Displacement of p130Cas from focal adhesions links actomyosin contraction to cell migration.

2014. Published by The Company of Biologists Ltd | Journal of Cell Science (2014) 127, 3440–3450 doi:10.1242/jcs.143438

RESEARCH ARTICLE

Displacement of p130Cas from focal adhesions links actomyosin contraction to cell migration

ABSTRACT Cell adhesion complexes provide platforms where cell-generated forces are transmitted to the extracellular matrix (ECM). Tyrosine phosphorylation of focal adhesion proteins is crucial for cells to communicate with the extracellular environment. However, the mechanisms that transmit actin cytoskeletal motion to the extracellular environment to drive cell migration are poorly understood. We find that the movement of p130Cas (Cas, also known as BCAR1), a mechanosensor at focal adhesions, correlates with actin retrograde flow and depends upon actomyosin contraction and phosphorylation of the Cas substrate domain (CasSD). This indicates that CasSD phosphorylation underpins the physical link between Cas and the actin cytoskeleton. Fluorescence recovery after photobleaching (FRAP) experiments reveal that CasSD phosphorylation, as opposed to the association of Cas with Src, facilitates Cas displacement from adhesion complexes in migrating cells. Furthermore, the stabilization of Src–Cas binding and inhibition of myosin II, both of which sustain CasSD phosphorylation but mitigate Cas displacement from adhesion sites, retard cell migration. These results indicate that Cas promotes cell migration by linking actomyosin contractions to the adhesion complexes through a dynamic interaction with Src as well as through the phosphorylation-dependent association with the actin cytoskeleton. KEY WORDS: Actomyosin, Cell migration, Focal adhesion, FRAP, p130Cas, Src

INTRODUCTION

Cell migration plays important roles in both physiological and pathological processes; from normal tissue development to cardiovascular disorders and cancer metastasis (Ridley et al., 2003). Actomyosin interactions generate contractile forces, which are transmitted to the extracellular substrate through focal adhesions. This subsequently drives cell migration (Chen, 2008; Ridley et al., 2003). Adhesion complexes incorporate a wide variety of molecules, including cytoskeletal proteins, adapter 1

Mechanobiology Institute, National University of Singapore, 117411 Singapore. Department of Biological Sciences, National University of Singapore, 117411 Singapore. 3Immunology Frontier Research Center, Osaka University, Suita, Osaka, 565-0871 Japan. 4Laboratory for Mechanical Medicine, Locomotive Syndrome Research Institute, Nadogaya Hospital, Kashiwa, Chiba, 277-0032 Japan. *Present address: Immunology Frontier Research Center, Osaka University, Suita, Osaka 565-0871 Japan. 2

`

Author for correspondence ([email protected])

Received 26 September 2013; Accepted 29 May 2014

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proteins, kinases, phosphatases, small GTPases and their modulators (Geiger et al., 2001). Focal adhesions also provide platforms where extracellular cues are converted into intracellular signals (Schwartz, 2010; Zaidel-Bar et al., 2007). Integrin– extracellular-matrix (ECM) binding initiates the tyrosine phosphorylation of focal adhesion proteins such as paxillin, FAK (also known as PTK2) and p130Cas (Crk-associated substrate, hereafter referred to as Cas; also known as BCAR1), which then promotes actin remodeling, actomyosin contraction and focal adhesion turnover (Kawauchi et al., 2012; Meenderink et al., 2010; Parsons, 2003). Continuous turnover of focal adhesions is crucial for rapid cell migration (Webb et al., 2002). Individual focal adhesions in fibroblasts have a typical lifespan of ,30 min (Delorme-Walker et al., 2011; Kuo et al., 2011; Meenderink et al., 2010). Both the assembly and disassembly of focal adhesions are reportedly modulated by tyrosine phosphorylation of several focal adhesion proteins including FAK, paxillin and Cas (Geiger et al., 2001). This suggests that the regulatory mechanisms of focal adhesion turnover involve tyrosine phosphorylation. Actomyosin contraction plays a crucial role in the dynamic regulation of focal adhesions (Geiger et al., 2001), and actomyosin-derived forces modulate the dissociation of molecules from focal adhesions (Lele et al., 2006; Wolfenson et al., 2011) in conjunction with their retrograde fluxes (Brown et al., 2006; Hu et al., 2007; Guo and Wang, 2007). By contrast, tyrosine phosphorylation of focal adhesion proteins is enhanced when forces are exerted onto the adhesion sites (Pasapera et al., 2010). However, it remains unclear how these effects of actomyosin force on molecular dissociation and phosphorylation are coordinated to achieve the regulated dynamics of focal adhesions in migrating cells. Among those aforementioned focal adhesion proteins, Cas, which we have previously reported to link mechanical forces to tyrosine phosphorylation signaling (Sawada et al., 2006), is of particular interest. Through phosphorylation of its substrate domain (CasSD), Cas participates in the regulation of cell migration. This has been revealed by observations made in Cas-deficient fibroblasts expressing several different forms of Cas variants, including a phosphorylation-defective mutant (Meenderink et al., 2010). Upon Cas recruitment to adhesion sites, CasSD is phosphorylated by Src family kinases (SFKs) (Fonseca et al., 2004). In turn, CasSD phosphorylation regulates focal adhesion turnover and cell migration (Meenderink et al., 2010). Cas associates with focal adhesions through its N-terminal SH3 and C-terminal Src-binding domains (CasSB) (Nakamoto et al., 1997), which have been reported to bind directly to FAK and Src, respectively (Nakamoto et al., 1997; Polte and Hanks, 1995). Src therefore has dual roles on Cas at focal adhesions; firstly, as a

Journal of Cell Science

Hiroaki Machiyama1,2,*, Hiroaki Hirata1, Xia Kun Loh1,2, Madhu Mathi Kanchi1, Hideaki Fujita3, Song Hui Tan1, Keiko Kawauchi1 and Yasuhiro Sawada1,2,4,`

kinase, it phosphorylates the tyrosine residues in CasSD, and secondly, it acts as a scaffolding protein that tethers the Cas molecule to the adhesion complex. In this study, we find that Cas molecules located at adhesion sites move inward along with filamentous actin. Using a total internal reflection fluorescence (TIRF) microscope in fluorescence recovery after photobleaching (FRAP) experiments, we demonstrate that the displacement of Cas molecules from adhesion sites is accelerated by CasSD phosphorylation as well as by actomyosin contraction. Our TIRF-FRAP analyses also reveal that Cas displacement is hampered by its association with Src, which occurs through the Src-binding domain. Furthermore, stabilization of the Src–Cas association by genetic engineering retards focal adhesion turnover and cell migration without decreasing CasSD phosphorylation. Collectively, these findings suggest that Cas displacement at adhesion sites plays a significant role in linking myosin-mediated centripetal fluxes of the actin cytoskeleton to cell–substrate contacts. We propose that the displacement of Cas molecules from adhesion sites reconciles the discrepancy in dynamics between the two cellular components; one of which is the inwardly moving actin filaments, with the other being the stationary adhesion complex. This might constitute a mechanism by which Cas transmits actomyosin-derived forces to adhesion sites, thereby driving cell migration. RESULTS Actomyosin-driven inward movement of Cas from adhesion sites

