99
REVIEW ARTICLE
Haemogregarina bigemina (Protozoa: Apicomplexa: Adeleorina)
– past, present and future
Angela J. Davies1
, Nico J. Smit2
, Polly M. Hayes1
, Alan M. Seddon1
and David Wertheim3
1
School of Life Sciences and 3
School of Computing, Kingston University, Penrhyn Road, Kingston upon Thames, Surrey, KT1
2EE, UK;
2
Department of Zoology, Rand Afrikaans University, P.O. Box 524, Auckland Park 2006, South Africa
Key words: Haemogregarina bigemina, haemogregarines, development, transmission, Gnathia, isopods
Abstract. This paper reviews past, current and likely future research on the fish haemogregarine, Haemogregarina bigemina
Laveran et Mesnil, 1901. Recorded from 96 species of fishes, across 70 genera and 34 families, this broad distribution for H.
bigemina is questioned. In its type hosts and other fishes, the parasite undergoes intraerythrocytic binary fission, finally forming
mature paired gamonts. An intraleukocytic phase is also reported, but not from the type hosts. This paper asks whether stages
from the white cell series are truly H. bigemina. A future aim should be to compare the molecular constitution of so-called H.
bigemina from a number of locations to determine whether all represent the same species. The transmission of H. bigemina
between fishes is also considered. Past studies show that young fish acquire the haemogregarine when close to metamorphosis,
but vertical and faecal-oral transmission seem unlikely. Some fish haemogregarines are leech-transmitted, but where fish
populations with H. bigemina have been studied, these annelids are largely absent. However, haematophagous larval gnathiid
isopods occur on such fishes and may be readily eaten by them. Sequential squashes of gnathiids from fishes with H. bigemina
have demonstrated development of the haemogregarine in these isopods. Examination of histological sections through gnathiids
is now underway to determine the precise development sites of the haemogregarine, particularly whether merozoites finally
invade the salivary glands. To assist in this procedure and to clarify the internal anatomy of gnathiids, 3D visualisation of
stacked, serial histological sections is being undertaken. Biological transmission experiments should follow these processes.
Haemogregarines are apicomplexan protozoa,
broadly distributed among vertebrate hosts, including
fishes (Davies and Johnston 2000). They are especially
common in marine fishes, where they are recognised in
circulating erythrocytes, but also in cells of the leukocytic
series (Davies 1995). Siddall (1995), in a partial
taxonomic revision of the haemogregarine complex,
placed fish haemogregarines in the genera Cyrilia
Lainson, 1981 (one species), Desseria Siddall, 1995 (41
species) and Haemogregarina (sensu lato) Danilewsky,
1885 (13 species). Haemogregarina (sensu stricto) was
reserved for species infecting chelonians. This classification
was later adopted by Barta (2000) in his general
account of the apicomplexan suborder Adeleorina
Léger, 1911, although at least some Desseria spp. are
now known to belong to other genera (see Negm-Eldin
1999, Davies and Johnston 2000, Smit et al. 2003a).
Cyrilia spp. and Haemogregarina spp. (sensu lato)
are characterised by intraerythrocytic merogony in the
fish host, whereas Desseria spp. lack this process (see
Barta 2000, Davies and Smit 2001). Cyrilia spp. and
Desseria spp. also undergo sporogony in leeches (see
Davies and Smit 2001). In Cyrilia spp., sporogonic
development produces numerous sporozoites and two
life cycles are reported (Lainson 1981, Negm-Eldin
1999). In Desseria spp., sporogony yields more than 16
sporozoites, this is followed by primary merogony in
the same leech hosts, and one life cycle is described in
detail (see Siddall 1995). Evidence for invertebrate
stages among Haemogregarina spp. (sensu lato) is,
according to Siddall (1995), limited to just one fish
haemogregarine, Haemogregarina (s.l.) uncinata (Khan,
1978) Siddall, 1995, which undergoes development in
leeches, although this species was considered a member
of the genus Cyrilia by Lainson (1981).
The marine fish haemogregarine Haemogregarina
(s.l.) bigemina Laveran et Mesnil, 1901 appeared first in
Siddall’s list of Haemogregarina (sensu lato) and its
development in an invertebrate host was not reported
(Siddall 1995). However, it is a remarkable haemogregarine
because of its apparent cosmopolitan distribution
among marine fishes. It also appears to be the only apicomplexan
of its type transmitted by arthropods rather
than leeches (Davies and Smit 2001). This paper
reviews past and current knowledge of H. bigemina,
highlights some anomalies concerning the organism,
and suggests what is still to be determined.
This paper was presented at the 6th International Symposium on Fish
Parasites in Bloemfontein, South Africa, 22–26 September 2003.
FOLIA PARASITOLOGICA 51: 99–108, 2004
Address for correspondence: A.J. Davies, School of Life Sciences, Kingston University, Penrhyn Road, Kingston upon Thames,
Surrey, KT1 2EE, UK. Phone: ++44 208 547 2000; Fax: ++44 208 547 7562; E-mail: ajdavies.russell@kingston.ac.uk
100
Fig. 1. Giemsa-stained erythrocytes from Lipophrys pholis with stages of Haemogregarina bigemina. A – trophozoite; B, C – developing
meronts; D–F – meronts undergoing transverse binary fission; G–I – longitudinal binary fission of meronts; J – mature
paired gamonts. Scale bar = 10 µm.
