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Journal of Cell Science 113, 1231-1239 (2000) Printed in Great Britain © The Company of Biologists Limited 2000 JCS0842
Morphology and behaviour of dinoflagellate chromosomes during the cell cycle and mitosis Yvonne Bhaud1, Delphine Guillebault1, Jean-François Lennon2, Hélène Defacque1, Marie-Odile Soyer-Gobillard1 and Hervé Moreau1,* 1Observatoire 2Observatoire
Océanologique de Banyuls, Laboratoire Arago, UMR CNRS 7628, BP44, F-66651 Banyuls-sur-Mer, France cedex Océanologique de Roscoff, UPR CNRS 9042, BP 74, F-29682 Roscoff, France cedex
*Author for correspondence (e-mail:
[email protected])
Accepted 30 January; published on WWW 7 March 2000
SUMMARY The morphology and behaviour of the chromosomes of dinoflagellates during the cell cycle appear to be unique among eukaryotes. We used synchronized and aphidicolinblocked cultures of the dinoflagellate Crypthecodinium cohnii to describe the successive morphological changes that chromosomes undergo during the cell cycle. The chromosomes in early G1 phase appeared to be loosely condensed with numerous structures protruding toward the nucleoplasm. They condensed in late G1, before unwinding in S phase. The chromosomes in cells in G2 phase were tightly condensed and had a double number of arches, as visualised by electron microscopy. During
prophase, chromosomes elongated and split longitudinally, into characteristic V or Y shapes. We also used confocal microscopy to show a metaphaselike alignment of the chromosomes, which has never been described in dinoflagellates. The metaphase-like nucleus appeared flattened and enlarged, and continued to do so into anaphase. Chromosome segregation occurred via binding to the nuclear envelope surrounding the cytoplasmic channels and microtubule bundles. Our findings are summarized in a model of chromosome behaviour during the cell cycle.
INTRODUCTION
proposed, but more data are needed before a definitive view can emerge (Spector, 1984). The permanently high degree of DNA compaction in the dinoflagellate genome, and the absence of nucleosomes, raise the problem of genome replication and/or transcription. In higher eukaryotes these processes depend largely on the dynamic structure of chromatin and nucleosomes (Wade et al., 1997). There is a distinct S phase in dinoflagellates and transient unwinding of chromosomes has been correlated with this phase (Spector et al., 1981). Experimental evidences strongly supports the view that transcription occurs on extrachromosomal DNA filaments protruding from the chromosome core in several species, although the molecular mechanisms underlying this phenomenon are unknown (Sigee, 1984, 1986; Soyer-Gobillard et al., 1990; Rizzo, 1991). Mitosis has several original characteristics in dinoflagellates, such as a persistent nuclear envelope and cytoplasmic invaginations that form channels crossing the nucleus and containing microtubular bundles (Loeblich and Hedberg, 1976; Fritz and Triemer, 1983; Perret et al., 1993). A major consequence of this structural arrangement is that the microtubular spindle is not in direct contact with chromosomes as it is in other eukaryotes, but is separated by the nuclear envelope. Condensed chromosomes appear bound to the nuclear envelope and electron-dense material has been observed at the contact point in several species (Oakley and
Dinoflagellates are unicellular micro-organisms that are widely distributed in marine and fresh waters. They are true eukaryotes with a G1-S-G2-M cell cycle, but have some unusual nuclear characteristics, such as a high genomic DNA content, a permanent nuclear envelope, an extranuclear mitotic spindle and chromosomes bound to the nuclear membrane for segregation (for reviews, see Raikov, 1995; Soyer-Gobillard, 1996). These oddities, as in many other protozoans, can be fruitful models for studying general cellular processes, or can supply the exception that proves the rule (Vickerman and Coombs, 1999). The most prominent feature of dinoflagellates is probably the morphology and structure of their chromosomes. DNA filaments are packaged in chromosomes varying in number from 4 to 200, depending on the species. The chromosomes of most dinoflagellates are described as permanently condensed throughout the cell cycle, and the nuclear filaments appear as a series of arches in ultrathin sections (Spector, 1984). Nuclear filaments lack histones and nucleosomes: no repeating subunit structures have been detected in dispersed genomic DNA, and the protein:DNA mass ratio is 1:10, while it is one in other eukaryotes. This absence of nucleosomes and histones from the chromatin is unique among eukaryotes (Rizzo, 1987, 1991). Several models of the chromosome structure have been
Key words: Chromosome, Dinoflagellate, Mitosis
1232 Y. Bhaud and others Dodge, 1974; Ris and Kubai, 1974; Fritz and Triemer, 1983). Although prophase and anaphase have been described, no metaphase alignment of chromosomes has been reported. The generally accepted view of dinomitosis is that segregation of chromosomes is mediated by their attachment to microtubules accross the nuclear envelope. This attachment to the membrane is reminiscent of the segregation of nucleoids in prokaryotes. In contrast to prokaryotes, however, the presence of a microtubular spindle in dinoflagellates suggests a different eukaryote-like segregation mechanism, driven by microtubule bundles. In spite of numerous published descriptions of the dinoflagellate cell cycle, the precise succession of changes in chromosome morphology during the cell cycle, and the alignment of chromosomes (metaphase) before their segregation, remain unclear. We therefore used synchronized cultures of Crypthecodinium cohnii to accurately describe the morphology of the chromosomes and their alignment before segregation during mitosis. These findings have been used to develop a model of chromosome behaviour during the cell cycle.
