Actin cytoskeleton dynamics and the cell division cycle

The International Journal of Biochemistry & Cell Biology 42 (2010) 1622–1633

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Review

Actin cytoskeleton dynamics and the cell division cycle Yi-Wen Heng, Cheng-Gee Koh ∗ School of Biological Sciences, Nanyang Technological University, 60 Nanyang Drive, Singapore 637551, Singapore

a r t i c l e

i n f o

Article history: Received 21 January 2010 Received in revised form 12 April 2010 Accepted 14 April 2010 Available online 20 April 2010 Keywords: Actin cytoskeleton Cell cycle Mitosis Rho GTPases Focal adhesion

a b s t r a c t The network of actin filaments is one of the crucial cytoskeletal structures contributing to the morphological framework of a cell and which participates in the dynamic regulation of cellular functions. In adherent cell types, cells adhere to the substratum during interphase and spread to assume their characteristic shape supported by the actin cytoskeleton. This actin cytoskeleton is reorganized during mitosis to form rounded cells with increased cortical rigidity. The actin cytoskeleton is re-established after mitosis, allowing cells to regain their extended shape and attachment to the substratum. The modulation of such drastic changes in cell shape in coordination with cell cycle progression suggests a tight regulatory interaction between cytoskeleton signalling, cell–cell/cell–matrix adhesions and mitotic events. Here, we review the contribution of the actin cytoskeleton to cell cycle progression with an emphasis on the effectors responsible for the regulation of the actin cytoskeleton and integration of their activities with the cell cycle machinery. © 2010 Elsevier Ltd. All rights reserved.

Contents 1. 2.

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Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1622 Actin cytoskeleton, myosin and the cell cycle . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1623 2.1. Actin cytoskeleton in cell cycle control . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1623 2.2. Actin, myosin and the regulation of the mitotic spindle . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1623 2.3. Septin, actin cytoskeleton, and the cell cycle . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1627 Rho GTPases, their regulators and the cell cycle . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1627 3.1. RhoA and partners in cytokinesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1627 3.2. RhoA and partners in other stages of mitosis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1628 3.3. Cdc42 and partners . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1629 3.4. Cyclin-dependent kinase and Rho GTPases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1629 Cell attachment and the cell cycle . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1629 4.1. Integrin signalling and the cell cycle . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1630 4.2. Cadherin signalling and the cell cycle . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1630 4.3. Focal adhesion proteins and the cell cycle . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1630 Conclusion and perspective . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1631 Acknowledgements . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1631 References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1631

1. Introduction

Abbreviations: APC, adenomatous polyposis coli; CDK1, cyclin-dependent kinase 1; ECM, extra cellular matrix; FAK, focal adhesion kinase; GAP, GTPase-activating protein; GEF, guanine nucleotide exchange factor; MLCK, myosin light chain kinase; PAK, p21-activate kinase; Plk1, polo-like kinase 1; ROCK, p160-Rho-associated coiled-coil-containing protein kinase. ∗ Corresponding author. Tel.: +65 63162854. E-mail address: [email protected] (C.-G. Koh). 1357-2725/$ – see front matter © 2010 Elsevier Ltd. All rights reserved. doi:10.1016/j.biocel.2010.04.007

The actin cytoskeleton of eukaryotic cells undergoes drastic changes and remodelling during cell division. Interphase cells usually contain an extensive actin network but this network is rapidly dismantled and rearranged when cells enter mitosis, giving mitotic cells their characteristic round shape. At the end of mitosis, actin rearranges at the cleavage furrows and forms part of the contractile ring, which is central to the process of cytokinesis. Another mitotic

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event in which the actin network plays an important role is the separation of centrosomes, which is dependent on the cortical flow of cortical actin and the myosin network. Disruption of the actin and myosin II networks by inhibitory drugs such as latrunculin, ROCK (p160-Rho-associated coiled-coil-containing protein kinase) inhibitor and myosin II RNA interference (RNAi) causes failure in centrosome separation and proper spindle assembly (Rosenblatt et al., 2004; Uzbekov et al., 2002). Thus, the regulation of the actin cytoskeleton and of cell cycle progression appears to be connected. However, the nature of their functional integration is not well understood. Here, we review the current state of knowledge concerning the regulatory links between these two activities in proliferating cells. The emphasis is on proteins known to regulate the actin cytoskeleton and are implicated in cell cycle control. Our discussion on the actin cytoskeleton will not be limited to the acto-myosin filament or filamentous actin (F-actin) but will also encompass proteins associated with the modulation of the actin cytoskeleton such as the Rho GTPases and their regulators. We will also discuss cell adhesions and their effect on the cell cycle.

2. Actin cytoskeleton, myosin and the cell cycle 2.1. Actin cytoskeleton in cell cycle control Actin is a highly conserved globular protein found in almost all eukaryotic cells. It forms cellular scaffold structures that provide cells with their shape, tension support, intracellular vesicular transport, cell attachment, adhesion properties and the ability to move. Apart from these well-studied mechanical functions, actin also plays a more subtle role in chemical signal transduction. It was once thought that the cell cycle machinery controls the state of actin organization within the cell via an “inside-out” signalling mechanism (Wang, 1991; Yamashiro et al., 1991). However, retrograde signalling where the state of actin organization within the cell controls cell cycle progression has proven to be important as well (Assoian and Zhu, 1997; Thery and Bornens, 2006). The significance of the actin cytoskeleton for cell cycle progression can be easily gleaned from experiments using drugs or chemicals that interfere with the actin filament in the cells. Depolymerization of actin filaments by toxins such as cytochalasin D and latrunculin B has been reported to delay progression of mitosis in primary cells and fission yeast, suggesting that an intact actin cytoskeleton may be required for the efficient onset of mitosis (Gachet et al., 2001; Lee and Song, 2007). A summary of the different drugs and chemicals and their effect on the actin cytoskeleton is listed in Table 1. While a morphogenesis checkpoint has been proposed in budding yeast which is activated in response to perturbation of the actin cytoskeleton leading to delays in chromosome segregation (McMillan et al., 1998), a similar actin regulated checkpoint control has not been established in mammalian cells. Apart from causing a delay in mitosis, disruption of actin filaments also leads to G1 arrest. This actin cytoskeleton dependent arrest has been linked to cyclin expression and cyclin-dependent kinase (CDK) activation (Reshetnikova et al., 2000). In a study in which disruption of the actin cytoskeleton was induced by the over-expression of cofilin, a member of the actin depolymerization factor (ADF)/cofilin family, more than 90% of H1299 lung carcinoma cells arrested at G1 phase of the cell cycle (Lee and Keng, 2005). Excessive polymerization of F-actin by a mutant WASP or the drug Jasplakinolide, which interferes with actin depolymerization, causes an increase in multinucleate cells suggesting a possible defect in cytokinesis (Moulding et al., 2007). Similarly, expression of mutant WASPI294T which mis-regulates the Arp2/3 complex and enhances F-actin polymerization, results in abnormal accumulation of F-actin around the mitotic chromo-

