Local actin nucleation tunes centrosomal microtubule nucleation during passage through mitosis
2019; Springer Nature; Volume: 38; Issue: 11 Linguagem: Inglês
10.15252/embj.201899843
ISSN1460-2075
AutoresFrancesca Farina, Nitya Ramkumar, Louise C. Brown, Dureen Samandar Eweis, Jannis Anstatt, Thomas Waring, Jessica Bithell, Giorgio Scita, Manuel Théry, Laurent Blanchoin, Tobias Zech, Buzz Baum,
Tópico(s)Advanced Fluorescence Microscopy Techniques
ResumoArticle23 April 2019Open Access Transparent process Local actin nucleation tunes centrosomal microtubule nucleation during passage through mitosis Francesca Farina Francesca Farina MRC-LMCB, UCL, London, UK IPLS, UCL, London, UK IFOM, the FIRC Institute of Molecular Oncology, University of Milan, Milan, Italy University of Grenoble, Grenoble, France Search for more papers by this author Nitya Ramkumar Corresponding Author Nitya Ramkumar [email protected] orcid.org/0000-0002-4086-4562 MRC-LMCB, UCL, London, UK IPLS, UCL, London, UK Search for more papers by this author Louise Brown Louise Brown Institute of Translational Medicine, Cellular and Molecular Physiology, University of Liverpool, Liverpool, UK Search for more papers by this author Dureen Samandar Eweis Dureen Samandar Eweis MRC-LMCB, UCL, London, UK IPLS, UCL, London, UKCorrection added on 3 June 2019 after first online publication: Author name was corrected from Durren Samander-Eweis to Dureen Samandar Eweis Search for more papers by this author Jannis Anstatt Jannis Anstatt MRC-LMCB, UCL, London, UK IPLS, UCL, London, UK Search for more papers by this author Thomas Waring Thomas Waring Institute of Translational Medicine, Cellular and Molecular Physiology, University of Liverpool, Liverpool, UK Search for more papers by this author Jessica Bithell Jessica Bithell Institute of Translational Medicine, Cellular and Molecular Physiology, University of Liverpool, Liverpool, UK Search for more papers by this author Giorgio Scita Giorgio Scita orcid.org/0000-0001-7984-1889 IFOM, the FIRC Institute of Molecular Oncology, University of Milan, Milan, Italy Department of Oncology and Hemato-Oncology, University of Milan, Milan, Italy Search for more papers by this author Manuel Thery Manuel Thery orcid.org/0000-0002-9968-1779 Hospital Saint-Louis, Paris, France Search for more papers by this author Laurent Blanchoin Laurent Blanchoin orcid.org/0000-0001-8146-9254 University of Grenoble, Grenoble, France Search for more papers by this author Tobias Zech Tobias Zech Institute of Translational Medicine, Cellular and Molecular Physiology, University of Liverpool, Liverpool, UK Search for more papers by this author Buzz Baum Corresponding Author Buzz Baum [email protected] MRC-LMCB, UCL, London, UK IPLS, UCL, London, UK Search for more papers by this author Francesca Farina Francesca Farina MRC-LMCB, UCL, London, UK IPLS, UCL, London, UK IFOM, the FIRC Institute of Molecular Oncology, University of Milan, Milan, Italy University of Grenoble, Grenoble, France Search for more papers by this author Nitya Ramkumar Corresponding Author Nitya Ramkumar [email protected] orcid.org/0000-0002-4086-4562 MRC-LMCB, UCL, London, UK IPLS, UCL, London, UK Search for more papers by this author Louise Brown Louise Brown Institute of Translational Medicine, Cellular and Molecular Physiology, University of Liverpool, Liverpool, UK Search for more papers by this author Dureen Samandar Eweis Dureen Samandar Eweis MRC-LMCB, UCL, London, UK IPLS, UCL, London, UKCorrection added on 3 June 2019 after first online publication: Author name was corrected from Durren Samander-Eweis to Dureen Samandar Eweis Search for more papers by this author Jannis Anstatt Jannis Anstatt MRC-LMCB, UCL, London, UK IPLS, UCL, London, UK Search for more papers by this author Thomas Waring Thomas Waring Institute of Translational Medicine, Cellular and Molecular Physiology, University of Liverpool, Liverpool, UK Search for more papers by this author Jessica Bithell Jessica Bithell Institute of Translational Medicine, Cellular and Molecular Physiology, University of Liverpool, Liverpool, UK Search for more papers by this author Giorgio Scita Giorgio Scita orcid.org/0000-0001-7984-1889 IFOM, the FIRC Institute of Molecular Oncology, University of Milan, Milan, Italy