Nu MA assemblies organize microtubule asters to establish spindle bipolarity in acentrosomal human cells
2019; Springer Nature; Volume: 39; Issue: 2 Linguagem: Inglês
10.15252/embj.2019102378
ISSN1460-2075
AutoresTakumi Chinen, Shohei Yamamoto, Yutaka Takeda, Koki Watanabe, Kanako Kuroki, Kaho Hashimoto, Daisuke Takao, Daiju Kitagawa,
Tópico(s)14-3-3 protein interactions
ResumoArticle29 November 2019free access NuMA assemblies organize microtubule asters to establish spindle bipolarity in acentrosomal human cells Takumi Chinen Division of Centrosome Biology, Department of Molecular Genetics, National Institute of Genetics, Mishima, Shizuoka, Japan Department of Physiological Chemistry, Graduate School of Pharmaceutical Science, The University of Tokyo, Bunkyo, Tokyo, Japan Search for more papers by this author Shohei Yamamoto Division of Centrosome Biology, Department of Molecular Genetics, National Institute of Genetics, Mishima, Shizuoka, Japan Department of Physiological Chemistry, Graduate School of Pharmaceutical Science, The University of Tokyo, Bunkyo, Tokyo, Japan Graduate Program in Bioscience, Graduate School of Science, University of Tokyo, Hongo, Tokyo, Japan Search for more papers by this author Yutaka Takeda Department of Physiological Chemistry, Graduate School of Pharmaceutical Science, The University of Tokyo, Bunkyo, Tokyo, Japan Search for more papers by this author Koki Watanabe Division of Centrosome Biology, Department of Molecular Genetics, National Institute of Genetics, Mishima, Shizuoka, Japan Department of Physiological Chemistry, Graduate School of Pharmaceutical Science, The University of Tokyo, Bunkyo, Tokyo, Japan Department of Genetics, School of Life Science, The Graduate University for Advanced Studies (SOKENDAI), Hayama, Kanagawa, Japan Search for more papers by this author Kanako Kuroki Department of Physiological Chemistry, Graduate School of Pharmaceutical Science, The University of Tokyo, Bunkyo, Tokyo, Japan Search for more papers by this author Kaho Hashimoto Department of Physiological Chemistry, Graduate School of Pharmaceutical Science, The University of Tokyo, Bunkyo, Tokyo, Japan Search for more papers by this author Daisuke Takao Division of Centrosome Biology, Department of Molecular Genetics, National Institute of Genetics, Mishima, Shizuoka, Japan Department of Physiological Chemistry, Graduate School of Pharmaceutical Science, The University of Tokyo, Bunkyo, Tokyo, Japan Search for more papers by this author Daiju Kitagawa Corresponding Author [email protected] orcid.org/0000-0003-2509-5977 Division of Centrosome Biology, Department of Molecular Genetics, National Institute of Genetics, Mishima, Shizuoka, Japan Department of Physiological Chemistry, Graduate School of Pharmaceutical Science, The University of Tokyo, Bunkyo, Tokyo, Japan Department of Genetics, School of Life Science, The Graduate University for Advanced Studies (SOKENDAI), Hayama, Kanagawa, Japan Search for more papers by this author Takumi Chinen Division of Centrosome Biology, Department of Molecular Genetics, National Institute of Genetics, Mishima, Shizuoka, Japan Department of Physiological Chemistry, Graduate School of Pharmaceutical Science, The University of Tokyo, Bunkyo, Tokyo, Japan Search for more papers by this author Shohei Yamamoto Division of Centrosome Biology, Department of Molecular Genetics, National Institute of Genetics, Mishima, Shizuoka, Japan Department of Physiological Chemistry, Graduate School of Pharmaceutical Science, The University of Tokyo, Bunkyo, Tokyo, Japan Graduate Program in Bioscience, Graduate School of Science, University of Tokyo, Hongo, Tokyo, Japan Search for more papers by this author Yutaka Takeda Department of Physiological Chemistry, Graduate School of Pharmaceutical Science, The University of Tokyo, Bunkyo, Tokyo, Japan Search for more papers by this author Koki Watanabe Division of Centrosome Biology, Department of Molecular Genetics, National Institute of Genetics, Mishima, Shizuoka, Japan Department of Physiological Chemistry, Graduate School of Pharmaceutical Science, The University of Tokyo, Bunkyo, Tokyo, Japan Department of Genetics, School of Life Science, The Graduate University for Advanced Studies (SOKENDAI), Hayama, Kanagawa, Japan Search for more papers by this