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Dynamic changes in microtubule organization during division of the primitive dinoflagellate Oxyrrhis marina
Biology of the Cell 92 (2000) 583−594 © 2000 Éditions scientifiques et médicales Elsevier SAS. All rights reserved S0248490000011060/FLA
Original article
Dynamic changes in microtubule organization during division of the primitive dinoflagellate Oxyrrhis marina Koichi H. Katoa*, Akihiko Moriyamaa, Tomohiko J. Itohb, Masayuki Yamamotoc, Tetsuya Horioc, Philippe Huitoreld a
Institute of Natural Sciences, Nagoya City University, Nagoya 467-8501, Japan
b
Division of Biological Sciences, Graduate School of Science, Nagoya University, Nagoya 464-8601, Japan
c
Department of Food Microbiology, School of Medicine, The University of Tokushima, Tokushima 770-0042,
Japan d
Laboratoire de biologie du développement, UMR 7009, CNRS/UPMC, Station zoologique, 06230 Villefranche-
sur-mer, France Received 18 September 2000; accepted 11 December 2000
The marine dinoflagellate Oxyrrhis marina has three major microtubular systems: the flagellar apparatus made of one transverse and one longitudinal flagella and their appendages, cortical microtubules, and intranuclear microtubules. We investigated the dynamic changes of these microtubular systems during cell division by transmission and scanning electron microscopy, and confocal fluorescent laser microscopy. During prophase, basal bodies, both flagella and their appendages were duplicated. In the round nucleus situated in the cell centre, intranuclear microtubules appeared radiating toward the centre of the nucleus from densities located in some nuclear pores. During metaphase, both daughter flagellar apparatus separated and moved apart along the main cell axis. Microtubules of ventral cortex were also duplicated and moved with the flagellar apparatus. The nucleus flattened in the longitudinal direction and became discoid-shaped close to the equatorial plane. Many bundles of microtubules ran parallel to the short axis of the nucleus (cell long axis), between which chromosomes were arranged in the same direction. During ana–telophase, the nucleus elongated along the longitudinal axis and took a dumbbell shape. At this stage a contractile ring containing actin was clearly observed in the equatorial cortex. The cortical microtubule network seemed to be cut into two halves at the position of the actin bundle. Shortly after, the nucleus divided into two nuclei, then the cell body was constricted at its equator and divided into one anterior and one posterior halves which were soon rebuilt to produce two cells with two full sets of cortical microtubules. From our observations, several mechanisms for the duplication of the microtubule networks during mitosis in O. marina are discussed. © 2000 Éditions scientifiques et médicales Elsevier SAS dinoflagellate / Oxyrrhis / mitosis / microtubule / actin
* Correspondence and reprints. E-mail address: [email protected] (K.H. Kato).
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1. INTRODUCTION Oxyrrhis marina is a heterotrophic free-living dinoflagellata and is commonly found in tide pools along the coast. The mode of nuclear division in dinoflagellates has been studied in a few species, including O. marina, among more than 6000 known species, and many differences from that of higher eukaryotic cells have been reported. In these organisms the nuclear envelope does not break down during mitosis, which implies that they have chromosomes in the nucleus throughout the cell cycle and exhibit intranuclear mitosis. However, at least some of them do not show the typical closed mitosis found in yeast for example. Several cytoplasmic channels containing microtubules are formed through the nucleus (Spector and Triemer, 1981). O. marina shows an exceptional type of mitosis with an intranuclear microtubular system which develops at the time of cell division (Triemer, 1982; Gao and Li, 1986; Kato et al., 1997). This characteristic has lead to an argument regarding the position of this organism in the phylogenetic tree. The cortical microtubular systems and the flagellar apparatus have been studied in several dinoflagellates including O. marina (Roberts and Roberts, 1991). In O. marina, the cortical cytoskeleton is known to be composed of bundles of several parallel microtubules (Dodge and Crawford, 1971; Roberts et al., 1993), and is rather flexible (Hohfeld and Melkonian, 1998). The flagellar apparatus of dinoflagellates presents several characteristic features. Two basal bodies give rise to two flagella and several sets of flagellar roots, made of microtubules and fibrous material. Besides the usual axonemal structure, flagella contain highly organized accessory fibrillar appendages, whose structure and function are quite distinct (Cachon et al., 1988a; Cachon et al., 1988b; Cosson et al., 1988; Roberts and Roberts, 1991; Roberts et al., 1993). In the dinoflagellate Crypthecodinium cohnii, both flagella are lost as cells enter cell division, and two daughter cells receive an equal half of the mother cell (Bhaud et al., 1991; Perret et al., 1993). In contrast, O. marina do not loose flagella during cell division and continue swimming throughout the cell cycle. During the early steps of cell division, the flagellar apparatus is duplicated and separated into two pairs. The cell keeps its antero–posterior as well as dorso–ventral polarity during cell division, and consequently each daughter cell receives either an anterior or a posterior half of the mother cell body. The other half has to be built de novo during the late steps of cell division. Therefore, while most higher eukaryotic cells loose their typical interphase cytoskeleton at the entry in mitosis, O. marina duplicates and reorganizes it as in many other protists (see Grell, 1973). In Tetrahymena (Ng, 1979) and Trypanosoma (Sherwin and Gull, 1989) Cell division of O. marina
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for instances, detailed studies were reported concerning the process of duplication of cytoskeletal microtubules, a complex but well regulated process. In the present work, we have studied three microtubular systems, the flagellar apparatus, cortical cytoskeletal microtubules and intranuclear microtubules during cell division in O. marina, using electron microscopy and immunolocalization by confocal laser microscopy. We also examined the participation of actin in cell division in this species. Based on our observations, new mechanisms of duplication of the microtubular cytoskeleton are discussed.
2. MATERIAL AND METHODS 2.1. Cell culture O. marina was cultured in the laboratory essentially by a method described previously, except that quail yolk was used in some experiments as food instead of E.coli. Quail yolk was prepared as follows. One mL of the yolk was suspended in 50 mL of culture medium, boiled for several minutes, homogenized with a Dounce homogenizer (clearance: 10 µm) and centrifuged at 3000 × g for 20 min. The pellet was resuspended in the same volume of culture medium and filtered through nylon mesh (pore size: 10 µm) and used as food. In order to prevent bacterial growth, penicillin G (100 U·mL–1) and streptomycin (100 µg·mL–1) were added to the culture medium. The quail yolk is comparably as effective as E. coli for the food. When the newly prepared yolk was added to the original culture during the stationary phase, most cells started dividing within 12 h.
2.2. Electron microscopy For transmission electron microscopy (TEM), samples were fixed with 4% glutaraldehyde in 0.1 M phosphate buffer (pH 7.0) containing 0.15 M sucrose, 10 mM MgCl2 and 10 mM EGTA, for 45 min at room temperature room temperature. After rinsing in 0.1 M phosphate buffer containing 0.2 M sucrose, they were postfixed with 0.5% OsO4 in the same buffer for 45 min at 4°C. They were then dehydrated through ethanol series and embedded in Spurr’s low viscosity resin on a flat plastic plate. Thin sections were cut using a RMC MT6000 ultramicrotome and examined in a Hitachi H-7100 electron microscope. For scanning electron microscopy (SEM), samples were mounted on 1% polylysine coated cover slips and fixed with the mixture of 3% glutaraldehyde and 0.2% OsO4 in the buffer solution (50 mM borate buffer (pH 8.2) and 250 mM NaCl) for 15 min at room temperature. They were then transferred into the buffer solution containing 3% glutaraldehyde and 1% tannic Kato et al.
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acid and fixed for 1.5 h, followed by post-fixation with 1% OsO4 in the buffer solution for 3 h at 4°C. After rinsing with distilled water, they were dehydrated with ethanol series, infiltrated with iso-amylacetate and critical point dried. A small piece of coverslip was fixed on the specimen stub and spatter coated with gold, and then observed with a JEOL JSM-T100 scanning electron microscope.
2.3. Immunocytochemistry For whole mount preparations, samples were placed on 0.5% polylysine coated cover slips and fixed with 4% formaldehyde in 50% seawater for 4 min. Then, an equal volume of Tris buffer solution (TBS) containing 1% Triton X-100 was added and mixed for 4 min to permeabilize the cells. TBS was composed of 20 mM Tris and 0.3 M NaCl, and adjusted to pH 7.6 with 1 N HCl. After permeabilization, samples were rinsed several times with TBS containing 0.1% Triton X-100 (TBST) followed by rinsing with TBST containing 1% donkey whole serum (TBSTD). Mouse monoclonal anti-β-tubulin antibody D66 (a generous gift of Dr. C. Gagnon, McGill University), mouse monoclonal anti-αtubulin DM1A (Sigma) were used to visualize microtubules. A mouse monoclonal anti-γ-tubulin antibody G19 (Horio et al., 1999) was used to visualize microtubule organizing centres (MTOCs). The G19 antibody has been raised against bacterially expressed Schizosaccharomyces pombe γ-tubulin. Epitope mapping using truncated γ-tubulin peptides revealed that the sequence recognized by G19 is localized within the peptide GGAGNNWANGYSHA (Horio et al., unpublished observation). This region of γ-tubulin is very well conserved among conventional γ-tubulins. Samples were incubated with these antibodies diluted to 1:100–1:1000 with TBSTD for 12 h at 4°C. After rinsing with TBST, they were incubated with FITCconjugated donkey anti-mouse IgG antibody (Amersham) diluted to 1:100–1:1000 with TBSTD for 45 min at room temperature. Samples were then rinsed with TBS and mounted in 50% glycerin containing 10% of DABCO. Preparations were observed on a confocal laser scanning microscope (Yokogawa CSU-10) equipped with an argon/argon–krypton laser at the wave length of 488 nm. Digital images were collected with a cooled CCD camera (Hamaphoto C5985) and saved on a computer. To reconstruct three-dimensional structures, serial images collected at different focal planes were processed on a workstation with imaging software (VoxelView, Vital Images, Inc., USA) by simple stacking of selected images. For semi-thin sectioning, samples were fixed with 3% paraformaldehyde in 0.1 M sodium phosphate buffer (pH 7.0) containing 0.2 M sucrose, for 1 h at room temperature. After rinsing with the same buffer solution, they were dehydrated with ethanol series and Cell division of O. marina
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embedded in methacrylate resin according to the method of Baskin and Gubler (1992). Semi-thin sections (0.5–1.0 µm in thickness) were prepared using an ultramicrotome and mounted on a glass slide. The resin was removed by propylene oxide prior to immunohistochemical staining. The staining procedure was almost the same as described for whole-mount preparations. DM1A was used to visualize microtubules and mouse monoclonal anti-actin antibody mAbC4 (Boehringer Mannheim) was used to visualize actin. Some sections were post-stained with DAPI to visualize chromatin. Preparations were observed with a microscope (Nikon LabPhoto.) equipped for epifluorescence microscopy, and images were collected with a CCD camera controlled by a personal computer.
