TY - JOUR
AU - Hasezawa, Seiichiro
AB - Abstract The roles of actin microfilaments (MFs) in the organization of microtubules (MTs) at the M/G1 interface were investigated in transgenic tobacco BY-2 cells stably expressing a GFP-tubulin fusion protein, using the MF-disrupting agent, Bistheonellide A (BA). When MFs were disrupted by BA treatment, cortical MTs (CMTs) did not become reorganized even 3 h after phragmoplast collapse, whereas non-treated cells completed CMT reorganization within 1 h. Furthermore, in the absence of MFs, the tubulin proteins did not show appropriate recruitment but remained at the site where the phragmoplast had existed, or extra-phragmoplasts were organized. These extra-phragmoplasts could functionally form extra-cell plates. This is the first observation of the formation of multiple cell plates during one nuclear division, and of phragmoplast generation irrespective of the position of the mitotic spindle or nuclei. The significance of these observations on the role of MFs at the M/G1 interface is discussed. (Received January 25, 2004; Accepted April 5, 2004) Introduction Higher plant cells possess four characteristic microtubule (MT) structures: cortical MTs (CMTs) that regulate the direction of cellulose-microfibril deposition; the preprophase band (PPB) that predicts the future division site; the mitotic spindle that segregates the chromosomes; and the phragmoplast that forms the cell plate. A phragmoplast consists of two “anti-parallel” sets of MT arrays on both sides of a mid-zone. In normal vegetative cells, the phragmoplast is initiated from the interzonal array of MTs between segregated sister chromosomes in anaphase/telophase, and spreads as a ring-like structure that surrounds the leading edge of the cell plate (Verma 2001). Cell plate formation begins with vesicle fusion, following callose and cellulose deposition (Samuels et al. 1995). The vesicles are thought to be transported along phragmoplast MTs (Palevitz and Hepler 1974, Kakimoto and Shibaoka 1988), probably by plus end-directed kinesin-like motor proteins (Lee et al. 2001, Nishihama et al. 2002). As well as MTs, actin microfilaments (MFs) are also aligned on the phragmoplast, and form a ring-like structure that surrounds the cell plate (Kakimoto and Shibaoka 1987, Kakimoto and Shibaoka 1988, Cleary et al. 1992, Hasezawa et al. 1998). Although some reports have argued for the involvement of MFs in vesicle transport (Kakimoto and Shibaoka 1988) or in guidance of the cell plate to the correct division site (Kakimoto and Shibaoka 1987, Valster and Hepler 1997, Palevitz and Hepler 1974, Mineyuki and Palevitz 1990), MFs on the phragmoplast appear not to be essential for cell plate formation, since the cell plate can enlarge and join the parental cell wall and cytokinesis can also be completed even when the MFs are disrupted by actin polymerizing inhibitors. Therefore, the functions of MFs on the phragmoplast are still unclear. The phragmoplast becomes organized in anaphase/telophase, and collapses at the M/G1 interface. Accompanying phragmoplast collapse, free tubulin protein, released from the phragmoplast, is first translocated to and accumulates on the surface of the daughter nuclei (Yoneda and Hasezawa 2003), followed by MT elongation from the nuclear surface to the cell cortex (Hasezawa et al. 2000, Granger and Cyr 2000, Yoneda and Hasezawa 2003). When the MTs reach the cell cortex, bright spots appear on the cell cortex, from where nascent CMTs parallel to the cell longitudinal axis become elongated, followed by transverse CMT reorganization (Kumagai et al. 2001, Yoneda et al. 2003, Yoneda and Hasezawa 2003). Although these processes from phragmoplast collapse to CMT reorganization have been clarified, the regulatory mechanisms of the process are still unclear. In this study, we have focused on the roles of MFs and, in particular, on the structural changes of MTs at the M/G1 interface in the absence of MFs. BY-GT16 cells, transgenic tobacco BY-2 suspension cultured cells stably expressing a GFP-α-tubulin fusion protein, were treated by the actin polymerization inhibitor, Bistheonellide A (BA), and then time-sequential observations by deconvolution microscopy or confocal laser scanning microscopy (CLSM) were performed. When the MFs were disrupted by BA treatment, the cells showed no signs of CMT reorganization. Although in about 80% of the BA-treated cells the tubulin did not move and remained at the site where the phragmoplast had previously existed, we found that in about 20% of BA-treated cells the tubulin moved irregularly and formed phragmoplast-like structures of MTs in the outer equatorial regions after the original phragmoplast had collapsed. These extra-phragmoplasts had a mono-layered appearance, that is a half of the “anti-parallel” MT set of normal phragmoplasts, but were functionally able to form the extra-cell plates. Furthermore, γ-tubulin, a component of the microtubule-organizing center (MTOC), was also localized on the extra-phragmoplast instead of on the cell cortex. Based on these data, we discuss the role of MFs in CMT reorganization and phragmoplast formation. Results Localization of MTs and MFs from metaphase to early G1 phase To investigate the precise localization of MTs and MFs, the BY-GT16 cells were stained with rhodamine-phalloidin and 4′,6-diamidino-2-phenylindole (DAPI), and observed by deconvolution microscopy (Fig. 1). At metaphase, when MTs formed the mitotic spindle (Fig. 1A, a-1), the MFs were not localized around the spindle and displayed the actin-depleted zone (ADZ; Fig. 1A, a-2, bracket) that is thought to participate