To examine whether Cas molecules behave as a component that moves in relation to actin fibers, we first analyzed the correlation between the movement of Cas molecules at focal adhesions and the myosin-II-driven flow of actin filaments. To achieve this, we monitored the movement of Cas molecules and actin filaments simultaneously in the presence or absence of Y-27632, a ROCKspecific inhibitor that downregulates actomyosin contraction (Uehata et al., 1997). Because the effects of Y-27632 on cell behavior can vary depending on experimental conditions (Nakayama et al., 2005; Olivero and Furcht, 1993), we tested various concentrations (0–20 mM) of Y-27632. Consistent with previous reports describing the dissociation of adhesion complexes upon inhibition of actomyosin contraction (Balaban et al., 2001; Hirata et al., 2008), treatment with 10 mM or 20 mM Y-27632 disrupted the distinct assembly of focal adhesions and actin cytoskeletons (supplementary material Fig. S1A,B). However, the shape of the focal adhesions and their connection to actin cytoskeletons were retained at a lower concentration (2 mM or 5 mM) of Y-27632 (supplementary material Fig. S1A,B). In addition, the punctate distribution of Cas molecules and their phosphorylation were also preserved in the regions close to the leading edges of cells treated with 5 mM Y-27632 (supplementary material Fig. S2). Despite this, from the significant decrease in FAK phosphorylation (Fig. 1A), treatment with 5 mM Y-27632 appeared to compromise actomyosin contractility (Pasapera et al., 2010). We therefore examined the role of actomyosin contraction in the dynamics of Cas at focal adhesions using 5 mM Y-27632. When we viewed mCherry-tagged actin and GFP-tagged wildtype Cas (GFP–CasWT) expressed in migrating Cas-deficient fibroblasts using a TIRF microscope, we observed that GFP– CasWT colocalized with mCherry–actin in the leading lamellae (Fig. 1B). Time-lapse imaging demonstrated that GFP–CasWT

Journal of Cell Science (2014) 127, 3440–3450 doi:10.1242/jcs.143438

moved inward along with the flux of mCherry–actin (Fig. 1C). Both GFP–CasWT and mCherry–actin movements were slowed down by inhibiting actomyosin contraction by treating cells with Y-27632 (Fig. 1D–F; supplementary material Movies 1, 2). Furthermore, treatment with 5 mM Y-27632 hindered cell migration (Fig. 1G; supplementary material Fig. S1C) and extended the lifetime of focal adhesions (Fig. 1H; supplementary material Fig. S1D). Similar results were obtained when we treated cells with a low concentration (10 mM) of blebbistatin (supplementary material Fig. S3), a myosin II-specific inhibitor that also impedes actomyosin contraction (Kovaćs et al., 2004). These results suggest that the inward movement of Cas molecules correlates with an actomyosin-driven actin retrograde flux that promotes focal adhesion disassembly and cell migration (Gupton and Waterman-Storer, 2006). The inward flux of Cas is dependent on its phosphorylation

Because phosphorylation of the CasSD reportedly plays a role in focal adhesion disassembly and cell migration (Meenderink et al., 2010), we explored whether or not the movement of Cas molecules was dependent on their phosphorylation. When we viewed GFP–CasWT or its phosphorylation-defective mutant GFP–Cas15YF (Sawada et al., 2006) coexpressed with mCherrytagged actin in migrating Cas-deficient fibroblasts (Fig. 2A), we found that the movement of GFP–CasWT was significantly faster than that of GFP–Cas15YF (Fig. 2B,C; supplementary material Movies 3, 4). These results suggest that the inward movement of Cas molecules is dependent on CasSD phosphorylation. By contrast, the velocity of mCherry–actin retrograde flow was not significantly different between GFP–CasWT-expressing and GFP–Cas15YF-expressing cells (Fig. 2B,D; supplementary material Movies 3, 4). Considering that the actin retrograde flow in the leading lamellae is driven by actomyosin contraction (Fig. 1D,F; Ponti et al., 2004), CasSD phosphorylation does not appear to upregulate actomyosin contraction. Taken together with the observation that neither Y-27632 nor blebbistatin treatment decreased CasSD phosphorylation (Fig. 1A; supplementary material Fig. S3E), it seems that CasSD phosphorylation is independent of actomyosin contraction (Hotta et al., 2014), yet links Cas molecules to actomyosin-driven actin retrograde flux. FRAP analysis reveals that actomyosin contraction drives the displacement of Cas molecules from adhesion sites

Because we identified a correlation between the movement of Cas molecules and actomyosin-driven actin flux (Fig. 1D–F), we examined the mobility of Cas molecules at the sites of cell–ECM adhesion with reference to actomyosin activity. To this end, we conducted FRAP experiments using a TIRF microscope. Combining TIRF and FRAP allowed us to exclusively analyze the events on the lowest plane of adherent cells (,100 nm from the surface of the glass coverslips), where focal adhesion complexes are located (Kanchanawong et al., 2010), as well as to selectively photobleach fluorescent proteins fused to the Cas proteins localized at focal adhesions but not those in the cytosol. We monitored the recovery of fluorescence emitted from the fluorescent protein tagged to wild-type Cas at individual focal adhesions located in the leading lamellae of migrating cells during wound healing (Fig. 3A). The recovery of the fluorescence of GFP–CasWT was significantly impeded by Y-27632 treatment (Fig. 3A,B). When we calculated the mobile fraction (M) and half-recovery time (T1/2) from the fluorescence recovery curves (see Materials and Methods), we found that Y-27632 treatment 3441


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Fig. 1. Myosin II drives both actin retrograde flow and inward movement of Cas molecules at adhesion sites. (A) Cas-deficient fibroblasts expressing GFP–CasWT treated with or without Y-27632 (5 mM) for 1 h were solubilized with SDS sample buffer and subjected to SDS-PAGE followed by immunoblotting for Cas (aCas3), phospho-Cas (pCas-165), FAK, phospho-FAK (pFAK-397) and actin. (B) Cas-deficient fibroblasts coexpressing GFP–CasWT (left) and mCherry–actin (center) treated with 5 mM Y-27632 (lower panels) or vehicle control (water, upper panels) were viewed with a TIRF microscope. The right panels show the merged images of GFP–CasWT and mCherry–actin. Scale bar: 5 mm. (C) Time-lapse images of GFP–CasWT were merged with the average-intensityprojection images of mCherry–actin. A single cluster of GFP–CasWT moving inward was tracked and is indicated by the arrows. Scale bar: 2 mm. (D) Kymographs representing the movement of GFP–CasWT and mCherry–actin in the cells treated with 5 mM Y-27632 (lower panels) or its vehicle control (water, upper panels) were generated along the dashed lines in B. The arrows highlight the retrograde movements of Cas and actin clusters. Vertical scale bar: 2 mm. Horizontal scale bar: 5 min. (E,F) The rates of Cas (E) and actin (F) movement in the absence (control) or presence (Y-27632) of 5 mM Y-27632 were quantified from the kymographs in D. (G) Monolayers of Cas-deficient fibroblasts expressing GFP–CasWT treated with or without Y-27632 (5 mM) were subjected to a wound healing assay. The migration velocity was defined by the distance over which nuclei moved during recording. (H) Monolayers of Casdeficient fibroblasts coexpressing GFP–CasWT and mCherry–paxillin treated with or without Y-27632 (5 mM) were subjected to wound healing experiments and the lifetimes of individual focal adhesions were determined. All statistical analyses were performed between samples with and without 5 mM Y-27632. Data show the mean6s.e.m.; **P,0.01 (n.20 in E and F, n.40 in G, n.40 in H); Student’s t-test.

led to a significant increase in T1/2 as well as a significant decrease in M (Fig. 3C,D). These results suggest that the exchange of Cas molecules at adhesion sites is accelerated by actomyosin contractions. We then tested blebbistatin in the TIRFFRAP analysis (Fig. 3E–G), and we found that the blebbistatin treatment decreased M without significantly changing T1/2 (Fig. 3F,G). The decrease in M, which was observed in both Y27632-treated (Fig. 3C) and blebbistatin-treated (Fig. 3F) cells, is 3442

likely a consequence of actomyosin inhibition. By contrast, the difference in the effect on T1/2 between Y-27632 and blebbistatin treatments might be due to the Y-27632-induced downregulation of a wide range of ROCK targets. These not only include actomyosin contraction, which would be affected through the myosin regulatory light chain and myosin phosphatase, but also cell acidification (Bourguignon et al., 2004), focal adhesion maturation (Tsuruta and Jones, 2003) and modulation of various


