Integrating actin dynamics, mechanotransduction and integrin activation: The multiple functions of actin binding proteins in focal adhesions

European Journal of Cell Biology 92 (2013) 339–348

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Review

Integrating actin dynamics, mechanotransduction and integrin activation: The multiple functions of actin binding proteins in focal adhesions Corina Ciobanasu, Bruno Faivre, Christophe Le Clainche ∗ Laboratoire d’Enzymologie et Biochimie Structurales CNRS, avenue de la terrasse, 91198 Gif-sur-Yvette, France

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Article history: Received 31 July 2013 Received in revised form 10 October 2013 Accepted 23 October 2013 Keywords: Actin Mechanotransduction Integrins Focal adhesions

a b s t r a c t Focal adhesions are clusters of integrin transmembrane receptors that mechanically couple the extracellular matrix to the actin cytoskeleton during cell migration. Focal adhesions sense and respond to variations in force transmission along a chain of protein-protein interactions linking successively actin filaments, actin binding proteins, integrins and the extracellular matrix to adapt cell-matrix adhesion to the composition and mechanical properties of the extracellular matrix. This review focuses on the molecular mechanisms by which actin binding proteins integrate actin dynamics, mechanotransduction and integrin activation to control force transmission in focal adhesions. © 2013 Elsevier GmbH. All rights reserved.

Introduction During migration, cells respond to chemical and mechanical cues by adapting their shape, dynamics and adhesion to the extracellular matrix (ECM). In this process, focal adhesions (FAs) play a critical role (Geiger et al., 2009). FAs are clusters of integrin transmembrane receptors that couple the ECM to the actin cytoskeleton, via actin binding proteins (ABPs) (Gardel et al., 2010; Le Clainche and Carlier, 2008). The integrin family comprises 24 ␣␤ heterodimers characterised by their specificity for a large repertoire of ECM molecules and their activation mechanism (Hynes, 2002; Hynes and Naba, 2012). In addition, FAs contain a variety of ABPs that are regulated by specific signalling pathways (ZaidelBar et al., 2007). FAs sense and respond to variations in force transmission along the actomyosin-integrin-ABP-ECM pathway to adapt cell-matrix adhesion to the composition and the properties of the ECM (Geiger et al., 2009; Ross et al., 2013). In this chain of protein-protein interactions, force transmission is determined by the mechanical properties of the proteins and the kinetic parameters of the interactions. The stretching and the fibrillogenesis of ECM molecules like fibronectin control the binding of the extracellular domain of integrins to the ECM and the resulting force transmission (Klotzsch et al., 2009; Roca-Cusachs et al., 2012; Sechler et al., 2001). Inside the cell, ABPs do not convey the force passively but modulate transmission efficiency by controlling the

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elongation of actin filaments and the activation of integrins. Finally, ABPs act as mechanosensitive switches to control the mechanical coupling between FAs and the actin cytoskeleton (Moore et al., 2010; Roca-Cusachs et al., 2012). Instead of providing a comprehensive description of the molecular components of FAs, this review focuses on the molecular mechanisms by which a subset of ABPs controls force transmission by integrating actomyosin dynamics, mechanotransduction and integrin activation. Actin structures connected to FAs FAs transmit the force generated by different actin networks. FAs interact with the retrograde flow of the lamellar actin network and, in many cell types, with actomyosin bundles called dorsal and ventral stress fibres (SFs) and transverse arcs (Burridge and Wittchen, 2013; Vallenius, 2013). SF formation results from the combined action of parallel pathways including ROCK (Rho-associated protein kinase) and mDia1, downstream of RhoA. ROCK activates myosin II, leading to the formation of contractile actomyosin fibres while the formin mDia1 induces the formation of thin non-contractile bundles (Watanabe et al., 1999). In migrating cells, mDia1 is required for the elongation of dorsal SFs (Hotulainen and Lappalainen, 2006). The formation of ventral SFs results from the assembly of two subpopulations of actin filaments. The first population is initiated in nascent adhesions or at the leading edge by the formin mDia2 and assembles with tropomyosin and myosin II filaments while the second is made of branched filaments nucleated by Arp2/3 at the leading edge and reorganised by the crosslinking protein ␣-actinin.