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somes and may possibly lead to the observed cytokinesis defects. These observations demonstrate actin’s involvement in cell cycle progression. Various proteins known to function in both the regulation of the actin cytoskeleton and the cell cycle progression are summarized in Table 2. Many of these proteins change their cellular localization at different phases of the cell cycle (Fig. 1 and Table 3). Recently, cortactin, an actin-binding protein, has been identified as an anchor between the centrosome and F-actin and is essential for F-actin driven centrosome separation during mitosis. The triply phosphorylated form (Tyr421-, Tyr466- and Tyr482-) of cortactin is found to be localized exclusively to the spindle poles during transition to anaphase. Truncated cortactin lacking its actin-binding domain inhibits centrosome separation (Wang et al., 2008). Interestingly, cortactin has also been identified as a substrate of CDK1 at serine 405 (Blethrow et al., 2008). Although the significance of this phosphorylation of cortactin during mitosis has not been examined, it raises the possibility of a link between CDK1 signalling and cortactin-mediated centrosomes separation during mitosis. Links between actin cytoskeleton and transcription control have also emerged. Disruption of the actin cytoskeleton during mitosis leads to changes in the G- to F-actin ratio and hence in transcription activities mediated by the myocardin-related transcription factor and serum response factor, MAL/SRF (Miralles et al., 2003). It is possible that cell cycle progression could be affected by the status of actin polymerization via MAL/SRF mediated transcription. A recent report shows that in human uterine leiomyosarcoma cells, downregulation of MAL leads to reduction of p21 CDK inhibitor (Kimura et al., 2010). However, work by Triesman’s group has shown that depletion of SRF or MAL affects cell spreading and adhesion without affecting cell proliferation or apoptosis (Medjkane et al., 2009). 2.2. Actin, myosin and the regulation of the mitotic spindle In mitosis, microtubules have hogged the limelight with their beautiful arrays and precisely choreographed functions in organising events from the establishment of the bipolar spindle to the capturing, alignment and accurate segregation of chromosomes. Most importantly, mitotic spindle assembly and chromosome segregation can be reconstituted in vitro in cell free extract. The studies on actin in mitosis have remained focused almost solely on its mechanical function during cytokinesis until recently, where actin’s role in the biogenesis of the mitotic spindle has gained increasing attention. Using different methods of interference with actin polymerization and actin-myosin at the cell cortex, two earlier papers clearly showed the requirement of myosin II and actin in centrosome separation in higher eukaryotic and mammalian cells (Rosenblatt et al., 2004; Uzbekov et al., 2002). Upon actin depolymerization with latrunculin treatment, centrosome separation is blocked and a proper spindle cannot be assembled (Uzbekov et al., 2002). Proper spindle assembly is also disrupted when myosin II is inhibited through the use of the ROCK inhibitor – Y26732, which prevents ROCK-mediated phosphorylation of myosin light chain phosphatase, eventually blocking myosin activity. A similar effect is seen in cells treated with blebbistatin which inhibits the ATPase activity of non-muscle myosin II (Rosenblatt et al., 2004). Silencing of non-muscle myosin II heavy or light chain using RNA interference also disrupts spindle formation. When the cortical flow of acto-myosin filaments is prevented by cross-linking the cell surface with lectins such as concanavalin A, centrosome separation and movement are impeded, which results in a lopsided spindle (Rosenblatt et al., 2004). More recent studies have illustrated a closer direct link between the actin filaments and the mitotic spindles where F-actin was found localized to the mitotic apparatus (Woolner et al., 2008; Yasuda et al., 2005). Additional reports have also suggested that the stability of the cortical actin network is crucial in establishing

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Fig. 1. Different localization of proteins at interphase and mitosis. HeLa cells were harvested at different stages of the cell cycle and immunostained with (A) Top panels: anti-␣-tubulin antibodies. Bottom panels: phalloidin for F-actin. (B) Non-muscle myosin II heavy chain antibodies (red), DAPI (blue). (C) Top panels: anti-phospho LIMK1 (T508)/LIMK2 (T505) antibodies (green). Bottom panels: Merged image with anti-␣-tubulin (red) and DAPI stain (blue). (D) Top panels: anti-phospho PAK1 (T423) antibodies (red). Bottom panels: merged image with anti-␣-tubulin (green) and DAPI (blue). All bars: 10 ␮m.

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Table 1 Summary of different reagents used to perturb the actin cytoskeleton and their effects on cell cycle progression. Drug

Target

Action

Phenotype(s) reported

Reference(s)

Clostridium difficile toxin B

RhoA, Rac1, Cdc42

G2 arrest

Ando et al. (2007)

Clostridium botulinum exoenzyme C3 transferase

RhoA, RhoB, RhoC

ROCK I, ROCK II

Slows down p21 degradation, at mitosis caused re-spreading of prometaphase cells Failure in centrosome separation

Olson et al. (1995), Yamamoto et al. (1993)

Y-27632-Rock inhibitor

ML-7, ML-9

MLCK

G1 arrest, affect spindle organization

2,3-Butanedione monoxime (BDM) Blebbistatin

Myosin

Glycosylates RhoA, Rac1 and Cdc42, thereby inactivating them. ADP-ribosylation on asparagine 41 in the effector binding domain of the GTPase Competitive binding with ATP to active site of ROCK I and ROCK II Competitive binding with ATP to active site of MLCK Myosin ATPase inhibitor

Deng et al. (2005), Bhadriraju and Hansen (2004) Forer et al. (2007), Bhadriraju and Hansen (2004), Huang et al. (1998) Rosenblatt et al. (2004)

Calyculin A

Protein phosphatases

CEP1347 WR-PAK18

PAK1

Cytochalasin D

Actin

Latrunculin A/B

Actin

Jasplakinolide

Actin

Myosin II

Binds to Myosin-ADP-Pi with high affinity and interferes with phosphate release process Binds to active site of PP1 and PP2A protein phosphatases Targets PAK1 ATP-binding site Binds SH3 domain of PIX, blocking PAK-PIX interaction Binds to the barbed, fast growing plus ends of microfilaments, inhibiting actin monomer assembly and disassembly Binds to actin monomers near their ATP-binding site, preventing actin polymerization Binds to F-actin, stabilizing and promoting actin polymerization

correct spindle orientation in mammalian cells. The activity of LIM Kinase-1 (LIMK1) was found to increase during mitosis resulting in the phosphorylation and inactivation of cofilin (Kaji et al., 2008). Knockdown of LIMK1 leads to more activation and mislocalization of cofilin, which in turn results in mis-orientation of the spindle. Similar defects can be induced by inhibition of actin polymerization using latrunculin A or by over-expression of a non-phophorylatable cofilin (S3A). In addition, phosphorylated LIM kinases, but not the unphosphorylated form, have been reported to colocalize and coimmunoprecipitate with ␥-tubulin during early stages of mitosis (Chakrabarti et al., 2007). During metaphase, LIMK1 localises to the centrosomes while LIMK2 associates mainly with the mitotic spindle (Sumi et al., 2006). These data suggest that the activity of LIM kinases may play an important role in the regulation of spindle activity during mitosis. Apart from the interactions between the astral microtubule and the cortical actin network, other sites of interaction between the spindle microtubules and acto-myosin filaments have long been reported (Maupin and Pollard, 1986; Wu et al., 1998). More recently, a myosin isoform Myo10, which binds to both actin and microtubules, has been found to localize to the poles of mitotic spindles in Xenopus embryos (Woolner et al., 2008). Knocking down Myo10 causes mitotic spindle defects which include fragmentation of the spindle poles and lengthening of the spindles. Dynamic F-actin cables are also found localized to the mitotic spindles and the spindle poles. The authors proposed that F-actin and Myo10 regulate spindle lengthening and shortening, respectively. While the F-actin-mediated spindle lengthening is independent of Myo10, spindle shortening induced by Myo10 requires F-actin. Thus F-actin and Myo10 have both overlapping and distinct roles in mitosis. These observations suggest that the actin and microtubule structures may act synergistically in the assembly and positioning of the mitotic apparatus.

