CC BY
27
Protistology 13 (2), 57-63 (2019) Protistology
Ultrastructural aspects of ecdysis in the naked dinoflagellate Amphidinium carterae
Mariia Berdieva, Pavel Safonov and Olga Matantseva
Institute of Cytology, Russian Academy of Sciences, St. Petersburg, Russia
\ Submitted April 09, 2019 | Accepted April 29, 2018 | Summary
The stressor-induced ecdysis takes a special place in dinoflagellate biology. During ecdysis, a cell loses the plasmalemma, outer amphiesmal vesicle membrane and, in armored species, thecal plates, becomes immotile, and then amphiesma regeneration occurs. Here we report the results of our study of cell covering rearrangement during ecdysis in the naked dinoflagellate species Amphidinium carterae Hulburt 1957. Ecdysis was induced by mechanical treatment (centrifugation). The changes in cell organization at the ultrastructural level were examined using transmission electron microscopy methods. Shedding of the plasma membrane and the outer amphiesmal vesicle membranes, fusion of the inner amphiesmal vesicle membranes were observed. The amorphous cytoplasm zone, which underlies inner amphiesmal vesicle membranes in motile cells, retains under the new plasma membrane in ecdysed cells. We showed accumulation of small vesicles and flattened tubules that apparently begin fusion to form juvenile amphiesmal vesicles in this zone. The absence of pellicle in Amphidinium dinoflagellates was suggested.
Key words: dinoflagellate, ecdysis, Amphidinium carterae, amphiesma, electron microscopy
Introduction
The presence of a complex cell covering (amphiesma) determines structural organization and physiology of dinoflagellate cells. Flattened alveoli, or amphiesmal vesicles, underlie the plasma membrane and are closely adjacent to it and to each other. Morphologically, two forms of dinoflagellates are distinguished depending on the content of alveoli — naked, or unarmored, and armored (Dodge and Crawford, 1970; Morrill and Loeblich, 1983). In the latter, amphiesmal vesicles contain cellulosic plates
(thecal plates). The naked dinoflagellates possess amphiesmal vesicles that either are empty or contain amorphous material. An additional pellicular layer (pellicle) may be present in the amphiesma in both cases (Morrill and Loeblich, 1983; Pozdnyakov and Skarlato, 2012). At the ultrastructural level, the dinoflagellate cell covering includes three membranes — plasmalemma, outer amphiesmal vesicle membrane (OAVM), and inner amphiesmal vesicle membrane (IAVM).
This type of cell covering organization makes it a labile system that can be re-formed in dinofla-
doi:10.21685/1680-0826-2019-13-2-2 © 2019 The Author(s)
Protistology © 2019 Protozoological Society Affiliated with RAS
gellate life history. The phenomenon of such a rearrangement is called ecdysis. Ecdysis is a process of shedding of the cell covering elements unique to dinoflagellates. An ecdysing cell loses plasmalemma, OAVM and, in armored species, thecal plates (Pozdnyakov and Skarlato, 2012). These radical changes are followed by the formation of the new covering. The former IAVM becomes the new plasma membrane, and the new amphiesmal vesicles are formed beneath it in the cortical cytoplasm zone. During this transformation, a cell remains immotile. In the vast majority of studied species, the pellicle becomes a well-developed layer and covers a cell until completion of the new amphiesma formation (Morrill, 1984; Bricheux et al., 1992; Hohfeld and Melkonian, 1992; Sekida et al., 2001, 2004). In the literature, the usually short-term immotile stage formed as a result of ecdysis is called "temporary", "ecdysal", or "pellicle" cyst (Bravo and Figueroa, 2014).
Ecdysis is employed at the different stages of the dinoflagellate life cycle whenever cell covering rearrangement is necessary. This process occurs during cytokinesis in species reproducing by eleu-theroschisis (Morrill and Loeblich, 1984; Pfiester, 1984). Shedding of the cell covering elements also takes place during the transition to the resting cysts stage/excystment (Kokinos and Anderson, 1995; Pfiester, 1989). Ecdysis is considered as a mechanism of the multimembrane cell covering formation in the symbiotic Symbiodinium genus (Wakefield et al., 2000; Wakefield and Kempf, 2001).
