[page 66] [International Journal of Plant Biology 2010; 1:e13]
A simplified method
for differential staining
of aborted and non-aborted
pollen grains
Ross Peterson,1
Janet P Slovin,2
Changbin Chen1
1
Department of Horticultural Science,
University of Minnesota, MN, USA;
2
Genetic Improvement of Fruits and
Vegetables Laboratory, United States
Department of Agriculture, Baltimore,
MD, USA
Abstract
The ability to use chemical staining to discriminate
aborted from non-aborted pollen
grains has well-known practical applications in
agriculture. A commonly used technique for
assessing pollen vitality, Alexander’s stain,
uses chloral hydrate, phenol and mercuric
chloride, all of which are highly toxic. We
describe here an improved pollen staining
technique that avoids the use of a regulated
chemical chloral hydrate and two extremely
toxic chemicals mercuric chloride and phenol,
and requires a much shorter time period for
sample preparation and staining. This simplified
method is very useful for field studies
without high-end equipments such as fluorescence
microscopes. Samples can be collected
and fixed in the fields and examined in a simple
laboratory that has light microscopes.
Introduction
A facile assay for determination of the
vitality of pollen grains is important for plant
breeders as well as for biologists interested
in development of the male gametophyte.
Pollen germination assays frequently require
optimization as well as being time consuming
and difficult to reproduce.1
A simple rapid
staining technique that clearly differentiates
aborted from non-aborted pollen would provide
needed information and improve
throughput. For many years, nuclear dyes
such as acetocarmine and cotton blue have
been used for staining pollen, as have the
vital dyes triphenyl tetrazolium chloride
(TTC) and tetrazolium red. TTC and other
vital dyes have been found to stain aborted
pollen grains to varying degrees.2-4
Some fluorescent
dyes have been used to differentiate
aborted and non-aborted pollen grains, which
include a most commonly used dye of fluorescein
diacetate that was initially developed to
examine the viability of mammalian cells.5,6
The uses of fluorescent dyes, however,
require special equipment, such as fluorescence
microscopes, which limit the application
for field studies and most agricultural
extension stations that do not have the available
equipment.
A major improvement in differential pollen
staining was reported by M.P. Alexander in
1969. Alexander’s stain colors aborted pollen
grains from most angiosperms and the
spores of gymnosperms blue-green, and nonaborted
pollen grains and spores stain
magenta-red.7
Unfortunately, three ingredients
of Alexander’s protocol exhibit possible
health risks. Two of these reagents are chloral
hydrate that is used in the stain solution
and the mercuric chloride that is used in the
fixative. Both are either highly regulated by
worldwide government standards on chemical
toxicities or threaten the health of those
who use the stain solution.8
The third harmful
reagent is phenol. It can be absorbed
through the skin and has shown to disrupt
the function of the central nervous system.9
Another drawback to Alexander’s protocol is
the long time it requires to obtain clear
results. Thus, it is not a very efficient method
and does not allow for acquisition of data in a
timely manner. In addition, the protocol calls
for optimization of the recipes to penetrate
oily, spiny, thick, or thin coatings around the
pollen and for staining pollen in non-dehiscent
anthers. Due to the protocol variations,
experimental results obtained in different
laboratories have proven to be inconsistent.
Some researchers have obtained excellent
results, with differential coloration reflecting
the results in Alexander’s original report.10-12
In some cases, the pollen grains were able to
be distinguished, but the images were not
clear due to non-transparent anther walls.13
In other cases, the anther wall was sufficiently
transparent and the non-aborted
pollen grains stained red, but the aborted
pollen grains were colorless or only very pale
green.14-16
Such variability can be explained by
differences in preparation of the tissues, differences
in local environmental conditions
during staining, or differences in the source
of chemical ingredients.