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The annealing of these two actin structures leads to the formation of transverse arcs that ultimately connect with dorsal SFs to form contractile ventral SFs between FAs (Tojkander et al., 2011). In cells that do not form prominent stress fibres, mDia2 controls the dynamics of FAs and the associated actin network (Gupton et al., 2007). In these cells, mDia2 is associated with an unidentified vesicular fraction and microtubules but importantly, the protein is absent from the leading edge and FAs. Although it is possible that a subpopulation of formins remains associated to actin filaments in FAs, the main linkage between FAs and SFs, or the lamellar retrograde flow, is provided by the ABPs described in the next sections (Fig. 1). Recent studies revealed that the focal adhesion plaque is a heterogeneous surface in which distinct domains are associated with actin structures characterised by different architectures and contractile properties. The composite architecture of the actin network anchored to a single FA was revealed by three-dimensional reconstruction using cryo-electron tomography (Patla et al., 2010). This study showed that unidentified ring-shaped particles anchor a first layer of short and randomly oriented actin filaments, while an upper layer is occupied by the tip of a SF made of parallel actin filaments. The upper layer only covers the central part while the first layer covers the entire surface of the adhesion plaque. The combination of super-resolution microscopy and singleparticle tracking showed that ␤3 integrins are enriched and immobile in mature FAs, while ␤1 integrins are present at a lower density and display a slow rearward motion, indicating that distinct pathways connecting actin to integrins coexist and transmit force differently inside FAs (Rossier et al., 2012). The recent development of genetically engineered cell lines, expressing either ␣5 ␤1 integrins or ␣v ␤3 and ␣v ␤5 integrins, provided the demonstration that integrins are segregated in distinct domains in FAs, where they are associated to distinct actin networks (Schiller et al., 2013). ␣5 ␤1 integrins are enriched in nascent adhesions where they are associated with the lamellipodial actin network. In contrast, ␣v integrins are enriched in FAs where prominent stress fibres are anchored. ␤1 integrins are associated with high myosin activity and traction force whereas, surprisingly, the large SFs associated with ␣v integrins apply a low traction force. Co-expression of both integrins revealed that ␣5 ␤1 and ␣v ␤3 segregate in partially overlapping areas. Interestingly, in these cells, ␣v ␤3 integrins are associated to prominent SFs that apply a high traction force. These observations, together with proteomic data, suggest that two distinct compartments in FAs made of ␣5 ␤1 and ␣v integrins cooperate to induce actomyosin contractility through the ROCK-myosin II pathway and actin assembly through mDia1 respectively.

Force transmission by actin binding proteins Force transmission efficiency is regulated by the kinetic parameters of the protein-protein interactions, the mechanical properties of the proteins and multiple biochemical reactions along the pathway linking successively actin filaments, ABPs, integrins and the ECM (Fig. 2). The view that a hierarchical network of proteins transmits force across FAs may not reflect the very dynamic nature of the mechanical coupling between the ECM and the actin cytoskeleton. Indeed, the observation of fluorescent speckles corresponding to various ABPs, moving centripetally at the interface between FAs and the retrograde flow of actin, revealed a complex slippage interface. Inside this interface, also called “molecular clutch”, the motion of ABPs, like talin and vinculin, is only partially coupled to actin (Hu et al., 2007). In addition to the protein-protein interactions identified in individual studies, recent proteomic studies provided a detailed cartography of the protein network that conveys the force between the actin cytoskeleton and the ECM (Kuo et al., 2011; Schiller and