Department of Oncology and Hemato-Oncology, University of Milan, Milan, Italy Search for more papers by this author Manuel Thery Manuel Thery orcid.org/0000-0002-9968-1779 Hospital Saint-Louis, Paris, France Search for more papers by this author Laurent Blanchoin Laurent Blanchoin orcid.org/0000-0001-8146-9254 University of Grenoble, Grenoble, France Search for more papers by this author Tobias Zech Tobias Zech Institute of Translational Medicine, Cellular and Molecular Physiology, University of Liverpool, Liverpool, UK Search for more papers by this author Buzz Baum Corresponding Author Buzz Baum [email protected] MRC-LMCB, UCL, London, UK IPLS, UCL, London, UK Search for more papers by this author Author Information Francesca Farina1,2,3,4,‡, Nitya Ramkumar *,1,2,‡, Louise Brown5, Dureen Samandar Eweis1,2, Jannis Anstatt1,2, Thomas Waring5, Jessica Bithell5, Giorgio Scita3,6, Manuel Thery7, Laurent Blanchoin4, Tobias Zech5 and Buzz Baum *,1,2 1MRC-LMCB, UCL, London, UK 2IPLS, UCL, London, UK 3IFOM, the FIRC Institute of Molecular Oncology, University of Milan, Milan, Italy 4University of Grenoble, Grenoble, France 5Institute of Translational Medicine, Cellular and Molecular Physiology, University of Liverpool, Liverpool, UK 6Department of Oncology and Hemato-Oncology, University of Milan, Milan, Italy 7Hospital Saint-Louis, Paris, France ‡These authors contributed equally to this work *Corresponding author. Tel: +44 20 7679 3040; E-mail: [email protected] *Corresponding author. Tel: +44 20 7679 3040; E-mail: [email protected] The EMBO Journal (2019)38:e99843https://doi.org/10.15252/embj.201899843 See also: D Inoue et al (June 2019) PDFDownload PDF of article text and main figures. 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 Cells going through mitosis undergo precisely timed changes in cell shape and organisation, which serve to ensure the fair partitioning of cellular components into the two daughter cells. These structural changes are driven by changes in actin filament and microtubule dynamics and organisation. While most evidence suggests that the two cytoskeletal systems are remodelled in parallel during mitosis, recent work in interphase cells has implicated the centrosome in both microtubule and actin nucleation, suggesting the potential for regulatory crosstalk between the two systems. Here, by using both in vitro and in vivo assays to study centrosomal actin nucleation as cells pass through mitosis, we show that mitotic exit is accompanied by a burst in cytoplasmic actin filament formation that depends on WASH and the Arp2/3 complex. This leads to the accumulation of actin around centrosomes as cells enter anaphase and to a corresponding reduction in the density of centrosomal microtubules. Taken together, these data suggest that the mitotic regulation of centrosomal WASH and the Arp2/3 complex controls local actin nucleation, which may function to tune the levels of centrosomal microtubules during passage through mitosis. Synopsis Centrosomes, the main interphase microtubule-organizing centers, were recently found to also nucleate actin filaments. Here, local actin nucleation is observed in anaphase, tuning centrosomal microtubule nucleation as cells passage through mitosis. Centrosomes function as both actin- and microtubule-organizing centers during mitotic exit. Actin transiently accumulates around centrosomes at early anaphase. In vitro reconstitution assays with isolated centrosomes show a burst in actin filament formation at early anaphase. Anaphase actin filament formation depends on Arp2/3 and WASH complex activity. The anaphase actin burst around centrosomes is accompanied by reduction in microtubule density suggesting cross-talk between the two systems. Introduction The microtubule (Zhai et al, 1996; Meraldi & Nigg, 2002) and actin cytoskeletons (Ramkumar & Baum, 2016) undergo profound parallel changes in dynamics and organisation as cells go through mitosis. These changes play a vital role in the control of animal cell division and begin as cells enter prophase. At this time, the interphase microtubule cytoskeleton is disassembled (Centonze & Borisy, 1990; Niethammer et al, 2007; Mchedlishvili et al, 2018), allowing microtubule nucleation to become focused at centrosomes (Zhai et al, 