the cell surface. Importantly, these oocysts were almost
indistinguishable from advanced developmental stages such
as meronts or gamonts enveloped by parasitophorous sacs,
which could be responsible for the potential incorrect deter-
minationofdevelopmentalstagesinstudiesbasedexclusively
on light and scanning electron microscopy. In this study, we
routinely used tilting and rotation of samples within the scanning
electron microscope in order to confirm the shape of the
cryptosporidian developmental stage and avoid misidentification
as much as possible.
The numerous reported failures to replicate successful
in vitro cultivation of cryptosporidia suggests that completion
of their life cycle on cell lines may be attributable to either
rare coincidence or the flexibility of specific cryptosporid-
J. Melicherová et al. / European Journal of Protistology 62 (2018) 101–121 115
Fig. 13. Scanning electron micrographs of HCT-8 cell lines inoculated with Cryptosporidium proliferans antigen-coated polystyrene microspheres
at 24, 48 and 72 HPI. (A) HCT-8 monolayer with an antigen-coated microsphere (white arrowhead) at 24 HPI. Note the slight
elongation of surrounding microvilli (white arrow). (B) A more detailed, lateral view of the microsphere shown in A. (C) Three antigen-coated
microspheres (white arrowhead) embraced by HCT-8 projections with elongated microvilli (white arrow) at 24 HPI. (D) HCT-8 cells with
microsphere (white arrowhead) at 48 HPI. Note the more intensive reaction, characterised by the presence of numerous elongated microvilli
(white arrows) enveloping the microsphere. (E) A detailed lateral view of the microsphere shown in D. (F) Three antigen-coated microspheres
(white arrowhead) completely covered with HCT-8 cell plasma membrane (white arrow) at 48 HPI. (G) An HCT-8 multilayer at 72 HPI, with
three microspheres (white arrowhead) completely covered with cultured cells (white arrows). (H, I) A detailed top (H) and lateral (I) view of
an antigen-coated microsphere (white arrowhead) enveloped by HCT-8 cell (white arrow) at 72 HPI.
ian isolates to develop under unusual conditions. Between
48 and 72 HPI, many previous studies have recorded a peak
in cryptosporidian development along with the presence of
both asexual and sexual stages, with subsequent decrease in
the parasite number (Aji et al. 1991; Flanigan et al. 1991;
McDonald et al. 1990; Rochelle et al. 2001; Wu et al. 2009).
Despite following the protocols and conditions specified in
previous studies (Choi et al. 2004; Hijjawi et al. 2001; King
et al. 2011; Rosales et al. 1993), we were only able to record
the initial phase of the C. proliferans life cycle, up to the
appearance of trophozoites and meront-/gamont-like stages,
and these were only present in extremely low numbers. We
detected few stages resembling oocysts enveloped by parasitophorous
sac, however, their sporadic occurrence and
location not suitable for SEM tilting/rotation did not allow a
reliable determination.
We used phalloidin staining and confocal laser scanning
microscopy in our study in order to reveal changes in actin filaments
distribution in parasitised cells or cultured cells with
oocysts attached. It is well known that epithelial cells respond
to the presence of cryptosporidia with cytoskeletal modulations
such as actin polymerisation and villin aggregation
(Elliott and Clark 2000). Previous studies described reorganisation
of host F-actin into a plaque-like structure at the
host-parasite interface during parasite invasion, the structure
persisting during parasite development. Similar accumulation
of polymerised actin was also documented at the base and
within parasitophorous sacs surrounding another epicellular
apicomplexan, protococcidian Eleutheroschizon duboscqi
116 J. Melicherová et al. / European Journal of Protistology 62 (2018) 101–121
Fig. 14. Scanning electron micrographs of HT-29 cell lines inoculated with Cryptosporidium proliferans antigen-coated microspheres at 24,
48 and 72 HPI. (A) An HT-29 cell line with a group of antigen-coated microspheres (white arrowhead), showing signs of reaction (white
arrows) to the foreign objects at 24 HPI. (B) Microspheres (white arrowheads) completely embraced by HT-29 cells at 24 HPI. (C) A detailed
top view of the two microspheres shown in B. (D, E) HT-29 cell line with microspheres (white arrowheads) at 48 HPI. Note the intensive
interaction of the HT-29 plasma membrane (white arrow) with the surface of the microspheres. (F) A detailed lateral view of two microspheres
enveloped by HT-29 cell plasma membrane (white arrow) at 48 HPI. (G) Microspheres completely overlain by HT-29 cells at 72 HPI (white
arrowheads). (H) A more detailed view of the microsphere (white arrowhead) shown in G. Note the HT-29 plasma membrane (white arrow)
covering the microsphere. (I) Another view of a microsphere (white arrowhead) completely covered with HT-29 cells (white arrow) at 72 HPI.
(Valigurová et al. 2015). Cryptosporidia, in contrast, showed
only low amounts of F-actin within their parasitophorous
sacs (Bonnin et al. 1999). Interestingly, this study revealed
that even contact with unexcysted oocysts or microspheres
coated with cryptosporidian antigens induces the F-actin
reorganisation in cultured cells, resulting in the formation
of F-actin network surrounding the foreign object. This reaction
was more evident in HT-29 cell lines with visibly shorter
microvilli.
A further topic of this study was to evaluate whether the
unusual enveloping of C. proliferans oocysts by cultured
cell projections was provoked by the parasite itself or represents
an innate reaction of cell lines to foreign objects in
general. For this purpose, we designed a test that simulated
parasitisation of cell lines using an experimental inoculation
of polystyrene microspheres. Pure microspheres failed to
induce an obvious reaction in both types of cell culture (HCT-
8 and HT-29); however, those covered with cryptosporidian
antigens were regularly enveloped by cultured cell plasma
membrane folds. Hence, we conclude that this behaviour of
cell culture is provoked by the parasite and is not innate.
It is usually assumed that a parasite actively manipulates
the target cells before and during invasion (Borowski et al.
2008; Lumb et al. 1988). While motility of apicomplexan
zoites is considered the main mechanism facilitating host
cell invasion, our observations show that motility of C. proliferans
sporozoites was limited and featureless (Melicherová
et al. 2016). This concurs with the study stating that, in
contrast to other apicomplexan zoites (e.g. Plasmodium
spp., Toxoplasma gondii), Cryptosporidium invasion is a
passive process that does not require actomyosin motility
machinery (Forney et al. 1998). Aggregation of Gal-GalNAc
glycoproteins in the plasma membrane of invaded cells at
the sporozoite attachment site initiates a signalling cascade
J. Melicherová et al. / European Journal of Protistology 62 (2018) 101–121 117
Fig. 15. Fluorescence visualisation of F-actin in HCT-8 cell lines inoculated with Cryptosporidium proliferans antigen-coated polystyrene
microspheres at 24, 48 and 72 HPI. (A, B) HCT-8 cells with their projections gradually enveloping the antigen-coated fluorescent microspheres
(white arrowheads) at 24 HPI. CSLM. (C) HCT-8 cells with fluorescent microspheres (white arrowheads) at 48 HPI. Note the cultured cell
membrane folds (white arrows) containing actin filaments that embrace the microspheres. CLSM. (D, E) HCT-8 cells with four deeply
embedded non-fluorescent microspheres (white arrowheads) at 72 HPI. CLSM with transmitted light (D) and CLSM (E). A and C are
composite views created by flattening a series of optical sections using the TRITC (F-actin stained with phalloidin) and FITC (fluorescent
microspheres) filter sets. B is a view created by flattening a series of optical sections using the TRITC (F-actin stained with phalloidin) filter
set only. D and E are composite views created by flattening a series of optical sections using the TRITC (F-actin stained with phalloidin) and
UV (nuclei stained with Hoechst) filter sets.
resultinginactin-dependentmembraneprotrusionandencapsulation
of the sporozoite in a parasitophorous sac (Nelson
et al. 2006). Earlier studies have confirmed the presence of
specific surface lectins on cryptosporidian sporozoites that
may facilitate the attachment of sporozoites to the host cell. In
a study focusing on C. parvum invasion/attachment proteins,
for example, 61 proteins including 27 previously reported,
were identified (Singh et al. 2015). Of these, two have been
widelystudied. The first is p30 (30 kDAGal/GalNAc-specific
lectin), which associates with the mucin-like microneme glycoprotein
gp900, localises on the apical region of C. parvum
and C. hominis sporozoites, binds to epithelial cells and
mediates sporozoite attachment to these cells. The second
is the apical complex glycoprotein CSL (1300-kDa) of C.
parvum sporozoites/merozoites with properties consistent
with being a sporozoite ligand for intestinal epithelial cells
(Bhat et al. 2007; Joe et al. 1994, 1998; Langer and Riggs
1999). Other data, however, suggest that pre-treatment of
parasites with Gal/GalNAc inhibits the entry of C. parvum
into HCT-8 cells and primary bovine cells but has no effect
on entry of either C. parvum or C. hominis into primary
human cells or C. hominis into HCT-8 cells (Hashim et al.
2006). These results confirm that success of in vitro cultivation
depends on the choice of the appropriate cell line
preferred by specific cryptosporidian species. Another work,
comparing the influence of FBS-enriched medium with and
without Gal/GalNAc oligosaccharides and medium without
FBS on trophozoite development, revealed that sporozoites
were positively affected by the presence of Gal/GalNAc in
FBS-medium, accelerating their transformation into trophozoites
(Edwinson et al. 2016).
The outer layer of cryptosporidian oocysts is also covered
by a carbohydrate matrix. This surface coat, the glycocalyx,
plays an important role in modulation of resistance to proteolysis,
antibody binding and adhesion to host cells (Nanduri
et al. 1999). Our experiments revealed that, in addition to
polybeads coated with an antigen ‘cocktail’ (obtained from
oocysts with sporozoites), intact (unexcysted) or even empty
oocysts were able to induce cultured cell plasma membrane
modification and potentially complete envelopment within a
cultured cell membrane-derived sac. Hence, oocyst surface
antigens might not only serve for cell adhesion (Yao et al.
2007) but also represent a likely passive means of host cell
manipulation at the oocyst stage. Furthermore, our data indicate
that the enclosing of oocysts by HT-29 and HCT-8 cells
was induced by the parasite antigens, and that this encapsu-
118 J. Melicherová et al. / European Journal of Protistology 62 (2018) 101–121
Fig. 16. Fluorescence visualisation of F-actin in HT-29 cell lines inoculated with Cryptosporidium proliferans antigen-coated polystyrene
microspheresat24,48and72HPI.(A–C)HT-29cellswithpartiallyimmersedantigen-coatednon-fluorescentmicrospheres(whitearrowheads)
at 24 HPI. CLSM (A, C) and CLSM with transmitted light (B). (D–F) HT-29 cells with deeply immersed fluorescent microspheres that were
partially enveloped by cultured cell actin filaments (white arrowheads) at 48 HPI. Note the crater (black arrowhead) from previously immersed
microsphere that was probably washed away. CLSM. (G–K) The reaction of HT-29 cells to antigen-coated fluorescent microsphere (white
arrowhead) at 72 HPI. The microsphere was completely encapsulated by the cultured cell (white arrow). CLSM. A-C, F and I-K are composite
views created by flattening a series of optical sections using the TRITC (F-actin stained with phalloidin) and UV (nuclei stained with Hoechst,
fluorescent microspheres) filter sets. D and G are composite views created by flattening a series of optical sections using the TRITC (F-actin
stained with phalloidin) and FITC (fluorescent microspheres) filter sets. E and H are views created by flattening a series of optical sections
using the TRITC (F-actin stained with phalloidin) filter set only.
lation of the parasite occurred independently of any active
invasion by motile stages.
Several previous studies reported the occurrence of
extracellular stages of cryptosporidia in cell-free cultures
(Aldeyarbi and Karanis 2015; Boxell et al. 2008; Hijjawi
et al. 2004; Yang et al. 2015). Moreover, some authors claim
that even unexcysted cryptosporidian sporozoites (within
oocysts) can continue to develop in cell-free culture systems
and transform into their next stages (Hijjawi et al.
2010). These authors also observed morphologically distinct
stages and considered them to be newly formed stages
of Cryptosporidium spp. occurring exclusively in cell-free
medium. Simultaneously, they conceded that the use of light
microscopy only made it almost impossible to identify the
exact developmental stage of cryptosporidia dispersed in
medium. Nevertheless, it is important to mention that other
studies have failed to propagate cryptosporidia in cell-free
culture and confirmed that some of the previously reported
extracellular cryptosporidia-like objects could, in fact, be
bacteria or debris (Girouard et al. 2006; Woods and Upton
2007). A study using molecular approaches has reported
the proliferation of C. parvum in cell-free culture with
a measurable, though limited, increase in the concentration
of parasite DNA (Zhang et al. 2009). The latest study
published on cell-free cultivation of cryptosporidia documented
asexual stages only detected under a transmission
electron microscope (Aldeyarbi and Karanis 2015). While a
cell-free culture system would be very helpful for research
focused on Cryptosporidium drug development, our data on
the gastric pathogen C. proliferans indicates that individual
cryptosporidian strains are variously susceptible to in vitro
conditions. Moreover, during our in vivo (Melicherová et al.
2014) and in vitro studies (this study) of C. proliferans, we
did not observe objects resembling the extracellular stages of
J. Melicherová et al. / European Journal of Protistology 62 (2018) 101–121 119
cryptosporidia. Nevertheless, it must be mentioned that light
microscopic observation of the medium within well plates
with inoculated cultures (performed in this study) might not
be sufficient to reach unequivocal conclusion about nonexistence
of extracellular stages in C. proliferans cultivated
in vitro.
Conclusions
In this study, we evaluated the development of the gastric
parasite Cryptosporidium proliferans strain TS03 using
in vitro culture systems HCT-8 and HT29. We documented
free zoites apparently searching for an appropriate infection
site and the presence of advanced stages enveloped by
parasitophorous sac as well as already emptied sacs. Moreover,
we recorded a curious reaction of cultured cells to the
presence of the parasite, when even the unexcysted oocysts
became enveloped by cultured cell projections. Therefore,
we designed an experiment using polystyrene microspheres
to evaluate the response of cell lines to simulated inoculation
with cryptosporidian oocysts to verify their innate and
parasite-induced behaviour. These observations revealed that
cultured cell encapsulation of oocysts is induced by parasite
antigens, independent of any active invasion or motility.
Acknowledgements
The establishment and maintenance of the HCT-8 and
HT-29 cell lines and primary in vitro cultivations of
cryptosporidia were funded through a postdoctoral grant
GPP506/10/P372 from the Czech Science Foundation.
Financial support for the subsequent experimental assays
summarised in this study was provided through project
GAP505/11/1163 of the Czech Science Foundation. The
authors are greatly indebted to Monika Nechvílová from
the Department of Pathological Morphology and Parasitology,
University of Veterinary and Pharmaceutical Sciences
in Brno, for her kind help with laboratory animals. Many
thanks go to Mgr. Kevin Roche BSc. CSc. for English correction
of this manuscript. The authors acknowledge support
from the Department of Botany and Zoology of the Faculty
of Science, Masaryk University, towards the preparation of
this manuscript.
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7.13
Valigurová A., Pecková R., Doležal K., Sak B., Květoňová D., Kváč M.,
Nurcahyo W., Foitová I.
2018
Limitations in the screening of potentially anti-cryptosporidial
agents using laboratory rodents with gastric cryptosporidiosis
Folia Parasitologica 65, 010
http://folia.paru.cas.cz
This is an Open Access article distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/by/4.0),
which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.
The phylum Apicomplexa comprises exclusively parasitic
protists infecting invertebrates and vertebrates,
including humans. One of the most significant and widespread
pathogens are coccidia of the genus Cryptosporidium
Tyzzer, 1907, causative agents of zoonotic disease
(cryptosporidiosis) of the gastrointestinal and respiratory
tracts. In healthy hosts, cryptosporidiosis is self-limiting;
nevertheless, in immunocompromised hosts, it represents a
chronic and debilitating condition (Chen et al. 2002).
Although gastric cryptosporidia have been reported in
fish, reptiles, amphibians, birds and mammals (Jirků et al.
2008, Ryan 2010, Nakamura and Meireles 2015), there is a
dearth of useful studies dealing with the treatment of gastric
cryptosporidiosis. In contrast to intestinal species, the
course of gastric cryptosporidiosis in both immunocompetent
and immunodeficient animals is an asymptomatic
and chronic infection (Kváč et al. 2008, 2011). In humans,
gastric involvement is reported to be very common in patients
with cryptosporidiosis when combined with severe
immunodepression (Rivasi et al. 1999).
The recently described species Cryptosporidium proliferans
Kváč, Havrdová, Hlásková, Daňková, Kanděra,
Ježková, Vítovec, Sak, Ortega, Xiao, Modrý, Chelladurai,
Prantlová et McEvoy, 2016 (previously known as strain
TS03 of Cryptosporidium muris Tyzzer, 1907), used in
this study, develops exclusively in the glandular part of the
stomach, similar to C. muris and Cryptosporidium andersoni
Lindsay, Upton, Owens, Morgan, Mead et Blagburn,
Research Article
Address for correspondence: Ivona Foitová, Department of Botany and Zoology, Faculty of Science, Masaryk University, Kotlářská 2, 611 37 Brno,
Czech Republic Phone: +420 549 494 447; Fax: +420 549 498 331; E-mail: foitova@sci.muni.cz
Institute of Parasitology, Biology Centre CAS
Folia Parasitologica 2018, 65: 010
doi: 10.14411/fp.2018.010
Limitations in the screening of potentially anti-cryptosporidial
agents using laboratory rodents with gastric cryptosporidiosis
Andrea Valigurová1
, Radka Pecková1
, Karel Doležal2
, Bohumil Sak3
, Dana Květoňová3
, Martin Kváč3,4
, Wisnu
Nurcahyo5
, Ivona Foitová1,5
1
	Department of Botany and Zoology, Faculty of Science, Masaryk University, Brno, Czech Republic;
2
	Department of Chemical Biology and Genetics & Laboratory of Growth Regulators, Centre of the Region Haná for Biotechnological
and Agricultural Research, Faculty of Science, Palacký University, and Institute of Experimental Botany, Academy of Sciences of
Czech Republic, Olomouc-Holice, Czech Republic;
3
	Institute of Parasitology, Biology Centre of the Czech Academy of Sciences, České Budějovice, Czech Republic;
4
	Department of Animal Husbandry Sciences, Faculty of Agriculture, University of South Bohemia in České Budějovice, Czech Republic;
5
	Department of Parasitology, Faculty of Veterinary Medicine, Gadjah Mada University, Yogyakarta, Indonesia
Abstract:The emergence of cryptosporidiosis, a zoonotic disease of the gastrointestinal and respiratory tract caused by Cryptosporidium
Tyzzer, 1907, triggered numerous screening studies of various compounds for potential anti-cryptosporidial activity, the majority
of which proved ineffective. Extracts of Indonesian plants, Piper betle and Diospyros sumatrana, were tested for potential anticryptosporidial
activity using Mastomys coucha (Smith), experimentally inoculated with Cryptosporidium proliferans Kváč, Havrdová,
Hlásková, Daňková, Kanděra, Ježková, Vítovec, Sak, Ortega, Xiao, Modrý, Chelladurai, Prantlová et McEvoy, 2016. None of the
plant extracts tested showed significant activity against cryptosporidia; however, the results indicate that the following issues should
be addressed in similar experimental studies. The monitoring of oocyst shedding during the entire experimental trial, supplemented
with histological examination of affected gastric tissue at the time of treatment termination, revealed that similar studies are generally
unreliable if evaluations of drug efficacy are based exclusively on oocyst shedding. Moreover, the reduction of oocyst shedding did
not guarantee the eradication of cryptosporidia in treated individuals. For treatment trials performed on experimentally inoculated
laboratory rodents, only animals in the advanced phase of cryptosporidiosis should be used for the correct interpretation of pathological
alterations observed in affected tissue. All the solvents used (methanol, methanol-tetrahydrofuran and dimethylsulfoxid) were shown to
be suitable for these studies, i.e. they did not exhibit negative effects on the subjects. The halofuginone lactate, routinely administered
in intestinal cryptosporidiosis in calves, was shown to be ineffective against gastric cryptosporidiosis in mice caused by C. proliferans.
In contrast, the control application of extract Arabidopsis thaliana, from which we had expected a neutral effect, turned out to have
some positive impact on affected gastric tissue.
Keywords: Cryptosporidium, gastric, oocyst, pathology, treatment
doi: 10.14411/fp.2018.010	 Screening of potentially anti-cryptosporidial agents
Folia Parasitologica 2018, 65: 010	 Page 2 of 17
2000, with a life cycle corresponding to that of C. muris
(see Tyzzer 1910, Melicherová et al. 2014, Kváč et al.
2016). Though C. proliferans had been considered identical
with C. muris in previous years, their clinical courses of
parasitisation in Mastomys coucha (Smith) differ considerably
(Kváč et al. 2016). Compared to C. muris, rodents
shed oocysts of C. proliferans for a much longer period
and at a greater intensity, and only C. proliferans induces
significant clinical and pathological changes, such as
weight loss and massive proliferation of the gastric mucosa
associated with a considerable increase in stomach weight
(Kváč et al. 2016). The change in the ratio of glandular to
non-glandular surfaces from 55 : 45 to 80 : 20, detected in
M. coucha infected with C. proliferans, was not observed
in C. muris infection (Kváč et al. 2016).
Cryptosporidiosis is recognised as a major medical concern
as there is no effective treatment for either intestinal
or gastric cryptosporidiosis (Fayer et al. 2000, Thompson
et al. 2005). Numerous compounds have been screened in
vitro and in vivo for potential anti-cryptosporidial activity,
but the majority turned out to be ineffective and only
a few agents have shown promise. The most commonly
used drugs used against cryptosporidiosis include antibiotics
(e.g. paromomycin or azithromycin) and halofuginone,
which are partially effective. In contrast to chemotherapeutics,
often with considerable side effects and a certain
level of toxicity, the use of natural products or dietary supplements
with anti-cryptosporidial activity could represent
a new and safe approach to the effective pharmacological
control of cryptosporidiosis. For example, L-arginine was
shown to have a protective role during infection with Cryptosporidium
parvum Tyzzer, 1912 in undernourished mice
(Castro et al. 2012). Several studies have found probiotics
to be effective against cryptosporidiosis in humans and animals,
reporting prompt clinical improvement and resolution
of the infection following treatment (Rotkiewicz et al.
2001, Pickerd and Tuthill 2004). Lactobacillus spp. significantly
reduced the viability of oocysts of C. parvum (see
Foster et al. 2003). Administration of exogenous agmatine
seems to alter the metabolism of C. parvum enough to interfere
with its ability to colonise the mammalian intestine
(Moore et al. 2001). Mangiferin, widely distributed in
higher plants and one of the constituents of folk medicines
(Yoshimi et al. 2001), has significant anti-cryptosporidial
activity comparable to the same dose (100 mg/kg/day) of
paromomycin (Tarantino et al. 2004, Perrucci et al. 2006).
Curcumin, which is active against a variety of diseases,
was found to be effective against C. parvum in cell cultures
(Shahiduzzaman et al. 2009).
Garlic (Allium sativum) appears to be a prophylactic and
a promising therapeutic agent, as it successfully eradicated
cryptosporidial oocysts from the faeces and intestines of
infected immunocompetent mice that had received garlic
two days before the experimental infection and continued
for two weeks (Gaafar 2012). The administration of
garlic to human HIV patients with chronic diarrhoea and
confirmed cryptosporidiosis resulted in complete or partial
remission (Fareed et al. 1996). Onion (Allium cepa) and
cinnamon (Cinnamomum zeylanicum) oils also turned out
to be effective against the infection with C. parvum in mice
(Abu El Ezz et al. 2011). In contrast, an in vitro study using
HCT-8 cells inoculated with C. andersoni did not confirm
the significant inhibition of parasite growth when exposed
to garlicin (antifungal component extracted from garlic)
(Wu et al. 2011). The authors concluded that garlicin inhibits
the growth of cryptosporidia in vivo by enhancing
macrophage activity, rather than by exerting direct effects
on the parasite. This study also showed anti-cryptosporidial
activity of ginkgolic acids extracted from maidenhair
tree (Ginkgo biloba) sarcotesta. With the exception of the
prophylactic effect of garlic, however, none of these natural
products were able to completely eradicate crypto-
sporidiosis.
As humans and orang-utans exhibit phylogenetic similarities
(Grehan and Schwartz 2009), we focused on orangutans’
feeding behaviour with an emphasis on specific
plants consumed that would lead to a reduction in parasite
infections. Recently we documented a secondary self-medication
(the external application of a medicinal substance)
in Bornean orang-utans (Morrogh-Bernard at al. 2017).
These findings validate the anti-inflammatory properties of
Dracaena cantleyi and its application to muscles and joints
by orang-utans, and may serve as the first evidence for the
deliberate external application of substances with bioactive
potential for self-medication in great apes.
We selected few plants including Piper betle and Diospyros
sumatrana with promising antiparasitic activity on
the basis of behavioural data and decreases in parasite load
(Foitová et al. 2010). Our analyses show a positive correlation
between the prevalence of these plant species in
orang-utan diets and the presence of parasites (based on
the Jaccard index of known frequency in nature) that cannot
be explained by their prevalence in the environment.
The betel, P. betle, has been used as a medicinal plant in
traditional medicine throughout South and South East Asia
since ancient times. Experimental studies have revealed its
wide and diverse biological and pharmacological effects
(Pecková et al. 2018). Diospyros sumatrana has not yet
been studied for pharmacological potential, but our study
shows its possible potential.
This study aimed to test extracts of Indonesian plants
selected by orang-utans for self-medication for potential
anti-cryptosporidial activity, using a rodent host that had
been experimentally inoculated with C. proliferans. The
extract of Arabidopsis thaliana (the Eurasian plant routinely
used as a model in research laboratories) was used
as a control with an expected neutral effect. Halofuginone
lactate (Halocur), an oral solution used for the treatment of
cryptosporidiosis in calves, was tested for its potentially
positive effect.
MATERIALS AND METHODS
Preparation of plant extracts
The dried leaves obtained from Piper betle (akar sirih), Diospyros
sumatrana (kayu hitam) and Arabidopsis thaliana (thale
cress) were homogenised to a fine powder in liquid nitrogen.
Portions of the ground material (0.33 g) were then extracted sep-
doi: 10.14411/fp.2018.010	 Screening of potentially anti-cryptosporidial agents
Folia Parasitologica 2018, 65: 010	 Page 3 of 17
arately in 10 ml of water, methanol (methanol) or methanol-tetrahydrofurane
(methanol-THF, 1 : 1). After 16 hours of extraction
(overnight) at -20 °C, the resulting homogenates were centrifuged
(26,000 g, 4 °C, 20 min); the sediments were then re-extracted
for one hour in the same way and centrifuged. Afterward, these
two supernatants were pooled and dried in a vacuum at 35 °C,
and then dissolved in 100 µl of pure Dimethylsulfoxid (DMSO),
except for samples dissolved in sterile water, which were further
diluted in sterile water.
The parasite used in this study
The gastric species Cryptosporidium proliferans used in this
study and our previous studies (Kváč et al. 2008, 2011, 2016
Melicherová et al. 2014, 2016) originated from a naturally infected
East African mole rat Tachyoryctes splendens (Rüppell) and
was kept in severe combined immunodeficiency (SCID) mice and
southern multimammate mice (Mastomys coucha) under laboratory
conditions.
Laboratory animals and experimental inoculations with
oocysts of Cryptosporidium proliferans.
Eight-week old M. coucha mice (Biology Centre, CAS, České
Budějovice) were used for this study. To prevent environmental
contamination with oocysts, each group of mice was housed in
plastic cages with sterile wood-chip bedding and supplied with
sterilised food and water ad libitum. The rearing of animals was
regulated by Czech legislation (Act No. 246/1992 Coll., on protection
of animals against cruelty); these documents are consistent
with legislation by the European Commission. All housing,
feeding, and experimental procedures were conducted under protocols
approved by the Institute of Parasitology, Biology Centre,
CAS and Institute and National Committees (Protocols No.
52/2014).
For the experimental inoculation of mice, oocysts collected
from faeces were purified using Sheather’s sugar flotation method
(Arrowood and Sterling 1987) and modified caesium chloride
gradient centrifugation (Kilani and Sekla 1987). Each mouse was
inoculated orally by an oesophagus tube with a dose of 106 viable
oocysts of C. proliferans. Afterwards, fresh mouse faeces were
collected daily in the morning and examined microscopically for
the presence of oocysts using staining according to Miláček and
Vítovec (1985). The intensity of oocyst excretion was assessed as
the number of oocysts per gram of faeces (OPG) as previously described
Kváč et al. (2007). In addition, faecal consistency, faecal
colour and general health status were examined daily.
The treatment of parasitised animals using plant extracts
The potential antiparasitic effect of Indonesian plants (P. betle
and D. sumatrana), was compared with the expected null effect
of A. thaliana, as well as with the potentially positive effect
of the Halocur oral solution (Intervet Production S.A., Rue de
Lyons, France). Mice infected with C. proliferans two or three
months before treatment with plant extracts were divided into
groups (three animals per group) and treated with the following
treatment doses administered per os. Treated non-infected and
untreated infected and non-infected control groups were included
in all experiments. The effect of administered extracts/drugs/
diluents on the course of parasitisation was evaluated as change
in the parasitisation intensity expressed by OPG in comparison to
parasitisation intensity of infected mice administered with only
distilled water. The coefficient of determination (r2
) was calculated
for each linear regression. All computation was carried out
with the SigmaPlot 13.0 (Systat Software Inc., San Jose, CA).
The histopathological changes of parasitised gastric mucosa were
evaluated post mortem.
Trial 1. Groups of mice were treated daily for 14 days, beginning
two months post inoculation with C. proliferans, with
12.5 mg per 100 g of body mass (BM) of either P. betle, D. sumatrana
or A. thaliana extracted in methanol, dissolved in DMSO,
and diluted with sterile water to obtain a final concentration of
0.5% DMSO. The effect of the Halocur (100 µg/kg BM) and the
diluent (0.5% DMSO in sterile water) was evaluated in infected
control mice.
Trial 2. A second trial was conducted after the completion of
the first trial. The treatments began three months post inoculation
with C. proliferans. Three extraction media were used: methanol,
methanol-THF and sterile water. Extracted material was dissolved
in DMSO (except for that dissolved in sterile water) and
diluted with sterile water to obtain a final concentration of 0.5%
DMSO. A dose of 40 mg of per 100 g BM of either P. betle, D. sumatrana
or A. thaliana extract was administered twice a day for
21 days. Additionally, the effect of the Halocur (100 µg/kg BM)
and the diluent (0.5% DMSO in sterile water) was evaluated in
infected control mice.
Parasitological dissection and tissue processing for microscopic
evaluation
After either 14 (Trial 1) or 21 (Trial 2) days of treatment, control
and treated animals were euthanised by cervical dislocation
and dissected according to protocols described by Melicherová
et al. (2014). For histological sectioning, gastric tissue was fixed
in AFA (Alcohol-Formalin-Acetic Acid) solution and processed
according to Valigurová et al. 2008. The blocks were cut using a
Zeiss Hyrax M 300 rotary microtome and the 7 µm thick sections
were stained with haematoxylin-eosin. Preparations were viewed
using an Olympus BX61 microscope.
For scanning electron microscopy, samples of gastric tissue
were fixed overnight at 4 °C in freshly prepared 2.5% glutaraldehyde
(v/v) in cacodylate buffer (0.1 M; pH 7.4), washed
3  ×  15  min in the buffer, postfixed in 2% OsO4
in cacodylate
buffer for two hour at room temperature, and washed again
3 × 15 min in buffer. After dehydration in a graded acetone series,
specimens were critical point-dried using CO2
, coated with gold,
and examined using a JEOL JSM-7401F – Field Emission Scanning
Microscope. Abbreviations used in Figs. 1–9: LM – light
microscopy; SEM – scanning electron microscopy
RESULTS
Histopathological observations of the gastric tissue
of uninfected and infected rodents
In the stomach of healthy (control) Mastomys coucha
individuals, the surface of the gastric mucosa was smooth
with a brain-like ornamentation (Fig. 1A). In histological
sections stained with haematoxylin-eosin, the gastric
mucosa, along with a thin layer of muscularis mucosae
and subjacent submucosa, appeared homogeneously
pink with well-demarcated blue nuclei (Fig. 1B). It was
doi: 10.14411/fp.2018.010	 Screening of potentially anti-cryptosporidial agents
Folia Parasitologica 2018, 65: 010	 Page 4 of 17
possible to distinguish quite easily the tubular glands
and necks ended by pits invaginating the luminal surface
of mucosa (Fig. 1B,C). The gastric pits and glands were
obviously constricted and contained within a thin lamina
propria (Fig. 1A–F). A tall simple columnar epithelium
lined the mucosal surface and gastric pits (Fig. 1B). Surface
mucous cells lining the pits as well as mucous neck
cells demarcating the necks of gastric glands appeared pale
(Fig. 1B), while the cells forming the base of glands were
stained darker with prominent, intensively stained nuclei
(Fig. 1C,F).
In mice infected with Cryptosporidium proliferans, parasite
endogenous stages were restricted to the epithelial
cells in the glandular part of the gastric mucosa. This species
primarily parasitises epithelial cells lining the gastric
pits and glands, though some parasites could be found attached
to cells lining the luminal surface of the gastric epithelium,
especially as parasitisation progressed (most likely
due to the increased space requirements). The duration
of the prepatent period (18–21 days) and the chronology of
pathological changes correspond to previously published
data (Melicherová et al. 2014).
In the first trial, the use of mice in a relatively early
stage of cryptosporidiosis (two months post inoculation
with C. proliferans) was shown to be unsuitable for the microscopic
evaluation of treatment effects, as affected gastric
mucosa exhibited only mild to moderate pathological
changes. The gastric tissue was also irregularly affected by
cryptosporidia in an island-like manner, where individual
parasitised gastric pits were surrounded by regions of
healthy epithelium. The affected gastric tissue showed no
obvious alterations and the pits appeared almost fully constricted
when evaluated under SEM (Fig. 2A). Despite the
mild character of pathological alterations visible by SEM,
Fig. 1. Healthy gastric mucosa of Mastomys coucha (Smith). A – general view of surface of the gastric mucosa exhibiting constricted
gastric pits; note the remnants of mucus not washed away during rinsing (SEM); B – general view of gastric mucosa showing the longitudinally
sectioned pits and glands (LM); C – general view of mucosa with cross-sectioned gastric glands (LM, histology); D – detailed
view of constricted gastric pits in longitudinal section (LM, histology); E – detailed view of constricted pits in cross section (LM, histology);
F – detailed view of constricted glands in cross sections (LM, histology); asterisk – gastric pit; m – mucosa; mm – muscularis
mucosae; sm – submucosa; white arrow – mucus with cell debris.
C
D
A B
FE
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Folia Parasitologica 2018, 65: 010	 Page 5 of 17
histological sectioning revealed the moderate dilatation
of gastric glands due to the presence of numerous cryptosporidia
(Fig. 2B).
Pathological changes in affected gastric tissue became
more prominent three months post inoculation with C. proliferans,
when the parasitisation entered a chronic phase
(Fig. 3A–I). Such mice, used in Trial 2, had the surface
of their gastric glandular epithelium markedly deformed
due to intense pathological changes. At the macroscopic
level, the gastric mucosa was typified by a cauliflower-like
appearance. The epithelium proliferated into the luminal
space, so that the stomach exhibited extensive folding that
was especially visible under SEM (Fig. 3A,C,D). This was
the result of an increase in the volume of the lamina propria,
which then increased the distance between individual
gastric glands and caused the longitudinal folds to become
twisted and obviously deformed. The progress of parasitisation
caused a distinctive form of diffuse mucosal hypertrophy
typified by the presence of enlarged/giant gastric
folds and intensive epithelial hyperplasia.
The gastric glands, packed with various developmental
stages of C. proliferans and necrotic material, were markedly
dilated and hypertrophied (Fig. 3B,E–I). In addition,
numerous cryptosporidia developed attached to the luminal
surface of the gastric mucosa outside the dilated pits (Fig.