G1 arrest, inhibits kinetochore-microtubule elongation Failure in centrosome separation

Rosenblatt et al. (2004)

Accelerate anaphase chromosome separation G1 arrest, aberrant spindle formation, delay in mitosis transition Mitosis delay, G1 arrest, inhibits kinetochore-microtubule elongation

Fabian et al. (2007)

Mitosis delay, G1 arrest, inhibits kinetochore-microtubule elongation, failure in centrosome separation

Rosenblatt et al. (2004), Uzbekov et al. (2002), Gachet et al. (2001), Lee and Song (2007)

Cytokinesis defect

Moulding et al. (2007)

Nheu et al. (2004) Gachet et al. (2001), Lee and Song (2007), Forer et al. (2007)

Besides spindle formation, actin and myosin can exert their effects at different stages of the cell division. For example, myosin light chain kinase (MLCK) has been shown to function in the phosphorylation of myosin II which is required for its bundling with actin for contractile ring formation and for actomyosin contractility forces that are necessary for cytokinesis (Mabuchi, 1986). MLCK activity also plays an important role in early mitotic events. Microinjection of the catalytic fragment of MLCK into prophase cells delays the transition from nuclear envelope breakdown to the onset of anaphase, but does not affect the duration between anaphase onset and mid cytokinesis. Unregulated MLCK activity is also found to cause reduced fluorescent staining of spindle microtubules in prometaphase and metaphase cells (Fishkind et al., 1991). Similarly, treatment of mouse eggs with MLCK inhibitor ML-7, or the auto-inhibitory Peptide 18, affects localization of actin cap on the metaphase II spindles and subsequent cortical reorganization activities (Deng et al., 2005). Although ROCK and myosin activities are required for the rounding up of cells during mitosis (Maddox and Burridge, 2003), the actual mechanism remains elusive. Additional evidence that the actin structures can affect cell morphology during mitosis came from the study of moesin. Moesin is a member of the ezrin/radixin/moesin (ERM) family of actin-binding proteins and has been implicated in cell rounding in mitosis. Phosphorylation of moesin by the Ste20-like protein kinase Slik in Drosophila S2 cells causes cell rounding in mitosis mainly via cross-linking actin to the membrane at the cortex to increase cortical rigidity (Carreno et al., 2008; Kunda et al., 2008). Knocking down of moesin and its upstream kinase Slik leads to defects in cell cortex organization as well as metaphase spindle stability. Increasing cell rigidity externally by using lectin to crosslink the cell membrane can partially rescue the moesin knockdown phenotype suggesting that

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Table 2 Summary of the role of different groups of proteins in actin cytoskeleton regulation and cell cycle progression. Protein Group

Protein

Roles in actin cytoskeleton regulation

Roles in cell cycle progression regulation

Reference(s)

Myosin II

Actin crosslinker; ATPase dependent acto-myosin force generation.

Rosenblatt et al. (2004), Uzbekov et al. (2002), Mabuchi (1986), Fabian et al. (2007)

Myosin X Anillin

Unconventional actin crosslinker. Actin and microtubule bundling.

Cortactin

Recruitment of Arp2/3 to F-actin.

Moesin

Actin binding; ERM protein.

Septin

RhoA

Actin binding; scaffold for non-muscle MyoII and its kinases. Stress fibres formation.

G1 -S progression; mitotic cell rounding; centrosome separation; mitotic spindle assembly; kinetochore-microtubule formation; anaphase chromosome movement; cytokinesis. Mitotic spindle shortening. Spindle associated cleavage specification. F-actin mediated centrosome separation. Mitotic cell rounding; mitotic cortical rigidity; metaphase spindle stability. Cytokinesis; chromosome congression and segregation at mitosis

Rac1

Lamellapodia formation.

G1 -S transition; cytokinesis.

Cdc42

Filopodia formation.

p190RhoGAP Ect2

RhoA GAP. RhoA GEF, Cdc42 GEF during mitosis.

G1 -S transition; kinetochore-microtubule stabilization; spindle biorientation; metaphase chromosome alignment; cytokinesis. Mitotic cell rounding. Contractile ring formation; contraction of contractile ring at cleavage furrow.

GEF-H1

RhoA GEF; microtubule dynamics. Rac1 and Cdc42 GAP; RhoA GAP during mitosis.

Actin binding

Rho GTPase

Rho GTPase Regulator

MgcRacGAP

Rho GTPase effector and down-stream target

Lfc MyoGEF

RhoA GEF. RhoA GEF; myosin II binding.

ROCK

RhoA effector; myosin regulation; actin bundling.

LIMK1 LIMK2 Cofilin

ROCK & PAK substrates; cofilin regulation; actin bundling. Downstream of RhoA pathway; actin severing. RhoA effector; actin nucleation and elongation.

mDia1

PRK2/PKN2 PAK1

RhoA and Rac effector. Rac and Cdc42 effector; cell motility; focal adhesion turnover.

N-WASP

Rac and Cdc42 effector; Arp2/3 activator; actin nucleation.

the maintenance of cell shape and rigidity is sufficient to stabilize the mitotic spindles. Moesin’s function in rounding up of cells appears to be independent of myosin II because cells expressing active T559D moesin but lacking myosin light chain are still able to round up during mitosis (Kunda et al., 2008). However, it is very likely that both moesin-actin and myosin-actin activities are required to establish the rounded cell shape and rigidity for proper spindle assembly and positioning. Actin and myosin structures also participate in generating the forces required for chromosome segregation. Treatment of crane-fly spermatocytes with actin depolymerization drugs such as cytochalasin D and lantrunculin A or myosin ATPase inhibitors like butanedione monoxime (BDM) causes inhibition of spindle microtubule elongation (Forer et al., 2007). Conversely, nonspecific inhibition of myosin light chain phosphatase by calyculin A

G1 -S transition; mitotic cell rounding; mitotic cortical rigidity; cytokinesis.

RhoA related cytokinesis. Formation of ingression furrow; interacts with Ect2. Mitotic spindle assembly. Localize RhoA and Ect2 to contractile ring. S phase progression, mitotic cell rounding; centrosome separation; mitotic spindle assembly; cytokinesis. Mitotic spindle orientation. Regulate cyclin A and p27 expression. G1 -S progression; mitotic spindle orientation. Spindle formation in early mitosis through an “Lfc-RhoA-mDia1” pathway. Mitosis entry; mitosis exit. G1 -S transition; G2 -M transition; centrosome maturation; regulation of Plk1 and Aurora-A activity; regulate astral microtubule dynamics; spindle orientation. Cytokinesis.