in demarcation of the future division site (Cleary 1995, Hoshino et al. 2003). At early telophase, when the phragmoplast developed, the MFs became co-localized at that site as described previously (Fig. 1A, b, arrows, Hasezawa et al. 1998). As the phragmoplast was gradually collapsing at late telophase, the MFs that were co-localized in the phragmoplast simultaneously began to disappear (Fig. 1A, c, arrows). At the early G1 phase, at about which time the CMTs became reorganized, the MFs no longer existed at the division site, but ran parallel to the cell longitudinal axis (Fig. 1A, d-2). To investigate the roles of MFs during cell cycle progression from the M phase to G1 phase, we disrupted the MFs in this period. BA, a dimeric macrolide that inhibits actin polymerization, caused rapid MF disruption within 30 min after application at a final concentration of 1 µM (Hoshino et al. 2003). Although the MFs were completely disrupted by BA treatment at anaphase/telophase (Fig. 1B, b-2), the arrays of phragmoplast MTs showed no disarrangement compared with normal untreated cells (Fig. 1B, a-1, b-1). Even if BA was added at metaphase, when the phragmoplast had not yet become organized, the phragmoplast could still be formed at anaphase/telophase (data not shown). Effect of BA on CMT reorganization To examine the roles of MFs on CMT reorganization at the M/G1 interface, the cell cortex was time-sequentially observed by CLSM (Fig. 2, cortex). In control cells, the CMTs were completely reorganized within 1 h after the collapse of the phragmoplast (Fig. 2, a, cortex). However, in BA-treated cells, the CMTs did not appear even by 3 h after the disappearance of the phragmoplast (Fig. 2, b, c, cortex). To investigate the reason why CMTs were not reorganized, the mid-plane of the cells were simultaneously observed (Fig. 2, mid-plane). In the control cell, MTs elongated from the nuclear surface to the cell cortex, after which the nucleus gradually moved away from the division plane (Fig. 2, a, mid-plane). In contrast, in about 80% of the BA-treated cells, the fluorescence of the GFP-tubulin showed no movement and remained near the division plane (Fig. 2, b, mid-plane). Nevertheless, in about 20% of these cells, phragmoplast-like structures could still become organized (Fig. 2, c, mid-plane, arrow) after collapse of the original phragmoplast (Fig. 2, c, mid-plane, arrowheads). Time-sequential observations of extra-phragmoplast formation To investigate how such extra-phragmoplasts became organized, we performed detailed time-sequential observations on the mid-plane of the BY-GT16 cells by CLSM. In control cells, the free tubulin that was released from the collapsing phragmoplast became translocated to and accumulated on the nuclear surface (Fig. 3, Control, 9–18 min), with subsequent MT elongation from the nuclear surface to the cell cortex (Fig. 3, Control, 27 min, arrow) as reported previously (Kumagai et al. 2001, Yoneda et al. 2003). However, in some BA-treated cells, some short MTs appeared at the final stage of collapse of the original phragmoplast (Fig. 3, BA, 42 min, arrow). These MTs did not reach the cell cortex, but instead gradually increased in number (Fig. 3, BA, 60 min, arrow) until they finally formed a phragmoplast-like structure (Fig. 3, BA, 114 min). Characterization of extra-phragmoplast To examine the features of the extra-phragmoplasts, the cells were observed by CLSM in 1 µm steps, and 3-D images were digitally reconstructed from these data (Fig. 4A). Front view (Fig. 4A, a-1, b-1) and rotated view (Fig. 4A, a-2, b-2) images were then examined. In a control cell, the phragmoplast appeared as a well-ordered circular structure, with the mid-region at the site of cell plate formation being depleted of GFP-fluorescence between the bi-layer MTs of the phragmoplast (Fig. 4A, a-1, a-2). The original phragmoplasts of BA-treated cells were indistinguishable from those of control cells (data not shown). At phragmoplast collapse in control cells, the tubulin proteins were translocated to and accumulated at the nuclear surface, from where the nascent MTs elongated, followed by CMT reorganization on the cell cortex (Kumagai et al. 2001, Yoneda et al. 2003). However, after collapse of the original phragmoplast, some BA-treated cells were found to have organized extra-phragmoplasts (Fig. 4A, b). These extra-phragmoplasts were organized independently from, and invariably after collapse of, the original phragmoplasts. Furthermore, the extra-phragmoplasts somehow possessed a disordered MT array, and seemed conspicuously “mono-layered”: the fluorescence-depleted zone was not observed, and appeared as one side of a normal phragmoplast (Fig. 4A, b). To examine whether the extra-phragmoplasts could function in forming cell plates, the BY-GT16 cells were stained with 0.05% aniline blue solution and photographed by deconvolution microscopy. In control cells, the cell plates were observed in the mid-zone of the phragmoplasts (Fig. 4B, a), and they showed centrifugal growth. In BA-treated cells, the extra-phragmoplasts formed extra-cell plates (Fig. 4B, b, arrows). As a result, three independent cell plates, one formed by the original phragmoplast (Fig. 4B, b-2, arrowheads) and the others formed by the extra-phragmoplasts (Fig. 4B, b-2, arrow), could be simultaneously observed. In this case, two extra-phragmoplasts were organized in the two daughter cells; however, in some cases, only one extra-phragmoplast appeared in one daughter cell and resulted in the appearance of only two cell plates (data not shown). In all cases, these extra-cell plates were distorted and