1996; Piehl et al, 2004; Mchedlishvili et al, 2018), where gamma-tubulin accumulates (Khodjakov & Rieder, 1999; Bettencourt-Dias & Glover, 2007; Sulimenko et al, 2017). With the loss of the nuclear/cytoplasmic compartment barrier at the onset of prometaphase, this is followed by a sudden change in microtubule organisation (Mchedlishvili et al, 2018) and dynamics (Zhai et al, 1996). During prometaphase, the short, dynamic centrosomal microtubules that remain capture chromosomes (Mitchison & Kirschner, 1985) drive bipolar spindle formation (Magidson et al, 2011) and interact with the cortex to guide positioning of the mitotic spindle (McNally, 2013). The actin cytoskeleton also undergoes changes over the same period. These begin in prophase when the interphase actin cytoskeleton is disassembled (Matthews et al, 2012). This likely frees up a pool of actin monomers (Kaur et al, 2014), which is then used to assemble a thin (Clark et al, 2013), mechanically rigid (Fischer-Friedrich et al, 2015), cortical actomyosin network that drives mitotic rounding (Reinsch & Karsenti, 1994; Ragkousi & Gibson, 2014; Sorce et al, 2015). While the mechanisms underlying this mitotic switch in actin organisation are not well understood, the process likely involves the following: (i) the loss of interphase focal adhesions (Dix et al, 2018; Lock et al, 2018), (ii) the loss of Arp2/3-dependent lamellipodia (Ibarra et al, 2005; Bovellan et al, 2014; Rosa et al, 2015) and (iii) the activation of formins downstream of Ect2/Pbl and the GTPase Rho (Maddox & Burridge, 2003; Matthews et al, 2012; Rosa et al, 2015; Chugh et al, 2017). Interestingly, these parallel changes in actin and microtubule organisation appear to be largely independent of one another (Mchedlishvili et al, 2018). Thus, mitotic rounding is not much altered in cells entering mitosis without microtubules. Conversely, mitotic spindle assembly occurs with relatively normal kinetics in spherical cells that have been treated with latrunculin to remove their actomyosin cortex (Lancaster et al, 2013). Thus, during mitotic entry, the two systems appear to be independently regulated. However, this changes at anaphase, where the behaviour of the two filament systems is tightly coordinated for proper cell division. The signals emanating from the anaphase spindle polarises the overlying actomyosin cortex (Rappaport, 1996). This is achieved mainly through the activity of the centralspindlin complex (White & Glotzer, 2012), which binds overlapping microtubules at the midzone. This complex, in turn, recruits Ect2 (Yüce et al, 2005; Su et al, 2011), leading to Rho activation and assembly of a contractile actomyosin ring (Rappaport, 1996; Fededa & Gerlich, 2012), which drives cytokinesis (Wagner & Glotzer, 2016; Basant & Glotzer, 2018). At the same time, as the anaphase spindle elongates, signals associated with the anaphase chromatin appear to aid relaxation of the polar cortical actomyosin network (Salmon & Wolniak, 1990; Motegi et al, 2006; von Dassow, 2009; Ramkumar & Baum, 2016). This leads to the de-phosphorylation of ERM proteins, which crosslink actin to the plasma membrane (Rodrigues et al, 2015), to the loss of anillin (Kiyomitsu & Cheeseman, 2013) and to activation of SCAR/WAVE and the Arp2/3 complex at opposing cell poles (Zhang & Robinson, 2005; King et al, 2010; Nezis et al, 2010; Bastos et al, 2012; Luo et al, 2014). In some instances, the process of polar relaxation and cell re-spreading is sufficient to drive division in cells that lack an actomyosin ring (Neujahr et al, 1997; Dix et al, 2018). In studies looking at the role of the actin cytoskeleton in division, the mitotic cortical actomyosin network has been subject to most scrutiny. This is because cortical cytoskeleton controls animal cell shape and is by far the brightest actin-based structure visible under the microscope. However, two groups have reported the existence of dynamic cytoplasmic actin-based structures in dividing HeLa cells (Mitsushima et al, 2010; Field & Lénárt, 2011; Fink et al, 2011). While the precise function of this pool of cytoplasmic actin remains unclear, it has been reported to play a role in spindle assembly and positioning in various systems (Woolner et al, 2008; Sabino et al, 2015). In