3. RESULTS Morphological changes taking place during cell division were observed by TEM, SEM and fluorescence microscopy. It was difficult to separate the time course of cell division of O. marina into well-defined phases as observed in higher eukaryotic cells. The cell size is quite variable, from 5 to 25 µm in length along the longitudinal axis. When cells are starved for long periods, all cells gradually stop dividing and take the rice grain shape of interphase cells. When new food is added to the culture medium, both larger and smaller cells begin to divide asynchronously after a lag time of 8–12 h corresponding to a non-feeding period. This means that the cell size cannot be related to cell cycle phases. Furthermore, there are great diversities in morphological features of mitosis from that of higher eukaryotes. Therefore, based on our observations of the morphological features of the nucleus and the cell shape, we conventionally separated the division cycle into four phases: interphase, prophase, metaphase, and ana–telophase. To visualize microtubules, we used mouse monoclonal anti-β-tubulin (D66) and monoclonal anti-α-tubulin (DM1A) antibodies both for whole-mount preparations and thin sections. Microtubules stained by D66 looked continuous, while DM1A staining was grainy. A monoclonal anti γ-tubulin antibody G19 was used to visualize MTOCs. It is known that G19 can weakly recognize β-tubulin and label the microtubules in several organisms (Horio et al., unpublished observation). However, G19 did not stain cortical microtubules in O. marina.
3.1. Interphase Cells at interphase were first observed for comparison to dividing cells. Interphase cells have an overall appearance like a rice grain as shown in figure 1a. The cell body is separated by an incomplete girdle into two parts: a large anterior epicone and a small posterior Kato et al.
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Figure 1. Overall appearance of O. marina in the course of cell division, observed by SEM. a. Interphase. Arrows point to extruded trychocyst filaments and arrowheads point to the girdle that defines the boundary of epicone and hypocone. b. Prophase. Both flagella have already duplicated. Longitudinal flagella (lf1 and lf2) have separated, while transverse flagella (tf1, 2) have not yet separated. c. Metaphase or early ana–telophase. Two flagellar apparatus and tentacles(arrows) separate to opposite direction along the long axis of the cell body. d. Ana–telophase. The furrowing is proceeding. Shapes of future daughter cells are different, as a result of unequal cell division. lf: longitudinal flagellum; tf: transverse flagellum; te: tentacle. Bar corresponds to 5 µm.
hypocone (arrowheads). Two flagella, a longitudinal flagellum and a transverse flagellum emanate from the indented basal portion of the hypocone or the neck of tentacle. At the basal portion of both flagella, complicated microtubular and fibrous structures are found. These structures have already been reported by previous authors (Roberts, 1985; Roberts et al., 1993), and therefore will not be shown in this paper. A TEM image in low magnification is shown in figure 2. A spherical nucleus with many chromosomes is observed at the central portion of the cell. A prominent nucleolus is always located at the centre of the nucleus. The chromosomes are composed of a bundle of fine filaments embedded in electron-dense material (figure 3). By observations of serial sections, it was confirmed that most of the chromosomes are attached to the
nuclear envelope by one or both ends (as in figure 3, arrows). Besides chromosomes, many electron-dense granules are scattered in the nucleoplasm (arrowheads). The cytoplasm is divided into two parts, the inner cytoplasm with a complicated reticular and vesicular system, and the peripheral thin cytoplasm. A layer of large vacuoles separates these two parts. In the inner cytoplasm, mitochondria with characteristic tubular cristae, trychosysts, and phagosomes of various size are observed (figure 2). In the peripheral cytoplasm (figure 4), a thin layer between flattened vesicles (asterisks) and the plasma membrane corresponds to the theca (between arrowheads). Bundles of microtubules (arrows) are observed just underneath. Each bundle is
Figure 2. A longitudinally sectioned interphase cell observed by TEM in a low magnification. m: mitochondria, no: nucleolus; ph: phagosomes; t: trychocyst; v: cortical vacuole. Bar: 2 µm. Figure 3. Enlargement of chromosomes in an interphase cell. Chromosomes are composed of fine filaments and closely apposed to the nuclear envelope with their ends (arrows). Arrowhead: electron-dense granule. Bar: 1µm. Figure 4. Structure of the lateral cell cortex. Bundles of microtubules (arrows) are observed just beneath flattened vesicles (asterisks). The layer between black arrowheads is the theca. m: mitochondrion; v: cortical vacuole. Bar: 0.2 µm. Figure 5. Two sets of basal bodies and appendages in a cell during prophase. Numbers at lower right corner indicate section number in serial sections cut vertically in the antero–posterior direction. Two sets are about 1 µm apart along the longitudinal axis of the cell. lb: longitudinal basal body; tb: transverse basal body; lmr: longitudinal microtubular root; ac: accessory centriole. Bar; 0.5 µm. Figure 6. A part of nucleus at early prophase. Note deposition of electron-dense material at some of nuclear pores (arrows). Bar: 0.5 µm. Figure 7. Localization of γ-tubulin at the periphery of a nucleus in prophase. Whole cells were stained with monoclonal anti-γ-tubulin (G19) by the method described in materials and methods. Image was taken with an ordinary epifluorescence microscope.
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Figure 8. Three dimensional organization of cortical and flagellar microtubules stained with D66 mAb in interphase cell. Images are reconstructed on a workstation from images obtained with a confocal laser microscope. a. Ventral surface view. b. Dorsal view; (a) and (b) are obtained from one specimen. Arrows indicate microtubular ridge of the girdle. c. Apical view. d. Bottom view. e. Microtubular desmose, which connects the nucleus and the flagellar base (arrow). Bars: 5 µm (a–d); 1 µm (e).
composed of 2–8 microtubules that run evenly spaced and parallel to each other, and represent the so-called cortical microtubules. Confocal images of cortical microtubules stained with D66 are shown in figure 8. The cell cortex contains many microtubular bundles of different thickness, probably a reflection of the number of microtubules contained in each bundle (figure 8a, ventral view and 8b, dorsal view of the same specimen). All cortical microtubule bundles in the epicone (anterior of the cell) converge at one focal point at the cell apex. The distal ends of these microtubules in the ventral and lateral halves join the microtubular rim, which represents the ridge of the girdle (arrows), while dorsal microtubules join the dorsal lip of the girdle, the posterior tip of the cell. A view from the top is shown in figure 8c. From the medial part of the cortex, microtubule bundles gradually fuse together and converge in one small focal area like the spokes of a wheel, so Cell division of O. marina
that the anterior cortex is covered by an even density of microtubules. At the cortex of the hypocone, bundles of microtubules, which are considerably thicker than that of epicone, start from the flagellar base and extend posteriorly to the dorsal lip of the girdle (figure 8d). They converge to the focal area wider than the focal point of the apical tip. The basal circumference of the tentacle is covered by a thick layer of microtubules, producing a brighter fluorescence. A thin microtubular desmose connects the flagellar base with the periphery of the nucleus (figure 8e, arrow).
3.2. Prophase The duplication of the flagellar apparatus is a landmark of early prophase. The overall appearance of the cell is the same as that at interphase except for the duplicated flagellar apparatus (figure 1b). New axonemes elongate along parent ones and then separate Kato et al.
