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First published online 20 November 2007
doi: 10.1242/jcs.015875
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Research Article |
Department of Biology, Faculty of Science, Yamaguchi University, Yamaguchi 753-8512, Japan
* Author for correspondence (e-mail: yumura{at}yamaguchi-u.ac.jp)
Accepted 9 October 2007
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Summary |
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 Top Summary IntroductionResultsDiscussionMaterials and MethodsReferences |
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Key words: Actin, Cell-substratum adhesion, Cytokinesis, GFP, Myosin
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Introduction |
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TopSummaryIntroductionResultsDiscussionMaterials and MethodsReferences |
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) or silenced by antisense RNA (Knecht and Loomis, 1987). However, these myosin-null cells can still divide on a substratum.
When multinucleated myosin-II-null cells in suspension culture are placed on the substratum, they adhere tightly to the substratum and divide without a preceding ordinary nuclear division. This phenomenon is called traction-mediated cytofission, and the traction force against the substratum generated by the two daughter cells is considered to mediate this type of cytokinesis (Spudich, 1989; Fukui et al., 1990). Interestingly, when myosin-II-null cells are cultured on the substratum, they can divide in a cell-cycle-dependent manner by means of an almost morphologically normal cytokinesis (Gerisch and Weber, 2000). This cell-cycle-dependent cytokinesis is distinguishable from traction-mediated cytofission and is referred to as cytokinesis B (Neujahr et al., 1997; Zang et al., 1997). Uyeda and colleagues (Uyeda et al., 2000) renamed these three modes of cytokinesis as follows: the myosin-II-dependent (contractile-ring-dependent) cytokinesis is cytokinesis A, cell-cycle-dependent and myosin-II-independent cytokinesis is cytokinesis B, and cell-cycle-independent and myosin-II-independent cytokinesis is cytokinesis C. The molecular mechanisms of cytokinesis B and C have not been clarified to date. Probably, these types of cytokinesis are mediated by the traction force generated by the two daughter cells. Recent research has shown similar substratum-dependent and contractile-ring-independent cytokinesis in mammalian cells (Kanada et al., 2005).
Several actin-containing structures have been described in Dictyostelium cells. Filopods, long thin extensions containing actin-bundles; microvilli, short thin extensions containing actin-bundles; and pseudopods or lamellipodia, large extensions containing dense actin-meshworks that are mainly observed at the anterior half of migrating cells and the polar regions of dividing cells (Yumura et al., 1984), which are common to many animal cells. Dictyostelium cells have no large actin bundles inside the cell, such as stress fibers. Actin filaments are localized mainly at the cortex to form an actin-rich cortex. During phagocytosis, the cell forms a phagocytic cup surrounding the solid food, which is formed by an actin mesh. Crown-like structures (De Hostos et al., 1991) are observed mainly in cells drinking fluid medium and sometimes behave as pseudopods. Actin filaments also form clusters at focal adhesions for cell migration (Uchida and Yumura, 2004).
In the present study, to investigate the role of actin in substratum-dependent cytokinesis, the dynamics of actin in myosin-II-null cells expressing GFP-actin were observed by total internal reflection fluorescence (TIRF) microscopy. We found novel dynamic actin structures, which appeared specifically during mitosis. We referred to these novel structures as mitosis-specific dynamic actin structures (MiDASes) and investigated their dynamics and functions. Our results demonstrate that MiDASes are formed beneath the centrosomes presumably as a result of some signals transmitted along astral microtubules and play an important role in cytokinesis as scaffolds for transmitting the traction force to the substratum.