incomplete compared to the original ones that had relatively flat and smooth structures (Fig. 4B, a-2, b-2). Localization of γ-tubulin Since MF disruption caused not only CMT reorganization but also extra-phragmoplast formation, it could conceivably also have affected components of the MTOC. To examine this possibility, we stained BY-GT16 cells using an anti-γ-tubulin antibody, and observed the localization of γ-tubulin by CLSM. At telophase, γ-tubulin was localized in the phragmoplast, especially on both outer edges of the phragmoplast MTs (Fig. 5, a). After phragmoplast collapse, the tubulin translocated to and accumulated on the nuclear surface, first to the proximal sides of the division site, and then to the distal sides where the translocation of γ-tubulin preceded that of α-tubulin (Fig. 5, b). Later, the γ-tubulin moved to the cell cortex along the MTs that elongated from the nuclear surface (Fig. 5, c). When the extra-phragmoplasts became organized in BA-treated cells, the γ-tubulin also localized to the extra-phragmoplast (Fig. 5, d, arrows), but did not reach the cell cortex. Discussion Dynamics of MTs and MFs at the M/G1 interface In higher plant cells, the MTs and MFs display dynamic structural changes during cell cycle progression, and play critical roles in multiple aspects of cellular events. Especially in cytokinesis, the phragmoplast acts to form the cell plate. When that phragmoplasts are organized at the interface between anaphase and telophase, the MFs co-localize with the phragmoplast MTs (Fig. 1A, Kakimoto and Shibaoka 1987, Kakimoto and Shibaoka 1988, Cleary et al. 1992, Hasezawa et al. 1998). After phragmoplast collapse at late telophase, the MFs that had co-localized at this site also disappear (Fig. 1A, c). Subsequently, the MFs become localized on the cell cortex, and the CMTs reorganize following the predeveloped cortical MFs (Hasezawa et al. 1998, Kumagai and Hasezawa 2001). Despite the obvious involvement of MFs in these events, their precise roles in phragmoplast organization and CMT reorganization are still unclear. To identify the roles of MFs throughout this period, we have examined the effects of MF disruption in synchronized BY-GT16 cells using an actin polymerization inhibitor, BA (Saito et al. 1998, Hoshino et al. 2003). Roles of MFs on CMT reorganization In this study, CMTs did not become reorganized at the M/G1 interface after disruption of MFs by BA treatment. Although CMTs were reorganized within 1 h after phragmoplast collapse in normal cells (Fig. 2, a), the CMTs were not reorganized in BA-treated cells even 3 h after disappearance of the original phragmoplast (Fig. 2, b, c). This suggests that MFs are indispensable for CMT reorganization. In about 80% of BA-treated cells, the tubulin remained at the site where the phragmoplast had existed (Fig. 2, b), suggesting that MFs are required for tubulin translocation from the collapsing phragmoplast to the nuclear surface. In addition, in about 20% of the MF-disrupted cells, the nascent MTs were nucleated from the nuclear surface, but they did not reach the cell cortex, and formed the extra-phragmoplasts (Fig. 2, c, 3, BA). This suggests that MFs are also required for MT elongation from the nuclear surface of the daughter nucleus to the cell cortex. The cortical MFs may act as a “guide” or a “target” of MT elongation towards the cell cortex. Furthermore, it has been reported that after the MTs reach the cell cortex, the CMTs elongate following the predeveloped MFs on the cell cortex (Hasezawa et al. 1998, Kumagai and Hasezawa 2001), implying that cortical MFs guide the direction of the organizing CMTs. The CMTs were not reorganized when the MFs were disrupted, probably because tubulin released from the collapsing phragmoplast was not translocated to the cell cortex in the absence of MFs, thus resulting in the lack of the source for CMT nucleation. These findings are in support of our previous hypothesis of tubulin recycling at the M/G1 interface (Yoneda and Hasezawa 2003). Because of the low level of gene expression at M phase, tubulin recycling may be important in the rapid reorganization of CMTs and in the directional determination of cell elongation. The MFs may therefore play important roles in tubulin recruitment and CMT reorganization at the M/G1 interface. MT nucleation on the nuclear surface In this study, the MTs were nucleated from the nuclear surface, even in BA-treated cells, when the tubulin proteins relocated there (Fig. 3, BA). This suggests that the nuclear surface acts as a MTOC, and that this MTOC capability is independent of MFs. Indeed, the nuclear surface of higher plant cells is thought to function as a MTOC site (Vantard et al. 1990, Mizuno 1993, Stoppin et al. 1994, Meier 2001, Baluška et al. 1997, Yoneda and Hasezawa 2003). In addition, γ-tubulin, a component of the MTOC, accumulated on the nuclear surface in non-treated cells (Fig. 5, Control), suggesting that γ-tubulin participates as MTOC component on the nuclear surface at the M/G1 interface. Other MTOC components on the nuclear surface, for example, EF-1α (Kumagai et al. 1995) and Spc98p (Erhardt et al. 2002), may show similar localization to γ-tubulin, and may act on MTOC components. Indeed, EF-1α was localized at the MT nucleation site on the nuclear surface (Kumagai et al. 1995). In cases where MTOC components and tubulin proteins accumulated on the nuclear surface, MT nucleation could occur in the absence of MFs. Extra-phragmoplast formation Interestingly, when MFs were disrupted by BA treatment, the MTs that were nucleated on the nuclear surface did not reach the cell cortex but