PAST AND PRESENT
Distribution and development of Haemogregarina
bigemina in fishes
Haemogregarina bigemina was first described from
intertidal blenniid fishes Lipophrys pholis (Linnaeus,
1758) and Coryphoblennius galerita (Linnaeus, 1758)
in northern France (Laveran and Mesnil 1901). In
Giemsa-stained blood films from the type hosts, the
smallest stages detectable are intraerythrocytic trophozoites.
These increase in size and mature into meronts
that undergo transverse or longitudinal pregamontic
binary fission and produce, finally, the paired intraerythrocytic
gamonts, characteristic of the species (Fig.
1 A–J). In L. pholis, transverse and longitudinal binary
fissions of H. bigemina can occur in erythrocytes of the
same blood smear, but the factors that govern these
events and their significance are not clear.
Subsequent to its observation in northern France, H.
bigemina has been recorded from a large number of
fishes, worldwide (see the lists of Becker 1970, Levine
1988, Siddall 1995, and the updated list in Table 1).
These reports include descriptions of H. bigemina from
the white as well as the red blood cell series of fishes,
and at face value they equate to the haemogregarine
having been reported from a remarkable 96 species of
host fishes, in 70 genera and 34 families (Table 1).
However, if these records are scrutinized, they are very
curious. In the type hosts (L. pholis and C. galerita),
only intraerythrocytic development has been reported
(Laveran and Mesnil 1901, Henry 1913, Davies and
Johnston 1976, Davies 1982, Sarasquete and Eiras 1985,
Eiras 1987, Eiras and Davies 1991, Davies et al. 1994)
(see Table 1). Such intraerythrocytic development has
also been observed in another member of the Blenniidae
(Parablennius cornutus) in South Africa (Smit et al.
2003a) and in other members of this family elsewhere
(Table 1). Two additional families of fishes (Clinidae,
Gobiesocidae) from South Africa (Table 1) also exhibit
H. bigemina that develops as in the type hosts (Smit and
Davies 1999, Davies and Smit 2001).
Intraleukocytic development of H. bigemina was first
reported by Laird (1953) from New Zealand fishes of
the Clinidae and Tripterygidae (Table 1). This process
was illustrated from Ericentrus rubrus and involved a
series of merogony and binary fission culminating in the
production of six to eight merozoites in small and large
lymphocytes, and monocytes. Intraerythroblastic and
intraerythrocytic development (illustrated from several
fishes), that apparently followed the intraleukocytic
phase, was like that described for H. bigemina by
Laveran and Mesnil (1901) in the type hosts. Subsequent
to his observations in New Zealand, Laird reported
(Table 1) intraleukocytic development of H.
bigemina from fishes in the South Pacific and New
England (Laird 1958, Laird and Bullock 1969).
By far the majority of sightings of H. bigemina are
those of Saunders (1955, 1958a, 1958b, 1959, 1960,
1964, 1966) (Table 1). While largely supporting Laird’s
observations, many of Saunders’ reports are of concern
because they do not record which stages were present,
or they identify H. bigemina from intraleukocytic forms,
or from immature stages in the red cell series. The
paired mature gamonts, characteristic of the species,
appear largely absent. For example, a report from the
Florida Keys does not record or illustrate the stages
present in this material (Saunders 1958a). The account
of H. bigemina from Bermuda notes development only
in large leukocytes (Saunders 1959). Two further reports
from Florida (Saunders 1955, 1964), and those
from the Bahamas (Saunders 1958b) and Puerto Rico
(Saunders 1966), record development in cells of the
intraleukocytic series and immature stages of the
haemogregarine, or “early gametocytes”, in erythrocytes.
Even among Red Sea samples (Saunders 1960)
the report stated, “more early stages of the parasite were
found than gametocytes”.
Davies et al.: Review of Haemogregarina bigemina
101
Fig. 2. Replete larvae of Gnathia pantherina Smit et Basson,
2002 attached to the gill septum and filaments of Haploblepharus
edwardsii (Voight, 1832). Scale bar = 4 mm.
While it is not impossible that Saunders found H.
bigemina in a wide range of hosts, it is clear that caution
is needed in interpreting her data. In our opinion intraleukocytic
and intraerythrocytic stages that are clearly
immature should not be positively identified as H. bigemina.
We have observed H. bigemina infections in
seven species and six genera of fishes (Blennioclinus
brachycephalus, Chorisochismus dentex, Coryphoblennius
galerita, Clinus cottoides, Clinus superciliosus,
Lipophrys pholis, Parablennius cornutus), across three
families (Blenniidae, Clinidae, Gobiesocidae), from the
UK, Portugal and South Africa (Table 1). We have also
seen H. bigemina-like infections in four species of the
Acanthuridae from Australia (unpublished data, not
recorded in Table 1). Our reports in Table 1 are based
on finding mature paired gamonts, morphometrically
identical to those seen in the type hosts, in erythrocytes.
Where stages similar to Laird’s (1958) intraleukocytic
stages have been observed in the absence of mature
paired gamonts in erythrocytes, as for example in the
intertidal wrasse Symphodus (Crenilabrus) melops from
Wales (see Davies 1982), the identification of H. bigemina
has not been confirmed.
Transmission
The apparent broad distribution of Haemogregarina
bigemina raises questions concerning its transmission
and it is interesting to note in Table 1 that most of its
vertebrate hosts appear to be intertidal or reef-associated
teleosts.