author Kanako Kuroki Department of Physiological Chemistry, Graduate School of Pharmaceutical Science, The University of Tokyo, Bunkyo, Tokyo, Japan Search for more papers by this author Kaho Hashimoto Department of Physiological Chemistry, Graduate School of Pharmaceutical Science, The University of Tokyo, Bunkyo, Tokyo, Japan Search for more papers by this author Daisuke Takao Division of Centrosome Biology, Department of Molecular Genetics, National Institute of Genetics, Mishima, Shizuoka, Japan Department of Physiological Chemistry, Graduate School of Pharmaceutical Science, The University of Tokyo, Bunkyo, Tokyo, Japan Search for more papers by this author Daiju Kitagawa Corresponding Author [email protected] orcid.org/0000-0003-2509-5977 Division of Centrosome Biology, Department of Molecular Genetics, National Institute of Genetics, Mishima, Shizuoka, Japan Department of Physiological Chemistry, Graduate School of Pharmaceutical Science, The University of Tokyo, Bunkyo, Tokyo, Japan Department of Genetics, School of Life Science, The Graduate University for Advanced Studies (SOKENDAI), Hayama, Kanagawa, Japan Search for more papers by this author Author Information Takumi Chinen1,2, Shohei Yamamoto1,2,3, Yutaka Takeda2, Koki Watanabe1,2,4, Kanako Kuroki2, Kaho Hashimoto2, Daisuke Takao1,2 and Daiju Kitagawa *,1,2,4 1Division of Centrosome Biology, Department of Molecular Genetics, National Institute of Genetics, Mishima, Shizuoka, Japan 2Department of Physiological Chemistry, Graduate School of Pharmaceutical Science, The University of Tokyo, Bunkyo, Tokyo, Japan 3Graduate Program in Bioscience, Graduate School of Science, University of Tokyo, Hongo, Tokyo, Japan 4Department of Genetics, School of Life Science, The Graduate University for Advanced Studies (SOKENDAI), Hayama, Kanagawa, Japan *Corresponding author. Tel: +81 03 5841 4750; E-mail: [email protected] EMBO J (2020)39:e102378https://doi.org/10.15252/embj.2019102378 PDFDownload PDF of article text and main figures.AM PDF Peer ReviewDownload a summary of the editorial decision process including editorial decision letters, reviewer comments and author responses to feedback. ToolsAdd to favoritesDownload CitationsTrack CitationsPermissions ShareFacebookTwitterLinked InMendeleyWechatReddit Figures & Info Abstract In most animal cells, mitotic spindle formation is mediated by coordination of centrosomal and acentrosomal pathways. At the onset of mitosis, centrosomes promote spindle bipolarization. However, the mechanism through which the acentrosomal pathways facilitate the establishment of spindle bipolarity in early mitosis is not completely understood. In this study, we show the critical roles of nuclear mitotic apparatus protein (NuMA) in the generation of spindle bipolarity in acentrosomal human cells. In acentrosomal human cells, we found that small microtubule asters containing NuMA formed at the time of nuclear envelope breakdown. In addition, these asters were assembled by dynein and the clustering activity of NuMA. Subsequently, NuMA organized the radial array of microtubules, which incorporates Eg5, and thus facilitated spindle bipolarization. Importantly, in cells with centrosomes, we also found that NuMA promoted the initial step of spindle bipolarization in early mitosis. Overall, these data suggest that canonical centrosomal and NuMA-mediated acentrosomal pathways redundantly promote spindle bipolarity in human cells. Synopsis Mitotic spindle formation in animal cells involves both centrosomal and acentrosomal pathways. Studies on acentrosomal human cells suggest that canonical centrosomal and NuMA/dynein-mediated mechanisms may redundantly promote spindle biploarity early in mitosis. NuMA promotes spindle bipolarity in the absence of centrosomes. Dynein and the clustering activity of NuMA facilitate acentrosomal spindle pole organization. NuMA organizes the radial array of microtubules, which incorporates kinesin Eg5 to facilitate spindle bipolarization. NuMA also promotes the initial step of spindle bipolarization in cells with centrosomes. Introduction The mitotic spindle, a large cytoskeletal structure, ensures proper chromosomal segregation (Prosser & Pelletier, 2017). In somatic animal cells, the mitotic spindle poles consist of centrosomes and several factors