addition, cytoplasmic actin appears to work together with an unconventional Myosin, Myo19, to aid the partitioning of mitochondria at anaphase (Rohn et al, 2014). Here, building on a previous study that identified the WASH/Arp2/3-dependent nucleation of actin at centrosomes in interphase cells (Farina et al, 2016), we have re-examined the dynamics and potential function of non-cortical actin at mitotic exit. Using a combination of cell biology and biochemistry, we report the identification of a pool of WASH/Arp2/3-dependent cytoplasmic actin that is nucleated around centrosomes in early anaphase, which appears to limit the nucleation of centrosomal microtubules. Results In order to explore the possibility that actin is nucleated at centrosomes during mitotic exit, as it is in interphase cells (Farina et al, 2016), we fixed a population of HeLa cells and examined the amount of F-actin (phalloidin) and microtubules in a region close to centrosomes at different cell cycle stages. This revealed an increase in the density of F-actin in a small region around the centrosomes during the passage from metaphase to early anaphase (Fig 1A and B). During this period, we observed no significant changes in the levels of non-centrosomal cytoplasmic actin (Fig EV1A and C). This increase in centrosomally associated actin was accompanied by a decrease in microtubules intensity in the same region (Figs 1A and EV1B). A similar increase in actin accumulation and a corresponding decrease in tubulin intensity were also observed around the centrosomes of early anaphase Jurkat cells, a T-lymphocyte cell line (Fig EV1E and F). To investigate the dynamics of actin during this period, we used a spinning disc confocal to image HeLa cells expressing Lifeact-GFP. We observed a dynamic pool of cytoplasmic actin in metaphase, as described previously (Mitsushima et al, 2010; Fink et al, 2011). At anaphase, this pool became concentrated sub-cortically around the opposing poles (Figs 1C and EV2, Movies EV1–EV3). In order to define the dynamics of actin accumulation around the centrosomes, we generated a stable HeLa cell line expressing RFP-Lifeact and GFP-tubulin and performed relatively high temporal resolution acquisition (Fig 1D). Shortly after anaphase onset, we observed an increase in the levels of actin around centrosomes (Fig 1D–F), while non-centrosomal cytoplasmic actin levels remained unchanged (Fig EV1A and D). This burst of actin filament formation was extremely transient, occurring within minutes of anaphase onset, and was over by the time the cytokinetic furrow became clearly visible (Figs 1C–F and EV2, Movies EV1–EV3). Further, we observed this transient accumulation of actin around centrosomes during anaphase using diverse actin reporters (siR-actin, Lifeact-GFP and RFP-Lifeact), albeit with different accumulation dynamics that we attribute to the nature of the probes and their differing actin-binding kinetics (Figs 1 and EV2). During the same period, the density of microtubules, measured as an integrated intensity in a small region around spindle poles, dropped (Fig 1D and G). Thus, this transient appearance of cytoplasmic actin close to centrosomes at mitotic exit is associated with a reduction in the density of centrosomal microtubules. Figure 1. Dynamics of actin and microtubule networks during mitotic exit Maximum projection (2 z-slices) of HeLa cells immunostained for F-actin (phalloidin), tubulin, pericentrin and DAPI at metaphase and early anaphase showing actin accumulation around centrosomes in early anaphase. Scale bar = 10 μm Quantification of F-actin (phalloidin) intensity around pericentrin-positive centrosomes in HeLa cells immunostained as in (A), showing the increase in F-actin around centrosomes at early anaphase. Mean actin metaphase = 1 ± 0.03996, n = 92; mean actin anaphase = 1.875 ± 0.08895, n = 121; Student's t-test, ****P < 0.0001. Time-lapse sequence from a representative HeLa cell expressing Lifeact-GFP transiting from metaphase to anaphase, showing accumulation of actin in the presumptive centrosomal region, red arrows. Images represent 4z-projection. Scale bar = 10 μm. Time-lapse sequence of a representative HeLa cell expressing GFP-alpha-tubulin and RFP-Lifeact transiting from metaphase to anaphase showing the