Fassler, 2013; Schiller et al., 2013; Zaidel-Bar et al., 2007; Zamir et al., 1999). Altogether, these studies revealed that four ABPs (talin, filamin, ␣-actinin and tensin) connect the actin cytoskeleton to integrins directly. Several ABPs (VASP, vinculin, and the ILK-PINCHParvin complex) are also indirectly connected to integrins via one or more adaptors. In the next paragraphs we discuss the contribution of several ABPs in force transmission across FAs. Talin Talin is one of the earliest ABPs to connect integrins and the actin cytoskeleton (Critchley, 2009). There are two talin genes encoding for talin1 and talin2. Talin1 is ubiquitously expressed while talin2 is absent in hematopoietic cells. Mice lacking talin1 die from gastrulation defects while the complete deletion of talin2 has little consequences. Talin1 and talin2 share 74% sequence identity and display the same domain organisation (Critchley, 2009; Debrand et al., 2012; Monkley et al., 2000). Because the function of talin1 has been extensively studied, while the role of talin2 remains unclear, this review focuses on talin1, simply referred to as talin. Talin is made of a head domain containing a F0 domain followed by a FERM (four-point-one, ezrin, radixin, moesin) domain divided into three subdmomains (F1, F2 and F3). F2 and F3 form an actin filament binding domain (ABD) while F3 interacts with the cytoplasmic tail of the ␤ subunit of integrins. This head domain is followed by a rod domain divided into 13 helix bundles. The rod contains 11 vinculin binding sites (VBSs), a central ABD and a C-terminal ABD (THATCH) (Fig. 1). A structural model of fulllength talin, derived from the combination of crystallographic data and electron microscopy reconstruction, suggests that both the head and the C-terminal ABD are masked in the dimeric inactive talin (Goult et al., 2013a). In the inactive state, F2 and F3 must be released from autoinhibitory contacts with the R1-R2 and R9 domains of the rod respectively (Banno et al., 2012; Goult et al., 2009) (Fig. 3A). The mechanism by which talin is initially activated is still unclear. The binding of PtdIns(4,5)P2 to basic residues at the surface of the F1, F2 and F3 domains could disrupt the autoinhibitory contacts (Elliott et al., 2010; Goult et al., 2010). The membrane-bound RAP1A-RIAM complex may also recruit and activate talin through a direct interaction between RIAM and the talin rod domain (Goult et al., 2013b; Han et al., 2006; Lee et al., 2009) (Fig. 1). Once activated, talin provides the early 2 pN bond between the actin cytoskeleton and the ECM (Giannone et al., 2003; Jiang et al., 2003). However, the molecular pathway that transmits the force is not known. The position of FA proteins along the vertical axis (perpendicular to the plasma membrane), determined by super-resolution microscopy, showed that the N-terminal domain of talin is associated with a membrane-proximal layer of adaptors while the C-terminal domain interacts with the actin network, 80 nm away from the membrane (Kanchanawong et al., 2010). Because the N-terminal head of talin interacts with the NPXY motif of integrin ␤-tails and the C-terminal domain contains the THATCH actin binding domain, it was suggested that talin mechanically couples integrins to actin filaments. However, in Drosophila, a mutation in the talin head domain, that abolishes its binding to the integrin NPXY motif, provokes adhesion defects but does not affect talin recruitment in adhesion complexes nor cytoskeleton anchoring (Tanentzapf et al., 2006). In addition, super-resolution microscopy showed that talin head, containing the first integrin binding site, is not enriched in FAs (Rossier et al., 2012). Altogether, these observations suggest that, although integrins are activated by talin, integrins do not anchor talin in FAs. However, the requirement of the C-terminal dimerisation domain of talin, absent in talin head, could explain some of the results mentioned above (Smith and McCann, 2007). By interacting directly with talin head and by synthesising PtdIns(4,5)P2 , PtdInsP Kinase I␥ could anchor and