Woolner et al. (2008) Piekny and Glotzer (2008), Gregory et al. (2008) Wang et al. (2008) Carreno et al. (2008), Kunda et al. (2008) Joo et al. (2007), Spiliotis et al. (2005) Coleman et al. (2006), Nishimura and Yonemura (2006), Birkenfeld et al. (2007) Canman et al. (2008), Klein et al. (2009) Yasuda et al. (2004, 2006), Jaffe et al. (2008), Mitsushima et al. (2009) Maddox and Burridge (2003) Nishimura and Yonemura (2006), Oceguera-Yanez et al. (2005) Birkenfeld et al. (2007) Minoshima et al. (2003), Zhao and Fang (2005), Miller and Bement (2009) Bakal et al. (2005) Asiedu et al. (2009) Rosenblatt et al. (2004), Uzbekov et al. (2002), Croft and Olson (2006) Kaji et al. (2008) Croft and Olson (2006) Lee and Keng (2005), Kaji et al. (2008) Bakal et al. (2005)

Schmidt et al. (2007) Maroto et al. (2008), Zhao et al. (2005), Vadlamudi et al. (2000), Nheu et al. (2004), Balasenthil et al. (2004) Moulding et al. (2007)

results in the stabilization of myosin II phosphorylation and the subsequent acceleration of pole-ward movement of chromosomes during anaphase (Fabian et al., 2007). These studies strongly imply that the formation of mitotic spindle is highly dependent on the actin and myosin networks within the cell. Actin reorganization during mitosis helps create the cellular environment required for the mitotic spindle to serve its function. Disruption of the cortical actin architecture during mitosis can severely affect spindle orientation which may result in cell cycle arrest. There may well exist in mammalian cells, like in the budding yeast, a morphogenesis checkpoint which is dependent on the actin cytoskeleton (Lew, 2003) or a spindle orientation checkpoint (Gachet et al., 2006) which hinges on the interaction between the microtubule and cortical actin, although the existence of a spindle orientation checkpoint

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Table 3 The localization of different proteins linked to the actin cytoskeleton in interphase and mitosis. Protein

F-actin Myosin II Anillin Septin Cortactin Ect2 GEF-H1 LIMK1 LIMK2 MyoGEF PAK1 RhoA HEF1 Integrin-linked kinase (ILK) Focal adhesion kinase (FAK) Pyk2 Paxillin Zyxin

Subcellular localization by immunofluorescence staining Interphase (adherent cell type)

M-phase

Stress fibers; actin structures Stress fibers; actin structures Ubiquitous Stress fibres Ubiquitous, actin structures Ubiquitous Ubiquitous Cell–cell contacts Ubiquitous Ubiquitous Ubiquitous; actin structures; centrosomes Ubiquitous Focal adhesions Focal adhesions Focal adhesions Focal adhesions Focal adhesions Focal adhesions

Contractile ring Contractile ring Contractile ring Contractile ring Centrosomes (phosphorylated from) Central spindle Mitotic spindle; midzone Centrosomes; equatorial cortex; contractile ring Mitotic spindle, contractile ring Central spindle Centrosomes; contractile ring; mid-body Contractile ring Mitotic spindle; mid-body Centrosomes Centrosones Centrosomes Centrosomes Mitotic spindle; central spindle

in the fission yeast is now questionable (Meadows and Millar, 2008). 2.3. Septin, actin cytoskeleton, and the cell cycle Septins are a family of GTPase which can polymerize to form filamentous structures. They were originally discovered in the screen for cell division mutants in the budding yeast (Hartwell, 1971). Their main role is in the control of cytokinesis. A ring of septin polymers is assembled during early stages of the cell cycle at the bud neck and remains till cytokinesis (Cid et al., 2001). Apart from cytokinesis, septins have also been implicated in the regulation of GIN4 kinase activation required for bud growth (Carroll et al., 1998). Mammalian septins are found to associate with the plasma membrane, actin cytosketon and the microtubules (Spiliotis and Nelson, 2006). Mammalian Septin2 colocalizes with actin stress fibres during the interphase and the contractile ring at cytokinesis. It has also been shown to bind to non-muscle myosin II (Joo et al., 2007). In HeLa and MDCK cells, septins are localized to the metaphase plate during mitosis (Spiliotis et al., 2005). Knocking down of septins results in loss of chromosomes from the metaphase plate. It has been suggested that the septins form a scaffold at the midplane of mitotic spindle to maintain CENP-E motor protein at the kinetochores thereby facilitating the congression of chromosomes at the metaphase plate. Interestingly, Kremer et al. have shown that septins present in the cytoplasm bind to and act as a reservoir for SOCS7 (suppressor of cytokine signalling 7). Knocking down of septins 2, 6 and 7 causes loss of stress fibres and also nuclear accumulation of NCK (Kremer et al., 2007). SOCS7 is responsible for the import of NCK into the nucleus. DNA damages also result in the nuclear accumulation of SOCS7 and NCK. Hence septins are also linked to the DNA damage checkpoint via the septin-SOCS7-NCK pathway. Given that NCK is associated with the control of actin cytoskeleton, there is a possibility that cell cycle progression and actin cytoskeleton can influence each other via this pathway. 3. Rho GTPases, their regulators and the cell cycle The small GTPases belonging to the Rho family have long been associated with the regulation and remodelling of the actin cytoskeleton and are important for cell motility, morphogenesis and neurite development (Etienne-Manneville and Hall, 2002; Koh, 2006). Active RhoA causes the formation of stress fibres by

Reference(s)

Schroeder (1968) Mabuchi and Okuno (1977) Piekny and Glotzer (2008) Joo et al. (2007) Wang et al. (2008) Nishimura and Yonemura (2006) Birkenfeld et al. (2007) Sumi et al. (2006) Sumi et al. (2006) Asiedu et al. (2009) Zhao et al. (2005) Yuce et al. (2005) Law et al. (1998) Fielding et al. (2008) Rodriguez-Fernandez et al. (1999) Rodriguez-Fernandez et al. (1999) Herreros et al. (2000) Hirota et al. (2000)

increasing acto-myosin contraction and bundling of actin filament whereas active Cdc42 and Rac1 increase filopodia and lamellipodia formation, respectively. Apart from their roles in modulating the actin cytoskeleton, Rho GTPases have also been found to regulate cell cycle progression with two main sites of action: one at the G1 /S transition and the other during cytokinesis. For example, inhibition of Cdc42, Rac1 and RhoA results in cell cycle arrest at G1 phase of the cell cycle (Olson et al., 1995; Yamamoto et al., 1993). In human capillary endothelial cells, active RhoA causes an increase in the expression of the F-box containing protein Skp2 which is required for ubiquitinylation-dependent degradation of the CDK inhibitor p27kip1 (Mammoto et al., 2004). p27kip1 binds to and inactivates the cyclin D1/CDK4 and cyclin E/CDK2 complexes. Absence of active RhoA leads to high levels of p27kip1 , resulting in cell cycle arrest in G1 . Active RhoA acts via the balance of activities of its two effectors, ROCK and mDia, to activate the Skp2-p27kip1 pathway. It has also been shown that inhibition of RhoA or disruption of F-actin drastically slows down the degradation of another CDK inhibitor p21Waf/Cip1 (Coleman et al., 2006). Specifically, ROCK has been shown to regulate cyclin A expression via the Ras/MAPK pathway and via LIMK2 (Croft and Olson, 2006). More recently, a study using the Clostridium difficile toxin B to inhibit the Rho GTPases at the G2 phase of the cell cycle also reveals possible involvement of the Rho GTPases in the control of multiple signalling pathways involved in the progression to mitosis (Ando et al., 2007). The different pathways regulated by Rho GTPases at different phases of the cell cycle are summarized in Fig. 2.