It has been known for some time that the infection
can be detected in very young intertidal fish measuring
under 4 cm in length (Laird 1953, Davies and Johnston
1976, Eiras 1987). In a study of almost 500 Lipophrys
pholis (one of the type hosts) collected at Aberystwyth,
Wales, UK the first patent infections of H. bigemina
were seen in 26% of group 0 fish measuring 3.5 cm TL
(total length) undergoing metamorphosis. Prevalence
was 46% of group 0 fish of 4.0 cm TL and in 94% of
group 0 fish of 4.5 cm TL (Davies and Johnston 1976).
Yearlings (fish of about 5 cm TL) and L. pholis from
group 1 and upwards were all infected with H. bigemina
(Davies and Johnston 1976). These same patterns of
prevalence at Aberystwyth have now been observed
many times at this site over a period spanning almost 30
years (Davies, personal observation). Similar prevalences
for H. bigemina have also been detected among
L. pholis from Foz do Douro, Portugal, except that the
smallest infected fish was 3.2 cm long, prevalence was
87% in fishes of 5–5.9 cm in length and 100% prevalence
was seen in fishes 7.0 cm long and upwards (Eiras
1987). Prevalence was, however, 100% in L. pholis of
6.0 cm and above at other sites in Portugal (Eiras 1987).
The question arises how transmission to such small
fish is effected. Vertical transmission from adult to
young fishes seems unlikely. When 234 hatchling L.
pholis caught at Aberystwyth were kept in isolation for
a period of two months, 42 that survived to reach 3.5 cm
TL or more, did not reveal H. bigemina (see Davies and
Johnston 1976). Faecal-oral transmission was also considered
unlikely at this site (see Davies and Johnston
1976).
Two candidate haematophagous vectors for H.
bigemina have been observed on L. pholis in the UK
(see Davies and Johnston 1976). These are the leech
Oceanobdella blennii (Knight-Jones, 1940) and the
isopod Gnathia maxillaris (Montagu, 1804). However,
O. blennii has never been observed at the study site in
Aberystwyth where H. bigemina occurs commonly.
Furthermore, O. blennii is not known to attach to fish as
small as 3.5 cm TL, and does not feed at a time of year
when young L. pholis are undergoing metamorphosis
(see Davies and Johnston 1976). A general dearth of
leeches has also been reported at other sites where H.
bigemina has been investigated (Eiras 1987, Eiras and
Davies 1991, Davies et al. 1994, Davies and Smit
2001). On the other hand, although Gnathia spp. adults
are free-living, their blood-sucking larvae seem almost
ideal vectors for H. bigemina.
Gnathia spp. larvae are isopods that behave much
like underwater ticks, feeding from the general body
surfaces, buccal cavity and the gills of fishes (Fig. 2).
They become swollen with fish blood when feeding,
normally drop off when replete and digest the blood
meal, moult and then re-attach to fishes, feeding three
times in this manner (see Smit et al. 2003b). Gnathiid
larvae are therefore capable of drawing blood with H.
bigemina from infected fishes and on at least three
occasions during their development. Many of these
isopods are widely distributed in marine environments,
especially intertidally and on reefs, are apparently nonseasonal,
and are readily eaten by many young and adult
fishes (Monod 1926, Davies and Johnston 1976, Cohen
and Poore 1994, Grutter and Poulin 1998, Arnal and
Cote 2000, Sikkel et al. 2000, Davies and Smit 2001).
102
Fig. 3. A – Replete larva of Gnathia sp. Note the region of the inflated anterior hindgut (h). B – Longitudinal histological section
through the posterior cephalosome and anterior pereon of larval gnathiid (Gnathia africana), stained with Masson’s trichrome.
Note the paired digestive glands (g) and the blood-filled anterior hindgut (h). C – Oocyst (arrow) from section through the
digestive glands of G. africana shown in B. Scale bars: A = 1 mm; B = 0.2 mm; C = 10 µm.
Evidence that these isopods actually sustain the development
of H. bigemina is supported by work on
gnathiids and L. pholis in Wales and Portugal (Davies
and Johnston 1976, Davies 1982, 1995, Davies et al.
1994). However, final proof has emerged from studying
H. bigemina, Gnathia africana Barnard, 1914 and the
intertidal fish Clinus superciliosus in South Africa
(Davies and Smit 2001).
Stages observed by sequentially squashing G. africana
that had fed on infected Cl. superciliosus revealed
gamonts of H. bigemina like those seen in the fish host,
normally from 1–6 days post feeding (d.p.f.). Syzygy
also occurred up to 6 d.p.f., immature oocysts from 7
d.p.f. and mature oocysts (sporonts) were observed from
14 d.p.f. Sporogony yielding at least five sporozoites
occurred around 11 d.p.f. and postsporogonic merogony
forming three generations of corresponding merozoites
also began close to 11 d.p.f., with second-generation
merozoites appearing at 18 d.p.f. This developmental
sequence in G. africana, with that observed in the fish
host, formed the basis of a proposed life cycle for H.
bigemina (see Davies and Smit 2001). However, questions
about the cycle remain. The precise sites of
haemogregarine development within gnathiids require
identification, particularly whether merozoites settle in
the salivary glands, and the mode of transmission from
gnathiid to fish needs to be established.