The stressor-induced ecdysis holds a special place in the dinoflagellate biology. Mechanical treatment, osmotic shock, temperature, and nutrient changes can act as such stressors (Morrill, 1984; Bricheux et al., 1992; Hohfeld and Melkonian, 1992; Smayda, 2010; Onda et al., 2014; Chan et al., 2019). As a variant of the stress response, ecdysis appears to provide rearrangements in the cell cover, possibly including alteration in its molecular composition which may be necessary for an adequate response to external conditions. Stressor-induced ecdysis is an appropriate model to study the mechanisms underlying shedding and regeneration of the amphiesmal elements. It should be noted that such studies have hitherto been focused on armored dinoflagellates (Morrill, 1984; Bricheux et al., 1992; Kwok and Wong, 2003; Sekida et al., 2001, 2004). The exception is a work of Hohfeld and Melkonian (1992) that included data on ecdysis in the naked species Amphidinium rhynchocephalum. Besides, dissociation of two outer membranes from
the rest of the cell covering in Noctiluca scintillans (referred as N. miliaris) was mentioned (Melkonian and Hohfeld, 1988). However, those results should be reconsidered in the light of newly accumulated information and revision ofthe model of amphiesma changes after ecdysis.
Here we present the first results of the study of the cell covering rearrangement during ecdysis in the naked dinoflagellate species Amphidinium carterae Hulburt 1957. Ecdysis was induced by mechanical treatment (centrifugation). The changes in cell organization at the ultrastructural level were examined using transmission electron microscopy methods. Our observations represent the basis for subsequent investigation of the membrane transformation and amphiesma regeneration.
Material and methods
Cell culture
The culture of the dinoflagellate isolated from the White Sea and designated as Amphidinium carterae was obtained from the collection at the Department of Hydrobiology, Lomonosov Moscow State University and is currently maintained in the protist collection at the Laboratory of Cytology of Unicellular Organisms (Institute of Cytology RAS). Cells were grown in 17 PSU f/2 medium without silicate prepared in artificial seawater (ASW) (Guillard and Ryther, 1962; Kester et al., 1967) at room temperature, pH 8.2 and 50 ^mol photons m-2 s-1 under a 12 h light : 12 h dark cycle.
DNA extraction, PCR amplification, electrophoresis and sequencing
Cells were lysed by freezing at —80 °C for 10 min and then thawed at room temperature. Total DNA was isolated using a DNA extraction kit (BioSilica Ltd., Russia) in accordance with manufacturer's instructions.
In order to verify the species identity of the dinoflagellates in the culture, several PCR reactions were conducted using 18S rRNA gene-specific primer pairs 18ScomF1/ Dino18SR1 (Zhang et al., 2005), RibA/ S20R, and RibA/ RibB (RibA
- ACCTGGTTGATCCTDCCAGT; RibB -TGATCCATCTGCAGGTTCACCTAC; S20R
- GACGGGCGGTGTGTACAA). Amplification was carried out in 30 ^l mixture containing 15 ^l 2X DreamTaq MasterMix (Thermo Fisher Scientific,
USA), 6 ^l DNase/RNase-free water, 3 ^l of forward and reverse primers and 3 ^l of genomic DNA template. PCR reaction started with pre-denaturation at 95 °C for 5 min followed by 39 cycles comprising denaturation at 94 °C for 30 s, primer annealing for 1 min (temperature differs for each primer pair, Table 1), and elongation at 72 °C for 2 min. The procedure was completed by the elongation step at 72 °C for 7 min.
Separation of PCR products was conducted in 1.5 % agarose gel in 1* TAE buffer. To estimate amplicon sizes, we used GeneRuler 1000 bp DNA ladder (Thermo Fisher Scientific, USA). After the separation, gels were stained with ethidium bromide, and amplified fragments were visualized under UV light. PCR products were then extracted from gels by means of BioSilica gel extraction kit (BioSilica Ltd., Russia) according to the manufacturer's instructions. DNA sequencing ofthe obtained amplicons was performed by Beagle Co. Ltd. with the use of the abovementioned primers, as well as the internal sequencing primer sAF (CTGGTTGATYCTGCCAG). NCBI nucleotide BLAST search was used to confirm the species identity.
Induction of ecdysis, light and transmission
electron microscopy
To induce ecdysis, cells were harvested by centrifugation (2000 g for 3 min), carefully re-suspended and incubated at room temperature for 6 h.
For intravital observations of A. carterae cells, a Leica DM2500 (Leica-Microsystems, Germany) light microscope equipped with phase and differential interference contrast optics was used.