rather formed the extra-phragmoplasts (Fig. 3B, BA). The extra-phragmoplasts appeared in a mono-layered form; that is half of the “anti-parallel” phragmoplast MT arrays (Fig. 2A, b). In normal mitotic cells, the phragmoplasts appear at the central region of the anaphase spindle, so that the anti-parallel phragmoplast MTs are thought to be due to the non-kinetochore spindle MTs overlapping between the daughter nuclei (Hepler and Wolniak 1984, Baskin and Cande 1990). Although phragmoplast formation necessarily follows spindle separation in vegetative cells, it is also known that phragmoplasts are organized in the absence of the spindle and nuclear division in reproductive cells (Brown and Lemmon 2001). In cellularization of syncytia, “adventitious phragmoplasts” were formed between neighboring, non-sister as well as sister nuclei (Shimamura et al. 1998, Brown and Lemmon 1988, Brown and Lemmon 2001). Adventitious phragmoplasts were initiated from overlapped MTs of opposing radial MT systems nucleated from two perinuclear areas (Brown and Lemmon 2001), resulting in the anti-parallel MT arrays. These adventitious phragmoplasts are independent of mitotic spindles, although they are necessarily organized between the nuclei as well as mitotic phragmoplasts. However, the extra-phragmoplasts, first identified in this study, did not become organized at the central region of the mitotic spindles or nuclei. This may suggest that if components involved in the phragmoplasts accumulate, then phragmoplast-like structures could be organized without the spindle, the nuclei, or MFs. Some phragmoplast-associated components had been reported: for example, kinesin-like proteins (Liu et al. 1996, Bowser and Reddy 1997, Asada and Shibaoka 1994, Asada et al. 1997, Lee et al. 2001), cytokinesis-specific syntaxin, KNOLLE (Waizenegger et al. 2000, Völker et al. 2001, Muller et al. 2003) and the MAP cascade (Nishihama and Machida 2001, Soyano et al. 2003). These may also contribute to the extra-phragmoplasts and the extra-cell plates. The MFs may therefore retain these proteins at the original phragmoplast. In addition, the extra-phragmoplasts, found in BA-treated cells, possess a mono-layered structure. This may suggest that the existence of the mitotic spindle or nuclei at both sides of the phragmoplast formation site is necessary to form anti-parallel phragmoplast MT arrays, or that the MFs are important in the formation of the anti-parallel MT structures. Conclusion In summary, our observations suggest that MFs are necessary for CMT reorganization at the M/G1 interface, and are in accordance with the finding that tubulin is translocated from the phragmoplast to the cell cortex (Fig. 6). This is the first observation that phragmoplasts appear irrespective of the position of the mitotic spindle or nuclei, and that multiple cell plates can be formed in one nuclear division. These findings also suggest that phragmoplasts can be organized independently of the spindle, nuclei, or MFs, if the components involved in phragmoplast formation are able to accumulate. Using this system, the mechanisms involved in phragmoplast formation, for example, the functions of the phragmoplast-associated kinesins, can be investigated in detail. Materials and Methods Plant material and synchronization Suspension cultures of a BY-GT16 (BY-2 cells expressing GFP-tubulin fusion protein clone 16) cell line were maintained as the original tobacco BY-2 cell suspensions (Nagata et al. 1992). Briefly, the BY-GT16 cell suspensions were diluted 80-fold with a modified Linsmaier and Skoog medium at weekly intervals, and agitated on a rotary shaker at 130 rpm at 27°C in the dark. For cell synchronization, 10 ml of 7-day-old BY-GT16 cells were transferred to 95 ml of fresh medium and cultured for 24 h with 5 mg liter–1 aphidicolin (Sigma Chemical Co., St. Louis, MO, U.S.A.). The cells were washed with 10 volumes of the fresh medium and then resuspended in the same medium. The peak of mitotic index of ca. 60% could be observed at 9.5–10 h after the release from aphidicolin (Kumagai et al. 2001). Drug treatment BA (Wako Pure Chemical Industries, Ltd., Osaka, Japan; Saito et al. 1998) was dissolved in dimethyl sulfoxide (DMSO) to 1 mM and stored at –20°C. The solution was thawed immediately before use and added to the cell suspension at a final concentration of 1 µM. The treated cell suspension was incubated on a rotary shaker at 130 rpm at 27°C in the dark. Time-sequence observations and spatial analysis of BY-GT16 cells using CLSM and deconvolution microscopy Co-localization of GFP fluorescence and tubulin molecules has been clearly observed in living BY-GT16 cells (Kumagai et al. 2001). For the time-sequence observations, synchronized or 3- to 4-day-old BY-GT16 cells were transferred into ϕ35 mm Petri dishes with poly-l-lysine-coated ϕ14 mm coverslip windows at the bottom (Matsunami Glass Ind., Ltd., Osaka, Japan). The dishes were placed onto the inverted platform of a fluorescence microscope (IX, Olympus Co. Ltd., Tokyo, Japan) equipped with a confocal laser scanning head system (GB-200, Olympus) or a cooled CCD camera head system (CoolSNAP HQ, PhotoMetrics Inc., Huntington Beach, Canada). Using the latter system, the Z-stack figures acquired were deconvoluted by AutoDeblur software (AutoQuant Imaging Inc., CA, U.S.A.) to digitally display the optical sections, which were then 3D-reconstructed using MetaMorph software (Universal Imaging Co., Downingtown, Panama). Subsequently, the images were digitally processed, using Photoshop software (Adobe Systems Inc., CA, U.S.A.). Immunocytochemistry and cell staining For