Current research
Attempts to locate the development sites of H. bigemina
in gnathiids now involve sequential histological
studies from conventional wax-embedded material
rather than serial squashes. Initial results indicate that
gamonts, as they are released from the fish erythrocytes
during digestion, accumulate in the inflated anterior
hindgut and some young oocysts, similar to those seen
Davies et al.: Review of Haemogregarina bigemina
103
Fig. 4. Serial histological sections through the cephalosome and anterior pereon of larva of Gnathia maxillaris, later in digestive
cycle than G. africana larva shown in Fig. 3B. A – Uppermost histological section stained with haematoxylin and eosin. Note
salivary glands (s), paired digestive glands (g) and anterior portion of anterior hindgut (h). B – Simple stack of five serial
histological sections (uppermost section, as in A) captured and visualised in IRIS Explorer. C – Rotation of stack imparts depth
to salivary and particularly digestive glands (arrow), beginning the 3D effect. Scale bar: A = 0.2 mm; B and C are not to scale.
by Davies and Johnston (1976) and Davies and Smit
(2001), appear in the paired digestive glands that lie
anterior to the hindgut (Fig. 3). However, if the digestive
processes in gnathiids are similar to those of typical
isopods, then the occurrence of oocysts in the digestive
glands is problematic. In a typical isopod, the contents
of the anterior hindgut (essentially a storage organ) pass
forward to the digestive glands as digestion proceeds.
However, only liquids or fine material can pass the
primary and secondary filtering system of the stomach
to reach the digestive glands (see Wägele 1992). If this
type of digestion also occurs in parasitic gnathiids, then
the developing stages of H. bigemina would be unlikely
to survive passage through such a filtering mechanism.
Clearly, careful study of the internal anatomy and digestive
processes within larval gnathiids is required to discover
how oocysts form in the paired digestive glands.
While the internal anatomy of one gnathiid, Paragnathia
formica (Hesse, 1864) is reasonably well understood
(see Monod 1926, Charmantier 1982), that of
Gnathia spp. is not well known and neither are their
digestive cycles. Serial and sequential histological sections,
used to locate H. bigemina, are therefore being
investigated by 3D visualisation through a system
developed using IRIS Explorer (NAG Ltd., Oxford,
UK). A series of contiguous images is stacked and a
‘render’ module allows adjustment of the angle of
viewing and zoom. A simple stack of five serial sections
is shown in Fig. 4 A, B and on rotation, this gives some
depth to the paired digestive glands (Fig. 4 C). Stacks of
20 or more serial histological sections are also currently
under investigation. These may well permit the entire
internal anatomy of the gnathiid to be represented in this
manner, allowing the sites of development of the
haemogregarine to be clearly defined. It is anticipated
that this technique will also help determine the mode of
passage of H. bigemina from gnathiid to fish. Although
young fishes are known to eat gnathiids, suggesting that
they acquire H. bigemina by predation on these isopods,
transmission to fish by bite from gnathiids cannot yet be
ruled out. It is therefore important to examine the salivary
glands of these isopods in histological sections and
3D representations (Fig. 4) for merozoites of H. bigem-
ina.
104
Table 1. Reports of Haemogregarina bigemina with current, valid names of fish hosts (according to Froese and Pauly 2000),
location in which each host was caught and associated habitat. Also noted are infections (Inf.) in host erythrocytic series (E),
leukocytic series (L), or both (E/L), and the author(s) who first reported the haemogregarine at each location.
Host fish Location Habitat Inf. Author(s)
A c a n t h u r i d a e
Ctenochaetus strigosus (Bennett, 1828) Red Sea Reef E/L Saunders 1960
B a l i s t i d a e
Balistes capriscus Gmelin, 1789 Florida Keys Reef Saunders 1958a
Balistes vetula Linnaeus, 1758 Bahamas Reef E/L Saunders 1958b
B e l o n i d a e
Strongylura notata notata (Poey, 1860) Bahamas;
Southwest Florida
Reef E/L Saunders 1958b, 1964
B l e n n i i d a e
Coryphoblennius galerita (Linnaeus, 1758) France; Portugal Intertidal E Laveran and Mesnil 19011
,
Davies et al. 1994
Ecsenius bicolor (Day, 1888) Heron Island, Australia Reef E Burreson 1989
Lipophrys pholis (Linnaeus, 1758) France; UK; Portugal Intertidal E Laveran and Mesnil 1901,
Henry 1910, Eiras 1984
Parablennius cornutus (Linnaeus, 1758) South Africa Intertidal E Fantham 19302
Parablennius gattorugine (Linnaeus, 1758) UK Intertidal E Henry 1913
Blenniella periophthalmus
(Valenciennes, 1836)
Fiji Reef E Laird 19513
C a r a n g i d a e
Carangoides bartholomaei (Cuvier, 1833) Puerto Rico Reef E Saunders 1966
Carangoides ruber (Bloch, 1793) Bermuda Reef L Saunders 1959