3D). The affected glands, lined with many undifferentiated
cells, lost their normal architecture; the atrophic epithelial
cells of the affected glands exhibited cuboidal or squamous
metaplasia (Fig. 3G–I). Besides the more intense staining
of affected tissue in histological sections, another feature
typical of advanced cryptosporidiosis was the thickening
of muscularis mucosae (Fig. 3B). Parasitised tissue exhibited
various degrees of oedema and the infiltration of
the lamina propria and submucosa with neutrophils (Fig.
3B,E). Stomach weight (due to proliferating mucosa) and
epithelial height were considerably greater than in non-infected
animals. Interestingly, despite the chronicity of infection,
no clinical signs were observed in infected rodents.
Treatment with plant extracts and Halocur
Trial 1 served as the primary screening of experimental
protocols and the selected treatment doses based on related
literature and empirical data on other unicellular parasites
used in our research (Pecková et al. 2018). On the basis of
the results obtained with daily doses of 12.5 mg/100 g BM
applied for 14 consecutive days, we decided to increase
the treatment doses to 40 mg per 100 g BM administered
twice daily for 21 consecutive days in Trial 2. Furthermore,
in Trial 2, we tested and compared the efficacy of selected
plant extracts obtained using various solvent media –
methanol, methanol-THF, and sterile water. In both trials,
an oral administration containing 100 µg/kg BM halofuginone
lactate (Halocur), a salt whose antiprotozoal properties
and efficacy against Cryptosporidium parvum have
been demonstrated under in vitro and in vivo conditions
(Giadinis et al. 2008, Petermann et al. 2014), was used as
a control treatment.
One of the parameters used to evaluate parasitisation intensity
was the monitoring of the number of shed oocysts
detected in faeces. Generally, the variations in oocyst shedding
were comparable in all experimental groups treated
with either plant extracts or Halocur. A decline in oocyst
number in Trial 1 can be observed in groups treated for
14 days with Diospyros sumatrana extracted by methaFig.
2. Pathological alterations to gastric mucosa induced by Cryptosporidium proliferans Kváč, Havrdová, Hlásková, Daňková,
Kanděra, Ježková, Vítovec, Sak, Ortega, Xiao, Modrý, Chelladurai, Prantlová et McEvoy, 2016 in control Mastomys coucha (Smith)
from Trial 1. A – superficial view of surface of the gastric mucosa exhibiting slightly enlarged gastric pits (SEM); B – general view of
the gastric mucosa with longitudinally sectioned pits. The gastric pits and glands exhibit moderate dilation when viewed in tangential
and cross sections (LM, histology); asterisk – gastric pit, black arrowhead – cryptosporidia, m – mucosa, white arrow – mucus with cell
debris, white arrowhead – cryptosporidia-free pit/gland.
A B
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Folia Parasitologica 2018, 65: 010	 Page 6 of 17
Fig. 3. Pathological alterations to gastric mucosa induced by Cryptosporidium proliferans Kváč, Havrdová, Hlásková, Daňková,
Kanděra, Ježková, Vítovec, Sak, Ortega, Xiao, Modrý, Chelladurai, Prantlová et McEvoy, 2016 in control Mastomys coucha (Smith)
from Trial 2. A – superficial view of the gastric mucosa exhibiting extensive folding and intense parasitisation (SEM); B – general
view of the gastric mucosa and submucosa in longitudinal section (LM, histology); C, D – detailed view of dilated pits filled with
numerous parasites (SEM); E – gastric mucosa showing the longitudinally sectioned pits and glands (LM, histology); F–H – detailed
view of parasitised pits and glands in longitudinal section (LM, histology); I – detailed view of cross-sectioned gastric glands filled
with parasites (LM, histology); asterisk – gastric pit; black arrowhead – cryptosporidia; m – mucosa; mm – muscularis mucosae; sm –
submucosa; white arrow – mucus with cell debris; white arrowhead – cryptosporidia-free pit/gland.
C
E
H
D
G
F
I
A B
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nol and Halocur (Fig. 4). In contrast, non-treated controls,
controls treated with pure DMSO and animals treated with
Piper betle and Arabidopsis thaliana extracted by methanol
exhibited an increase in oocyst shedding (Fig. 4). In
Trial 2, a decline in oocyst shedding occurred in groups
treated for 21 days with A. thaliana by methanol-THF, D.
sumatrana by methanol-THF and Halocur (Fig. 5). An increase
occurred in non-treated controls and animals treated
with pure DMSO, P. betle extracted by methanol, P. betle
by methanol-THF, P. betle by sterile water, A. thaliana by
methanol A. thaliana by sterile water, D. sumatrana by
methanol, and D. sumatrana by sterile water (Fig. 5). The
coefficient of determination, however, was low in all experimental
groups, with the exception of non-treated controls
and controls treated with pure DMSO in Trial 2 (Fig. 5).
Despite an obvious decrease in oocyst shedding in
some animals, post mortem histological examinations at
the end of both trials revealed heavy cryptosporidiosis in
all non-treated (Figs. 1–3) and treated animals (Figs. 6–9).
Only histological sections from animals treated for 21 days
(Trial 2) are shown to demonstrate the status of parasitised
gastric tissue at the end of the trial. Histopathological data
show that, independently of the solvent medium used, extracts
from P. betle (Fig. 6A–I) and D. sumatrana (Fig.
7A–I) neither helped to eradicate the parasites nor cured
the pathological changes induced by C. proliferans. When
compared to non-treated controls (Fig. 3A–I), no significant
difference was observed in parasitisation intensity or
in associated pathological alterations to the gastric mucosa
in treated animals examined in SEM or histological prepa-
rations.
Surprisingly, the application of a control with no expected
effect, the extract of A. thaliana (Fig. 8A–I), seemed
to have a positive impact on parasitised gastric mucosa,
especially when dissolved in methanol-THF (Fig. 8G–I).
This was obvious especially in SEM preparations, where
the pathological folding of parasitised gastric tissue appeared
less intense (Fig. 8G) when compared to those in
non-treated animals (Fig. 3A,C,D) and the effects of other
treatments (Figs. 7A,D,G, and 8A,D). Some degree of
this effect of A. thaliana on parasitised gastric mucosa was
also visible in histological sections; i.e. the architecture of
the gastric glands was closer to the normal state and the
epithelial cells lining the gastric glands were slightly less
atrophic (Fig. 8I).
Although it slightly decreased oocyst excretion, even
prolonged treatment with Halocur was insufficiently effective
in treating gastric cryptosporidiosis (Fig. 9A–C). The
Fig. 4. Infection dynamics of Cryptosporidium proliferans Kváč, Havrdová, Hlásková, Daňková, Kanděra, Ježková, Vítovec, Sak, Ortega,
Xiao, Modrý, Chelladurai, Prantlová et McEvoy, 2016 in Trial 1. Groups of eight-week-old Mastomys coucha (Smith) inoculated
with a dose of 106 oocysts two months before treatment and subsequently treated daily for 14 days with 12.5 mg per 100 g of body mass
of either Piper betle, Diospyros sumatrana or Arabidopsis thaliana extracted in methanol. The linear regression including regression
coefficient is included for each experimental group.
2,200,000
2,000,000
1,800,000
1,600,000
1,400,000
1,200,000
1,000,000
2,200,000
2,000,000
1,800,000
1,600,000
1,400,000
1,200,000
1,000,000
Day of treatment
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Fig. 5. Infection dynamics of Cryptosporidium proliferans Kváč, Havrdová, Hlásková, Daňková, Kanděra, Ježková, Vítovec, Sak, Ortega,
Xiao, Modrý, Chelladurai, Prantlová et McEvoy, 2016 in Trial 2. Groups of eight-week-old Mastomys coucha (Smith) inoculated
with a dose of 106 oocysts three months previously were treated twice a day for 21 days with a dose of 40.0 mg per 100 g of body mass
of either Piper betle, Diospyros sumatrana or Arabidopsis thaliana extracted in methanol, ethanol-tetrahydrofuran (methanol-THF) or
sterile water. The linear regression including regression coefficient is included for each experimental group.
3,500,000
3,000,000
2,500,000
2,000,000
1,500,000
1,000,000
3,500,000
3,000,000
2,500,000
2,000,000
1,500,000
1,000,000
3,500,000
3,000,000
2,500,000
2,000,000
1,500,000
1,000,000
3,500,000
3,000,000
2,500,000
2,000,000
1,500,000
1,000,000
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Fig. 6. The effect of Piper betle extract on Mastomys coucha (Smith) gastric mucosa parasitised with Cryptosporidium proliferans
Kváč, Havrdová, Hlásková, Daňková, Kanděra, Ježková, Vítovec, Sak, Ortega, Xiao, Modrý, Chelladurai, Prantlová et McEvoy, 2016
in Trial 2. A–C – P. betle in sterile water: A – superficial view of the gastric mucosa (SEM); B – general view of the gastric mucosa in
longitudinal section (LM, histology); C – detailed view of cross-sectioned glands filled with parasites (LM, histology); D–F – P. betle
in methanol: D – superficial view of the gastric mucosa (SEM); E – general view of the gastric mucosa in longitudinal section (LM,
histology); F – detailed view of cross-sectioned glands filled with parasites (LM, histology); G–I – P. betle in methanol-THF: G –
superficial view of the gastric mucosa (SEM); H – general view of the gastric mucosa and submucosa in longitudinal section (LM,
histology); I – detailed view of cross-sectioned glands filled with parasites (LM, histology); asterisk – gastric pit; black arrowhead
– cryptosporidia; m – mucosa; mm – muscularis mucosae; sm – submucosa; white arrow – mucus with cell debris; white arrowhead –
cryptosporidia-free pit/gland.
C
E
H
D
G
F
I
A B
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Fig. 7. The effect of Diospyros sumatrana extract on Mastomys coucha (Smith) gastric mucosa parasitised with Cryptosporidium
proliferans Kváč, Havrdová, Hlásková, Daňková, Kanděra, Ježková, Vítovec, Sak, Ortega, Xiao, Modrý, Chelladurai, Prantlová et
McEvoy, 2016 in Trial 2. A–C – D. sumatrana in sterile water: A – superficial view of the gastric mucosa (SEM); B – general view of
the gastric mucosa in longitudinal section (LM, histology); C – detailed view of tangentially-sectioned glands filled with parasites (LM,
histology); D–F – D. sumatrana in methanol: D – superficial view of the gastric mucosa (SEM); E – general view of the gastric mucosa
in longitudinal section (LM, histology); F – detailed view of cross-sectioned glands filled with parasites and neighbouring empty glands
(LM, histology); G–I – D. sumatrana in methanol-THF: G – superficial view of the gastric mucosa (SEM); H – general view of the
gastric mucosa in longitudinal section (LM, histology); I – detailed view of tangentially-sectioned glands filled with parasites (LM,
histology); asterisk – gastric pit, black arrowhead – cryptosporidia; m – mucosa; mm – muscularis mucosae; sm – submucosa; white
arrow – mucus with cell debris; white arrowhead – cryptosporidia-free pit/gland.
C
E
H
D
G
F
I
A B
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Fig. 8. The effect of Arabidopsis thaliana extract on Mastomys coucha (Smith) gastric mucosa parasitised with Cryptosporidium
proliferans Kváč, Havrdová, Hlásková, Daňková, Kanděra, Ježková, Vítovec, Sak, Ortega, Xiao, Modrý, Chelladurai, Prantlová et
McEvoy, 2016 in Trial 2. A–C – A. thaliana in sterile water: A – superficial view of the gastric mucosa (SEM); B – general view of
the gastric mucosa in longitudinal section (LM, histology); C – detailed view of cross-sectioned glands filled with parasites (LM,
histology); D–F – A. thaliana in methanol: D – superficial view of the gastric mucosa (SEM); E – general view of the gastric mucosa
in longitudinal section (LM, histology); F – detailed view of cross-sectioned glands filled with parasites (LM, histology); G–I – A.
thaliana in methanol-THF: G – superficial view of the gastric mucosa (SEM); H – general view of the gastric mucosa in longitudinal
section (LM, histology); I – detailed view of cross-sectioned glands filled with parasites (LM, histology); asterisk – gastric pit; black
arrowhead – cryptosporidia; m – mucosa; mm – muscularis mucosae; sm – submucosa; white arrow – mucus with cell debris; white
arrowhead – cryptosporidia-free pit/gland.
C
E
H
D
G
F
I
A B
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generally recommended dose for intestinal cryptosporidiosis,
corresponding to 100 µg/kg BM of halofuginone administered
for seven consecutive days, was reported to be
effective in the past (Giadinis et al. 2008, Petermann et al.
2014). In our study, the period of Halocur administration
was extended to 14 (Trial 1) and 21 days (Trial 2). Treated
animals showed no symptoms of toxicity (no clinical
signs of overdosing). However, SEM observations of the
intensively folded surface of the gastric mucosa, with the
presence of various developmental stages of C. proliferans,
confirmed heavy infection (Fig. 9A). Histopathological
changes (Fig. 9A–C) were very similar to those found
in untreated individuals (Fig. 3A–I). The only difference,
noticeable at higher magnification, was the better preservation
of epithelial cells lining the glands (Fig. 9C). These
cells appeared less atrophic with more preserved nuclei
compared to non-treated animals or those treated with
P. betle and D. sumatrana.
DISCUSSION
Histopathological changes related to gastric
cryptosporidiosis
Studies on animal gastric cryptosporidiosis usually report
only mild histopathological changes represented by
the dilatation and epithelial metaplasia of gastric glands.
These are generally reported without obvious alterations to
host health status, and with no or only insignificant inflammatory
responses in the lamina propria, although inflammatory
infiltrates are occasionally seen (e.g. Taylor et al.
1999, Masuno et al. 2006, Kváč et al. 2008). Nevertheless,
there are obvious differences in histopathological changes
induced by different gastric species, load of parasite inoculum
or immunological status of their host.
In addition, other pathogens might escalate the impact
of cryptosporidiosis on host tissue. For example, in mice
simultaneously infected with Cryptosporidium muris and
Helicobacter felis, the gastric glands were more severely
parasitised by cryptosporidia, and their stomachs showed
more severe cellular infiltrates (Tatar et al. 1995). The fundus
glands of nude mice inoculated with strain RN 66 of
C. muris showed dilatation with mild epithelial changes
(Taylor et al. 1999), and only mice receiving an inoculum
of 1,000,000 oocysts showed an inflammatory reaction.
A study using SCID mice reported only mild gastric
cryptosporidiosis without gross pathologic findings
(Jalovecká et al. 2010). The maximum peak of parasitisation
intensity was 21 DPI, whereas the highest numbers of
immune cells occurred in the gastric epithelium at 28 DPI,
a period when the majority of mice had already been cured
of the infection. Immunocompetent mice inoculated with
either C. muris CB03 or Cryptosporidium proliferans developed
T cell responses leading to a clearance of the primary
infection and complete resistance to re-infection with
the same strain (Jalovecká et al. 2010, Kváč et al. 2011).
In contrast, the intensity of infection with C. proliferans
in experimentally infected Mastomys coucha continued
to increase throughout experiments with maximum oocysts’
shedding at 126 DPI and animals developed lifelong
(chronic) infection (Melicherová et al. 2014, Kváč et al.
2016).
Despite the considerable gross pathology of gastric epithelium
documented in this and previous studies (Melicherová
et al. 2014, Kváč et al. 2016), neither clinical
signs of cryptosporidiosis, nor weight lost were observed
in southern multimammate mice. In contrast to previous
studies (Melicherová et al. 2014, Kváč et al. 2016), we observed
the inflammatory infiltration of muscularis mucosae
Fig. 9. The effect of Halocur on Mastomys coucha (Smith) gastric mucosa parasitised with Cryptosporidium proliferans Kváč, Havrdová,
Hlásková, Daňková, Kanděra, Ježková, Vítovec, Sak, Ortega, Xiao, Modrý, Chelladurai, Prantlová et McEvoy, 2016 in Trial 2.
A – superficial view of the gastric mucosa, halocur in PBS (SEM); B – general view of the gastric mucosa in longitudinal section,
halocur in PBS (LM, histology); C – detailed view of cross-sectioned glands filled with parasites, halocur in PBS (LM, histology);
asterisk – gastric pit, black arrowhead – cryptosporidia, m – mucosa, mm – muscularis mucosae, sm – submucosa, white arrow – mucus
with cell debris, white arrowhead – cryptosporidia-free pit/gland.
CA B
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and submucosa with neutrophils. Great variations in oocyst
shedding accompanied by heavy cryptosporidiosis revealed
in histological sections of southern multimammate
mice stomach in this study are also rather inconsistent with
the recent study by Kváč et al. (2016), in which high numbers
of cryptosporidian developmental stages were typically
associated with high oocyst shedding.
In cattle parasitised by Cryptosporidium andersoni, despite
the lack of apparent clinical signs, severe infection
was observed in the abomasum with prominent hyperplasia
of mucosa, along with a moderate degree of inflammatory
infiltration of lamina propria (Masuno et al. 2006).
The number and length of gastric pits increased considerably
because of the increasing number of epithelial cells.
Thickening and granulation of abomasal mucosa were
often reported (Anderson 1998). The epithelium of the
stomach antrum in an immunocompetent human patient
with isolated gastric cryptosporidiosis was shown to be
disorganised, fragile, and infiltrated by neutrophils (Ramsay
et al. 2007). Gastric involvement in AIDS patients is
usually considered to be secondary to retrograde spread
from the small intestine (Val-Bernal et al. 2013). Related
to Cryptosporidium gastropathy, patients suffer from
vomiting and epigastric pain. The gastric wall might exhibit
a lack of distensibility, stiffness, thickening, distortion
or erosions of the mucosal folds involving the antrum
region (Val-Bernal et al. 2013). Regularly in the same biopsy,
areas with cryptosporidia were contiguous to negative
ones. Various degrees of mucosal alterations were
observed, even in the same individual. Besides hyperplastic
reactive changes, high intensity of infection correlates
with erosions and acute inflammation. Commonly, individuals
with gastric cryptosporidiosis show no significant
endoscopic alteration to the gastric mucosa, even though
histological features are highly modified. In this and previous
studies (Melicherová et al. 2014), we also observed
a patchy (island-like) distribution of C. proliferans. Similarly
to the course of cryptosporidiosis in other homoiotherm
vertebrates, the rodents in our study did not show
any clinical signs of cryptosporidiosis.
Trends in drug development and screening studies
testing potentially anti- cryptosporidial compounds
Cryptosporidium spp. represent a highly problematic
target for drug development. One of the self-protective
strategies of cryptosporidia against the harsh conditions
of the host’s gastrointestinal tract is their unique epicellular
localisation within a parasitophorous sac of host cell
origin (Valigurová et al. 2007, 2008). The oocysts of cryptosporidia
sporulate inside the host and infective oocysts
are transmitted by the faecal-oral route. Besides the most
commonly used antibiotics and halofuginone, numerous
compounds have been screened for potential anti-cryptosporidial
activity, but the majority were ineffective.
Although some drugs have shown promise in calves and
lambs, they are too expensive (paramomycin) or highly
toxic at effective doses (halfuginone lactate and lasalocid)
(Tzipori 1998). Therefore, along with antibiotics administered
to control secondary bacterial infections, it has
been recommended to treat intestinal cryptosporidiosis in
calves with fluid therapy and the correction of acid-base
disturbances (Tzipori 1998). Colostrum containing anti-Cryptosporidium
antibodies also appears to be beneficial
(O’Donoghue 1995).
More recent studies have reported halofuginone
(Halocur) administration at the recommended dose of
100 µg/ kg for 7–10 consecutive days as very effective in
stopping diarrhoea and preventing deaths without side effects
(Giadinis et al. 2008, Petermann et al. 2014). At this
dose, it appears to inhibit the reproduction of cryptosporidia
within the host and encourages the development of immunity
in lambs (Causapé et al. 1999). Nitazoxanide, though
not effective in immunocompromised patients, significantly
shortens the duration of diarrhoea and decreases mortality
in adults and malnourished children (Gargala 2008).
Similarly, newly synthesised nitro- or non nitro- thiazolide
compounds, derived from nitazoxanide, have
been shown to be effective against Cryptosporidium parvum
(see Gargala 2008). Furthermore, compounds active
against protein disulphide isomerases (PDI2 and PDI4), the
epidermal growth factor (EGF) receptor, pp60v-src, and
pp110gag-fes, as well as new isoflavone derivatives, seem
to represent promising targets (Ortega-Pierres et al. 2009).
SCID mice orally administered with the chicken egg yolk
antibody against C. parvum infection demonstrated partial
reduction in oocyst shedding (Kobayashi et al. 2004).
Whilst the majority of these studies have been performed
on intestinal cryptosporidia, usually C. parvum,
there is still a lack of experimental work dealing with the
treatment of gastric cryptosporidiosis and most of the few
published works deal with the treatment of AIDS patients.
One of these papers has shown a positive effect of paramomycin
on reducing inflammation of the gastric mucosa
and mild relief from pain and diarrhoea, despite parasite
persistence in mucosa (Ventura et al. 1997). Another study
reported the eradication of AIDS-related gastric cryptosporidiosis
with azithromycin and suggested long-term
treatment (Dı́az Peromingo et al. 1999).
In our study, none of the Indonesian plant extracts were
shown to be effective against gastric cryptosporidiosis, despite
their proven activity against other protists parasitising
the small intestine (Pecková et al. 2018). For the first trial,
the dosage was calculated based on behavioural observation
of self-medication of wild animals (e.g. the number of
plant leaves consumed), but the dosage for the second trial
was increased to 40 mg to increase the potential antiparasitic
effect. This dosage was calculated based on maximum
concentrations of extracts reported in the literature (e.g.
Bin-Hafeez et al. 2003, Squires et al. 2011). Although additional
assays with different doses in a wide scale would
be of interest, such an extensive experiment would be too
demanding of time and material, especially in the number
of laboratory rodents required. Hence, respecting the rules
for breeding animals, regulated by Czech legislation and
the legislation of the European Commission on protection
of animals against cruelty, we have designed our experiments
so that we do not use too many laboratory rodents
unnecessarily, as do most of the world’s laboratories.
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The oral administration of Halocur – even for a prolonged
period – was not sufficiently effective either, though
it slightly decreased the degree of oocyst shedding and
seemed to facilitate the regeneration of epithelial cells lining
the gastric pits. Nevertheless, in contrast to previous reports,
it did not seem to inhibit the reproduction of cryptosporidia,
although some slight decrease might be observed
in the trendline showing oocyst excretion (Figs. 4, 5). Similarly,
halofuginone treatment did not produce a satisfactory
therapeutic outcome for infection with Cryptosporidium
serpentis Levine, 1980 affecting the gastric mucosa of
snakes (Graczyk et al. 1996), suggesting its ineffectiveness
against gastric cryptosporidiosis.
Of special interest, however, was the positive impact of
Arabidopsis thaliana on the gastric mucosa pathologically
altered by chronic cryptosporidiosis. Arabidopsis thaliana
is a small Eurasian annual flowering plant routinely used
as an important model plant in molecular biology research
and is reported to be edible (Lindh et al. 2008, Hansson et
al. 2016). The extract from the green parts (leaves) of this
plant was used as a control and we expected it to have a
neutral effect on animal health and its parasitised gastric
tissue, as we found only a few studies reporting the positive
effect of A. thaliana seed extract.
The use of plant-derived products as antimicrobial
agents has been investigated in depth. Isothiocyanates
(ITCs) are bioactive products resulting from enzymatic
hydrolysis of glucosinolates (GLs), the most abundant secondary
metabolites in the plant order Brassicales.Although
the antimicrobial activity of ITCs against foodborne and
plant pathogens has been well documented, little is known
about their antimicrobial properties against human pathogens
(Romeo et al. 2018). Concurrently, during the trial
finalisation in this study, an unexpected positive effect of
A. thaliana plant extract on the reduction of spores of microsporidian
Encephalitozoon cuniculi Levaditi, Nicolau
et Schoen, 1923 in the tissues of experimentally inoculated
BALB/c mice was documented (Mynářová 2015). Different
genetic programs, activated upon pathogen recognition
and leading to the production of inducible antimicrobial
compounds, have been identified in this plant (Tierens et
al. 2001). Using the fungus Neurospora crassa as a test
organism, Tierens et al. (2001) analysed the antimicrobial
compounds from aqueous extracts of leaves of Arabidopsis
and suggested their role in the plant’s protection
against some pathogens. The treatment of mice with Alloxan-induced
diabetes with A. thaliana at a dose of 200
mg/kg BM led to a significant reduction in blood glucose
levels and an improvement in insulin resistance (Rashid
et al. 2013, 2014; Taha et al. 2014). The consumption of
A. thaliana reversed most of the histological changes in
the liver of the diabetic mice, stimulated protein synthesis
by increasing the number of ribosomes and significantly
reduced oxidative stress in diabetes (= antioxidant effect)
(Rashid et al. 2014). Moreover, as A. thaliana plants have
a close relationship with species of Brassica eaten by humans
it is of particular interest with respect to further in-
vestigations.
Methods used for treatment efficacy evaluation:
limitations in the screening of anti-cryptosporidial
drugs.
During the course of our experiments, we found a number
of unexpected difficulties and limitations. One of these
is the need for sufficient (preferably more than estimated)
stocks of tested plant extracts for individual trials, which
can be a problem in screening studies using exotic plants
where their availability and quantity might be strictly limited
(such as those used in our study). Primary screening
of experimental protocols, therapeutic doses and the effectiveness
of selected plant extracts in various solvent media
consumes a lot of material before starting the animal experiments.
Therefore, for pilot screening of the antiparasitic
effect, it is preferable to use an in vitro system, at least
for parasites where cultivation is possible (e.g. C. parvum).
In vitro studies require smaller volumes of plant extracts
and this approach helps to minimise the number of animals
used and to reduce their distress during experiments.
Another issue was the variability of the course of infection
in tested animals inoculated at the same doses, resulting
in variations in oocyst shedding. Similarly to Sréter et
al. (1995) relatively small numbers of animals are required
for the estimation of the length of the prepatent period, but
large numbers of animals are needed for the estimation of
the mean of oocyst excretion. Additionally, histological
observations of treatment effects during early stage cryptosporidiosis
can be misleading, due to mild histopathological
changes and the patchy occurrence of the parasite.
Promising reports from studies focusing on anti-cryptosporidial
drug development are usually based on a reduction
in oocyst shedding. The results of this study, however,
indicate that the evaluation of parasitisation intensity based
exclusively on the number of oocysts shed in faeces can be
misleading. For example, despite a decline in oocyst shedding
in some treatment groups (including those administered
with Halocur), all known developmental stages of
C. proliferans, from early stages invading epithelial cells,
or freshly attached to the epithelium surface to oocysts enveloped
by a parasitophorous sac, were observed in corresponding
SEM and histological preparations. This means
that despite the fact that fewer oocysts were excreted in
host faeces, their development did not stop. Recently, it has
been shown that rats infected with Cryptosporidium occultus
Kváč, Vlnatá, Ježková, Horčičková, Konečný, Hlásková,
McEvoy et Sak, 2018 shed fewer oocysts than would
be predicted from the massive infection of the colonic epithelium
(Kváč et al. 2018). This could be explained by
the presence of two types of oocysts, i.e. thick- and thinwalled,
in life cycle of C. proliferans and other species/
genotypes (Current and Reese 1986, Uni et al. 1987, Melicherová
et al. 2014). It is likely that the treatment simply
induces increased production of thin-walled oocysts, which
excyst once they separate from host epithelium, are not
usually excreted in faeces and appear to be responsible for
autoinfection. The multiplication of cryptosporidia inside
the same host via autoinfective oocysts (= sexual stage)
appears to be beneficial for increasing parasite genetic variability,
and thereby fitness and infectivity (Melicherová et
doi: 10.14411/fp.2018.010	 Screening of potentially anti-cryptosporidial agents
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al. 2014). Alternatively, the drug-affected parasite could
begin investing in asexual multiplication and undergo multiple
rounds of merogony I to produce high numbers of invasive
merozoites. Autoinfection and the recycling of type
I merogony provide an explanation for persistent chronic
infections (Bouzid et al. 2013).
A further problem in similar studies is the correct choice
of microscopic techniques for screening for parasite presence
and morphopathological changes of the parasitised
tissue. The surface topology detectable by SEM is insufficient
for gastric cryptosporidia if not supplemented by histological
sectioning as the mucus, a thick substance naturally
produced by surface cells and cells of the gland necks
to prevent self-digestion of the gastric mucosa, might
hamper the view inside the gastric gland. This relatively
thick layer, forming a non-transparent film after chemical
fixation, is usually almost impossible to wash away, despite
the careful and repetitive rinsing of stomach tissue
(Melicherová et al. 2014). In addition, SEM analyses did
not prove helpful in evaluating parasitisation intensity of
gastric tissue, as it did not enable close examination of
constricted or only slightly dilated pits. Transmission electron
microscopy represents a powerful tool for evaluating
pathological aspects along with the presence of parasites,
but it is too expensive and time consuming for studies not
focusing on ultrastructural aspects. Hence, to more accurately
assess anti-cryptosporidial treatment efficacy in laboratory-housed
animals, the most reliable and economic
approach seems to be the monitoring of oocyst shedding
accompanied by histological (gastric tissue) or SEM analysis
(applicable for intestinal tissue) of the parasitised epithelium
post mortem. Although such an approach is not
applicable for medical purposes or for studies dealing with
livestock, experimental studies on small laboratory animals
based on the most accurate evaluation would provide important
information on the actual effect of the drug tested.
Acknowledgements. The authors would like to acknowledge
support from the Department of Botany and Zoology, Faculty
of Science, Masaryk University, towards the preparation of this
manuscript. The authors also to thank the State Ministry of Research
and Technology (RISTEK) and the Directorate General of
Forest Protection and Nature Conservation (PHKA) for their cooperation
and for their permission to conduct research in Gunung
Leuser National Park. The study was financially supported by the
ʻUMI – Saving of Pongidaeʼ Foundation project ʻParasites and
Natural Antiparasitics in the Orang-utanʼ and by the Czech Science
Foundation (Grant No. P505/11/1163), by the Ministry of
Education, Youth, and Sports, Czech Republic (Grant LO1204
from the National Program of Sustainability I). The authors
would also like to thank Mallory Abel for editing and formatting
the final draft.
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Cite this article as: Valigurová A., Pecková R., Doležal K., Sak B., Květoňová D., Kváč M., Nurcahyo W., Foitová I. 2018: Limitations
in the screening of potentially anti-cryptosporidial agents using laboratory rodents with gastric cryptosporidiosis. Folia
Parasitol. 65: 010
Received 19 September 2017 Accepted 31 May 2018 Published online 16 August 2018
7.14
Kováčiková M., Vaškovicová N., Nebesářová J.,
Valigurová A.
2018
Effect of jasplakinolide and cytochalasin D on cortical elements
involved in the gliding motility of the eugregarine
Gregarina garnhami (Apicomplexa)
European Journal of Protistology 66, 97-114
Available online at www.sciencedirect.com
ScienceDirect
European Journal of Protistology 66 (2018) 97–114
Effect of jasplakinolide and cytochalasin D on cortical elements involved in
the gliding motility of the eugregarine Gregarina garnhami (Apicomplexa)
Magdaléna Kováˇcikováa,∗, Nadˇeˇzda Vaˇskovicováb, Jana Nebesáˇrovác, Andrea Valigurováa
a
Department of Botany and Zoology, Faculty of Science, Masaryk University, Kotláˇrská 2, 611 37 Brno, Czech Republic
b
The Czech Academy of Sciences, Institute of Scientific Instruments, Královopolská 147, 612 64 Brno, Czech Republic
c
University of South Bohemia, Faculty of Science and Biology Centre of the ASCR, Institute of Parasitology, ˇCeské Budˇejovice, Czech Republic
Received 4 May 2018; received in revised form 7 August 2018; accepted 28 August 2018
Available online 31 August 2018
Abstract
Since apicomplexans represent exclusively parasitic unicellular organisms with medical and economic impacts, the principles
of their motility have been studied intensively. By contrast, the movement in apicomplexan basal groups, such as gregarines,
remains to be elucidated. The present study focuses on Gregarina garnhami parasitising the digestive tract of the locust
Schistocerca gregaria, and investigates the involvement of cytoskeletal elements (the ectoplasmic network and myonemes)
and the secretion of mucosubstances during eugregarine gliding motility. Combined microscopic analyses were used to verify
the role of actin filaments and membranes’ organisation in G. garnhami motility. A freeze-etching analysis of membranes
revealed the size, density, and arrangement of intramembranous particles along with the distribution and size of pores and ducts.
Experimental assays using actin-modifying drugs (jasplakinolide, cytochalasin D) confirmed that actin most likely plays a role
in cell motility, principally in its filamentous form (=F-actin). Myonemes, localised in the border between the ectoplasm and
endoplasm, correspond to the concentric bundles of F-actin. Microscopic analyses confirmed that changes in gamonts motility
corresponding to the changes in the organisation and density of myonemes and the ectoplasmic network in drug-treated cells,
suggesting that these structures might serve as contractile elements facilitating gliding motility in G. garnhami.
© 2018 Elsevier GmbH. All rights reserved.
Keywords: Ectoplasmic network; F-actin; Gregarine; Motility; Myonemes; Ultrastructure
Abbreviations: CLSM, confocal laser scanning microscopy; CYT D,
cytochalasin D; DMSO, dimethyl sulfoxide; EF, exoplasmic fracture face;
FE, freeze-etching; IMC, inner membrane complex; IMP(s), intramembranous
particle(s); JAS, jasplakinolide; Kp, partition coefficient; LM, light
microscopy; PBS, phosphate buffered saline; PF, protoplasmic fracture
face; SD, standard deviation; SE, standard error; SEM, scanning electron
microscopy; TEM, transmission electron microscopy; TRITC, tetramethylrhodamine
isothiocyanate.
∗Corresponding author.
E-mail address: kovacikova@sci.muni.cz (M. Kováˇciková).
Introduction
Gregarines are a highly diversified, basal lineage of
eukaryotic unicellular organisms belonging to the parasitic
group Apicomplexa. Eugregarines are widespread in marine,
freshwater, and terrestrial hosts and their development is traditionally
considered to be restricted to invertebrate hosts
(Schrével and Desportes 2015).
The gregarine pellicle comprises a plasma membrane
beneath which an inner membrane complex (IMC), consisting
of two closely apposed cortical cytomembranes,
is located. The pellicle covering the surface of intestinal
https://doi.org/10.1016/j.ejop.2018.08.006
0932-4739/© 2018 Elsevier GmbH. All rights reserved.
98 M. Kováˇciková et al. / European Journal of Protistology 66 (2018) 97–114
eugregarines creates numerous epicytic folds arranged in longitudinal
lines separated by grooves (Schrével and Desportes
2015). The organisation of these folds shows an undulating
pattern in some species (Valigurová and Koudela 2008;
Valigurová et al. 2013; Vávra and Small 1969; Vivier 1968).
The usually dilated tips of the epicytic folds comprise 12 nm
filaments and ripple dense structures, thought to have a
supportive scaffolding function or possibly representing a
component of the motility motor (Kováˇciková et al. 2017;
Schrével et al. 1983; Valigurová et al. 2013; Walker et al.
1984).
In comparison to apicomplexan invasive stages (zoites),
in which gliding motility is defined as substrate-dependent
and facilitated by an actomyosin motor associated with their
pellicle (Heintzelman 2015; Kappe et al. 2004; Keeley and
Soldati 2004; Matuschewski and Schüler 2008; Sibley et al.