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actin-binding proteins (Kimura et al., 1998; Fukata et al., 1999). Whereas T1/2 represents the rate of Cas molecule exchange between the adhesion site and the cytosol, which occurs quickly (over a timescale of ,20 seconds) (Fig. 3D,G), M represents the extent of occupancy of Cas molecules at focal adhesions (Fig. 3C,F) (Carisey et al., 2013). The results shown in Fig. 3B–G therefore suggest that myosin II activity is responsible for the displacement of ‘stably residing’ Cas molecules from focal adhesions but not for the fast exchange of Cas. Considering the rate of inward movement of Cas molecules (0.0860.01 mm/min, Fig. 1E) and the size of individual focal adhesions, which was measured along their long axes (typically 1–5 mm; Kim and Wirtz, 2013; Oakes et al., 2012; Wells et al., 2010), actomyosin-driven Cas displacement from focal adhesions appears to be an event with a timescale in the order of (tens of) minutes, but not seconds. Based on these notions, we analyzed the M value in our TIRF-FRAP experiments to further study the role of Cas displacement in regulating cell migration (Figs 4–6). CasSD phosphorylation facilitates the displacement of Cas molecules from adhesion complexes, whereas Src binding tethers Cas to adhesion sites

It has been reported that both CasSD phosphorylation and the association of Cas with Src are important for Cas to regulate focal adhesion disassembly and cell migration (Meenderink et al., 2010). We therefore examined the mobility of Cas molecules at adhesion complexes in migrating cells, with particular reference to CasSD phosphorylation and Src binding, by investigating the TIRF-FRAP dynamics of Cas variants in which either or both of these two elements were altered (Fig. 4A). We first tested whether displacement of Cas from adhesion sites was modulated by the phosphorylation status of CasSD. When compared with GFP–CasWT, the phosphorylation-defective

mutant GFP–Cas15YF showed remarkably slower fluorescence recovery (Fig. 4B) with a smaller mobile fraction (Fig. 4C). This indicates that the mobility of GFP–Cas15YF at focal adhesions is attenuated as compared with that of GFP–CasWT, and suggests that CasSD phosphorylation accelerates the displacement of Cas molecules from adhesion sites. We next examined the role of the Src–Cas association in the mobility of Cas molecules at focal adhesions. To preclude the effects of CasSD phosphorylation, we compared GFP–Cas15YF with GFP–Cas15YF/mPR, both of which were phosphorylation defective (Fig. 4D). mPR is a Cas mutation in which the prolinerich region of the C-terminal Src-binding domain in Cas is mutated (Fig. 4A), therefore the association with Src is attenuated in CasmPR (Ruest et al., 2001). We found that the mobile fraction of GFP–Cas15YF/mPR was larger than that of GFP–Cas15YF (Fig. 4C). This indicates that a reduction in Src–Cas binding by mPR mutation expedites the displacement of Cas molecules from focal adhesions independently of CasSD phosphorylation and, thus, that the Src–Cas association tethers Cas molecules to adhesion complexes. Fluorescence recovery was also compared between GFP– CasWT and GFP–CasmPR, but no significant difference was observed in the mobile fraction (Fig. 4C). This is probably due to the offset between two opposing elements in CasmPR, where dissociation from adhesion complexes was hampered by decreased CasSD phosphorylation (Fig. 4D), yet enhanced by attenuation of the Src–Cas interaction. Dual roles for Src, an enzyme that phosphorylates CasSD and tethers Cas molecules to adhesion sites

To examine the role of CasSD phosphorylation and Src–Cas association in Cas mobility at focal adhesions under different experimental settings, we used SYF cells, mouse embryonic 3443


Fig. 2. Cas moves inwardly, depending on CasSD phosphorylation. (A) Cas-deficient fibroblasts coexpressing GFP–Cas (wild-type or 15YF) (left) and mCherry-actin (center) were viewed with a TIRF microscope. The right panel shows the merged images of GFP–Cas and mCherry–actin. Scale bar: 5 mm. (B) Kymographs representing Cas (left) and actin (right) movement were generated along the dashed lines in A. The arrows highlight the retrograde movements of Cas and actin clusters. Vertical scale bar: 2 mm. Horizontal scale bar: 5 min. (C,D) The rates of GFP–Cas (C) and mCherry–actin (D) movement were quantified based on the kymographs in B. Statistical analyses were performed between the expression of GFP–CasWT and GFP– Cas15YF in Cas-deficient fibroblasts. Data show the mean6s.e.m.; **P,0.01 (n.20); Student’s t-test.

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Fig. 3. Actomyosin contraction involves the displacement of Cas molecules from adhesion sites. (A) Representative FRAPs of GFPtagged Cas molecules expressed in migrating Cas-deficient fibroblasts treated with (lower panels) or without (upper panels) 5 mM Y-27632 are presented as pseudocolor images before (Pre) and immediately after (0 s), or 15 s, 30 s, 40 s or 60 s after photobleaching. Scale bar: 1 mm. Pseudocolors correspond to the mean fluorescence intensities of individual focal adhesions normalized against their mean fluorescence intensity before photobleaching. (B) Mean fluorescence intensities I(t)/I0 of individual focal adhesions are plotted against time for GFP–CasWT with (diamond) and without (circle) 5 mM Y27632. (C,D) Mobile fraction (C) and half-recovery time (D) were obtained following equations 1 and 2, as described in Materials and Methods. (E) Trajectories of FRAP of mCherry– CasWT expressed in Cas-deficient fibroblasts with (diamond) or without (circle) 10 mM blebbistatin treatment are shown. Because blebbistatin is inactivated when exposed to the blue light that is needed to excite GFP (Sakamoto et al., 2005), we used Casdeficient fibroblasts expressing mCherry–CasWT, instead of GFP– CasWT, for this analysis. (F,G) Mobile fraction (F) and half-recovery times (G) were obtained from the results in E. All statistical analyses were performed between samples with and without 5 mM Y-27632 or 10 mM blebbistatin. Data show the mean6s.e.m.; **P,0.01 (n.40 in C,D,F,G); Student’s t-test.

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significantly hindered by the exogenous expression of Src (Fig. 5D,E). Furthermore, CGP77675 treatment did not significantly alter the effects of exogenous Src expression on the recovery of GFP–Cas15YF fluorescence (Fig. 5D,E). These findings support the notion that Src possesses dual roles in regulating the mobility of Cas molecules at focal adhesions. Stabilization of Src–Cas binding by genetic engineering retards cell migration

The mPR mutation in Cas that attenuated the Src–Cas association decreased CasSD phosphorylation (Fig. 4D), as well as the rates of focal adhesion turnover and cell migration (compare wild type and mPR in supplementary material Fig. S4). The decreased CasSD phosphorylation observed in CasmPR-expressing cells compared with that in CasWT-expressing cells (Fig. 4D) indicates that CasSD phosphorylation is dependent on the Src–Cas association. By contrast, there appeared to be a CasSD-phosphorylationindependent mechanism by which the Src–Cas association contributed to focal adhesion turnover and cell migration (compare 15YF and 15YF/mPR in supplementary material Fig. S4). Furthermore, the TIRF-FRAP analysis of Cas15YF and Cas15YF/mPR revealed a distinction between Src–Cas association


fibroblasts that do not express the major SFKs – c-Src, c-Yes and Fyn (Klinghoffer et al., 1999). As we reported previously (Sawada et al., 2006), CasSD phosphorylation was not detectable in SYF cells, but it was restored by the exogenous expression of Src (Fig. 5A). By contrast, when we conducted the TIRF-FRAP analysis using SYF cells, we found that the expression of exogenous Src did not significantly alter the recovery of GFP–CasWT fluorescence (Fig. 5B,C). This is probably because the expression of exogenous Src in SYF cells generated two opposing effects – CasSD phosphorylation and Src–Cas association. As observed from CasmPR expression in Cas-deficient fibroblasts (Fig. 4), these effects might offset each other in the regulation of Cas mobility. This notion is supported by the observation that treatment with the SFK inhibitor CGP77675 (Missbach et al., 1999), which reduced CasSD phosphorylation (Fig. 5A), impeded the recovery of GFP– CasWT fluorescence in Src-expressing SYF cells (Fig. 5B,C). To dissect the role of Src–Cas association from that of CasSD phosphorylation, we tested the phosphorylation-defective mutant Cas15YF, by expressing it in SYF cells with and without exogenous Src expression. When we conducted TIRF-FRAP analysis on GFP–Cas15YF, the fluorescence recovery was