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into two flagella. Longitudinal flagella seem to separate earlier than transverse ones do. At the base of these flagella, two pairs of basal bodies are present, which must have been duplicated at an earlier stage. Figure 5 shows two adjacent flagellar apparatus in a single serial section viewed from the anterior side. These apparati are separated by about 1 µm calculated from the number of sections. Two basal bodies baring flagella are arranged perpendicular to each other in close vicinity. From the wall of the longitudinal basal body, sheets of microtubules (longitudinal microtubular root; lmr) extend toward the surface of the hypocone. This appendage structure has the same structure and orientation in the two duplicated sets of flagellar apparatus. The lmr of posterior flagellar apparatus (lmr2) overlies that of anterior flagellar apparatus (lmr1). Other flagellar appendages have been also duplicated exactly in the same manner. Flagellar axonemes have been dissected, a very frequent phenomenon in this species, probably because of unsuitable fixation. In addition, two short accessory centrioles are observed near these basal bodies in this specimen, presumably representing duplicating basal bodies in the preparation of the next cell cycle. At this stage, cortical microtubule bundles do not change their interphase arrangement. The nucleus itself is still spherical (figure 6) with a nucleolus located at its centre. Multiple depositions of electron-dense material about 100 nm wide become apparent in the nuclear envelope (arrows). Due to their size and location, these densities are likely to be accumulated at some of the nuclear pores. Figure 7 shows localization of γ-tubulin at the periphery of the nucleus at this stage. Bright spots probably represent the densities identified by electron microscopy in nuclear pores. The confocal image of a nucleus shortly after this stage is shown in figure 12a. In the nucleus, many radially arranged microtubules appear with one end at the nuclear envelope. The desmose (arrow) present in interphase still connects basal bodies to the periphery of the nucleus.
3.3. Metaphase Cells gradually elongate in the antero–posterior direction and take oblong shapes. Duplicated flagellar apparatus separate and move in opposite directions along the longitudinal axis. Concomitant with the separation of the flagellar apparatus, tentacles are also duplicated and separated in the same way (figure 1c). At this stage, the nucleus is flattened along the longitudinal axis and takes on a discoid shape (figure 12b). From nuclear pores of flattened faces of the nuclear envelope, bundles composed of 6–9 microtubules run to the opposite side of the nucleus through chromosome clusters (figure 9). Each microtubule grows more than 3/4 of the short axis of the disk, but do not attach Cell division of O. marina
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to the opposite side of the nuclear envelope. Microtubule bundles originating from opposite faces of the disc-shaped nucleus seem to be paired, in other words, two densities in nuclear pores facing each other in opposite parts of the nuclear membrane become paired and organize two presumably antiparallel microtubule bundles. These constitute a kind of elementary mitotic spindle. Chromosomes are also arranged longitudinally along the short axis of the discoid nucleus although no equatorial plate can be recognized. Connections between chromosomes and microtubules, through kinetochores or some other special structures, are not recognized. The close relation between chromosomes and nuclear envelope that is observed in interphase nucleus has disappeared at this stage. A sagittal section double-stained with DM1A and DAPI (figure 13a) shows a parallel array of microtubule bundles in a flattened nucleus in which chromosomes are arranged in a plate. A transverse section of the nucleus at this stage shows that the number of microtubule bundles is over 70 (figure 13b).
3.4. Ana–telophase At ana–telophase, the flagellar apparatus and the hypocones separate further along the main cell axis. The cell body gradually elongates and then begins to be constricted at the circumference of its meridian plane between the anterior and the posterior parts (figure 1d). The microtubular cytoskeleton of early ana–telophase cells is reconstructed from confocal images (figure 12c, d; the same specimen). Two flagellar apparatus are separated by several microns (figure 12d, arrows). In the equatorial cortex, microtubule bundles appear to be bisected all around the cell surface (figure 12c, arrow). Probably at the same position of the cortex, a band of actin (contractile ring) is detected by immunostaining with anti-actin antibody mAbC4 (figure 13d, arrows). The presence of actin is observed on the nuclear envelope at the same time (arrowheads). In the nucleus, chromosomes separate into two plates, which indicates the beginning of ana–telophase (figure 13c, d). Many parallel microtubule bundles that run across the nucleus along its short axis are evident at this stage. Each bundle seems to be almost continuous across the nucleus. At this time, elongation and/or sliding of microtubule bundles must occur, which causes elongation of the nucleus along the antero–posterior axis and simultaneous separation of the chromosomes. The nucleolus takes an oblong shape with both ends apposed to the nuclear envelope (figure 13c, arrow). Then the equator of the disc-shaped nucleus becomes narrower and takes on a dumbbell shape (figure 10 and figure 12d). At the constricted area of the nucleus, microtubules are densely packed crossing each other in a close vicinity (figure 11), which proKato et al.
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Figure 9. View of a part of a nucleus at metaphase. Note microtubules emanating from a dense deposition of nuclear pore (arrows). Bar: 0.5 µm. Figure 10. A dumbbell-shaped nucleus at ana–telophase. An elongated nucleolus is at the centre of the nucleus (no). Bar: 1 µm. Figure 11. Microtubules at the constricted portion of an ana–telophase nucleus. Arrowheads indicate the limit of nuclear envelope. Bar: 0.2 µm.
duces quite a strong fluorescence signal in that area (figure 12d). Each microtubule runs rather straight than curved, suggesting their twisting around the longitudinal axis. At late ana–telophase, the constriction of the cell body proceeds. During constriction and elongation of the cell body, the future anterior daughter cell has an incomplete posterior part of the cytoskeleton and vice versa for the future posterior daughter cell (figure 12e). This might indicate that the posterior part of the anterior cell and that the anterior part of the posterior cell are rebuilt de novo during and after ana–telophase. The nucleus has already divided and positioned at the centre of the future daughter cells at this stage (figure 12f, the same specimen as figure 12e). Intranuclear microtubules remained until that time and then disappear. Considerably before the completion of cytokinesis, two daughter nuclei are formed (figure 13e), then the Cell division of O. marina
actin ring disappears, leaving a broad cytoplasmic connection between the two daughter cells (figure 13f, between two arrows). After completion of cytokinesis, the two daughter cells remain connected by a thin cytoplasmic bridge, which persists for more than 15 min. Occasionally, the next division cycle progresses before the final separation of the two daughter cells.
4. DISCUSSION In some dinoflagellates, new mitotic microtubules appear in the cytoplasm at the time of nuclear division, while the nuclear envelope persists and mitosis is intranuclear (Spector and Triemer, 1981). From an evolutionary point of view, this represents an intermediate process of mitosis between yeast and higher eukaryotes (Alberts et al., 1994). We confirm earlier reports showing that in O. marina, the microtubular mitotic Kato et al.
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Figure 12. Three dimensional organization of microtubular systems in dividing cells. a. Intranuclear microtubules in a spherical nucleus at prophase. The arrow indicates the microtubular desmose between the nucleus and the flagellar base. b. Intranuclear microtubules of a metaphase cell. Bundles of microtubules are arranged in parallel to the short axis of a disc-shaped nucleus. c. Cortical microtubules of an early ana–telophase cell. Microtubular bundles are separated at the cell body equator (arrow). d. Intranuclear microtubules in the same cell as (c). Arrows and arrowheads indicate flagellar base and microtubular desmose respectively. e. Ventro–lateral view of cortical microtubules and flagellar apparatus of an ana–telophase cell. f. Intranuclear microtubules in the same cell as (e). They appear to have been separated into two daughter nuclei. Bar: 5 µm.
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Figure 13. Arrangement of microtubules and localization of actin in semi-thin sections. Sections were stained with an anti-α-tubulin mAb (DM1A) and DAPI (a–c), or an anti-actin mAb (C4) and DAPI (d–f). Black and white images were incorporated in a personal computer, artificially colored and then overlapped. The brightness of FITC signals were intensified in (d–f) in order to see the boundary of cytoplasm. a. A longitudinal section of a metaphse cell. Chromosomes (blue) and microtubules (yellow) are overlapped in the disc-shaped nucleus. b. A transverse section of a metaphase cell. c. A longitudinal section of an early ana–telophase cell. Chromosomes are separated into two planes. Arrow indicates the elongated nucleolus. d. An early ana–telophase cell. Bright yellow spots (arrows) indicate actin bundle at cell cortex. The nuclear envelope is also stained by anti-actin antibody (arrowheads). e. An ana–telophase cell. The contractile ring (arrows) at the constricted cell cortex and actins around the nuclear envelope are apparent. f. A late ana–telophase cell. Two daughter cells are still connected by broad cytoplasm (between two arrows), while the actin ring has already disappeared. Bar: 5 µm.
apparatus is intranuclear (Triemer, 1982; Gao and Li, 1986; Kato et al., 1997). Such an intranuclear microtubular division system is exceptional among dinoflagellates as described to date. In many ciliates, intranuclear microtubules are frequently observed. In Paramecium for instance, microtubules appear both in the macro- and the micronucleus during vegetative division, and in the micronucleus during mitosis (Ishida et al., 1999). In this regard, O. marina is more similar to ciliates than other dinoflagellates. Since the nuclear envelope does not break down during nuclear division, translocation of tubulin molecules from the cytoplasm to the nucleus is a prerequisite to allow microtubule polymerization in the nucleus. Thus, it appears that this organism can segregate tubulin molecules into a cytoplasmic and an intranuclear pool devoted to different functions. Whether these tubulin pools have the same isotype and isoform composition remains to be established, since several isotypes have been reported (Laferrière et al., 1997; Huitorel et al.; 1999). Furthermore, the presCell division of O. marina
ence of a MTOC is a prerequisite to microtubule polymerization in yeast and animal cells. In the present paper, we describe the mitotic microtubule nucleation sites inserted at several loci of the nuclear envelope (figure 7 and figure 9). Their structure is comparable to the spindle pole body of budding yeast (Donaldson and Kilmartin, 1996) and that of fission yeast (Ding et al., 1997; Hagen and Yanagida, 1997) especially in terms of the appearance and the size. The number of microtubules nucleated per site is also similar. However, yeast spindle pole bodies are present throughout the cell cycle, either embedded in the nuclear envelope or at its outer surface, and simultaneously nucleates microtubules on both sides of the nuclear envelope at mitosis. We could not detect any cytoplasmic microtubules nucleated from the density in the nuclear pore. There is a possibility that this might be due to the labile nature of cytoplasmic microtubules in this species. On the other hand, it is clear that the density appears only at the beginning of mitosis. Despite the apparent resemblance, these structures may be different in moKato et al.