3.1. RhoA and partners in cytokinesis Although RhoA activities have been reported to be required for cell cycle progression at different phases, most studies have concentrated on the role of RhoA in cytokinesis where the activity of RhoA is essential. RhoA organises the assembly of the contractile ring and induces the acto-myosin-driven constriction of the cleavage furrow [reviewed by Barr and Gruneberg, 2007]. ECT2, a guanine nucleotide exchange factor (GEF) which activates RhoA, localizes and activates RhoA at the cleavage furrow (Nishimura and Yonemura, 2006). GEF-H1, a microtubule-regulated GEF for RhoA, has also been shown to modulate RhoA activity during cytokinesis (Birkenfeld et al., 2007). These GEFs are, in turn, activated by CDK1/cyclin B and Aurora-A/B kinases (Birkenfeld et al., 2007; Hara et al., 2006; Niiya et al., 2006). Through the use of chemical genetics and specific inhibitors, Polo-like kinase Plk1, originally thought to

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Fig. 2. Functional connections between proteins involved in the regulation of Rho GTPases, the actin cytoskeleton and cell cycle progression. The activities of different proteins and how they can influence one another at different phases of the cell cycle are summarized.

regulate only spindle assembly, has been found to also control the localization of ECT2 to the central spindle and RhoA at the equator in anaphase (Brennan et al., 2007; Burkard et al., 2007; Petronczki et al., 2007). Inhibition of Plk1 blocks the interaction of ECT2 with the midzone anchor HsCyk4 and the assembly of the contractile ring, resulting in the inhibition of cytokinesis and the formation of binucleate cells. Recently, centrosome/spindle pole-associated protein (CSPP) has been found to target MyoGEF (Myosin II-interacting GEF) to the central spindle during anaphase. MyoGEF also interacts with ECT2. Knockdown of MyoGEF results in mislocalization of ECT2 and RhoA during cytokinesis (Asiedu et al., 2009). Centralspindlin which is localized to both the central spindle microtubules and the tips of astral microtubules near the equatorial cortex is reported to recruit ECT2 to the central spindle (Nishimura and Yonemura, 2006). Knockdown of centralspindlin component, MKLP1, causes failure of ECT2 to localize to the equatorial cell cortex (Yuce et al., 2005). Another Rho GTPase regulator which participates in the control of cytokinesis and contractile ring assembly is MgcRacGAP (also known as HsCyk4). MgcRacGAP is a GTPase-activating protein for Rac and Cdc42 but is converted to a GAP for RhoA when phophorylated by Aurora-B in M phase of the cell cycle (Minoshima et al., 2003). Silencing of MgcRacGAP by RNAi results in the loss of ingression of the cleavage furrow and hence failure of the cells to undergo cytokinesis (Zhao and Fang, 2005). MgcRacGAP interacts with ECT2 and therefore may exert its effect via ECT2. A more recent report proposes a model where the GAP activities of MgcRacGAP and GEF activity of ECT2 act concurrently to promote a flux of RhoA

activities, thereby maintaining a RhoA-rich zone at the cell equator (Miller and Bement, 2009). Anillin, a 124 kDa RhoA binding protein, has been shown to accumulate at the cleavage furrow in a RhoA dependent manner (Piekny and Glotzer, 2008). Anillin was first isolated as an actin binding and bundling protein in Drosophila embryo (Field and Alberts, 1995). It was later shown to interact with myosin regulatory light chain (MLC) and septin (Kinoshita et al., 2002; Oegema et al., 2000; Straight et al., 2005). A recent report reveals that in Drosophila the RhoGEF pebble controls the formation of a filamentous structure containing Rho1, anillin and septin at the cleavage furrow, as well as the interaction of this filamentous structure with the plasma membrane and microtubules (Hickson and O’Farrell, 2008). Interestingly, anillin also interacts with RacGAP50C, which is a spindle associated protein that specifies the cleavage site (Gregory et al., 2008). Depletion of anillin results in the loss of RacGAP50C from the equator of cleavage and the collapse of the cleavage furrow. Thus anillin acts as a linker between the contractile ring and the spindle. 3.2. RhoA and partners in other stages of mitosis Other than cytokinesis, RhoA activities are also required in other phases of the cell cycle. Microtubule-associated Rho GEF Lfc has been implicated in spindle formation in early mitosis in Rat2 fibroblast cells (Bakal et al., 2005). Lfc knockdown or microinjection of anti-Lfc antibody causes spindle assembly defects in the early stages of mitosis. Both active RhoA and the formin mDia1 can res-

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cue these defects, suggesting that an “Lfc-RhoA-mDia1” pathway is working in Rat2 cells for spindle formation during early mitosis. Additionally, via the use of novel covalent capturing method of kinase-specific phosphopeptides, mDia1 was found to contain an optimal CDK1 phosphorylation consensus sequence (Blethrow et al., 2008), suggesting a possible direct connection between CDK1 and mDia1 during mitosis. In addition to mDia1 and ROCK, another RhoA effector, PRK2/PKN2, has also been implicated in the onset of mitosis and the completion of cytokinesis (Schmidt et al., 2007). Silencing of PRK2 in HeLa S3 cells leads to accumulation of bi-nucleated cells and these cells also show delay in G2 /M progression. PRK2 contributes positively towards the phosphorylation of Cdc25B, leading to the activation of CDK1/cyclin B. These effectors appear to constitute a feedback loop as active CDK1/cyclinB can further phosphorylate PRK2 which can then participate in the abscission process during cytokinesis. 3.3. Cdc42 and partners The level of the guanosine 5 -triphosphate (GTP) bound form of Cdc42 changes during cell cycle progression. The abundance of Cdc42-GTP is low during pre-metaphase, peaks during metaphase and declines again at telophase. However, there is no change in Rac-GTP levels throughout the cell cycle (Oceguera-Yanez et al., 2005). The level of RhoA-GTP, on the other hand, peaks at telophase. Over-expression of dominant-negative ECT2 and MgcRacGAP as well as silencing of ECT2 by RNAi prevented the change in the level of Cdc42-GTP in mitosis (Oceguera-Yanez et al., 2005). These observations suggest that both ECT2 and MgcRacGAP can regulate both RhoA and Cdc42 at different stages of the cell cycle. More specifically, while ECT2 catalyses the formation of Cdc42-GTP at metaphase, MgcRacGAP increases the hydrolysis of Cdc42-GTP at prometaphase. The spatial and temporal function of Cdc42-GTP is required for the progression through mitosis. Reduction in the activity of Cdc42 by RNAi causes a delay in mitotic progression. When Cdc42 RNAi is combined with the knockdown of other Cdc42-like GTPases, a high proportion of the cells showed misalignment of chromosomes (Yasuda et al., 2006). It has been suggested that one of the major roles of Cdc42 in mitosis is to control biorientation and stabilization of the kinetochore-microtubule attachment via its association and activation of the formin protein mDia3 (Yasuda et al., 2004), rather than the modulation of the actin cytoskeleton. Another formin protein mDia2 is reported to have microtubule stabilizing activities, which is independent of its actin nucleation activity (Bartolini et al., 2008). mDia2 has been shown to bind to microtubules directly as well as to the microtubule tip proteins such as EB1 and APC. The actin cytoskeleton related role for mDia2 in the regulation of cell cycle has been attributed to the stabilization of the actin scaffold for the contractile ring during cytokinesis (Watanabe et al., 2008). The p21-activated kinases (PAKs), a family of serine-threonine kinases which are effectors of Cdc42 and Rac1, have been implicated in the control of G2 /M transition. It has been shown that PAK1 regulates Plk1 activity. Inhibition of PAK1 activity leads to a delay in G2 /M and aberrant spindle assembly which are also the phenotypes that result from Plk1 inactivation (Maroto et al., 2008). PAK1 has also been shown to activate Aurora-A kinase at the centrosome (Zhao et al., 2005). PAK1 is targeted to the centrosome by the PIX-GIT1 complex. Studies have also proposed that PAK1 localization to the centrosome during metaphase to anaphase transition requires its kinase activity as the expression of a kinase inhibitory domain, PAK1-KID, causes a drastic reduction in centrosomal targeting (Li et al., 2002). When PAK1 is activated at the centrosome, it dissociates from PIX-GIT1 and is able to phosphorylate and activate Aurora-A. Aurora-A activation is required for the maturation