1998), the exact mechanism of motility in gregarines still
remains to be elucidated (Valigurová et al. 2013). Intestinal
eugregarines are usually capable of unidirectional progressive
gliding with or without obvious changes of cell shape
(King 1981, 1988; Kováˇciková et al. 2017). The extrusion of
a mucous material left to trail behind the gliding gregarines
suggests this material to be a part of the gliding machinery
(Mackenzie and Walker 1983; Valigurová et al. 2013;
Walker et al. 1979). Ultrastructural analysis revealed diverse
cortical filamentous structures (e.g. myonemes, ectoplasmic
network), assumed to be involved in the motility and cell contraction
(Beams et al. 1959; Hildebrand 1980; Valigurová
et al. 2013; Walker et al. 1979). A more recent study on
Gregarina representatives, comprising biochemical analysis,
showed these ectoplasmic myonemes to be actin and myosin
rich (Heintzelman 2004). Both proteins were also found in
the gregarines’ cortex (cell envelope comprising the three layered
pellicle and associate cytoskeletal structures), following
the pattern of longitudinally arranged epicytic folds (Ghazali
et al. 1989; Ghazali and Schrével 1993; Heintzelman 2004;
Valigurová et al. 2013). Nevertheless, only a few experimental
studies have been performed to verify the role of actin
filaments in eugregarine gliding motility, using cytoskeletal
drugs such as jasplakinolide (inducing actin polymerisation)
and cytochalasins (blocking the association or eventual dissociation
of actin subunits) (King 1988; Valigurová et al. 2013;
Walker et al. 1979).
The present study provides a complex microscopic analysis
of the cell cortex in the eugregarine Gregarina garnhami
(Gregarinidae Labbé, 1899) parasitising the intestine of
the desert locust Schistocerca gregaria (Forskål, 1775)
(Orthoptera, Acrididae). Using the combined approaches of
light, electron and confocal microscopy, supplemented by
experimental motility assays, we focused on structures that
appear to be responsible for gliding motility in G. garnhami.
A gliding force derived from the putative actomyosin system
was already proposed by King (1988); however, the exact
mechanism involved in eugregarine motility is still poorly
characterised (Heintzelman 2004; Schrével and Desportes
2015; Valigurová et al. 2013). The main purpose of this study
was to perform experimental analyses comparable to those
published on other apicomplexan species, to evaluate newly
acquired data using different microscopic techniques and to
link these data with already known fragmentary information
about motility and related structures in G. garnhami. We have
chosen this species for experimental purposes because it represents
a model parasite from a widely available laboratory
insect.Dataobtainedonthisspeciesisthereforeverifiableand
suitable for comparison with data published on other Gregarina
spp. (e.g. Heintzelman 2004; Valigurová et al. 2013). For
the first time, gamonts of G. garnhami were experimentally
treated with jasplakinolide and cytochalasin D to observe
their effect on eugregarine survival, motility and changes in
cortical filaments of actin nature (myonemes and ectoplasmic
network). Phalloidin, in comparison to actin antibodies,
specifically binds to F-actin and provides the proof of druginduced
changes in organisation of actin filaments. This study
also deals with a drug-induced changes of epicytic folds, the
undulating pattern of which remains preserved, and with the
role of mucus in gregarine gliding. Detailed FE analysis of
gregarine cortex facilitated the visualisation and identification
of structures that are difficult to observe under TEM (e.g.
ectoplasmic network) and allowed us to distinguish between
the types of pores. Additionally, complex statistical analysis
of size/distribution of IMPs and pores’ diameters was
performed and compared with other Gregarina representatives
(personal non-published data obtained during previous
studies).
Material and Methods
Gamonts of G. garnhami were collected from the midgut
and caeca of S. gregaria (Insecta, Orthoptera). After
narcosis, decapitation, and dissection of the host, parasites
were isolated from the host intestine using Ringer’s saline
solution (0.75% [w/v] NaCl, 0.035% [w/v] KCl, 0.021%
[w/v] CaCl2; pH = 7.2). Gregarine gamonts were then transferred
to embryo dishes and carefully washed with Ringer’s
solution. The manipulation and observation of parasites
were performed using an Olympus SZX7 stereomicroscope
(Olympus).
For experimental assays, gregarines were divided among
four embryo dishes with a 30 mm cavity. Afterwards, the parasites
were treated with commercial membrane-permeable
drugs influencing the polymerisation of actin: jasplakinolide
(JAS, Invitrogen, Czech Republic) and cytochalasin D (CYT
D, Invitrogen, Czech Republic). Both drugs were reconstituted
in dimethyl sulfoxide (DMSO, Sigma-Aldrich, Czech
Republic) to prepare a 1 mM stock solution and, prior to
each experimental assay, diluted in Ringer’s saline to prepare
a working solution with a final concentration of 30 ␮M.
For controls and each experimental assay with cytoskeletal
drug, approximately 200 individuals of G. garnhami were
used. The monitoring of parasite motility was performed
hourly using a stereomicroscope. After 7 h of each experi-
M. Kováˇciková et al. / European Journal of Protistology 66 (2018) 97–114 99
mental assay, a portion of the gregarines (around 50 gamonts)
was separated for the observation of mucus shedding, while
the remaining portion was divided into thirds and fixed for
electron (SEM and TEM) and confocal microscopic analyses.
Mucus shedding was observed on living drug-treated and
control gregarines put on microscopic slides covered by a
thin layer of microbiological LB agar (LB Broth with agar,
Lennox; Sigma-Aldrich, Czech Republic) and slightly moistened
with Ringer’s solution. Observations were performed
using an Olympus BX51 microscope (Olympus) equipped
with phase contrast and an ND 25 filter.
For transmission electron microscopy (TEM), specimens
were fixed in an ice bath in 2.5% (v/v) glutaraldehyde in phosphate
buffered saline (PBS). Fixed samples were washed 3×
for20 minandpost-fixedin2%(w/v)OsO4 for2 hinthesame
buffer. After rinsing 3× for 20 min in PBS, samples were
dehydrated in acetone series and embedded in Epon (Polybed
812). Ultrathin sections were cut with diamond knives using
a Leica EM UC6 ultramicrotome (Leica Microsystems) and
stained with uranyl acetate and lead citrate. Observations
were made using a TEM-1010 (JEOL).
For scanning electron microscopy (SEM), specimens were
fixed in 2.5% (v/v) glutaraldehyde in PBS, washed 3× for
15 min and post-fixed in 2% (w/v) OsO4 for 2 h, and finally
washed 3× for 15 min in the same buffer. After dehydration
in an acetone series, parasites were critical point-dried with
CO2, coated with gold, and observed using a JSM-7401F
(JEOL).
For freeze fracture, the cell suspension was fixed overnight
in 2.77% (v/v) glutaraldehyde in 0.2 M cacodylate buffer
(pH = 7.4). Afterwards, specimens were washed for 1 h in
PBS and saturated with 20% glycerol (w) overnight in a
refrigerator for cryprotection. Prior to further processing, the
suspension was concentrated on a clock glass and the dense
pellet was placed on a gold carrier using tweezers. Agar was
used to ensure the better adhesion of the pellet to the gold
carrier. Pellet was frozen in liquid nitrogen, mounted on a
gold holder, and then processed in a BAF 060 freeze-etching
system (BAL-TEC). The temperature for manipulation of
the samples was set to −100 ◦C and the pressure inside the
chamber was 10−5 Pa. Subsequently, samples were fractured
with a microtome knife and etched for 5 min. The surfaces
of the fractured structures were coated with layers of platinum
(2.4 nm at an angle 45◦) and carbon (22.4 nm at an
angle 90◦). Afterwards, samples were removed from the
BAF 060 device and melted at room temperature. Replicas
were cleaned with 5% sodium hypochlorite and 70% sulfuric
acid, then washed in distilled water and transferred to
copper grids for examination using a Morgagni 268 D (FEI)
transmission electron microscope. Statistical evaluation of
intramembranous particles (IMP) per unit area (1 ␮m2) was
performed in ImageJ software; histograms illustrating the
IMP size distribution were prepared in Microsoft Excel. The
nomenclature follows that proposed in Branton et al. (1975)
and used in Schrével et al. (1983) and Valigurová et al.
(2013, 2017). Statistical data on gregarines from mealworms
shown in Tables 2 and 4 were obtained from replicas used
for study of Valigurová et al. (2013) and were not shown
previously.
For confocal laser scanning microscopy (CLSM), specimens
were fixed for 1 h in freshly prepared 4%
paraformaldehyde in 0.1 M PBS at room temperature and
washed 3× for 15 min in 0.1 M PBS before further processing.
Afterwards, the parasites were permeabilised in
0.5% Triton X-100 (Sigma-Aldrich, Czech Republic) for 1 h.
For the direct fluorescent staining of F-actin, samples were
incubated overnight at room temperature with phalloidintetramethylrhodamine
B isothiocyanate (phalloidin-TRITC;
Sigma-Aldrich, Czech Republic) and then washed 3× for
10 min in 0.1 M PBS. Controls were incubated using the same
protocol but without phalloidin. Preparations were mounted
in VECTASHIELD Hard Set Mounting Medium (Vector laboratories,
USA). Samples were examined under an Olympus
IX81 FVBF-2 microscope equipped with a laser-scanning
Fluo View 500 confocal unit (Fluo View 3.4 software;
Olympus) and DP70 digital camera. Fluorescence was visualised
using the TRITC (phalloidin, anti-myosin/544 nm)
laser set.
Results
Cell motility in control and drug-treated
gregarines
Gamonts of G. garnhami exhibited continuous progressive
gliding motility without obvious abrupt changes in
movement direction or speed (see Supplementary Video
S1 for a video example in the online version at DOI:
10.1016/j.ejop.2018.08.006). The average of their gliding
rate was 12.2 ␮m/s. After the application of 30 ␮M jasplakinolide
(JAS) the gamonts were able to glide until
the end of experiment (7 h) without obvious cell deformations
or gliding deceleration (see Supplementary Video
S2 for a video example in the online version at DOI:
10.1016/j.ejop.2018.08.006). The speed of movement of
JAS-treated gamonts in the first half of the experiment (up to
4 h after drug application) was even slightly higher than in
controls (15.5 ␮m/s in average) and gradually decreased to
a normal gliding rate (12.4 ␮m/s in average). After washing
the drug out and replacing it with Ringer’s saline solution,
gregarines still exhibited active forward gliding. In contrast,
gregarines treated with 30 ␮M cytochalasin D stopped
moving almost immediately after drug application (see Supplementary
Video S3 for a video example in the online
version at DOI: 10.1016/j.ejop.2018.08.006). At the end
of the experiment (7 h after drug application) and after
the careful washing out of cytochalasin D, their motility
recovered, confirming that the gregarines were alive,
but had suffered complete immobility (see Supplementary
Video S4 for a video example in the online version at
100 M. Kováˇciková et al. / European Journal of Protistology 66 (2018) 97–114
Table 1. The treatment of living gamonts of Gregarina garnhami with actin-modifying drugs.
Changes/time after drug
application
Drug (concentration)
Jasplakinolide
(30 ␮M)
Cytochalasin
D (30 ␮M)
Cytochalasin
D (10 ␮M)
Cytochalasin
D (5 ␮M)
Control in
DMSO (30 ␮M)
Initial increase of gliding speed ≥10 min (100) – – – –
Decrease of gliding speed to a
normal, regular movement
≥240 min (95) – – – –
Deceleration of gliding motility – – – ≤60 min (100) –
Complete stoppage of gliding – ≥5 min (98) ≥5 min (95) ≥100 min (90) –
Full recovery of gliding motility
in majority of gregarines after
washing in Ringer
gregarines
exhibited gliding
motility during
entire experiment
(95)
≥20 min (80) ≥20 min (80) ≥20 min (90) gregarines
exhibited active
gliding motility
during entire
experiment (95)
The symbol ‘–‘ indicates no obvious changes; ≥changes appeared after the noted time period; ≤changes appeared only during the noted time period. The
numbers in parenthesis represent the percentage of gamonts exhibiting the described motility changes.
NOTE: In control gregarines incubated with Ringer’s saline solution the movement was regular with a constant gliding speed during the entire experiment
duration.
Table 2. Summary of diameters of pores in Gregarina garnhami and three different Gregarina species.
Species TEM SEM Replicas
SP MP LP SP MP LP SP MP LP
Mean ± SE Mean ± SE Mean ± SE Mean ± SE Mean ± SE Mean ± SE Mean ± SE Mean ± SE Mean ± SE
Gregarina garnhami – 39.8 ± 1.8 129.8 ± 6.3 – – 41.9 ± 6.2 20.1 ± 0.6 47.8 ± 0.4 141.5 ± 1.9
Gregarina cuneata – 24.5 ± 8.0 48.6 ± 3.2 13.4 ± 0.4 30.2 ± 1.0 48.6 ± 3.2 – 41.8 ± 1.2 120.0 ± 6.1
Gregarina polymorpha – – 42.8 ± 3.7 – – – 15.2 ± 2.7 40.5 ± 0.6 132.8 ± 4.1
Gregarina steini – 39.6 ± 2.8 56.5 ± 3.3 14.1 ± 0.9 34.4 ± 2.2 87.5 ± 4.7 – 44.5 ± 1.1 128.9 ± 2.5
The mean diameter (nm) was calculated from measurements taken at the plasma membrane and cortical cytomembranes.
SP — small pore; MP — medium pore; LP — large pore/micropore.
SE — standard error.
− The character of the specimen did not allow the pore diameter to be reliably measured.
DOI: 10.1016/j.ejop.2018.08.006). In both cases (incubation
with JAS and cytochalasin D), the monitoring of gliding in
recovered cells for the next one hour showed that the majority
of gregarines had survived the experiment and retained
the ability to glide after the drugs had been washed out
(Table 1).
The surface topology and ultrastructural
organisation in control and drug-treated
gregarines
Gamonts were divided into an anterior protomerite and
posteriorly situated deutomerite, separated by a septum
(Fig. 1a, c, e). The cell surface of G. garnhami gamonts
was covered by a pellicle arranged into numerous, longitudinal
epicytic folds (Figs. 1 a–f, 2 a–d, 3 a–f, 4 b, d, f, 6 a,
b, e, f, 7 d–i). Individual folds clustering on several projections
distributed regularly throughout the gregarine periphery
created the so-called superfolds (Figs. 1 b, 2 a, b, d). Superficial
SEM analysis revealed the presence of mucus drops
on the apical and lateral parts of the epicytic folds, and
deeper within the grooves (Figs. 1 a, b, 2 b, c). In addition,
the site of the gregarine pellicle where the superfolds
grouped together exhibited a denser secretion and accumulation
of mucus, suggesting this part to be the gliding site
(Fig. 1a, b; see Supplementary Fig. S1 for an example in the
online version at DOI: 10.1016/j.ejop.2018.08.006). Either
the epicytic folds of gregarines treated with JAS (Fig. 1c, d)
or cytochalasin D (Fig. 1e, f) showed any significant changes
in their arrangement under SEM when compared to control
gregarines. Mucus drops were present on the surface of
individuals in both drug-treated groups (Fig. 1d, f).
Thepelliclewascomposedofthreemembranes:theplasma
membrane and the underlying inner membrane complex
(IMC; membrane system formed by a flattened alveolus)
(Figs.2e,g,6a,b).Incontrolsamples,bundlesoffilamentous
M. Kováˇciková et al. / European Journal of Protistology 66 (2018) 97–114 101
Fig. 1. General morphology of control and drug-treated Gregarina garnhami gamonts.
(a–b) The general (a) and detailed (b) view of epicytic folds and superfolds in control gregarines. Note epicytic folds grouped markedly at
one side, with the increased secretion of a dense mucosubstance in this region. SEM. (c–d) General (c) and detailed (d) view of epicytic folds
in an individuals incubated with 30 ␮m JAS for 7 h. SEM. (e–f) General (e) and detailed (f) view of epicytic folds in an individuals incubated
with 30 ␮m cytochalasin D for 7 h. SEM.
black arrow — mucus drops, dm — deutomerite, double white arrowhead — constriction in the area of septum separating the protomerite
from the deutomerite, pm — protomerite, s — superfolds, white arrow — epicytic folds.
102 M. Kováˇciková et al. / European Journal of Protistology 66 (2018) 97–114
Fig. 2. Ultrastructure and morphology of Gregarina garnhami gamonts.
(a) Transversal section of deutomerite comprising the prominent nucleus, showing the organisation of superfolds. TEM. (b–c) Superficial
view of longitudinal epicytic folds clustering together and forming the superfolds (b) and detail on their organisation at higher magnification
(c) SEM. (d) Detail of superfolds and cytoplasm comprising the bundles of myonemes. TEM. (e) The three-layered pellicle and duct-like
structures. TEM. (f) Superficial section of ectoplasm revealing the ectoplasmic filaments forming an anastomosing network. Inset shows the
network in detail. TEM. (g) Organisation of the ectoplasm, comprising the filaments corresponding to the ectoplasmic network. FE TEM.
am — amylopectin, black arrow — mucus drops, black arrowhead — medium-sized pores, bm — bundles of myonemes, double black
arrowhead — duct-like structure, ec — ectoplasm, ecn — ectoplasmic network, en — endoplasm, imc — inner membrane complex, n —
nucleus, plm — plasma membrane, s — superfolds, white arrow — epicytic folds, white arrowhead — myonemes, white asterisk — cytoplasm.
M. Kováˇciková et al. / European Journal of Protistology 66 (2018) 97–114 103
Fig. 3. Ultrastructure of cell cortex in control and drug-treated Gregarina garnhami gamonts.
(a–b) Cross section of cell cortex showing the ectoplasmic network and annular myonemes in control gamonts. TEM. (c–d) Cell cortex in
gamonts incubated with 30 ␮m JAS for 7 h. TEM. (e–f) Cell cortex in gamonts incubated with 30 ␮m cytochalasin D for 7 h. Note the absence
of annular myonemes and the fading ectoplasmic network. TEM.
am — amylopectin, black arrow — duct-like structure, black asterisk — area of usual occurrence of myonemes bundles in control gamonts, bm
— bundles of myonemes, cc — cell coat, de — dense material, double black arrowhead — micropore, ec — ectoplasm, ecn — ectoplasmic
network, en — endoplasm, white arrow — epicytic folds, white arrowhead — myonemes.
104 M. Kováˇciková et al. / European Journal of Protistology 66 (2018) 97–114
Fig. 4. Phalloidin staining of epicytic folds in control and drug-treated gamonts of Gregarina garnhami.
(a–b) Control gamonts incubated with Ringer’s saline solution. (c–d) Gamonts treated for 7 h with 30 ␮m JAS. (e–f) Gamonts treated for
7 h with 30 ␮m cytochalasin D. a, c, e — overviews; b, d, f — composite views created by flattening a series of optical sections. CLSM,
phalloidin-TRITC.
white arrow — epicytic folds.
annular myonemes (21 ± 1 nm thick) were located beneath
the pellicle at the interface between the ectoplasm and endoplasm,
and running perpendicular to the longitudinal cell axis
(Figs. 2 d, f, 3 a, b). The ectoplasmic network was situated
betweenthemyonemesandtheIMC(Fig.2e)andwasformed
by anastomosing filaments, visible in superficial ultrathin
sections (Fig. 2f). These correspond to the filamentous structures
occupying the cytoplasm beneath the epicytic folds in
replicas (Fig. 2g). Cytoskeletal structures showed differences
in drug-treated gamonts. Under the pellicle of JAS-treated
M. Kováˇciková et al. / European Journal of Protistology 66 (2018) 97–114 105
Fig. 5. Phalloidin staining of myonemes in control and drug-treated gamonts of Gregarina garnhami.
(a–b) Control gamonts incubated with Ringer’s saline solution. (c–d) Gamonts treated for 7 h with 30 ␮m JAS. (e–f) Gamonts treated for 7 h
with 30 ␮m cytochalasin D. a, c, e — composite views created by flattening all optical sections in the region of the myonemes; b, d, f —
composite views created by flattening a series of optical sections. CLSM, phalloidin-TRITC.
white arrowhead — myonemes.
gregarines, the dense ectoplasmic network occurred in the
form of filaments running in different angles to annular
myonemes (Fig. 3c, d). The density of these structures was
only slightly higher than in non-treated control parasites. In
contrast, the gregarines incubated with cytochalasin D exhibited
a different density of subpellicular structures (myonemes
andectoplasmicnetwork),whoseoccurrencewaslessevident
and they seemed to vanish (Fig. 3e, f).
106 M. Kováˇciková et al. / European Journal of Protistology 66 (2018) 97–114
Fig. 6. Architecture of epicytic folds, pores and ducts in Gregarina garnhami.
(a) Longitudinally-sectioned micropore and cross-sectioned epicytic folds. Insets show the micropores in different sections in detail. TEM.
(b) Small-sized pores occurring in cytomembranes covering the lateral sides of epicytic folds. Inset shows the small-sized pores in detail. FE
TEM. (c) Replica showing the duct and micropore. FE TEM. (d) Fracture face of IMC showing bases of the grooves between epicytic folds.
Note the presence of ducts, micropores and medium-sized pores. FE TEM. (e) Cross section of epicytic folds comprising dense material on
their lateral sides and clusters of dense material in ectoplasm. Inset shows position of dense material between the plasma membrane and IMC.
TEM. (f) Tangential section of epicytic folds with apically situated 12 nm filaments and a dense material on their lateral sides. Inset shows
apical filaments in detail. TEM.
M. Kováˇciková et al. / European Journal of Protistology 66 (2018) 97–114 107
The direct fluorescent labelling of F-actin with phalloidin
for CLSM revealed the presence of F-actin located superficially,
in longitudinal arrangement copying the pattern of
epicytic folds (Fig. 4a–f). No significant changes were monitored
between the control (incubated with Ringer’s saline
solution) and drug-treated gregarines (Fig. 4b, d, f). Second
structures visualised after phalloidin labelling were the
filaments running in circles, perpendicularly to the longitudinal
cell axis. These annular structures, localised between the
parasite pellicle (under the epicytic folds) and endoplasm,
correspond to the myonemes (Fig. 5a–f). Detailed analysis
of myonemes revealed differences in the fluorescent signals
exhibitedbycontrolanddrug-treatedgamonts.InJAS-treated
individuals,theprominentlayerofannularmyonemes,organised
in a dense network of filaments running close to each
other and linked lengthwise, exhibited very intense staining
(Fig. 5c, d). A different situation occurred in gregarines
treated with cytochalasin D. While traces of myonemes lying
in the plane between the gregarine ectoplasm and endoplasm
were still obvious, their density decreased and they seemed
to be less distinct (Fig. 5e, f).
Epicytic folds, pores and ducts in G. garnhami
Electron microscopic analyses revealed three types of
pores at the bottom of the grooves and on the lateral sides
of the epicytic folds: micropores, medium-sized pores and
small-sized pores (Fig. 6a–d). Typical apicomplexan micropores
were situated between two adjacent epicytic folds
(Figs. 3 b, 6 a, c, d). The central duct of each micropore,
interrupting the pellicle, was circumscribed by a prominent
collar and connected to a vacuolar structure located within
the cell ectoplasm (Fig. 6a, c). The irregularly distributed
micropores and medium-sized pores were clearly visible on
freeze-fractured pellicle membranes (Fig. 6d). Medium-sized
pores together with the small-sized pores were also present
in both cytomembranes on the lateral sides of the epicytic
folds, but did not reach the plasma membrane (Fig. 6b). In
ultrathin sections, the medium-sized pores were noticeable as
interruptions of the IMC (Figs. 2 e, 7 d). Measurements of the
pore diameters in G. garnhami and another three Gregarina
species(takenfromourspecimens)usingseveralmicroscopic
techniques are shown in Table 2.
Duct-like structures of wavy appearance and terminating
at the bottom of the grooves between adjacent folds
were detected in the ectoplasm (Figs. 2 e, 6 c). The openings
of these ducts in the external cytomembrane formed
Table 3. The sizes of IMP in individual fracture faces of pellicle
membranes in Gregarina garnhami.
Membrane Face Size of IMP (nm)
Mean Median SD SE Min Max
Plasma
membrane
PF 4.5 3.9 2.7 0.1 0.5 15.2
EF 4.0 3.7 2.1 0.1 0.5 15.0
External
cytomembrane
PF 4.4 4.0 2.3 0.1 0.5 16.1
EF 3.6 3.2 1.8 0.1 0.5 12.8
Internal
cytomembrane
PF 4.9 4.1 3.0 0.1 0.5 17.4
EF 2.8 2.6 1.4 0.1 0.5 9.1
SD — standard deviation; SE — standard error.
extensive depressions (Fig. 6d). The cross-fractured termination
of these duct-like structures in exoplasmic fracture face
(EF) of the external cytomembrane measured 43.4 ± 1.4 nm
in diameter, while in longitudinally fractured ducts it
was 54.9 ± 2.5 nm in diameter. The interconnection of the
ductus with the internal cortical cytomembrane measured
78.3 ± 2.4 nm in diameter (measured in replicas). In several
ultrathin sections it seemed that the duct-like structures
crossed the pellicle and opened to the external environment.
Ducts were connected with vesicles located in the ectoplasm.
In addition, clusters of a dense material in gregarine ectoplasm
and filling the space between the plasma membrane
and IMC on the lateral part of the epicytic folds and between
two adjacent folds were observed (Figs. 3 a, b, 6 e, f).
In the apical part of the epicytic folds, linear IMP
arrays were observed in EF of both cortical cytomembranes
corresponding with their localisation to the 12 nm
filaments observed under TEM (Fig. 6a, e, f). Analysis of
the supramolecular organisation of the plasma membrane
in G. garnhami showed similar sizes of intramembranous
particle(s) (IMPs) in the protoplasmic (PF) and in the exoplasmic
(EF) fracture faces. The size of IMPs varied from 0.5
to 17.4 nm, depending on the membrane type and its fractured
face(Table3,seeSupplementaryFig.S2foranexampleinthe
online version at DOI: 10.1016/j.ejop.2018.08.006). Unlike
the plasma membrane, in cortical cytomembranes the density
of IMPs was higher in the EF than in the PF of fractured
faces (Table 4). Only particles ranging from 6 to 14 nm were
used for statistical calculations of Kp (partition coefficient) in
order to obtain data comparable with those so far published on
other apicomplexans (Table 5). The statistical values differed
considerably when including all visible IMPs. Therefore, the
values listed in Table 3 include the total number of IMPs in the
black arrow — small-sized pores, black arrowhead — medium-sized pores, black asterisk — mucosubstance, c — micropore collar, cc —
cell coat, d — micropore duct, de — dense material, double black arrowhead — micropore, double white arrowhead — duct-like structure, ee
— EF of the external cytomembrane, ei — EF of the internal cytomembrane, ep — EF of the plasma membrane, fil — 12 nm apical filaments,
imc — inner membrane complex, pe — PF of the external cytomembrane, plm — plasma membrane, pp − PF of the plasma membrane, v −
vesicle, white arrow — epicytic folds, white arrowhead — myonemes, white asterisk — cytoplasm, white circle — termination of duct-like
structure.
108 M. Kováˇciková et al. / European Journal of Protistology 66 (2018) 97–114
Fig. 7. Secretion in control and drug-treated gamonts of Gregarina garnhami.
(a–c) Mucous trail left behind gliding gregarine. a − Ringer’s saline solution, b − 30 ␮m JAS, c − 30 ␮m cytochalasin D. LM, phase
contrast. (d) Dense material observed between the epicytic folds. Note the medium-sized pores and a prominent layer of cell coat on the
gregarine surface. Inset shows a dense material passing through the IMC. TEM. (e) Secretion of mucous material. FE TEM. (f) Mucus drops
observed between the epicytic folds. SEM. (g) A view of ectoplasm and epicytic folds in a gregarine treated with 30 ␮m cytochalasin D. Note
the presence of lipid-like droplets in ectoplasm. Inset shows the duct connecting to the lipid-like droplet. TEM. (h-i) Putative secretion of
electron-lucent vesicles throughout the pore-like structures. h — 30 ␮m DMSO, i — 30 ␮m JAS. TEM.
black arrow — mucus trail, black arrowhead — medium-sized pores, cc — cell coat, double black arrowhead — termination of pore-like
structure, ld — lipid-like droplets, white arrow — epicytic folds, white arrowhead — mucus, white asterisk — cytoplasm.
M. Kováˇciková et al. / European Journal of Protistology 66 (2018) 97–114 109
Table4.Summaryofdensity(particles/␮m2
)andsizesofIMP(nm)inGregarinagarnhamiandthreeotherGregarinaspecies.
SpeciesPlasmamembraneExternalcytomembraneInternalcytomembrane
EFPFPFEFEFPF
Density
ofIMP
Sizeof
IMP
Density
ofIMP
Sizeof
IMP
Density
ofIMP
Sizeof
IMP
Density
ofIMP
Sizeof
IMP
DensityofIMPSizeof
IMP
Density
ofIMP
Sizeof
IMP
Mean±SEMin–MaxMean±SEMin–MaxMean±SEMin–MaxMean±SEMin–MaxMean±SEMin–MaxMean±SEMin–Max
Gregarinagarnhami6115±6660.5–157138±8510.5–15.26741±5430.5–16.19832±9090.5–12.89273±6070.5–9.17365±4390.5–17.4
Gregarinacuneata2950±1563.3–27.03621±2743.7–20.62633±991.1–17.42681±852.2–16.48297±4901.5–12.95991±4473.7–19.1
Gregarinapolymorpha3278±423.2–11.75294±2252.98–11.61731±1583.7–13.73477±3472.1–10.51884±992.7–9.72652±862.7–12.7
Gregarinasteini2485±992.1–15.12531±1272.1–14.13268±2022.2–14.24170±372.1–14.43368±2301.7–13.33033±952.1–9.8
ThetotalnumberofallsizesofIMPinamembranefracturewasusedforcalculation.
SE—standarderror.
fracture face of individual pellicle membranes as well as the
maximum and minimum size of IMPs in specimens that were
used in our previous studies. The actual IMP size distribution
in each fractured membrane is shown in histograms (see
Supplementary Fig. S2 for an example in the online version
at DOI: 10.1016/j.ejop.2018.08.006).
Mucus secretion in control and drug-treated
gregarines
Following treatment with cytoskeletal drugs, the gliding
of gregarines on agar was examined to confirm the presence
of secreted mucus trails. Clearly evident long and regular
mucous paths were left behind gliding gamonts in all experimental
groups (Fig. 7a–c); however, the paths of drug-treated
gregarines were less noticeable (Fig. 7b, c).
Ultrastructural observations showed differences in cytoplasm
organisation between the drug-treated and control
gregarines. Control gregarines incubated with Ringer’s saline
solution exhibited a higher accumulation of dense granules in
ectoplasm (Figs. 2 e, 3 a, b, 6 e). This dense material seemed
to pass throughout both cortical cytomembranes to the space
between the plasma membrane and IMC and reach the external
environment via exocytosis (Figs. 6 e, f, 7 d). Presumably,
the content of the abovementioned dense granules is secreted
on the gregarine surface and forms the cell coat (Figs.6 a, 7 d).
Secreted globular or conical structures, situated in-between
and on the top of the epicytic folds, and observed in our
SEM and EF TEM preparations, correspond to mucus drops
(Figs. 1a, b, d, f, 2 b, c, 7 e, f). In contrast, the ectoplasm
of gregarines incubated with chemicals (pure DMSO, JAS,
and cytochalasin D, the latter two both diluted in DMSO)
exhibited a less pronounced concentration of dense granules,
while the more prominent presence of lipid-like droplets was
observed (Fig. 7g). Ducts were observed to connect cytoplasmic
vesicles containing this lipid-like substance; some
vesicles were already partially empty (Fig. 7g inset). The
putative secretion of vesicles comprising an electron-lucent
material was observed throughout the pore-like structures
(similar to micropores but lacking a collar; 35–50 nm in diameter)
located between two adjacent epicytic folds (Fig. 7h,
i). While a prominent filamentous cell coat was observed
in control gregarines, the drug-treated cells showed only an
inconspicuous glycocalyx layer (Figs. 3 b, 6 a, 7 d vs. 3 c–f,
7 g–i).
Discussion
The supramolecular organisation of the pellicle
in Gregarina representatives
Gamonts of G. garnhami analysed in this and a previous
study (Valigurová and Koudela 2008) exhibited a pellicle
of typical eugregarine organisation, consisting of the
110 M. Kováˇciková et al. / European Journal of Protistology 66 (2018) 97–114
Table 5. Density of IMP (particles/␮m2
) in different apicomplexan species.
Species
Plasma membrane External cytomembrane Internal cytomembrane
EF PF Kp PF EF Kp EF PF Kp
Gregarina garnhami 3095 ± 242 4219 ± 249 1.4 3468 ± 213 1572 ± 247 2.2 2106 ± 212 3822 ± 241 1.8
Gregarina blaberaea
977 ± 235 1469 ± 233 1.5 285 ± 39 133 ± 34 2.1 158 ± 72 297 ± 33 1.9
Gregarina cuneata 2770 ± 96 2244 ± 283 0.8 1420 ± 190 1260 ± 211 1.1 1502 ± 273 1993 ± 253 1.3
Gregarina polymorpha 2473 ± 147 1446 ± 158 0.6 602 ± 265 863 ± 202 0.7 814 ± 246 1276 ± 200 1.6
Gregarina steini 1783 ± 233 2265 ± 154 1.3 2588 ± 189 3820 ± 211 0.7 1886 ± 274 2339 ± 132 1.2
Eimeria nieschulzib
218 ± 21 648 ± 73 3.0 2360 ± 133 29 ± 7 81.4 146 ± 31 1780 ± 97 12.2
Plasmodium knowlesic
185 ± 25 2198 ± 528 11.9 1751 ± 228 38 ± 15 46.1 48 ± 28 574 ± 200 12.0
Siedleckia nematoidesd
183 ± 8 2926 ± 135 16.0 2745 ± 220 458 ± 15 6.0 797 ± 60 3342 ± 128 4.2
The size of IMP is in range 6–14 nm.
Kp partition coefficient defined as number of particles per ␮m2 in the PF face/number of particles per ␮m2 in the EF face.
aValues taken from Schrével et al. (1983).
bValues taken from Dubremetz and Torpier (1978).
cValues taken from McLaren et al. (1979).
dValues taken from Valigurová et al. (2017).
plasma membrane and IMC, folded into numerous, longitudinal
epicytic folds (Desportes and Schrével 2013; Schrével
et al. 1983; Valigurová et al. 2013; Vávra and Small 1969;
Walker et al. 1984). The values of Kp coefficient (represents
an objective comparative factor characterising the membranes
skeleton) in the plasma membrane and both cortical
cytomembranes of G. garnhami were comparable with those
of all investigated species of the genus Gregarina Dufour,
1828 (Schrével et al. 1983; Valigurová et al. 2013). Nevertheless,
it needs to be emphasized that the IMP density is
apparentlyhighestinG.garnhami comparedtootherapicomplexans
investigated thus far (Table 5). Variability of IMPs
(mostly the presence of small particles in all membranes)
in G. garnhami significantly differs from other Gregarina
representatives (Valigurová et al. 2013). Because the nature
of documented IMPs remains unknown, we cannot speculate
about their potential involvement in gregarine gliding
motility.
Pores, ducts and mucus secretion in Gregarina
representatives
In agreement with previous studies (Valigurová and
Koudela 2008; Walker et al. 1984), we observed typical
micropores consisting of a central duct lined by a dense
collar. In general, the apicomplexan micropores (also called
“cytostomes”) were thought to have a role in the feeding
of zoites by ingesting host cell cytoplasm via the formation
of pinocytic vesicles (Aikawa 1966; Nichols et al. 1994;
Scholtyseck and Mehlhorn 1970). A nutritive function has
also been attributed to micropores in gregarines detached
from host tissue (Desportes and Schrével 2013; Warner
1968). In addition, the freeze-fractured IMC in eugregarines
showed the presence of medium-sized pores situated in both
cortical cytomembranes on the lateral sides of epicytic folds
and in the grooves between the folds (Walker et al. 1984;
this study). Moreover, this study revealed the presence of
small-sized pores (with diameters more than 50% smaller
compared to medium-sized pores; Table 2) occupying the
lateral sides of epicytic folds, which were not described in
freeze-etching study performed by Walker et al. (1984). To
compare the sizes of micropores and different pores in several
apicomplexan species (Valigurová et al. 2013, 2017), measurements
were taken using different microscopic techniques
(Table 2).