the simultaneous observation of MTs and MFs or γ-tubulin, BY-GT16 cells were treated with 250 µM m-maleimidobenzoyl-N-hydroxysuccinimide ester (Pierce Chemical Co., Rockford, IL, U.S.A.) for 30 min prior to fixation. They were fixed with 3.7% (w/v) formaldehyde solution for 1 h and then placed onto coverslips coated with poly-l-lysine. The cells were then treated with an enzyme solution containing 0.1% pectolyase Y23 and 1% cellulase Y-C (both from Seishin Co., Tokyo, Japan) for 5 min, followed by washing with PMEG solution (50 mM PIPES, 1 mM MgSO4, 5 mM EGTA, and 1% Glycerol, pH 6.8). Treatment with a detergent solution containing 1% IGEPAL CA-630 was then performed for 15 min, followed by washing with phosphate-buffered saline (PBS; 20 mM Na-phosphate and 150 mM NaCl, pH 7.0). The cells were subsequently treated with a glycine solution (0.1 M glycine, 1% bovine serum albumin and 0.005% Triton X-100 in PBS) for 10 min, followed by washing with PBS. To stain γ-tubulin, the cells were incubated with clone G9 antibody (mouse monoclonal antibody raised against Schizosaccharomyces pombe γ-tubulin; Kumagai et al. 2003) for 1 h, washed with PBS, and then incubated with a rhodamine-conjugated goat anti-mouse antibody (Cappel Co., West Chester, PA, U.S.A.) for 1 h, followed by washing with PBS. To stain MFs, the cells were incubated with a rhodamine-phalloidin solution [3.3 µM rhodamine-phalloidin (Sigma), 100 mM PIPES, 1 mM MgSO4, 2 mM EGTA (pH 6.9), 5% DMSO, 0.1% IGEPAL CA-630] for 30 min. After DNA staining with 20 mg liter–1 DAPI for 5 min, the cells were finally embedded in a glycerol solution containing an antifading reagent (SlowFade Light; Molecular Probes, Oregon, U.S.A). To observe the newly synthesized cell plate, BY-GT16 cells were stained with 0.05% aniline blue (Biosupplies Australia, Parkville, Victoria, Australia) for 30 min. All procedures described above were performed at room temperature. The cells were subsequently observed by CLSM or deconvolution microscopy as described above. Acknowledgments The authors thank Prof. N. Kondo of Teikyo University of Science & Technology for critical reading of the manuscript, and this paper dedicated to him on the occasion of his happy retirement from the University of Tokyo. This study was supported in part by a Grant-in-Aid for Scientific Research on Priority Areas (Grant No. 15301209) from the Ministry of Education, Culture, Sports, Science and Technology, Japan, to S.H. 1 Corresponding author: E-mail, hasezawa@k.u-tokyo.ac.jp; Fax, +81-4-7136-3706. View largeDownload slide Fig. 1 Localization of MTs and MFs at the M/G1 interface. BY-GT16 cells were stained with rhodamine-phalloidin and DAPI. GFP-tubulin (A, a-1, b-1, c-1, d-1; B, a-1, b-1); rhodamine-phalloidin (A, a-2, b-2, c-2, d-2; B, a-2, b-2); DAPI (A, a-3, b-3, c-3, d-3; B, a-3, b-3); and their merged images (A, a-4, b-4, c-4, d-4; B, a-4, b-4) were obtained by deconvolution microscopy. (A) At metaphase, the MTs formed the mitotic spindle (a-1), and the actin-depleted zone (ADZ) surrounded it (a-2, bracket). When the phragmoplast appeared at telophase, the MFs co-localized with the phragmoplast MTs, especially in the central region of the phragmoplast (b-1, b-2, arrows). At the collapse of the phragmoplast MTs, the MFs that were co-localized there also disappeared (c-1, c-2, arrows). At the early G1 phase, the MFs no longer existed near the division site, and ran parallel to the longitudinal axis of the daughter cell (d-2). (B) BY-GT16 cells were treated with BA at a final concentration of 1 µM for 30 min. The MFs were completely disrupted by this treatment (b-2). The images from each fluorescent dye were photographed using the same exposure time and were processed digitally at the same level of correction. Bars indicate 10 µm. View largeDownload slide Fig. 1 Localization of MTs and MFs at the M/G1 interface. BY-GT16 cells were stained with rhodamine-phalloidin and DAPI. GFP-tubulin (A, a-1, b-1, c-1, d-1; B, a-1, b-1); rhodamine-phalloidin (A, a-2, b-2, c-2, d-2; B, a-2, b-2); DAPI (A, a-3, b-3, c-3, d-3; B, a-3, b-3); and their merged images (A, a-4, b-4, c-4, d-4; B, a-4, b-4) were obtained by deconvolution microscopy. (A) At metaphase, the MTs formed the mitotic spindle (a-1), and the actin-depleted zone (ADZ) surrounded it (a-2, bracket). When the phragmoplast appeared at telophase, the MFs co-localized with the phragmoplast MTs, especially in the central region of the phragmoplast (b-1, b-2, arrows). At the collapse of the phragmoplast MTs, the MFs that were co-localized there also disappeared (c-1, c-2, arrows). At the early G1 phase, the MFs no longer existed near the division site, and ran parallel to the longitudinal axis of the daughter cell (d-2). (B) BY-GT16 cells were treated with BA at a final concentration of 1 µM for 30 min. The MFs were completely disrupted by this treatment (b-2). The images from each fluorescent dye were photographed using the same exposure time and were processed digitally at the same level of correction. Bars indicate 10 µm. View largeDownload slide Fig. 2 Effect of BA treatment on CMT reorganization. The cortex and mid-plane of BY-GT16 cells were simultaneously observed by CLSM at 15-min intervals. In control cells, transverse CMTs were organized within 60 min after the disappearance of the phragmoplast (a, cortex). However, in BA-treated cells, the CMTs were not reorganized even 3 h after phragmoplast collapse (b, c, cortex). In contrast, in about 80% of the BA-treated cells, fluorescence from the GFP-tubulin did not move and remained at the division plane (b, mid-plane), whereas in about 20% of these