MATERIALS AND METHODS Cell cultures and synchronization Crypthecodinium cohnii (ATCC strain 50050), a heterotrophic dinoflagellate, was grown in MLH medium (Tuttle and Loeblich, 1975) in the dark at 27°C. The cell cycle lasts for 8 hours under these conditions, and cells were synchronized as described in Bhaud et al. (1991, 1994). Briefly, dividing cells lack flagella and are encysted, while interphase cells are biflagellated and swimming; these morphological differences were used to separate the phases of the cell cycle (Bhaud et al., 1994). Synchronized cultures of C. cohnii cells at S phase are difficult to obtain uncontaminated with late G1 and/or G2 and early M cells, because the S phase is short. It probably begins when the cells are still swimming and ends in young encysted cells. We overcame this problem by adding 30 µM aphidicolin (10 µg/ml) (Sigma, Saint Quentin Fallavier, France) to the culture medium. Aphidicolin blocks many eukaryotic cells in early S phase by inhibiting DNA polymerase activity (Decker et al., 1986). Flow cytometry analysis The cell cycle was analysed by flow cytometry essentially as described by Taroncher-Oldenburg et al. (1997) for Alexandrium fundyense and by Wong and Whiteley (1996) for C. cohnii. At least 1×106 cells were collected by centrifugation (5 minutes at 3000 g, 4°C) and incubated in 4% paraformaldehyde for at least 1 hour at 25°C. The fixed cells were washed twice in phosphate-buffered saline (PBS) and placed in methanol at 4°C for at least 1 hour. DNA was stained by washing fixed cells three times in PBS, resuspending them in PBS containing propidium iodide (50 µg/ml) (Sigma, Saint Quentin Fallavier, France) and RNAse I (100 µg/ml) (Promega, Madison, USA) in the dark for at least 2 hours. 10,000 events were measured using a FACScan (Becton Dickinson, San Jose, CA, USA) with an argon ion laser at 488 nm for excitation. The emitted fluorescence was collected in the FL2 channel with a 586 nm bandpass filter. Acquisition and analysis of the results were performed with the CELLQuest software (Beckton Dickinson). Debris were excluded from the analysis by gating on the Forward Scatter versus Side Scatter parameters. Nuclear fluorescence intensity was recorded on a linear scale of the FL2-A and FL2-H signals, which distinguished dividing cells from cell doublets that had the same fluorescence intensity. Unless specified, deconvolution of FL2-H intensity histograms to yield the proportion of cells in G1, S
and G2+M phases was performed using the Modfit software (Beckton Dickinson), in which G1 and G2+M populations were modelled as Gaussian distributions, whereas cells in S phase were analysed as a sum of broadened rectangles. Cysts that produced four daughter cells (defined as those populations with a DNA content above that of the G2+M peak) were excluded for this analysis. Fluorescence microscopy Crypthecodinium cohnii cultures were centrifuged at 800 g and pellets were treated for cryofixation and cryosectioning (Perret et al., 1993). Cryosections (1.5 µm thick) were post-fixed for 10 minutes in 3% paraformaldehyde and rinsed three times in PBS-0.1% Tween 20. Sections were stained with DAPI (0.1 µg/ml in water) for 10 minutes, rinsed with water and mounted in Mowiol containing 5% N-propyl gallate as anti-fading agent. Confocal microscopy Intact cells fixed in 3% paraformaldehyde were stained in DAPI as described above. Cells labelled for chromosome fluorescence were imaged on an Olympus Fluoview confocal microscope (Oceanological Center of Roscoff, France), modified for use in twophoton mode (Denk et al., 1990). This involved replacing the Argon/Krypton CW laser by a pulsed infra-red laser (Mira 900 pumped by a 5 watts Verdi, both from Coherent) and adapting a dichroic filter for the new excitation wavelength for the DAPI (750 nm in the experiments described here). The objective was an Olympus 60× UplanFI (NA=1.25, oil immersion). The 3-D structures of the nuclei were first recorded as stacks of planes at 0.5 µm intervals and then rotated over 180°, by 20° steps, around a horizontal axis. Electron microscopy Two methods of fixation were used to obtain well preserved nucleic acids (chemical fixation) or proteins (fast freeze fixation). Crypthecodinium cohnii cells were collected by centrifugation and fixed for DNA preservation in paraformaldehyde/glutaraldehyde/ Pipes buffer, then post-fixed in osmium tetroxide in Pipes buffer, dehydrated and embedded in Epon (Soyer, 1977). For protein preservation a drop of concentrated cell suspension was placed on filter paper (10 mm2) and mounted on a specimen holder. The sample was slammed onto a metal-mirror block of pure copper cooled with liquid helium to −269°C on a cryovacublock (Escaig, 1982), and stored in liquid nitrogen. Freeze-substitution in acetone and 2% OsO4 was carried out in a cryocool apparatus for 3 days at −80°C. The temperature was gradually raised to −30°C and kept there for 2 hours. Finally, the samples were thawed at room temperature for 1 hour, washed in pure acetone followed by absolute ethanol and propylene oxide, and embedded in Epon. Sections were stained with uranyl acetate and lead citrate and examined in a Hitachi H-600 transmission electron microscope.