Duct-like structures of wavy appearance detected under
the pellicle were seen to originate between two adjacent
folds and pass inside the cytoplasm. Ectoplasmic ducts or
channel-like structures previously observed in gregarines
from mealworms and in G. garnhami (Valigurová et al. 2013;
Walker et al. 1984) were postulated to pass to the exterior
between the folds and to have a role in mucus secretion
(Walker et al. 1984). In G. cuneata, some of the ducts were
connected to the pore-like structures in the IMC (Valigurová
2012). In the present study, ultrathin sections did not reveal
secretion of mucus on the gregarine surface throughout the
duct-like structures, but the ducts were connected to cytoplasmic
vesicles. In drug-treated gregarines, these cytoplasmic
vesicles were filled with lipid-like content, suggesting their
role in metabolic processes. In general, lipid droplets in apicomplexans
play roles in lipid metabolism, cell signalling,
and intracellular vesicle trafficking (Sonda and Hehl 2006).
Ultrathin sectioning also revealed the presence of unknown
pore-like structures (similar to micropores but without a
collar). The drug-treated gregarines exhibited an intensive
secretion of vesicles comprising electron-lucent material
throughout these pores, pointing to their role in the excretion
of waste products or toxic material out of the cell. We also
hypothesize their possible involvement in the transportation
of material necessary for the restoration of natural conditions
in the environment surrounding the gregarine.
The presence of mucus drops on the top of, and between the
epicyticfoldsofG.garnhamiwasconfirmedbySEManalysis
M. Kováˇciková et al. / European Journal of Protistology 66 (2018) 97–114 111
(Valigurová and Koudela 2008; this study). In eugregarines,
the mucus trail left behind gliding gamonts was previously
investigated as a potential lubricant for gliding locomotion
(Schwiakoff1894;Valigurováetal.2013;Walkeretal.1979).
Although mucus flow accompanies the gliding, there is no
evidence to show whether it is the cause or a consequence
of gliding movement (Mackenzie and Walker 1983). In our
research, the presence of mucus trails left behind gliding
gamonts in control and both experimentally affected groups
was monitored. Importantly, the mucus trails in drug-treated
gamonts indicate this substance to have a supportive function,
maintaining an environment suitable for gregarine movement
onasolidsurface.Itcanbeexpectedthatthecellcoatcovering
theentiresurfaceofthegregarineiscontinuouslyreformedby
the secretion of its components from the cell. In G. garnhami,
a dense material occupying the space between the plasma
membrane and IMC on the lateral sides of epicytic folds and
in-between the epicytic folds, appears to be transported from
gregarine cytoplasm to the intramembranous space through
the cytomembranes. We hypothesize that this dense matter is
subsequently secreted by exocytosis to the cell surface and
represents the core component for the formation of the cell
coat. Similarly, dense inclusions localised mainly in the ectoplasm
of G. blaberae (Schrével 1972) were suggested to be
associated with the dense filamentous cell coat rich in glycoconjugates
and covering the plasma membrane (Philippe
et al. 1979; Schrével 1972; Schrével et al. 1983).
Cytoskeletal organisation in Gregarina
representatives
The actomyosin motor in apicomplexan zoites (considered
to be the key component of the parasites’ motor responsible
for gliding and host invasion) is assumed to be embedded
between the plasma membrane and the IMC and connected to
transmembrane adhesin complexes contacting the substrate
(Opitz and Soldati 2002). In gregarines, the actomyosin
system was proposed as one of the potential mediators
causing lateral undulations of epicytic folds and, in this
way, contributing to the gliding movement (Desportes and
Schrével 2013). In our SEM analysis of drug-treated gamonts
with ceased motility, however, the reorganisation (undulation
or striking of epicytic folds) was not substantiated. The rows
of IMPs in both cytomembranes occurring in the apical parts
of epicytic folds correspond to the position of the 12-nm
filaments. Potential connections between IMPs anchoring
these 12 nm filaments to the IMC at the tips of epicytic
folds were detected in G. garnhami previously (Walker
et al. 1984). The interaction between these structures might
facilitate gregarine gliding (Dallai and Talluri 1983; Walker
et al. 1984), nevertheless, their real functions is still poorly
understood. It has been shown that the number of 12-nm
filaments, exhibiting the properties of intermediate filaments,
does not influence the gregarine gliding speed, but rather
seems to control the direction of movement (Valigurová
et al. 2013). However, while the number of detected 12-nm
filaments was the same in G. garnhami and in G. cuneata
(=7), the gliding of gamonts in latter one was characterised
by multiple and prolonged stops and changes of direction,
i.e. linear vs. semi-circular gliding path (Valigurová et al.
2013). The frequent changes of gliding path in G. cuneata
appear to be additionally influenced by the bending movements
of its protomerite (Valigurová et al. 2013). Majority
of eugregarines investigated in our long-term research
(Kováˇciková et al. 2017; Valigurová et al. 2013, and personal
nonpublished data) exhibited more or less discontinuous
gliding and we cannot provide any explanation of the occurrence
in the stops in G. cuneata gliding. Considering the fact
that all these observations were performed using parasites
isolated from their host, it could simply represent a reaction
of various intensity (differing among species) to in vitro
conditions, or different sensitivity of gregarine species to the
quality of the substrate and barriers in their gliding path.
For a deeper understanding of the actomyosin system in
Gregarina spp., several biochemical and molecular investigations
were performed (Baines and King 1989; Ghazali
et al. 1989; Ghazali and Schrével 1993; Heintzelman
2004; Heintzelman and Mateer 2008; Philippe et al. 1982;
Valigurová 2012; Valigurová et al. 2013). As observed in
other terrestrial eugregarines, the cytoplasm of G. garnhami
is divided into inner finely granular endoplasm and outer
ectoplasm (Canning 1956), the latter lying under the pellicle
and possessing cytoskeletal filamentous structures. Crawley
(1905) described a layer of fibrils encircling the gregarine and
joined together by connectives forming a network, which he
called the myocyte. The myocyte is supposed to be formed
by contractile elements that are responsible for gregarine
motility and bending. In more recent publications (Beams
et al. 1959; Hildebrand 1980; Valigurová and Koudela 2008;
Valigurová et al. 2013; Walker et al. 1979) these filaments
in eugregarines were divided into two types depending on
their thickness and position relative to the gregarine axis.
The ectoplasmic network underlies the inner cytomembrane,
while deeper in the edge of the ectoplasm and endoplasm,
annular rib-like myonemes are localised. In Gregarina representatives,
the myonemes and ectoplasmic network were
suggested to be the major contractile elements providing the
driving force for cell locomotion and/or protomerite bending
(Beams et al. 1959; Valigurová et al. 2013). The staining of
actin for indirect immunofluorescence showed it to be confined
to the cortical region, lacking a fibrillar pattern (Baines
and King 1989; Valigurová et al. 2013). Surprisingly, contradicting
the observations on G. cuneata (Valigurová et al.
2013), the presence of actin stained with antibody was not
observed in ghosts (=cell cortex) of G. garnhami (Mackenzie
1980). A study, in which actin was specifically labelled with
antibody generated to G. polymorpha actin, revealed its bilaminar
staining pattern (Heintzelman 2004). The outer layer
of actin was organised into longitudinal lines, copying the
arrangement of epicytic folds, while the inner layer corresponded
to rib-like myonemes oriented perpendicular to the
longitudinal cell axis (Heintzelman 2004).
112 M. Kováˇciková et al. / European Journal of Protistology 66 (2018) 97–114
In G. garnhami, a dense layer of annular myonemes
running perpendicular to the cell axis along with a network
of ectoplasmic filaments were also observed under electron
and confocal microscopy. Although Walker et al. (1979)
described in G. garnhami circularly arranged microtubules
in the transition area between the ectoplasm and endoplasm,
these structures most likely correspond to the myonemes
observed in the present study, since the presence of microtubules
in G. garnhami by the negative staining of ␣-tubulin
for immunofluorescent microscopy was not monitored
(data not shown). The study on Gregarina species from
mealworms came to the same conclusion (Valigurová et al.
2013). Though several biochemical analyses of Gregarina
representatives confirmed the actin nature of myonemes, the
form of present actin (monomeric vs. polymerised into filaments)
was not determined. In gregarines from mealworms,
the F-actin accumulated in the apical end of the protomerite
and in the area of the membrane fusion site (i.e. osmiophilic
ring); in addition, it was restricted to the cell cortex, fibrillar
septum, and nucleus; however, the recognition of individual
annular myonemes was not possible (Valigurová et al. 2009,
2013). In our study, labelling of F-actin with fluorescent
phalloidin, revealed numerous subpellicular actin filaments
running perpendicularly to the cell axis, which correspond
in their organisation and localisation to myonemes detected
in ultrathin sections. According to these observations,
myonemes are most likely formed by the bundles of actin
filaments and our experiments showed them to be essential
in G. garnhami gliding motility. The association of actin filaments
with different types of myosins (A, B and F) appears to
be likely, because these myosins were immunoflurescently
detected in other Gregarina spp. in similar localisation
(restricted to the epicytic folds and annular myonemes)
and organisation pattern (Heintzelman 2004; Heintzelman
and Mateer 2008) as F-actin in G. garnhami. Actomyosin
motor was proposed to participate in gliding motility, as
well as myonemes-mediated bending of G. polymorpha
(Heintzelman 2004; Heintzelman and Mateer 2008).
Effect of actin-modifying drugs on motility and
cytoskeleton of Gregarina representatives
Experimental studies dealing with drugs influencing the
actin polymerisation, namely jasplakinolide and cytochalasins,
demonstrated their significant impact on eugregarine
motility (King 1988; Valigurová et al. 2013; Walker et al.
1979). Jasplakinolide (JAS) is known to stabilise actin filaments
and induce the polymerisation of monomeric actin
(Bubb 2000). Concentrations of JAS ranging from 5 ␮M to
30 ␮M were used in motility experiments on eugregarines
from mealworms (Valigurová et al. 2013). These eugregarines
were able actively to glide for up to 150 min after the
beginning of their incubation with JAS, and their movement
recovered fully after the drug was washed out (Valigurová
et al. 2013). Experiments with G. garnhami showed even
greater tolerance of individuals to high concentrations of this
drug. After 7 h (420 min) of incubation with 30 ␮M JAS,
gamonts were still able to glide and no superficial changes or
abnormalities in their motility were noticed. We attribute the
weak effect of JAS on G. garnhami gamonts to the assumption
that the majority of actin in this species is already present
in its polymerised form (even before incubation with JAS as
an actin stabilising drug). This hypothesis is further supported
by our CLSM data. The presence and density of myonemes
in ultrathin sections after treatment with JAS was concordat
with CLSM analysis.
An opposite effect on motility was documented in gregarines
treated with cytochalasin D. Cytochalasins are fungal
metabolites binding to actin filaments, which inhibit the association
and dissociation of subunits at the barbed end (Cooper
1987). A previous study performed on G. garnhami described
the irreversible cessation of gliding motility induced by 0.1%
cytochalasin B, with no recovery after completion of the
experiment (Walker et al. 1979). However, it is important
to note that cytochalasin B binds to both actin filaments
and glucose transporters. In contrast, cytochalasin D has a
high affinity to actin filaments and specifically binds to them
only, thereby reducing the probability that motility could be
blocked by another mechanism (Cooper 1987). Our experiments
on G. garnhami resulted in the rapid cessation of
motility almost immediately after the application of 30 ␮M
cytochalasin D. Subsequent microscopic analyses demonstrated
less dense arrangements of ectoplasmic network and
myonemes with fading connections. The gregarines were
treated for 7 h, during which they showed no sign of gliding;
however, after washing with Ringer’s saline solution,
the recovery of gliding motility was achieved in the majority
of individuals. Lower cytochalasin D concentrations of 5
and 10 ␮M were also applied in order to compare the effects
of various drug concentrations on G. garnhami motility, but
no significant differences were observed (Table 1). In eugregarines
from mealworms, the highest applied concentration
of cytochalasin D (30 ␮M) blocked their motility from 10 to
75 min after beginning of experiment, and full recovery was
observed 10 min after washing in Ringer’s saline solution
(Valigurová et al. 2013). It can be assumed that cytochalasin
D did not lead to the mortality of gregarines (a fact demonstrated
by their recovery), despite the fact that their motility
was completely blocked. Further investigation is needed to
specify the exact mechanism of this drug on actin filaments
and their role in gregarine motility.
Conclusions
The present study revealed that the wavy pattern of epicytic
folds in G. garnhami is similar in treated and non-treated
gregarines, even in gregarines incubated with cytochalasin
D, where the gliding motility was completely blocked for a
long period (Figs. 1 a, b, 2 b, c vs. Fig. 1c–f). This observation
contradicts the expectation that the lateral undulation
of epicytic folds provides the force behind gregarine glid-
M. Kováˇciková et al. / European Journal of Protistology 66 (2018) 97–114 113
ing, an idea previously proposed by other authors (Schrével
and Philippe 1993; Vávra and Small 1969; Vivier 1968). In
addition, our results on drug-treated gamonts demonstrate
the importance of subpellicular structures such as the ectoplasmic
network and myonemes in G. garnhami motility.
Changes in gliding motility, i.e. cessation of movement in
cytochalasin D-treated gamonts, were accompanied with partial
degradation of myonemes and their fading in CLSM
micrographs. After the treatment with JAS, the changes
in gliding motility were inconspicuous despite denser network
of myonemes was observed after phalloidin labelling.
Based on microscopic observations we conclude that annular
myonemes, occurring at the border between the ectoplasm
and endoplasm, consist of bundles of actin filaments and that
the majority of actin is present in polymerised form. The
organisation of myonemes and ectoplasmic network changed
during the experiments with actin-modifying drugs in accordance
with gamonts gliding activity. The actin filaments
most likely represent contractile elements facilitating gliding
motility in G. garnhami gamonts, while mucus secretion
has only supportive function. In addition, our results suggest
that the dynamic process of actin polymerisation and subsequent
rapid depolymerisation, proposed for apicomplexan
zoites using the so-called “glideosome” system to move over
surfaces (Opitz and Soldati 2002), is not essential for gliding
motility in studied eugregarines. In contrast, only the polymerised
form of actin seems to be the main leading motor
structure responsible for gliding movement in gamonts of G.
garnhami and other Gregarina representatives. This conclusion
is supported by the fact that the incubation with JAS,
inducing further stabilisation of F-actin already present in
treated gamonts, did not significantly change their motility.
In contrast, treatment with cytochalasin D almost immediately
blocked eugregarines motility due to depolymerisation
of existing actin filaments.
Acknowledgements
Authors acknowledge support from the Department of
Botany and Zoology of the Faculty of Science at Masaryk
University towards the preparation of this manuscript. Financial
support was provided by MEYS CR (LO1212) and
EC (CZ.1.05/2.1.00/01.0017). The Laboratory of Electron
Microscopy, Biology Centre CAS was supported by the
Czech-BioImaging large RI project (LM2015062 funded by
MEYS CR). MK, AV and NV were also supported by the
project 7AMB14FR013 from the Ministry of Education,
Youth and Sports of the Czech Republic.
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7.15
Simdyanov T.G., Paskerova G.G., Valigurová A., Diakin A.,
Kováčiková M., Schrével J., Guillou L., Dobrovolskij A.A., Aleoshin V.V.
2018
First ultrastructural and molecular phylogenetic evidence from
the blastogregarines, an early branching lineage of plesiomorphic
Apicomplexa
Protist 169(5), 697-726
Protist, Vol. 169, 697–726, November 2018
http://www.elsevier.de/protis
Published online date 23 April 2018
ORIGINAL PAPER
First Ultrastructural and Molecular
Phylogenetic Evidence from the
Blastogregarines, an Early Branching
Lineage of Plesiomorphic Apicomplexa
Timur G. Simdyanova,1
, Gita G. Paskerovab
, Andrea Valigurovác
, Andrei Diakinc
,
Magdaléna Kováˇcikovác
, Joseph Schréveld,e
, Laure Guillouf
,
Andrej A. Dobrovolskijb
, and Vladimir V. Aleoshing,h
aFaculty of Biology, Lomonosov Moscow State University, Leninskiye Gory 1-12, 119 234
Moscow, Russian Federation
bDepartment of Invertebrate Zoology, Faculty of Biology, Saint Petersburg State University,
Universitetskaya emb. 7/9, 199 034 St. Petersburg, Russian Federation
cDepartment of Botany and Zoology, Faculty of Science, Masaryk University, Kotláˇrská 2,
611 37 Brno, Czech Republic
dCNRS 7245, Molécules de Communication et Adaptation Moléculaire (MCAM), CP 52,
rue Cuvier, 75005 Paris, France
eSorbonne Universités, Muséum National d’Histoire Naturelle (MNHN), UMR 7245, CP 52,
rue Cuvier, 75005 Paris, France
fSorbonne Université, Université Pierre et Marie Curie—Paris 6, CNRS, UMR 7144,
Station Biologique de Roscoff, Place Georges Teissier, CS90074, 29688 Roscoff Cedex,
France
gBelozersky Institute of Physico-Chemical Biology, Lomonosov Moscow State University,
Leninskiye Gory 1-40, 119 234 Moscow, Russian Federation
hInstitute for Information Transmission Problems, Russian Academy of Sciences, Bolshoy
Karetny per. 19-1, 127 051 Moscow, Russian Federation
Submitted March 16, 2018; Accepted April 13, 2018
Monitoring Editor: Frank Seeber
Blastogregarines are poorly studied parasites of polychaetes superficially resembling gregarines, but
lacking syzygy and gametocyst stages in the life cycle. Furthermore, their permanent multinuclearity
and gametogenesis by means of budding considerably distinguish them from other parasitic Apicomplexa
such as coccidians and hematozoans. The affiliation of blastogregarines has been uncertain:
different authors considered them highly modified gregarines, an intermediate apicomplexan lineage
between gregarines and coccidians, or an isolated group of eukaryotes altogether. Here, we report
the ultrastructure of two blastogregarine species, Siedleckia nematoides and Chattonaria mesnili, and
provide the first molecular data on their phylogeny based on SSU, 5.8S, and LSU rDNA sequences.
1
Corresponding author;
e-mail simdyanov@mail.bio.msu.ru (T.G. Simdyanov).
https://doi.org/10.1016/j.protis.2018.04.006
1434-4610/© 2018 Elsevier GmbH. All rights reserved.
698 T.G. Simdyanov et al.
Morphological analysis reveals that blastogregarines possess both gregarine and coccidian features.
Several traits shared with archigregarines likely represent the ancestral states of the corresponding
cell structures for parasitic apicomplexans: a distinctive tegument structure and myzocytotic feeding
with a well-developed apical complex. Unlike gregarines but similar to coccidians however, the nuclei of
male blastogregarine gametes are associated with two kinetosomes. Molecular phylogenetic analyses
reveal that blastogregarines are an independent, early diverging lineage of apicomplexans. Overall,
the morphological and molecular evidence congruently suggests that blastogregarines represent a
separate class of Apicomplexa.
© 2018 Elsevier GmbH. All rights reserved.
Key words: Apicomplexa; blastogregarines; ultrastructure; plesiomorphic traits; molecular phylogeny; 18S and
28S ribosomal DNAs.
Introduction
The name Apicomplexa was first introduced by
Levine (Levine 1970) in order to unite “genuine”
Sporozoa (gregarines, coccidians, and haemosporidians)
and Piroplasmida relying on ultrastructural
characters because the life cycles of
the latter were poorly studied at that time and
any molecular phylogenetic evidence was absent.
Through the years, the composition of the group
changed: in 1980, Levine moved piroplasms into
Sporozoa, but simultaneously expanded the taxon
(phylum) Apicomplexa, which comprised two subphyla
that time: Perkinsezoa (with Perkinsus) and
Sporozoa. Later, Perkinsezoa were consistently
removed from Apicomplexa (e.g., Perkins et al.
2000), especially when Perkinsus was revealed
to be an earliest branch of Dinozoa (Goggin and
Barker 1993; Kuvardina et al. 2002), so that “Apicomplexa”
was eventually reduced to “Sporozoa”
and successively substituted this name, despite
the fact that it was conventional during many
decades before (e.g., Grassé 1953a,b). Thus,
recently Apicomplexa is a junior synonym of
Sporozoa in terms of the International Code of
Zoological Nomenclature and consequently should
be abolished. However, we support the original
approach of Levine to combine Sporozoa and
their closest relatives in a single taxon, and therefore,
also following some recent viewpoints (e.g.,
Cavalier-Smith 2014; Vot´ypka et al. 2016), we
consider Apicomplexa in this paper as a large
phylogenetic clade comprising Chrompodellida (or
Apicomonada in Cavalier-Smith’s terminology) and
Sporozoa (or Sporozoasida in Levine’s terminology).
Chrompodellids include free-living predatory
flagellates (colpodellids) and symbiotic photosynthetic
organisms (chromerids) closely related to
each other (Janouˇskovec et al. 2015), whereas
sporozoans are obligate parasites: many of them
are pathogens of humans and domestic animals
causing serious diseases (Perkins et al. 2000).
The major sporozoan groups of high practical
importance are coccidians (e.g., Toxoplasma and
Eimeria), cryptosporidians, haemosporidians (e.g.,
Plasmodium causing malaria), and piroplasms
(e.g., Babesia), therefore these organisms are popular
subjects of scientific research while gregarines
and other early branching invertebrate parasites
remain understudied.
Phylogenetic relationships and evolutionary history
of sporozoans within Apicomplexa are still an
open question. The primary divergence of sporozoans
occurred most likely in marine invertebrates
(Cox 1994; Leander 2008; Théodoridès 1984),
which are the hosts of early branching gregarines
and coccidians, as well as of some sporozoans
incertae sedis that could be a source of important
evidence for reconstructing the ancestral states for
the Apicomplexa as a whole.
One of such unusual and poorly studied organisms
are the blastogregarines – a tiny group
of uncertain taxonomic affiliation encompassing
intestinal parasites of marine polychaetes of
the family Orbiniidae. This group comprises four
species formally belonging to the single genus
Siedleckia: S. nematoides Caullery and Mesnil,
1898, S. mesnili Chatton and Dehorne, 1929,
S. caulleryi Chatton and Villeneuve, 1936, and
S. dogieli Chatton and Dehorne, 1929. The type
species S. nematoides was described from the
intestine of the polychaete Scoloplos armiger
(Caullery and Mesnil 1898). Typical features of
blastogregarines are epicellular parasitism like gregarines,
persistent multinuclearity in trophozoites
(unlike gregarines, which are uninuclear), bending
motility like archigregarines – the most plesiomorphic
gregarine group (Schrével and Desportes
2015; Schrével et al. 2013), and the capacity to
produce globular buds, putative stages of game-
Blastogregarines: Plesiomorphic Apicomplexa 699
togenesis, from their posterior end (Chatton and
Dehorne 1929; Chatton and Villeneuve 1936a).
This latter feature is unknown from other sporozoans
and led to the name “blastogregarines”
(i.e., “budding gregarines”) (Chatton and Villeneuve
1936a). The life cycle of blastogregarines (Fig. 1)
was proposed by Chatton and co-authors relying
on evidence from studies on S. mesnili and
S. caulleryi (Chatton and Dehorne 1929; Chatton
and Villeneuve 1936a). Although the sexual process
was studied incompletely (gamete formation,
especially microgametogenesis, and the development
of zygote, i.e., sporogony, were not traced),
Chatton and Villeneuve suggested that blastogregarines
are more similar to coccidians than to
gregarines due to the absence of the syzygy and
gametocyst (characteristic for gregarines) and their
extremely pronounced anisogamy (a coccidian feature)
(Chatton and Villeneuve 1936b).
The persistent multinuclearity unusual for sporozoans
and the peculiar life cycle (absence of
so-called Leuckart’s triade: a cyclic sequence of
merogony, gamogony, and sporogony, which is
characteristic for many sporozoans, e.g., coccidians,
haemosporidians, and a part of gregarines
(Perkins et al. 2000)) gave rise to discrepant
and changeable interpretations of the taxonomic
position of blastogregarines following their initial
discovery. Different authors considered them as
aberrant gregarines (Dogiel 1910), a group incertae
sedis within sporozoans (Léger 1909; Léger
and Duboscq 1910) or a protistean lineage unrelated
to sporozoans at all (Caullery and Mesnil
1899). After more detailed studies on the life cycles
of blastogregarines (see above), Chatton and
Villeneuve (1936a,b) concluded that the permanent
gametogenesis distinguished them from both gregarines
and coccidians and suggested considering
them as an independent group of the same rank
(the order) within Sporozoa (Telosporidia). Grassé
(1953a) supposed that blastogregarines should be
included in the class Gregarinomorpha. Krylov and
Dobrovolskij challenged the assignment of the blastogregarines
to the phylum Sporozoa altogether
and considered them only as an addendum to
that (Krylov and Dobrovolskij 1980). Conversely,
de Puytorac et al. (1987) adopted and developed
the standpoint of Chatton and co-authors
and assigned the blastogregarines as the separate
sporozoan class Blastogregarinea along with
the classes of gregarines, coccidians, and haemosporidians
(de Puytorac et al. 1987). Finally, despite
the absence of syzygy, Levine and his followers
(Levine 1985; Perkins et al. 2000) assigned
the blastogregarines to the order Eugregarinorida
Figure 1. Diagram of the life cycle of blastogregarines
according to Chatton and co-authors (Chatton and
Dehorne 1929; Chatton and Villeneuve 1936a). (A)
Young mononuclear vermicular individual similar to
sporozoites of coccidians and gregarines. (B) Nondifferentiated
trophozoites attached to the intestinal
epithelium of the host; the number of the nuclei
increases during trophozoite growth. (C, D) The epicellular
trophozoites develop into the gamonts of
two types: macrogamonts (C) and microgamonts (D);
the macrogamonts (female gamonts) have the nuclei
arranged in a single row; the size of the nuclei
increases towards the rear end of the cell; the microgamonts
(male gamonts) have a similar arrangement
and size of the nuclei in the anterior third of the cell,
after that the distribution of the nuclei, which perform
multiple divisions, becomes random and they
decrease in size towards the rear end of the cell. (E–H)
The gamonts attached to the intestinal epithelium
produce either numerous uninuclear (E) or multinuclear
(F) globular buds from their posterior ends.
Chatton and Villeneuve (1936a) considered this process
as gamogony giving rise to macrogametes (E,
H) or multinuclear microgametocytes (F), which presumably
release small microgametes (G). Chatton
and colleagues detected stages in the hindgut content
that were interpreted as gamete copulation (H) and
zygotes (I). (J) Oocysts, found in the feces of the hosts,
contain 10-16 free banana-shaped sporozoites (without
sporocyst envelopes), and a residual body shifted
to one of the oocyst poles. Adapted from (Caullery
and Mesnil 1898) (A), and (Chatton and Villeneuve
1936a) (B–J).
700 T.G. Simdyanov et al.
Figure 2. General morphology of the blastogregarine Siedleckia cf. nematoides ((A–C), LM, phase contrast
or bright field; (D–F), SEM). (A) A living macrogamont individual. (B) and (C) The fixed young macrogamont
and microgamont, respectively, stained by Böhmer’s hematoxylin, flat views; three regions of the cell are conspicuous:
#1 (mucron), #2 (“asexual”), and #3 (“sexual”) – see next figure (Fig. 3) and Discussion for further
explanations. (D) Individuals attached to the intestinal epithelium of the host. (E) An individual attached to the
intestinal epithelium (under higher magnification) shows the smooth surface of the cell. (F) The mucron (arrow)
of an individual dislodged from the gut epithelium.
Blastogregarines: Plesiomorphic Apicomplexa 701
Figure 3. General ultrastructure of the blastogregarine Siedleckia cf. nematoides (TEM). (A, B) Cross sections
through the cortical region of the cell showing the trimembrane pellicle (pe) consisting of the plasma membrane
(pm) covered by the cell coat (glycocalyx, gly) and inner membrane complex (imc), longitudinal subpellicular
microtubules (smt) chiefly arranged in a layer beneath the pellicle; mitochondria (m) and the granules of amylopectin
(ap) are present in the cytoplasm. (C) Micropore (mp). (D) The longitudinal section through a parietal
part of the anterior end (region 1–see below) of an individual attached to the host cell showing longitudinal
subpellicular microtubules (smt) and numerous putative micronemes (mn). (E) two combined micrographs of
the neighbored longitudinal sections of the anterior half of a macrogamont cell showing the subdivision of the
cell into three regions: (1) mucronal region with rhoptries (rh) and putative micronemes (mn), (2) region of the
linear arrangement of nuclei (n), which is rich in channels of endoplasmic reticulum (er), putative micronemes
(mn), and mitochondria (m), and (3) posterior part of the cell (the section covered only its anterior part) rich in
amylopectin granules (ap); see Figures 4–6 for details.
that was formally explained only by the absence
of merogony in their life cycle. This taxonomic
scheme has recently been accepted in the WoRMS
and NCBI databases. The more recent reviews
(e.g., Adl et al. 2012) largely ignored this group
likely because of the absence of ultrastructural and
molecular phylogenetic evidence that makes their
phylogenetic relationships and taxonomic position
actually obscure. It should be emphasized that
the structure and biology of blastogregarines were
not addressed since 1936 (Chatton and Villeneuve
1936a). Thus, the uncertain affiliation of these
unusual organisms required efforts to be clarified
with the use of modern methods. This work
presents the first ultrastructural and molecular
phylogenetic evidence from two blastogregarine
species, Siedleckia nematoides and Chattonaria
mesnili (formerly S. mesnili) gen. n., comb. n., which
lead us to redefine the phylogenetic and taxonomic
position of the group.
Results
Morphology and Ultrastructure
Siedleckia cf. nematoides Caullery et Mesnil,
1898. Although the morphology of these blastogregarines
collected by us matched the first description
of Siedleckia nematoides (Caullery and Mesnil
1898) and excellent drawings in a later paper about
it (Caullery and Mesnil 1899), we use “cf.” (Lat. confer
– compare with) in the species name because
the sampling was performed quite far from the
type locality (Gulf of Wimereux, the English Chan-
702 T.G. Simdyanov et al.
Figure 4. Ultrastructure of the attachment apparatus of the blastogregarine Siedleckia cf. nematoides
(TEM). (A) Longitudinal section through the anterior part of a blastogregarine attached to the host
intestinal epithelium showing the large mucronal vacuole (mv), rhoptries (rh), and nuclei (n). (B–F) Longitudinal
sections of the mucron showing the details of its organization: the conoid (co), the mucronal
vacuole (mv), the rhoptries (rh) with the ducts (rd), external and internal parts of the polar ring (epr and ipr,
Blastogregarines: Plesiomorphic Apicomplexa 703
nel, France) and, additionally, due to the possibly
existence of cryptic species within this morphospecies
(see below). Individuals of S. cf. nematoides
were isolated from the intestine of the polychaete
worm Scoloplos cf. armiger (O.F. Müller, 1776)
from two localities in the White Sea, Russia: near
Marine Biological Station of Saint Petersburg State
University (MBS) and near White Sea Biological
Station of Moscow State University (WSBS). All
dissected worms (more than 200 individuals) from
both localities were infected. The parasites were
observed in the midgut (the area behind the stomach)
among epithelial cells bearing microvilli and
cilia. The intensity of the infection varied from few
parasites per host up to 20 cells per 0.01 mm2
in
some loci of the host intestinal epithelium (SEM
data on 30 samples). Neither light nor electron
microscopy revealed any appreciable differences
between individuals of S. cf. nematoides from the
both sampling localities. The cells of S. cf. nematoides
were elongated and flattened with pointed
anterior and rounded posterior ends; their smooth
surface lacked any grooves and folds (Fig. 2A–E).
The cell size varied broadly among individuals:
length 5–200 ␮m (av. 70 ␮m; mode 60 ␮m; n = 139),
width 3–17 ␮m (av. 9 ␮m; mode 8 ␮m; n = 139)
across flattened side, and 1–3 ␮m (n = 139) across
narrow side. Living blastogregarines attached or
dislodged from the intestinal epithelium were continuously
motile with active bending, twisting, and
squirming movements (for details, see: Valigurová
et al. 2017).
In female gamonts (=macrogamonts), nuclei
were arranged in a row along the cell’s length
(Fig. 2A, B). They were located closer to each
other in the anterior half of the body and appeared
compressed and ranged in size from 0.5 × 1.3 up
to 0.7 × 2.2 ␮m (av. 0.9 × 1.3 ␮m, n = 60 nuclei in
10 individuals). The distance between the nuclei
increased and they became a little larger towards
the posterior end (from 0.5 × 1.6 up to 1.0 × 2.9 ␮m;
av. 1.1 × 1.7, n = 32, the same 10 individuals).
The nuclei in the male gamonts (=microgamonts)
were much more numerous (Fig. 2C) with a linear
arrangement only in the anterior part of the
cell. Like the nuclei of macrogamonts in this region,
they were slightly compressed and ranged in size
from 0.5 × 1.1 up to 1.8 × 2.2 ␮m (av. 1.1 × 1.5 ␮m,
n = 56, 10 individuals). In the posterior part of
microgamonts, the nuclei were distributed randomly;
their shape became irregular and the
size decreased (0.5 × 0.7 up to 1.1 × 1.4 ␮m; av.
0.7 × 0.9, n = 58, the same 10 individuals). The border
between these patterns of distribution lay nearly
in the middle of the body in smaller (younger) indviduals
(Fig. 2C) or moved to the anterior third of
the body in larger (elder) ones (not shown, see:
Caullery and Mesnil 1899). Compared with younger
gamonts, the nuclei were more numerous in elder
ones, both female and male.
Individuals of S. cf. nematoides embedded their
apical end (mucron) into the brush border of enterocytes
bearing microcilia and microvilli (Fig. 2D, E).
When dislodged after fixation, some of them, with
an intact attachment site, exhibited a small apical pit
in its center (Fig. 2F). No additional structures providing
attachment, e.g., hooks or other projections,
were found.
The tegument of S. cf. nematoides cells (Fig. 3A,
B) was represented by a trimembrane pellicle,
32 nm thick. It consisted of the plasma membrane
with a well-developed glycocalyx, and two
closely adjacent cytomembranes forming the inner
membrane complex, IMC. The internal lamina,
an electron-dense layer just beneath the IMC,
which is characteristic for gregarines (Schrével
et al. 2013) was not detected. A single layer of
numerous regularly arranged longitudinal subpellicular
microtubules arose from the anterior end
and passed along the whole cell (Fig. 3D); each
microtubule appeared to be surrounded by an
electron-translucent area (Fig. 3A, B). Few additional
microtubules were located just beneath this
layer (Fig. 3A). Micropores (Fig. 3C) were detected
rarely, in the anterior part of the cell. The cell of
S. cf. nematoides is highly polarized and the ultrastructure
allowed to define three regions (Fig. 3D,
E): (1) the mucron (attachment and feeding apparatus)
containing organelles of the apical complex
and lacking nuclei, (2) the region of linear arrangerespectively),
longitudinal subpellicular microtubules (smt) arising from epr, putative micronemes (mn), and
mitochondria (m); the mucron is covered by the pellicle (pe) forming a septate cell junction (sj) with the host
cell; hm, plasma membrane of the host cell, pm, plasma membrane of the parasite cell, imc, inner membrane
complex, gly, glycocalyx; arrows mark the modified part of the host cell surface facing parasite cytostome.
Image (F) also shows longitudinal thick fibrils (f) around the posterior part of the mucronal vacuole. (G) Cross
section through the anterior part of a blastogregarine cell behind the mucronal vacuole showing a crystalloid
structure (cr). (C, E, F) fixed in the presence of ruthenium red (see “Methods”).
704 T.G. Simdyanov et al.
ment of nuclei rich in endoplasmic reticulum (ER),
and (3) the posterior half of the body (in full-sized
individuals) with significant differences between the
macro- and microgamonts in the structure, number,
and arrangement of nuclei that will be detailed
below.
TEM studies revealed that the attachment apparatus
of S. cf. nematoides is a mucron. It was
embedded in the host enterocyte between the
microvilli (Fig. 4A) and contained the mucronal vacuole,
the conoid, two (internal and external) polar
rings, and numerous rhoptries (Fig. 4B–E). The
conoid (Fig. 4B–E) had upper and basal diameters
of nearly 290 and 400 nm, respectively, and
its height was about 130 nm. It was built of coiled
microtubules: six microtubules were visible on longitudinal
sections (Fig. 4B, D); the anterior three
of them were closely adjacent to each other, while
the rear three were spaced 6–7 nm apart (Fig. 4D).