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and CasSD phosphorylation in Cas displacement from adhesion sites (compare 15YF and 15YF/mPR in Fig. 4B,C). We therefore dissected the roles of the Src–Cas association in the regulation of Cas dynamics at adhesion sites during cell migration. To achieve this, we employed an approach different from the use of CasmPR, because experiments using the latter did not allow us to distinguish between the effects of Src–Cas association and those of CasSD phosphorylation. To this end, we induced stable Src–Cas binding by genetic engineering. We took advantage of the bimolecular fluorescence complementation (BiFC) technique based on stable complex formation by YFP moieties (Hu et al., 2002). For BiFC to occur, two fragments of a fluorescent protein, such as Venus, a modified YFP (Nagai et al., 2002), need to be brought together through the interaction of the proteins to which they are fused, but not through the affinity of the fluorescent protein fragments themselves (Hu et al., 2002). Once a BiFC complex is formed, dissociation of the two component proteins in the complex is hampered by the stable association of the fluorescent protein fragments (Kerppola, 2009). The ability of BiFC complexes to stabilize protein complexes can be used to investigate the functional consequences of stabilizing interactions between specific proteins (Kerppola, 2009). We fused the N-terminal (VN, amino acids 1–172) and Cterminal (VC, amino acids 155–238) fragments of Venus to the

C-termini of Src and mCherry–Cas, respectively, for our BiFC experiments (Fig. 6A, upper panel). This was because fusing any extra sequence to the N-terminus of Src prohibits its N-terminal myristoylation and normal function (Buss et al., 1986) and the Src-binding domain of Cas lies in its C-terminal region. When Src–VN and mCherry–Cas–VC were coexpressed in Casdeficient fibroblasts, Venus fluorescence was observed in a peripheral punctate pattern (Fig. 6B, upper panels), indicating that these molecules formed BiFC-based stable complexes at focal adhesions. By contrast, the combination of Src–VC and mCherry–Cas–VN (Fig. 6A, lower panels) did not appear to form BiFC complexes, as Venus fluorescence was not detectable (Fig. 6B, lower panels). This indicates that stable binding is not produced between Src–VC and mCherry–Cas–VN. Although the linker sequences between the Venus moieties (VN or VC) and Src or Cas are identical between these BiFC-positive and BiFCnegative combinations, association between the Venus moieties of the latter appears to be hindered by steric constraints (Kerppola, 2009). We confirmed that the Cas molecules with Venus moieties were phosphorylated and therefore still functional as Src substrates (Fig. 6C). To examine whether BiFC-based stable binding between Src and Cas affected the mobility of Cas molecules at adhesion sites, we conducted TIRF-FRAP 3445


Fig. 4. CasSD phosphorylation but not Src–Cas association facilitates dissociation of Cas molecules from focal adhesions. (A) Schematic representation of Cas and its mutant variants. GFP-tagged Cas variants used in this study were CasWT (wild-type full-length mouse Cas), Cas15YF (all fifteen YxxP motifs in the substrate domain are mutated to FxxPs), CasmPR (proline-rich motif RPLPSPP in the Src-binding domain is mutated to RAAASPP) and Cas15YF/mPR (combination of 15YF and mPR mutations). (B) Mean fluorescence intensities I(t)/I0 of individual focal adhesions are plotted against time for GFP–CasWT (circle), Cas15YF (square), CasmPR (triangle) and Cas15YF/mPR (inverted triangle). (C) The mobile fraction was determined by using equation 1, as described in Materials and Methods. Statistical analysis was performed between cells expressing CasWT (black bar) and other Cas variants or expressing Cas15YF and Cas15YF/mPR. Data show the mean6s.e.m.; **P,0.01 (n.40); Student’s t-test. (D) Expression and phosphorylation of GFP-tagged Cas molecules were analyzed by immunoblotting with antibodies against Cas (aCas3) , phospho-Cas (pCas-165) and actin. Note that the mutant Cas variants display either absent (15YF and 15YF/mPR) or significantly reduced (mPR) CasSD phosphorylation.

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Fig. 5. Src tethers Cas to adhesion complexes. (A) SYF cells, with or without the restoration of Src expression, were subjected to combinations of co-infection with retroviruses that mediated the expression of either GFP– CasWT or GFP–Cas15YF. Immunoblotting was conducted to analyze the expression and phosphorylation of Cas proteins (GFP–CasWT or GFP–Cas15YF) (upper two panels). The expression of exogenous Src and actin was also analyzed (lower two panels). Note the phospho-Cas (pCas-165) blot showing Srcdependent CasSD phosphorylation of GFP– CasWT (lane 2) and its reduction by treatment with CGP77675 (lane 3), neither of which was seen in cells expressing GFP–Cas15YF (lanes 5, 6). (B) FRAP analysis of GFP–CasWT expressed in SYF cells (circle, refer to lane 1 in A) and those expressing exogenous Src in the presence (square) or absence (diamond) of 10 mM CGP77675 (refer to lanes 3 and 2 in A, respectively). (C) The mobile fraction was determined from the results in B. (D) FRAP analysis of GFP–Cas15YF expressed in SYF cells (circle, refer to lane 4 in A) and those expressing exogenous Src in the presence (square) or absence (diamond) of 10 mM CGP77675 (refer to lanes 6 and 5 in A, respectively). (E) The mobile fraction was determined from the results in D. All statistical analyses were performed between the control condition [black bar, SYF cells expressing GFP– CasWT (C) or SYF cells expressing GFP– Cas15YF without exogenous Src expression and CGP77675 treatment (E)] and another condition. Data show the mean6s.e.m.; **P,0.01 (n.50); Student’s t-test.

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DISCUSSION Dissociation of Cas from Src at adhesion sites is a crucial step in the regulation of cell migration

We have demonstrated that both the Src–Cas association and CasSD phosphorylation are factors that contribute to the regulation of focal adhesion turnover and cell migration (supplementary material Fig. S4). Cas binding to Src through CasSB supports CasSD phosphorylation (Fig. 4D; Meenderink et al., 2010; Nakamoto et al., 1997), which facilitates focal adhesion turnover and cell migration (supplementary material Fig. S4). CasSD phosphorylation is known to influence regulatory proteins with downstream consequences on cell migration. For example, CasSD phosphorylation activates the Crk–DOCK180– Rac signaling pathway (Hsia et al., 2003), which, in turn, affects Pak1, a downstream effector protein of Rac. Pak1 has been shown to increase the rates of focal adhesion turnover and cell migration (Delorme-Walker et al., 2011). It is therefore possible that Src– Cas associations contribute to the acceleration of cell migration merely by enhancing CasSD phosphorylation. To address this, we used the BiFC technique and, interestingly, the comparison between the BiFC-positive and BiFC-negative combinations of Src and Cas reveals that interrupting Cas dissociation from Src at


experiments on mCherry-Cas molecules (Fig. 6D). The BiFCpositive combination of Src and mCherry–Cas gave a significantly smaller mobile fraction compared to the BiFCnegative combination (Fig. 6E). These results indicate that BiFCbased stabilization of Src–Cas binding impedes Cas displacement from adhesion complexes. The expression levels of exogenous Src molecules fused with Venus moieties (Src–VN and Src–VC) were much higher than that of endogenous Src in both BiFCpositive and BiFC-negative combinations (Fig. 6C). As such, the effects of endogenous Src on the behavior of Cas molecules (Cas–VC and Cas–VN) were presumably minor compared with those of exogenous Src. The BiFC technique was therefore instrumental in dissecting the role of Src–Cas association or dissociation from that of CasSD phosphorylation. We then analyzed the effects of stabilized Src–Cas binding on the rates of cell migration and focal adhesion turnover. Casdeficient fibroblasts expressing the BiFC-positive combination of Src and Cas molecules showed slower migration with a longer focal adhesion lifetime compared with the cells expressing the BiFC-negative combination (Fig. 6F,G), suggesting that dissociation from Src is vital for Cas to accelerate focal adhesion turnover and facilitate cell migration.

