1231
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
1234 Y. Bhaud and others
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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