The polar ring, located slightly higher than the
upper opening of the conoid (Fig. 4D, E), gave
rise to the longitudinal subpellicular microtubules,
i.e., it is their MTOC (the microtubule organization
center). The polar ring appeared to be subdivided
into two parts of different electron density:
external and internal. The external part having moderate
electron density was about 40 nm thick with
the external diameter about 450 nm. The internal
part (Fig. 4D) of high electron density is thinner
(∼20 nm thick) and narrower (ext. diam ∼360 nm,
int. diam. ∼340 nm) than the external one. The
majority of longitudinal sections showed a voluminous
mucronal vacuole with a loose fibrous
material and a wide duct passing through the
conoid and opening outside, i.e. into the space
between the parasite and host cells (Fig. 4A–F).
Several large rhoptries were distributed around
and behind the mucronal vacuole; they formed
ducts passing through the conoid and, apparently,
opened outside the cell (Fig. 4C, E). Few putative
micronemes and mitochondria were detected
in the parietal area of the anterior region of the
mucron (Fig. 4B–E). The number of the putative
micronemes increased in the region behind the
mucronal vacuole where they had a regular distribution
on the periphery of cytoplasm (Figs 3D,
4G). The mucron was covered by the trimembrane
pellicle excepting the region against the conoid
with the wide inlet opening (diam. ∼130 nm) of the
mucronal vacuole: we consider it to be a cytostomecytopharyngeal
complex performing myzocytosis
(Fig. 4D, E). The cytostome was opened into a
gap (∼20 nm) between the parasite and host cell
plasma membranes, which had the appearance of
the septate cell junction; the “septa” were putatively
formed by both parasite and host cell coats (Fig. 4D,
E). The region of host plasma membrane facing
the parasite cytostome was of higher electron
density than the rest of the membrane and had uniformly
spaced electron-dense structures appeared
as bold dots on the external surface of the host cell
(Fig. 4D, E); it might be a perforated or modified
host cell coat. No other significant modifications of
the host cell were detected. Some additional structures
were observed in the mucron and the region
just behind it: thick fibrils around the posterior part
of the mucronal vacuole, and a crystalloid structure
(Fig. 4G).
The main feature of the anterior half of the cell
behind the mucron (region 2) is the linear arrangement
of nuclei (Fig. 3E). Another conspicuous
characteristic is the abundance of the channels
or cisterns of the endoplasmic reticulum (ER),
arranged uniformly in the cytoplasm, chiefly perpendicularly
to the longitudinal cell axis (Fig. 5).
Occasionally, they were connected to the nuclear
envelopes (Fig. 5A). In the subcortical layer of the
cytoplasm, mitochondria and putative micronemes
were numerous (Figs 3E, 5); the latter were concentrated
in the lateral areas (Fig. 5A). Golgi apparatus,
few small granules of amylopectin (the storage carbohydrate),
and multimembrane structures were
also observed in this cell region (Fig. 5).
In the macrogamont cells, the number and size
of the amylopectin granules dramatically increased
from the middle of the cell (region 2) towards its rear
end (region 3) (Fig. 3E). In the region 3 of the cell,
the ER was displaced with these granules (Figs 3E,
6A). In addition, rare large electron-dense globules
(not observed in the anterior half of the cell) were
present here (Fig. 6A). All nuclei in the macrogamont
cells had similar structure: they contained
small clods of heterochromatin and a single nucleolus,
however, the nuclei in the posterior region were
slightly larger than those in the region 2 of the cell:
about 1-2 × 2 ␮m and 1.2 × 1.7 ␮m, respectively
(Figs 3E, 6A).
Unlike macrogamonts, the structure of nuclei in
the microgamont cells changed from the anterior
to the posterior end of the body, with a gradual
increase in chromatin condensation (Fig. 6B, C). In
the middle of the cell (region 2), the nuclei were
completely filled by highly condensed chromatin
(Fig. 6B). Raikov called such nuclei “of spermal
type” in his classification of protist nuclei (Raikov
1982). From the border between regions 2 and 3
and further towards the rear, the linear arrangement
of the nuclei became disordered and the
dividing nuclei were repeatedly observed; some of
them were equipped with kinetosomes (Fig. 6C–E).
Blastogregarines: Plesiomorphic Apicomplexa 705
Figure 5. Cytoplasmic organelles of the blastogregarine Siedleckia cf. nematoides (TEM). (A) Cross section
through the region 2 of the cell (linearly arranged nuclei) showing putative micronemes (mn) in the lateral regions
of the cell and a connection of the endoplasmic reticulum (er) with the nuclear envelope (arrow). (B) Fragment
of the longitudinal section of the same region as in (A) showing Golgi apparatus (g), abundant channels of
the endoplasmic reticulum (er), numerous mitochondria (m), and putative micronemes (mn). Other abbreviations:
ms, the multimembrane structure; n, nucleus; pe, pellicle; ap, amylopectin granules; smt, longitudinal
subpellicular microtubules.
Electron-translucent areas within the dividing nuclei
contained microtubules, probably the elements of a
mitotic spindle (Fig. 6D, E). The connection of these
microtubules with the kinetosome or any other
possible MTOC was not observed. The nuclear
divisions in the region 3 of the microgamont cells
obviously correspond to progamic mitoses resulting
in the production of microgametes based on
the the appearance of two adjacent kinetosomes
(Fig. 6D, E). The cytoplasm of the posterior region
in the microgamont cells had a completely different
structure than that of the macrogamont cells due
to the presence of the ER channels and only few
small amylopectin granules (Fig. 6D, E).
Chattonaria (Syn. Siedleckia) mesnili (Chatton
et Dehorne, 1929) gen. n., comb. n. Individuals
of Chattonaria mesnili were isolated from the
intestine of the polychaete worms Orbinia (Syn. Aricia)
latreillii (Audouin et H. Milne Edwards, 1833)
(Orbiniidae) collected on littoral zone of English
Channel, France (see “Methods” for details). We
managed to sample only 12 individuals of the host
species Orbinia latreillii, four of them were free
of parasites and eight were infected. The parasites
were found in the stomach – the dilated
part of the intestine situated immediately after the
esophagus and covered by glandular cells lacking
cilia. The intensity of infection varied from few to
tens of parasites (30–40) per host (the latter was
observed only in two worms). As a direct consequence
of difficulties to sample hosts with a high
prevalence of the blastogregarine parasites, the
number of C. mesnili cells analyzed under SEM,
and TEM was significantly lower than in S. cf. nematoides.
LM studies were not performed especially
as the collected cells fully fit the first description
of Siedleckia mesnili and its drawings of excellent
quality (Chatton and Dehorne 1929). The collected
cells were elongated (48.8 × 7.6 to 220 × 13 ␮m;
av. 97.4 × 9.1 ␮m; n = 8) and cylindric with a roundly
pointed posterior end. Their tegument formed longitudinal
folds (Fig. 7A–E). These originated behind
the mucron, where pairs of adjacent folds shared
a common origin (Fig. 7D), and ran towards the
posterior end of the cell, where they merged into
a smooth terminal region (Fig. 7E). The number
of the folds was 28 and 34 on the cross-sections
of two different individuals. The anterior ends of
the parasites were embedded in the cells of the
host stomach (Fig. 7A). One of the individuals,
artificially dislodged during mounting of the SEM
preparation, exhibited the anterior end covered with
a remnant of the host tissue, therefore we were not
able to observe the superficial structure and the
shape of the attachment apparatus (Fig. 7B, D),
however, the first description suggested it equipped
with hooks. In contrast to S. cf. nematoides, the
living individuals of C. mesnili observed under a
stereomicroscope immediately after the host dissection
showed only weak motility by slow and
intermittent bending movements.
706 T.G. Simdyanov et al.
Figure 6. Nuclear apparatus of the blastogregarine Siedleckia cf. nematoides (TEM). (A) The cross section
through the rear part (region 3) of a macrogamont cell showing the abundance of amylopectin granules
of large size (ap), a mitochondrion (m), putative micronemes (mn), an electron-dense globule (dg), and a
nucleus (n) containing heterochromatin and a nucleolus. (B) The longitudinal section of a microgamont cell in the
Blastogregarines: Plesiomorphic Apicomplexa 707
The cells of C. mesnili were covered by the trimembrane
pellicle about 50 nm thick and organized
in longitudinal folds with flattened tops (Fig. 7F,
G). The size of the folds was about 1.1 ␮m high
and 0.9 ␮m wide at their bases. The height was
more and the width was less in the anterior part
of the cell: about 2 and 0.3 ␮m at the bases,
respectively, so the shape varied depending on
the section site (compare Fig. 7F and G). Just
beneath the pellicle, numerous longitudinal subpellicular
microtubules were observed, with a regular
distribution and arrangement: two layers in the
tops of the folds and a single layer on their lateral
sides and between them (Fig. 7G). Although
no typical micropores were observed, microporelike
structures interrupting the IMC and the layer
of subpellicular microtubules and connected to
multimembrane vesicles were present (Fig. 7G,
compare with Fig. 5A). Similarly to S. cf. nematoides,
the cell of C. mesnili can be subdivided
into three regions: (1) the mucronal region, (2) the
region of the linearly arranged nuclei, and (3) the
posterior region with developing gametic nuclei.
TEM studies revealed that the attachment apparatus
of C. mesnili is a strongly modified mucron
lacking the conoid and anchored in the host cell
with peripheral bulges (several or the only circular
one) formed by large alveoli between the cytomembranes
of the IMC (Figs 7B, E; 8A–D, G). One
observation suggested that a hook-like cytoplasmic
projection jutted into the alveolus (Fig. 8A,
C). The flat top of the mucron was covered by
the pellicle of varying thickness (∼14 to 27 nm)
with a loose layer of fibrils just beneath it. In
this region, the middle and inner membranes of
the IMC terminated around an external opening
(cytostome) of the mucronal vacuole. The diameter
of the cytostome was about 110 nm (Fig. 8E).
Similarly to S. cf. nematoides, a gap of varying
width (∼20 to 45 nm) was present between the
parasite pellicle and the host cell membrane, but
that was not a septate cell junction because of the
absence of “septa”. The host plasma membrane
had an increased electron density in front of the
cytostome (Fig. 8D–F). The frontal region of the
mucron cytoplasm was free of organelles, with an
exception of the mucronal vacuole connected to the
cytostome by a wide duct (Fig. 8D, E). Near the IMC
terminus, a structure similar to an apical polar ring
was observed (Fig. 8E, F). This putative polar ring
(∼27 nm thick, ext. diam. ∼240 nm) was not subdivided
into any parts, as it was observed in S. cf.
nematoides, and no microtubule contacting with it
were detected, even though they were abundant
within the mucron. The microtubules arose immediately
from the fibrillar matter lying beneath the
pellicle in the frontal region of the mucron. Numerous
longitudinal microtubules were located in the
cytoplasm behind the mucronal vacuole (Fig. 8D,
E). Microneme-like bodies were detected in the
parietal region of the mucron (Fig. 8D), but no
obvious rhoptries or rhoptry ducts were observed.
However, large electron-dense globules (∼300 nm
in diameter) without ducts were present in the
region behind the mucron and around the first
nucleus (Figs 8A, B; 9A).
The first non-differentiated nuclei of both macroand
microgamonts cells were slightly ellipsoid in
shape (up to ∼1.5 × 1 ␮m) and contained 1 nucleolus
(Figs 8A–B; 9A–B). The channels of the ER,
dense bodies (likely micronemes), and putative
mitochondria were also present.
The region of linearly arranged nuclei (region
2) was studied only in the macrogamonts.
The nuclei (up to ∼2 × 1.5 ␮m) contained one
nucleolus and heterochromatin that tended to
congregate in a single lump (Fig. 9C). Other
observed organelles include the small micronemelike
bodies (Fig. 9C, D), few small amylopectin
granules, putative mitochondria, and well developed
ER; however, unlike S. cf. nematoides,
the ER did not exhibit the regular arrangement
(Fig. 9C).
The posterior end (region 3) of the macrogamont
cells showed nuclei (up to ∼1.7 × 1.2 ␮m)
with the circular arrangement of the heterochromatin
and excentric nucleolus. The cytoplasm
appeared vacuolated and contained numerous
middle of the body (posterior part of region 2) and (C) the diagonal section of a microgamont cell through the
border between regions 2 and 3: abundant nuclei (n) with the highly condensed chromatin are arranged in
the raw in region 2; they acquire kinetosomes (k) near the border between regions 2 and 3 and then become
arranged randomly; (C) fixation in the presence of ruthenium red. (D, E) Cross sections through the rear part of
a microgamont cell (region 3) showing the channels of endoplasmic reticulum (er), the small and rare granules
of amylopectin (ap), a putative progamic mitosis, and the nuclei of future microgametes (mgn) associated with
the kinetosomes (k). Note microtubules, probably of the mitotic spindle: (mt) and compare with the subpellicular
microtubules (smt).
708 T.G. Simdyanov et al.
Figure 7. General morphology and cortex organization of the blastogregarine Chattonaria mesnili ((A–E) SEM;
(F, G) TEM, cross sections). (A) Two individuals attached to the stomach epithelium of the host. (B) An individual
artificially dislodged from the epithelium during mounting of the SEM preparation exhibiting the attachment
apparatus (aa) embedded in the fragment of the intestinal lining. (C–E) Details of (B) under higher magnifications:
surface with longitudinal pellicular folds (C), the attachment apparatus (aa) covered by the remnant of
the host tissue (D), and the smooth posterior end (E). (F) Cross section of an individual through anterior region
of the cell showing transversally cut longitudinal pellicular folds (pf) and a nucleus (n) with a single nucleolus.
(G) A fragment of another cross section under higher magnification showing trimembrane pellicle (pe) coated
by glycocalyx (gly), longitudinal subpellicular microtubules (smt) located just beneath it, and a micropore-like
structure (mps).
Blastogregarines: Plesiomorphic Apicomplexa 709
Figure 8. Attachment apparatus (mucron) of the blastogregarine Chattonaria mesnili (TEM). (A–B) Longitudinal
sections of anterior parts of two individuals attached to the host cells: a microgamont (A) and a macrogamont
(B) showing nuclei (n) and dense globules (dg) in the cytoplasm; two marked regions are corresponding those
of S. cf. nematoides (see Fig. 3). (C) Longitudinal section of the same individual that in (A) showing a large
alveolus (alv) with a protrusion of the cytoplasm and microtubules (mt) arising from the pellicle (pe) of the
mucron; hm and pm are host and parasite plasma membranes, respectively. (D–F) Longitudinal sections of
the mucron and its parts of the same individual as in (B) showing myzocytosis through the cytostome facing
the electron-dense region of the host cell plasma membrane (white arrow), mucronal vacuole (mv), putative
polar ring (pr?) closely adjacent to endings of the IMC (imc), subpellicular microtubules (smt) without any visible
MTOC, microtubules (mt) arising from the frontal zone of the mucron, microneme-like bodies (mn?), the cell
junction, which is a non-septate gap between parasite and host plasma membranes (pm and hm, respectively).
(G) A detail of (D) showing the structure of the large alveolus (alv) formed as space between the middle (mm)
and inner (im) membranes of the pellicle; hm and pm are the plasma membranes of the host and parasite cells,
respectively.
710 T.G. Simdyanov et al.
Figure 9. Nuclear apparatus and cytoplasm of the blastogregarine Chattonaria mesnili (TEM). (A) The first
nucleus of a microgamont (the same cell as in Fig. 8A) containing heterochromatin (hcr) and a single nucleolus
(nl) and surrounded by large dense globules (dg). (B) Fragment of a cross section through the anterior part
of a macrogamont (the same cell as in Fig. 7F) showing a nucleus with a single nucleolus (nl), but lacking
heterochromatin; the channels of the endoplasmic reticulum (er), microneme-like bodies (mn?), and a putative
mitochondrion (m) are visible. (C) Longitudinal section of the middle part of a macrogamont cell (region 2,
corresponding that of S. cf. nematoides) showing nuclei with the large lumps of the heterochromatin (hcr)
and a single nucleolus (nl, visible in the lowest nucleus), the channels of the endoplasmic reticulum (er),
putative mitochondria (m), rare small granules of the amylopectin (ap), and microneme-like bodies (mn?). (D)
Fragment of the cytoplasm (the same region as in (C)) showing microneme-like bodies (mn?) under a higher
magnification. (E) Longitudinal section of the posterior end of a macrogamont (region 3, corresponding to that
of S. cf. nematoides) showing nuclei with the large lumps of the heterochromatin (hcr) and weakly developed
nucleoli (nl, visible in the left lower nucleus); the cytoplasm is vacuolated (v, vacuoles) and contains numerous
globules (gl), putatively of protein, and grains of amylopectin (ap). (F–I) Diagonal sections of the posterior
parts of microgamonts (region 3) under different magnifications showing numerous randomly distributed male
gametes’ nuclei (n) associated with two kinetosomes (k) and connected to the main cytoplasm with a stalk (black
arrow); the cytoplasm contains numerous microneme-like bodies (mn?, in (H)). (J) cross-section through two
kinetosomes connected with an electron-dense link (white arrow). (K) cross section of the top of a kinetosome
(the transitive zone to the future flagellum) showing 9 groups (doublets) of microtubules.
Blastogregarines: Plesiomorphic Apicomplexa 711
globules (putatively proteins), and amylopectin
granules not abundant in this region (Fig. 9E).
The posterior end of the microgamont cells contained
numerous spherical (∼0.6 ␮m in diameter)
nuclei of “spermal” type according to the classification
by Raikov (1982) with highly condensed
chromatin in the whole volume (Fig. 9F – I).
Each nucleus was encircled by an electrontranslucent
zone; some with a stalk connected
them to the main cytoplasm (Fig. 9H). Most
nuclei were associated with two kinetosomes containing
nine peripheral groups of microtubules,
apparently triplets or doublets depending on
the kinetosome region; the kinetosomes were
connected to each other by an electron-dense
link (Fig. 9J, K). The cytoplasm appeared vacuolated,
with abundant microneme-like bodies
(Fig. 9H); the amylopectin granules and putative
protein globules were not detected. No
mitochondria were observed in the posterior
regions of both macrogamont and microgamont
cells.
Molecular Phylogenetic Analyses
For S. cf. nematoides, the contiguous sequence
of the near-completed ribosomal operon (SSU
rDNA + ITS1 + 5.8S rDNA + ITS2 + LSU rDNA) was
obtained from the WSBS sample, whereas only a
partial sequence of SSU rDNA was obtained from
the MBS sample. For the sample of C. mesnili,
the obtained contiguous sequence covered nearcomplete
gene of SSU rRNA, ITS1, 5.8S rDNA,
ITS2, and a large part (∼2,000 bp) of LSU rDNA
(Table 1). All these sequences were involved in
phylogenetic analyses with the use of Bayesian
inference (BI) and Maximum likelihood (ML) methods
(see “Methods” for details).
Analyses of 18S (SSU) rDNA. Both Bayesian
inference and Maximum likelihood (ML) analyses
resulted in almost identical tree topologies with
some differences in the branching within the coccidia
+ hematozoa clade (not shown). The Bayesian
tree appeared more accurate than the ML one: in
the ML tree Plasmodium spp. grouped not with piroplasms,
but with adeleid coccidians, although with
low support (BP = 38%), i.e. hematozoans were
split up. We consider this an artifact, which is
absent in the Bayesian tree. This point does not
affect the position of blastogregarines and neighboring
branches. The higher accuracy of Bayesian
inference than ML bootstrap analysis was also
revealed previously (Alfaro et al. 2003). Overall
the newly obtained phylogenies matched molecular
phylogenetic evidence from alveolates and apicomplexans
published recently (e.g., Cavalier-Smith
2014; Janouˇskovec et al., 2015; Lepelletier et al.
2014; Rueckert and Horák 2017; Schrével et al.
2016). The Bayesian tree inferred from the dataset
of 110 taxa and 1,550 sites (Fig. 10) showed the
monophyly of major alveolate groups, although with
moderate or low statistical support. The backbone
of the apicomplexan region in the newly obtained
tree was poorly resolved by both Bayesian and
ML analyses. The three obtained blastogregarine
sequences together with environmental sequence
D3P05D06 formed a clade with full PP (posterior
probabilities in Bayesian analysis) and high BP
(bootstrap percentage in ML analysis) supports (1.0
and 90%, respectively). This robust blastogregarine
clade was located between two archigregarine lineages,
but all node supports in this region of the
tree backbone were extremely low (Fig. 10). The
archigregarine split had been already recovered
before, also with very weak supports (Rueckert
et al. 2015; Rueckert and Horák 2017; Schrével
et al. 2016; Wakeman et al. 2014; Wakeman and
Horiguchi 2017; Wakeman and Leander 2013), i.e.
this is not an effect of the addition of blastogregarines
to the taxon sampling. SSU rDNA identities
between the two S. cf. nematoides samples and
the environmental sequence D3P05D06 were at
about 90% (MBS vs. D3P05D06 = 90%, WSBS
vs. D3P05D06 = 92.1%, WSBS vs. MBS = 93.7%),
whereas those between C. mesnili and S. cf.
nematoides were lower, at 82.8%–83.9% (see Supplementary
Material Table S1 for details).
Analyses of 28S (LSU) rDNA and the ribosomal
DNA operon. All phylogenies based on
these phylogenetic markers resulted in identical
topologies both in the Bayesian (Fig. 11) and ML
(not shown) analyses. Overall, they recovered the
major alveolate clades that agreed the phylogenies
inferred from SSU rDNA – both already published
(see above) and newly obtained – but with the
higher resolution of all-alveolate and myzozoan
deep branching than in SSU rDNA trees.
In the LSU rDNA-alone-based phylogeny
(Fig. 11A), the monophyletic apicomplexan clade
comprised the moderately supported chrompodellid
lineage and sporozoans; the latter were
subdivided into the clades of firmly supported
coccidiomorphs (coccidians + hematozoans)
and moderately supported by BP cryptosporidians
+ gregarines. All presented (available)
gregarine sequences formed a monophyletic
clade, although the BP support was only
moderate (PP = 1.0, BP = 84%). Similar val-
712 T.G. Simdyanov et al.
ues of BP supports were also obtained for
other nodes in the backbone of this clade.
The monophyly of eugregarines was broken
due to the archigregarine Selenidium sp. In
contrast to the SSU rDNA phylogenies, the
blastogregarine LSU rDNA sequences showed
no affinity to gregarines, but formed a sister
branch to the coccidiomorph clade, although with
rather low PP and BP supports (0.67 and 52%,
respectively).
Compared with the LSU rDNA, phylogenies using
the near complete ribosomal DNA operons (concatenated
SSU, 5.8S, and LSU rDNAs) showed
higher BP supports for the dinozoans and for the
sporozoans (Fig. 11B). In contrast, BP supports
for the cryptosporidians + gregarines clade and
its internal branching decreased. The sistership of
the blastogregarines and coccidiomorphs was also
recovered, although with lower support (PP = 0.50,
BP = 37%).
Testing alternative topologies. The alternative
topologies of phylogenetic trees were analyzed with
the use of the set of six widespread tests (see
“Methods” for details). Among 12 alternative SSU
rDNA phylogenies (Supplementary Material Fig.
S1), six were found to reject by no test (Fig. 12A);
four of them represented the blastogregarines as a
member of the clade cryptosporidians + gregarines
and two as a sister group either to this clade or
to the sporozoans as a whole. However, approximately
unbiased test and majority of others did not
reject any alternative topology including the direct
association of the blastogregarines with the coccidiomorph
clade (see Supplementary Material Table
S2 and Fig. S1, topologies #7 and 10). However, the
boostrap probability rejected this location (topologies
#7 and 10 in the Supplementary Material Fig.
S1), even though it was the best in the LSU rDNA
and the ribosomal operon phylogenies (Fig. 11, the
reference topologies #0 and alternative topologies
#1 in Fig. 12B and C). In contrast, among LSU
rDNA and ribosomal operon phylogenies, all tests
rejected any position of the blastogregarines within
the cryptosporidian + gregarine clade, as chiefly
preferred in SSU rDNA phylogenies (Supplementary
Material Table S2; Fig. 12B, C). However,
no test rejected the blastogregarines as the sister
group to all other sporozoans. In summary,
the blastogregarines as the earliest branch of the
sporozoans was the only case permitted by all tests
in all three genetic markers examined (i.e., SSU
rDNA, LSU rDNA, and ribosomal operon phyloge-
nies).
Discussion
The general morphology and ultrastructure of blastogregarines
represent a fanciful combination of
the features characteristic to different far-related
taxa of sporozoans (Fig. 13). This is congruent
with the uncertain position of blastogregarines
within the sporozoans provided by the conflicting
SSU rDNA and LSU rDNA ribosomal operon
molecular phylogenies. On the one hand, blastogregarines
share many ultrastructural features
with archigregarines, the most plesiomorphic group
of the sporozoans known to date (Schrével 1971b;
Schrével and Desportes 2015; Schrével et al.
2013) – that agrees with SSU rDNA-based phylogenies
placing these organisms in the close
neighborhood to each other, although with low
supports. First, both blastogregarines and archigregarines
possess longitudinally folded or smooth
pellicle – in archigregarines, the latter is rare, but
sometimes exists (Simdyanov 1992) – and longitudinal
subpellicular microtubules arranged in layer(s)
just beneath it (Schrével 1971a,b; Simdyanov and
Kuvardina 2007). Second, the mucron and apical
complex in both these groups does not disappear
in the early developmental stages as in
majority of sporozoans, but persists over a long
period of time: during the larger part of their life
cycle indeed. The trophic stages of both blastogregarines
and archigregarines attach to host cells
with the mucron that contains the mucronal vacuole
and well-developed components of the apical
complex (at least in S. cf. nematoides among blastogregarines)
and performs myzocytotic feeding
(Schrével 1968, 1971b; Simdyanov and Kuvardina
2007). In blastogregarines, this mucronal complex
(the apical complex and mucronal vacuole)
remains active (myzocytosis) during the trophozoite
lifespan. Unlike blastogregarines, archigregarines
have non-feeding mature gamonts (syzygy stage),
but the conoid and rhoptries persist in them until
starting progamic mitoses at least (Schrével et al.
2013; Simdyanov and Kuvardina 2007). Despite
generally plesiomorphic body plan, the studied
blastogregarines exhibit modifications of the cortex
and mucron, which can be considered autapomorphies,
and they show a mosaic distribution.
Thus, the folded pellicle characteristic for both
C. mesnili and many Selenidium spp. appears to
be a plesiomorphic trait, but the major modification
of its mucron (loss of conoid and rhoptries)
is a distinct apomorphy. On the contrary, S. cf.
nematoides, has a plesiomorphic mucron with
the complete apical complex; however, its smooth
pellicle appears rather an apomorphy. The more
Blastogregarines:PlesiomorphicApicomplexa713
Table 1. The main characteristics of the obtained blastogregarine sequences: the source and total length of assembled contiguous sequences,
the length of overlapping PCR-amplified fragments used for their creation, and PCR primers used for the amplification of the fragments.
Sources and total
lengths of
assembled
sequences
Amplified fragmentsa Fragment
length
PCR primers: forward (F) and reverse
(R); annealing temperature used in the
PCRs
Siedleckia cf.
nematoides from
MBS (1,645 bp)
MH061197
SSU rDNA (part) 1,645 bp (F) 5 -GTATCTGGTTGAT
CCTGCCAGT-3
(R) 5 -GCGACGGGCGGTGTGTAC-3
t◦ = 48 ◦C
Siedleckia cf.
nematoides from WSBS
(5,617 bp) MH061198
(I) SSU rDNA (part) 1,765 bp (F) 5 -GTATCTGGTTGAT
CCTGCCAGT-3
(R) 5 -GAATGATCCWTC
MGCAGGTTCACCTAC-3
t◦ = 48 ◦C
(II) SSU rDNA (part),
ITS1, 5.8S rDNA, ITS2,
LSU rDNA (part)
2,004 bp (F) 5 -GCATGGCCGTTCT
TAGTTGGTGG-3
(R)
5 -CCTTGGTCCGTGTTTCAAGAC-3
t◦ = 48 ◦C
(III) LSU rDNA (part) 1,021 bp (F) 5 -ACCCGCTGAAYTT
AAGCATAT-3
(R) 5 -GCTATCCTGAGGG
AAACTTCGG-3
t◦ = 53 ◦C
(IV) LSU rDNA (part) 1,549 bp (F) 5 -GTCTTGAAACACG
GACCAAGG-3
(R) 5 -CAGAGCAGTGGGC
AGAAATC-3
t◦ = 53 ◦C
714T.G.Simdyanovetal.
Table 1 (Continued)
Sources and total
lengths of
assembled
sequences
Amplified fragmentsa Fragment
length
PCR primers: forward (F) and reverse
(R); annealing temperature used in the
PCRs
(V) LSU rDNA (part) 1,001 bp (F) 5 -GTAACTTCGGGAW
AAGGATTGG CT-3
(R) 5 -GTCTAAACCCAGC
TCACGTTCC CT-3
t◦ = 53 ◦C
Chattonaria mesnili
(4,560 bp) MH061199
(VI) SSU rDNA (part) 1,782 bp (F) 5 -GTATCTGGTTGAT
CCTGCCAGT-3
(R) 5 -GATCCTTCTGCAG
GTTCACCTAC-3
t◦ = 48 ◦C
(VII) SSU rDNA (part),
ITS1, 5.8S rDNA, ITS2,
LSU rDNA (part)
∼1,600 bp (F) 5 -GTCCCTGCCCTTT
GTACACACCGCCCG-3
(R) 5 -CCTTGGTCCGTGT
TTCAA GAC-3
t◦ = 53 ◦C
(VIII) LSU rDNA (part) ∼2,000 bp (F) 5 -ACCCGCTGAAYTT
AAGCATAT-3
(R) 5 -AGCCAATCCTTWTCCCGAAG
TTAC-3
t◦ = 53 ◦C
aThe sequence overlap between fragments I and II is about 480 bp, II and III—about 720 bp, III and IV—about 280 bp, IV and V—about 230 bp.
The overlap between fragments VI and VII is about 180 bp, between VII and VIII—about 750 bp.
Blastogregarines: Plesiomorphic Apicomplexa 715
Figure 10. Bayesian inference tree of alveolates obtained by using the GTR + + I model from the dataset
of 110 SSU rDNA sequences (1,550 sites). Numbers at the nodes indicate Bayesian posterior probabilities
(numerator) and ML bootstrap percentage (denominator). Black dots on the branches indicate Bayesian posterior
probabilities and bootstrap percentages of 0.95 and 90%, respectively, and higher. The blastogregarine
clade is highlighted by gray. The newly obtained sequences of blastogregarines (Siedleckia nematoides and
Chattonaria mesnili) are given on black background.
716 T.G. Simdyanov et al.
Figure 11. Bayesian inference trees of alveolates obtained by using the GTR + + I model from two rDNA
datasets of the same 54-taxon sampling. (A) LSU rDNA dataset (2,912 sites); (B) Ribosomal operon dataset
(4,618 sites). Numbers at the nodes indicate Bayesian posterior probabilities (numerator) and ML bootstrap
Blastogregarines: Plesiomorphic Apicomplexa 717
Figure 12. Diagrams of the reference (“best”) phylogenies and alternative topologies rejected by none test: (A)
SSU rDNA (110-taxon dataset); (B) LSU rDNA (54-taxon dataset); (C) ribosomal operon (54-taxon dataset).
Only tree regions comprising parasitic apicomplexans are shown, the major clades correspond to those in the
phylogenies obtained by the phylogenetic analyses (Figs 10 and 11) and are shown schematically. The reference
trees are designated as (0). The alternative topologies are arranged from (1) and further on the decrease of the
test values (see Table S3): the phylogenies having number (1) are the most likely of all alternative topologies
in each category (A, B, C) and the last tree topologies are the least likely. Abbreviations: B, blastogregarines;
Ag1 and 2, archigregarine clades (see Fig. 10); Eg1 and 2, eugregarine clades (see Figs 10 and 11); Cr,
cryptosporidians; C, coccidiomorphs (coccidians + hematozoans). In the reference LSU rDNA phylogeny (B0)
the archigregarine Selenidium pygospionis is grouped with the eugregarine Ancora sagittata belonging to the
clade Eg2 in the phylogenies inferred from SSU rDNA and ribosomal operon (compare Figs 10 and 11), where
they are separated; however, no one test did rejected they separation in the LSU rDNA phylogeny too (topology
B1; the P-value of the AU test is even higher than for (0), see Supplementary Material Table S3).
primitive morphology of the mucron in this species
could be the result of something like the paedomorphosis,
i.e. the retention of “juvenile” features
in an adult individual. The third common feature
is shared by blastogregarines not only with archigregarines,
but with all other gregarines too: their
sporozoites lie in the oocysts freely (Chatton and
Villeneuve 1936a), without additional internal cysts
(sporocysts) characteristic for core coccidians, both
eimeriids and adeleids; however, in some divergent
blood-parasitic coccidians (e.g., Lanketerella)
as well as in the majority of haemosporidians the
sporocysts are absent too.
Apart from obvious similarities, there is a conspicuous
difference between blastogregarine and
archigregarine ultrastructure: the extensive development
of the ER in the anterior half of the
blastogregarine cell vs. secondary food vacuoles
in the medullary part of the cytoplasm in the archigregarine
cell (Schrével 1968, 1971b; Schrével
et al. 2013; Simdyanov and Kuvardina 2007). One
possible explanation for this is that the numerous
nuclei in blastogregarines obstruct the traffic of
food vacuoles along the axial microtubules from the
mucron along the cell towards its rear. Such feeding
mechanism was previously suggested for the archipercentage
(denominator). Black dots on the branches indicate Bayesian posterior probabilities and bootstrap
percentages of 0.95 and 90%, respectively, and higher. The blastogregarine clade is highlighted by gray. The
newly obtained sequences of blastogregarines (Siedleckia nematoides and Chattonaria mesnili) are given on
the black background. Accession numbers in (B) are arranged in following order: SSU rDNA, 5.8S (if exists),
LSU rDNA.
718 T.G. Simdyanov et al.
Figure 13. Key features of the organization of the blastogregarines. (A, B) Scheme of the general ultrastructure
of a macrogamont cell and the egenerative region of a microgamont cell, respectively. (C, D) Siedleckia
nematoides: the schemes of the longitudinal and cross sections of the mucron and tegument, respectively.
(E, F) Chattonaria mesnili: the schemes of the longitudinal and cross sections of the mucron and tegument,
respectively. Abbreviations: alv, alveoles between the cytomembranes of the IMC; ap, amylopectin granules; co,
conoid; cs, cytostome; er, endoplasmic reticulum; hm, plasma membrane of the host cell; imc, inner membrane
complex of pellicle; pm, plasma membrane; mgn, microgamete nuclei; mt, microtubules; mv, mucronal vacuole;
mu, mucron; nu, nuclei; pgm, progamic mitoses in the microgamont; pm, plasma membrane of the parasite; pr,
polar ring (gives rise to the longitudinal subpellicular microtubules (smt) in Siedleckia); rh, rhoptries; sj, septate
cell junction between the parasite and host cells.
Blastogregarines: Plesiomorphic Apicomplexa 719
gregarine Selenidium orientale (Simdyanov and
Kuvardina 2007). In the absence of that system,
the ER may substitute the function of nutrient distribution.
One more possible explanation is that the
“overdeveloped” ER provides membranes necessary
for blastogregarine budding.