involved in their organization. The centrosomes consist of a pair of centrioles surrounded by pericentriolar material (PCM; Bornens, 2002), which nucleates and anchors microtubules. Centrosomes act as the major microtubule-organizing centers and organize the radial array of microtubules during early mitosis (Kaseda et al, 2012). At the G2/M transition, centrosomes move away from each other through the activity of centrosome separation pathways (Rattner & Berns, 1976; Waters et al, 1993; Whitehead & Rattner, 1998; Toso et al, 2009; Raaijmakers et al, 2012) in order to assemble a bipolar mitotic spindle. At the onset of centrosome separation, the evolutionary conserved kinesin Eg5 (Enos & Morris, 1990; Hagan & Yanagida, 1990; Heck et al, 1993; Blangy et al, 1995) is loaded onto centrosomes. Therefore, the centrosomal microtubules promote the initial steps establishing mitotic spindle bipolarity (Megraw et al, 2001; Bertran et al, 2011; Smith et al, 2011; Kaseda et al, 2012; Hata et al, 2019). In addition to centrosomes, spindle pole organization is regulated by several proteins, such as nuclear mitotic apparatus protein (NuMA) and the dynein–dynactin complex. The functions of these proteins for spindle pole organization and spindle positioning have been well studied in meiotic and mitotic spindles. It has been shown that NuMA is recruited to the minus-ends of microtubules dependent on (Merdes et al, 2000) or independently of dynein (Hueschen et al, 2017). During prophase, NuMA begins to form aggregates in the nucleus (Kisurina-Evgenieva et al, 2004). These small aggregates of NuMA are incorporated around centrosomes within a few minutes after nuclear envelope breakdown (NEBD; Kisurina-Evgenieva et al, 2004). Specific inhibition of NuMA or dynein disrupts spindle pole formation (Merdes et al, 1996, 2000). Thus, the spindle pole is organized by NuMA and the dynein–dynactin complex (Merdes et al, 1996, 2000; Hueschen et al, 2017). In addition to their functions in spindle pole organization, NuMA and the dynein–dynactin complex cooperate to regulate spindle positioning by forming dynein–dynactin–NuMA clusters at the mitotic cell cortex through the clustering activity of NuMA (Okumura et al, 2018). In contrast, oocytes do not have centrosomes; instead, they exhibit acentrosomal spindle formation. In mouse oocytes, the meiotic spindle poles are organized by NuMA, and the microtubule binding activity of NuMA is required for female fertility (Kolano et al, 2012). It has been shown that NuMA localizes at the meiotic spindle poles during meiosis I and II in human oocytes (Xu et al, 2011). In addition to meiotic spindles, previous studies have shown that vertebrate and fly somatic cells can divide continuously, even after the removal of centrosomes (Bonaccorsi et al, 2000; Khodjakov et al, 2000; Basto et al, 2006; Hornick et al, 2011; Sir et al, 2013) and, in green monkey fibroblasts, NuMA accumulates at the acentrosomal spindle poles (Khodjakov et al, 2000). Although these studies suggest that meiotic and mitotic spindle poles are organized by NuMA, the exact functions of NuMA in the initial steps of spindle bipolarization remain to be elucidated. The microtubule organization activity of NuMA has been studied in detail (Nachury et al, 2001; Wiese et al, 2001). It has been shown that NuMA possesses clustering properties through its C-terminus (Harborth et al, 1999) and forms microtubule-independent insoluble complexes that are resistant to nocodazole treatment (Dionne et al, 1999; Merdes et al, 2000; Kisurina-Evgenieva et al, 2004; Hueschen et al, 2017). It has also been reported that the C-terminus of NuMA interacts with microtubules (Du et al, 2002; Haren & Merdes, 2002), presumably mediating centrosome-independent aster formation (Gaglio et al, 1995; Du et al, 2002; Haren & Merdes, 2002). The properties of NuMA, which assemble and organize an aster-like microtubule array, are similar to those of centrosomes in early mitosis. However, the role of NuMA in the initial step of spindle bipolarization, rather than spindle pole organization or spindle positioning, is not fully understood yet. In this study, using live-cell imaging and auxin-inducible degradation (AID) experiments, we revealed that acentrosomal human somatic cells organized bipolar mitotic spindles