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focal adhesions decreases the rates of focal adhesion turnover and cell migration. This suggests that the dissociation of Cas from Src is a crucial step in the process regulating cell migration and demonstrates the importance of the Src–Cas association besides its effect on CasSD phosphorylation. Roles of actomyosin contraction in adhesion dynamics and cell migration

Our results show that actomyosin-driven displacement of Cas from adhesion sites accelerates cell migration. Consistently,

inhibition of actomyosin contraction retarded cell migration (Fig. 1G; supplementary material Fig. S3F; Hopkins et al., 2007; Ivkovic et al., 2012). However, there are other studies that report that myosin inhibition either does not decrease or actually increases cell migration speed (Kuo et al., 2011). Although actomyosin-based contractile force is crucial for the maturation of focal adhesions, contractility-dependent phosphorylation of FAK and paxillin facilitates the disassembly of focal adhesions (Pasapera et al., 2010; Webb et al., 2004). These opposing effects of actomyosin on the turnover dynamics of focal 3447


Fig. 6. Stabilizing the Src–Cas association at focal adhesions impedes cell migration. (A) Schematic representation of the BiFC-positive and BiFCnegative combinations. VN and VC, which indicate the N- and the C-terminal moieties of Venus, respectively, were fused with the C-termini of mCherry–Cas or Src. (B) DIC (left), TIRF for mCherry (middle) and TIRF for Venus (right) images of Cas-deficient fibroblasts expressing Cas and Src molecules of either BiFCpositive (upper panels) or BiFC-negative (lower panels) combination. Scale bar: 20 mm. (C) Cas-deficient fibroblasts expressing Cas and Src molecules of either the BiFC-positive or BiFC-negative combination were solubilized with SDS sample buffer and subjected to SDS-PAGE followed by immunoblotting with antibodies against Cas (aCas3), phospho-Cas (pCas-165), Src, GFP and actin. (D,E) TIRF-FRAP analysis of the N-terminal mCherry of Cas molecules was carried out for the BiFC-positive and BiFC-negative combinations. Trajectories of FRAP in BiFC-positive (circle) and BiFC-negative (square) combinations are shown (D). The mobile fraction (E) was determined from the results in D. (F) Monolayers of Cas-deficient fibroblasts expressing Cas and Src molecules of BiFCpositive or BiFC-negative combinations were subjected to wound healing experiments. Cell migration velocity was measured as in Fig. 1G. (G) Monolayers of Cas-deficient fibroblasts coexpressing Cas and Src molecules of the BiFC-positive or BiFC-negative combination together with mCherry–paxillin were subjected to wound healing experiments. The lifetimes of individual focal adhesions were calculated as in Fig. 1H. All statistical analyses were performed between the BiFC-positive and BiFC-negative combinations. Data show the mean6s.e.m.; **P,0.01 (n.40 in E and F, n.80 in G); Student’s t-test.

RESEARCH ARTICLE

adhesions potentially influence cell migration. Because there is a large variation in the reported lifetime of focal adhesions (Lee et al., 2010; Meenderink et al., 2010; Stricker et al., 2013), we speculate that the maturation and disassembly of focal adhesions might be modulated and coordinated by actomyosin contraction in a manner that differs between different cell types and/or under different conditions. In line with this idea, we found that treatment with a higher concentration (10 or 20 mM) of Y-27632 increased the cell migration rate (supplementary material Fig. S1C), suggesting that a difference in the extent of actomyosin inhibition shifts the regime of adhesion dynamics and cell migration. Cas displacement at adhesion sites – a link between inwardmoving actin filaments and stationary components of adhesion complexes

Based on the results and ideas described above, we propose a model of Cas displacement at adhesion sites (Fig. 7). In this model, unphosphorylated Cas molecules are distributed diffusely throughout the cytosol. When a Cas molecule is recruited to an adhesion complex, Src binds to the CasSB and tethers Cas to the adhesion complex. CasSD is subsequently phosphorylated, and this phosphorylation facilitates the formation of a molecular complex that links Cas molecules at focal adhesions to the actin cytoskeleton. The actin cytoskeleton continues to move inwards, leading to the displacement of Cas from the adhesion site. Actomyosin-based forces that are transmitted to extracellular substrates through focal adhesions drive cell migration (Chen, 2008; Ridley et al., 2003). Force transmission from the actin cytoskeleton, which is undergoing retrograde flow, to the stationary integrin–ECM complexes has been described as a ‘slippage clutch’ (Wang, 2007; Gardel et al., 2010), where the force is transmitted through dynamic transient connections between focal adhesion components (Brown et al., 2006; Hu et al., 2007). If CasSD phosphorylation facilitates the formation of a molecular complex that connects actin filaments to adhesion complexes as discussed above, it follows that such Cas complexes might be able to transmit the inward motion of the actin cytoskeleton to the adhesion complexes. In this case, Src–Cas binding might serve as a means for the transmission of forces from actin-associated Cas to Src-containing adhesion complexes. Meanwhile, the transient nature of this link would enable coupling of the actin cytoskeleton to ECM-adhesion sites


without abrogating actin movement. Because the dynamic actin-adhesion connection is crucial to allow cells to sense different substrate rigidities (Chan and Odde, 2008), Cas might be involved in the rigidity-sensing mechanism through the formation of the actin–Cas–Src link that is tuned by CasSD phosphorylation (Kostic and Sheetz, 2006). Further studies would be required to decipher the biophysical aspects of Cas in force transmission within the cytoskeleton–ECM link. MATERIALS AND METHODS Expression vectors

To construct retroviral expression vectors for GFP–CasWT and GFP– Cas15YF, we cloned the coding regions for wild-type Cas, Cas15YF (Shin et al., 2004) and monomeric EGFP (Murakoshi et al., 2008) into pBabehygromycin by PCR. The retroviral expression vectors for GFP–CasmPR and GFP–Cas15YF/mPR were constructed by site-directed mutagenesis using the vectors for GFP–CasWT and GFP–Cas15YF expression as templates, following a procedure described previously (Machiyama et al., 2009). All Cas mutants used in this study were cloned to the retroviral expression vector pBabe-hygromycin together with the coding sequence for mEGFP or mCherry. The retroviral expression vector for paxillin– mCherry, paxillin–GFP, c-Src and mCherry–actin were constructed by PCR using pBabe-blasticidin (for paxillin–mCherry, paxillin–GFP and cSrc) and pBabe-puromysin (for mCherry–actin) as parent vectors. To prepare the expression vectors used in the BiFC experiments, we first constructed mCherry-pBabe-hygromycin by PCR. The coding sequences for the N-terminal (amino acids 1–172) and the C-terminal (amino acids 155–238) fragments of Venus fused to the C-termini of wild-type Cas and c-Src were cloned by PCR using pcDNA3 as a parent vector and subcloned to the retroviral expression vectors mCherrypBabe-hygromycin and pBabe-puromycin, respectively. All PCR-based constructions were subject to DNA sequence confirmation. Cell culture and retroviral infection

Cas-deficient fibroblasts and SYF cells were used for microscopic analyses, and HEK293T cells were used for retroviral infection. The cells used in this study were cultured in Dulbecco’s Modified Eagle’s Medium (DMEM; Nissui Pharmaceutical, Tokyo, Japan) containing 10% fetal bovine serum (FBS; Gibco, Grand Island, NY) supplemented with 100 units of penicillin, 100 mg/ml streptomycin (Gibco), 2 mM L-glutamine (Gibco), and 0.18% sodium bicarbonate at 37 ˚C under 5% CO2. Retroviral infection was performed as described previously (Kawauchi et al., 2008). Infected cells were selected with 1.5 mg/ml puromycin, 300 mg/ml hygromycin or 5 mg/ml blasticidin for 2–3 days, depending upon the vector used. Antibodies

The antibodies used were against Cas (aCas3) (Sakai et al., 1994), phosphorylated (phospho)-Cas (Tyr165) (Cell Signaling Technology, Danvers, MA), FAK (Abcam, Cambridge, UK), phospho-FAK (Tyr397) (Millipore, Billerica, MA), Src (Millipore) and b-actin (Sigma-Aldrich, St Louis, MO).