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lecular composition and in their mechanisms of regulation. Interestingly, plant cells do not have such a well-defined MTOC, but the microtubule nucleation material is dispersed around the outer face of the nuclear envelope. In O. marina, the nucleating material may also be dispersed, then well focused during mitosis. In euplotes, the presence of γ-tubulin in the nucleus throughout the cell cycle has been reported (Liang et al., 1996). In O. marina, we have detected γ-tubulin on the nuclear envelope using the monoclonal anti γ-tubulin antibody G19 (figure 7). Therefore, the electron-dense material accumulated at nuclear pores might correspond to the structure that contains γ-tubulin, which is a component of MTOCs. Further detailed study using immuno-electron microscopy is awaited. The presence of an actin band at furrowing cell surface reported in this paper confirms the presence of actin previously described in other dinoflagellates (Soyer-Gobillard et al., 1996). The cytokinesis is likely to be mediated by the contraction of this band. On the other hand, the constriction of the nucleus precedes cytokinesis. This means that the constriction of the nucleus cannot be caused by contraction of the cytoplasmic contractile ring. There is a possibility that the intranuclear microtubule system itself produces the physical power to constrict the nucleus and separate the two daughter nuclei. We observed the elongation of intranuclear microtubules during nuclear division. If the elongation and the sliding of microtubules of opposite polarities occur, then the nucleus would inevitably become weaker at its equator, and be constricted and twisted. Our observations shown in figure 11 and figure 12d strongly suggest this possibility. In addition, there is a possibility that actin also participates in this process. The presence of actin on the nuclear envelope at metaphase (figure 13d) may suggest a potential role of actin polymers in the constriction of the nucleus. Therefore, if both microtubules and actin fibres participate in the nuclear division in harmony, the mechanism of nuclear division of O. marina may be comparable to the cell division of higher eukaryotic cells. The polar region of the nucleus is connected to the flagellar apparatus by a thin desmose containing microtubules. However, it seems impossible to restrict the physical force necessary for nuclear elongation to the desmose, because it is connected almost vertically to the polar region. More likely, the desmose may act to define the position of polar regions, and possibly of the nucleus itself. In Chlamydomonas, for instance, a centrin filament-based desmose connects the basal bodies to the nucleus and positions it at the time of mitosis (Salisbury, 1995). In several species of dinoflagellates, chromosomes are connected to nuclear pores and are separated by Cell division of O. marina
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the movement of the nuclear envelope (see Spector, 1984). In those cases, extranuclear microtubules seem to render a shearing force to the nuclear envelope, and provide an orientation to the movement of the nuclear envelope and chromosomes. In dividing cells of O. marina, we did not detect any direct connection between chromosomes and the nuclear envelope or microtubules. The mechanism of chromosome separation is still uncertain in this species and remains a matter for further investigations. The cell division of O. marina is an unequal division; the dividing cell keeps its antero–posterior polarity and the division plate is perpendicular to this axis. However, after completion of cytokinesis, the two daughter cells show equivalent structures. Therefore, during the last stage of cell division, a new posterior half has to be rebuilt along the old anterior half and vice versa for the posterior half, according to yet unknown mechanisms. Basal bodies and flagella have been duplicated at an early stage of prophase. Microtubular and fibrous appendages are also duplicated until this stage. Two sets of flagellar apparatus separate at metaphase as the cell body elongates. It is clear that the new hypocone and tentacle are duplicated and separated along with the separation of flagella. When daughter flagellar apparatus have separated by several microns, a belt-like gap appears at the midst of the cortical microtubule arrays. It is possible that the cortical microtubules are cut at the equatorial area, which leads to the gap formation. Microtubule severing factors, especially active in M phase, have been identified in higher eukaryotes (Shiina et al., 1995). Similar factors might be involved in the gap formation of microtubule arrays in O. marina. Since the position of the gap correlates with the location of the actin ring, the localization of such factors might be actin-dependent. Double staining using anti-actin and anti-tubulin antibodies will help to get better understanding about the process of gap formation. At this time and later, cortical microtubules must be elongated to reconstruct two full cell body structures from both halves. The polymerization of tubulin occurs preferentially at nucleation centres or at pre-existing microtubule ends in vivo. The absolute polarity of microtubules in dinoflagellates and the precise location of cortical microtubule nucleation sites are still unknown. In this study, we also could not detect any special structures at the anterior or posterior focal areas, or at the cell equator at the site and time where the ‘actin belt’ is observed. There is a possibility that a recently cut end may be a preferential elongation site, regardless of its polarity, provided the microtubule is stable enough. If this is the case, microtubules can simply elongate to rebuild a complete cortical network. There is an alternative possibility that cortical microtubules elongate at their plus ends with their original Kato et al.
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polarities. In this case cortical microtubules of one daughter cell would re-grow by simple elongation. Cortical microtubules of the other daughter cell would elongate from some new nucleating site and the preexisting cortical network should gradually disappear. A candidate for new nucleating sites would be the newly formed ridge that separates the epicone and the hypocone. The ridge must be organized by the basal bodies at the time of flagellar duplication and separation. Therefore, one may propose that the basal bodies and their associated structures, may contain all the microtubule nucleating material necessary to build the interphase microtubule networks, i.e. the flagellar apparatus and appendages and cortical microtubules. On the other hand, the cytoskeletal microtubules may not necessarily be continuous throughout cortical microtubular bundles. The longitudinal microtubular band of Tetrahymena is composed of stacks of short and slightly angled microtubules (Ng, 1979). These microtubules could be regenerated by a lateral nucleation mechanism from preexisting microtubules reported in trypanosomes (Sherwin and Gull, 1989). In both cases the polarity of microtubules is preserved. Which model applies to O. marina is a matter for future investigations. Acknowledgments. We wish to warmly thank Pr. H. Hotani, Nagoya University, for his welcome in his laboratory to run the confocal microscopy. We are indebted to Dr. C. Gagnon, McGill University, for the use of the D66 monoclonal antibody. Part of this work was supported by a grant from the ‘Centre national de la recherche scientifique’ (CNRS, France) to P.H. P.H. also acknowledges receipt of the fund of Nagoya City University for invited scholars.
REFERENCES Alberts, B., Bray, D., Lewis, J., Raff, M., Roberts, K., Watson, J.D., 1994. Molecular Biology of the Cell. Garland Publisher, New York, pp. 941–943. Baskin, T., Gubler, F., 1992. Improvements in immunostaining samples embedded in methacrylate: localization of microtubules and other antigens throughout developing organs in plants of diverse taxa. Planta 187, 405–413. Bhaud, Y., Salmon, J.M., Soyer-Gobillard, M.O., 1991. The complex cell cycle of the dinoflagellate protoctist Crypthecodinium cohnii as studied in vivo and by cytofluorimetry. J. Cell Sci. 100, 675–682. Cachon, J., Cachon, M., Cosson, M.P., Cosson, J., 1988a. The flagellar rod: a structure in search of a function. Biol. Cell 63, 169–181. Cachon, M., Cosson, J., Cosson, M.P., Huitorel, P., Cachon, J., 1988b. Ultrastructure of the flagellar apparatus of Oxyrrhis marina. Biol. Cell 63, 159–168. Cosson, J., Cachon, M., Cachon, J., Cosson, M.P., 1988. Swimming behavior of the unicellular biflagellate Oxyrrhis marina: in vivo and in vitro movement of the two flagella. Biol. Cell 63, 117–126. Donaldson, A.D., Kilmartin, J.V., 1996. Spc42p: a phosphorylated component of the S. serevisiae spindle pole body (SPB) with an essential function during SPB duplication. J. Cell Biol. 132, 887–901.