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Results |
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100 nm above the coverslip. Consistent with observations by conventional fluorescence microscopy (Yumura et al., 1984), actin localized to polar pseudopods of dividing cells (arrowheads in Fig. 1A). TIRF microscopy revealed other major structures containing actin (circles in Fig. 1A), which were not observed in interphase cells (Fig. 1B) but were present only in the mitotic cells. These structures are referred to as MiDASes hereafter. A MiDAS emerged as an irregularly shaped and large flat sheet in the middle of the ventral cell cortex when the spindle began to elongate. The MiDAS split in two in accord with the nuclear division and moved to each daughter cell. The MiDASes dynamically changed their shape and size from 2-5 µm in diameter (see also supplementary material Movie 1). The trajectories of two MiDASes during cell division are shown in Fig. 1C. Fig. 1D shows the time-course of the area of MiDASes, showing that they disappeared shortly after the completion of cytokinesis. When the cells were treated with latrunculin A, which depolymerizes actin filaments, MiDASes disappeared within 40 seconds (data not shown). Furthermore, MiDASes stained with tetramethyl rhodamine-phalloidin, which binds specifically to actin filaments, as shown below. Therefore, MiDASes comprise actin filaments. MiDASes were also observed in dividing wild-type cells, although their appearance was very transient (circles in Fig. 1E). We conclude that the MiDAS is a novel mitosis-specific actin-containing structure.
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Astral microtubules are required for the formation and maintenance of MiDASes
Fig. 2A-F shows a series of optical sections by confocal microscopy of fixed multinucleated cells expressing GFP-histone1, which shows the position of nuclei, and stained with tetramethyl rhodamine-phalloidin. Multinucleated cells were prepared in suspension culture as myosin-null cells cannot divide in suspension. Fig. 2H-M shows multinucleated cells expressing GFP–
-tubulin stained with tetramethyl rhodamine-phalloidin. MiDASes were located underneath the nuclei and the centrosomes. The latter two organelles associated with each other. The number of MiDASes was equal to that of nuclei and centrosomes. These observations suggest the possibility that nuclei and/or centrosomes might induce the formation of MiDASes. Centrosomes especially are plausible as inducers because astral microtubules emerging from them can directly reach the ventral cortex to induce formation of MiDASes.
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-tubulin by confocal microscopy (Fig. 3A-C). Consistent with the observation of fixed cells, MiDASes were located underneath the centrosomes and, interestingly, relocated following the movement of centrosomes (Fig. 3B,C). As TIRF microscopy can limit illumination to 100 nm above the coverslip, which corresponds to the thickness of the ventral cortex, including the actin cortex and MiDASes, it is possible to observe the correlation between the distal ends of each microtubule and the MiDASes. Fig. 3D shows typical images of the relationship between the distal ends of microtubules and MiDASes that were captured by an exposure of 100 milliseconds. Fig. 3E shows a sum of traces of microtubules (green) and outlines of MiDAS (red) for 15 seconds, indicating that the distal ends of microtubules reached mainly into the MiDAS regions. These observations strongly suggest that astral microtubules emerging from the centrosomes determine the position of MiDASes, probably by sending signals to the ventral cortex.
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To examine further the possible role of microtubules in the formation of MiDASes, myosin-null cells were exposed to thiabendazole, which specifically disrupts microtubules in a manner similar to that of nocodazole (Kitanishi et al., 1984). Shortly after the addition of thiabendazole, the astral microtubules shortened (Fig. 4A) and MiDASes gradually regressed in size and finally disappeared (Fig. 4B). When thiabendazole was removed by washing with buffer, MiDASes reappeared underneath the centrosomes (arrows in Fig. 4B). These results strongly suggest that the astral microtubules carry some signals to the cortex that are involved in the formation and maintenance of MiDASes.
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MiDASes are required for substratum-dependent cytokinesis
Myosin-null cells fail to complete cytokinesis on a substratum with a frequency of about 10% (Neujahr et al., 1997). In such cases, one half of the dividing cell was resorbed by the other half, probably owing to an imbalance in the motile activities between the two halves. We frequently observed abortive cytokinesis when MiDASes disappeared by accident. When one of the MiDASes disappeared (arrows in Fig. 5A), this half of the cell was finally resorbed by the other. This phenomenon was observed in 21 out of 24 cells examined. The other three cells also showed abortive cytokinesis, leaving only small fragments. Quantitative analysis shows that the area of the MiDAS decreased coincidentally with that of the resorbed half (Fig. 5B).