One additional problem with Alexander’s
protocol is limited access to chloral hydrate,
which requires restricted federal and state
licenses for purchase in the United States,
that also incur additional costs to the
research laboratory. We report here a simplified
staining protocol that does not use chloral
hydrate, mercuric chloride or phenol and
yet rapidly produces a clear differentiation
between aborted and non-aborted pollen,
even within non-dehiscent anthers.
Materials and Methods
Plants
A total of 22 plant species and 3 different
Arabidopsis lines were tested (Table 1).
Arabidopsis, strawberry (Fragaria vesca
Coville), tomato (Solanum lycopersicum L.)
and rice (Oryza sativa L.) plants were grown in
the greenhouse or growth chambers. Other
plants tested were grown outdoors under ambient
conditions in Minnesota, USA.
Bud fixation
Flower buds or free anthers were collected
before anthesis, when pollen was mature but
anthers non-dehiscent (stage 12 flower buds) ,17
International Journal of Plant Biology 2010; volume 1:e13
Correspondence: Changbin Chen, Department
of Horticultural Science, University of Minnesota
380 Alderman Hall, 1970 Folwell Avenue, St. Paul,
MN 55108, USA. E-mail: chenx481@umn.edu
Key words: pollen staining, Alexander’s stain,
chloral hydrate, phenol.
Contributions: RP did most of experiments and
contributed to the writing; JPS did further test
and contributed to edit the manuscript; CC initiated
this study, supervised RP for his bench-work
and data collection and wrote this manuscript.
Acknowledgements: we deeply appreciated the
comments from all three reviewers, which greatly
improved the final protocol with the removal of
phenol. The authors are grateful to Dr. Luca Comai
for providing Arabidopsis autotetraploid seeds and
Dr. Dong-Hoon Jeong for providing rice
Nipponbare seeds. Our sincere thanks also go to
Tao Li and Asmita Batajoo for plant care, Dr.
Junhua Li and the former colleagues, Drs. Cary H
Hord, Yoshitaka Azumi and Wuxing Li at the
Pennsylvania State University for sharing their
experiences with us. This study was supported by
funds from the American Society of Plant Biologists
SURF program and the University of Minnesota
UROP program to RP and by funds from the USDABRDC,
Dow Agrosciences, and University of
Minnesota Academic Health Center to CC.
Conflict of interest: the authors report no conflicts
of interest.
Received for publication: 19 March 2010.
Revision received: 16 July 2010.
Accepted for publication: 16 July 2010.
This work is licensed under a Creative Commons
Attribution 3.0 License (by-nc 3.0).
©Copyright R. Peterson et al., 2010
Licensee PAGEPress, Italy
International Journal of Plant Biology 2010; 1:e13
doi:10.4081/pb.2010.e13
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[International Journal of Plant Biology 2010; 1:e13] [page 67]
and fixed in Carnoy’s fixative (6 alcohol:3 chloroform:1
acetic acid)18
for a minimum of 2
hours. Buds can be stored in fixative for 12
months at either room temperature or cold
room. For large buds, such as Rhododendron,
Tulipa or Magnolia, individual anthers were dissected
from the stage 13 flower bud and fixed.
Stain solution
The final stain solution we used was prepared
by adding the following constituents in
the order given below and stored in the dark.
10 mL 95% alcohol
1 mL Malachite green (1% solution in
95% alcohol)
50 mL Distilled water
25 mL Glycerol
5 mL Acid fuchsin (1% solution in water)
0.5 mL Orange G (1% solution in water)
4 mL Glacial acetic acid
Add distilled water (4.5 mL) to a total of 100 mL.
Staining
Following at least two hours of fixation, the
bud can be placed on a microscope slide and
the fixative’s liquid was thoroughly and carefully
dried from the plant material with
absorbent paper. Proper safety gloves should
be worn to avoid the risk of chloroform from
being absorbed through the skin. Apply 2-4
drops of the stain solution before the sample
completely dries. If flower buds have been collected
instead of free anthers, the buds should
be dissected to release the anthers and pollen.