For the electron microscopy studies, cells were fixed immediately after centrifugation, 30 min, 3 h and 6 h after the treatment. Non-centrifuged cells were fixed for the control. No additional harvesting was carried out prior to fixation. The cell suspensions were incubated with the fixative mixture added directly to the culture medium. The mixture contained the following components with final concentrations after combining with a sample: 3% (v/v) glutaraldehyde (Sigma-Aldrich, USA), 1% (w/v) osmium tetraoxide (TAAB Laboratories Equipment, UK), 1 mM CaCl2, 17 PSU ASW (pH 8.0-8.5). After 30 min fixation, 17 PSU ASW was added and samples were gently harvested by centrifugation (600 g for 3 min). Then cells were rinsed in 17 PSU ASW thrice, embedded in 2% agar,
Table 1. Annealing temperature for the used primer pairs.
Primer pair Annealing temperature (°C)
18ScomF1/ Dino18SR1 58
RibA/ S20R 52
RibA/ RibB 52
dehydrated and embedded in Epon 812 — Araldite M (Fluka, Switzerland) resin mixture. Ultrathin sections were cut using an Ultracut E (Reichert Jung, Austria) ultramicrotome, contrasted with uranyl acetate and lead citrate and examined in a Libra 120 (Carl Zeiss, Germany) microscope.
Results
Species identification
The studied dinoflagellate monoculture was tentatively identified as Amphidinium carterae according to the morphological similarity with the original description (Fig. 1, A) (Hulburt, 1957). To confirm the species identity, we conducted pCR amplification with total DNA extracted from the culture and 18S rRNA gene-specific primers. Five products were obtained and sequenced. The best BLAST hits showed that all of them were fragments of18S RNA gene of A. carterae.
Cell morphology and ultrastructure after mechanical treatment
Some A. carterae cells lost motility almost immediately after centrifugation (Fig. 1, B). At the light microscope level, we did not observe any distinguishable morphological changes in immotile cells as compared to the untreated motile cells (Fig. 1, A, B). They retained elongate-elliptical cell shape with a small conical epicone.
We examined changes in the organization of A. carterae amphiesma and cortical cytoplasm zone at the ultrastructural level using transmission electron microscopy. The intact amphiesma of amphidinoid vegetative cells consisted of flattened empty (filled with electron-transparent content) amphiesmal vesicles beneath the plasma membrane (Fig. 2, A). The OAVMs adjoined to the plasma membrane, and lateral membranes of adjacent vesicles were tightly appressed in sutures. The cortical cytoplasm zone under IAVMs was organized as an amorphous layer (20-30 nm in width) underlain by a discontinuous
Fig. 1. Morphology of Amphidinium carterae cells (light microscopy): A — motile vegetative cells in culture; B — immotile cells after centrifugation. Scale bars: 10 ^m.
row of microtubules (Fig. 2, A, B). The membrane compartment consisting of large vacuoles with electron-transparent content (cytoplasmic vacuoles) that contained smaller vesicles and tubules was localized beneath microtubules.
After centrifugation the integrity of the plasma membrane and OAVMs was disrupted (Fig. 2, C). These membranes separated from the sutures, vesiculated and then were shedded by the cell (Fig. 2, C). Some cells underwent ecdysis immediately after treatment. The sutures were disrupted, IAVMs fused covering the entire cell, and it was the only membrane surrounding the cell. The amorphous zone of cytoplasm and microtubules were retained in immotile cells (Fig. 2, D). Such a pattern of cell covering organization was maintained at that stage. Finally, the new amphiesma began to form in the dinoflagellate cells. The small vesicles and flattened tubules were observed in the amorphous cytoplasm zone between the former IAVM and large cytoplasmic vacuoles that also contained small vesicles and membrane structures (Fig. 2, E, F). The vesicles in the amorphous cytoplasm zone apparently began fusion to formjuvenile amphiesmal vesicles that would be structural elements ofthe new amphiesma.
Discussion
The ability for ecdysis is one of a number of specific features inherent to this amazing group — the dinoflagellates. This process and the concurrent amphiesma rearrangements ensure protection ofthe dinoflagellate cells during unfavorable conditions or the reproduction period. However, a number of issues concerning different aspects of ecdysis,
including changes that occur at the molecular level, still need to be resolved.