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activate talin (Legate et al., 2011). As mentioned previously, the binding of RIAM to the R2-R3 bundles of the rod may also contribute to anchor talin (Goult et al., 2013b). Finally, the second integrin binding site located in the rod is not involved in integrin activation and could also target talin to the plasma membrane (Ellis et al., 2011; Rodius et al., 2008). Vinculin Vinculin is a ubiquitously expressed ABP. This protein is made of a head domain (Vh) and a tail domain (Vt) separated by a flexible linker (Fig. 1). In muscle cells, vinculin is co-expressed with a splice variant called metavinculin. This splice variant is characterised by the presence of a 68 amino acids insertion in Vt. Metavinculin reinforces the mechanical coupling between actin filaments and muscles-specific adhesion sites (costameres) (Thoss et al., 2013). In the inactive state of vinculin, the intramolecular interaction between Vh and Vt prevents Vt to interact with actin filaments (Ziegler et al., 2006). The activation of vinculin requires the concomitant binding of talin to Vh and actin filaments to Vt (Chen et al., 2006). The use of a FRET-based mechanosensor revealed that vinculin transmits force across FAs (Grashoff et al., 2010). The current view suggests that force is applied between Vh, bound to membrane-anchored talin, and Vt, bound to the actomyosin cytoskeleton. In support to this hypothesis, the time during which full-length vinculin remains associated with FAs correlates with the force applied. This correlation requires the presence of Vt (Dumbauld et al., 2013). The observation that vinculin recruitment is abolished in cells lacking talin also supports this pathway (Zhang et al., 2008). Finally, this model is compatible with the relative position of the VBS-containing talin rod domain and vinculin along the vertical axis (Kanchanawong et al., 2010). An alternative model has been proposed in which the force-dependent recruitment of vinculin involves the Src/FAK-dependent phosphorylation of paxillin (Pasapera et al., 2010). However, vinculin and paxillin do not colocalise in FAs (Humphries et al., 2007). In addition, Vh, that does not contain the paxillin binding site, is efficiently targeted to FAs (Carisey et al., 2013). The binding of PtdIns(4,5)P2 to Vt, that inhibits the binding of actin filaments (Steimle et al., 1999), may regulate force transmission in FAs. In agreement with this hypothesis, vinculin increases the turnover of FAs in regions of high PtdIns(4,5)P2 density (Chandrasekar et al., 2005; Saunders et al., 2006). ˛-Actinin ␣-Actinin is an actin filament crosslinking protein. There are four ␣-actinin genes encoding for ␣-actinin-1, -2, -3 and -4. The isoforms 1 and 4 are associated with FAs and SFs in non-muscle cells while the isoforms 2 and 3 are found in skeletal and smooth muscles respectively (Sjoblom et al., 2008). The four isoforms interact with actin filaments through two N-terminal calponin homology (CH) domains and form anti-parallel dimers through four spectrin repeats (R1–R4) (Fig. 1). Surprisingly, the selective removal of SFs in ␣-actinin-1 siRNA-treated cells, results in a higher traction force, indicating that the crosslinking activity of ␣-actinin impedes the actomyosin contractility (Oakes et al., 2012). In agreement with this observation, the higher traction force is not measured at SF-FA attachment site (Plotnikov et al., 2012). Altogether, these observations suggest that ␣-actinin stabilises SFs to promote the establishment of a persistent isometric tension rather than a high tension. In addition to this long distance effect on force generation, ␣-actinin is thought to control force transmission in FAs between integrins and SFs through a direct interaction with integrins (Otey et al., 1990; Roca-Cusachs et al., 2013). The binding site for the integrin ␤-tail is located between the spectrin repeats