Caranx crysos (Mitchell, 1815) Florida Keys; Bahamas Reef E/L Saunders 1958a, b
Caranx hippos (Linnaeus, 1766) Florida Keys;
Puerto Rico
Reef E Saunders 1958a, 1966
Seriola dumerili (Risso, 1810) Florida Keys Reef Saunders 1958a
Trachinotus falcatus (Linnaeus, 1758) Bermuda Reef L Saunders 1959
C l i n i d a e
Blennioclinus brachycephalus
(Valenciennes, 1836)
South Africa Intertidal E Davies et al.4
Clinus cottoides Valenciennes, 1836 South Africa Intertidal E Smit and Davies 1999
Clinus superciliosus (Linnaeus, 1758) South Africa Intertidal E Smit and Davies 1999
Ericentrus rubrus (Hutton, 1872) North Island,
New Zealand
Intertidal E/L Laird 1953
Heteroclinus perspicillatus
(Valenciennes, 1836)
Norfolk Island,
South Pacific
Reef E/L Laird 1958
C o r y p h a e n i d a e
Coryphaena hippurus Linnaeus, 1758 Florida Keys; Bermuda Pelagic L Saunders 1958a, 1959
C o t t i d a e
Artedius fenestralis Jordan et Gilbert, 1883 Vancouver Island,
Canada
Intertidal E Laird 1961
G e r r e i d a e
Eucinostomus gula
(Quoy et Gaimard, 1824)
Bahamas Reef E/L Saunders 1958b
Gerres cinereus (Walbaum, 1792) Southwest Florida Reef E/L Saunders 1964
G o b i e s o c i d a e
Chorisochismus dentex (Pallas, 1769) South Africa Intertidal E Davies and Smit 2001
Trachelochismus melobesia Phillips, 1927 North Island,
New Zealand
Intertidal E Laird 1953
G o b i i d a e
Amblygobius albimaculatus (Rüppell, 1830) Red Sea Reef E/L Saunders 1960
Bathygobius soporator
(Valenciennes, 1837)
Bahamas Intertidal E/L Saunders 1958b
H a e m u l i d a e
Haemulon album Cuvier, 1830 Florida Keys; Bahamas Reef E/L Saunders 1958a, b
Haemulon aurolineatum Cuvier, 1830 Puerto Rico Reef E Saunders 1966
Haemulon flavolineatum (Desmarest, 1823) Florida Keys Reef Saunders 1958a
Haemulon plumierii (Lacépède, 1801) Florida Keys Reef Saunders 1958a
Haemulon sciurus (Shaw, 1803) Florida Keys; Bahamas Reef E/L Saunders 1958a, b
Davies et al.: Review of Haemogregarina bigemina
105
Table 1. Continued.
Host fish Location Habitat Inf. Author(s)
H e m i r a m p h i d a e
Hemiramphus brasiliensis (Linnaeus, 1758) Puerto Rico Reef E Saunders 1966
Hyporhamphus unifasciatus (Ranzani, 1842) Bermuda Reef L Saunders 1959
I s t i o p h o r i d a e
Istiophorus albicans (Latreille, 1804) Florida Keys Pelagic Saunders 1958a
K y p h o s i d a e
Kyphosus bigibbus Lacépède, 1801 Red Sea Reef E/L Saunders 1960
L a b r i d a e
Cheilinus trilobatus Lacépède, 1801 Red Sea Reef E/L Saunders 1960
Halichoeres bivittatus (Bloch, 1791) Bermuda Reef L Saunders 1959
Lachnolaimus maximus (Walbaum, 1792) Florida Keys Reef Saunders 1958a
Pteragogus pelycus Randall, 1981 Red Sea Reef E/L Saunders 1960
Thalassoma bifasciatum (Bloch, 1791) Bahamas Reef E/L Saunders 1958b
Thalassoma purpureum (Forsskål, 1775) Red Sea Reef E/L Saunders 1960
L e t h r i n i d a e
Lethrinus mahsena (Forsskål, 1775) Red Sea Reef E/L Saunders 1960
Lethrinus nebulosus (Forsskål, 1775) Red Sea Reef E/L Saunders 1960
Lethrinus variegatus Valenciennes, 1830 Red Sea Reef E/L Saunders 1960
L u t j a n i d a e
Lutjanus apodus (Walbaum, 1792) Bahamas; Puerto Rico Reef E/L Saunders 1958b, 1966
Lutjanus bohar (Forsskål, 1775) Red Sea Reef E/L Saunders 1960
Lutjanus griseus (Linnaeus, 1758) Florida Keys Reef Saunders 1958a
Lutjanus synagris (Linnaeus, 1758) Florida Keys; Bahamas Reef E/L Saunders 1958a, b
Ocyurus chrysurus (Bloch, 1791) Florida Keys; Bahamas;
Puerto Rico
Reef E/L Saunders 1958a, b, 1966
M a l a c a n t h i d a e
Malacanthus plumieri (Bloch, 1786) Florida Keys; Bahamas Reef E/L Saunders 1958a, b
M u g i l i d a e
Mugil trichodon Poey, 1875 Bahamas Marine, brackish
and freshwater
E/L Saunders 1958b
M u l l i d a e
Parupeneus forsskali
(Fourmanoir et Guézé, 1976)
Red Sea Reef E/L Saunders 1960
Upeneus tragula Richardson, 1846 Red Sea Reef E/L Saunders 1960
M u r a e n i d a e
Gymnothorax funebris Ranzani, 1840 Florida Keys Reef Saunders 1958a
P i n g u i p e d i d a e
Parapercis hexophtalma (Cuvier, 1829) Red Sea Reef E/L Saunders 1960
P o m a c a n t h i d a e
Pomacanthus maculosus (Forsskål, 1775) Red Sea Reef E/L Saunders 1960
P o m a c e n t r i d a e
Abudefduf saxatilis (Linnaeus, 1758) Bahamas Reef E/L Saunders 1958b
S c a r i d a e
Chlorurus sordidus (Forsskål, 1775) Red Sea Reef E/L Saunders 1960
Scarus iseri (Bloch, 1789) Puerto Rico Reef E Saunders 1966
Sparisoma aurofrenatum
(Valenciennes, 1840)
Puerto Rico Reef E Saunders 1966
S c i a e n i d a e
Bairdiella chrysoura (Lacépède, 1802) Southwest Florida Marine, brackish
and freshwater
E/L Saunders 1964
Menticirrhus littoralis (Holbrook, 1855) Atlantic Coast Florida;
Southwest Florida
Marine and
brackish
E/L Saunders 1955, 1964
S c o m b r i d a e
Auxis thazard thazard (Lacépède, 1800) Puerto Rico Pelagic E Saunders 1966
Scomberomorus regalis (Bloch, 1793) Florida Keys Reef Saunders 1958a
Scomberomorus cavalla (Cuvier, 1829) Florida Keys Reef Saunders 1958a
106
Table 1. Continued.