in a NuMA-dependent manner. In acentrosomal cells, NuMA started to assemble and formed microtubule asters shortly after NEBD. Depletion of NuMA induced umbrella-like monopolar spindles in acentrosomal cells, suggesting that NuMA organizes the microtubule array and Eg5 localization to promote proper separation of the poles. In addition, we found that the depletion of NuMA delayed spindle bipolarization in cells with centrosomes. Collectively, based on the present data, we propose the following model: (i) Canonical centrosomal and NuMA-mediated acentrosomal pathways redundantly promote spindle bipolarization in early mitosis of cells with centrosomes; and (ii) NuMA can compensate for the function of centrosomes in the initial step of bipolar spindle formation in acentrosomal human cells. Results NuMA comprises a structure specific to acentrosomal human cells in the process of bipolar spindle assembly We initially induced the formation of acentrosomal spindles in human cells to understand the mechanism through which human somatic cells can establish spindle bipolarity without centrosomes. For this purpose, we treated HeLa cells with centrinone, a specific PLK4 inhibitor (Wong et al, 2015), to remove the centrosomes. We subsequently examined the mechanism through which the two separate spindle poles were established in these acentrosomal cells. Immunofluorescence was used to analyze the localization patterns of proteins known to accumulate at the mitotic spindle poles (Fig 1A–F, Appendix Figs S1 and S2). We confirmed that, in cells with two centrosomes, the spindle poles consist of NuMA/p150Glued and ASPM–katanin complexes (Fig 1A–C). Interestingly, the acentrosomal cells assembled a bipolar spindle with a dense NuMA structure between the spindle poles (Fig 1A and B). In addition, p150Glued, a dynactin subunit that forms a complex with NuMA, also showed this dense structure (Fig 1C). In contrast, in these spindles, the ASPM–katanin complex (Jiang et al, 2017) was distributed only near the spindle poles (Fig 1B and C). We subsequently observed the localization pattern of the PCM components in acentrosomal cells. Several PCM components (e.g., pericentrin, CDK5RAP2, and Cep192) were not detected at most acentrosomal spindle poles in HeLa cells (Appendix Fig S1A and B). Furthermore, pericentrin was consistently undetectable at the acentrosomal spindle poles of various human cell lines (Appendix Fig S1C and D). Collectively, these data indicate that the organization of NuMA/p150Glued and ASPM–katanin complexes at acentrosomal spindle poles differs. Figure 1. Acentrosomal human cells form a specific structure of NuMA in the process of bipolar spindle assembly A–C. Distribution of spindle pole factors in centrosomal and acentrosomal spindles. DMSO-treated control mitotic spindles (2-centrosomes) and centrinone-treated acentrosomal spindles (0-centrosome) of HeLa cells. (A) Red, gray, green, and blue represent NuMA, α-tubulin, Cep152, and DNA, respectively. Z-projections of eight sections, every 0.3 μm. Scale bar, 5 μm. (B) Red, gray, and blue represent NuMA, katanin p60, and DNA, respectively. Z-projections of five sections, every 0.3 μm. Scale bar, 5 μm. (C) Red, gray, and blue represent p150Glued, ASPM, and DNA, respectively. Z-projections of 10 (2-centrosomes) or five (0-centrosome) sections, every 0.3 μm. Scale bar, 5 μm. D. The structure of NuMA and microtubules in centrinone-treated acentrosomal spindles of HeLa cells. Two-way arrows indicate the bipolarity of elongated NuMA structures. Gray, red, green, and blue represent α-tubulin, NuMA, Cep152, and DNA, respectively. Scale bar, 5 μm. E. The localization of p150Glued in centrinone-treated acentrosomal spindles of HeLa cells. Red, gray, green, and blue represent NuMA, p150Glued, Cep152, and DNA, respectively. Scale bar, 5 μm. F. NuMA distribution in 3D. Images in (D) of NuMA (Red) and chromosomes (blue) were reconstructed in 3D and surface-rendered. G. Representative images of NuMA (arrowheads) in centrosomal and acentrosomal cells before and after photobleaching. DMSO- or centrinone-treated HCT116 TetOsTIR1 NuMA–mAID–mClover–FLAG cells were observed. Time after photobleaching (s) is indicated. Scale bar, 5 μm. H. Normalized fluorescence intensities of NuMA–mClover at centrosomal poles (control, 