On the other hand, the blastogregarines share
some features with the coccidiomorphs that agrees
with the topologies of the LSU rDNA and ribosomal
operon phylogenies (although the blastogregarine
position is also weakly supported there). First, like
eimeriid coccidians and hematozoans, they lack
one of the most characteristic features of the gregarine
life-cycle: the stage of syzygy followed by
the formation of the gametocyst (Grassé 1953a;
Levine 1971; Perkins et al. 2000; Schrével and
Desportes 2015; Schrével et al. 2013) – at least
it was never reported in the literature and also
did not detected by us throughout more than 10
years of observations. It should be noted here that
the syzygy is characteristic not only for the gregarines,
but for the adeleid coccidians too, but the
gametocyst is still absent in them (Grassé 1953b;
Perkins et al. 2000). Thus, the absent of the gametocyst
is a common feature of the blastogregarines
and coccidiomorphs. Second, judging by the two
kinetosomes associated with the spermal nuclei,
the male gametes (=microgametes) of blastogregarines
must bear two flagella as in the majority
of coccidians (Grassé 1953b; Perkins et al. 2000),
whereas gregarine male gametes are chiefly uniflagellated
or, sometimes (in Monocystidae), lack
flagella at all (Grassé 1953a; Martinucci et al. 1981;
Perkins et al. 2000; Schrével et al. 2013). Third,
blastogregarines exhibit an extremely pronounced
difference in size between male (micro-) and female
(macro-) gametes (oogamy; Fig. 1H) that is a
characteristic feature of all coccidians and haemosporidians
(Chatton and Villeneuve 1936a). Thus,
the atypical life cycle of blastogregarines is more
similar to coccidians – as it was noted by Chatton
and Villeneuve (1936b). For example, the multiple
rounds of mitosis without immediate cytokinesis,
that leads to multinuclearity (Fig. 1B–D), is characteristic
not only for blastogregarines (Caullery and
Mesnil 1899; Chatton and Dehorne 1929; Chatton
and Villeneuve 1936a), but for the growing meronts
of eucoccidians and haemosporidians too, e.g., in
Eimeria, Adelea, and Plasmodium (Grassé 1953b;
Perkins et al. 2000), although the multinuclearity
is temporary in them. The only difference here
is seemingly that these “merogonic” mitoses in
blastogregarines do not result in the formation of
merozoites (Chatton and Villeneuve 1936b).
The microgametogenesis of blastogregarines
has not been traced, but the budding-off of already
multinucleate spherical cells from microgamonts
(Fig. 1E–F) has been previously reported for all
examined blastogregarine species: S. cf. nematoides,
S. caullery, and S. mesnili (Caullery and
Mesnil 1899; Chatton and Dehorne 1929; Chatton
and Villeneuve 1936a). These “buds” are supposed
by Chatton and Dehorne (1929) to produce
microgametes and, if so, they may be considered
as microgametocytes or microgametoblasts – the
latter is a peculiarity known in Protococcidia (e.g.,
Myriospora), which lack merogony (Grassé 1953b;
Perkins et al. 2000). The formation of the microgametoblasts
in blastogregarines and, probably, in the
aforementioned protococcidians, may be considered
as the deferred merogonic divisions of the
cell. In blastogregarine macrogamonts, the suggested
“deferred merogony” results in budding-off
uninucleate putative macrogametocytes supposed
to mature into macrogametes directly, i.e. without
nuclear and cell divisions (again a coccidian feature)
(Chatton and Dehorne 1929; Chatton and
Villeneuve 1936a). To corroborate or dismiss these
putative homologies in the blastogregarine and coccidian
life cycles, the “asexual” nuclear divisions
(not leading directly to the gamete formation) in
the anterior part of the cell should be studied
in details and the fate of these nuclei should be
revealed: whether their “merogonic” divisions stop
with the gamogony starting or they may continue
within the lifespan of an individual. Another question
is: whether the female nuclei really descend
directly from the “asexual” (vegetative) nuclei of
region 2 or there are additional progamic mitoses
on the border between regions 2 and 3 as in
microgamonts. The fate of the individuals after
gametogenesis is also uncertain: whether they disintegrate
totally during the gamete formation or the
gametogenesis stops at some point and “merogonic”
mitoses start again? Anyway, the merogonic
(asexual) nuclear divisions and the formation of
gamete nuclei appear to run in blastogregarines
within the same cell, although in its different places
– in region 2 and in region 3, respectively – therefore
we introduce the name “merogamont” for blastogregarine
individuals. The merogamont cell is
differentiated into three regions specialized on different
functions: mucron or feeding region (#1),
merogonic or vegetative region (#2) where vegetative
nuclei proliferate, and gamogonic or generative
region (#3) where sexual (gametic, generative)
nuclei arise and develop (Figs 2B, C; 3C, and
13).
720 T.G. Simdyanov et al.
Although the peculiarities of the nuclear apparatus
and life cycle create a certain affinity between
the blastogregarines and coccidians, the former
also retain a set of plesiomorphic morphological
characteristics shared with the archigregarines
(see above), which should be therefore considered
as common ancestral features of the sporozoans.
Thus, we have two plesiomorphic groups
of the Sporozoa with a quite similar organization,
but one of them possesses the gregarine
life cycle (archigregarines) while the other (blastogregarines)
– the coccidia-like one. Summing
up the morphological and molecular phylogenetic
data, the blastogregarines should be considered
as a relatively isolated group of plesiomorphic
sporozoans. This conclusion appear to correlate
with the results of the alternative topology testing
that revealed the most basal position of blastogregarines
within sporozoans as the only possible
consensus between conflicting SSUrDNA and LSU
rDNA ribosomal operon phylogenies. The mismatch
of the “best” topologies might be referred
to the twice less taxon sampling of LSU rDNA
(54 vs. 110 taxa), but the comparison with the
SSU rDNA trees computed on the reduced 54taxon
dataset (see Supplementary Material Fig.
S2) showed the same features as in 110-taxon
sampling SSU rDNA phylogeny: the split of gregarines
and the affiliation blastogregarines to them.
On the other hand, these reduced SSUrDNA phylogenies
were considerably worse resolved and
less consistent in the region of the apicomplexan
backbone than both the LSU rDNA/operon-based
trees (Fig. 11) and the SSU rDNA phylogenies
inferred from the 110-taxon sampling (Fig. 10).
This point rather corroborates the previously published
opinion that SSU rDNA is likely not the
most eligible marker for the study of apicomplexan
deep branching (Simdyanov et al. 2015, 2017). The
expanded taxon sampling of LSU rDNA and multigene
analyses may resolve this ambiguity in the
future.
Molecular phylogenetic analyses have failed to
assign blastogregarines firmly to either gregarines
or coccidiomorphs, but unequivocally indicate them
as a robust clade affiliated to sporozoans that
is in full agreement with the ultrastructural evidence.
Apart from that, putative cryptic species
within the morphospecies S. cf. nematoides were
revealed by means of the molecular approach
that may be inferred from significant differences
between the sequences obtained from WSBS
and MBS samples. The single available environmental
blastogregarine-like sequence, D3P05D06
from oxygen-depleted sediment from littoral of
Greenland (Stoeck et al. 2007), is also closely
related to both sequences of S. cf. nematoides.
To indicate these putative cryptic species more
firmly, the molecular datasets must cover highly
variable regions of the genome such as the internal
transcribed spacers of the ribosomal operon,
ITS1 and ITS2 (Müller et al. 2007). The hosts
of the cryptic species might be cryptic species
revealed within the Scoloplos armiger (Bleidorn
et al. 2006) and the polychaetes closely related to
it.
Conclusion
From all diversity of the contradicting to each other
opinions reviewed in the Introduction, the results of
this study corroborate the viewpoint on the taxonomic
position of the blastogregarines of Chatton
and co-authors and their pioneering interpretation
of the blastogregarine life cycle (Chatton and
Dehorne 1929; Chatton and Villeneuve 1936a,b)
and dismiss the more recent and widespread
taxonomic scheme of Levine and his followers,
which consider blastogregarines a part of eugregarines
(Levine 1985; Perkins et al. 2000; WoRMS).
First, our ultrastructural data confirm the formation
of microgamete nuclei in the formerly
hypothetical microgamonts. Second, both molecular
phylogenies and morphological data indicate
that the blastogregarines definitely belong to the
Apicomplexa and, more exactly, to the sporozoans.
Third, molecular phylogenetic analyses fail
to assign blastogregarines firmly either to the gregarines
or coccidiomorphs (combined coccidians
and hematozoans): these “best” SSU and LSU
rDNA phylogenies conflict with each other and,
additionally, are weakly supported in both cases;
the morphological data reveal a plesiomorphic
status blastogregarines as the relatively isolated
group with the mixed features of coccidians and
plesiomorphic gregarines. Proceeding from the
aforesaid we classify the blastogregarines as a
separate class Blastogregarinea (also see: de
Puytorac et al. 1987) within the phylum Apicomplexa,
subphylum Sporozoa. This class includes
a single order Sidleckiida comprising two families,
which separation from each other is based
on the conspicuous differences in the structure
of the attachment apparatus in S. cf. nematoides
and C. mesnili manifested in the loss of the apical
complex, change of the cell junction type,
and development of additional attachment devices
(alveolar bulge(s)) in the latter species. These differences
are comparable with those between archi-
Blastogregarines: Plesiomorphic Apicomplexa 721
and eugregarines possessing mucron or epimerite,
respectively, and this is in use as a relevant
taxonomic criterion for the separation of the aforementioned
orders within the genuine gregarines
(Schrével and Desportes 2013a,b, 2015; Schrével
et al. 2013; Simdyanov et al. 2017). Applying
this morphological approach to the blastogregarines,
however in the “limited mode” because
of the sparsity of the group, we establish the
new genus Chattonaria for Siedleckia mesnili (in
honour of Édouard Chatton, who described this
species and contributed significantly to the field
of blastogregarine research), as well as the new
family Chattonariidae, which, together with the
family Siedleckiidae, compose the order Blastogregarinida,
single in the class. We expect future
increasing the species composition of the family
Chattonariidae due to the addition of other
named blastogregarine species as, e.g., Siedleckia
caulleryi and the poorly described S. dogieli.
Presumably, these organisms with longitudinal striations
and complex attachment apparatus (as far
as it may be inferred from light-microscopic data)
can be members of the same genus Chattonaria.
The composition of the family Siedleckiidae is
expected to increase rather due to future recognizing
cryptic species in the morphotype S.
nematoides (see above). The consequent formal
taxonomical actions are stated in the summary
below.
Taxonomic Summary
Fixing the taxonomic position of the blastogregarines
we leave the taxonomic ranks of Apicomplexa
(phylum) and Sporozoa (subphylum) as
originally established by Levine (Levine 1970, 1985;
Levine et al. 1980) and update their diagnoses taking
into consideration recent evidence.
Phylum APICOMPLEXA Levine, 1970
Diagnosis. The apical complex initially fulfilling
myzocytosis and giving rise to longitudinal
subpellicular microtubules arranged in layer(s);
micropores.
Subphylum Sporozoa Leuckart, 1879 (Syn. Sporozoasida
Levine, 1985)
Diagnosis. Parasitic Apicomplexa largely with the
complex life cycle (Leuckart’s triade) resulting in the
oocysts performing sporogony and finally containing
sporozoites; reduced plastid (apicoplast).
Class Blastogregarinea Chatton et Villeneuve,
1936, emend.
Diagnosis. Sporozoa. Epicellular parasites with
permanent multinuclearity and gametogenesis:
nuclear divisions of merogony and gamogony proceed
within the same individual (merogamont)
throughout its lifespan. The merogamonts with
plesiomorphic ultrastructure: the well-developed
apical (mucronal) complex performing myzocytotic
feeding, regularly arranged longitudinal subpellicular
microtubules arising from the mucron. The
merogamonts are motile (bending), and sexually
differentiated: in female individuals, the nuclei lie in
a row along the cell axis, in male individuals they lie
linearly only in the anterior part, but are scattered
randomly in the posterior part of the cell. Oogamy
is characteristic: female gamogony is realized by
continuous budding of mononuclear macrogametocytes
or macrogametes from the posterior part of
female merogamonts, while male gamogony is realized
by budding of multinuclear microgametocytes
or microgametoblasts apparently followed by their
dissociation into small putatively biflagellated male
gametes with spermal nuclei. According to Chatton
and co-workers (1929, 1936a) oocysts with many
(10–16) free sporozoites (no sporocysts). Intestinal
parasites of the polychaetes Orbiniidae.
Order Siedleckiida nom. nov.
With the characteristics of the class.
Family Siedleckiidae Chatton et Villeneuve, 1936,
emend.
Diagnosis. With the characteristics of the order.
Merogamonts with smooth surface lacking any
grooves or folds. Mucron contains all components
of the apical complex and performs myzocytotic
feeding; apical ring(s) is likely MTOC of subpellicular
microtubules. Monotypic.
Type genus. Siedleckia Caullery et Mesnil, 1898,
emend.
Diagnosis. With the characteristics of the family.
Merogamonts elongate and flattened with pointed
anterior and rounded posterior ends. Monotypic,
although the complex of cryptic species is suggested
(molecular-phylogenetic data).
Type species. Siedleckia nematoides Caullery et
Mesnil, 1898
Family Chattonariidae, fam. nov.
Diagnosis. With the caracteristics of the order. The
mucron is modified: it performs myzocytotic feeding,
but the apical complex is significantly reduced:
722 T.G. Simdyanov et al.
it retains only mucronal vacuole and, probably, polar
ring, which is not connected with microtubules; no
conoid and rhoptries. Monotypic.
Type genus. Chattonaria, gen. nov.
Diagnosis. With the characteristics of the family.
Merogamonts elongate, cylindric, with roundly
pointed posterior end; the surface with large longitudinal
folds; mucron bearing voluminous alveoli
with protuberances of the cytoplasm.
Type species. Chattonaria mesnili (Chatton et
Dehorne, 1929), comb. nov. for Siedleckia mesnili
Chatton et Dehorne, 1929
Etymology. Named in honour of Édouard Chatton,
the eminent protistologist.
Note. Siedleckia caulleryi Chatton et Villeneuve,
1936 and Siedleckia dogieli Chatton et Dehorne,
1929 likely belong to the family Chattonariidae and
to the genus Chattonaria, judging by their general
morphology studied with LM. Ultrastructural and
molecular-phylogenetic studies are required for a
more precise conclusion. The species S. dogieli
was established relying solely on a draft drawing
by Dogiel with the indication of the host (Chatton
and Dehorne 1929), i.e. it does not have a valid
description and should be considered a nomen
nudum.
Methods
Sampling: The hosts of S. cf. nematoides were collected
from two littoral sites located in the Kandalaksha Gulf of the
White Sea, Russia: on Bolshoy Gorely Island (66◦
18 46 N,
33◦
37 40 E) near Marine Biological Station of Saint Petersburg
State University (MBS) and from the coast of Velikaya Salma
Straight (66◦
33 11 N, 33◦
06 33 E) near White Sea Biological
Station of Moscow State University (WSBS). The hosts of Chattonaria
mesnili were collected on the lowest littoral zone of a
sandy beach (48◦
41 23 N, 4◦
04 17 W) near Moguériec, coastal
zone of English Channel, France.
The blastogregarine individuals were isolated by tearing
apart the intestine of the hosts with fine tip needles under a
stereomicroscope (MBS-1 or MBS-10 (LOMO, Russia) for S. cf.
nematoides, or Olympus SZ40 (Olympus, Japan) for C. mesnili).
The released parasites and small fragments of the host
gut with the attached individuals were rinsed three times in
filtered seawater using fine glass pipettes and then prepared
for light microscopy (S. cf. nematoides only), scanning electron
microscopy, transmission electron microscopy, and further DNA
extraction.
Light microscopy: After rinsing in seawater, the living parasites
were examined under light microscopes Leica DM2500,
and Leica DM5000B (Leica Microsystems, Germany). Digital
images of the living S. cf. nematoides were acquired under an
MBR-1 microscope (LOMO, Russia) in phase-contrast mode
with a Canon EOS 300D camera (Canon, Japan). Also wet
smears of small pieces of the host intestine content were fixed
by Bouin’s fluid, stained by Böhmer’s hematoxylin, and examined
under a light microscope Zeiss AxioImager A1 with a digital
camera AxioCam MRc5 (Carl Zeiss, Germany).
Electron microscopy: For SEM study, individual blastogregarines
and small fragments of the host gut with attached
blastogregarines were fixed with 2.5% (v/v) glutaraldehyde
in 0.05 M cacodylate buffer (pH = 7.4) containing NaCl 1.28%
(w/v) (for White Sea samples) or 2.3% (w/v) (for English
Channel samples): two replacements of the fixative for 1 h
each, in the ice bath in the dark, then rinsed three times
(20 min each) with filtered seawater, and post-fixed with 2%
(w/v) OsO4 in the same buffer (room temperature, 2 h).
Fragments of dissected gut containing the parasites were
dehydrated in a graded series of ethanol up to 96% (v/v),
transferred to a 96% ethanol/pure acetone mixture (1:1,
v/v), rinsed three times with pure acetone, and critical point
dried with CO2. Alternatively, after 96% ethanol, the samples
were rinsed three times with 100% ethanol and then
critical point dried with CO2. The samples were mounted on
stubs, sputter coated with gold/palladium, and examined under
a LEO-420 scanning electron microscope (Carl Zeiss, Germany)
or a JSM-6380LA scanning electron microscope (JEOL,
Japan).
The same fixation protocol as for SEM was used for the
majority of TEM samples. To visualize the cell coat, some
specimens of S. cf. nematoides were fixed with the mixture
of glutaraldehyde and ruthenium red (3% (v/v) and 0.05%
(w/v), respectively) in 0.2 M cacodylate buffer (pH = 7.4) and
post-fixed with the mixture of OsO4 and ruthenium red (1%
(v/v) and 0.05% (w/v), respectively) in the same buffer – all
under the same conditions as described above (Luft 1971a,b).
After dehydration through ascending series of ethanol, the
fixed samples were transferred into embedding mediums (Epon
(Sigma-Aldrich, USA) or Spurr (Ted Pella, USA)) according
to manufacturer’s protocols, using acetone or isopropanol as
intermediate dehydrating reagents for Epon and Spurr, respectively.
Ultrathin sections (40 to 50 nm) were obtained using an
LKB-III (LKB, Sweden), Reichert-Jung Ultracut E (C. Reichert,
Austria) or Leica EM UC6 (Leica Microsystems, Germany)
ultramicrotomes and then contrasted with uranyl acetate and
lead citrate (Reynolds 1963) and examined under a JEM-
100B, JEM-1010 or JEM-1011 electron microscopes (JEOL,
Japan).
DNA isolation, PCR, cloning, and sequencing: After rinsing
in seawater, blastogregarine individuals were collected
into 1.5-ml microcentrifuge tubes: ∼10 individuals of S. cf.
nematoides from MBS (2003), ∼100 individuals of S. cf. nematoides
from WSBS (2004), and ∼25 individuals C. mesnili
(2010). For S. cf. nematoides, the material was lysed by
an alkaline procedure (Floyd et al. 2002), and directly used
for PCR amplification. For C. mesnili, the material was preserved
in the “RNAlater” reagent (Life Technologies, USA),
then stored at −20 ◦
C until DNA extraction. The DNA extraction
was performed using the “Diatom DNA Prep 200” kit (Isogen
Laboratory, Russia). The resulting ribosomal DNA (rDNA)
sequences were deduced from a combination of shorter fragments
individually amplified using different pairs of primers
(Table 1) and represented 18S or small subunit (SSU), 5.8S,
28S or large subunit (LSU) rDNAs, and internal transcribed
spacers 1 and 2 (ITS 1 and ITS 2, respectively). All fragments
were PCR amplified with an Encyclo PCR kit (Evrogen,
Russia) in 25 ␮L of the reaction mixture prepared according
to the manufacturer’s protocol and contained 1 ␮L of the
DNA extract using a DNA Engine Dyad thermocycler (Bio-Rad
Laboratories, USA) and the following protocol: initial denatu-
Blastogregarines: Plesiomorphic Apicomplexa 723
ration at 95 ◦
C for 3 min, 40 cycles of 95 ◦
C for 30 s, 48 ◦
C
or 53 ◦
C (see Table 1) for 30 s, and 72 ◦
C for 1.5 min, and a
final extension at 72 ◦
C for 10 min. The PCR products of the
expected size were cut from the gel and extracted by using
a Cytokine DNA isolation kit (Cytokine, Russia). The PCR
products were directly sequenced for fragments obtained from
C. mesnili and S. cf. nematoides from MBS. The fragments
obtained from S. cf. nematoides from WSBS were heterogeneous
and therefore were cloned by using InsTAclone PCR
Cloning Kit (Fermentas, Lithuania). Sequencing was performed
using an ABI PRISM BigDye Terminator v. 3.1 reagent kit on
an automatic sequencer Applied Biosystems 3730 DNA Analyzer
(Applied Biosystems, USA). All sequences were tested
with BLAST in order to detect the matches for apicomplexans
and retain them for further analyses. For the sample of S.
cf. nematoides from WSBS, the contiguous sequence of the
near-completed ribosomal operon (SSU rDNA + ITS1 + 5.8S
rDNA + ITS2 + LSU rDNA) was assembled from 5 overlapping
PCR-amplified and cloned fragments, whereas only partial
sequence of SSU rDNA alone was obtained from the sample
of S. cf. nematoides from MBS. For the sample of C. mesnili,
the assembled contiguous sequence (from 3 overlapped
PCR-amplified fragments) covered near-complete gene of SSU
rRNA, ITS1, 5.8S rDNA, ITS2, and a large part (∼2,000 bp) of
LSU rDNA (Table 1). The contiguous sequences were built with
the use of BioEdit 7.0.9.0 (Hall 1999). The overlapped regions
(180–750 sites, see Table 1) revealed 100% of matches. All
obtained sequences were deposited in NCBI GenBank (accession
numbers: MH061197-9).
Phylogenetic analyses: Three alignments were prepared
for phylogenetic analyses: SSU rDNA (110 sequences, 1,550
sites), LSU rDNA (54 sequences, 2,913 sites), and the ribosomal
operon (concatenated SSU, 5.8S, and LSU rDNAs: 54
sequencess, 4,618 sites). The alignments were generated in
MUSCLE 3.6 (Edgar 2004) under default parameters and then
manually adjusted with BioEdit 7.0.9.0 (Hall 1999); columns
containing few nucleotides and hypervariable regions were
removed. The taxon sampling of SSU rDNA alignment was
designed in order to maximize the phylogenetic diversity and
completeness of sequences in alignments, by preferentially
selecting taxa having their SSU and LSU rDNA both sequenced.
Representatives of heterokonts and rhizarians were used as
outgroups.
The final alignment of SSU rDNA included 110 representative
sequences (1,550 sites). To assess similarities among
the SSU rDNA sequences within the blastogregarine clade, we
calculated the percentage of identities as it is implemented in
NCBI BLAST: the ratio of the matching sites to the total amount
of unambiguous sites in the overlapping regions of each pair
of aligned sequences: b/a × 100%, where a = total number of
the aligned unambiguous sites, b = number of matches between
them.
For the LSU rDNA and ribosomal operon (concatenated
SSU, 5.8S and LSU rDNA sequences) analyses, the taxon
sampling of only 54 sequences was used due to the limited
availability of data for LSU rDNA, and, especially, 5.8S rDNA.
Therefore, the 5.8S rDNA (155 sites in the alignment) was
not represented in the analysis of concatenated rDNA genes
for seven sequences (Chromera velia, Colponema vietnamica,
Goussia desseri, Stentor coeruleus, and 3 environmental
sequences: Ma131 1A38, Ma131 1A45, and Ma131 1A49):
the corresponding positions were replaced with “N” in the
alignment. The resulting multiple alignments contained 54
sequences (2,913 sites) for the LSU rDNA, and the same 54
sequences (4,618 sites) for the concatenated rDNAs (ribosomal
operon). Thus, both taxon sampling comprised an identical set
of species, all of which were also represented in the alignment
of the 110 SSU rDNA sequences.
Maximum-likelihood (ML) analyses were performed with the
RAxML 8.2.9 program (Stamatakis 2006) under the GTR +
model and CAT approximation (25 rate categories per site).
The procedure included 100 alternative runs of the ML analysis
and 1,000 replicates of multiparametric bootstrap. Bayesian
inference (BI) analyses were conducted using MrBayes 3.2.6
program (Ronquist et al. 2012) under GTR + + I model with
8 discrete categories of gamma distribution. The program
was set to operate using the following parameters: nst = 6,
ngammacat = 8, rates = invgamma, covarion = yes; parameters
of Metropolis Coupling Markov Chains Monte Carlo (mcmc):
nchains = 4, nruns = 4, temp = 0.2, ngen = 7,000,000, samplefreq
= 1,000, burninfrac = 0.5 (first 50% of 7,000 sampled trees,
i.e. first 3,500 generations were discarded in each run). The
following average standard deviations of split frequencies were
reached at the end of calculations: 0.009432 for the SSU rDNA
analysis, 0.002976 for the LSU rDNA analysis, and 0.001371
for the ribosomal operon analysis.
Alternative tree topologies were manually created and edited
using TreeView 1.6.6 program (Page 1996). The reference tree
topology of SSU rDNA phylogeny (110-taxon dataset) was
copied from the Bayesian tree (Fig. 10) that appeared more
accurate in point of branching order within coccidiomorphs’
clade (coccidians and hematozoans), but was identical with
the corresponding ML tree in other respects. The reference
topologies of LSU rDNA and ribosomal operons (54-taxon
datasets) were copied from the trees showed in Figure 11;
their ML and Bayesian phylogenies were identical. Alternative
topologies were constructed by positioning the blastogregarines
as a sister group successively to the major sporozoan
clades, which were picked out as either high-resolved (high
statistical supports) molecular phylogenetic lineages in the
reference trees and published phylogenies or, in respect of
low-resolved gregarines in SSU rDNA phylogenies, additionally
relying on relevant morphological evidence (Simdyanov
et al. 2017). As a result, we examined the relations of the
blastogregarines to coccidiomorphs, cryptosporidians, different
gegarine lineages, combined gregarine-cryptosporidian
clade and sporozoa as a whole. Topology tests were performed
with TREE-PUZZLE 5.3.rc16 and CONSEL 0.1j
programs (Schmidt et al. 2002; Shimodaira and Hasegawa
2001). The following tests were used: Bootstrap Probability
(Felsenstein 1985), Expected-Likelihood Weights (Strimmer
and Rambaut 2002), Kishino-Hasegawa test (Kishino and
Hasegawa 1989), Shimodaira-Hasegawa test (Shimodaira
and Hasegawa 1999), Weighted Shimodaira-Hasegawa test
(Shimodaira and Hasegawa 1999), and approximately unbiased
test (Shimodaira 2002).
Acknowledgements
The authors thank the staff of the White Sea
Biological Station of Lomonosov Moscow State
University (WSBS MSU) and the Marine Biological
Station of Saint Petersburg State University
(MBS SPbSU) for providing facilities for field
sampling and material processing. Light microscopic
studies were conducted using equipment
of the Center of microscopy WSBS MSU, Dept.
of Invertebrate zoology of MSU, and Dept. of
724 T.G. Simdyanov et al.
Invertebrate zoology of SPbSU. Electron microscopic
studies were performed in the Electron
microscopy laboratory of the Faculty of Biology,
Lomonosov Moscow State University (Russia),
Center of Electron Miscroscopy of Papanin Institute
for Biology of Inland Waters, Russian Academy of
Sciences, Electron microscopy laboratory of Dresden
Technical University (Germany), Research
Recourse Centers of St Petersburg State University
(“Molecular and Cell Technology” and “Environmental
Safety Observatory”), and in Laboratory
of Electron Microscopy of Institute of Parasitology,
Academy of Sciences of the Czech Republic,
ˇCeské Budˇejovice. Authors are deeply grateful to
the teams of these labs for technical support and
help. This study used CYPRES Science Gateway
(Miller et al. 2010) and the Supercomputer
Center of Lomonosov Moscow State University
(http://parallel.ru/cluster) to make phylogenetic
computations. DNA sequencing was performed at
the DNA sequencing center “Genome” (Engelhardt
Institute of Molecular Biology, Russian Academy of
Sciences, www.genome-centre.ru).
We would like to thank Kirill V. Mikhailov
(Belozersky Institute of Physico-Chemical Biology,
Lomonosov Moscow State University) for the
assistance in several experiments and for his
help in the calculations of sequence identities,
Dr. Anna Zhadan (White Sea Biological Station)
for her expert commentary on Scoloplos spp.
biodiversity, and Dr. Nikolay Mugue (Institute of
Developmental Biology, Russian Academy of Sciences)
for his help in sequencing and valuable
comments.
This work was supported by the Russian
Foundation for Basic Research (projects No. 15-
29-02601 and 18-04-00324), ECO-NET project
2131QM (Égide, France), the French governmental
ANR Agency under ANR-10-LABX-0003
BCDiv, ANR-11-IDEX-0004-02, and ANR HAPAR
(ANR-14-CE02-0007), the Interdisciplinary Program
of the MNHN (ATM-Emergence des clades,
des biotes et des cultures), and the Czech Science
Foundation, project No. GBP505/12/G112
(ECIP). The phylogenetic analyses of SSU rDNA
presented in this study were supported by the
Russian Science Foundation, project No. 14-50-
00029.
Appendix A. Supplementary Data
Supplementary data associated with this article
can be found, in the online version, at
https://doi.org/10.1016/j.protis.2018.04.006.
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Paskerova G.G., Miroliubova T.S., Diakin A., Kováčiková M.,
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Fine structure and molecular phylogenetic position of two marine
gregarines, Selenidium pygospionis sp. n. and S. pherusae sp. n.,
with notes on the phylogeny of Archigregarinida (Apicomplexa)
Protist 169(6), 826-852
Protist, Vol. 169, 826–852, December 2018
http://www.elsevier.de/protis
Published online date 28 June 2018
ORIGINAL PAPER
Fine structure and Molecular
Phylogenetic Position of Two Marine
Gregarines, Selenidium pygospionis sp.
n. and S. pherusae sp. n., with Notes on
the Phylogeny of Archigregarinida
(Apicomplexa)
Gita G. Paskerovaa,1
, Tatiana S. Miroliubovaa
, Andrei Diakinb
,
Magdaléna Kováˇcikováb
, Andrea Valigurováb
, Laure Guillouc
,
Vladimir V. Aleoshind
, and Timur G. Simdyanove
aDepartment of Invertebrate Zoology, Faculty of Biology, Saint Petersburg State University,
Universitetskaya emb. 7/9, St Petersburg 199 034, Russian Federation
bDepartment of Botany and Zoology, Faculty of Science, Masaryk University, Kotláˇrská 2,
611 37 Brno, Czech Republic
cSorbonne Université, Université Pierre et Marie Curie – Paris 6, CNRS, UMR 7144,
Station Biologique de Roscoff, Place Georges Teissier, CS90074, 29688 Roscoff Cedex,
France
dBelozersky Institute for Physico-Chemical Biology, Lomonosov Moscow State University,
Leninskie Gory, Moscow 119 234, Russian Federation
eDepartment of Invertebrate Zoology, Faculty of Biology, Lomonosov Moscow State
University, Leninskie Gory, Moscow 119 234, Russian Federation
Submitted October 30, 2017; Accepted June 19, 2018
Monitoring Editor: Frank Seeber
Archigregarines are a key group for understanding the early evolution of Apicomplexa. Here we report
morphological, ultrastructural, and molecular phylogenetic evidence from two archigregarine species:
Selenidium pygospionis sp. n. and S. pherusae sp. n. They exhibited typical features of archigregarines.
Additionally, an axial row of vacuoles of a presumably nutrient distribution system was revealed in S.
pygospionis. Intracellular stages of S. pygospionis found in the host intestinal epithelium may point
to the initial intracellular localization in the course of parasite development. Available archigregarine
SSU (18S) rDNA sequences formed four major lineages fitting the taxonomical affiliations of their
hosts, but not the morphological or biological features used for the taxonomical revision by Levine
(1971). Consequently, the genus Selenidioides Levine, 1971 should be abolished. The branching order
of these lineages was unresolved; topology tests rejected neither para- nor monophyly of archigregarines.
We provided phylogenies based on LSU (28S) rDNA and near-complete ribosomal operon
1
Corresponding author; fax +7 812 3289703
e-mail gitapasker@yahoo.com, g.paskerova@spbu.ru (G.G. Paskerova).
https://doi.org/10.1016/j.protis.2018.06.004
1434-4610/© 2018 Elsevier GmbH. All rights reserved.
Selenidium spp. n.: Structure and Phylogeny 827
(concatenated SSU, 5.8S, LSU rDNAs) sequences including S. pygospionis sequences. Although being
preliminary, they nevertheless revealed the monophyly of gregarines previously challenged by many
molecular phylogenetic studies. Despite their molecular-phylogenetic heterogeneity, archigregarines
exhibit an extremely conservative plesiomorphic structure; their ultrastructural key features appear to
be symplesiomorphies rather than synapomorphies.
Key words: Unicellular parasites; polychaetes; ultrastructure; 18S rDNA; 28S rDNA; molecular phylogeny.
© 2018 Elsevier GmbH. All rights reserved.
Introduction
Apicomplexa is a large group of unicellular parasites
infecting a wide range of invertebrate and
vertebrate hosts. Some apicomplexans, such as
the human pathogens Plasmodium, Toxoplasma,
and Cryptosporidium, are well studied. At the
same time, basal apicomplexans, archigregarines,
agamococcidia, blastogregarines, and protococcidia
inhabiting exclusively marine invertebrate
hosts and being crucial for our understanding of
the evolution of parasitism and evolutionary paths
of apicomplexans in general, are still poorly inves-
tigated.
Archigregarines (Archigregarinida Grassé, 1953,
Apicomplexa Levine, 1970) are unicellular parasites
inhabiting marine invertebrates, mostly
polychaetes. They are thought to have retained
a number of plesiomorphic characteristics from
the most recent ancestor of all apicomplexans
(Cavalier-Smith and Chao 2004; Cox 1994;
Desportes and Schrével 2013; Grassé 1953;
Leander 2008a; Leander and Keeling 2003). The
most often encountered stage of their life cycle
is a trophozoite. It is a relatively large cell usually
attached to the host cell by the mucron, an
attachment apparatus with organelles of the apical
complex typical of the invasive stages (zoites).
The mucron participates in myzocytosis, feeding by
sucking out the cytoplasmic contents of the host cell
into food vacuoles (Cavalier-Smith and Chao 2004;
Desportes and Schrével 2013; Schrével 1968;
Schrével et al. 2016; Simdyanov and Kuvardina
2007; Wakeman and Horiguchi 2018; Wakeman
et al. 2014). The pellicle of archigregarines is organized
as a three-layered membrane complex. It
is supported by microtubules arranged in one or
more subpellicular layers (Desportes and Schrével
2013).
The life cycle of archigregarines, a sequence
of gamogony (gamete production) and sporogony
(zygote fission for producing sporozoites),
is often thought to include asexual cell multiplication
in the trophozoite stage – merogony (Adl
et al. 2012; Desportes and Schrével 2013; Grassé
1953). Levine (1971) and his followers placed great
importance on the presence/absence of merogony
in the life cycle for the classification of
archigregarines, transferring species without merogony
to eugregarines (Levine 1971, 1985; Perkins
et al. 2000). On the contrary, Schrével with coauthors
considered merogony to be non-important
for the high-level classification of archigregarines
(Desportes and Schrével 2013; Schrével 1970,
1971a,b; Schrével et al. 2016). This point of view
was shared by several contemporaneous authors
(Kuvardina and Simdyanov 2002; Leander 2006,
2007; Rueckert and Leander 2009; Rueckert and
Horák 2017; Simdyanov and Kuvardina 2007). It
should be noted that the absence of merogony is
difficult to prove. In this context, taxa delineation
should be based on the presence of the easiest
observable stage (trophozoites) and morphological
characteristics. At present, taxonomy and determination
of basic taxonomic characters are routinely
determined by electron microscopy and molecular
phylogenies.
SSU rDNA-based phylogenetic trees obtained
recently are in good accordance with the interpretation
that archigregarines are a paraphyletic
stem group from which other gregarine lineages
evolved (Cavalier-Smith 2014; Cavalier-Smith and
Chao 2004; Grassé 1953; Rueckert and Leander
2009; Rueckert and Horák 2017; Schrével et al.
2016; Wakeman and Horiguchi 2018; Wakeman
and Leander 2012, 2013; Wakeman et al. 2014).
To date, there are more than 70 species of archigregarines.
Most of them belong to the genus
Selenidium (Desportes and Schrével 2013; Levine
1971; Rueckert and Horák 2017; Rueckert and
Leander 2009; Wakeman and Horiguchi 2018;
Wakeman and Leander 2012, 2013; Wakeman
et al. 2014; WoRMS 2018).