RESULTS Characterization of Crypthecodinium cohnii cultures by flow cytometry analysis We first analysed the DNA content of an unsynchronized culture of C. cohnii by propidium iodide staining and flow cytometry (Fig. 1A,C). The first well-defined DNA peak was due to cells in G1 phase (61% of the population); the second peak (having twice the DNA fluorescence as cells in G1) was formed by G2+M phase cells (27% of the population). Between these two peaks, 12% of cells were in S phase. G1 cells are biflagellated and swimming in C. cohnii, whereas dividing cells (G2+M) lack flagella and are encysted (Bhaud et al., 1991). Since swimming-G1 cells and encysted-
Dinoflagellate chromosomes 1233
Fig. 1. Cell cycle arrest induced by aphidicolin (A) and cell cycle synchronisation (B), as detected by flow cytometry analysis. Percentages of individual cells in G1, S and G2+M (treated as in A and B) and measured by Modfit software are shown in (C and D). In A, cells were treated for 15 hours with 30 µM aphidicolin (solubilised in DMSO) or DMSO alone (control). In B, cells were synchronized as described previously (Bhaud et al., 1994): top panel, interphasic swimming cells released from plates 5 hours after synchronisation; bottom panel, encysted cells recovered from plates. (1) in these samples, the percentage of cells in G1, S or G2+M phases was visibly determined using the CELLQuest software, by gating individual cells in G1, S or G2+M phases (as determined from control cultures).
dividing cells differed in their morphology and behaviour, we used these properties to synchronize and separate them. Flow cytometry analysis of swimming cells released after synchronisation revealed that 72% of the cells were in G1 phase, and only 8% could be considered to be in G2+M phases (Fig. 1B,D). In contrast, 60% of encysted cells were in G2+M phases; this sample also had few cells in G1 phase (9%). The results confirm previous observations made by us (Bhaud et al., 1991, 1994) and others (Wong and Whiteley, 1996) on this dinoflagellate species after synchronisation. We have previously shown (Bhaud et al., 1991) that the S phase is short (1 hour) in C. cohnii, which makes it difficult to efficiently isolate cells at this stage. We added aphidicolin to the culture medium to increase the proportion of cells in S phase. This drug inhibits DNA polymerase, and consequently
inhibits DNA synthesis in many systems (Decker et al., 1986). Crypthecodinium cohnii cells treated with aphidicolin stopped growing to give swimming and encysted cells in roughly equal proportions. Flow cytometry indicated that a synchronized aphidicolin-treated culture contained many more cells in S phase (80%) than a control culture (12%) (Fig. 1A,C). Only a few cells (18%) were in G1 phase. Ultrastructural morphology of chromosomes during the cell cycle Early and late swimming (G1) cells (released after 1 or 4 hours), aphidicolin-blocked (S) cells and encysted (S+G2+M) cells of C. cohnii were fixed, embedded in Epon and examined at the EM level (Figs 2, 3). The vast majority of the early swimming cells (at least 80%) had chromosomes that appeared
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Dinoflagellate chromosomes 1235 Fig. 2. Crypthecodinium. cohnii: ultrastructural study of chromosome organization during the cell cycle. (A,B) Longitudinal (A) and tranverse (B) sections of a G1 phase chromosome, 2 hours after release from the cyst. Arrows indicate outwardly protruding structures; see text. (C,D) Longitudinal (C) and transverse (D) sections of a late G1 phase in aphidicolin-blocked cells. (E-G) Different times of the S phase from the beginning to almost complete chromosomal deconsensation. Early S phase (E) was observed in aphidicolin-blocked cells, whereas later S phases (F and G) came from classically synchronized cultures. (H) Longitudinal section of G2 chromosome showing abundant, compact genomic DNA. There are twice as many arches as in the G1 phase (A-D). All sections are at the same magnification. Cells were fixed in formaldehyde/glutaradehyde. Bar, 0.2 µm.