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R1 and R2 (Kelly and Taylor, 2005). However, in mature FAs, ␣actinin is absent from the membrane proximal layer of proteins where integrin tails are concentrated (Kanchanawong et al., 2010). In addition, ␣-actinin is strongly coupled to actin retrograde flow and no immobile fraction was reported in mature FAs (Hu et al., 2007). Finally, after cytoschalasin D treatment, ␣-actinin dissociates from FAs and follows the remaining actin cytoskeleton (Carisey et al., 2013). Although the role of the integrin-␣-actinin interaction in mature FAs remains to be clarified, this direct interaction could explain another possible role of ␣-actinin in the elongation of nascent adhesions (Choi et al., 2008). Tensin Tensin is a family of ABPs that connect integrins to the actin cytoskeleton. These proteins are absent in nascent adhesions but appear in FAs and are enriched in fibrillar adhesions (FBs). The three isoforms of tensin (tensin1, tensin2 and tensin3) contain a Nterminal ABD that binds to actin filaments, a Src Homology 2 (SH2) domain and a C-terminal phosphotyrosine binding domain (PTB) that interacts with integrin ␤-tails. The C-terminal part of tensin, made of the SH2 and the PTB domains, targets tensin to FAs (Lo, 2004). Tensin1 is the only isoform that contains a central ABD that caps actin filament barbed ends (Lo et al., 1994). Although tensinrich FBs are not linked to actomyosin SFs, they are connected to thinner actin fibres and they control fibronectin fibrillogenesis in an actomyosin-dependent manner (Zamir et al., 2000; Zhong et al., 1998). Whether force transmission across tensin plays a role in this process is not known. ILK/PINCH/Parvin (IPP) complex The IPP complex made of ILK (Integrin Linked Kinase), PINCH and parvin may also play a role in force transmission across FAs by stabilising the link between the actin cytoskeleton and integrins (Legate et al., 2006; Stanchi et al., 2009). Protein-protein interaction studies showed that the N-terminal ankyrin repeats of ILK interact with the first LIM (Lin11, Isl-1 and Mec-3) domain of PINCH while the C-terminal pseudokinase domain of ILK interacts with the second CH domain of parvin (Fig. 1). In this complex the two CH domains of parvin, also known as actopaxin, interact with actin filaments (Nikolopoulos and Turner, 2000). Although ILK interacts with integrin ␤1 (Hannigan et al., 1996), it is not clear whether the targeting of the IPP complex to FAs is mediated by this direct interaction or by an interaction with kindlins (Widmaier et al., 2012). There is only one gene encoding for ILK. In contrast, the parvin family comprises three isoforms (␣-parvin, ␤-parvin and ␥-parvin) and two isoforms of PINCH have been identified (PINCH1 and PINCH2). Many of these isoforms are co-expressed in the same tissues. It is therefore possible that distinct IPP complexes coexist in cells (Legate et al., 2006). The recent SAXS (Small angle X-ray scattering) analysis of a minimal IPP complex revealed that the limited flexibility of the structure is compatible with efficient force transmission (Stiegler et al., 2013). A recent report showed that ILK-null cells are characterised by adhesion defects and a disorganisation of the actin cytoskeleton (Elad et al., 2013), suggesting that the IPP complex could regulate directly or indirectly the inside-out activation of integrins through ILK and actin dynamics through parvin. Filamin The filamin family comprises filamin A (ABP-280), filamin B and filamin C (Razinia et al., 2012). Filamin A and B are expressed in many cell types while filamin C expression is limited to skeletal muscles. Filamins interact with actin filaments through two N-terminal CH domains. This ABD is followed by a rod made of

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Fig. 1. Domain organisation and binding partners of the actin binding proteins of focal adhesions. The name of the domains are written in black and the major binding partners are written in grey. The following abbreviations are listed below by order of appearance. Talin: ABD, actin binding domain; FERM, four-point-one, ezrin, radixin, moesin; IBS, integrin binding site; PTB, phosphotyrosine binding domain; VBS, vinculin binding site (11 VBSs are indicated in red); THATCH, Talin/Hip1R/Sla2p Actin Tethering C-terminal Homology domain; DD, dimerisation domain. Filamin A: CH, calponin homology; Ig1, immunoglobulin domain 1. Vinculin: Vh, vinculin head; D1–D4, vinculin head subdomains 1–4; D5/Vt, vinculin tail. ␣-Actinin: R1–R4, spectrin repeats 1–4; EF, EF hand motif. VASP: EVH1, Ena/VASP homology domain 1; G, G-actin (monomeric actin) binding domain; F, F-actin (actin filament) binding domain; PRD, Proline-rich domain; TD, tetramerisation domain. Tensin: SH2, Src Homology 2 domain. IPP complex: ILK, Integrin Linked Kinase; ANK 1–4, Ankyrin repeats 1–4; PK, Pseudokinase domain; L1–L5, LIM (Lin11, Isl-1 and Mec-3) domain 1–5. The numerous binding partners of the IPP complex are not indicated in this figure. A comprehensive list can be found in Widmaier et al. (2012). Note that the abbreviation ABD refers to a domain that interacts with actin filaments. Some ABDs correspond to conserved domains, as mentioned on the figure (CH and THATCH for example). Others are not related to conserved domains and have not been named (ABD2 in talin and ABD1 and ABD2 in tensin1). Unless mentioned in the figure, ABDs interact with the side of actin filaments. ABD2 in tensin1 and Vt in vinculin also interact with the barbed end of actin filaments.