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MiDASes function as scaffolds for substratum-dependent cytokinesis
Since MiDASes exist in the ventral cell cortex and are necessary for substratum-dependent cytokinesis, they might transmit the traction force generated by the cell to the substratum. If this is the case, the cell membrane underneath MiDASes must be attached to the substratum. To assess this possibility, myosin-null cells were observed by interference reflection microscopy (IRM), which is generally used to detect the adhesion sites of cells. Fig. 6A shows dual images of GFP-actin and IRM, which shows the existence of a darker tone at the MiDAS regions in comparison with the tone at other ventral membrane areas, suggesting that MiDAS regions are closer to the substratum than the other regions of the ventral membrane.
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To assess further whether the dividing cells attach to the substratum at the MiDAS regions, cell bodies were blown away by a jet of buffer from a pipette under confocal microscopy. Fig. 6B,C shows MiDASes before and after blowing away the cell bodies. In every case, only MiDAS regions remained attached to the substratum, although other parts of the cells were detached by the blowing, indicating that dividing cells were strongly attached to the substratum mainly at the MiDAS regions. These observations strongly suggest that MiDASes are scaffolds for substratum-dependent cytokinesis and platforms for the cell to transmit mechanical force to the substratum.
Dynamics of actin in MiDASes
The size and shape of MiDASes were not constant but changed dynamically (see supplementary material Movie 1). To investigate the dynamics of actin in MiDASes, a part of the MiDAS region was photobleached by scanning laser illumination. The fluorescence of the bleached regions recovered immediately (Fig. 7A,B, half time of the recovery, 2.15±0.89 seconds, mean ± s.d., n=17), which indicates that actin in the MiDASes undergoes continuous rapid turnover. Fig. 7C,D shows high magnifications of the fluorescence recovery processes. After photobleaching, fluorescence appeared as individual small dots and finally became an aggregate of these dots.
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; Uchida and Yumura, 2004). Actin foci also appeared and disappeared with a duration similar to that of actin dots in MiDASes. However, as actin foci still exist even after the disappearance of MiDASes induced by the disruption of microtubules, the formation of actin foci does not seem to be regulated by microtubules (Fig. 5C).
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Discussion |
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Because astral microtubules were required for the formation of MiDASes, there might be some signals along the astral microtubules that induce the formation and maintenance of MiDASes. In vertebrate animal cells, CLIP 170, EB1 and APC (collectively termed +Tips), which associate to the plus-ends of microtubules, regulate the actin cytoskeleton at the leading edges (Fukata et al., 2002; Kawasaki et al., 2000; Wen et al., 2004). EB1 has been identified also in Dictyostelium cells and colocalizes with actin (Rehberg and Graf, 2002), but whether DdEB1 induces the expansion of pseudopods is still unknown. In the case of 3T3 fibroblasts, the ruffling is induced by Rac1, a small GTPase that is activated by polymerizing microtubules (Waterman-Storer et al., 1999). As the plus-ends of astral microtubules are concentrated at the polar regions of a dividing cell, they might activate ruffling there (Neujahr et al., 1998). In our observations, MiDASes sometimes fused with actin of pseudopods (data not shown), suggesting that similar signals by the plus-ends of astral microtubules might induce actin polymerization both in pseudopods at the leading edges and in MiDASes in the ventral cell cortex.