to be well-defined entities (Fig. 2A,B), with the typical archshape organization generally described for dinoflagellate chromosomes (Soyer-Gobillard, 1996). These were the cells
with a minimum DNA content, the G1 cells. The boundaries between chromosomes and the nucleoplasm were ruffled, and there were numerous structures protruding from the chromosomes toward the nucleoplasm (Fig. 2A,B, arrows). There was dense material inside the chromosome cores, usually in the domains defined by arches. The nucleoplasm was dense and granular. Swimming cells sampled 4 hours later (the G1 phase lasts 6 hours) had the same flow cytometry fluorescence pattern (not shown). The chromosomes in these late G1 cells were similar to those of early G1 cells, but the material inside the chromosomes appeared more densely packed into arch domains (Fig. 2C,D). The ruffled aspect of the chromosome/ nucleoplasm boundaries was often absent and there were very few protruding structures. Two cell populations were observed in aphidicolin-blocked cells, having two chromosome morphologies. The arch
Fig. 3. Crypthecodinium cohnii: chromosome organization during mitosis. Progressive splitting of chromosomes during prophase from beginning (A, arrow) to typical Y and V shapes (B, arrow and C). Formaldehyde/glutaraldehyde fixation. (D,E) Relationship between chromosomes and cytoplasmic channels. No distinct structure was found at the point of contact between chromatid and channel membrane, but a discrete thickening of the membrane is visible (D and E, arrows), and there is thin fiber between the channel membrane and the chromatid (D). Bars, 0.2 µm (A), 0.25 µm (B), 0.3 µm (C,E), 0.8 µm (D).
1236 Y. Bhaud and others
Fig. 4. Progression of mitosis monitored by epifluorescence of DAPI-stained cryosections. G1(A) and mitotic (B-H) cells. Prophase (B), transverse (C-D) and longitudinal (E) sections of a metaphase-like plate with thin chromatids arranged around the cytoplasmic channels (arrow in F). Early (F) and late (G) anaphase with the appearance of two discs migrating toward opposite poles of the cell, and the end of chromosome migration (telophase) (H). Bar, 10 µm.