The studies of membrane rearrangement at the molecular level should be linked to the morphological and ultrastructural data. For this purpose, we started a revision of the ecdysis process in a representative of the naked dinoflagellates Amphidinium carterae using light and transmission electron microscopy methods. Fixation was performed based on the modified protocols offered by Shigenaka et al. (1973) for ciliates and by McFadden and Melkonian (1986) for microalgae. The outer membranes in the Amphidinium amphiesma are very vulnerable to the conventional double fixation (Klut et al., 1985). Simultaneous treatment with a mixture of glutaraldehyde and osmium tetraoxide allowed to preserve their intact structure. Divalent cations, e.g. Ca2+, were included in fixatives to improve the preservation of microtubules and other cytoskeletal elements (Shigenaka et al., 1973).
The initial stages of ecdysis were previously described in the other Amphidinium species, A. rhynchocephalum (Höhfeld and Melkonian, 1992). Based on the results of observations of changes in amphiesma ultrastructure in A. rhynchocephalum and armored species Heterocapsa niei, Höhfeld and Melkonian (1992) proposed their ecdysis model. They described the membrane that covered a cell after shedding of two outer membranes as a pellicle membrane. It is formed by IAVMs fusion. The amorphous layer beneath this membrane was therefore referred to as pellicular. In A. rhyn-chocephalum this layer exists in the vegetative cells and does not undergo structural changes during amphiesma rearrangement. In H. niei it forms during ecdysis and has an additional honeycomb-patterned layer. However, the model of Höhfeld and Melkonian (1992) did not take into consideration the issue of the origin of the new plasmalemma. Pozdnyakov and Skarlato (2012) pointed this out in the paper revising and generalizing data concerning the amphiesma rearrangement stages.
Here it is necessary to mention the modern concept of a pellicle. Currently, the consensus appears to have been reached that a pellicle is an external layer covering a cell over the former IAVM after ecdysis (Bricheux et al., 1992; Kwok and Wong, 2003; Sekida et al., 2001, 2004; Chan et al., 2019). It is a temporary covering that bounds an immotile cell in the period of vulnerability and is to be removed after amphiesma renovation/completion of division. Consequently, the membrane that
Fig. 2. Ultrastructure of cell covering in Amphidinium carterae cells (transmission electron microscopy). A, B — Motile vegetative cells; C — ecdysing cell; integrity of the plasma membrane and OAVMs is disrupted; they separate from the sutures (arrowheads); D — ecdysed cell; after sutures disruption IAVMs fuse and form continuous membrane that covers a cell (new plasma membrane); amorphous cytoplasm zone and microtubules (inset, arrowheads) are retained; E, F — ecdysed cells, 6 h after centrifugation; small vesicles and flattened tubules (arrows) accumulate in the amorphous cytoplasm zone. Abbreviations: az — amorphous cytoplasm zone, av — amphiesmal vesicles, iavm — inner amphiesmal vesicle membrane, lv — large cytoplasmic vesicles, mt — microtubules, npm — new plasmalemma, oavm — outer amphiesmal vesicle membrane, pm — plasmalemma, s - sutures. Scale bars: A, B, E, F — 500 nm, C-D — 1 ^m.
retains after pellicle shedding should be a new plasma membrane. Sekida with co-authors (Sekida et al., 2004) investigated the changes in the membrane polysaccharide staining pattern and thus demonstrated that transformation of IAVM into the new plasmalemma occurred after the development of a pellicle layer in the armored dinoflagellate Scrippsiella hexapraecingula.
For the Amphidinium dinoflagellates, we suppose that Höhfeld and Melkonian's (1992) pellicle
membrane should be considered as the new plasmalemma derived from fused IAVMs. The amorphous cytoplasm zone is an internal layer and therefore it cannot be a pellicle in the sense that is described above. Otherwise, another continuous membrane would be required under this layer in the ecdysed cell. The outer membrane of the cytoplasmic vacuoles is localized in this zone but this compartment is permanently present in cells ofmany dinoflagellates and is likely to remain stable during the membrane
rearrangements (e.g. Wetherbee, 1975; Morrill, 1984; Klut et al., 1985; Lucas and Vesk, 1990; Bricheux et al., 1992). Moreover, we consider the presence of small vesicles and flattened tubules in the amorphous zone as evidence in favor of our hypothesis. They accumulate freely in this area and appear to begin fusion to form juvenile amphiesmal vesicles. Besides, A. carterae, A. corpulentum, and al. operculatum were among 45 dinoflagellate species tested for the presence of a pellicle that is acetolysis-resistant and positively reacts with carbohydrate-specific stains (Morrill and Loeblich, 1981), and the results ofthis analysis were negative for Amphidinium species.