changes in actin and microtubules. Images represent 4 z-projection of movies taken every 1 min. Bottom—higher magnification view of the centrosomes C1 and C2 from time lapse above showing a transient increase in actin around the centrosomes in early anaphase. Scale bar = 10 μm, 1 μm in zoom. T = 0 is one frame before anaphase onset. Dotted circle shows centrosome position. Integrated fluorescence intensity of GFP-alpha-tubulin and RFP-Lifeact around the centrosomes for a population of cells exiting mitosis, showing actin accumulation around the centrosomes and a simultaneous decrease in tubulin over time. N = 30 centrosomes (15 cells). T = 0 is one frame before anaphase onset. Graph shows mean ± SD, and values were normalised to intensity at T = 0. Quantification and comparison of actin (RFP-Lifeact) and tubulin (a-tubulin-GFP) fluorescence intensity around the centrosomes at metaphase (−4 to −1 min), early anaphase (2–5 min) and late anaphase (7–10 min) for a population of cells. Mean actin intensity at metaphase = 1.002 ± 0.0239, n = 97; early anaphase = 1.023 ± 0.03047, n = 101; late anaphase = 0.9703 ± 0.05004, n = 98. Tubulin intensity at metaphase = 1.008 ± 0.03356, n = 106; early anaphase = 0.9748 ± 0.04124, n = 116; late anaphase = 0.9276 ± 0.05382, n = 101; one-way ANOVA. Download figure Download PowerPoint Click here to expand this figure. Figure EV1. Actin dynamics during mitotic exit in fixed cells Schematic representation of metaphase and anaphase cell explaining the pools of actin measured in fixed cells. 1 and 2 refer to centrosomal actin, while regions 3 and 4 measure non-centrosomal or cytoplasmic actin. Quantification of tubulin intensity around pericentrin-positive centrosomes in HeLa cells immunostained as in (Fig 1A), showing the decrease in tubulin intensity around centrosomes at early anaphase. Mean tubulin metaphase = 1 ± 0.02363, n = 90; mean tubulin anaphase = 0.89 ± 0.02937, n = 120; Student's t-test, P = 0.006. Quantification of non-centrosomal actin during metaphase and anaphase in fixed HeLa cells (as in Fig 1A and B, Scheme in EV1A) shows that there is no significant increase in the amount of cytoplasmic actin during this period. Mean actin intensity at metaphase = 1.403 ± 0.086, n = 52; mean actin intensity at anaphase = 1.574 ± 0.08301, n = 70, P = 0.1567. All values were normalised to mean actin intensity around centrosomes at metaphase. Error bars indicate standard deviation. Quantification of non-centrosomal actin during metaphase and early anaphase in live HeLa cells (as in Fig 1D and E, Scheme EV1A) shows that there is no significant increase in the amount of non-centrosomal actin during this period. Mean actin intensity at metaphase = 0.9994 ± 0.003416, n = 108 and mean actin intensity at anaphase = 1.007 ± 0.004158, n = 120. P = 0.1561, Welch's t-test. All values were normalised to mean actin intensity around centrosomes at metaphase. Error bars indicate standard deviation. Maximum projection (2 z-slices) of Jurkat cells immunostained for F-actin (phalloidin), tubulin, pericentrin and DAPI at metaphase and early anaphase showing the actin accumulation around centrosomes in early anaphase. Scale bar = 10μm Quantification of F-actin (phalloidin) and tubulin intensity around pericentrin-positive centrosomes in Jurkat cells immunostained as in (D), showing the increase in F-actin around centrosomes at early anaphase with a decrease in tubulin. Mean actin metaphase = 1 ± 0.03351, n = 70; mean actin anaphase = 1.315 ± 0.04993, n = 79; mean tubulin metaphase = 1.006 ± 0.02017, n = 56; mean tubulin anaphase = 0.7905 ± 0.02144, n = 78, Student's t-test for both, ****P < 0.0001. Download figure Download PowerPoint Click here to expand this figure. Figure EV2. Actin dynamics during mitotic exit in live cells Representative images of time lapse of HeLa cells imaged with siR-actin, every 1 min. Red arrows point to actin in the presumptive centrosomal region. Note that with siR-actin, this pool of actin around centrosomes persists for a longer time. Representative images of time lapse of HeLa cells expressing Lifeact-GFP, imaged every 1 min. Red arrows point to presumptive centrosomal region. This reporter shows a more widespread accumulation of cytoplasmic actin, with frequent enrichment