Despite their significant molecular-phylogenetic
heterogeneity, species of Selenidium possess a
similar and extremely conservative morphology.
It is represented by the morphostasis, a set of
characters typical of the invasive stages (zoites)
828 G.G. Paskerova et al.
Figure 1. General morphology and motility of Selenidium pygospionis sp. n. Differential interference contrast
(DIC) and phase-contrast (PH) light micrographs. A. Attached large and detached small trophozoites.
In the large trophozoite, note the axial streak along the longitudinal cell axis and radial threads running
from the axial streak towards the cell periphery. DIC. B. Young trophozoite showing bending motility; the
composition of two micrographs of the same cell. DIC. C. Large trophozoite lying on one of the narrow
sides and slightly pressed with the coverslip. Note the axial streak. PH. D–E. Large trophozoite lying on
one of the flattened sides and pressed with the coverslip; D and E—different optical sections of the same
cell. The axial streak is not visible. DIC. F–I. Medium-sized trophozoite lying on one of the narrow sides;
Selenidium spp. n.: Structure and Phylogeny 829
of parasitic apicomplexans (Leander and Keeling,
2003). Additionally, molecular phylogenetic studies
have repeatedly demonstrated a host-parasite
coevolution when closely related archigregarines
parasitize closely related hosts (Desportes and
Schrével 2013; Rueckert and Horák 2017; Schrével
et al. 2016; Wakeman and Leander 2013). Whether
these two observations are linked is unknown.
In this study, we report the discovery of two
new archigregarine species, Selenidium pygospionis
sp. n. isolated from spionid polychaetes
Pygospio elegans and Polydora glycymerica, and
S. pherusae sp. n. isolated from flabelligerid polychaetes
Pherusa plumosa. We examined the new
species using light and electron microscopy, conducted
phylogenetic analyses based on the SSU
rDNA and obtained the first LSU rDNA sequences
of archigregarines.
Results
Selenidium pygospionis sp. n.
Occurrence
The gregarine Selenidium pygospionis sp. n. was
found in the intestine of the polychaete Pygospio
elegans (Spionidae) collected at the silty-sand
intertidal zone of the White Sea. There were 109
infected polychaetes out of the 302 dissected.
The intensity of infection usually varied from 1 to
50 (mode = 1, average = 9.8) gregarines per host;
in two cases, the number of parasites reached
100 and 150 cells per host. Both attached and
non-attached trophozoites of different sizes were
found in the host intestine (Fig. 1A). Syzygies were
extremely rare (a few in all dissected polychaetes).
In squash preparations (see Methods) of more than
100 examined polychaetes, no other stages of the
life cycle (gametocysts or stages of merogony)
were observed.
We also found very similar parasites in the
intestine of the shell-boring polychaete Polydora
glycymerica (Spionidae) inhabiting the bivalve
Glycymeris yessoensis from the Sea of Japan.
The intensity of infection was about 30 parasites
per host.
Parasites isolated from polychaetes of both
species were identical in their morphology, fine
structure, and DNA sequences. Therefore, we considered
them to belong to the same species.
Further description of S. pygospionis was predominantly
based on the evidence obtained from the
White Sea samples as the most representative
ones.
General Morphology
As shown with light microscopy (LM) and scanning
electron microscopy (SEM), trophozoites S. pygospionis
were anchored in the host tissue with their
anterior end (Figs 1 A, 2 A, B). The parasites were
easy to dislodge from the intestinal epithelium during
the dissection of the hosts.
The smallest trophozoites, presumably young
trophozoites not long after their transformation from
zoites (most likely, sporozoites), were observed
occasionally. They were spindle-shaped with a
pointed anterior end and a rounded posterior end.
Their length varied from 15 to 23 ␮m (n = 2), and
their width, from 6 to 7 ␮m (n = 2) in the middle part
of the cell. A single rounded nucleus was located
in the anterior half of the cell. The parasites could
bend slightly in one plane but never glided (Fig. 1B).
Well-developed trophozoites were elongated,
vermiform and slightly flattened (Figs 1 C–M, 2 C,
D). Their length varied from 34 to 288 ␮m (average
144 ␮m, mode 146 ␮m, n = 79); their maximum
width (4–25 ␮m, average 12 ␮m, mode 11 ␮m,
n = 76) was in the middle of the cell where an oval
nucleus [6–22 ␮m (av. 17 ␮m, n = 40) × 5–11 ␮m
(av. 8.4 ␮m, n = 26)] was located. The nucleus
was elongated along the longitudinal axis of the
cell (Fig. 1A, C, D). A single spherical nucleolus
(3.1–6.3 ␮m, n = 6) was commonly situated at the
anterior pole of the nucleus (Fig. 1D). The anterior
end of the parasites was usually hook-like, bent in
the median plane (the plane perpendicular to the
flattened sides) towards one of the flattened sides
of the cell (Figs 1 C–E, 2 C, D). The mucron was
dome-shaped with a smooth surface. In some individuals,
it had a small pit in the center (Fig. 2C).
The posterior end was rounded (Fig. 2D). The entire
surface bore 22–30 (n = 12), usually 28, broad and
low folds separated by grooves (Figs 1 E, 2 B–E).
a series of micrographs illustrating bending motility (1–2 bends along the cell). DIC. J–M. Large trophozoite
lying on one of the narrow sides; a series of micrographs illustrating bending motility (2–3 bends along the cell).
Note the coiling of the anterior end (K). DIC. N–Q. Syzygy; a series of micrographs illustrating bending motility
(up to 4 bends along the cell) and the coiling of the anterior end. The axial streak is not visible. DIC. *, anterior
end; ax, axial streak; f, folds; N, nucleus; n, nucleolus; r, radial threads.
830 G.G. Paskerova et al.
Figure 2. Surface morphology of Selenidium pygospionis sp. n. Scanning electron micrographs. A. Trophozoite
attached to a fragment of the host intestinal epithelium by its anterior end. B. Attached trophozoite with three
bends of its cell. Note small transversal compression folds of the parasite’s pellicle at the inner surface of each
bend. Inset. Transversal compression folds at high magnification. C–D. Detached trophozoites lying on one
of the narrow sides (C) and one of the wide sides (D) of their cells. Note a hook-bent anterior end with a
smooth, dome-shaped mucron, and folds at the cell surface. The arrowhead points to the pit at the center of the
mucron in C. E. Micrograph of the trophozoite surface showing the number and location of micropores (mp).
F. Micropore at high magnification. cf, transversal compression folds; f, folds; h, host intestinal epithelial tissue;
mu, mucron; mp, micropores; p, parasite.
Selenidium spp. n.: Structure and Phylogeny 831
The width of the folds varied from 0.5 to 1.5 ␮m (av.
0.9, n = 21) in archigregarines fixed according to different
protocols (Fig. 2D, E). Numerous micropores
(10–20 per 50 ␮m2
, n = 2) were observed at the bottom
of the grooves; their edges were slightly raised
above the cell surface (Fig. 2E, F). The outer and
the inner diameter of the micropores was 134–287
and 38.5–74.6 nm (n = 6), respectively.
LM studies showed that well-developed trophozoites
had an intracellular axial streak of optically
distinct cytoplasm extending from the anterior end
to the posterior end and forming an expansion
around the nucleus. Numerous radial threads ran
from the axial streak towards the cell periphery. The
axial streak was easy to observe in trophozoites
lying on one of the narrow sides (Fig. 1A, C, H,
J–M).
Both attached and non-attached cells performed
very active bending movements in the median
plane (Fig. 1A, C, F–M, and Supplementary
Material Video S1). Non-attached archigregarines
usually moved non-progressively on one of their
narrow sides along the substrate. Medium-sized
trophozoites usually formed 1–2 bending sections
along the cell, while large-sized ones combined 2–4
bends with coiling of their anterior end (Fig. 1F–I vs
J–Q). Bending and coiling of the cell always started
at the hook-like anterior end. Fixed archigregarines
usually retained the bends of their body resulting
from their motility (Fig. 2A–D). There were small
transversal compression folds of the pellicle at the
inner surface of each bend (Fig. 2B, C). In addition,
the trophozoites of S. pygospionis never shortened
along their anterior-posterior axis.
Syzygies were caudal when two gamonts coupled
with their posterior ends. The syzygy partners
moved asynchronously (Fig. 1N–Q, Supplementary
Material Video S2).
Fine Structure
Transmission electron microscopic (TEM) studies
showed that the trophozoite tegument was represented
by a trimembrane pellicle [the plasma
membrane and the inner membrane complex
(IMC)] (Fig. 3 A). The IMC consisted of the external
and the internal cortical cytomembrane separated
from each other by an electron-transparent space
(8–10 nm thick), while the plasma membrane was
separated from the IMC by a space of higher electron
density (12–14 nm thick). The thickness of the
pellicle varied from 36 to 40 nm. The cell coat (glycocalyx)
covering the parasite surface was poorly
visible. The pellicle was underlain by longitudinally
oriented subpellicular microtubules (Fig. 3A–D).
They were arranged in a single layer, the continuity
of which was interrupted under the grooves
where micropores were usually located. Additionally,
groups of irregularly arranged microtubules
were present under the main layer. Each microtubule
was surrounded by an electron-transparent
sheath of the cell cytoplasm (Fig. 3A). Micropores
were present as invaginations of the plasma membrane.
Each invagination was surrounded with a
thick cylindrical structure, formed by the cytomembranes
of the IMC, and some electron-dense
substance (Fig. 3A, inset). Numerous vesicles with
some multi-membranous whorls or dense material
inside were incorporated in the microtubule layer
and the inner membrane complex of the pellicle;
they were connected with the plasma membrane
(Fig. 3C, D).
It could be seen in ultrathin sections of
trophozoite-infected intestines that the anterior end
of S. pygospionis was inserted between folds of
the host intestinal epithelium (Fig. 3E, F). There
was no direct contact between the host cell and
the attached parasite in any of the examined cases.
Several rhoptries and numerous groups of putative
micronemes were present in the cytoplasm of the
anterior end (Fig. 3E). The basal part of the conoid,
several ducts of rhoptries within the conoid, and
a large mucronal vacuole containing some loose
fibrillar material could be seen in some tangential
sections through the mucron (Fig. 3F).
The cytoplasm of the trophozoites was indistinctly
differentiated into two areas: the ectoplasm
and the endoplasm. The former was a narrow cortical
region containing subpellicular microtubules,
numerous mitochondria arranged at the peripheral
layer and vesicles with some multi-membranous
whorls or dense material under grooves. The endoplasm,
the rest of the cytoplasm, contained a
large nucleus, numerous amylopectin granules,
lipid droplets, some small electron-dense bodies,
and a few mitochondria (Fig. 3G, H). The distribution
of organelles and inclusions in the endoplasm
was irregular. Narrow, electron-light spaces without
any visible organelles could be seen around
the nucleus, along the cell axis and perpendicular
to it (Figs 3 G–I, 4 B, C). In addition, a series of
differently-sized vacuoles was arranged along the
cell axis in front and behind the nucleus in the endoplasm.
Some of them were linked to each other
by membrane tunnels. The content of these vacuoles
was mainly electron-transparent with a small
amount of some loose filamentous material (Fig. 4
A–C). Small vacuoles of similar appearance also
surrounded the nucleus alongside with an electrontransparent
area of the cytoplasm (Fig. 4B, inset).
832 G.G. Paskerova et al.
Figure 3. Fine structure of Selenidium pygospionis sp. n. Transmission electron micrographs. A. Transverse
section showing details of the cortex organization. Inset. Micropore at high magnification. B. Superficial
longitudinal section of a trophozoite showing the layer of longitudinal microtubules and the layer of mitochondria
under the pellicle. C–D. Transversal sections of the pellicle showing vesicles inserted in the microtubule layer
and the inner membrane complex. E. Superficial longitudinal section of the anterior end of a trophozoite inserted
Selenidium spp. n.: Structure and Phylogeny 833
As shown in a SEM study of de-paraffinized histological
sections of P. elegans, some gregarines
were deeply embedded in the intestinal epithelium
almost reaching the basal lamina (Fig. 4D–E).
TEM observations of ultrathin sections revealed
that some small single trophozoites (up to 40 ␮m
in length) were localized intracellularly within parasitophorous
vacuoles (Fig. 4F–G). The organization
of these individuals was generally identical to
that of well-developed trophozoites (Fig. 4F, G).
The internal space of the parasitophorous vacuole
was electron-transparent and filled with filamentous
material and electron-dense granules. The density
of their arrangement increased towards the periphery.
In some sections, there were agglomerations
of the electron-dense granules near the parasitophorous
vacuole membrane (Fig. 4G, inset).
Parasitophorous vacuoles were surrounded on
the outside by electron-dense fibrillar material,
membranes of endoplasmic reticulum, numerous
mitochondria, and cytoplasmic vesicles with
multilaminar inclusions, whereas the rest of the
host cytoplasm was electron-transparent with rare
organelles and vesicles (Fig. 4F–G).
Selenidium pherusae sp. n.
Occurrence
Five polychaetes Pherusa plumosa collected at the
sublittoral zone of the Sea of Japan were dissected.
All of them were infected with S. pherusae sp. n.
Trophozoites were all localized in the host midgut.
Up to several tens of trophozoites per host were
found in four of the worms, while the fifth harbored
more than 100 trophozoites. No other stages were
observed.
General Morphology
S. pherusae trophozoites were attached to the host
intestinal epithelium by their anterior end, but were
easy to dislodge during the dissection of the hosts.
The trophozoites were elongated, vermiform,
38–269 ␮m (n = 6) in length and 10–18 ␮m (n = 4)
in maximum width. The anterior end was narrowed
and slightly truncated (Fig. 5A, B), while the
posterior one was usually rounded in large individuals
(Fig. 5A, B) or pointed in small trophozoites
(Fig. 5C, D). A spherical nucleus (11–12 ␮m, in two
large trophozoites of 215 and 269 ␮m in length) was
located in the posterior half of the trophozoite, in the
widest part of the cell. It contained one nucleolus,
3–5 ␮m (Fig. 5A, B). Trophozoites had neither longitudinal
pellicular folds at the surface nor the axial
streak in the cytoplasm (Fig. 5A–C).
Both attached and non-attached gregarines
showed a bending motility of the entire cell, usually
one bend along the cell at a time. During bending,
transient transverse folds formed on the inner surface
of the bent part of the cell (Fig. 5A, B). Some
fixed gregarines were slightly helically twisted along
the longitudinal cell axis (Fig. 5D).
Fine Structure
The tegument of S. pherusae had almost the
same structure as that of S. pygospionis, but the
pellicle seemed smooth, without any grooves or
folds (Figs 5, 6 ). Its thickness varied from 33 to
44 nm. The plasma membrane and both cortical
cytomembranes were separated from each other by
electron-transparent spaces of a similar thickness
(7–11 nm). The cell coat was poorly visible (Fig. 6A,
C–E). Though the trophozoite surface seemed to
be smooth (Fig. 5C, D), micropores were sometimes
observed in transverse sections (Fig. 6D,
E). They were also similar in structure and size,
about 80 ␮m wide at the surface and 150 ␮m deep
(Fig. 6E). Although the subpellicular microtubules
were usually poorly preserved in the ultrathin sections
because of fixation artefacts, it could be seen
in some sections that they were organized in an
uninterrupted layer under the pellicle (Fig. 6C).
between the folds of the host intestinal epithelium. F. Oblique-transverse section of the mucron of a trophozoite
inserted between the folds of the host intestinal epithelium. Inset. Another section of the cell shown in F. G–H.
Transverse sections of the pre-nuclear (G) and post-nuclear (H) parts of trophozoites showing an indistinct
division of the cytoplasm into the ectoplasm at the periphery and the endoplasm occupying the remaining cell
volume. Note an electron-light space of the cytoplasm (pc) not containing any visible organelles. Total number
of folds in G is 25. I. Longitudinal section showing the nucleus. Note an electron-light area of the cytoplasm
around the nucleus. *, anterior end; a, amylopectin granules; c, conoid; db, dense bodies; ecto, ectocyte; em,
external cytomembrane; gmt, group of microtubules; gly, glycocalyx; h, host tissue; im, inner cytomembrane;
ld, lipid droplets; mic, micronemes; mit, mitochondria; mt, microtubules; mu, mucronal vacuole; N, nucleus; n,
nucleolus; p, pellicle; pc, electron-light area of the cytoplasm; pm, plasma membrane; rd, ducts of rhoptries; rh,
rhoptries; vac, vacuole; v, vesicles with multi-membrane whorls or dense material.
834 G.G. Paskerova et al.
Figure 4. Selenidium pygospionis sp. n.: organization of the cytoplasm, localization in the host epithelium,
intracellular stage. Transmission (TEM) and scanning (SEM) electron micrographs. A–C. Longitudinal thin
sections of the trophozoites in the regions where the nucleus is localized showing a series of connected
vacuoles along the cell axis. TEM. Inset in B. Electron-light area of the cytoplasm around the nucleus
with small vacuoles. TEM. D. Sagittal histological section of an infected host showing well-developed
trophozoites and the degree of their embedding in the host intestinal epithelium. SEM. E. Fragment of
D at higher magnification. SEM. F–G. Superficial oblique thin section (F) and nearly longitudinal thin
section (G, inset) showing intracellular stages of the trophozoite development. TEM. Inset in G. Agglomeration
of electron-dense granules in the parasitophorous vacuole and a vesicle with myelin-like structures in the
Selenidium spp. n.: Structure and Phylogeny 835
Figure 5. General morphology and motility of Selenidium pherusae sp. n. Light microscopic (LM) and scanning
electron (SEM) micrographs. A. Detached trophozoite showing bending motility; a composition of two micrographs
of the same cell. LM. B. Detached trophozoite fixed and stained with Carazzi’s hematoxylin. LM. C–D.
Differently sized trophozoites attached to the host intestinal epithelium. SEM. *, anterior end; cf, transversal
compression folds; N, nucleus; n, nucleolus.
A truncated and asymmetrical mucron of the
parasites formed an extended (up to 4 ␮m long)
zone of cell junction with the host cell (Fig. 6A, C).
This junction was represented by a gap 10–30 nm
wide. The polar ring, the conoid, several rhoptries
and micronemes were observed (Fig. 6A, B). The
conoid was a truncated hollow cone consisting of
6–7 spirally arranged microtubules. It measured
about 200 nm in height, 286 nm in apical diameter,
and 514 nm in basal diameter. The IMC terminated
near the apical end of the conoid, where there was
a polar ring, adjacently located to the IMC. Subhost
cell cytoplasm near the membrane of the parasitophorous vacuole. TEM. *, anterior end; arrowhead,
membrane tunnel between two vacuoles; bl, basal lamina of the host intestinal epithelium; epr, membranes
of the rough endoplasmic reticulum in the host cell; g, archigregarine; gr, electron-dense granules; h, host
intestinal epithelium; hc, host cell; ic, intestinal contents; N, nucleus; n, nucleolus; mit, host cell mitochondria; f,
fibrils in the host cell; pc, electron-light area of the cytoplasm; pv, parasitophorous vacuole; vac, vacuoles; vm,
vesicles with multilaminar inclusions.
836 G.G. Paskerova et al.
Figure 6. Fine structure of Selenidium pherusae sp. n. Transmission electron micrographs. A. Longitudinal
section of the anterior end of a trophozoite. B. Detail of A showing the mucron structure. Numerals indicate
the ordering numbers of the conoid microtubules. C. Superficial longitudinal section of the anterior end of
another trophozoite. D. Transversal section of the anterior third of the trophozoite showing its cytoplasmic
organization. E. Detail of D showing the micropore and pellicle structure. a, amylopectin granules; c, conoid; d,
Selenidium spp. n.: Structure and Phylogeny 837
pellicular microtubules arose from the polar ring
and ran along the cell (Fig. 6B, C). A voluminous
mucronal vacuole of irregular shape was present
in the basal part of the conoid (Fig. 6A). It contained
unidentified heterogeneous material and a
few vesicles with electron-translucent content. The
mucronal vacuole was surrounded by numerous
rhoptries and micronemes. The ducts of the rhoptries
extended to the apical pole of the parasite and
closely adjoined the mucronal vacuole. The content
of the ducts was less electron-dense than that of the
rhoptries (Fig. 6A–C).
Numerous putative micronemes and several
vesicles with an electron-dense material within
multi-membranous whorls were present in the cytoplasm
of the anterior third of the parasite (Fig. 6A).
Similarly to S. pygospionis, the entire cytoplasm
of S. pherusae trophozoites was indistinctly differentiated
into the ectoplasm and the endoplasm,
which had a similar content (Fig. 6D). In contrast
to S. pygospionis, only a few mitochondria
were observed near the pellicle, under subpellicular
microtubules (Fig. 6D).
Molecular Phylogeny
Characteristics of DNA sequences: The contiguous
sequence of S. pygospionis from Pygospio
elegans (White Sea) was generated from four
overlapping fragments and comprised SSU (small
subunit or 18S), 5.8S, LSU (large subunit or 28S)
rDNAs, and the internal transcribed spacers ITS 1
and 2 (4,910 bp totally). For S. pygospionis from
Polydora glycymerica (Sea of Japan), only the
near complete SSU rDNA sequence (1,610 bp) was
obtained; it was nearly identical with that from the
White Sea sample (only 2 substitutions). The contiguous
sequence of S. pherusae (2,551 bp) was
generated from two overlapping fragments and
comprised SSU, 5.8S, the first ∼600 nucleotides of
the LSU rDNA, and the internal transcribed spacers
ITS 1 and ITS 2 (Table 1 and Supplementary
Material Fig. S1).
Phylogenies inferred from SSU rDNA: Both
Bayesian inference (BI) and Maximum likelihood
(ML) analyses resulted in similar tree topologies
differing from each other by the position of platyproteids
(“squirmids”): they were the earliest branch of
Myzozoa in the Bayesian tree (Fig. 7), but the sister
group of Apicomplexa in the ML tree (data not
shown). Overall, the newly obtained phylogenies
matched recent molecular phylogenetic evidence
from alveolates and apicomplexans (e.g., CavalierSmith
2014; Janouˇskovec et al., 2015; Lepelletier
et al. 2014; Rueckert and Horák 2017; Schrével
et al. 2016). The resulting Bayesian tree inferred
from the dataset of 128 taxa and 1,550 sites
(Fig. 7) showed the monophyly of major alveolate
groups, although chiefly with moderate or low
support, especially in the ML analysis. The backbone
of the apicomplexan region in the obtained
trees was poorly resolved by both BI and ML
analyses. Within the sporozoans (parasitic apicomplexans),
the cryptosporidians were consistently
located as the sister group of the “short-branching”
eugregarine clade Eg1 (Actinocephaloidea and
Stylocephaloidea) in both analyses, although with
moderate Bayesian posterior probabilities (PP) and
low ML bootstrap percentage (BP) supports. The
top of the phylogenetic tree was formed by several
long branches of eugregarines grouping into
the loose clade Eg2 (Fig. 7); thus, eugregarines
and, consequently, gregarines in general were not
monophyletic, but polyphyletic.
Archigregarines branched after the cryptosporidians
+ Eg1 clade; they were not monophyletic either,
being split into four firmly supported major lineages
of greatly variable lengths (Ag1–Ag4), which arose
successively from the backbone of the phylogenetic
tree (Fig. 7). The earliest lineage Ag4 (PP = 0.99,
BP = 40%) encompassed a robust clade comprising
four parasites from polychaetes of the family Terebellidae
and its sister group (PP = 99, BP = 62%)
consisting of two environmental sequences. In contrast
to the results obtained by Rueckert and Horák
(2017), the archigregarine Selenidium fallax from
the cirratulid polychaete Cirriformia tentaculata represented
the isolated lineage Ag3 located not
as sister to the cryptosporidia + gregarines clade
but between the archigregarine clades Ag4 and
Ag2 + Ag1 in the resulting phylogenies from both
BI and ML analyses, though its nodal support
was weak (Fig. 7). The robust lineages Ag2 and
Ag1 formed a common clade, although weakly
supported. This clade was located as a sister to
the very weakly supported eugregarine clade Eg2
electron-dense droplet; ecto, ectocyte; em, external cytomembrane; epr, membranes of the rough endoplasmic
reticulum; gly, glycocalyx; hc, host cell; hp, host cell plasmalemma; im, inner cytomembrane; mic, micronemes;
mit, mitochondria; mp, micropore; mt, microtubules; mu, mucronal vacuole; pp, parasite plasmalemma; pr, polar
ring; rd, ducts of rhoptries; rh, rhoptries; v, vesicles with multi-membrane whorls or dense material.
838G.G.Paskerovaetal.
Table 1. Main characteristics of the archigregarine sequences obtained in this study.
Sample name, length of
resulting sequence and its
accession number
Amplified fragment Length Primers: forward (F) and reverse (R); annealing
temperature used in the PCRs
Selenidium pygospionis
from Pygospio elegans
(White Sea)
(I) SSU rDNA (part) ∼1,640 bp
(F)a 5 -GTATCTGGTTGATCCTGCCAGT-3
4,910 bp (R) 5 -GGAAACCTTGTTACGACTTCTC-3
MH061278 t◦ = 45 ◦C
(II) SSU rDNA (part), ITS1, 5.8S
rDNA, ITS2, LSU rDNA (part)
∼1,110 bp
(F) 5 -CCGTTCTTAGTTGGTGG-3
(R)b 5 -CRGTACTTGTBBDCTATCG-3
t◦ = 45 ◦C
(III) LSU rDNA (part) ∼1,770 bp
(F)b 5 -ACCCGCTGAAYTTAAGCATAT-3
(R)b 5 -GCCAATCCTTATCCCGAAGTTAC-3
t◦ = 50 ◦C
(IV) LSU rDNA (part) ∼1,740 bp
(F)b 5 -TCCGCTAAGGAGTGTGTAACAAC-3
(R)b 5 -TTCTGACTTAGAGGCGTTCAG-3
t◦ = 50 ◦C
Selenidium pygospionis
from Polydora glycymerica
(Sea of Japan)
(V) SSU rDNA (part) ∼1,610 bp
(F)a 5 -GTATCTGGTTGATCCTGCCAGT-3
1,610 bp (R) 5 -GGAAACCTTGTTACGACTTCTC-3
MH061279 t◦ = 45 ◦C
Selenidium pherusae
(VI) SSU rDNA (part) ∼1,600 bp
(F)a 5 -GTATCTGGTTGATCCTGCCAGT-3
2,551 bp (R) 5 -GGAAACCTTGTTACGACTTCTC-3
MH061280 t◦ = 45 ◦C
(VII) SSU rDNA (part), ITS1,
5.8S rDNA, ITS2, LSU rDNA
(part)
∼1,040 bp
(F)b 5 -TCCGCTAAGGAGTGTGTAACAAC-3
(R)b 5 -CCTTGGTCCGTGTTTCAAGAC-3
t◦ = 50 ◦C
aThe primer sequence was based on Medlin et al., 1988.
bThe primer sequences were based on Van der Auwera et al., 1994.
Selenidium spp. n.: Structure and Phylogeny 839
(see above), i.e. archigregarines were paraphyletic
in the resulting SSU rDNA-based phylogenies;
however, the nodal support of this grouping was
low (Fig. 7). The archigregarine paraphyly has
been repeatedly reported before but always with
weak support (Rueckert and Horák 2017; Rueckert
et al. 2011; Schrével et al. 2016; Wakeman et al.
2014; Wakeman and Horiguchi 2018; Wakeman
and Leander 2013). Thus, the deep branching of
archigregarine lineages had remained unresolved.
Therefore, we tested alternative phylogenies (see
below). The lineage Ag2 comprised two parasites
of sipunculids and one environmental sequence.
The lineage Ag1 was the largest and comprised
parasites of polychaetes from the families Cirratulidae,
Flabelligeridae, Opheliidae, Sabellidae,
Sabellariidae, Serpulidae, and Spionidae, including
the newly obtained archigregarine sequences,
and a number of environmental sequences, only
two of which were involved in the final phylogenetic
analyses. This was the longest archigregarine
branch, which had full support in both BI and ML
analyses. Number “1” was assigned to this clade
because Selenidium pendula, the type species
of the genus Selenidium, belonged to it (Fig. 7).
Within the clade, the parasites of Serpulidae and
Sabellariidae formed robust subclades, whilst the
subclade of the parasites of Spionidae had full
BP and moderate BP support. The two available
sequences of the parasites of Sabellidae did not
form a subclade, although they occupied neighbouring
positions in the tree with moderate or
low nodal supports; this indicates that their positions
are actually unresolved. Both sequences of
S. pygospionis (from Pygospio elegans and Polydora
glycymerica) grouped with the sequences of S.
boccardiellae and S. pendula within the subclade of
parasites of spionid polychaetes (see above). The
sequence of S. pherusae branched earlier, after S.
opheliae from the polychaete Ophelia roscoffensis
(Opheliidae); both these branches had moderate or
low nodal supports.
Analyses of LSU rDNA and the ribosomal
operon: All phylogenies based on these phylogenetic
markers resulted in identical topologies both
in the BI (Fig. 8) and the ML (not shown) analysis.
Overall, they recovered the major alveolate clades
that agreed with the phylogenies inferred from SSU
rDNA, both already published (see above) and
newly obtained, but with a higher resolution of allalveolate
and myzozoan deep branching.
Unlike SSU rDNA-based phylogenies, all analyses
of the LSU rDNA dataset (53 sequences, 2,913
sites) resulted in the well-supported monophyly of
gregarines (PP = 1.0, BP = 91). The only archigregarine
sequence (S. pygospionis from P. elegans)
formed a long branch (PP = 1.0, BP = 70) immediately
after the clade Eg1 containing sequences
of the “short-branching” gregarines Ascogregarina
taiwanensis and Neogregarinida sp. OPPPC1
AB748927 and before the clade Eg2 including
sequences of the “long-branching” eugregarines.
Therefore, it broke down the monophyly of eugregarines
(Fig. 8A).
The resulting phylogenetic trees inferred from
the ribosomal operon dataset (alignment of 53
sequences, 4,618 sites) showed the same topology
as the LSU rDNA-based phylogenetic tree with
increased support for several branches (Fig. 8B).
Within the sporozoan clade, all the studied gregarines
were monophyletic, and the support was
almost the same as in the LSU rDNA phylogenies.
However, the BP support for the position of
S. pygospionis, splitting eugregarine monophyly,
was somewhat lower (BP = 64% vs 70% in the LSU
rDNA tree).
Testing alternative phylogenies: Alternative
topologies of phylogenetic trees were analyzed
together with the topologies of the resulting tree
yielded by the phylogenetic analyses (the reference
tree, Fig. 7) with the use of the set of six
widespread tests (see Methods). The results are
presented in Figure 9 and Supplementary Material
Table S1. For SSU rDNA phylogenies based on 128
sequences, the hypotheses on the monophyly of
archigregarines and various positions of this combined
lineage (Ag1 + Ag2 + Ag3 + Ag4) within the
cryptosporidian-gregarine clade and sisterly to it
were tested under the assumptions of monophyly
or polyphyly of eugregarines (Fig. 9A). In addition
to the resulting phylogenetic trees exhibiting the
archigregarine paraphyly (see above), two of the
four alternative SSU rDNA phylogenies containing
monophyletic archigregarines were found not to be
rejected by any test; this contradicts the results
obtained by Rueckert and Horák (2017). Both these
phylogenies included monophyletic eugregarines:
(i) monophyletic archigregarines as a sister group
either to monophyletic eugregarines (which means
the monophyly of the gregarines as a whole) or
(ii) to the clade comprising cryptosporidians and
monophyletic eugregarines. All the tests rejected
simultaneous monophyly of archigregarines and
polyphyly of eugregarines (Fig. 9A and Supplementary
Material Table S1).
For the LSU rDNA- and ribosomal operon-based
phylogenies, the phylogenetic position of the only
available archigregarine sequence, S. pygospionis
840 G.G. Paskerova et al.
Figure 7. Bayesian inference tree of alveolates inferred from the dataset of 128 SSU rDNA sequences and
1,550 sites under the GTR + + I model. Numbers at the nodes indicate Bayesian posterior probabilities (numerator)
and ML bootstrap percentage (denominator). Black dots on the branches indicate Bayesian posterior
probabilities and bootstrap percentages of 0.95 and 90% or more, respectively. The newly obtained sequences
Selenidium spp. n.: Structure and Phylogeny 841
from P. elegans, was examined, and the same set of
topologies was used (Fig. 9B and C, and Supplementary
Material Table S1). Congruently to SSU
rDNA-based phylogenies, the hypothesis of its sister
position to monophyletic eugregarines was not
rejected by any test among the phylogenies based
both on LSU rDNA (Fig. 9B) and on the ribosomal
operon (Fig. 9C). All the other topologies were
rejected by all tests among LSU rDNA-based phylogenies
but among operon-based phylogenies the
polyphyly of eugregarines was not rejected in the
case when the archigregarine was a sister lineage
to the eugregarine clade Eg2 and cryptosporidians
were a sister group to the clade Eg1, i.e. as in the
SSU rDNA reference tree.
Summing up, the monophyletic archigregarines
as a sister group to the monophyletic eugregarines
was the only possible alternative topology (permitted
by all the tests) shared by the phylogenies
based on all the three genetic markers used (SSU
rDNA, LSU rDNA, and ribosomal operon phylogenies).
However, the number of sequences in the
LSU rDNA/ribosomal operon database is significantly
lower than in the SSU rDNA database and
is still insufficient to make valid comparisons and
meaningful conclusions.
Discussion
Justification of Newly Described Species
Selenidium pygospionis sp. n.
There are currently eleven species of archigregarines
inhabiting polychaetes of the family
Spionidae (Dibb 1938; de Faria et al. 1917;
Fowell 1936a,b; Ganapati 1946; Giard 1884; Levine
1971; Ray 1930; Reichenow 1932; Wakeman and
Leander 2012). They are described in varying
degree of detail; in particular, only three species
have been examined by electron microscopy:
Selenidium pendula, S. boccardiellae, and S.
pygospionis sp. n. (Rueckert and Horák 2017;
Schrével et al. 2016; Wakeman and Leander 2012;
this study). Most of these archigregarines belong to
the genus Selenidium, one archigregarine belongs
to the genus Selenocystis (S. foliata). However,
Selenidium foliatum Ray, 1930 is suggested to be
a synonym of S. foliata Dibb, 1938 (Desportes
and Schrével 2013; Dibb 1938; Schrével 1970).
We also suspect that Selenidium intraepitheliale
Reichenow, 1932 may be a synonym of S. spionis
(Kolliker, 1945) Ray, 1930 as these archigregarines
parasitizing the same host are identical (Levine
1971; Ray 1930; Reichenow 1932; Schrével 1970).
The resulting combinations of archigregarines from
spionid polychaetes are presented in Supplementary
Material Table S2. Additionally, three
Selenidium species were reported from the spionid
polychaetes Dipolydora coeca (Caullery and Mesnil
1899), Spio filicornis (Caullery and Mesnil 1901),
and Pygospio elegans (Caullery and Mesnil 1899;
Reichenow 1932) without species descriptions.
The archigregarines from spionids are usually
characterized by a high frequency and a middle
intensity of parasite infection (Douglass and Jones
1991). Merogony was reliably shown only in S.
axiferens (Fowell 1936b). At the same time, the
presence of numerous individuals of an archigregarine
in a spionid polychaete was considered as
an indirect evidence of the presence of merogony
in their life cycle (Schrével et al. 2016). They usually
have a vermiform, more or less flattened cell, with
a knob-shaped (dome-shaped) mucron. Some of
them, including the type species S. pendula, have
an optically distinct cytoplasm arranged along the
cell axis, around the nucleus and in the radials running
to the cell periphery (Desportes and Schrével
2013; Fowell 1936a,b; Ray 1930; Rueckert and
Horák 2017).
Trophozoites of Selenidium pygospionis sp. n.
isolated from Pygospio elegans polychaetes in
this study were somewhat similar with S. spionis,
S. intraepitheliale, S. martinensis, parasitizing
in various hosts, in cell size, cell shape, and in
the number of longitudinal folds (Levine 1971;
Ray 1930; Reichenow 1932; Supplementary Material
Table S2). However, the trophozoites of S.
pygospionis were easy to distinguish from other
species of Selenidium by their hook-like anterior
end. Therefore, we established a new species
for these archigregarines using host-specific and
morphological characteristics as well as molecularphylogenetic
markers.