cells, phragmoplast-like structures were organized (c, mid-plane, arrow) after collapse of the original phragmoplasts (c, arrowheads). Bars indicate 10 µm. View largeDownload slide Fig. 2 Effect of BA treatment on CMT reorganization. The cortex and mid-plane of BY-GT16 cells were simultaneously observed by CLSM at 15-min intervals. In control cells, transverse CMTs were organized within 60 min after the disappearance of the phragmoplast (a, cortex). However, in BA-treated cells, the CMTs were not reorganized even 3 h after phragmoplast collapse (b, c, cortex). In contrast, in about 80% of the BA-treated cells, fluorescence from the GFP-tubulin did not move and remained at the division plane (b, mid-plane), whereas in about 20% of these cells, phragmoplast-like structures were organized (c, mid-plane, arrow) after collapse of the original phragmoplasts (c, arrowheads). Bars indicate 10 µm. View largeDownload slide Fig. 3 Time-lapse observations of extra-phragmoplast formation. BY-GT16 cells were time-sequentially observed by CLSM at 3-min intervals. In control BY-GT16 cells, tubulin proteins released from the collapsing phragmoplast were translocated to the nuclear surface (control, 9–18 min), from where MTs elongated to the cell cortex (Control, 27 min, arrow). In BA-treated BY-GT16 cells, as the original phragmoplast collapsed, the MTs became organized near the nuclear surface (BA, 42–60 min, arrows), and gradually formed a phragmoplast-like structure (BA, 78–114 min). Bars indicate 10 µm. View largeDownload slide Fig. 3 Time-lapse observations of extra-phragmoplast formation. BY-GT16 cells were time-sequentially observed by CLSM at 3-min intervals. In control BY-GT16 cells, tubulin proteins released from the collapsing phragmoplast were translocated to the nuclear surface (control, 9–18 min), from where MTs elongated to the cell cortex (Control, 27 min, arrow). In BA-treated BY-GT16 cells, as the original phragmoplast collapsed, the MTs became organized near the nuclear surface (BA, 42–60 min, arrows), and gradually formed a phragmoplast-like structure (BA, 78–114 min). Bars indicate 10 µm. View largeDownload slide Fig. 4 Features of the extra-phragmoplast by BA treatment. (A) Each focal plane was obtained by CLSM every 1 µm. The data were digitally processed to reconstruct 3-D images, that are presented in front view (a-1, b-1) and rotated view (a-2, b-2). Control cells had circular and bi-layered phragmoplasts (a-1, a-2). When BA was applied at metaphase/telophase, an extra-phragmoplast appeared after collapse of the original phragmoplast (b-1, b-2). The extra-phragmoplast appeared disordered and in a mono-layered form. (B) BY-GT16 cells were stained with 0.05% aniline blue solution and then photographed by deconvolution microscopy. GFP-tubulin (a-1, b-1), aniline blue (a-2, b-2), and their merged images (a-3, b-3) are shown. In the control cell, a flat cell plate was formed (a-2). In the BA-treated cell, an extra-phragmoplast (b-1, arrow) formed an extra-cell plate with a somewhat distorted structure (b-2, arrow). Arrowheads indicate the original division plane. Bars indicate 10 µm. View largeDownload slide Fig. 4 Features of the extra-phragmoplast by BA treatment. (A) Each focal plane was obtained by CLSM every 1 µm. The data were digitally processed to reconstruct 3-D images, that are presented in front view (a-1, b-1) and rotated view (a-2, b-2). Control cells had circular and bi-layered phragmoplasts (a-1, a-2). When BA was applied at metaphase/telophase, an extra-phragmoplast appeared after collapse of the original phragmoplast (b-1, b-2). The extra-phragmoplast appeared disordered and in a mono-layered form. (B) BY-GT16 cells were stained with 0.05% aniline blue solution and then photographed by deconvolution microscopy. GFP-tubulin (a-1, b-1), aniline blue (a-2, b-2), and their merged images (a-3, b-3) are shown. In the control cell, a flat cell plate was formed (a-2). In the BA-treated cell, an extra-phragmoplast (b-1, arrow) formed an extra-cell plate with a somewhat distorted structure (b-2, arrow). Arrowheads indicate the original division plane. Bars indicate 10 µm. View largeDownload slide Fig. 5 Localization of γ-tubulin on the original- and extra-phragmoplasts. BY-GT16 cells were stained with anti-γ-tubulin antibody and rhodamine-conjugated secondary antibody. GFP-α-tubulin (a-1, b-1, c-1, d-1), anti-γ-tubulin (a-2, b-2, c-2, d-2), and their merged images (a-3, b-3, c-3, d-3) were obtained by CLSM. The γ-tubulin stained the outer edges of phragmoplast MTs (a-1, a-2, a-3). As the phragmoplast collapsed, the α-tubulin and γ-tubulin were translocated to the nuclear surface (b-1, b-2, b-3). The γ-tubulin then moved to the cell cortex along the MTs nucleated from the nuclear surface (c-1, c-2, c-3, arrows). When BA was applied and an extra-phragmoplast was organized, the γ-tubulin co-localized there (d-1, d-2, d-3, arrow). Bars indicate 10 µm. View largeDownload slide Fig. 5 Localization of γ-tubulin on the original- and extra-phragmoplasts. BY-GT16 cells were stained with anti-γ-tubulin antibody and rhodamine-conjugated secondary antibody. GFP-α-tubulin (a-1, b-1, c-1, d-1), anti-γ-tubulin (a-2, b-2, c-2, d-2), and their merged images (a-3, b-3, c-3, d-3) were obtained by CLSM. The γ-tubulin stained the outer edges of phragmoplast MTs (a-1, a-2, a-3). As the phragmoplast collapsed, the α-tubulin and γ-tubulin were translocated to the nuclear surface (b-1, b-2, b-3). The γ-tubulin then moved to the cell cortex along the MTs nucleated from the nuclear surface (c-1, c-2, c-3, arrows). When BA was applied and an extra-phragmoplast was