2-centrosomes) or at acentrosomal spindle poles (0-centrosome) during the FRAP time course (N = 20 cells for each condition). The first three time points are before photobleaching. Error bars show standard deviation (SD). I. Mobile fractions from the normalized recovery curves of NuMA–mClover at centrosomal spindle poles (control, 2-centrosomes) or at acentrosomal spindle poles (0-centrosome). Mobile fractions (roughly corresponds to normalized fluorescence intensities at the last time point of (H)) were determined by single exponential fitting. Line and error bars represent the median and interquartile range (N = 20 cells for each condition). The Mann–Whitney U-test (two-tailed) was used to obtain a P value. Download figure Download PowerPoint Subsequently, we observed the localization patterns of NuMA, α-tubulin, and chromosomes during mitosis to further analyze the establishment of spindle bipolarity in acentrosomal cells. Acentrosomal cells formed a unique pattern of NuMA and p150Glued in the center of the condensed chromosome ring (monopolar-like; Fig 1D–F). These cells organized the radial array of microtubules around the NuMA structure (Fig 1D). In acentrosomal cells, NuMA also showed a bundle-like pattern, which colocalized with microtubules (bundle-like; Fig 1D and F). It has been shown that microtubule bundles form within the meiotic spindle through the assembly of central spindle components (So et al, 2019). However, the bundle-like structure in acentrosomal human cells did not contain the central spindle components, such as PRC1 (Mollinari et al, 2002) and KIF4 (Kurasawa et al, 2004; Appendix Fig S2A and B). The structure of NuMA in acentrosomal cells was similarly formed in various cell lines derived from different human tissues, including nontransformed and hTERT-immortalized human retinal pigment epithelium cells (RPE1; Appendix Fig S2C and D). It has been reported that PLK4 contributes to microtubule nucleation in oocytes (Bury et al, 2017). Thus, we examined whether the observed structure of NuMA could be attributed to the inhibition of PLK4 or to the low microtubule density induced by the removal of centrosomes. Monopolar- and bundle-like NuMA structures were similarly observed in centrosome-eliminated cells after Sas6 depletion using the AID system (Appendix Fig S2E; Yoshiba et al, 2019), suggesting that the aforementioned phenotype was not the result of PLK4 inhibition. We further confirmed that the microtubule density in the acentrosomal spindle was higher than that in the centrosomal spindle (Appendix Fig S2F). This result suggests that the NuMA structure in acentrosomal cells is not formed due to low microtubule density. In addition, we performed fluorescence recovery after photobleaching (FRAP) analysis using HCT116 cells, in which endogenous NuMA was tagged with mClover to further characterize the properties of NuMA in the acentrosomal spindle pole. Prior to photobleaching, the fluorescence intensity of NuMA in acentrosomal spindle poles was significantly higher than that measured in centrosomal poles (Fig 1G, Appendix Fig S3A, and Movies EV1 and EV2). In control cells, the fluorescence of NuMA recovered to 54% on average within 3 min (Fig 1G–I, Movie EV1). In contrast, the fluorescence of NuMA in acentrosomal cells recovered to only 29% on average (Fig 1G–I, Movie EV2), suggesting that a large proportion of NuMA is more static in the acentrosomal spindle poles than in the centrosomal poles. However, the half-time for the recovery and absolute value of the mobile fraction in acentrosomal spindle poles were comparable with those determined in centrosomal poles (Appendix Fig S3B–D). Thus, these data suggest that the acentrosomal spindle pole consists of two populations of NuMA: a dynamic population, which is also present in centrosomal poles; and a static fraction, which is specific to acentrosomal spindle poles. Collectively, these results suggest that a bipolar spindle in acentrosomal human cells is established in two steps: (i) organization of the radial array of microtubules around the dense NuMA structure and (ii) separation of the two spindle poles. This system may be conserved for the formation of bipolar mitotic spindles in acentrosomal human cells. NuMA organizes small asters during NEBD and assembles a small