As shown in Fig. 8A, small actin-containing dots increased in number and formed a nascent MiDAS in early anaphase. Conversely, when MiDASes disappeared after the completion of cytokinesis, the actin dots decreased in number (Fig. 8B). These observations suggest that MiDASes comprise actin-containing dots. Interestingly, the individual actin dots did not change their positions: they appeared and disappeared at the same positions. These actin dots were indistinguishable from actin foci – the duration of their appearance and their sizes were identical. In a previous study, we showed that actin foci are fixed in position relative to the substratum and are a candidate for the focal adhesions of Dictyostelium during cell migration (Uchida and Yumura, 2004). It would seem that MiDASes changed their position so as to coordinate with the position of the centrosomes (Figs 1 and 3). The change in the position of MiDASes can be explained by this rapid turnover of actin dots; their appearance in the front leading edge of the MiDAS and their disappearance in the rear result in a gradual change in the location of the MiDAS as a whole. A recent TIRF microscopy study revealed mobile actin clusters, the sizes of which were similar to actin foci, during the reorganization of actin networks after they were depolymerized by latrunculin A and after drug removal (Gerisch et al., 2004). The actin foci and the actin dots in the MiDASes seem to be different to actin clusters because the former are stationary with respect to the substratum.
Do wild-type Dictyostelium cells perform cytokinesis by dual mechanisms, both myosin-II-dependent and -independent? MiDASes appear transiently, even in wild-type cells (Fig. 1E). When wild-type cells divide on a substratum, most of them round up, suggesting that most of the contact sites are detached from the substratum. However, some of the population of cells divides when adhering strongly to the substratum, maintaining their flat morphology. In this case, the cellular shape during mitosis is similar to that of myosin-II-null cells (Neujahr et al., 1997; Nagasaki et al., 2001). Probably, wild-type cells divide through both myosin-II-dependent and -independent mechanisms. It is also plausible that the appearance of MiDASes might be a result of adaptation in myosin-null cells, which is transitory and less important in wild-type cells because they can divide in the absence of a substratum. Myosin-II-independent cell division has also been observed in mammalian cells. Epitheloid kidney cells injected with antibodies against myosin to diminish the levels of myosin II in the equatorial region can still divide (Zurek et al., 1990). Normal rat kidney cells can divide in the presence of blebbistatin, a potent inhibitor of the nonmuscle myosin II ATPase, in a manner similar to that of Dictyostelium myosin-null cells (Kanada et al., 2005). Normal kidney cells and 3T3 fibroblasts that were microinjected with C3 ribosyltransferase, an inhibitor of Rho, could not accumulate myosin at the cleavage furrow but still divided (O'Connell et al., 1999). This cleavage is considered cell-substratum dependent because HeLa cells, which detach from the substratum during cell division, cannot divide when injected with C3 enzyme. The same authors described how, when fluorescent beads were attached to the surface of C3-injected normal rat kidney cells, they moved towards the chromosomes, and the resultant cluster of beads followed the movement of chromosomes and split in two as if they were tethered to the chromosomes. At that time, clusters of cortical F-actin were localized under the chromosomes, which is reminiscent of MiDASes (O'Connell et al., 1999). More recently, Guha and colleagues (Guha et al., 2005) showed that large actin structures similar to MiDASes appeared in dividing normal rat kidney cells after local treatment with the myosin II inhibitor blebbistatin. It should be noted that MiDASes can be observed even in myosin-null Dictyostelium cells expressing motorless myosin II (G.I and S.Y., unpublished). Therefore, the loss of myosin ATPase activities seems to induce or enhance the emergence of MiDASes.
Myosin-II-independent and adhesion-dependent cell division is probably an ancient mechanism that emerged before the evolution of myosin and is still available in various cells (Uyeda and Nagasaki, 2004). Therefore, further investigation of this mechanism is required for a complete understanding of the mechanism of cytokinesis. In future studies, it will be important to clarify the putative signals transmitted along the astral microtubules that regulate the formation and maintenance of MiDASes.
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Materials and Methods |
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-tubulin and GFP-histone1 were transformed into cells by electroporation (Yumura et al., 1995). Briefly, cells that grew to confluence in plastic dishes were washed twice with 15 mM Na/K phosphate buffer, and they were centrifuged at 350 g for 90 seconds at 4°C. After centrifugation, an equal volume of the phosphate buffer was added to the pellet, which was then suspended by mild tapping. 30 µl of the cell suspension and 3 µl of plasmid DNA solution were mixed and charged at 380 V, which was repeated twice. After electroporation, the solution in the cuvette was suspended in 10 ml of HL5 medium including 40 µg/ml of streptomycin in a plastic Petri dish. After 12 hours, 10 µg/ml of G418 (Sigma) and/or blasticidin (Kaken) was added for selection.