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Biology of the Cell 92 (2000) 583–594 Ding, R., West, R.R., Morphew, M., McIntosh, J.R., 1997. The spindle pole body of Schizosaccharomyces pombe enters and leaves the nuclear envelope as the cell cycle proceeds. Mol. Biol. Cell 8, 1461–1479. Dodge, J.D., Crawford, R.M., 1971. Fine structure of the dinoflagellate Oxyrrhis marina. I – The general structure of the cell. Protistologica 7, 295–304. Gao, X.P., Li, J.Y., 1986. Nuclear division in the marine dinoflagellate Oxyrrhis marina. J. Cell Sci. 85, 161–175. Grell, K.G., 1973. Protozoology. Springer-Verlag, Berlin, Heiderberg, New York. Hagen, I., Yanagida, M., 1997. Evidence for cell cycle-specific, pole body-mediated nuclear positioning in the fission yeast Schizosaccharomyces pombe. J. Cell Sci. 110, 1851–1866. Hohfeld, I., Melkonian, M., 1998. Lifting the curtain? The microtubular cytoskeleton of Oxyrrhis marina, Dinophyceae, and its rearrangement during phagocytosis. Protist 149, 75–88. Horio, T., Basaki, A., Takeoka, A., Yamato, M., 1999. Lethal level overexpression of γ-tubulin in fission yeast causes mitotic arrest. Cell Motil. Cytoskel. 44, 284–295. Huitorel, P., Audebert, S., White, D., Cosson, J., Gagnon, C., 1999. Role of tubulin epitopes in the regulation of flagellar motility. In: Gagnon, C. (Ed.), The male gamete: from basic science to clinical applications. Cache River Press, Vienna, IL, pp. 475–491. Ishida, M., Nakajima, Y., Kurokawa, K., Mikami, K., 1999. Nuclear behavior and differentiation in Paramecium caudatum, analyzed by immunofluorescence with anti-tubulin antibody. Zool. Sci. 16, 915–926. Kato, K.H., Moriyama, A., Huitorel, P., Cosson, J., Cachon, M., Sato, H., 1997. Isolation of the major basic nuclear protein and its localization on chromosomes of the dinoflagellate, Oxyrrhis marina. Biol. Cell 89, 43–52. Laferrière, N.B., MacRae, T.H., Brown, D.L., 1997. Tubulin synthesis and assembly in differentiating neurons. Biochem. Cell Biol. 75, 103–117. Liang, A., Ruiz, F., Heckmann, K., Klotz, C., Tollon, Y., Beisson, J., Wright, M., 1996. Gamma-tubulin is permanently associated with basal bodies in ciliates. Eur. J. Cell Biol. 70, 331–338. Ng, S.F., 1979. Unidirectional regeneration is an intrinsic property of longitudinal microtubules in Tetrahymena – an in vivo study. J. Cell Sci. 36, 109–119. Perret, E., Davoust, J., Albert, M., Besseau, L., Soyer-Gobillard, M.O., 1993. Microtubule organization during the cell cycle of the primitive eukaryote dinoflagellate Crypthecodinium cohnii. J. Cell Sci. 104, 639–651. Roberts, K.R., 1985. The flagellar apparatus of Oxyrrhis marina. Pyrrophyta. J. Phycol. 21, 641–655. Roberts, K.R., Rusche, M.L., Taylor, F.J.R., 1993. The cortical microtubular cytoskeleton of Oxyrrhis marina, Dinophyceae, observed with immunofluorescence and electron microscopy. J. Phycol. 29, 642–649. Roberts, K.R., Roberts, J.E., 1991. The flagellar apparatus and cytoskeleton of the dinoflagellates, a comparative overview. Protoplasma 164, 105–122. Salisbury, J.L., 1995. Centrin, centrosomes, and mitotic spindle poles. Curr. Opin. Cell Biol. 7, 39–45. Sherwin, T., Gull, K., 1989. Visualization of detyrosination along single microtubules reveals novel mechanisms of assembly during cytoskeletal duplication in trypanosomes. Cell 57, 211–221. Shiina, N., Gotoh, Y., Nishida, E., 1995. Microtubule-severing activity in M phase. Trend Cell Biol. 5, 283–286. Soyer-Gobillard, M.O., Ausseil, J., Geraud, M.L., 1996. Nuclear and cytoplasmic actin in dinoflagellates. Biol. Cell 87, 17–35. Spector, D.L., Triemer, R.E., 1981. Chromosome structure and mitosis in the dinoflagellates: an ultrastructural approach to an evolutionary problem. BioSystems 14, 289–298. Spector, D.L., 1984. Dinoflagellate nuclei. In: Spector, D.L. (Ed.), Dinoflagellates. Academic Press, New York, pp. 107–147. Triemer, R.E., 1982. A unique mitotic variation in the marine dinoflagellate Oxyrrhis marina (pyrrophyta). J. Phycol. 18, 399–411.
Kato et al.
Original article
Dynamic changes in microtubule organization during division of the primitive dinoflagellate Oxyrrhis marina Koichi H. Katoa*, Akihiko Moriyamaa, Tomohiko J. Itohb, Masayuki Yamamotoc, Tetsuya Horioc, Philippe Huitoreld a
Institute of Natural Sciences, Nagoya City University, Nagoya 467-8501, Japan
b
Division of Biological Sciences, Graduate School of Science, Nagoya University, Nagoya 464-8601, Japan
c
Department of Food Microbiology, School of Medicine, The University of Tokushima, Tokushima 770-0042,
Japan d
Laboratoire de biologie du développement, UMR 7009, CNRS/UPMC, Station zoologique, 06230 Villefranche-
sur-mer, France Received 18 September 2000; accepted 11 December 2000
The marine dinoflagellate Oxyrrhis marina has three major microtubular systems: the flagellar apparatus made of one transverse and one longitudinal flagella and their appendages, cortical microtubules, and intranuclear microtubules. We investigated the dynamic changes of these microtubular systems during cell division by transmission and scanning electron microscopy, and confocal fluorescent laser microscopy. During prophase, basal bodies, both flagella and their appendages were duplicated. In the round nucleus situated in the cell centre, intranuclear microtubules appeared radiating toward the centre of the nucleus from densities located in some nuclear pores. During metaphase, both daughter flagellar apparatus separated and moved apart along the main cell axis. Microtubules of ventral cortex were also duplicated and moved with the flagellar apparatus. The nucleus flattened in the longitudinal direction and became discoid-shaped close to the equatorial plane. Many bundles of microtubules ran parallel to the short axis of the nucleus (cell long axis), between which chromosomes were arranged in the same direction. During ana–telophase, the nucleus elongated along the longitudinal axis and took a dumbbell shape. At this stage a contractile ring containing actin was clearly observed in the equatorial cortex. The cortical microtubule network seemed to be cut into two halves at the position of the actin bundle. Shortly after, the nucleus divided into two nuclei, then the cell body was constricted at its equator and divided into one anterior and one posterior halves which were soon rebuilt to produce two cells with two full sets of cortical microtubules. From our observations, several mechanisms for the duplication of the microtubule networks during mitosis in O. marina are discussed. © 2000 Éditions scientifiques et médicales Elsevier SAS dinoflagellate / Oxyrrhis / mitosis / microtubule / actin
* Correspondence and reprints. E-mail address: [email protected] (K.H. Kato).
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1. INTRODUCTION Oxyrrhis marina is a heterotrophic free-living dinoflagellata and is commonly found in tide pools along the coast. The mode of nuclear division in dinoflagellates has been studied in a few species, including O. marina, among more than 6000 known species, and many differences from that of higher eukaryotic cells have been reported. In these organisms the nuclear envelope does not break down during mitosis, which implies that they have chromosomes in the nucleus throughout the cell cycle and exhibit intranuclear mitosis. However, at least some of them do not show the typical closed mitosis found in yeast for example. Several cytoplasmic channels containing microtubules are formed through the nucleus (Spector and Triemer, 1981). O. marina shows an exceptional type of mitosis with an intranuclear microtubular system which develops at the time of cell division (Triemer, 1982; Gao and Li, 1986; Kato et al., 1997). This characteristic has lead to an argument regarding the position of this organism in the phylogenetic tree. The cortical microtubular systems and the flagellar apparatus have been studied in several dinoflagellates including O. marina (Roberts and Roberts, 1991). In O. marina, the cortical cytoskeleton is known to be composed of bundles of several parallel microtubules (Dodge and Crawford, 1971; Roberts et al., 1993), and is rather flexible (Hohfeld and Melkonian, 1998). The flagellar apparatus of dinoflagellates presents several characteristic features. Two basal bodies give rise to two flagella and several sets of flagellar roots, made of microtubules and fibrous material. Besides the usual axonemal structure, flagella contain highly organized accessory fibrillar appendages, whose structure and function are quite distinct (Cachon et al., 1988a; Cachon et al., 1988b; Cosson et al., 1988; Roberts and Roberts, 1991; Roberts et al., 1993). In the dinoflagellate Crypthecodinium cohnii, both flagella are lost as cells enter cell division, and two daughter cells receive an equal half of the mother cell (Bhaud et al., 1991; Perret et al., 1993). In contrast, O. marina do not loose flagella during cell division and continue swimming throughout the cell cycle. During the early steps of cell division, the flagellar apparatus is duplicated and separated into two pairs. The cell keeps its antero–posterior as well as dorso–ventral polarity during cell division, and consequently each daughter cell receives either an anterior or a posterior half of the mother cell body. The other half has to be built de novo during the late steps of cell division. Therefore, while most higher eukaryotic cells loose their typical interphase cytoskeleton at the entry in mitosis, O. marina duplicates and reorganizes it as in many other protists (see Grell, 1973). In Tetrahymena (Ng, 1979) and Trypanosoma (Sherwin and Gull, 1989) Cell division of O. marina
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for instances, detailed studies were reported concerning the process of duplication of cytoskeletal microtubules, a complex but well regulated process. In the present work, we have studied three microtubular systems, the flagellar apparatus, cortical cytoskeletal microtubules and intranuclear microtubules during cell division in O. marina, using electron microscopy and immunolocalization by confocal laser microscopy. We also examined the participation of actin in cell division in this species. Based on our observations, new mechanisms of duplication of the microtubular cytoskeleton are discussed.
2. MATERIAL AND METHODS 2.1. Cell culture O. marina was cultured in the laboratory essentially by a method described previously, except that quail yolk was used in some experiments as food instead of E.coli. Quail yolk was prepared as follows. One mL of the yolk was suspended in 50 mL of culture medium, boiled for several minutes, homogenized with a Dounce homogenizer (clearance: 10 µm) and centrifuged at 3000 × g for 20 min. The pellet was resuspended in the same volume of culture medium and filtered through nylon mesh (pore size: 10 µm) and used as food. In order to prevent bacterial growth, penicillin G (100 U·mL–1) and streptomycin (100 µg·mL–1) were added to the culture medium. The quail yolk is comparably as effective as E. coli for the food. When the newly prepared yolk was added to the original culture during the stationary phase, most cells started dividing within 12 h.