Under a dissecting microscope the leftover
plant debris can be carefully removed. To save
stain solution, samples can be dissected prior
to putting individual anthers into stain. Some
anthers, such as those of Magnolias, are too
large to be viewed intact and must be dissected
further.
Once the sample is in the stain, slowly heat
the slide over an alcohol burner in a fume hood
until the stain solution is near boiling (~30
seconds). A more moderate rate of heating
allows better penetration of the dye into the
cellulose and protoplasm of the pollen.
Extremely high temperatures resulting in
smoking or bubbling of the stain can burn the
dye and the sample. Heating can be adjusted by
briefly moving the slide in and out of the flame.
To ensure stain has been completely absorbed
into the pollen grains, 10 to 15 minutes should
be allowed for some species such as Lonicera
tatarica, Ginkgo biloba, Pinus resinosa and
Rhododendron mucronulatum.
Imaging
Place a cover-slip over the sample and apply
even pressure on the cover-slip to ensure that
all plant components converge to one plane.
The cover-slip can be sealed using nail polish
or wax. Slides were examined using a Leitz
microscope (Ernst Leitz Wetzlar GmbH,
Germany). Micrographs were taken using a
Spot Insight digital camera (Diagnostic instruments,
Inc. Sterling Heights, MI, USA) and
edited with Adobe Photoshop CS2 (Adobe
Systems Inc. CA, USA).
Results and Discussion
The simplified stain resulted in excellent
differentiation of pollen in all 22-plant species
tested (Table 1). Photographs of 13 out of 22
examined species are included in Figure 1, all
of which demonstrated clear differentiation of
aborted and non-aborted pollen grains, except
the Arabidopsis Ler ecotype that showed all
non-aborted pollen grains (Figure 1D). To represent
different types of pollen and/or anther
structures, both Angiosperms and Gymno sperms
were studied. To exemplify these different
morphologies certain species are shown
in Figure 1. Of the Angiosperm species, strawberry
[Fragaria vesca (Rugen)] has thick
anther walls and small pollen grains (Figure
1H); woody honeysuckle (Lonicera tatarica)
has thick pollen walls (Figure 1I); azalea
(Rhododendron mucronulatum) shows tetrad
pollen structure (Figure 1L); rice [Oryza sativa
(Nipponbare)] has thin pollen walls (Figure
1J); dandelion (Taraxacum officinale) has
thick and oily pollen walls (Figure 1N) and
tulip (Tulipa spp) has larger pollen grains
(Figure 1O). In all tested samples, non-aborted
pollen grains stained magenta-red and aborted
pollen grains stained blue-green. This differentiation
can be seen in Arabidopsis autotetraploid
(Figure 1E). No significant differences
in the stain clarity were observed with
thick or thin walled pollen grains, or with oily
or waxy pollen. Pollen within non-dehiscent
anthers (Figure 1 C, D, E, H, K, N) stained as
well as free pollen (Figure 1 A, B, G, I), even
without the mercuric chloride and phenol used
in the original Alexander’s staining prepara-
tion.7,19
As a supplementary improvement for the
safety and accessibility of this improved staining
technique, phenol was avoided. The origi-
Article
Table 1. A list of plant species tested.
Divisions Species name Fig. Non-aborted Non-aborted
(family) pollen (%)* pollen (%)**
Gymnosperm Ginkgo biloba L. (Ginkgoaceae) 1A 94.5 97.5
Pinus resinosa Ait. (Pinaceae) 1B 95.5 93.0
Angiosperm Acer negundo L. (Aceraceae) N/A 95.5 96.0
Acer rubrum L. (Aceraceae) 1C, 1F 99.0 98.5
Alnus incana (L.) Moench. (Betulaceae) N/A 92.5 90.5
Arabidopsis thaliana (L.)