Host fish Location Habitat Inf. Author(s)
S e r r a n i d a e
Centropristis striata (Linnaeus, 1758) Eastern Canada;
New England
Reef E/L Fantham et al. 1942,
Laird and Bullock 1969
Cephalopholis hemistiktos (Rüppell, 1830) Red Sea Reef E/L Saunders 1960
Cephalopholis miniata (Forsskål, 1775) Red Sea Reef E/L Saunders 1960
Epinephelus adscensionis (Osbeck, 1765) Florida Keys Reef Saunders 1958a
Epinephelus fasciatus (Forsskål, 1775) Red Sea Reef E/L Saunders 1960
Epinephelus fuscoguttatus (Forsskål, 1775) Red Sea Reef E/L Saunders 1960
Epinephelus guttatus (Linnaeus, 1758) Florida Keys; Bermuda Reef L Saunders 1958a, 1959
Epinephelus morio (Valenciennes, 1828) Florida Keys Reef Saunders 1958a
Epinephelus striatus (Bloch, 1792) Florida Keys; Bermuda Reef L Saunders 1958a, 1959
Epinephelus summana (Forsskål, 1775) Red Sea Reef E/L Saunders 1960
Epinephelus tauvina (Forsskål, 1775) Red Sea Reef E/L Saunders 1960
Mycteroperca bonaci (Poey, 1860) Florida Keys; Bermuda Reef L Saunders 1958a, 1959
Mycteroperca microlepis
(Goode et Bean, 1879)
Florida Keys Reef Saunders 1958a
Plectropomus maculatus (Bloch, 1790) Red Sea Reef E/L Saunders 1960
Variola louti (Forsskål, 1775) Red Sea Reef E/L Saunders 1960
S p a r i d a e
Acanthopagrus bifasciatus (Forsskål, 1775) Red Sea Reef E/L Saunders 1960
Argyrops spinifer (Forsskål, 1775) Red Sea Range of marine
habitats
E/L Saunders 1960
Calamus bajonado
(Bloch et Schneider, 1801)
Florida Keys; Bahamas Reef E/L Saunders 1958a, b
Rhabdosargus haffara (Forsskål, 1775) Red Sea Reef E/L Saunders 1960
Lagodon rhomboides (Linnaeus, 1766) Southwest Florida Marine, brackish
and freshwater
E/L Saunders 1964
S p h y r a e n i d a e
Sphyraena barracuda (Walbaum, 1792) Florida Keys; Bahamas;
Bermuda
Reef E/L Saunders 1958a, b, 1959
S y n o d o n t i d a e
Synodus japonicus5
Red Sea E/L Saunders 1960
T r i p t e r y g i i d a e
Bellapiscis medius (Günther, 1861) North Island,
New Zealand
Intertidal E Laird 1953
Enneapterygius rufopileus (Waite, 1904) Norfolk Island,
South Pacific
Reef E/L Laird 1958
Forsterygion varium (Forster, 1801) North Island,
New Zealand
Intertidal E Laird 1953
Notoclinus fenestratus (Forster, 1801) North Island,
New Zealand
Intertidal E Laird 1953
Z o a r c i d a e
Zoarces americanus
(Bloch et Schneider, 1801)
Atlantic Coast, Canada Intertidal Fantham et al. 1942 6
Zoarces viviparus (Linnaeus, 1758) UK Intertidal Bentham 19177
1
Laveran and Mesnil (1901) reported H. bigemina from Blennius pholis (valid name Lipophrys pholis) and Blennius gattorugine.
However, Laveran and Mesnil (1902) amended their original B. gattorugine to Blennius montagui (valid name Coryphoblennius
galerita);
2
Described by Fantham (1930) as Haemogregarina fragilis, this was formally recognised as H. bigemina by Smit et al. (2003);
3
Described by Laird (1951) as Haemogregarina salariasi, this was formally recognised as H. bigemina by Siddall (1995);
4
Paired mature gamonts of H. bigemina in the erythrocytes of B. brachycephalus were observed by AJD, NJS and PMH at De
Hoop Nature Reserve, South Africa in 2003, a new host record for the haemogregarine;
5
Appears to be a nomen nudum;
6
A haemogregarine, probably Haemogregarina bigemina, according to Fantham et al. (1942);
7
Reported in Laird (1953), but we were unable to trace the reference.
Davies et al.: Review of Haemogregarina bigemina
107
FUTURE RESEARCH
Future developments in research on Haemogregarina
bigemina will obviously centre on continued attempts to
reconstruct the internal anatomy of larval gnathiids from
histological sections and to locate the development sites
of the haemogregarine. However, does this haemogregarine
really develop in as many fish hosts as Table 1
suggests? Biological transmission to clean fishes under
laboratory conditions should be, therefore, another
logical aim for the future. The apparent broad distribution
of H. bigemina among fishes, the ease with which
gnathiids can be persuaded to feed on these hosts (see
Smit et al. 2003b) and the fact that many fishes eat
gnathiids, may all aid attempts at transmission. Another
important question is, does H. bigemina truly have an
intraleukocytic phase or are mixed infections involved?