2.2. Electron microscopy For transmission electron microscopy (TEM), samples were fixed with 4% glutaraldehyde in 0.1 M phosphate buffer (pH 7.0) containing 0.15 M sucrose, 10 mM MgCl2 and 10 mM EGTA, for 45 min at room temperature room temperature. After rinsing in 0.1 M phosphate buffer containing 0.2 M sucrose, they were postfixed with 0.5% OsO4 in the same buffer for 45 min at 4°C. They were then dehydrated through ethanol series and embedded in Spurr’s low viscosity resin on a flat plastic plate. Thin sections were cut using a RMC MT6000 ultramicrotome and examined in a Hitachi H-7100 electron microscope. For scanning electron microscopy (SEM), samples were mounted on 1% polylysine coated cover slips and fixed with the mixture of 3% glutaraldehyde and 0.2% OsO4 in the buffer solution (50 mM borate buffer (pH 8.2) and 250 mM NaCl) for 15 min at room temperature. They were then transferred into the buffer solution containing 3% glutaraldehyde and 1% tannic Kato et al.
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acid and fixed for 1.5 h, followed by post-fixation with 1% OsO4 in the buffer solution for 3 h at 4°C. After rinsing with distilled water, they were dehydrated with ethanol series, infiltrated with iso-amylacetate and critical point dried. A small piece of coverslip was fixed on the specimen stub and spatter coated with gold, and then observed with a JEOL JSM-T100 scanning electron microscope.
2.3. Immunocytochemistry For whole mount preparations, samples were placed on 0.5% polylysine coated cover slips and fixed with 4% formaldehyde in 50% seawater for 4 min. Then, an equal volume of Tris buffer solution (TBS) containing 1% Triton X-100 was added and mixed for 4 min to permeabilize the cells. TBS was composed of 20 mM Tris and 0.3 M NaCl, and adjusted to pH 7.6 with 1 N HCl. After permeabilization, samples were rinsed several times with TBS containing 0.1% Triton X-100 (TBST) followed by rinsing with TBST containing 1% donkey whole serum (TBSTD). Mouse monoclonal anti-β-tubulin antibody D66 (a generous gift of Dr. C. Gagnon, McGill University), mouse monoclonal anti-αtubulin DM1A (Sigma) were used to visualize microtubules. A mouse monoclonal anti-γ-tubulin antibody G19 (Horio et al., 1999) was used to visualize microtubule organizing centres (MTOCs). The G19 antibody has been raised against bacterially expressed Schizosaccharomyces pombe γ-tubulin. Epitope mapping using truncated γ-tubulin peptides revealed that the sequence recognized by G19 is localized within the peptide GGAGNNWANGYSHA (Horio et al., unpublished observation). This region of γ-tubulin is very well conserved among conventional γ-tubulins. Samples were incubated with these antibodies diluted to 1:100–1:1000 with TBSTD for 12 h at 4°C. After rinsing with TBST, they were incubated with FITCconjugated donkey anti-mouse IgG antibody (Amersham) diluted to 1:100–1:1000 with TBSTD for 45 min at room temperature. Samples were then rinsed with TBS and mounted in 50% glycerin containing 10% of DABCO. Preparations were observed on a confocal laser scanning microscope (Yokogawa CSU-10) equipped with an argon/argon–krypton laser at the wave length of 488 nm. Digital images were collected with a cooled CCD camera (Hamaphoto C5985) and saved on a computer. To reconstruct three-dimensional structures, serial images collected at different focal planes were processed on a workstation with imaging software (VoxelView, Vital Images, Inc., USA) by simple stacking of selected images. For semi-thin sectioning, samples were fixed with 3% paraformaldehyde in 0.1 M sodium phosphate buffer (pH 7.0) containing 0.2 M sucrose, for 1 h at room temperature. After rinsing with the same buffer solution, they were dehydrated with ethanol series and Cell division of O. marina
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embedded in methacrylate resin according to the method of Baskin and Gubler (1992). Semi-thin sections (0.5–1.0 µm in thickness) were prepared using an ultramicrotome and mounted on a glass slide. The resin was removed by propylene oxide prior to immunohistochemical staining. The staining procedure was almost the same as described for whole-mount preparations. DM1A was used to visualize microtubules and mouse monoclonal anti-actin antibody mAbC4 (Boehringer Mannheim) was used to visualize actin. Some sections were post-stained with DAPI to visualize chromatin. Preparations were observed with a microscope (Nikon LabPhoto.) equipped for epifluorescence microscopy, and images were collected with a CCD camera controlled by a personal computer.
3. RESULTS Morphological changes taking place during cell division were observed by TEM, SEM and fluorescence microscopy. It was difficult to separate the time course of cell division of O. marina into well-defined phases as observed in higher eukaryotic cells. The cell size is quite variable, from 5 to 25 µm in length along the longitudinal axis. When cells are starved for long periods, all cells gradually stop dividing and take the rice grain shape of interphase cells. When new food is added to the culture medium, both larger and smaller cells begin to divide asynchronously after a lag time of 8–12 h corresponding to a non-feeding period. This means that the cell size cannot be related to cell cycle phases. Furthermore, there are great diversities in morphological features of mitosis from that of higher eukaryotes. Therefore, based on our observations of the morphological features of the nucleus and the cell shape, we conventionally separated the division cycle into four phases: interphase, prophase, metaphase, and ana–telophase. To visualize microtubules, we used mouse monoclonal anti-β-tubulin (D66) and monoclonal anti-α-tubulin (DM1A) antibodies both for whole-mount preparations and thin sections. Microtubules stained by D66 looked continuous, while DM1A staining was grainy. A monoclonal anti γ-tubulin antibody G19 was used to visualize MTOCs. It is known that G19 can weakly recognize β-tubulin and label the microtubules in several organisms (Horio et al., unpublished observation). However, G19 did not stain cortical microtubules in O. marina.
3.1. Interphase Cells at interphase were first observed for comparison to dividing cells. Interphase cells have an overall appearance like a rice grain as shown in figure 1a. The cell body is separated by an incomplete girdle into two parts: a large anterior epicone and a small posterior Kato et al.
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Figure 1. Overall appearance of O. marina in the course of cell division, observed by SEM. a. Interphase. Arrows point to extruded trychocyst filaments and arrowheads point to the girdle that defines the boundary of epicone and hypocone. b. Prophase. Both flagella have already duplicated. Longitudinal flagella (lf1 and lf2) have separated, while transverse flagella (tf1, 2) have not yet separated. c. Metaphase or early ana–telophase. Two flagellar apparatus and tentacles(arrows) separate to opposite direction along the long axis of the cell body. d. Ana–telophase. The furrowing is proceeding. Shapes of future daughter cells are different, as a result of unequal cell division. lf: longitudinal flagellum; tf: transverse flagellum; te: tentacle. Bar corresponds to 5 µm.
hypocone (arrowheads). Two flagella, a longitudinal flagellum and a transverse flagellum emanate from the indented basal portion of the hypocone or the neck of tentacle. At the basal portion of both flagella, complicated microtubular and fibrous structures are found. These structures have already been reported by previous authors (Roberts, 1985; Roberts et al., 1993), and therefore will not be shown in this paper. A TEM image in low magnification is shown in figure 2. A spherical nucleus with many chromosomes is observed at the central portion of the cell. A prominent nucleolus is always located at the centre of the nucleus. The chromosomes are composed of a bundle of fine filaments embedded in electron-dense material (figure 3). By observations of serial sections, it was confirmed that most of the chromosomes are attached to the
nuclear envelope by one or both ends (as in figure 3, arrows). Besides chromosomes, many electron-dense granules are scattered in the nucleoplasm (arrowheads). The cytoplasm is divided into two parts, the inner cytoplasm with a complicated reticular and vesicular system, and the peripheral thin cytoplasm. A layer of large vacuoles separates these two parts. In the inner cytoplasm, mitochondria with characteristic tubular cristae, trychosysts, and phagosomes of various size are observed (figure 2). In the peripheral cytoplasm (figure 4), a thin layer between flattened vesicles (asterisks) and the plasma membrane corresponds to the theca (between arrowheads). Bundles of microtubules (arrows) are observed just underneath. Each bundle is
Figure 2. A longitudinally sectioned interphase cell observed by TEM in a low magnification. m: mitochondria, no: nucleolus; ph: phagosomes; t: trychocyst; v: cortical vacuole. Bar: 2 µm. Figure 3. Enlargement of chromosomes in an interphase cell. Chromosomes are composed of fine filaments and closely apposed to the nuclear envelope with their ends (arrows). Arrowhead: electron-dense granule. Bar: 1µm. Figure 4. Structure of the lateral cell cortex. Bundles of microtubules (arrows) are observed just beneath flattened vesicles (asterisks). The layer between black arrowheads is the theca. m: mitochondrion; v: cortical vacuole. Bar: 0.2 µm. Figure 5. Two sets of basal bodies and appendages in a cell during prophase. Numbers at lower right corner indicate section number in serial sections cut vertically in the antero–posterior direction. Two sets are about 1 µm apart along the longitudinal axis of the cell. lb: longitudinal basal body; tb: transverse basal body; lmr: longitudinal microtubular root; ac: accessory centriole. Bar; 0.5 µm. Figure 6. A part of nucleus at early prophase. Note deposition of electron-dense material at some of nuclear pores (arrows). Bar: 0.5 µm. Figure 7. Localization of γ-tubulin at the periphery of a nucleus in prophase. Whole cells were stained with monoclonal anti-γ-tubulin (G19) by the method described in materials and methods. Image was taken with an ordinary epifluorescence microscope.