around one or both centrosomes. Stills from time lapse of HeLa cells expressing Lifeact-GFP, showing actin enrichment around both centrosomes. Download figure Download PowerPoint In order to better visualise centrosomal actin in vivo, and as a method by which to isolate centrosomes for the in vitro experiments (see below), we also carried out similar analysis during monopolar cytokinesis (Hu et al, 2008). For these experiments, we arrested HeLa cells in prometaphase using STLC (Mayer et al, 1999; DeBonis et al, 2004)—a treatment that inhibits the Eg5 motor to prevent the assembly of a bipolar spindle (Fig 2). Actin nucleation was then followed as these prometaphase-arrested cells were forced to exit mitosis through the addition of a Cdk1 inhibitor (RO-3306) (Hu et al, 2008). Importantly, under these conditions, the analysis of centrosomal actin is facilitated by the fact that the monopole contains both centrosomes and remains far from the cell cortex (Fig 2A)—even though many other aspects of cytokinesis appear similar (Karayel et al, 2018). In this experiment, we observed little centrosomal-associated actin in the cells arrested in prometaphase with STLC (Figs 2A and EV3, Movie EV4). However, within ~6 min of Cdk1 inhibitor addition we observed a burst of actin filament formation close to centrosomes (Figs 2A and EV3, Movie EV4). Strikingly, these actin filaments formed parallel bundles that lay in between the astral microtubules emanating from the large monopole present in these cells (Fig 2A and B). To confirm this finding, cells arrested in prometaphase and cells after forced exit were fixed and stained to visualise actin filaments (phalloidin). We observed a similar increase in actin around centrosomes during forced exit in multiple cell lines: in HeLa cells (Fig 2C–E), the Jurkat T-cell line (Fig EV4A and B) and the MAVER1 B-cell line (Fig EV4C and D). Further, this actin accumulation was accompanied by a reduction in the density of centrosomal microtubules (Fig 2E). Thus, cytoplasmic actin appears to transiently accumulate around centrosomes during mitotic exit in both monopolar and bipolar divisions, and this is accompanied by a local decrease in microtubule density. Figure 2. Actin dynamics during forced mitotic exit Stills from time lapse of HeLa cells expressing GFP-alpha-tubulin and RFP-Lifeact arrested at prometaphase with STLC (t = 0) and forced to exit mitosis with Cdk1 inhibition (RO-3306) imaged every 90 s. Scale bar = 10 μm and for zoom = 4 μm. n = 21 cells from four independent experiments. Quantification of actin around centrosomes when cells are forced to exit mitosis using Cdk1 inhibition as in (A), showing a similar accumulation of actin as observed in bipolar divisions. Graph shows mean actin accumulation normalised to t = 0, and errors bars indicate standard deviation. HeLa cells expressing GFP-centrin 1 arrested at prometaphase or forced mitotic exit (RO-3306 5′) and stained with phalloidin. Scale bar = 5 μm and for zoom = 2 μm. The level of actin around the centrosome in (C) was quantified and normalised relative to metaphase and shows an increase during forced mitotic exit. STLC arrest 1 ± 0.01317, n = 210; STLC+RO-5 min = 1.251 ± 0.02441, n = 144; unpaired t-test with Welch correction, ****P < 0.0001. Quantification of tubulin intensity around centrosomes during prometaphase-arrested cells or cells forced to exit with Cdk1 inhibitor. Tubulin intensity decreases during forced mitotic exit. STLC-DMSO = 1 ± 0.02407, n = 180; STLC-RO-3306 = 0.6516 ± 0.02337, n = 136. P < 0.0001, Welch's t-test. Download figure Download PowerPoint Click here to expand this figure. Figure EV3. Actin dynamics during forced exit in HeLa cells Stills from time lapse of HeLa cells expressing Lifeact-GFP and labelled with siR-tubulin during monopolar exit, imaged every 1.5 min, showing actin accumulation around centrosomes. High-resolution image of Lifeact-GFP localisation in cells arrested in prometaphase (+DMSO) or cell forced to exit with Cdk1 inhibitor, for 15 min, showing actin accumulation around the centrosomes. Download figure Download PowerPoint Click here to expand this figure. Figure EV4. Actin dynamics during forced exit in Jurkat and MAVER1 cell lines Maximum projection view of Jurkat cells