Klut with co-authors (1985) conducted an examination of A. carterae cell surface using cytoche-mical methods. Notably, they mentioned a so-called amorphous layer, but they did not hypothesize about its nature. The authors showed that along with the acid mucopolysaccharide coat overlying the plasma membrane, there was a "fuzz" material on the inner surface of amphiesmal vesicles (Klut et al., 1985). It can be assumed that this material is a precursor of glycocalyx, and therefore IAVMs can be ready to form the new plasmalemma in the case of ecdysis. As we mentioned above, Sekida with co-authors (Sekida et al., 2004) showed the transformation of IAVM into the new plasmalemma after the development of a pellicle layer. Such a transformation can probably occur faster in non-pelliculate species like Amphidinium.
Our hypothesis about nature of the amorphous cytoplasm zone does not exclude specificity of its structural organization and, apparently, the peculiar molecular composition. Such a layer (interpreted as a pellicle) was also shown in Noctiluca cells (Melkonian and Höhfeld, 1988). Interestingly, the authors noted the dependence of amphiesmal vesicles shape on the amorphous layer thickness. It can be assumed that this zone (layer) is responsible for cell rigidity in the absence of other strengthening amphiesmal elements. The special study of the zone structure and its molecular composition is required to elucidate this issue.
The genus Amphidinium is considered as the basal group of core dinoflagellates according to the eight-gene phylogeny (Orr et al., 2012). The abovementioned Noctiluca, which possesses a similar layer, was also often positioned among basal core dinoflagellates (Ki, 2010; Hoppenrath et al., 2017). This similarity allows assuming that pellicle can be absent in the basal groups. Given that the origin of theca was in the focus of the phylogenetic
study (Orr et al., 2012), the origin of pellicle can also be a promising albeit complicated issue for such analysis. Further investigations of these aspects in amphidinioid dinoflagellates and involvement of more species in the analysis can shed new light on this basic problem of dinoflagellate biology.
Acknowledgments
The research was funded by the Russian Science Foundation, project No 18-74-10093.
References
Bravo I. and Figueroa R.I. 2014. Towards an ecological understanding of dinoflagellate cyst functions. Microorganisms. 2, 11—32.
Bricheux G., Mahoney D.G. and Gibbs S.P. 1992. Development of the pellicle and thecal plates following ecdysis in the dinoflagellate Glenodinium foliaceum. Protoplasma. 168, 159—71.
Chan W.S., Kwok A.C.M. and Wong J.T.Y. 2019. Knockdown of dinoflagellate cellulose synthase CesA1 resulted in malformed intracellular cellulosic thecal plates and severely impeded cyst-to-swarmer transition. Front. Microbiol. 10, 546.
Dodge J.D. and Crawford R.M. 1970. A survey of thecal fine structure in the Dinophyceae. Bot. J. Linn. Soc. 63, 53—67.
Guillard R.R.L. and Ryther J.H. 1962. Studies of marine diatoms I. Cyclistella nana Hustedt and Detonula confervacea (Cleve) Gran. Can. J. Microbiol. 8, 229-39.
Höhfeld I. and Melkonian M. 1992. Amphiesmal ultrastructure of dinoflagellates: a reevaluation of pellicle formation. J. Phycol. 28, 82-89.
Hoppenrath M. 2017. Dinoflagellate taxonomy—a review and proposal ofa revised classification. Mar. Biodivers. 47, 381-403.
Hulburt E.M. 1957. The taxonomy ofunarmored Dinophyceae of shallow embayments on Cape Cod, Massachusetts. Biol. Bull. 112, 196-219.
Kester D.R., Duedall I.W., Connors D.N. and Pytkowicz R.M. 1967. Preparation of artificial seawater. Limnol. Oceanogr. 12, 176-179.
Ki J.S. 2010. Nuclear 28S rDNA phylogeny supports the basal placement of Noctiluca scintillans (Dinophyceae; Noctilucales) in dinoflagellates. Eur. J. Protistol. 46, 111-120.
Klut M.E., Bisalputra T. and Antia N.J. 1985. Some cytochemical studies on the cell surface of
Amphidinium carterae (Dinophyceae). Protoplasma. 129, 93-99.
Kokinos J.P. and Anderson D.M. 1995. Morphological development ofresting cysts in cultures of the marine dinoflagellate Lingulodinium polyedrum (= L. machaerophorum). Palynology. 19, 143-166.