The molecular-phylogenetic analyses revealed
an almost full identity (two substitutions per
1,610 bp) of sequences from archigregarines isolated
from the polychaetes P. elegans of the
of Selenidium spp. are highlighted by black. Single asterisks indicate “Selenidioides” spp. and double asterisks
– “Selenidium” spp. as proposed in the revision by Levine (1971). The information about the host taxonomical
affiliations is given in gray.
842 G.G. Paskerova et al.
Figure 8. Bayesian inference trees of alveolates inferred from the datasets of 53 taxa under GTR + + I model
for, A, LSU rDNA (2,913-sites dataset); B, ribosomal operon (4,618-sites dataset). Numbers at the nodes
indicate Bayesian posterior probabilities (numerator) and ML bootstrap percentage (denominator). Black dots
on the branches indicate Bayesian posterior probabilities and bootstrap percentages of 0.95 and 90% or more,
Selenidium spp. n.: Structure and Phylogeny 843
Figure 9. Tested alternative topologies of phylogenetic trees. A, testing archigregarine monophyly on alternative
topologies for the SSU rDNA phylogenies (the dataset of 128 taxa and 1,550 sites). The scheme below the
raw of the alternative topologies shows the structure of the artificially composed monophyletic archigregarine
clade. B and C, testing phylogenetic position of the archigregarine Selenidium pygospionis from Pygospio
elegans on alternative topologies for the (B) LSU rDNA phylogenies (the dataset of 53 taxa and 2,913 sites)
and (C) ribosomal operon phylogenies (the dataset of 53 taxa and 4,618 sites). Permissible topologies (not
discarded by all tests) are marked by numbers within circles. Abbreviations: Ag1, archigregarines (the subclade
numbers in the scheme correspond those in Fig. 8); Eg1 and 2, eugregarine clades (short- and long-branching,
respectively; see Figs 8 and 9); Cr, cryptosporidia; C, coccidiomorphs (Coccidia and Hematozoa); S.p., the
sequences of S. pygospionis.
respectively. The newly obtained sequences of the archigregarine Selenidium pygospionis is highlighted by
black. Accession numbers in the tree B are arranged in following order: SSU rDNA, 5.8S (if not available then
“–”), LSU rDNA.
844 G.G. Paskerova et al.
White Sea and Polydora glycymerica of the Sea
of Japan. This means that these archigregarines,
though isolated from hosts from geographically
distinct regions, belong to the same species, S.
pygospionis. We also suspect that Selenidium sp.
from P. elegans (former P. seticornis) collected
near Plymouth, the English Channel (Caullery and
Mesnil 1899; Reichenow 1932), is S. pygospionis
described in the present study. These data indicate
that the biogeographic distribution of archigregarines
may be extensive and that the systematic
principle “new host – new species”, sometimes
used for gregarines (Levine 1971), should be
applied to archigregarines with caution.
We believe that the archigregarine recently found
to harbor the microsporidium Metchnikovella dogieli
(Paskerova et al. 2016) is likely to be S. pygospionis
since it had been sampled at the same
site where samples for the present study were
taken. Metchnikovellidean microsporidia, inhabiting
both archigregarines and eugregarines (Desportes
and Schrével 2013; Mikhailov et al. 2017; Rotari
et al. 2015; Sokolova et al. 2013, 2014), appear to
possess a universal complex of adaptations (mechanisms
of invasion, life cycles, metabolic strategies,
etc.) allowing them to parasitize in structurally different
gregarines.
Selenidium pherusae sp. n.
Only four archigregarine species have been known
to parasitize polychaetes of the family Flabelligeridae
(Bogolepova 1953; Castellon and Gracia
1988; Kuvardina and Simdyanov 2002; Simdyanov
1992; Tuzet and Ormières 1958; Tuzet and
Ormières 1965). The resulting combinations of
archigregarines from flabelligerid polychaetes are
presented in Supplementary Material Table S3.
All of them belong to the genus Selenidium, and
almost all have a longitudinally folded cortex,
except S. pennatum which has two lateral vanes or
ridges (Simdyanov 1992). Only one archigregarine
species has been described from the polychaete
Pherusa plumosa, S. curvicollum (Bogolepova
1953). Its trophozoites have 13–15 cortical grooves
per side and a large, curved proboscis-like anterior
end. The archigregarine described in the present
study parasitizes the intestine of the polychaete Ph.
plumosa; however, it lacks any folds at the surface
and has a small, slightly truncated mucron.
Hence, we consider it as a new species, Selenidium
pherusae sp. n.
Nutrition of Archigregarines
Myzocytotic feeding (myzocytosis), i.e. sucking the
host cell cytoplasm via a well-developed apical
complex, was found in S. hollandei, S. pendula and
S. orientale (Schrével 1968, 1971b; Schrével et al.
2016; Simdyanov and Kuvardina 2007). It is thought
that myzocytosis is common in archigregarines
(Desportes and Schrével 2013; Schrével et al.
2016; Simdyanov and Kuvardina 2007; Wakeman
and Horiguchi 2018; Wakeman et al. 2014). In the
mucron of S. pygospionis and S. pherusae, we
observed the organelles of the apical complex such
as the conoid, the mucronal vacuole, rhoptries, and
micronemes. Additionally, a series of connected
vacuoles arranged along the cell axis and around
the nucleus was present in S. pygospionis. We suggest
that the axial streak of S. pygospionis, well
visible in living archigregarines under a light microscope,
is a system of the observed (presumably
digestive) vacuoles originating in the mucron during
the myzocytosis and transporting nutrients from
the anterior to the posterior end along the cell axis.
To note, numerous vacuoles around the nucleus
have also been found in the cytoplasm of S. pendula
(Schrével 1970, 1971a). These vacuoles may
be a component of a similar system of digestive
vacuoles.
Different cytoplasmic vesicles containing some
multi-membranous whorls or dense material and
located in the ectoplasm near the pellicle or
inserted in the inner membrane complex were
previously observed in several Selenidium spp.
(Schrével 1971a; Schrével et al. 2016; Wakeman
and Horiguchi 2018). It was suggested that these
structures might be involved in a surface-mediated
nutrition present alongside with the nutrition via
the typically organized micropores (Wakeman and
Horiguchi 2018).
The heterogeneity of the organelles’ distribution
(narrow electron-translucent spaces lacking any
visible organelles) observed in the cytoplasm of S.
pygospionis may point to the presence of microfilaments
similar to those demonstrated around the
nucleus in S. pendula and S. hollandei (Schrével
1971a). Putative digestive vacuoles and coexisting
microfilaments may play a role in the transport of
nutrients from the anterior to posterior end during
myzocytotic feeding.
Selenidium spp. n.: Structure and Phylogeny 845
Motility of Archigregarines
In general, archigregarines show different types
of movement such as bending, twisting, coiling,
rolling, and pendular motility. Some of them can
also contract their cell. These movements were
often likened to those of nematode worms (Fowell
1936a,b; Gunderson and Small 1986; Leander
2007, 2008b; Mellor and Stebbings 1980; Schrével
1971a,b; Schrével et al. 1974, 2016; Stebbings
et al. 1974; Wakeman et al. 2014). Both archigregarines
described in the present study can move
by forming 1–4 (in S. pygospionis) or only one
(in S. pherusae) bending sections along the cell
but never contract. Additionally, as it can be easily
seen in flattened trophozoites of S. pygospionis,
their bending motility is generated only in one cell
plane. We propose to refer to this type of motility
as a nematode-like bending, where the alternate
sides (flattened sides in S. pygospionis or body
halves in S. pherusae) act antagonistically in the
bending sections forming in a single plane of the
cell. It is similar to the pendular motility of S. pendula,
where one bend is generated in the anterior
part of the cell and runs to the posterior one, but
considerably different from the bending motility of
other Selenidium spp. (e.g. S. hollandei, S. sabellariae),
where bends, generated in different cell
planes, can combine with contraction and twisting
of the cell in different cell sections (Desportes and
Schrével 2013; Rueckert and Horák 2017; Schrével
1967, 1970). The character of archigregarine motility
may reflect the dynamics and architecture of the
cytoskeleton.
Nematode-like bending is an evidence in favour
of the general hypothesis about motility of Selenidium
spp. postulating that the three-membrane
pellicle and the longitudinal microtubules are
skeletal and motile units representing together
a unicellular analogue of the musculocuticular
system of nematodes (Leander 2007, 2008b;
Stebbings et al. 1974). A similar motility mechanism
was also demonstrated in the blastogregarine
Siedleckia nematoides. It performs pendular, twisting,
undulating movements and possesses the
longitudinal microtubules organized in a layer or layers
under the trimembrane pellicle (Valigurová et al.
2017). To note, the axial streak together with putative
microfilaments may also be involved in the cell
motility as an additional skeletal element helping
to reverse movement and maintenance of the cell
shape in bends as suggested by the concept of the
statomotor system (Fowell 1936b).
According to the previous authors, the dynamic
motility of archigregarines and blastogregarines
is correlated with the number of mitochondria
located directly beneath the subpellicular microtubules
(Desportes and Schrével 2013; Leander
2006, 2007; Mellor and Stebbings 1980; Schrével
1971b; Stebbings et al. 1974; Valigurová et al.
2017). Our observations of the ultrastructure of both
studied archigregarines may lend further support
to this idea. An actively bending (up to 4 bends
at a time) S. pygospionis has numerous mitochondria
arranged in a layer beneath the subpellicular
microtubules, while a less pronouncedly bending
(only one bend at a time) S. pherusae has fewer
mitochondria.
Intracellular Development in Selenidium
pygospionis sp. n.
We observed small cells of putative S. pygospionis
localized within the parasitophorous vacuoles
in the host enterocytes. Intracellular young trophozoites
were also found in other archigregarines
such as Selenidium spionis, S. mesnili, S. foliatum,
and S. cauleryi (Ray 1930). An intracellular localization
may be an initial stage of the trophozoite
development in some archigregarines. As trophozoites
grow, the host cells are destroyed, and the
location of the parasites becomes extracellular. In
the case of S. pygospionis, well-developed and
extracellularly localized trophozoites are anchored
between the host intestinal epithelium folds by their
hook-like anterior end. We do not know whether
they form any attachment site with the host cell
at some period of their development. Intra-tissue
stages mentioned in the description of S. spionis
and S. foliatum (Caullery and Mesnil 1899, 1901;
Ray 1930) as a few full-grown individuals embedded
in the intestine wall tissue under the epithelial
layer and lying parallel to the host longitudinal axis
can be regarded as an abnormal and occasional
development of trophozoites.
Notes on the Taxonomy and Phylogeny of Archi-
gregarines
Levine (1971) proposed to divide the genus
Selenidium into two genera based on the presence/absence
of merogony in the life cycle. He
established a new genus, Selenidioides, for gregarines
with merogony and assigned it to the
archigregarines. The species without merogony (or
in which merogony was unknown) were transferred
to the eugregarines within the genus Selenidium.
846 G.G. Paskerova et al.
In his opinion (Levine 1971), the data on ultrastructure
and life cycles of archi- and eugregarines
available at that time (MacGregor and Thomasson
1965; Schrével 1966, 1970, 1971a,b; Vávra 1969;
Vivier and Schrével 1964, 1966; Vivier et al. 1970;
etc.) were insufficient for the separation of these two
groups. Levine’s classification has become popular
and has been applied in some revisions on protists
and taxonomic databases (Perkins et al. 2000;
WoRMS).
Available molecular phylogenetic evidence
(Rueckert and Horák 2017; Rueckert and Leander
2009; Schrével et al. 2016; Wakeman and
Horiguchi 2018; Wakeman and Leander 2012,
2013; Wakeman et al. 2014) and the results of this
study show that the archigregarines are separated
into several phylogenetic lineages. However, this
separation does not correspond to the taxonomical
action proposed by Levine. On one hand, the
type species of the genus Selenidium, S. pendula,
belong to the clade Ag1 (“true Selenidiidae” as
proposed by Schrével et al. (2016)) together with
representatives of the “Selenidioides” group, S.
mesnili and S. hollandei. On the other hand, some
gregarines of the “Selenidium” group, S. terebellae,
S. fallax, and S. orientale, belong to the clades Ag2,
Ag3, and Ag4 respectively, not to the clade Ag1
(Fig. 7). Thus, the taxonomical approach proposed
by Levine (1971) for archigregarines should be
abolished together with the genus Selenidioides
Levine, 1971 as has already been suggested
by different authors (Rueckert and Horák 2017;
Rueckert and Leander 2009; Schrével et al. 2016;
Wakeman and Leander 2012).
Overall, the molecular phylogeny of the archigregarines
based on the available DNA sequences is
largely congruent with the taxonomical affiliations of
their hosts (Schrével et al. 2016; this study). Indeed,
archigregarines of the lineage Ag4 inhabit polychaetes
of the family Terebellidae, archigregarines
of the lineage Ag2 occur in sipunculid hosts, while
most Selenidium spp. of the lineage Ag1 parasitize
in different sedentary polychaetes (Fig. 7), however,
their grouping in subclades within this clade
also agrees well with the taxonomical affiliations
(families) of hosts. We consider that this reflects in
various degrees some aspects of host-parasite coevolution,
which may become an important subject
of research in the future.
The macrosystem of archigregarines is questionable
because of the issue of their monophyly,
unresolved both in terms of the molecular phylogeny
and the classic cladistic approach based
on morphology. The morphology-based hypothesis
about the monophyly of Selenium-like archigregarines
is difficult to substantiate in terms of
cladistics, as all their ultrastructural key features
(the ultrastructure of the cortex and mucron)
appear to be symplesiomorphies (the aforementioned
morphostasis) rather than synapomorphies.
Apart from Selenidium spp., morphologically different
representatives of the genera Ditrypanocystis,
Exoschizon, Merogregarina, Meroselenidium, and
Selenocystis have also been affiliated to archigregarines
(Desportes and Schrével 2013; Perkins
et al. 2000). They are intestinal parasites of polychaetes
and sipunculids as well as of colonial
ascidians and oligochaetes. No molecular phylogenetic
evidence from these parasites is currently
available (Desportes and Schrével 2013) and
electron-microscopic data are extremely scanty
(Butaeva et al. 2006). Some of these organisms
may be neither archigregarines nor even gregarines.
A demonstrative example is the case of
Platyproteum (formerly Selenidium) vivax and Filipodium
fascolosomae, gregarine-like organisms,
which have been shown to be an independent
lineage of Myzozoa, so-called “squirmids”
(Cavalier-Smith 2014; Rueckert and Leander
2009). Both parasites are capable of very dynamic
cellular deformations referred to as the peristaltic
motility (metaboly) (Gunderson and Small 1986;
Leander 2006; Rueckert and Leander 2009). Since
this is dissimilar to the real squirming, we prefer to
call this lineage “platyproteids” (Fig. 7) instead of
“squirmids” proposed by Cavalier-Smith (2014).
The SSU rDNA-based phylogenies (CavalierSmith
2014; Cavalier-Smith and Chao 2004;
Grassé 1953; Rueckert and Leander 2009;
Rueckert and Horák 2017; Schrével et al. 2016;
Wakeman and Horiguchi 2018; Wakeman and
Leander 2012, 2013; Wakeman et al. 2014; this
study) reveal archigregarines as a paraphyletic
group, although their deep branching is actually
unresolved and shows weak nodal supports.
The test of alternative topologies, provided in this
study, did not reject archigregarine monophyly,
but only within the framework of the hypothesis
that eugregarines were monophyletic too (Fig. 9A).
Thus, the phylogenetic analyses of the SSU rDNA
yield ambiguous results. As explained previously
(Simdyanov et al. 2017), SSU rDNA-based phylogenies
appear to be of little use for resolving the
deep branching order of gregarines and apicomplexans
altogether. This is likely a consequence
of an explosive evolutionary radiation of gregarines
and/or rapid evolution of their SSU rDNA
sequences. The LSU rDNA and near-complete
rDNA operon provide an increased phylogenetic
resolution over SSU rDNA and could be useful in
Selenidium spp. n.: Structure and Phylogeny 847
advancing the phylogeny and taxonomy of archigregarines
and gregarines in general (Simdyanov et al.
2015, 2017). Unfortunately, only one archigregarine
LSU rDNA sequence is now available (this study),
and it even forms a long branch, so that its position
in the obtained phylogenies might be affected by the
long branch attraction artifact. An enhanced taxon
sampling of archigregarine LSU rDNA sequences
is necessary for more substantial conclusions, with
special attention to short-branching species, representatives
of the lineages Ag2, Ag3, and Ag4.
Multigenic phylogenies including a broad representative
sampling of archigregarines could provide the
ultimate test of the hypotheses about the evolution
within Apicomplexa.
Taxonomic Summary
Phylum Apicomplexa Levine, 1970
Subphylum Sporozoa Leuckart, 1879
Class Gregarinomorpha Grassé, 1953, emend.
Simdyanov et al., 2017
Order Archigregarinida Grassé, 1953
Family Selenidiidae Brasil, 1907
Genus Selenidium Giard, 1884
Selenidium pygospionis sp. n.
Diagnosis. Trophozoites aseptate, elongated and
slightly flattened with narrowed ends, embedded
in the host intestinal epithelium (extracellular or
intracellular location) or freely localized in the
intestinal lumen. Anterior end usually hook-like,
bent towards one of the wide sides. Mucron
naked, dome-shaped. Trophozoites measuring
34 to 288 ␮m (average 144 ␮m, mode 146 ␮m,
n = 79) in length, 4 to 25 ␮m (average 12 ␮m,
mode 11 ␮m, n = 76) in width. Cell surface with
22–30 (usually 28, n = 12) broad and low folds
separated by grooves. 10–12 grooves per flattened
side and 1–3 grooves per narrow side. Nucleus
oval, 6–22 ␮m (av. 17 ␮m, n = 40) × 5–11 ␮m (av.
8.4 ␮m, n = 26), located in the widest part and
expanding along the longitudinal axis of the cell.
Single nucleolus situated in nucleus part facing
anterior end. Intracellular axial streak of optically
distinct cytoplasm expanding from anterior to
posterior end and forming expansion around
nucleus and numerous radial threads toward cell
periphery. Syzygy caudo-caudal. Attached or
non-attached trophozoites and syzygy partners
moving by nematode-like bending (formation of
bends in a single plane of cell).
DNA sequences. SSU, ITS1, 5.8 S, ITS2, and
LSU rDNA sequences for individuals, isolated from
the polychaetes Pygospio elegans (White Sea)
(GenBank MH061278) and SSU rDNA – from the
polychaetes Polydora glycymerica (Sea of Japan)
(GenBank MH061279).
Type material (syntypes). Resin blocks and fixed
slides containing archigregarines and pieces of
infected host intestine deposited in the collection of
Department of Invertebrate Zoology, St Petersburg
State University; Figures 1–4 (this publication)
show some of the syntypes.
Hosts and localities. Polychaetes Pygospio
elegans Claparède, 1863 (Spionidae, Polychaeta);
Bolshoy Goreliy Island, Keret’ Archipelago, Chupa
Inlet, Kandalaksha Bay, White Sea, 66◦
18.770 N;
33◦
37.715 E; Velikaja Salma, Kandalaksha Bay,
White Sea, 66◦
33.200 N, 33◦
6.283 E. Polychaetes
Polydora glycymerica Radashevsky, 1993 (Spionidae,
Polychaeta); Peter the Great Bay, Sea of
Japan, 42◦
53 29 N, 132◦
44 07 E.
Location within host. Intestine (midgut and
hindgut).
Etymology. Species name, pygospionis, refers to
the genus name of one of the hosts.
Selenidium pherusae sp. n.
Diagnosis. Trophozoites aseptate, vermiform.
Anterior end narrowed, slightly truncated; posterior
end pointed in young or rounded in
mature individuals. Cell surface smooth, without
well-developed folds or grooves. Trophozoites
measuring 38–269 ␮m (n = 6) in length, 10–18 ␮m
(n = 4) in width. Nucleus spherical (11–12 ␮m,
n = 2), located in the widest part of the posterior half
of the cell, with the single nucleolus. Attached and
detached trophozoites exhibiting bending motility.
DNA sequences. SSU rDNA sequences (GenBank
MH061278).
Type material (syntypes). Resin blocks and fixed
slides containing archigregarines and pieces of
infected host intestine deposited in the collection of
the author TGS, Department of Invertebrate Zoology,
Lomonosov Moscow State University; Figures
5–6 (this publication) show some of the syntypes.
848 G.G. Paskerova et al.
Host and locality. Polychaetes Pherusa plumosa
(Müller, 1776) (Flabelligeridae, Polychaeta); Peter
the Great Bay, Sea of Japan, 42◦
53 29 N,
132◦
44 07 E.
Location within host. Midgut.
Etymology. The species name, pherusae, refers
to the genus name of the host.
Methods
Collection of polychaete hosts and isolation of gregarines:
Polychaetes Pygospio elegans Claparède, 1863 (Spionidae,
Polychaeta) were collected at two sites of the littoral zone
near the Marine Biological Station of St Petersburg State
University (Bolshoy Goreliy Island, Keret’ Archipelago, Chupa
Inlet, Kandalaksha Bay, White Sea, 66◦
18.770 N; 33◦
37.715 E)
and the White Sea Biological Station of Lomonosov Moscow
State University (Velikaja Salma, Kandalaksha Bay, White Sea,
66◦
33.200 N, 33◦
6.283 E) during the summers of 2002–2015
years.
Polychaetes Polydora glycymerica Radashevsky, 1993 (Spionidae,
Polychaeta) boring shell walls of the living bivalves
Glycymeris yessoensis (G. B. Sowerby III, 1889) and polychaetes
Pherusa plumosa (Müller, 1776) (Flabelligeridae,
Polychaeta) inhabiting druses of the Far East mussels Crenomytilus
grayanus (Dunker, 1853) were collected by SCUBA
divers in 2007 near Vostok biological station, National Scientific
Center of Marine Biology, Russian Academy of Sciences (Peter
the Great Bay, Sea of Japan, near Nakhodka, 42◦
53 29 N,
132◦
44 07 E).
Prior to dissection, the examined animals were stored in
small containers (about 50 worms per 250 ml container) at
+10 ◦
C with periodically changed seawater. The polychaetes
were cultured up to a month. Dissection of polychaetes and
isolation of parasites were performed with the help of fine needles
and hand-drawn glass pipettes under a stereomicroscope
(MBS-10, Russia). The released parasites or small fragments
of the host intestine with attached gregarines were rinsed thrice
with seawater filtered through Millipore (0.22 ␮m), then prepared
for light microscopy or fixed for electron microscopy.
Individual cells were also subjected to DNA extraction.
Light microscopy: More than 100 polychaetes of P. elegans
were investigated in squash preparations [compressing
of a polychaete specimen between the object and cover slides
before microscopic analysis] (Fig. 1A–B, D–M). Separate archigregarines
isolated from the intestines of P. elegans (Fig. 1C,
N–Q), P. glycymerica (data not shown) and Ph. plumosa
(Fig. 5A) were also investigated in living preparations. All preparations
were investigated with the use of a series of light
microscopes equipped with different digital cameras: a MBR-1
microscope (LOMO, Russia) equipped with phase contrast and
connected to a Canon EOS 300D digital camera; a Zeiss microscope
(Carl Zeiss, Germany) connected to a Nikon Coolpix
7900 camera; a Zeiss Axio Imager.A1 connected to an AxioCam
MRc5 digital camera (Carl Zeiss, Germany); a Leica DM
2500 microscope equipped with DIC optics and Plan-Apo objective
lenses and connected to a DFC 295 digital camera (Leica,
Germany).
Several individuals of S. pherusae from polychaetes Ph.
plumosa were fixed with 3% formaldehyde in seawater, stained
with Carazzi’s hematoxylin and examined under a Zeiss microscope
connected to a Nikon Coolpix 7900 camera (Fig. 5B).
Electron microscopy: Small pieces of the polychaete intestine
with attached archigregarines or free archigregarines
released from the host gut lumen were fixed in 2.5% glutaraldehyde
in 0.2 M cacodylate buffer with/without 0.05% MgCl2 (pH
7.4, final osmolarity 720 mOsm) for 1–4 h, washed in filtered
seawater and postfixed in 1–2% osmium tetroxide in the same
buffer for 1–2 h. The entire procedure of fixation was performed
at +4 ◦
C. Fixed samples were dehydrated in an ascending
ethanol series. Some of them were additionally transferred to
an ethanol/acetone mixture and rinsed in pure acetone before
the following procedure. For SEM, the fixed and dehydrated
samples were critical point dried in liquid CO2 and then coated
with gold or platinum. The samples were investigated with a Tescan
MIRA3 LMU scanning electron microscope (TESCAN Brno,
Czech Republic), JSM-7401F (JEOL, Japan), FEI Quanta 250
(Thermo Fisher Scientific, Netherlands) and Hitachi S-405A
scanning electron microscope (Hitachi, Japan) (Figs 2, 5C, D).
For TEM, fixed and dehydrated samples were embedded in
Epon-Araldite or Epon blocks. They were sectioned with ultramicrotomes
Leica EM UC6 and Leica EM UC7 (Leica, Germany).
Ultra-thin sections were stained according to standard protocols
(Reynolds, 1963) and examined using a JEM 2100 (JEOL,
Japan), TEM–1010 (JEOL, Japan), and LEO 910 (Carl Zeiss,
Germany) electron microscopes equipped with a digital or film
cameras. In total, more than 20 gregarines from P. elegans
(Figs 3, 4A–C, F, G), about five gregarines from P. glycymerica
(data not shown) and five gregarines from Ph. plumosa (Fig. 6)
were sectioned and examined by TEM.
Ten entire worms of P. elegans were fixed in Bouin’s solution.
The material was dehydrated in a graded alcohol series,
infiltrated in a graded series of chloroform-Histomix and finally
embedded in Histomix paraffin wax (BioVitrum, Russian Federation).
Serial sagittal or coronal sections were prepared on
a Microm HM 360 rotary microtome (0.1–1 mm in thick). Sections
were mounted on the object slides. The preparations were
deparaffinized in xylol and then washed in acetone. After critical
point drying in liquid CO2 and coating with gold, they were
observed under a Tescan MIRA3 LMU scanning electron microscope
(TESCAN Brno, Czech Republic) (Fig. 4D, E).
DNA isolation, PCR and sequencing: Trophozoites of the
gregarine Selenidium pygospionis, about 100 cells isolated
from the polychaete P. elegans (Bolshoy Goreliy Island, Kandalaksha
Bay, White Sea, 2009), were fixed and stored in
RNAlater stabilization solution (Life Technologies, USA). DNA
extraction of this sample was performed with the Diatom DNA
Prep 200 kit (Isogen, Russia). About ten trophozoites and one
syzygy of S. pygospionis isolated from polychaetes P. glycymerica
(Peter the Great Bay, Sea of Japan, 2007) were rinsed with
seawater, deposited into 1.5-ml microcentrifuge tubes and then
were lyzed by the alkaline procedure (Floyd et al. 2002). In a
similar manner, 25 trophozoites of the gregarine S. pherusae
isolated from polychaete Ph. plumosa (Peter the Great Bay,
Sea of Japan, 2007) were lysed. The lysates obtained in two
last cases were directly used in PCR.
The nucleotide sequences of S. pygospionis from P. elegans
and S. pherusae were amplified in several PCRs with different
pairs of primers (Table 1 and Supplementary Material Fig.
S1). A set of overlapping fragments (I–IV and VI–VII, respectively;
see Supplementary Material Fig. S1) encompassing SSU
rDNA, ITS 1 and 2, 5.8S rDNA, and LSU rDNA was obtained.
For S. pygospionis from P. glycymerica, only one fragment (V)
containing the near complete SSU rDNA was PCR-amplified
(Table 1 and Supplementary Material Fig. S1). All PCRs were
performed with an Encyclo PCR kit (Evrogen, Russia) in a total
Selenidium spp. n.: Structure and Phylogeny 849
volume of 25 ␮l using a DNA Engine Dyad thermocycler (BioRad)
and the following protocol: initial denaturation at 95 ◦
C for
3 min; 40 cycles of 95 ◦
C for 30 sec, annealing at 45 ◦
C (fragments
I, II, V, and VI) or 50 ◦
C (fragments III, IV, and VII) for
30 sec, and extension at 72 ◦
C for 1.5 min; and final extension
at 72 ◦
C for 10 min. PCR products of the expected size were gelisolated
by a Cytokine DNA isolation kit (Cytokine, Russia). For
fragments I, II, and IV–VII, the PCR products were sequenced
directly. Fragment III was cloned by using an InsTAclone PCR
Cloning Kit (Fermentas, Lithuania) because the corresponding
PCR product was heterogeneous. Sequences were obtained by
using an ABI PRISM BigDye Terminator v. 3.1 reagent kit and an
Applied Biosystems 3730 DNA Analyzer for automatic sequencing.
After assembling the read fragments with the use of the
BioEdit 7.0.9.0 program (Hall 1999), the resulting contiguous
sequences were deposited in GenBank (accession numbers:
MH061278–80).
Molecular phylogenetic analysis: Four nucleotide alignments
were prepared for phylogenetic analyses: SSU (18S)
rDNA (128 and 53 sequences), LSU (28S) rDNA, and ribosome
operon (concatenated SSU, 5.8S, and LSU rDNA sequences).
The alignments were generated in the MUSCLE 3.6 program
(Edgar 2004) and manually adjusted with the use of the BioEdit
7.0.9.0 program (Hall 1999): gaps, columns containing few
nucleotides and hypervariable regions were removed. The final
length of the alignments was 1,550 bp. The taxon sampling of
128 sequences alignment was designed as to maximize the
phylogenetic diversity and the completeness of sequences in
the alignments. Representatives of heterokonts and rhizarians
were used as outgroups. The “reduced” SSU rDNA alignment
(53 sequences, 1,550 sites) was used as a constituent part of
the concatenated ribosomal operon dataset and, consequently,
covered the same taxon sampling.
For the LSU rDNA and ribosomal operon (concatenated
SSU, 5.8S, and LSU rDNAs) analyses, taxon sampling of
only 53 sequences were used due to the limited availability
of data for LSU rDNA and, especially, 5.8S rDNA. Therefore,
the 5.8S rDNA (155 sites in the alignment) was rejected
from the analysis of concatenated rDNA genes for seven
sequences (Chromera velia, Colponema vietnamica, Goussia
desseri, Stentor coeruleus, and 3 environmental sequences:
Ma131 1A38, Ma131 1A45, and Ma131 1A49): the corresponding
155 positions were replaced with “N” in the final ribosomal
operon dataset. The resulting datasets contained 53 sequences
(2,913 sites) for the LSU rDNA and the concatenated rDNA
sequences (4,618 sites) of the same 53 taxa for the ribosomal
operon. Thus, both taxon samplings comprised an identical set
of species, all of which were also represented in the alignment
of the 128 SSU rDNA sequences.
Maximum-likelihood (ML) analyses were performed with the
RAxML 7.2.8 program (Stamatakis 2006) under the GTR +
model and CAT approximation (25 rate categories per site).
The procedure included 100 independent runs of the ML analysis
and 1,000 replicates of multiparametric bootstrap. Bayesian
inference (BI) analyses were conducted with the MrBayes 3.2.6
program (Ronquist and Huelsenbeck 2003) under GTR + + I
model with eight categories of discrete gamma distribution.
The program was set to operate under the following parameters:
nst = 6, ngammacat = 6, rates = invgamma; the parameters
of Metropolis Coupling Marcov Chains Monte Carlo (mcmc):
nchains = 4, nruns = 4, temp = 0.2, ngen = 10,000,000, samplefreq
= 1,000, burninfrac = 0.5 (the first 50% of the 20,000
sampled trees, i.e., the first 5,000, were discarded in each
run). The following averages and standard deviations of split
frequencies were obtained: 0.014059 for the SSU rDNA analysis,
0.001084 for the LSU rDNA analysis, and 0.001113 for
the ribosomal operon concatenated analysis. The calculations
of bootstrap support for the resulting Bayesian trees were performed
with the use of the RAxML 7.2.8 program under the
same parameters as for the ML analyses (see above).
Alternative tree topologies were manually created and edited
with the use the TreeView 1.6.6 program (Page 1996). The reference
trees topologies of SSU rDNA (128-taxon dataset), LSU
rDNA, and ribosomal operons (53-taxon datasets each) were
copied from the trees showed in Figures 7 and 8. Alternative
topologies of SSU rDNA phylogenies were constructed by combining
all Selenidium-like archigregarines in a single clade (see
Fig. 9A) followed by positioning this clade within the sporozoans
clade successively as a sister group to the cryptosporidians
and eugregarine clades Eg1 and Eg2 under assumptions of
either their monophyly or polyphyly, or as a sister group to
the combined clade cryptosporidians + eugregarines (monophyletic
or polyphyletic variants). The eugregarine clades Eg1
and Eg2 were picked out as repeatedly recovered in the
reference trees and published phylogenies. Alternative topologies
of LSU rDNA and ribosomal operon phylogenies were
constructed in the same way, but with the use of the only
archigregarine sequence (S. pygospionis from P. elegans) available
to date. Topology tests were performed in Moscow State
University with TREE-PUZZLE 5.3.rc16 and CONSEL 0.1j
programs (Schmidt et al. 2002; Shimodaira and Hasegawa
2001). The following tests were used: bootstrap probability
(Felsenstein 1985); expected-likelihood weights (Strimmer
and Rambaut 2002); Kishino–Hasegawa test (Kishino and
Hasegawa 1989); Shimodaira–Hasegawa test (Shimodaira
and Hasegawa 1999); weighted Shimodaira–Hasegawa Test
(Shimodaira and Hasegawa 1999); approximately unbiased test
(Shimodaira 2002).
Acknowledgements
This work was supported by St Petersburg
State University [grant numbers 1.42.723.2017,
1.42.739.2017]; the Russian Foundation for Basic
Research [grant numbers 15-29-02601, 15-04-
08870, 15-34-51175, 16-34-50265, 18-04-00324,
18-04-01359]; the Czech Science Foundation
[project number GBP505/12/G112 (ECIP – Centre
of excellence)]. Ultrastructural and molecular studies
of S. pherusae were supported by the Russian
Science Foundation [project number 18-04-00123];
the molecular study of S. pygospionis – by the
Russian Science Foundation [project number 14-
50-00029]. To make phylogenetic computations,
the CIPRES Science Gateway (Miller et al. 2010)
and the Supercomputer Center of Lomonosov
Moscow State University (http://parallel.ru/cluster)
were used in this study. The authors would like to
thank the staff of the Marine Biological Station of St
Petersburg State University, the White Sea Biological
Station of Moscow State University, and Vostok
biological station of the National Scientific Center
of Marine Biology, Russian Academy of Sciences,
for providing facilities for field sampling and material
processing, as well as for their kind and friendly
850 G.G. Paskerova et al.
approach. TGS is deeply grateful to Dr. Vasily I.
Radashevsky (National Scientific Center of Marine
Biology, Russian Academy of Sciences), who first
detected the parasites of Polydora glycymerica and
helped with the sampling of the hosts, and to Profs.
Andrey V. Adrianov and Vladimir. V. Yushin from
the same institute for their help in organizing the
trip to Vostok biological station. GGP and TSM utilized
equipment of the core facility centers of St
Petersburg State University: “Molecular and Cell
Technologies” for electron microscopy and “Observatory
of Environmental Safety” for culturing of the
marine invertebrates. AD, MK and AV are grateful
to the Laboratory of Electron Microscopy, Biology
Centre CAS, an institution supported by the CzechBioImaging
large RI project (LM2015062 funded
by MEYS CR), for their help with obtaining some
EM data. TGS utilized equipment of the Electron
microscopy laboratory of the Faculty of Biology and
the Center of microscopy of the White Sea Biological
station, Lomonosov Moscow State University.
Appendix A. Supplementary Data
Supplementary data associated with this article
can be found, in the online version, at
https://doi.org/10.1016/j.protis.2018.06.004.
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