structure of the chromosomes was altered in at least 80% of the cells and the chromosome cores were unwound. Fiber bundles were still visible in most nuclei and chromosomes were only partially decondensed (Fig. 2E,F). The second cell population (around 20% of the cells) had the G1 chromosome morphology described above. In agreement with the flow cytometry histogram obtained from aphidicolin-treated cells, we interpreted cells with decondensed chromosomes as being in S phase. Encysted cells had three populations. One had unwound chromosome S phase cells (25% of the cells), with some almost totally decondensed (Fig. 2G). The second population represented mitotic cells (50% of the cells), identified either by the splitting of their chromosomes or by the presence of cytoplasmic channels crossing the nucleus. The third population (20% of the cells) had very condensed chromosomes with almost twice as many arches as in G1 (Fig. 2H). The overall chromosome size, particularly their thickness, was much greater than in the G1 phase. The nucleoplasmchromosome boundary at this stage was sharp (without fibers protruding toward nucleoplasm), and the nucleoplasm did not appear granular. We interpreted those cells as having twice as many arches as G2 cells. In mitotic cells, split-chromosomes (Fig. 3A,B, arrows) appeared before the membrane invaginations leading to the formation of cytoplasmic channels. The two daughter chromatids began to split at one end to give typical Y or V shapes (Fig. 3C). The chromatids then became elongated and were attached to the membrane of the cytoplasmic channels. We examined the link between chromosomes and the channel membrane using fast frozen sections (Fig. 3D,E). Although nucleic acids (and consequently chromosomes) were not well preserved by this method of fixation, the membrane and protein structures were well preserved. No distinct structure was seen at the contact point between the chromatids and the channel membrane, but there was often a discrete opaque material on the nuclear envelope (Fig. 3D,E, arrows). About 40 microtubules were counted within a transverse section of a channel (Fig. 3E). The chromatids in late mitotic cells were typically elongated and attached to the channel membranes by thin fibers (Fig. 3E). General dynamic of Crypthecodinium cohnii mitosis The synchronized cell samples were used to analyse mitosis
under the light miscroscope. Both cryosections and intact cells were analysed, using DAPI staining conventional fluorescence microscopy (Fig. 4) or a biphoton confocal microscope (Fig. 5). Crypthecodinium cohnii nuclei appeared spherical in 80% of swimming/G1 cells, with a diameter of around 6 µm (Figs 4A, 5B). Encysted cells contained four main mitotic figures in roughly equal proportions. The first group of cells had blurred chromosomes (Fig. 4B), reminiscent of the chromosome splitting seen by electron microscopy. Cytoplasmic invaginations were often visible inside the nuclei, producing the typical cytoplasmic channels crossing the nucleus, and around 10 (8-12) channels were counted. There was no organized distribution of chromosomes, either along the external nuclear envelope, or around the cytoplasmic channels. We interpreted this kind of mitotic figure as the prophase. In the second group of cells, the shape of nuclei had changed and they became flat discs 11-12 µm in diameter and 3 µm thick (Figs 4C-D, 5A), whereas the cells themselves remained spherical (visible in Fig. 5A). The chromosomes were found to be ordered and bound around the channels: 6 to 8 were found around each channel (Fig. 4C,D). Discs observed edge-on appeared to have chromosomes distributed at several levels (Fig. 4E). These superimposed chromosomes could reflect the steric hindrance of chromatids inside the nucleus rather than a real superimposition of the membrane-chromosome contact points. This phase appeared as a metaphase-like alignment, although a true plate in a single plane was never observed. In the third group of cells, the channels lengthened and the chromosomes were visualized as being carried along, giving two separated thick discs (Fig. 4F,G). During chromosome migration (anaphase) the cytoplasmic channels around which the chromosomes were still ordered crossed the two discs (Fig. 4F, arrow). The last group of cells was characterized by formation of two nuclei (telophase) (Fig. 4H). DISCUSSION Dinomitosis was first described by Chatton (1920) and displays several unusual features that were revealed later by electron microscopy. The main features are: (1) the nuclear envelope persists throughout mitosis, (2) the chromosomes, which remain condensed throughout the cell cycle, attach to the inner
Dinoflagellate chromosomes 1237
Fig. 5. Biphoton confocal microscopy observation of mitotic stages of DAPI-stained intact cells of C. cohnii. To show the shape of the nucleus, the basic 3-D information provided by a stack of optical slices (with 1 µm intervals) was processed to rotate the image of the cell by 20° steps. The ray-casting algorithm from the Fluoview software used the ‘brightest’ rendering mode: that means that brightness value retained for each point and each orientation is the brightest met by the projection ray. Associated with the low thickness of the metaphase nucleus (3 µm, i.e. 3 sections), this explains why the images corresponding to 0° and 180° (the first and the last ones) are very similar in (A). These sequences gave a visualization of the 3-D shapes of a metaphase-like (A) and a G1 (B) cell. The G1 spherical nucleus had a diameter of 6 µm while the metaphase nucleus was a disc with a diameter of 12 µm and a thickness of 3 µm. Bars, 6 µm.