Kwok A.C.M. and Wong J.T.Y. 2003. Cellulose synthesis is coupled to cell cycle progression at G1 in the dinoflagellate Crypthecodinium cohnii. Plant Physiol. 131, 1681-1691.
Lucas I.A. and Vesk M. 1990. The fine structure of two photosynthetic species of Dinophysis (Dino-physiales, Dinophyceae). J. Phycol. 26, 345-357.
McFadden G.I. and Melkonian M. 1986. Use of Hepes buffer for microalgal culture media and fixation for electron microscopy. Phycologia. 25. 551-557.
Melkonian M. and Höhfeld I. 1988. Amphiesmal ultrastructure in Noctiluca miliaris Suriray (Dinophyceae). Helgol. Meeresunters. 42, 601-612.
Morrill L.C. 1984. Ecdysis and the location of the plasma membrane in the dinoflagellate Hetero-capsa niei. Protoplasma. 119, 8-20.
Morrill L.C. and Loeblich A.R., Ill. 1981. The dinoflagellate pellicular wall layer and its occurrence in the division Pyrrhophyta. J. Phycol. 17, 315-323.
Morrill L.C. and Loeblich A.R. 1983. Ultrastructure of the dinoflagellate amphiesma. Int. Rev. Cytol. 82, 151-80.
Morrill L. and Loeblich A.R., III. 1984. Cell division and reformation of the amphiesma in the pelliculate dinoflagellate, Heterocapsa niei. J. Mar. Biol. Assoc. U. K. 64, 939-953.
Onda D.F.L., Lluisma A.O. and Azanza R.V. 2014. Development, morphological characteristics and viability of temporary cysts of Pyrodinium bahamense var. compressum (Dinophyceae) in vitro. Eur. J. Phycol. 49, 265-275.
Orr R.J., Murray S.A., Stbken A., Rhodes L. and Jakobsen K.S. 2012. When naked became armored: an eight-gene phylogeny reveals monophyletic origin of theca in dinoflagellates. PloS one. 7, e50004.
Pfiester L.A. 1984. Sexual reproduction. In: Dinoflagellates (Ed: Spector D.L). Academic Press, pp. 181-199.
Pfiester L.A. 1989. Dinoflagellate sexuality. Int. Rev. Cytol. 114, 249-272.
Pozdnyakov I. and Skarlato S. 2012. Dinoflagellate amphiesma at different stages ofthe life cycle. Protistology. 7, 108-115.
Sekida S., Horiguchi T. and Okuda K. 2001. Development of the cell covering in the dinoflagellate Scrippsiella hexapraecingula (Peridinales, Dinophyceae). Phycol. Res. 49, 163-176.
Sekida S., Horiguchi T. and Okuda K. 2004. Development of thecal plates and pellicle in the dinoflagellate Scrippsiella hexapraecingula (Peri-diniales, Dinophyceae) elucidated by changes in stainability of the associated membranes. Eur. J. Phycol. 39, 105-114.
Shigenaka Y., Watanabe K. and Kaneda M. 1973. Effects of glutaraldehyde and osmium tetroxide on hypotrichous ciliates, and determination of the most satisfactory fixation methods for electron microscopy. J. Protozool. 20, 414-420.
Smayda T.J. 2010. Adaptations and selection of harmful and other dinoflagellate species in upwelling systems 1. Morphology and adaptive polymorphism. Prog. Oceanogr. 85, 53-70.
Wakefield T.S., Farmer M.A. and Kempf S.C. 2000. Revised description of the fine structure of in situ "Zooxanthellae" genus Symbiodinium. Biol. Bull. 199, 76-84.
Wakefield T.S. and Kempf S.C. 2001. Development of host-and symbiont-specific monoclonal antibodies and confirmation of the origin of the symbiosome membrane in a cnidarian-dinoflagel-late symbiosis. Biol. Bull. 200, 127-143.
Wetherbee R. 1975. The fine structure of Cera-tium tripos, a marine armored dinoflagellate: III. Thecal plate formation. J. Ultrastruct. Res. 50, 77-88.
Zhang H., Bhattacharya D. and Lin S. 2005. Phylogeny of dinoflagellates based on mitochondrial cytochrome b and nulear small subunit rDNA sequence comparisons. J. Phycol. 41, 411—420.
Address for correspondence: Mariia Berdieva. Institute of Cytology of the Russian Academy of Sciences, Laboratory of Unicellular Organisms, Tikhoretsky Ave. 4, 194064 St. Petersburg, Russia; e-mail: maria.berd4@yandex.ru.