Heynh. (Ler ecotype: diploid, 1D 98.5 95.5
tetraploid, and dmc1 mutants) 1E 66.5 72.5
(Brassicaceae) N/A 15.5 22.0
Berberis thunbergi DC. (Berberidaceae) N/A 87.0 98.5
Betula populifolia Marsh. (Betulaceae) 1G 97.5 99.0
Fragaria vesca Coville cv. Rugen (Rosaceae) 1H 83.5 82.0
Lonicera tatarica L. (Caprifoliaceae) 1I 90.5 93.5
Magnolia stellata (Siebold & Zucc.) N/A 90.5 95.0
Maxim. (Magnoliaceae)
Malus floribunda Siebold ex Van Houtte N/A 94.0 94.5
(Rosaceae)
Narcissus spp. L. (Amarylliaceae) N/A 95.0 92.5
Oryza sativa L. cv. Nipponbare (Poaceae) 1J 93.0 91.0
Populus deltoids W. Bartram ex Marshall N/A 96.5 97.0
(Salicaceae)
Prunus padus L. (Rosaceae) 1K 90.5 88.5
Quercus rubra L. (Fagaceae) N/A 81.0 80.0
Rhododendron mucronulatum Turcz. 1L 64.5 82.5
(Ericaceae)
Scilla siberica Haw. (Liliaceae) N/A 92.0 91.0
Solanum lycopersicum L. (Solanaceae) 1M 76.5 83.5
Taraxacum officinale F.H. Wigg. (Asteraceae) 1N 73.5 80.0
Tulipa spp. L. (Liliaceae) 1O 52.5 64.0
*The stain without phenol; **the stain with phenol.
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[page 68] [International Journal of Plant Biology 2010; 1:e13]
nal use of phenol (5%) was to help make the
stain on the slides more transparent. As a
result of this transparency, the differential
staining would theoretically become clearer.
However, once quantitative and qualitative
experiments were done in the absence of phenol,
it became evident that phenol was unnecessary
for clear differential staining (Figure 1
C, F). A minor difference was noted only for
pollens inside the non-dehisced anthers.
Despite this small difference, no significant
change of the staining of non-aborted pollen
grains was observed when comparing the
stains with or without phenol (as shown in
Table 1). These data agree with the notion that
the stain without phenol is as credible as the
stain with phenol. Thus, the omission of phenol
in this pollen staining technique is recom-
mended.
Alexander’s protocol for differential staining
of aborted and non-aborted pollen is a frequently
cited technique because of its usability
for many different applications. However,
the availability of chemicals used for the stain,
the variability inherent in protocols that need
to be optimized for each plant and the amount
of time required to obtain results, limit the
protocol’s applicability and the reproducibility.
In particular, the need to use chloral hydrate
requires licenses that may be difficult or
expensive to obtain in some countries. The
simplified stain described here, using the
stain solution with Carnoy’s fixative, allows for
the use of the protocol in both laboratory and
field studies. The procedure can be used for all
types of pollen grains. The 2-hour incubation
in the chloroform and acetic acid containing
fixative sufficiently removes oils or sticky
materials and clears pollen and anther walls.7
Additional heating (1-2 min.) or incubation
time (10-15 min.) in the stain, for certain
species listed previously, may be necessary for
optimal stain absorption. There is no additional
chemicals (chloral hydrate or phenol) needed
to clear the sample, and no specific fixative
needed either. The simplification to using
Carnoy’s fixative also allows for longer fixation
prior to staining, facilitating field studies or
studies with large numbers of samples. The
entire procedure takes a little more than 2
hours, eliminating the long incubations and
the optimization of steps required in the
Alexander’s stain.7,20
References
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Figure 1. Differential staining of aborted and non-aborted pollen grains. (A) Ginkgo biloba.
(B) Pinus resinosa. (C) Acer rubrum, stained without phenol. (D) Arabidopsis
thaliana Ler (diploid). (E) Arabidopsis thaliana Ler (autotetraploid). (F) Acer rubrum,
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