Fig. 7. A model of Cas-mediated acceleration of cell migration – Cas serves as a molecular clutch at focal adhesions. Following CasSD phosphorylation at focal adhesions, displacement of Cas molecules links the inward motion of the actin cytoskeleton to adhesion complexes through Src– Cas association. This facilitates cell migration by transmitting actomyosin contraction to the ECM.

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A TIRF analysis system was set up on an IX81 inverted microscope (Olympus, Tokyo, Japan) equipped with an electron multiplying chargecoupled device camera with a 5126512-pixel chip (Evolve 512, Photometrics, Tucson, AZ) and fiber-coupled 488-nm and 559-nm lasers to excite GFP and mCherry, respectively. A 206NA 0.45 objective lens was used for the wound healing assay. A 606NA 1.45 objective lens was used in the Cas localization analysis (mCherry and Venus) and in the measurement of focal adhesion lifetime (mCherry). A 1006 NA 1.49 objective lens was used to detect GFP and mCherry fluorescence in the TIRF-FRAP experiments. Metamorph software (Molecular Devices, Sunnyvale, CA) was used for image acquisition, and ImageJ software (National Institutes of Health) was used for image processing and final figure preparation.


Microscope settings


To record the retrograde flow of actin and the inward movement of Cas simultaneously, we used a TIRF microscope (Ti-E, Nikon, Tokyo Japan) equipped with an electron multiplying charge-coupled device camera with a 5126512-pixel chip (iXon3 897, Andor Technology, Belfast, UK), 1006 NA 1.49 objective lens and fiber-coupled 488-nm and 559-nm lasers to excite GFP and mCherry. The TIRF images of GFP and mCherry were recorded every 10 s for 30 min. NIS elements software (Nikon) was used for image acquisition, and ImageJ software was used for image processing and final figure preparation.

Author contributions

Wound healing assay

Supplementary material

Cells were grown overnight in DMEM containing 10% FBS. A Chamlide magnetic chamber (Live Cell Instrument, Seoul, Korea) with a collagen (50 mg/ml)-coated glass bottom was used to form a monolayer that was scratched with a pipette tip and viewed with a microscope. Differential interference contrast (DIC) images were taken every 10 min for 8 h. Analysis of migration speed was performed using ImageJ software with a manual tracking plug-in. The displacement of the nuclei at each timepoint was plotted, and the migration rate was quantified as the slope of the plotted points.

Supplementary material available online at http://jcs.biologists.org/lookup/suppl/doi:10.1242/jcs.143438/-/DC1

Measurement of the lifetime of focal adhesion

Monolayers of Cas-deficient fibroblasts expressing mCherry-tagged paxillin were scratched 1 h before the initiation of observation, and mCherry–paxillin was monitored as a focal adhesion marker every 2 min for 4 h with TIRF microscopy. The lifetimes of individual focal adhesions were obtained by counting the number of sequential frames from the appearance of individual focal adhesions in the leading lamellae to their disappearance. Image acquisition and photobleaching in the TIRF-FRAP experiments

The prebleach images of GFP- or mCherry-tagged Cas molecules were acquired by illuminating them every 100 ms for 5 s with 6% of the maximum power of the 488-nm or 559-nm laser, respectively. Fluorophores were then photobleached by illuminating them with the maximum laser power for 10 s. The images of fluorescence recovery were acquired by illuminating the field every 100 ms for 60 s with 6% of the maximum laser power. Photobleaching of fluorophores between the prebleach and the postbleach acquisitions was negligible. TIRF-FRAP analysis

For TIRF-FRAP analysis, we measured the fluorescence from individual focal adhesions in the leading lamellae of a scratched monolayer and plotted the relative intensity against time. Time zero was set as the time when photobleaching was terminated. The plots were fitted by using equation 1, I ðtÞ=I0 ~M ½1{expð{t=tÞzc,

ð1Þ

where I(t) represents the mean intensity value of individual focal adhesions normalized against the mean intensity before photobleaching (I0). The mobile fraction (M), time constant (t) and constant (c) were used as fitting parameters. The half-recovery time T1/2 was calculated using equation 2; T1=2 ~lnð2Þ|t:

ð2Þ

All numerical data in this study are presented as the mean6s.e.m. Acknowledgements We thank Ryohei Yasuda (Max Planck Florida Institute, FL) and Hideji Murakoshi (National Institute for Physiological Science, Okazaki, Japan) for providing the mEGFP construct, and Steven Wolf (National University of Singapore) for assistance in manuscript preparation.

Competing interests The authors declare no competing interests.

H.M., H.H. and Y.S. conceived the project and designed the experiments; H.M. performed the experiments with assistance from X.K.L., M.M.K. and S.H.T.; H.M. analyzed the data; H.F. and K.K. provided the experimental materials and equipment; H.M., H.H. and Y.S. wrote the manuscript.

Funding This work was supported by the Biomedical Research Council [grant number R154-000-423-305]; and the Seed Fund [grant number R-714-004-007-271] from the Mechanobiology Institute, Singapore.