spindle prior to spindle elongation and chromosome congression Next, we further investigated the time course of mitotic events in acentrosomal cells using live-cell imaging. We used time-lapse fluorescence microscopy to track the localization and dynamics of endogenous NuMA tagged with mCherry or mClover in HeLa cells. In control cells with centrosomes, the centrosomes are dissociated through degradation of the centrosomal linker before mitosis (Wang et al, 2014). In these cells, NuMA was rapidly recruited to the two separated spindle poles during NEBD (Appendix Fig S4A and B) as previously described (Kisurina-Evgenieva et al, 2004). In contrast, acentrosomal cells formed several aggregates of NuMA after NEBD (4 min: Fig 2A; 0 min: Fig 2B, Movies EV3 and EV4). Subsequently, the bipolarity of the acentrosomal mitotic spindle, which was judged based on two separate NuMA structures, was established on average within 11.8 min after NEBD (Fig 2B and C). After the establishment of bipolarity of the acentrosomal mitotic spindle, the chromosomes started to be aligned on the metaphase plate (Fig 2B and C). This observation was also confirmed through immunofluorescence analysis with fixed cells, which showed that chromosomes and kinetochores localized around the elongating bundle-like structure of NuMA (Fig 2D). These acentrosomal cells showed a delay in chromosome alignment after NEBD and in the transition from chromosome alignment to the onset of anaphase versus control cells with centrosomes (Appendix Fig S4C and D), suggesting that both processes of spindle bipolarization and spindle assembly checkpoint satisfaction were delayed in acentrosomal spindle formation. In addition, a high frequency of chromosome segregation errors was detectable in acentrosomal cells as previously described (Sir et al, 2013; Appendix Fig S4E). Collectively, these findings indicate that bipolarity within the acentrosomal mitotic spindle is established after NEBD and that the process precedes chromosomal alignment. Figure 2. NuMA organizes small asters at the time of NEBD and assembles a small spindle before spindle elongation and chromosome congression A. Time-lapse observation of the establishment of bipolarity in acentrosomal cells. HeLa cells expressing EGFP–centrin1 and mCherry–NuMA were observed with a 63× objective. Magenta and green represent mCherry–NuMA and EGFP–centrin1, respectively. Arrowheads indicate the assembling NuMA after NEBD. Two-way arrows indicate the bipolarity. Z-projections of 20 sections, every 1.2 μm. Scale bar, 10 μm. Time zero corresponds to NEBD. B. Time-lapse observation of the dynamics of NuMA and chromosomes. Centrinone-treated HeLa cells expressing mCover–NuMA were observed with a 63× objective. Magenta and green represent SiR-DNA and mClover–NuMA, respectively. Arrowheads and two-way arrows indicate the assembling NuMA after NEBD and bipolarity, respectively. Z-projections of 20 sections, every 1.2 μm. Scale bar, 10 μm. Time zero corresponds to NEBD. C. The timing of bipolarity establishment in the acentrosomal pole, chromosome alignment, and anaphase onset in (B). Each plot shows the cumulative percentage of each event at individual time points (N = 49 cells from two independent experiments). D. The distribution of kinetochores during acentrosomal spindle pole separation was confirmed in fixed cells. Two-way arrow indicates the bipolarity of elongated NuMA structures. Red, gray, green, and blue represent NuMA, CENP-C, GT335, and DNA, respectively. Scale bar, 5 μm. E, F. Time-lapse observation of the structure of NuMA and microtubules upon centrosome removal. Centrinone-treated HCT116 TetOsTIR1 NuMA–mAID–mClover–FLAG cells were observed with a 60× objective. Red and green represent NuMA and SiR-tubulin, respectively. Z-projections of 17 sections, every 1 μm. Scale bar, 5 μm. Time zero corresponds to NEBD. (F) An example of NuMA formed several asters (arrowheads) at the time of NEBD. Download figure Download PowerPoint We subsequently examined the mechanism through which separation of the two spindle poles is achieved in acentrosomal cells, with a particular focus on the structure of NuMA and microtubules. We used a spinning disk confocal microscopy SpinSR10 system with a sCMOS camera to track the structure of NuMA and microtubules. Acentrosomal