Treatment with thiabendazole
HS1 cells null for the myosin II heavy chain and expressing GFP-actin or GFP–-tubulin that grew in HL5 medium were washed with BSS (10 mM NaCl2, 10 mM KCl, 3 mM CaCl2, 2 mM MES, pH 6.3). After incubation for 3 hours, cells were placed in a chamber that was made by attaching a silicon sheet with a hole (10 mm in diameter) to a 24x50 mm coverslip. An equal amount of thiabendazole (2x10–4 M in BSS) was added to the chamber. HS1 cells took 4-7 minutes to divide. Microtubules disappeared within 30 seconds, as shown in Fig. 4A. Next, thiabendazole was removed by replacing it with BSS. The cells were observed by confocal microscopy (LSM510, Carl Zeiss).
Fixation and staining
HS1 cells expressing GFP-histone1 or GFP–-tubulin that grew in HL5 medium were suspended and then placed on an 18x18 mm coverslip. After 20 minutes, fixation was performed with ethanol containing 1% (w/v) formaldehyde at –17°C for 5 minutes. After washing three times with PBS (135 mM NaCl2, 2.7 mM KCl, 1.5 mM KH2PO4, 80 mM NaH2PO4, pH 7.3) at intervals of 5 minutes, cells were incubated with tetramethyl rhodamine (TRITC)-phalloidin for 30 minutes and then washed three times with PBS at intervals of 5 minutes. After mounting the samples in mounting medium [10% polyvinyl alcohol in PBS containing 50% (v/v) glycerol], the cells were observed by confocal microscopy.
Confocal microscopy
Fluorescence images of HS1 cells expressing GFP-actin or GFP–-tubulin were obtained with the confocal microscope system, a LSM 510 microscope (Carl Zeiss) equipped with a 100x Plan Neofluar objective (NA 1.3). For excitation of GFP and TRITC, an argon laser (488 nm line) and HeNe laser (543 nm line) were used, respectively. Time-lapse microscopy of HS1 cells expressing GFP-actin was preformed at an optical section of 1.5 µm, with focusing on the ventral surface of the cell. To acquire sequential fluorescence images from bottom to top, confocal sections were scanned up at distances of 0.3 µm in the Z-axis. IRM images and fluorescence images were acquired simultaneously, as described previously (Uchida and Yumura, 2004). Photobleaching was performed as described previously (Yumura, 2001). Measurement of fluorescence intensity was performed by Scion software (Scion Corporation). To perform an assay blowing away the cell bodies, the cell bodies were blown away by a jet of the buffer from a pipette while under confocal microscopy.
TIRF microscopy
A TIRF microscopy system (based on I X70, Olympus) was constructed as described previously (Tokunaga et al., 1997). The fluorescence of the cells expressing GFP-actin was observed with the TIRF microscope equipped with a PlanApo oil objective (60x, NA 1.45, Olympus). The wild-type cells were slightly compressed by an agar sheet to observe the ventral surface, as described previously (Yumura et al., 1984). Illumination was performed by an argon laser (40 mW, the National Laser) and a HeNe laser (2 mW, Research Electro Optics). The fluorescence images were captured by a digital cooled CCD camera (C4742-95-12 ER, Hamamatsu) and digitized with IP lab software for MS Windows (Scanalytics).
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Acknowledgments |
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-tubulin, GFP-histone1 and mCherry-actin. We also thank T. Kitanishi-Yumura for helpful discussions. A part of this work was supported by Grants-in-aid for Scientific Research in Priority Areas from the Japan MEXT. |
Footnotes |
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References |
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