References Balaban, N. Q., Schwarz, U. S., Riveline, D., Goichberg, P., Tzur, G., Sabanay, I., Mahalu, D., Safran, S., Bershadsky, A., Addadi, L. et al. (2001). Force and focal adhesion assembly: a close relationship studied using elastic micropatterned substrates. Nat. Cell Biol. 3, 466-472. Bourguignon, L. Y., Singleton, P. A., Diedrich, F., Stern, R. and Gilad, E. (2004). CD44 interaction with Na+-H+ exchanger (NHE1) creates acidic microenvironments leading to hyaluronidase-2 and cathepsin B activation and breast tumor cell invasion. J. Biol. Chem. 279, 26991-27007. Brown, C. M., Hebert, B., Kolin, D. L., Zareno, J., Whitmore, L., Horwitz, A. R. and Wiseman, P. W. (2006). Probing the integrin-actin linkage using highresolution protein velocity mapping. J. Cell Sci. 119, 5204-5214. Buss, J. E., Kamps, M. P., Gould, K. and Sefton, B. M. (1986). The absence of myristic acid decreases membrane binding of p60src but does not affect tyrosine protein kinase activity. J. Virol. 58, 468-474. Carisey, A., Tsang, R., Greiner, A. M., Nijenhuis, N., Heath, N., Nazgiewicz, A., Kemkemer, R., Derby, B., Spatz, J. and Ballestrem, C. (2013). Vinculin regulates the recruitment and release of core focal adhesion proteins in a forcedependent manner. Curr. Biol. 23, 271-281. Chan, C. E. and Odde, D. J. (2008). Traction dynamics of filopodia on compliant substrates. Science 322, 1687-1691. Chen, C. S. (2008). Mechanotransduction – a field pulling together? J. Cell Sci. 121, 3285-3292. Delorme-Walker, V. D., Peterson, J. R., Chernoff, J., Waterman, C. M., Danuser, G., DerMardirossian, C. and Bokoch, G. M. (2011). Pak1 regulates focal adhesion strength, myosin IIA distribution, and actin dynamics to optimize cell migration. J. Cell Biol. 193, 1289-1303. Fonseca, P. M., Shin, N. Y., Bra´bek, J., Ryzhova, L., Wu, J. and Hanks, S. K. (2004). Regulation and localization of CAS substrate domain tyrosine phosphorylation. Cell. Signal. 16, 621-629. Fukata, Y., Oshiro, N., Kinoshita, N., Kawano, Y., Matsuoka, Y., Bennett, V., Matsuura, Y. and Kaibuchi, K. (1999). Phosphorylation of adducin by Rhokinase plays a crucial role in cell motility. J. Cell Biol. 145, 347-361. Gardel, M. L., Schneider, I. C., Aratyn-Schaus, Y. and Waterman, C. M. (2010). Mechanical integration of actin and adhesion dynamics in cell migration. Annu. Rev. Cell Dev. Biol. 26, 315-333. Geiger, B., Bershadsky, A., Pankov, R. and Yamada, K. M. (2001). Transmembrane crosstalk between the extracellular matrix – cytoskeleton crosstalk. Nat. Rev. Mol. Cell Biol. 2, 793-805. Guo, W. H. and Wang, Y. L. (2007). Retrograde fluxes of focal adhesion proteins in response to cell migration and mechanical signals. Mol. Biol. Cell 18, 4519-4527. Gupton, S. L. and Waterman-Storer, C. M. (2006). Spatiotemporal feedback between actomyosin and focal-adhesion systems optimizes rapid cell migration. Cell 125, 1361-1374. Hirata, H., Tatsumi, H. and Sokabe, M. (2008). Mechanical forces facilitate actin polymerization at focal adhesions in a zyxin-dependent manner. J. Cell Sci. 121, 2795-2804. Hopkins, A. M., Pineda, A. A., Winfree, L. M., Brown, G. T., Laukoetter, M. G. and Nusrat, A. (2007). Organized migration of epithelial cells requires control of adhesion and protrusion through Rho kinase effectors. Am. J. Physiol. 292, G806-G817. Hotta, K, Ranganathan, S, Liu, R, Wu, F, Machiyama, H, Gao, R, Hirata, H, Soni, N, Ohe, T, Hogue, C. W et al. (2014). Biophysical properties of intrinsically disordered p130Cas substrate domain – implication in mechanosensing. PLoS Comput Biol. 10, e 1003532. Hsia, D. A., Mitra, S. K., Hauck, C. R., Streblow, D. N., Nelson, J. A., Ilic, D., Huang, S., Li, E., Nemerow, G. R., Leng, J. et al. (2003). Differential regulation of cell motility and invasion by FAK. J. Cell Biol. 160, 753-767. Hu, C. D., Chinenov, Y. and Kerppola, T. K. (2002). Visualization of interactions among bZIP and Rel family proteins in living cells using bimolecular fluorescence complementation. Mol. Cell 9, 789-798. Hu, K., Ji, L., Applegate, K. T., Danuser, G. and Waterman-Storer, C. M. (2007). Differential transmission of actin motion within focal adhesions. Science 315, 111-115. Ivkovic, S., Beadle, C., Noticewala, S., Massey, S. C., Swanson, K. R., Toro, L. N., Bresnick, A. R., Canoll, P. and Rosenfeld, S. S. (2012). Direct inhibition of myosin II effectively blocks glioma invasion in the presence of multiple motogens. Mol. Biol. Cell 23, 533-542.

3449


RESEARCH ARTICLE

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Olivero, D. K. and Furcht, L. T. (1993). Type IV collagen, laminin, and fibronectin promote the adhesion and migration of rabbit lens epithelial cells in vitro. Invest. Ophthalmol. Vis. Sci. 34, 2825-2834. Parsons, J. T. (2003). Focal adhesion kinase: the first ten years. J. Cell Sci. 116, 1409-1416. Pasapera, A. M., Schneider, I. C., Rericha, E., Schlaepfer, D. D. and Waterman, C. M. (2010). Myosin II activity regulates vinculin recruitment to focal adhesions through FAK-mediated paxillin phosphorylation. J. Cell Biol. 188, 877-890. Polte, T. R. and Hanks, S. K. (1995). Interaction between focal adhesion kinase and Crk-associated tyrosine kinase substrate p130Cas. Proc. Natl. Acad. Sci. USA 92, 10678-10682. Ponti, A., Machacek, M., Gupton, S. L., Waterman-Storer, C. M. and Danuser, G. (2004). Two distinct actin networks drive the protrusion of migrating cells. Science 305, 1782-1786. Ridley, A. J., Schwartz, M. A., Burridge, K., Firtel, R. A., Ginsberg, M. H., Borisy, G., Parsons, J. T. and Horwitz, A. R. (2003). Cell migration: integrating signals from front to back. Science 302, 1704-1709. Ruest, P. J., Shin, N. Y., Polte, T. R., Zhang, X. and Hanks, S. K. (2001). Mechanisms of CAS substrate domain tyrosine phosphorylation by FAK and Src. Mol. Cell. Biol. 21, 7641-7652. Sakai, R., Iwamatsu, A., Hirano, N., Ogawa, S., Tanaka, T., Mano, H., Yazaki, Y. and Hirai, H. (1994). A novel signaling molecule, p130, forms stable complexes in vivo with v-Crk and v-Src in a tyrosine phosphorylation-dependent manner. EMBO J. 13, 3748-3756. Sakamoto, T., Limouze, J., Combs, C. A., Straight, A. F. and Sellers, J. R. (2005). Blebbistatin, a myosin II inhibitor, is photoinactivated by blue light. Biochemistry 44, 584-588. Sawada, Y., Tamada, M., Dubin-Thaler, B. J., Cherniavskaya, O., Sakai, R., Tanaka, S. and Sheetz, M. P. (2006). Force sensing by mechanical extension of the Src family kinase substrate p130Cas. Cell 127, 1015-1026. Schwartz, M. A. (2010). Integrins and extracellular matrix in mechanotransduction. Cold Spring Harb. Perspect. Biol. 2, a005066. Shin, N. Y., Dise, R. S., Schneider-Mergener, J., Ritchie, M. D., Kilkenny, D. M. and Hanks, S. K. (2004). Subsets of the major tyrosine phosphorylation sites in Crk-associated substrate (CAS) are sufficient to promote cell migration. J. Biol. Chem. 279, 38331-38337. Stricker, J., Beckham, Y., Davidson, M. W. and Gardel, M. L. (2013). Myosin IImediated focal adhesion maturation is tension insensitive. PLoS ONE 8, e70652. Tsuruta, D. and Jones, J. C. (2003). The vimentin cytoskeleton regulates focal contact size and adhesion of endothelial cells subjected to shear stress. J. Cell Sci. 116, 4977-4984. Uehata, M., Ishizaki, T., Satoh, H., Ono, T., Kawahara, T., Morishita, T., Tamakawa, H., Yamagami, K., Inui, J., Maekawa, M. et al. (1997). Calcium sensitization of smooth muscle mediated by a Rho-associated protein kinase in hypertension. Nature 389, 990-994. Wang, Y. L. (2007). Flux at focal adhesions: slippage clutch, mechanical gauge, or signal depot. Sci. STKE 2007, pe10. Webb, D. J., Parsons, J. T. and Horwitz, A. F. (2002). Adhesion assembly, disassembly and turnover in migrating cells – over and over and over again. Nat. Cell Biol. 4, E97-E100. Webb, D. J., Donais, K., Whitmore, L. A., Thomas, S. M., Turner, C. E., Parsons, J. T. and Horwitz, A. F. (2004). FAK-Src signalling through paxillin, ERK and MLCK regulates adhesion disassembly. Nat. Cell Biol. 6, 154-161. Wells, C. M., Whale, A. D., Parsons, M., Masters, J. R. and Jones, G. E. (2010). PAK4: a pluripotent kinase that regulates prostate cancer cell adhesion. J. Cell Sci. 123, 1663-1673. Wolfenson, H., Bershadsky, A., Henis, Y. I. and Geiger, B. (2011). Actomyosingenerated tension controls the molecular kinetics of focal adhesions. J. Cell Sci. 124, 1425-1432. Zaidel-Bar, R., Itzkovitz, S., Ma’ayan, A., Iyengar, R. and Geiger, B. (2007). Functional atlas of the integrin adhesome. Nat. Cell Biol. 9, 858-867.