Transmission studies may solve this problem, but another
future aim should be to compare the molecular
constitution of so-called H. bigemina from a number of
locations to determine whether all samples represent the
same species.
Haemogregarina bigemina is clearly an extraordinary
and intriguing apicomplexan, seemingly unlike
other fish haemogregarines in its distribution and transmission.
Its known development most closely resembles
that of Haemogregarina (sensu stricto), but the match is
not perfect (see Davies and Smit 2001). When answers
to some of the questions posed in this paper have been
resolved, it may be that H. bigemina, instead of its
current placing with Haemogregarina (sensu lato), will
be deserving of a genus in its own right.
Acknowledgements. AJD is immensely grateful to Dr. Mike
Johnston and Prof. Jorge Eiras for their friendship, good humour
and support in research on H. bigemina over many
years. AJD also wishes to record her gratitude for current
funding from a Leverhulme Trust Research Fellowship and a
study abroad grant from The Royal Society. NJS acknowledges
funding from the Claude Harris Leon Foundation.
REFERENCES
ARNAL C., COTE I.M. 2000: Diet of broadstripe cleaning
gobies on a Barbadian reef. J. Fish Biol. 57: 1075–1082.
BARTA J.R. 2000: Suborder Adeleorina Léger, 1911. In: J.J.
Lee, G.F. Leesdale and P. Bradbury (Eds.), The Illustrated
Guide to the Protozoa. Vol. I. Second Edition. Allen Press,
Lawrence, Kansas, pp. 305–318.
BECKER C.D. 1970: Haematozoa of fishes, with emphasis on
North American records. In: S.F. Sniesko (Ed.), A Symposium
on Diseases of Fishes and Shellfishes. Am. Fish.
Soc. Spec. Publ. No. 5, pp. 82–100.
BURRESON E.M. 1989: Haematozoa of fishes from Heron I.,
Australia, with the description of two new species of
Trypanosoma. Aust. J. Zool. 37: 15–23.
CHARMANTIER G. 1982: Les glandes céphaliques de Paragnathia
formica (Hesse, 1864) (Isopoda, Gnathiidae):
localization et ultrastructure. Crustaceana 42: 179–193.
COHEN B.F., POORE G.C.B. 1994: Phylogeny and biogeography
of the Gnathiidae (Crustacea: Isopoda) with descriptions
of new genera and species, most from southeastern
Australia. Mem. Mus. Victoria 54: 271–397.
DAVIES A.J. 1982: Further studies on Haemogregarina bigemina
Laveran & Mesnil, the marine fish Blennius pholis
L., and the isopod Gnathia maxillaris Montagu. J. Protozool.
29: 576–583.
DAVIES A.J. 1995: The biology of fish haemogregarines.
Adv. Parasitol. 36: 118–203.
DAVIES A.J., EIRAS J.C., AUSTIN R.T.E. 1994: Investigations
into the transmission of Haemogregarina bigemina
Laveran & Mesnil, 1901 (Apicomplexa: Adeleorina) between
intertidal fishes in Portugal. J. Fish Dis. 17: 283–
289.
DAVIES A.J., JOHNSTON M.R.L. 1976: The biology of
Haemogregarina bigemina Laveran & Mesnil, a parasite
of the marine fish Blennius pholis Linnaeus. J. Protozool.
23: 315–320.
DAVIES A.J., JOHNSTON M.R.L. 2000: The biology of
some intraerythrocytic parasites of fishes, amphibia and
reptiles. Adv. Parasitol. 45: 1–107.
DAVIES A.J., SMIT N.J. 2001: The life cycle of Haemogregarina
bigemina (Adeleina: Haemogregarinidae) in
South African hosts. Folia Parasitol. 48: 169–177.
EIRAS J.C. 1984: Virus infection of marine fish: prevalence
of viral erythrocytic necrosis (VEN) in Mugil cephalus L.,
Blennius pholis L. and Platichthys flesus L. in coastal
waters of Portugal. Bull. Eur. Assoc. Fish Pathol. 4: 52–
56.
EIRAS J.C. 1987: Occurrence of Haemogregarina bigemina
(Protozoa: Apicomplexa) in Blennius pholis (Pisces:
Blenniidae) along the Portuguese west coast. J. Fish Biol.
30: 597–603.
EIRAS J.C., DAVIES A.J. 1991: Haemogregarina bigemina
Laveran & Mesnil, 1901 (Protozoa, Apicomplexa) from
Blennius pholis L. (Osteichthyes, Blenniidae): an investigation
of seasonality in Portugal. J. Fish Dis. 14: 683–687.
FANTHAM H.B. 1930: Some parasitic protozoa found in
South Africa. – XIII. S. Afr. J. Sci. 27: 376–390.
FANTHAM H.B., PORTER A., RICHARDSON L.R. 1942:
Some haematozoa observed in vertebrates in Eastern
Canada. Parasitology 32: 199–266.
FROESE R., PAULY D. 2000: FishBase 2000: Concepts,
Design and Data Sources. ICLARM, Los Baños, Laguna,
Philippines, 344 pp.
GRUTTER A.S., POULIN R. 1998: Intraspecific and
interspecific relationships between host size and the
abundance of parasitic larval gnathiid isopods on coral
reef fishes. Mar. Ecol. Prog. Ser. 164: 263–271.
HENRY H. 1910: On the haemoprotozoa of British sea-fish.