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Figure 8. Three dimensional organization of cortical and flagellar microtubules stained with D66 mAb in interphase cell. Images are reconstructed on a workstation from images obtained with a confocal laser microscope. a. Ventral surface view. b. Dorsal view; (a) and (b) are obtained from one specimen. Arrows indicate microtubular ridge of the girdle. c. Apical view. d. Bottom view. e. Microtubular desmose, which connects the nucleus and the flagellar base (arrow). Bars: 5 µm (a–d); 1 µm (e).
composed of 2–8 microtubules that run evenly spaced and parallel to each other, and represent the so-called cortical microtubules. Confocal images of cortical microtubules stained with D66 are shown in figure 8. The cell cortex contains many microtubular bundles of different thickness, probably a reflection of the number of microtubules contained in each bundle (figure 8a, ventral view and 8b, dorsal view of the same specimen). All cortical microtubule bundles in the epicone (anterior of the cell) converge at one focal point at the cell apex. The distal ends of these microtubules in the ventral and lateral halves join the microtubular rim, which represents the ridge of the girdle (arrows), while dorsal microtubules join the dorsal lip of the girdle, the posterior tip of the cell. A view from the top is shown in figure 8c. From the medial part of the cortex, microtubule bundles gradually fuse together and converge in one small focal area like the spokes of a wheel, so Cell division of O. marina
that the anterior cortex is covered by an even density of microtubules. At the cortex of the hypocone, bundles of microtubules, which are considerably thicker than that of epicone, start from the flagellar base and extend posteriorly to the dorsal lip of the girdle (figure 8d). They converge to the focal area wider than the focal point of the apical tip. The basal circumference of the tentacle is covered by a thick layer of microtubules, producing a brighter fluorescence. A thin microtubular desmose connects the flagellar base with the periphery of the nucleus (figure 8e, arrow).
3.2. Prophase The duplication of the flagellar apparatus is a landmark of early prophase. The overall appearance of the cell is the same as that at interphase except for the duplicated flagellar apparatus (figure 1b). New axonemes elongate along parent ones and then separate Kato et al.
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into two flagella. Longitudinal flagella seem to separate earlier than transverse ones do. At the base of these flagella, two pairs of basal bodies are present, which must have been duplicated at an earlier stage. Figure 5 shows two adjacent flagellar apparatus in a single serial section viewed from the anterior side. These apparati are separated by about 1 µm calculated from the number of sections. Two basal bodies baring flagella are arranged perpendicular to each other in close vicinity. From the wall of the longitudinal basal body, sheets of microtubules (longitudinal microtubular root; lmr) extend toward the surface of the hypocone. This appendage structure has the same structure and orientation in the two duplicated sets of flagellar apparatus. The lmr of posterior flagellar apparatus (lmr2) overlies that of anterior flagellar apparatus (lmr1). Other flagellar appendages have been also duplicated exactly in the same manner. Flagellar axonemes have been dissected, a very frequent phenomenon in this species, probably because of unsuitable fixation. In addition, two short accessory centrioles are observed near these basal bodies in this specimen, presumably representing duplicating basal bodies in the preparation of the next cell cycle. At this stage, cortical microtubule bundles do not change their interphase arrangement. The nucleus itself is still spherical (figure 6) with a nucleolus located at its centre. Multiple depositions of electron-dense material about 100 nm wide become apparent in the nuclear envelope (arrows). Due to their size and location, these densities are likely to be accumulated at some of the nuclear pores. Figure 7 shows localization of γ-tubulin at the periphery of the nucleus at this stage. Bright spots probably represent the densities identified by electron microscopy in nuclear pores. The confocal image of a nucleus shortly after this stage is shown in figure 12a. In the nucleus, many radially arranged microtubules appear with one end at the nuclear envelope. The desmose (arrow) present in interphase still connects basal bodies to the periphery of the nucleus.
3.3. Metaphase Cells gradually elongate in the antero–posterior direction and take oblong shapes. Duplicated flagellar apparatus separate and move in opposite directions along the longitudinal axis. Concomitant with the separation of the flagellar apparatus, tentacles are also duplicated and separated in the same way (figure 1c). At this stage, the nucleus is flattened along the longitudinal axis and takes on a discoid shape (figure 12b). From nuclear pores of flattened faces of the nuclear envelope, bundles composed of 6–9 microtubules run to the opposite side of the nucleus through chromosome clusters (figure 9). Each microtubule grows more than 3/4 of the short axis of the disk, but do not attach Cell division of O. marina
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to the opposite side of the nuclear envelope. Microtubule bundles originating from opposite faces of the disc-shaped nucleus seem to be paired, in other words, two densities in nuclear pores facing each other in opposite parts of the nuclear membrane become paired and organize two presumably antiparallel microtubule bundles. These constitute a kind of elementary mitotic spindle. Chromosomes are also arranged longitudinally along the short axis of the discoid nucleus although no equatorial plate can be recognized. Connections between chromosomes and microtubules, through kinetochores or some other special structures, are not recognized. The close relation between chromosomes and nuclear envelope that is observed in interphase nucleus has disappeared at this stage. A sagittal section double-stained with DM1A and DAPI (figure 13a) shows a parallel array of microtubule bundles in a flattened nucleus in which chromosomes are arranged in a plate. A transverse section of the nucleus at this stage shows that the number of microtubule bundles is over 70 (figure 13b).
3.4. Ana–telophase At ana–telophase, the flagellar apparatus and the hypocones separate further along the main cell axis. The cell body gradually elongates and then begins to be constricted at the circumference of its meridian plane between the anterior and the posterior parts (figure 1d). The microtubular cytoskeleton of early ana–telophase cells is reconstructed from confocal images (figure 12c, d; the same specimen). Two flagellar apparatus are separated by several microns (figure 12d, arrows). In the equatorial cortex, microtubule bundles appear to be bisected all around the cell surface (figure 12c, arrow). Probably at the same position of the cortex, a band of actin (contractile ring) is detected by immunostaining with anti-actin antibody mAbC4 (figure 13d, arrows). The presence of actin is observed on the nuclear envelope at the same time (arrowheads). In the nucleus, chromosomes separate into two plates, which indicates the beginning of ana–telophase (figure 13c, d). Many parallel microtubule bundles that run across the nucleus along its short axis are evident at this stage. Each bundle seems to be almost continuous across the nucleus. At this time, elongation and/or sliding of microtubule bundles must occur, which causes elongation of the nucleus along the antero–posterior axis and simultaneous separation of the chromosomes. The nucleolus takes an oblong shape with both ends apposed to the nuclear envelope (figure 13c, arrow). Then the equator of the disc-shaped nucleus becomes narrower and takes on a dumbbell shape (figure 10 and figure 12d). At the constricted area of the nucleus, microtubules are densely packed crossing each other in a close vicinity (figure 11), which proKato et al.
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Figure 9. View of a part of a nucleus at metaphase. Note microtubules emanating from a dense deposition of nuclear pore (arrows). Bar: 0.5 µm. Figure 10. A dumbbell-shaped nucleus at ana–telophase. An elongated nucleolus is at the centre of the nucleus (no). Bar: 1 µm. Figure 11. Microtubules at the constricted portion of an ana–telophase nucleus. Arrowheads indicate the limit of nuclear envelope. Bar: 0.2 µm.
duces quite a strong fluorescence signal in that area (figure 12d). Each microtubule runs rather straight than curved, suggesting their twisting around the longitudinal axis. At late ana–telophase, the constriction of the cell body proceeds. During constriction and elongation of the cell body, the future anterior daughter cell has an incomplete posterior part of the cytoskeleton and vice versa for the future posterior daughter cell (figure 12e). This might indicate that the posterior part of the anterior cell and that the anterior part of the posterior cell are rebuilt de novo during and after ana–telophase. The nucleus has already divided and positioned at the centre of the future daughter cells at this stage (figure 12f, the same specimen as figure 12e). Intranuclear microtubules remained until that time and then disappear. Considerably before the completion of cytokinesis, two daughter nuclei are formed (figure 13e), then the Cell division of O. marina
actin ring disappears, leaving a broad cytoplasmic connection between the two daughter cells (figure 13f, between two arrows). After completion of cytokinesis, the two daughter cells remain connected by a thin cytoplasmic bridge, which persists for more than 15 min. Occasionally, the next division cycle progresses before the final separation of the two daughter cells.
4. DISCUSSION In some dinoflagellates, new mitotic microtubules appear in the cytoplasm at the time of nuclear division, while the nuclear envelope persists and mitosis is intranuclear (Spector and Triemer, 1981). From an evolutionary point of view, this represents an intermediate process of mitosis between yeast and higher eukaryotes (Alberts et al., 1994). We confirm earlier reports showing that in O. marina, the microtubular mitotic Kato et al.
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Figure 12. Three dimensional organization of microtubular systems in dividing cells. a. Intranuclear microtubules in a spherical nucleus at prophase. The arrow indicates the microtubular desmose between the nucleus and the flagellar base. b. Intranuclear microtubules of a metaphase cell. Bundles of microtubules are arranged in parallel to the short axis of a disc-shaped nucleus. c. Cortical microtubules of an early ana–telophase cell. Microtubular bundles are separated at the cell body equator (arrow). d. Intranuclear microtubules in the same cell as (c). Arrows and arrowheads indicate flagellar base and microtubular desmose respectively. e. Ventro–lateral view of cortical microtubules and flagellar apparatus of an ana–telophase cell. f. Intranuclear microtubules in the same cell as (e). They appear to have been separated into two daughter nuclei. Bar: 5 µm.