Kanchanawong, P., Shtengel, G., Pasapera, A. M., Ramko, E. B., Davidson, M. W., Hess, H. F. and Waterman, C. M. (2010). Nanoscale architecture of integrin-based cell adhesions. Nature 468, 580-584. Kawauchi, K., Araki, K., Tobiume, K. and Tanaka, N. (2008). p53 regulates glucose metabolism through an IKK-NF-kappaB pathway and inhibits cell transformation. Nat. Cell Biol. 10, 611-618. Kawauchi, K., Tan, W. W., Araki, K., Abu Bakar, F. B., Kim, M., Fujita, H., Hirata, H. and Sawada, Y. (2012). p130Cas-dependent actin remodelling regulates myogenic differentiation. Biochem. J. 445, 323-332. Kerppola, T. K. (2009). Visualization of molecular interactions using bimolecular fluorescence complementation analysis: characteristics of protein fragment complementation. Chem. Soc. Rev. 38, 2876-2886. Kim, D. H. and Wirtz, D. (2013). Focal adhesion size uniquely predicts cell migration. FASEB J. 27, 1351-1361. Kimura, K., Fukata, Y., Matsuoka, Y., Bennett, V., Matsuura, Y., Okawa, K., Iwamatsu, A. and Kaibuchi, K. (1998). Regulation of the association of adducin with actin filaments by Rho-associated kinase (Rho-kinase) and myosin phosphatase. J. Biol. Chem. 273, 5542-5548. Klinghoffer, R. A., Sachsenmaier, C., Cooper, J. A. and Soriano, P. (1999). Src family kinases are required for integrin but not PDGFR signal transduction. EMBO J. 18, 2459-2471. Kostic, A. and Sheetz, M. P. (2006). Fibronectin rigidity response through Fyn and p130Cas recruitment to the leading edge. Mol. Biol. Cell 17, 26842695. Kovaćs, M., To´th, J., Heteńyi, C., Ma´lna´si-Csizmadia, A. and Sellers, J. R. (2004). Mechanism of blebbistatin inhibition of myosin II. J. Biol. Chem. 279, 35557-35563. Kuo, J. C., Han, X., Hsiao, C. T., Yates, J. R., 3rd and Waterman, C. M. (2011). Analysis of the myosin-II-responsive focal adhesion proteome reveals a role for b-Pix in negative regulation of focal adhesion maturation. Nat. Cell Biol. 13, 383393. Lee, H. H., Tien, S. C., Jou, T. S., Chang, Y. C., Jhong, J. G. and Chang, Z. F. (2010). Src-dependent phosphorylation of ROCK participates in regulation of focal adhesion dynamics. J. Cell Sci. 123, 3368-3377. Lele, T. P., Pendse, J., Kumar, S., Salanga, M., Karavitis, J. and Ingber, D. E. (2006). Mechanical forces alter zyxin unbinding kinetics within focal adhesions of living cells. J. Cell. Physiol. 207, 187-194. Machiyama, H., Tatsumi, H. and Sokabe, M. (2009). Structural changes in the cytoplasmic domain of the mechanosensitive channel MscS during opening. Biophys. J. 97, 1048-1057. Meenderink, L. M., Ryzhova, L. M., Donato, D. M., Gochberg, D. F., Kaverina, I. and Hanks, S. K. (2010). P130Cas Src-binding and substrate domains have distinct roles in sustaining focal adhesion disassembly and promoting cell migration. PLoS ONE 5, e13412. Missbach, M., Jeschke, M., Feyen, J., Mu¨ller, K., Glatt, M., Green, J. and Susa, M. (1999). A novel inhibitor of the tyrosine kinase Src suppresses phosphorylation of its major cellular substrates and reduces bone resorption in vitro and in rodent models in vivo. Bone 24, 437-449. Murakoshi, H., Lee, S. J. and Yasuda, R. (2008). Highly sensitive and quantitative FRET-FLIM imaging in single dendritic spines using improved non-radiative YFP. Brain Cell Biol. 36, 31-42. Nagai, T., Ibata, K., Park, E. S., Kubota, M., Mikoshiba, K. and Miyawaki, A. (2002). A variant of yellow fluorescent protein with fast and efficient maturation for cell-biological applications. Nat. Biotechnol. 20, 87-90. Nakamoto, T., Sakai, R., Honda, H., Ogawa, S., Ueno, H., Suzuki, T., Aizawa, S., Yazaki, Y. and Hirai, H. (1997). Requirements for localization of p130cas to focal adhesions. Mol. Cell. Biol. 17, 3884-3897. Nakayama, M., Amano, M., Katsumi, A., Kaneko, T., Kawabata, S., Takefuji, M. and Kaibuchi, K. (2005). Rho-kinase and myosin II activities are required for cell type and environment specific migration. Genes Cells 10, 107-117. Oakes, P. W., Beckham, Y., Stricker, J. and Gardel, M. L. (2012). Tension is required but not sufficient for focal adhesion maturation without a stress fiber template. J. Cell Biol. 196, 363-374.


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Super-resolution links vinculin localization to function in focal adhesions.

Anisotropic forces from spatially constrained focal adhesions mediate contact guidance directed cell migration.

KSHV-TK is a tyrosine kinase that disrupts focal adhesions and induces Rho-mediated cell contraction.

Focal adhesion kinase-dependent focal adhesion recruitment of SH2 domains directs SRC into focal adhesions to regulate cell adhesion and migration.

Extracellular cell wall β(1,3)glucan is required to couple septation to actomyosin ring contraction.

Effects of substrate stiffness and actomyosin contractility on coupling between force transmission and vinculin-paxillin recruitment at single focal adhesions.

Dynamics of Actin Stress Fibers and Focal Adhesions during Slow Migration in Swiss 3T3 Fibroblasts: Intracellular Mechanism of Cell Turning.

Cancer Biology: APC Delivers Kiss of Death to Focal Adhesions.

Single and collective cell migration: the mechanics of adhesions.

Epstein-Barr virus LMP1 induces focal adhesions and epithelial cell migration through effects on integrin-α5 and N-cadherin.

Oligodendroglial p130Cas is a target of Fyn kinase involved in process formation, cell migration and survival.

Focal adhesions, stress fibers and mechanical tension.

Ouabain affects cell migration via Na,K-ATPase-p130cas and via nucleus-centrosome association.

Curvature-induced expulsion of actomyosin bundles during cytokinetic ring contraction.

Eardrum displacement following stapedius muscle contraction.

Disentangling Membrane Dynamics and Cell Migration; Differential Influences of F-actin and Cell-Matrix Adhesions.

A Case of Contraction of the Lower Jaw from Adhesions (Internal) of Twelve Years' Standing.

Induction of focal adhesions and motility in Drosophila S2 cells.

Step-by-step quantitative analysis of focal adhesions.

Tuning the material-cytoskeleton crosstalk via nanoconfinement of focal adhesions.

Src-mediated tyrosine phosphorylation of Protein Kinase D2 at focal adhesions regulates cell adhesion.

Vinculin-p130Cas interaction is critical for focal adhesion dynamics and mechano-transduction.

Talin determines the nanoscale architecture of focal adhesions.

ATM regulation of IL-8 links oxidative stress to cancer cell migration and invasion.