immunostained for F-actin (phalloidin) and pericentrin arrested at prometaphase with STLC and forced to exit mitosis with RO-3306 (10 min) showing the increase in F-actin around centrosomes during forced exit. Scale bar = 5 μm and for zoom = 1 μm. The level of actin around the centrosome in (A) was quantified and normalised relative to metaphase and shows an increase during forced mitotic exit. STLC arrest = 1.000 ± 0.08018, N = 29; STLC+RO-5 min = 1.713 ± 0.2361, N = 29, Welch's t-test, P = 0.0072. Maximum projection image of MAVER1 cells arrested in prometaphase with STLC and forced to exit with STLC+RO-3306 addition (10 min), immunostained with pericentrin (for centrosomes) and phalloidin (F-actin), showing the increase in actin around the centrosomes during the forced exit. Scale bar = 5 μm Quantification for MAVER1 cells stained as above, showing the increase in actin during forced mitotic exit. STLC arrested = 1.000 ± 0.08336, N = 35; STLC+RO-5 min = 3.647 ± 0.3894, N = 26; Welch's t-test, ****P < 0.0001. Error bars indicate standard error of the mean. Download figure Download PowerPoint Previous work demonstrated that the actin formed at interphase centrosomes is nucleated by a local pool of Arp2/3 (Farina et al, 2016). To investigate the role of Arp2/3 complex in actin accumulation during mitotic exit, we treated cells with either DMSO or the Arp2/3 complex inhibitor, CK666, and determined the amount of centrosomal actin in fixed (Fig 3) and live cells (Fig EV5). While the DMSO control behaved as described above, the CK666 Arp2/3 inhibitor was effective in eliminating the formation of cytoplasmic actin, including the actin emanating from the centrosome (Fig 3A–D). This was the case in both regular bipolar mitosis (Fig 3A and B), forced monopolar exit (Fig 3C and D) and live monopolar exit (Fig EV5A and B). While the actin accumulation decreased in cells treated with the Arp2/3 inhibitor, interestingly we found that Arp2/3 inhibition prevented the reduction in the density of microtubules associated with centrosomes during normal mitotic exit (Figs 3E and EV6A) and in monopolar exit (fixed cells–Figs 3F and 6B; live cells—EV5C). This suggests that the pool of actin associated with the centrosomes may influence centrosomal microtubules in early anaphase. This could aid division, as we observed a small percentage of cells with spindle oscillations (data not shown). Figure 3. Arp2/3-dependent actin accumulation at the centrosome Maximum projection (2 z-slices) view of HeLa cells pre-treated with DMSO and 0.2 mM CK666 for 15 min during their mitotic exit showing that treatment with CK666 leads to reduced accumulation of actin around the centrosomes during anaphase. Scale bar = 10 μm. Quantification of actin around centrosomes for cells treated with DMSO or 0.2 mM CK666, showing the reduction in actin accumulation around the centrosomes following CK666 treatment. DMSO-metaphase = 1 ± 0.4325, n = 54; CK666-metaphase = 0.936 ± 0.4604, n = 43; DMSO-anaphase = 1.8 ± 0.9736, n = 76; CK666-anaphase = 1.087 ± 0.4597, n = 73; one-way ANOVA, P < 0.0001. Date pooled from three independent experiments. Z-projection of HeLa cells expressing GFP-centrin 1 pre-treated with DMSO or 0.2 mM CK666 during prometaphase arrest and forced mitotic exit and stained with phalloidin for F-actin. Quantification of the level of actin around the centrosome from (C), which shows the reduction in actin accumulation around centrosomes following CK666 pre-treatment. DMSO-STLC = 1 ± 0.02768, n = 99; CK666 STLC = 0.776 ± 0.02186, n = 87; DMSO-RO-3306 = 1.339 ± 0.03048, n = 127; CK666-RO-3306 = 0.7699 ± 0.02246, n = 70; one-way ANOVA, P < 0.0001. Quantification of tubulin around centrosomes for cells treated with DMSO or 0.2 mM CK666, showing the failure to reduce tubulin density around centrosomes following CK666 treatment during bipolar exit. DMSO-metaphase = 1 ± 0.1968, n = 54; DMSO-anaphase = 0.867 ± 0.2345, n = 78; CK666-metaphase = 0.7995 ± 0.1275, n = 43; CK666-anaphase = 0.7407 ± 0.1859, n = 74; one-way ANOVA, P < 0.0001. Data pooled from three independent experiments. Error bars indicated standard deviation. Quantification of tubulin around centrosomes for cells treated with DMSO or 0.2 m
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