organized, the γ-tubulin co-localized there (d-1, d-2, d-3, arrow). Bars indicate 10 µm. View largeDownload slide Fig. 6 A scheme of MF-dependent tubulin recruitment. MTs involving tubulins, MFs, MTOC sites, and putative phragmoplast-associated proteins are shown in green, red, yellow, and blue, respectively. Tubulins, released from the collapsing phragmoplast, translocates to the nuclear surface, and nascent MTs elongate, from the MTOC site on the nuclear surface far away from the division site, to the cell cortex with the guide of MFs (Control). In the absence of MFs, the tubulin transferred to the nuclear surface organizes an extra-phragmoplast, probably with the aid of phragmoplast-associated proteins that are inadequately translocated. View largeDownload slide Fig. 6 A scheme of MF-dependent tubulin recruitment. MTs involving tubulins, MFs, MTOC sites, and putative phragmoplast-associated proteins are shown in green, red, yellow, and blue, respectively. Tubulins, released from the collapsing phragmoplast, translocates to the nuclear surface, and nascent MTs elongate, from the MTOC site on the nuclear surface far away from the division site, to the cell cortex with the guide of MFs (Control). In the absence of MFs, the tubulin transferred to the nuclear surface organizes an extra-phragmoplast, probably with the aid of phragmoplast-associated proteins that are inadequately translocated. Abbreviations ADZ actin-depleted zone BA Bistheonellide A CLSM confocal laser scanning microscopy CMT cortical microtubule DAPI 4′,6-diamidino-2-phenylindole GFP green fluorescent protein MF actin microfilament MT microtubule MTOC microtubule organizing center PPB preprophase band. References Asada, T. and Shibaoka, H. ( 1994) Isolation of polypeptides with microtubule-translocating activity from phragmoplasts of tobacco BY-2 cells. J. Cell Sci.  107: 2249–2257. Google Scholar Asada, T., Kuriyama, R. and Shibaoka, H. ( 1997) TKRP125, a kinesin-related protein involved in the centrosome-independent organization of the cytokinetic apparatus in tobacco BY-2 cells. J. Cell Sci.  110: 179–189. Google Scholar Baluška, F., Volkmann, D. and Bariow, P.W. ( 1997) Nuclear components with microtubule-organizing properties in multicellular eukaryotes: Functional and evolutionary considerations. Int. Rev. Cytol.  175: 91–135. Google Scholar Baskin, T.I. and Cande, W.Z. ( 1990) The structure and function of the mitotic spindle in flowering plants. Annu. Rev. Plant Physiol. Plant Mol. Biol.  41: 277–315. Google Scholar Bowser, J. and Reddy, A.S.N. ( 1997) Localization of a kinesin-like calmodulin-binding protein in dividing cells of Arabidopsis and tobacco. Plant J.  12: 1429–1437. Google Scholar Brown, R.C. and Lemmon, B.E. ( 1988) Cytokinesis occurs at boundaries of domains delimited by nuclear-based microtubules in sporocytes of Conocephalum conicum (Bryophyta). Cell Motil. Cytoskeleton  11: 139–146. Google Scholar Brown, R.C. and Lemmon, B.E. ( 2001) Phragmoplasts in the absence of nuclear division. J. Plant Growth Regul.  20: 151–161. Google Scholar Cleary, A.L. ( 1995) F-actin redistributions at the division site in living Tradescantia stomatal complexes as revealed by microinjection of rhodamine-phalloidin. Protoplasma  185: 152–165. Google Scholar Cleary, A.L., Gunning, B.E.S., Wasteneys, G.O. and Hepler, P.K. ( 1992) Microtubule and F-action dynamics at the division site in living Tradescantia stamen hair cells. J. Cell Sci.  103: 977–988. Google Scholar Erhardt, M., Stoppin-Mellet, V., Campagne, S., Canaday, J., Mutterer, J., Fabian, T., Sauter, M., Muller, T., Peter, C., Lambert, A.M. and Shimit, A.C. ( 2002) The plant SPC98p homologue colocalizes with γ-tubulin at microtubule nucleation sites and is required for microtubule nucleation. J. Cell Sci.  115: 2423–2431. Google Scholar Granger, C.L. and Cyr, R.J. ( 2000) Microtubule reorganization in tobacco BY-2 cells stably expressing GFP-MBD. Planta  210: 502–509. Google Scholar Hasezawa, S., Sano, T. and Nagata, T. ( 1998) The role of microfilaments in the organization and orientation of microtubules during the cell cycle transition from M phase to G1 phase in tobacco BY-2 cells. Protoplasma  202: 105–114. Google Scholar Hasezawa, S., Ueda, K. and Kumagai, F. ( 2000) Time-sequence observations of microtubule dynamics throughout mitosis in living cell suspensions of stable transgenic Arabidopsis – direct evidence for the origin of cortical microtubules at M/G1 interface. Plant Cell Physiol.  41: 244–250. Google Scholar Hepler, P.K. and Wolniak, S.M. ( 1984) Membranes in the mitotic apparatus: their structure and function. Int. Rev. Cytol.  90: 169–238. Google Scholar Hoshino, H., Yoneda, A., Kumagai, F. and Hasezawa, S. ( 2003) Roles of actin-depleted zone and preprophase band in determining the division site of higher-plant cells, a tobacco BY-2 cell line expressing GFP-tubulin. Protoplasma  222: 157–165. Google Scholar Kakimoto, T. and Shibaoka, H. ( 1987) Actin filaments and microtubules in the preprophase band and phragmoplast of tobacco cells. Protoplasma  140: 151–156. Google Scholar Kakimoto, T. and Shibaoka, H. ( 1988) Cytoskeletal ultrastructure of phragmoplast-nuclei complex isolated from cultured tobacco cells. Protoplasma (Suppl.) 