HCT116 NuMA–mAID–mClover cells showed several patterns of NuMA structures (Fig 2E and F, Movies EV5 and EV6). In one example, NuMA started to form two aggregates after NEBD (10 min, Cell 1, Fig 2E). These aggregates subsequently assembled into one structure (30 min, Cell 1, Fig 2E). After the assembly of the monopolar-like spindle (30 min, Cell 1, Fig 2E), the spindle elongated with a bundle-like structure of NuMA (60–70 min Cell 1, Fig 2E). This NuMA structure was eventually separated into two poles (80 min, Cell 1, Fig 2E). In other examples, NuMA formed several asters during NEBD (Fig 2F) or after NEBD (10 min, Cell 2, Fig 2E). These asters then assembled into two aggregates of NuMA to form a small bipolar spindle (40 min, Cell 2, Fig 2E), and this small spindle subsequently started to elongate (40, 70 min, Cell 2, Fig 2E). These observations suggest that NuMA organizes small asters during NEBD and assembles into two separate aggregates that eventually function as the two spindle poles. Microtubules and dynein are required for NuMA assembly to form two acentrosomal spindle poles We next addressed the mechanisms underlying NuMA assembly for acentrosomal spindle pole formation. It has been established that NuMA is transported toward the spindle poles in a microtubule-based motor dynein-dependent manner (Merdes et al, 1996, 2000; Hueschen et al, 2017). Therefore, we validated the requirement of microtubules and dynein for NuMA assembly at acentrosomal spindle poles. Nocodazole, a microtubule polymerization inhibitor, induced multiple asters with NuMA, suggesting that NuMA was assembled into two acentrosomal spindle poles in a microtubule-dependent manner (Fig 3A). Figure 3. Microtubule, dynein, and clustering activity of NuMA are required for the assembly of NuMA aggregates in acentrosomal human cells NuMA structure upon microtubule depolymerization. Nocodazole-treated HCT116 CMVOsTIR1 CEP152–mClover–mAID cells were treated with 100 ng/ml nocodazole. Gray, red, green, and blue represent α-tubulin, NuMA, CEP152 (mClover), and DNA, respectively. Scale bar, 5 μm. Immunostaining of DHC1 and NuMA in centrinone-treated HCT116 TetOsTIR1 DHC1–3X–mAID–mClover cells. Gray, green, red, and blue represent DHC1 (mClover), GT335, NuMA, and DNA, respectively. Scale bar, 5 μm. Schematic illustration of target protein depletion using the AID system to analyze the requirement of dynein for acentrosomal spindle pole formation. NuMA structure upon DHC1 depletion. Centrinone-treated HCT116 TetOsTIR1 DHC1–3X–mAID–mClover cells were treated with 1 μg/ml doxycycline (Dox) and 500 μM indole-3-acetic acid (IAA). Arrowhead indicates the spindle pole. Green, red, gray, and blue represent DHC1 (mClover), NuMA, GT335, and DNA, respectively. Scale bar, 5 μm. Frequency of acentrosomal pole patterns upon DHC1 depletion in (D). Values are mean percentages ± SD from three independent experiments (N ≧ 24 spindles in each experiment). The distribution of NuMA upon DHC1 depletion. Arrowheads indicate the presence of NuMA on microtubules. Green, red, and blue represent α-tubulin, NuMA, and DNA, respectively. Scale bars, 5 μm. Structure of acentrosomal spindle poles upon replacement of endogenous NuMA–mAID–mClover–FLAG with either mCherry–NuMA WT or 5A-3. Arrowheads indicate the assembled NuMA structure. Green, red, gray, and blue represent endogenous NuMA (mClover), expressed NuMA (mCherry), GT335, and DNA, respectively. Z-projections of 21 sections, every 1 μm. Scale bar, 5 μm. The number of NuMA structure in (G). Values are mean percentages ± SD from four independent experiments (N ≧ 15 spindles in each experiment). One-way ANOVA with Tukey's multiple comparisons test was used to obtain a P value. Download figure Download PowerPoint Next, time-lapse fluorescence microscopy revealed localization patterns of dynein during mitosis by tracking endogenous dynein heavy chain 1 (DHC1) tagged with 3X–mAID–mClover. In control cells with centrosomes, DHC1 initially showed a kinetochore-like distribution and was rapidly recruited into the spindle poles (Appendix Fig S5A). On the other hand, in acentrosomal cells, DHC1 showed a kinetochore-like distribution before being redistributed into the proximity of NuMA structures in the center of the condensed chromosome ring (Appen
Referência(s)