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of the centrosome in the late G2 phase. PAK1 activity may also play an important role in the regulation of astral microtubule dynamics during mitosis since over-expression of active PAK1 resulted in multiple spindle orientations (Vadlamudi et al., 2000). Studies have also linked PAK1 signalling to G1 to S phase transition via the regulation of the cyclin D1 machinery. It has been shown that in Ras transformed NIH 3T3 cells, the addition of two distinct and specific PAK1-3 inhibitors, CEP-1347 and WR-PAK18 was able to block malignant growth by down-regulation of cyclin D1 (Nheu et al., 2004). Consistent with this, perturbation of PAK1 activity by PAK1-KID, or knockdown of PAK1 by siRNA resulted in a marked decrease in cyclin D1 expression (Balasenthil et al., 2004). So far, a direct linkage of Cdc42 or Rac1 activity to these activities of PAK has not been documented, even though a concomitant increase in Cdc42 and PAK1 activity has been reported (Oceguera-Yanez et al., 2005). Thus PAK1 may exhibit its function in a GTPase-dependent or independent pathway. Two recent papers have illustrated that Cdc42 is important in controlling spindle orientation in mitotic cells (Jaffe et al., 2008; Mitsushima et al., 2009). Deletion of Cdc42 did not affect cell polarity but instead caused mis-orientation of the spindle leading to inappropriate positioning of the apical surfaces after cell division (Jaffe et al., 2008). Mitsushima et al. (2009) went further to demonstrate that two independent pathways downstream of Cdc42 are involved in regulating spindle orientation: one involving Cdc42PAK2-␤PIX and the other involving phosphatidylinositol 3 kinase (PI3K). It appears that the two independent pathways collectively affect the phosphatidyl 3,4,5 triphosphate (PIP3) levels and the cortical actin structures (Mitsushima et al., 2009). Not much has been reported about Rac1 and its role in the control of the cell cycle. One recent report suggests that the inhibition of Rac by the CYK-4/MgcRacGAP of the centralspindlin complex is essential for cytokinesis in C. elegans (Canman et al., 2008). Depletion of Rac but not RhoA can rescue the cytokinesis defect of a CYK-4 GAP mutant. This work suggests a parallel inhibition of Rac and activation of RhoA during cytokinesis, most probably to prevent the activation of Arp2/3 complex by WASP or WAVE which is downstream of Rac. The net result is the prevention of the formation of other actin networks which may interfere with the contractile ring. 3.4. Cyclin-dependent kinase and Rho GTPases That the onset of mitosis requires activation of CDK1 and is accompanied by drastic rearrangement of the actin cytoskeleton leading to the rounding up of the cell, suggests an intimate regulatory connection between CDK1 and actin cytoskeleton. While the exact signalling pathways of CDK1 activity leading to mitotic cytoskeletal changes remain poorly understood, p190 Rho GTPaseactivating protein (GAP) has been proposed as a major downstream effector of CDK1 (Maddox and Burridge, 2003). Activated CDK1 phosphorylates p190RhoGAP, down regulating its activity and thus decreases GTP hydrolysis by RhoA. This triggers a signalling cascade through ROCK and MLC phosphatase, that regulates cytoskeleton rearrangement observed in mitosis (Amano et al., 1996; Maddox and Burridge, 2003). 4. Cell attachment and the cell cycle Attachment of cells to the extracellular matrix (ECM) or other cells has long been implicated in cell cycle regulation. During cell division, the cells undergo extensive cell shape changes to detach from and reattach to the ECM. While cell–matrix adhesions have been reported to reduce during mitosis, cell–cell adhesions via the desmosomes, tight junctions and zonulae adherentes in epithelial cells are maintained throughout cell divisions (Baker and Garrod,