2: 95–103. Google Scholar Kumagai, F. and Hasezawa, S. ( 2001) Dynamic organization of microtubules and microfilaments during cell cycle progression in higher plant cells. Plant Biol.  3: 4–16. Google Scholar Kumagai, F., Hasezawa, S., Takahashi, Y. and Nagata, T. ( 1995) The involvement of protein synthesis elongation factor 1α in the organization of microtubules on the perinuclear region during the cell cycle transition from M phase to G1 phase in tobacco BY-2 cells. Bot. Acta  108: 467–473. Google Scholar Kumagai, F., Nagata, T., Yahara, N., Moriyama, Y., Horio, T., Naoi, K., Hashimoto, T., Murata, T. and Hasezawa, S. ( 2003) γ-Tubulin distribution during cortical microtubule reorganization at the M/G1 interface in tobacco BY-2 cells. Eur. J. Cell Biol.  82: 43–51. Google Scholar Kumagai, F., Yoneda, A., Tomida, T., Sano, T., Nagata, T. and Hasezawa, S. ( 2001) Fate of nascent microtubules organized at the M/G1 interface, as visualized by synchronized tobacco BY-2 cells stably expressing GFP-tubulin: Time-sequence observations of the reorganization of cortical microtubules in living plant cells. Plant Cell Physiol.  42: 723–732. Google Scholar Lee, Y.-R.J., Giang, H.M. and Liu, B. ( 2001) A novel kinesin-related protein specifically associates with the phragmoplast organelles. Plant Cell  13: 2427–2439. Google Scholar Liu, B., Cyr, R.J. and Palevitz, B.A. ( 1996) A kinesin-like protein, KatAp, in the cells of Arabidopsis and other plants. Plant Cell  8: 119–132. Google Scholar Meier, I. ( 2001) The plant nuclear envelope . Cell Mol. Life Sci.  58: 1774–1780. Google Scholar Mineyuki, Y. and Palevitz, B.A. ( 1990) Relationship between preprophase band organization, F-actin and the division site in Allium. Fluorescence and morphometric studies on cytochalashin-treated cells. J. Cell Sci.  97: 283–295. Google Scholar Mizuno, K. ( 1993) Microtubule-nucleation sites on nuclei of higher plant cells. Protoplasma  173: 77–85. Google Scholar Muller, I., Wagner, W., Volker, A., Schellmann, S., Nacry, P., Kuttner, F., Schwarz-Sommer, Z., Mayer, U. and Jurgens, G. ( 2003) Syntaxin specificity of cytokinesis in Arabidopsis. Nat. Cell Biol.  5: 531–534. Google Scholar Nagata, T., Nemoto, Y. and Hasezawa, S. ( 1992) Tobacco BY-2 cell line as the “Hela” cell in the cell biology of higher plants. Int. Rev. Cytol.  132: 1–30. Google Scholar Nishihama, R. and Machida, Y. ( 2001) Expansion of the phragmoplast during plant cytokinesis: a MAPK pathway may MAP it out. Curr. Opin. Plant Biol.  4: 507–512. Google Scholar Nishihama, R., Shoyano, T., Ishikawa, M., Araki, S., Tanaka, H., Asada, T., Irie, K., Ito, M., Terada, M., Banno, H., Yamazaki, Y. and Machida, Y. ( 2002) Expansion of the cell plate in plant cytokinesis requires a kinesin-like protein/MAPKKK complex. Cell  109: 87–99. Google Scholar Palevitz, B.A. and Hepler, P.K. ( 1974) Control of plane of division during stomatal differentiation in Allium. 1. Spindle reorientation. Chromosoma  46: 327–341. Google Scholar Saito, S., Watabe, S., Ozaki, H., Kobayashi, M., Suzuki, T., Kobayashi, H., Fusetani, N. and Karaki, H. ( 1998) Actin-depolymerizing effect of dimeric macrolides, bistheonellide A and swinholide A. J. Biochem.  123: 571–578. Google Scholar Samuels, A.L., Giddings, T.H. and Staehelin, L.A. ( 1995) Cytokinesis in tobacco BY-2 and root tip cells: a new model of cell plate formation in higher plants. J. Cell Biol.  130: 1345–1357. Google Scholar Shimamura, M., Deguchi, H. and Mineyuki, Y. ( 1998) Meiotic cytokinetic apparatus in the formation of the linear spore tetrads of Conocephalum japonicum (Bryophyta). Planta  206: 604. Google Scholar Soyano, T., Nishihama, R., Morikiyo, K., Ishikawa, M. and Machida, Y. ( 2003) NQK1/NtMEK1 is a MAPKK that acts in the NPK1 MAPKKK-mediated MAPK cascade and is required for plant cytokinesis. Genes Dev.  17: 1055–1067. Google Scholar Stoppin, V., Vantard, M., Schmit, A.C. and Lambert, A.M. ( 1994) Isolated plant nuclei nucleate microtubule assembly: The nuclear surface in higher plants has centrosome-like activity. Plant Cell  6: 1099–1106. Google Scholar Valster, A.H. and Hepler, P.K. ( 1997) Caffeine inhibition of cytokinesis: effect on the phragmoplast cytoskeleton in living Tradescantia samen hair cells. Protoplasma  196: 155–166. Google Scholar Vantard, M., Levilliers, N., Hill, A.M., Adoutte, A. and Lambert, A.M. ( 1990) Incorporation of Paramecium axonemal tubulin into higher plant cells reveals functional sites of microtubule assembly. Proc. Natl Acad. Sci. USA  87: 8825–8829. Google Scholar Verma, D.P.S. ( 2001) Cytokinesis and building of the cell plate in plants. Annu. Rev. Plant Physiol. Plant Mol. Biol.  52: 751–784. Google Scholar Völker, A., Stierhof, Y.-D. and Jürgens, G. ( 2001) Cell cycle-independent expression of the Arabidopsis cytokinesis-specific syntaxin KNOLLE results in mistargeting to the plasma membrane and is not sufficient for cytokinesis. J. Cell Sci.  114: 3001–3012. Google Scholar Waizenegger, I., Lukowitz, W., Assaad, F., Schwarz, H., Jurgens, G. and Mayer, U. ( 2000) The Arabidopsis KNOLLE and KEULE genes interact to promote vesicle fusion during cytokinesis. Curr. Biol.  10: 1371–1374. Google Scholar Yoneda, A. and Hasezawa, S. ( 2003) Origin of cortical microtubules organized at M/G1 interface: recruitment of tubulin from phragmoplast to nascent microtubules. Eur. J. Cell Biol.  82: 461–471. Google Scholar Yoneda, A., Kutsuna, N. and Hasezawa, S. ( 2003) Dynamic organization of microtubules and vacuoles visualized by GFP in living plant cells. Recent Res. Dev. Plant Mol. Biol.  1: 127–137. Google Scholar
TI - Disruption of Actin Microfilaments Causes Cortical Microtubule Disorganization and Extra-Phragmoplast Formation at M/G1 Interface in Synchronized Tobacco Cells
JF - Plant and Cell Physiology
DO - 10.1093/pcp/pch091
DA - 2004-06-15
UR - https://www.deepdyve.com/lp/oxford-university-press/disruption-of-actin-microfilaments-causes-cortical-microtubule-r0MXAj9VjY
SP - 761
EP - 769
VL - 45
IS - 6
DP - DeepDyve
ER -