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Figure 13. Arrangement of microtubules and localization of actin in semi-thin sections. Sections were stained with an anti-α-tubulin mAb (DM1A) and DAPI (a–c), or an anti-actin mAb (C4) and DAPI (d–f). Black and white images were incorporated in a personal computer, artificially colored and then overlapped. The brightness of FITC signals were intensified in (d–f) in order to see the boundary of cytoplasm. a. A longitudinal section of a metaphse cell. Chromosomes (blue) and microtubules (yellow) are overlapped in the disc-shaped nucleus. b. A transverse section of a metaphase cell. c. A longitudinal section of an early ana–telophase cell. Chromosomes are separated into two planes. Arrow indicates the elongated nucleolus. d. An early ana–telophase cell. Bright yellow spots (arrows) indicate actin bundle at cell cortex. The nuclear envelope is also stained by anti-actin antibody (arrowheads). e. An ana–telophase cell. The contractile ring (arrows) at the constricted cell cortex and actins around the nuclear envelope are apparent. f. A late ana–telophase cell. Two daughter cells are still connected by broad cytoplasm (between two arrows), while the actin ring has already disappeared. Bar: 5 µm.
apparatus is intranuclear (Triemer, 1982; Gao and Li, 1986; Kato et al., 1997). Such an intranuclear microtubular division system is exceptional among dinoflagellates as described to date. In many ciliates, intranuclear microtubules are frequently observed. In Paramecium for instance, microtubules appear both in the macro- and the micronucleus during vegetative division, and in the micronucleus during mitosis (Ishida et al., 1999). In this regard, O. marina is more similar to ciliates than other dinoflagellates. Since the nuclear envelope does not break down during nuclear division, translocation of tubulin molecules from the cytoplasm to the nucleus is a prerequisite to allow microtubule polymerization in the nucleus. Thus, it appears that this organism can segregate tubulin molecules into a cytoplasmic and an intranuclear pool devoted to different functions. Whether these tubulin pools have the same isotype and isoform composition remains to be established, since several isotypes have been reported (Laferrière et al., 1997; Huitorel et al.; 1999). Furthermore, the presCell division of O. marina
ence of a MTOC is a prerequisite to microtubule polymerization in yeast and animal cells. In the present paper, we describe the mitotic microtubule nucleation sites inserted at several loci of the nuclear envelope (figure 7 and figure 9). Their structure is comparable to the spindle pole body of budding yeast (Donaldson and Kilmartin, 1996) and that of fission yeast (Ding et al., 1997; Hagen and Yanagida, 1997) especially in terms of the appearance and the size. The number of microtubules nucleated per site is also similar. However, yeast spindle pole bodies are present throughout the cell cycle, either embedded in the nuclear envelope or at its outer surface, and simultaneously nucleates microtubules on both sides of the nuclear envelope at mitosis. We could not detect any cytoplasmic microtubules nucleated from the density in the nuclear pore. There is a possibility that this might be due to the labile nature of cytoplasmic microtubules in this species. On the other hand, it is clear that the density appears only at the beginning of mitosis. Despite the apparent resemblance, these structures may be different in moKato et al.
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lecular composition and in their mechanisms of regulation. Interestingly, plant cells do not have such a well-defined MTOC, but the microtubule nucleation material is dispersed around the outer face of the nuclear envelope. In O. marina, the nucleating material may also be dispersed, then well focused during mitosis. In euplotes, the presence of γ-tubulin in the nucleus throughout the cell cycle has been reported (Liang et al., 1996). In O. marina, we have detected γ-tubulin on the nuclear envelope using the monoclonal anti γ-tubulin antibody G19 (figure 7). Therefore, the electron-dense material accumulated at nuclear pores might correspond to the structure that contains γ-tubulin, which is a component of MTOCs. Further detailed study using immuno-electron microscopy is awaited. The presence of an actin band at furrowing cell surface reported in this paper confirms the presence of actin previously described in other dinoflagellates (Soyer-Gobillard et al., 1996). The cytokinesis is likely to be mediated by the contraction of this band. On the other hand, the constriction of the nucleus precedes cytokinesis. This means that the constriction of the nucleus cannot be caused by contraction of the cytoplasmic contractile ring. There is a possibility that the intranuclear microtubule system itself produces the physical power to constrict the nucleus and separate the two daughter nuclei. We observed the elongation of intranuclear microtubules during nuclear division. If the elongation and the sliding of microtubules of opposite polarities occur, then the nucleus would inevitably become weaker at its equator, and be constricted and twisted. Our observations shown in figure 11 and figure 12d strongly suggest this possibility. In addition, there is a possibility that actin also participates in this process. The presence of actin on the nuclear envelope at metaphase (figure 13d) may suggest a potential role of actin polymers in the constriction of the nucleus. Therefore, if both microtubules and actin fibres participate in the nuclear division in harmony, the mechanism of nuclear division of O. marina may be comparable to the cell division of higher eukaryotic cells. The polar region of the nucleus is connected to the flagellar apparatus by a thin desmose containing microtubules. However, it seems impossible to restrict the physical force necessary for nuclear elongation to the desmose, because it is connected almost vertically to the polar region. More likely, the desmose may act to define the position of polar regions, and possibly of the nucleus itself. In Chlamydomonas, for instance, a centrin filament-based desmose connects the basal bodies to the nucleus and positions it at the time of mitosis (Salisbury, 1995). In several species of dinoflagellates, chromosomes are connected to nuclear pores and are separated by Cell division of O. marina
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the movement of the nuclear envelope (see Spector, 1984). In those cases, extranuclear microtubules seem to render a shearing force to the nuclear envelope, and provide an orientation to the movement of the nuclear envelope and chromosomes. In dividing cells of O. marina, we did not detect any direct connection between chromosomes and the nuclear envelope or microtubules. The mechanism of chromosome separation is still uncertain in this species and remains a matter for further investigations. The cell division of O. marina is an unequal division; the dividing cell keeps its antero–posterior polarity and the division plate is perpendicular to this axis. However, after completion of cytokinesis, the two daughter cells show equivalent structures. Therefore, during the last stage of cell division, a new posterior half has to be rebuilt along the old anterior half and vice versa for the posterior half, according to yet unknown mechanisms. Basal bodies and flagella have been duplicated at an early stage of prophase. Microtubular and fibrous appendages are also duplicated until this stage. Two sets of flagellar apparatus separate at metaphase as the cell body elongates. It is clear that the new hypocone and tentacle are duplicated and separated along with the separation of flagella. When daughter flagellar apparatus have separated by several microns, a belt-like gap appears at the midst of the cortical microtubule arrays. It is possible that the cortical microtubules are cut at the equatorial area, which leads to the gap formation. Microtubule severing factors, especially active in M phase, have been identified in higher eukaryotes (Shiina et al., 1995). Similar factors might be involved in the gap formation of microtubule arrays in O. marina. Since the position of the gap correlates with the location of the actin ring, the localization of such factors might be actin-dependent. Double staining using anti-actin and anti-tubulin antibodies will help to get better understanding about the process of gap formation. At this time and later, cortical microtubules must be elongated to reconstruct two full cell body structures from both halves. The polymerization of tubulin occurs preferentially at nucleation centres or at pre-existing microtubule ends in vivo. The absolute polarity of microtubules in dinoflagellates and the precise location of cortical microtubule nucleation sites are still unknown. In this study, we also could not detect any special structures at the anterior or posterior focal areas, or at the cell equator at the site and time where the ‘actin belt’ is observed. There is a possibility that a recently cut end may be a preferential elongation site, regardless of its polarity, provided the microtubule is stable enough. If this is the case, microtubules can simply elongate to rebuild a complete cortical network. There is an alternative possibility that cortical microtubules elongate at their plus ends with their original Kato et al.
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polarities. In this case cortical microtubules of one daughter cell would re-grow by simple elongation. Cortical microtubules of the other daughter cell would elongate from some new nucleating site and the preexisting cortical network should gradually disappear. A candidate for new nucleating sites would be the newly formed ridge that separates the epicone and the hypocone. The ridge must be organized by the basal bodies at the time of flagellar duplication and separation. Therefore, one may propose that the basal bodies and their associated structures, may contain all the microtubule nucleating material necessary to build the interphase microtubule networks, i.e. the flagellar apparatus and appendages and cortical microtubules. On the other hand, the cytoskeletal microtubules may not necessarily be continuous throughout cortical microtubular bundles. The longitudinal microtubular band of Tetrahymena is composed of stacks of short and slightly angled microtubules (Ng, 1979). These microtubules could be regenerated by a lateral nucleation mechanism from preexisting microtubules reported in trypanosomes (Sherwin and Gull, 1989). In both cases the polarity of microtubules is preserved. Which model applies to O. marina is a matter for future investigations. Acknowledgments. We wish to warmly thank Pr. H. Hotani, Nagoya University, for his welcome in his laboratory to run the confocal microscopy. We are indebted to Dr. C. Gagnon, McGill University, for the use of the D66 monoclonal antibody. Part of this work was supported by a grant from the ‘Centre national de la recherche scientifique’ (CNRS, France) to P.H. P.H. also acknowledges receipt of the fund of Nagoya City University for invited scholars.
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