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1993; Jinguji and Ishikawa, 1992; Reinsch and Karsenti, 1994). In mammalian cells, inter-dependence of cell attachment and cell cycle signalling can be attributed to integrin and cadherin signals (Pugacheva et al., 2006). Knocking-out or silencing of focal adhesion proteins vinculin, paxillin and the adaptor protein CRK has been shown to result in the fusion of daughter cells leading to binucleate cells, demonstrating that these proteins are essential for the completion of cytokinesis (Nagasaki et al., 2009; Shafikhani et al., 2008). A recent paper reported a positive correlation between ECM stiffness and progression into S phase of the cell cycle in mammary epithelial cells and vascular smooth muscle cells (Klein et al., 2009). Using hydrogel to simulate physiological stiffness, it has been shown that an increase in matrix stiffness results in selective integrin activation, leading to localization and activation of focal adhesion kinase (FAK) which eventually induces RAC1 activation and cyclin D1 expression. 4.1. Integrin signalling and the cell cycle Integrins associate with numerous proteins and localise at the focal contacts. Upon engagement with ECM, they activate proximal signalling proteins such as FAK, SRC and CAS family members, which in turn signal through several signalling cascades, e.g. the RAP and B-RAF pathway, the PI3K, RAC, AKT and PAK pathway, and the SHC, GRB2, RAS and RAF pathway. These three pathways converge to activate MEK and ERK kinases and together activate G1 -specific cyclins D and E (Pugacheva et al., 2006). In addition, integrin-activated FAK can directly phosphorylate transcription factor KLF8, leading to its nuclear translocation and the activation of cyclin D1 promoter (Zhao et al., 2003). Integrins also activate different GEFs, which lead to the activation of the Rho GTPases. However, it has been reported that focal adhesion signalling and cell spreading are dispensable for progression through the cell cycle as long as there is sufficient cyclin D1 (Margadant et al., 2007). Integrin mediated cell adhesion and the ECM can also control the orientation of the spindles and hence determine the spindle axis and the plane of cell division (Thery et al., 2005; Toyoshima and Nishida, 2007). ␤1-Integrin knockout results in random spindle orientation and a high incidence of binucleate cells (Aszodi et al., 2003; Lechler and Fuchs, 2005). The use of an inactive ␤1-integrin mutant reveals a role for integrin in bipolar spindle assembly and cytokinesis (Reverte et al., 2006). In addition to its role in focal adhesion regulation, integrin-linked kinase (ILK) has also been localized to the centrosome and is implicated in mitotic spindle assembly and chromosome segregation (Fielding et al., 2008). However, it is not clear if the roles of ILK at the focal adhesions and the centrosomes are dependent on each other. Integrin signalling may also serve as an important link between the small GTPase Rap1 and mitosis. The activity of Rap1 is regulated during mitosis. Inhibition of Rap1 is required for focal adhesion disassembly at the onset of mitosis whereas Rap1 activation is needed for cell spreading after mitosis (Dao et al., 2009). Rap1 has also been shown to activate many integrins (Bos, 2005; Caron, 2003). It is very likely that Rap1 modulates integrin signalling and thereby influences the cell shape changes which accompany mitosis. 4.2. Cadherin signalling and the cell cycle Cadherins are Ca2+ -dependent transmembrane proteins that participate in cell–cell adhesion. In stable adherens junctions, cadherin assembles with ␣-catenin, ␤-catenin and actin filaments to form stable quaternary complexes that limit cell growth via contact-inhibition. In the absence of cell–cell contacts, ␤-catenin is displaced from cadherin and translocates to the nucleus thereby activating cyclin D1 transcription (Yamada et al., 2005). Interest-

ingly, ␤-catenin has also been shown to have an unexpected role as a component of the inter-centrosomal linker and is essential for the establishment of bipolar spindle (Bahmanyar et al., 2008). Although ␤-catenin is thought to be the major effector of cadherin engagement for cell cycle regulation, ␣-catenin has also been reported to influence cell cycle regulation. ␣-catenin has been shown to bind to various actin-binding proteins such as ␣-actinin (Knudsen et al., 1995), vinculin (Hazan et al., 1997; Watabe-Uchida et al., 1998; Weiss et al., 1998), Ajuba (Marie et al., 2003), spectrin (Pradhan et al., 2001), ZO-1 (Itoh et al., 1997), formin (Kobielak et al., 2004) and afadin (Pokutta et al., 2002), suggesting these actin-binding proteins may play associative roles in the modulation of adherens junctions-mediated cell cycle control. Another component of the adherens junctions, p120 catenin, is also implicated in the regulation of mitosis. Loss of p120 catenin results in mitotic defects leading to extended M phase and binucleate cells (Perez-Moreno et al., 2008). This has been attributed to the abnormally high RhoA activity in the p120 catenin conditional-knockout cells. Additional evidence that cell–cell adhesion plays an important role in determining spindle orientation comes from the work of den Elzen et al. (2009). They have reported that E-Cadherin can provide cues to orient the mitotic spindle during symmetric cell divisions in mammalian epithelia (den Elzen et al., 2009). When dominant-negative E-Cadherin is introduced into MDCK cells, more than half of the cells showed mis-oriented spindles. Moreover, cortical APC (adenomatous polyposis coli) staining, which is consistently localized to the cell cortex at the apicolateral region, is lost in cells expressing dominant-negative E-Cadherin. Silencing of APC by siRNA abolishes junctional staining of APC and causes spindle misorientation, suggesting that APC may mediate cadherin signalling to orient the mitotic spindle. In another study using Drosophila neuroepithelial cells, symmetric mode of cell division can be converted into asymmetric division upon the disruption of the adherens junctions. The APC protein which is localized at the adherens junctions is involved in the maintenance of the symmetric mode of division. The APC and the microtubule-associated EB1 proteins function together to orient the mitotic spindle to provide the polarity cues for symmetric division (Lu et al., 2001). 4.3. Focal adhesion proteins and the cell cycle In adherent cell types, cell rounding upon entry into mitosis is accompanied by a reduction in the focal contacts and an increase in cortical rigidity. After cytokinesis, cells reattach to their substratum and re-establish cytoskeletal networks. The concerted changes in cell shape and adhesion as cells prepare to enter mitosis suggest intense cross-signalling events between focal contacts and cell cycle signalling. This is evident from the observation that focal contacts are more densely distributed in the proximity of signalling molecules such as protein kinase C (Liao and Jaken, 1993), tyrosine kinases (Hanks et al., 1992; Schaller et al., 1992) and tyrosine phosphatases (Serra-Pages et al., 1995; Shen et al., 1998). It is possible that post translational modifications play a major role in eliciting such drastic changes within a few minutes. Indeed, HEF1, FAK, actopaxin, paxillin and PAK are phosphorylated at mitosis-specific sites (Clarke et al., 2004; Law et al., 1998; Yamakita et al., 1999) during early M phase. In addition, proteins associated with focal adhesion contacts such as FAK (Rodriguez-Fernandez et al., 1999), Pyk2 (Rodriguez-Fernandez et al., 1999), paxillin (Herreros et al., 2000) and zyxin (Hirota et al., 2000) are also found to associate with the mitotic spindle or the microtubules-organizing centre during mitosis (Table 3). The dual roles focal adhesion proteins play in adhesion-mediated signalling and mitosis-associated events suggest a precise temporal control of molecular events ensuring that

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one set of events is completed before the next phase of the cell cycle is initiated. FAK serves as an important signalling protein at focal adhesion sites to mediate focal adhesion formation, cell migration and cell cycle progression. Over-expression of FAK has been associated with invasiveness in a variety of human tumours (Owens et al., 1995). In interphase, FAK mediates cyclin D1 and p21 CDK inhibitor expression downstream of integrin engagement. Over-expression of dominant-negative FAK causes G1 arrest in mouse fibroblasts (Zhao et al., 1998). During mitosis, FAK is targeted to the centrosome (Rodriguez-Fernandez et al., 1999), suggesting that FAK may regulate centrosome functions. This is consistent with the observation that deletion of FAK in endothelial cells leads to multiple centrosome formation, multi-polar and disorganized mitotic spindles and misalignment of chromosomes during metaphase (Park et al., 2009). At the end of mitosis, integrins function to reattach the cells to its substratum, followed by the establishment of a spread-out shape and cytoplasmic tension. Similar to cell rounding during mitosis, these processes are likely regulated by post translational modifications of the attachment proteins. Activation of the anaphase promoting complex/cyclosome (APC/C) during late mitosis may degrade proteins that target the HEF1, zyxin and other attachment proteins to mitotic structures, thereby allowing focal contacts to reform. However, the exact mechanism of focal contact re-establishment and reformation of stress fibers following cytokinesis is poorly understood. It remains unclear if the formation of focal contacts precedes cytoskeletal tension or vice versa. Cells treated with trypsin also show loss of focal contacts and stress fibers accompanied by cell rounding. However, when replated, these cells re-establish focal contacts and spreading. Mitotic cells, on the other hand, stay detached until the completion of mitotic events. It is likely that cells use distinct mechanisms to promote or inhibit cytoskeletal events at different stages of cell cycle.