(A preliminary note.). J. Pathol. Bacteriol. 14: 463–465.
HENRY H. 1913: A summary of the blood parasites of British
sea-fish. J. Pathol. Bacteriol. 18: 218–223.
108
LAINSON R. 1981: On Cyrilia gomesi (Neiva & Pinto, 1926)
gen. nov. (Haemogregarinidae) and Trypanosoma bourouli
Neiva & Pinto, in the fish Synbranchus marmoratus:
simultaneous transmission by the leech Haementeria lutzi.
In: E.U. Canning (Ed.), Parasitological Topics. A presentation
volume to P.C.C. Garnham, FRS on the occasion of
his 80th birthday 1981. Allen Press for the Society of
Protozoologists (Special Publications No. 1), Lawrence,
Kansas, pp. 150–158.
LAIRD M. 1951: A contribution to the study of Fijian haematozoa.
Zool. Publ. Victoria Univ. Coll., Wellington, No.
10, 1–15.
LAIRD M. 1953: The protozoa of New Zealand intertidal
zone fishes. Trans. R. Soc. N. Z. 81: 79–143.
LAIRD M. 1958: Parasites of South Pacific fishes. 1. Introduction,
and haematozoa. Can. J. Zool. 36: 153–165.
LAIRD M. 1961: Trichodinids and other parasitic Protozoa
from the intertidal zone at Nanaimo, Vancouver Island.
Can. J. Zool. 39: 833–844.
LAIRD M., BULLOCK W.L. 1969: Marine fish haematozoa
from New Brunswick and New England. J. Fish. Res.
Board Can. 26: 1075–1102.
LAVERAN A., MESNIL F. 1901: Deux hémogrégarines
nouvelles des poissons. C. R. Acad. Sci. Paris 133: 572–
577.
LAVERAN A., MESNIL F. 1902: Sur les hématozoaires des
poissons marins. C. R. Acad. Sci. Paris 135: 567–570.
LEVINE N.D. 1988: The Protozoan Phylum Apicomplexa.
Vol. 1. CRC Press, Boca Raton, Florida, 203 pp.
MONOD T. 1926: Les Gnathiidae. Essai monographique
(Morphologie, Biologie, Systematique). Mem. Soc. Sci.
Nat. Maroc 13: 1–668.
NEGM-ELDIN M.M. 1999: Life cycle, host restriction and
longevity of Cyrilia nili (Haemogregarina nili Wenyon,
1909) n. comb. Dtsch. Tierärztl. Wochenschr. 106: 191–
199.
SARASQUETE M.C., EIRAS J.C. 1985: Hemoprotozoosis en
una población de Blennius pholis (L., 1758) de las costas
de Oporto (Portugal). Invest. Pesq. 49: 627–635.
SAUNDERS D.C. 1955: The occurrence of Haemogregarina
bigemina Laveran and Mesnil and H. achiri n. sp. in
marine fish from Florida. J. Parasitol. 41: 171–176.
SAUNDERS D.C. 1958a: Blood parasites of the marine fishes
of the Florida Keys. Year Book, Amer. Philos. Soc. 1958:
261–266.
SAUNDERS D.C. 1958b: The occurrence of Haemogregarina
bigemina Laveran and Mesnil, and H. dasyatis n. sp. in
marine fish from Bimini, Bahamas, B.W.I. Trans. Am.
Microsc. Soc. 77: 404–412.
SAUNDERS D.C. 1959: Haemogregarina bigemina Laveran
and Mesnil, from marine fishes of Bermuda. Trans. Am.
Microsc. Soc. 78: 374–379.
SAUNDERS D.C. 1960: A survey of the blood parasites in the
fishes of the Red Sea. Trans. Am. Microsc. Soc. 79: 239–
252.
SAUNDERS D.C. 1964: Blood parasites of marine fishes of
southwest Florida, including a new haemogregarine from
the menhaden, Brevoortia tyrannus (Latrobe). Trans. Am.
Microsc. Soc. 83: 218–225.
SAUNDERS D.C. 1966: A survey of the blood parasites of
the marine fishes of Puerto Rico. Trans. Am. Microsc.
Soc. 85: 193–199.
SIDDALL M.E. 1995: Phylogeny of adeleid blood parasites
with a partial systematic revision of the haemogregarine
complex. J. Eukaryot. Microbiol. 42: 116–125.
SIKKEL P.C., FULLER C.A., HUNTE W. 2000: Habitat/sex
differences in time at cleaning stations and ectoparasite
loads in a Caribbean reef fish. Mar. Ecol. Prog. Ser. 193:
191–199.
SMIT N.J., BASSON L., VAN AS J.G. 2003b: Life cycle of
the temporary fish parasite, Gnathia africana (Crustacea:
Isopoda: Gnathiidae). Folia Parasitol. 50: 135–142.
SMIT N.J., DAVIES A.J. 1999: New host records for Haemogregarina
bigemina from the coast of southern Africa. J.
Mar. Biol. Assoc. U.K. 79: 933–935.
SMIT N.J., VAN AS J.G., DAVIES A.J. 2003a: Taxonomic
re-evaluation of the South African fish haemogregarine,
Desseria fragilis. J. Parasitol. 89: 151–153.
WÄGELE J.-W. 1992: Chapter 11. Isopoda. In: F.W. Harrison
and A.G. Humes (Eds.), Microscopic Anatomy of Invertebrates.
Vol. 9, Crustacea. Wiley-Liss, New York, pp. 529–
618.
Received 1 December 2003 Accepted 27 April 2004
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