face of the nuclear envelope before segregation, and (3) cytoplasmic channels containing the extra-nuclear microtubular spindle traverse the nucleus during mitosis (Kubai and Ris, 1969; Spector, 1984). There have been many descriptions of dinoflagellate chromosomes at both the light and electron microscope levels in different species. Chromosomes are usually described as permanently condensed during the cell cycle, with a constant compact morphology (Raikov, 1978). Several unusual morphologies have also been reported, but these observations were made on unsynchronized or partially synchronized cultures, without firm correlation with a particular stage of the cell cycle (except for mitotic steps, when separation of the chromosomes or cytoplasmic channels were visible (Kubai and Ris, 1969). Spector et al. (1981) reported chromosome unwinding coinciding with a peak in the uptake of 3[H]thymidine by semi-synchronous cultures of Peridinium cinctum. The present description of dinomitosis is based on controlled synchronized cultures, and allows complete description of the successive appearances of chromosomes during the cell cycle (see model in Fig. 6). This confirms previous observations of G1 and S phases, and describes the appearance of chromosomes in the G2 phase for the first time. DNA polymerase activity of a dinoflagellate (unlike RNA polymerase) has never been described, and the inhibition of the C. cohnii DNA polymerase by aphidicolin in the current study indicates (at least for this criterion) that this enzyme has eukaryotic rather than prokaryotic features (Decker et al., 1986). Fig. 6. Diagram of chromosome morphology during the cell cycle of C. cohnii. Each representation is a drawing of the microscopic observations in Figs 1-3 (left), with interpretations on the right. Representations of chromosome morphology during G1 (A), late G1 (B), S (C), G2 (D), prophase/metaphase-like (E) and anaphase (F).
The description of dinomitosis usually begins with the appearance of the channels crossing nuclei. In light of our observations, prophase begins prior to this with partial splitting
1238 Y. Bhaud and others authors to suggest that they may be integrated into the nuclear membrane and perhaps may have evolved from some membrane components (Spector, 1984). The thickening of the cytoplasmic channel membrane at the chromosome attachment point and the end of a microtubule bundle at this level could reflect such a transmembrane structure. Mitosis is usually believed to have evolved from the primitive binding of the nucleoid to the membrane in prokaryotes towards the more complex microtubule- dependent system of higher eukaryotes (Alberts et al., 1994). Dinomitosis, lacking a clear metaphase alignment of chromosomes and direct contact between chromosomes and microtubules, represents one of the intermediate steps. However, several recent data question this particular evolutionary concept of mitosis. For example, the current view that the prokaryote chromosome segregates by attaching to the membrane is no longer considered to be an established fact (Gordon and Wright, 1998). The inferred primitive status of dinoflagellates is also questioned. Our description of a metaphase stage in dinomitosis draws together the different mechanisms of mitosis in eukaryotes, and it is now difficult to maintain this linear evolutionary concept of mitosis from a simple primitive form towards a more complex process. The different (although related) mitotic mechanisms may be viewed as several independent evolutionary attempts in different classes of organisms to produce a (more) efficient segregation of daughter genomes. Fig. 7. Diagram illustrating mitosis in C. cohnii in late prophase (A), metaphase-like (B), anaphase (C) and telophase (D). In metaphaselike (B), an inset shows the detail of chromosome binding to the nuclear membrane channel, in a transverse (upper) and a longitudinal (lower) view.
of the chromosomes. The appearance of flat nuclei before chromosome segregation strongly suggests a metaphase-like alignment of chromosomes in several planes around the cytoplasmic channels. Previous descriptions of dinomitosis were unclear about the existence of a metaphase, and it was generally only said that chromosome segregation occurred by chromosomes becoming attached to the nuclear channel membranes, without direct contact with microtubule bundles. The model shown in Fig. 7, for C. cohnii, illustrates the general scheme of dinomitosis, and shows a metaphase-like alignment of chromosomes, although the chromosomes are not arranged in a single plane as in a classical metaphase plate. Other eukaryotic cells, such as budding yeasts, also lack a true metaphase plate (Straight et al., 1997). Permanent nuclear envelopes during mitosis have also been described in several eukaryotic species including yeasts, diatoms and euglenids (Raikov, 1978), but here the mitotic spindle is intranuclear. In these cases, there is direct contact between chromosomes and microtubules, and the chromosomes are segregated in much the same way as in mammalian cells, but inside the nucleus. The only other known organisms having a permanent nuclear membrane and an extranuclear spindle are hypermastigotes (Raikov, 1978; Dyer, 1989). However, these cells have distinct kinetochore structures linking the chromosomes to the microtubules across the nuclear membrane. The absence of clearly visible kinetochore-like structures in most dinoflagellates led several
The authors thank C. Courtiss for his help with the flow cytometry, L. Besseau and M.L. Géraud for preparing cryofixed samples, M. Albert and D. Saint-Hilaire for technical assistance, M.J. Bodiou for drawing Figs 6 and 7, Pr. T. M. Preston for critically reading the manuscript and INIST for correcting the English. This work was supported by the CNRS (UMR 7628).
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