5. Conclusion and perspective Despite the extensive evidence suggesting that the disruption of the actin cytoskeleton can lead to cellular arrest, many aspects of the regulatory relationship between cytoskeletal integrity and cell cycle progression remain to be elucidated. In particular, it is unclear if a checkpoint-like mechanism is responsible to ensure the coordination of these two sets of events. More detailed studies of the direct targets of some key cell cycle regulators such as CDK1, Plk1, Aurora kinases which are also actin cytoskeleton regulators should provide further insight into how cell cycle progression can regulate the actin cytoskeleton and vice versa. Cdc28, the CDK responsible for cell cycle progression in Saccharomyces cerevisiae and Candida albicans, has been reported to phosphorylate and regulate the activity of a Rho GEF and IQGAP1, respectively (Kono et al., 2008; Li et al., 2008). Since CDK1 alone is sufficient to drive cell division in mammalian cells (Santamaria et al., 2007), it is possible that some regulatory proteins implicated in maintaining the actin cytoskeleton could be direct targets of CDK1. Many possible CDK1(Cdc28) targets have been identified in S. cerevisiae (Ubersax et al., 2003) and in HeLa cells (Blethrow et al., 2008). A thorough analysis of the roles of these CDK1 targets could lead to the identification of possible regulators of the actin cytoskeleton. Other key effectors, such as polo kinase, Aurora-A and ubiquitin ligases that play important roles in mitotic progression, may also serve as critical regulatory links between the actin cytoskeleton and the mitotic machinery. A connection between Plk1 and the actin cytoskeleton regulators has been found in a screen for the Plk1 interactome in U2OS cells at different stages of the cell cycle (Lowery et al., 2007). The interactome represents partners for a particular protein, isolated through

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affinity chromatography and characterized by mass spectrometry. In this screen, ROCK2 (an effector of RhoA) was found to be an interacting protein of Plk1 as well as being its substrate. Plk1 can control the local activation of RhoA during cytokinesis and phosphorylate ROCK2 directly and stimulate ROCK2 activity (Lowery et al., 2007; Yoshida et al., 2006). It is very likely that CDK1 plays a central role in integrating the mechanisms regulating the actin cytoskeleton and mitosis. CDK1, through its activity on GEFs and GAPs, could affect the activities of RhoGTPases. These GTPases can further regulate their effector proteins such as PAK which in turn can regulate Plk1 and Aurora-A kinases to modulate spindle dynamics. The activities of the RhoGTPases and actin-myosin filaments have been shown to be essential for spindle formation and for cytokinesis. The same proteins may be required at different stages of the cell cycle and may function differently to ensure successful cell division (Fig. 2). A better understanding of the mutual regulatory relationship between the cell cycle and the actin cytoskeleton will have important implications for many diseases. This is especially so for cancer biology as many therapeutic interventions involve interference with cell division of cancer cells. Acknowledgements We thank the Academic Research Fund, MOE Tier 2 and the Biomedical Research Council, ASTAR Singapore for research funding. References Amano M, Ito M, Kimura K, Fukata Y, Chihara K, Nakano T, et al. Phosphorylation and activation of myosin by Rho-associated kinase (Rho-kinase). J Biol Chem 1996;271:20246–9. Ando Y, Yasuda S, Oceguera-Yanez F, Narumiya S. Inactivation of Rho GTPases with Clostridium difficile toxin B impairs centrosomal activation of Aurora-A in G2 /M transition of HeLa cells. Mol Biol Cell 2007;18:3752–63. Asiedu M, Wu D, Matsumura F, Wei Q. Centrosome/spindle pole-associated protein regulates cytokinesis via promoting the recruitment of MyoGEF to the central spindle. Mol Biol Cell 2009;20:1428–40. Assoian RK, Zhu X. Cell anchorage and the cytoskeleton as partners in growth factor dependent cell cycle progression. Curr Opin Cell Biol 1997;9:93–8. Aszodi A, Hunziker EB, Brakebusch C, Fassler R. Beta1 integrins regulate chondrocyte rotation, G1 progression, and cytokinesis. Genes Dev 2003;17:2465–79. Bahmanyar S, Kaplan DD, Deluca JG, Giddings Jr TH, O’Toole ET, Winey M, et al. beta-Catenin is a Nek2 substrate involved in centrosome separation. Genes Dev 2008;22:91–105. Bakal CJ, Finan D, LaRose J, Wells CD, Gish G, Kulkarni S, et al. The Rho GTP exchange factor Lfc promotes spindle assembly in early mitosis. Proc Natl Acad Sci USA 2005;102:9529–34. Baker J, Garrod D. Epithelial cells retain junctions during mitosis. J Cell Sci 1993;104(Pt 2):415–25. Balasenthil S, Sahin AA, Barnes CJ, Wang RA, Pestell RG, Vadlamudi RK, et al. p21-activated kinase-1 signaling mediates cyclin D1 expression in mammary epithelial and cancer cells. J Biol Chem 2004;279:1422–8. Barr FA, Gruneberg U. Cytokinesis: placing and making the final cut. Cell 2007;131:847–60. Bartolini F, Moseley JB, Schmoranzer J, Cassimeris L, Goode BL, Gundersen GG. The formin mDia2 stabilizes microtubules independently of its actin nucleation activity. J Cell Biol 2008;181:523–36. Bhadriraju K, Hansen LK. Extracellular matrix dependent myosin dynamics during G1-S phase cell cycle progression in hepatocytes. Exp Cell Res 2004;300:259–71. Birkenfeld J, Nalbant P, Bohl BP, Pertz O, Hahn KM, Bokoch GM. GEF-H1 modulates localized RhoA activation during cytokinesis under the control of mitotic kinases. Dev Cell 2007;12:699–712. Blethrow JD, Glavy JS, Morgan DO, Shokat KM. Covalent capture of kinase-specific phosphopeptides reveals Cdk1-cyclin B substrates. Proc Natl Acad Sci USA 2008;105:1442–7. Bos JL. Linking Rap to cell adhesion. Curr Opin Cell Biol 2005;17:123–8. Brennan IM, Peters U, Kapoor TM, Straight AF. Polo-like kinase controls vertebrate spindle elongation and cytokinesis. PLoS ONE 2007;2:e409. Burkard ME, Randall CL, Larochelle S, Zhang C, Shokat KM, Fisher RP, et al. Chemical genetics reveals the requirement for Polo-like kinase 1 activity in positioning RhoA and triggering cytokinesis in human cells. Proc Natl Acad Sci USA 2007;104:4383–8. Canman JC, Lewellyn L, Laband K, Smerdon SJ, Desai A, Bowerman B, et al. Inhibition of Rac by the GAP activity of centralspindlin is essential for cytokinesis. Science 2008;322:1543–6.

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