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24 immunoglobulin (Ig) domains. The 24th Ig domain (IgFLNa 24) mediates filamin dimerisation, allowing the protein to crosslink orthogonal actin filaments (Nakamura et al., 2011) (Figs. 1 and 3B). The 21st Ig domain (IgFLNa 21) interacts directly with the integrin ␤-tails (Loo et al., 1998; Sharma et al., 1995). In non-muscle cells, filamins are associated with several actin-rich structures including lamellipodia, filopodia, stress fibres and focal adhesions (Lynch et al., 2013). Several observations support a role for filamins in the control of force transmission and rigidity sensing (Byfield et al., 2009; Lynch et al., 2013). Actin polymerisation and force transmission The observation that dorsal SFs elongate processively from FAs suggests that several factors control the elongation of actin filaments to modulate force transmission. Formins have been studied extensively for their role in the processive elongation of actin filaments (Goode and Eck, 2007; Romero et al., 2004). However, as mentioned previously, they have never been localised to FAs. Proteins from the Ena/VASP family, including Vasodilator-stimulated phosphoprotein (VASP) (Fig. 1), Mena (mammalian enabled) and the Ena-VASP-like protein (Evl) are clearly localised to FAs (Krause et al., 2003). In vitro reconstitution experiments showed that clusters of VASP track growing barbed ends in a processive-like manner (Breitsprecher et al., 2008). In addition to these “elongators”, barbed end capping proteins are present in FAs. Vinculin tail (Vt) does not only bind to actin filaments but also inhibits barbed end growth by a capping-like mechanism (Le Clainche et al., 2010). As mentioned previously, tensin is a high-affinity barbed end capping protein (Lo et al., 1994). In contrast with a stable anchorage, the processive elongation of actin filaments at the anchoring sites of a contractile bundle should release the tension. Therefore, in FAs, barbed end cappers should increase force transmission while elongators should decrease force transmission (Fig. 2). The presence of proteins that anchor actin filament barbed ends in FAs may have another function. By orienting the filaments in a uniform parallel manner, these proteins could favour their crosslinking by ␣-actinin. The resulting cohesive and stiff bundle has the adequate orientation and mechanical properties to be pulled by myosin filaments. This model explains why actin filaments attached to FAs display a uniform polarity with barbed ends facing the focal adhesion plaque (Cramer et al., 1997). The role of actin binding proteins in mechanotransduction Cellular observations and recent proteomic studies provided a detailed description of the force-dependent changes in adhesome composition (Kuo et al., 2011; Schiller and Fassler, 2013; Schiller et al., 2013; Zaidel-Bar et al., 2007; Zamir et al., 1999). The list of proteins recruited in FAs in a myosin-dependent manner includes signalling proteins such as numerous LIM-domain containing proteins (zyxin, paxillin, etc.) and cytoskeletal proteins (vinculin, filamin, VASP). However, only a small number of these proteins are mechanosensitive switches that convert a mechanical stimulus into a biochemical response. In addition to force-induced dissociation or reinforcement (catch bond) of protein-protein interactions (Wolfenson et al., 2011), exposure of cryptic binding domains upon protein stretching contributes to the mechanosensitive compositional changes observed in FAs (Moore et al., 2010; Roca-Cusachs et al., 2012). Talin stretching induces vinculin binding In talin, most of the 11 VBSs are buried inside helix bundles (Critchley, 2009). This observation, together with the fact that vinculin is recruited in FAs in a force-dependent manner (Galbraith

Fig. 2. Regulation of force transmission along the Actomyosin-ABP-Integrin-ECM pathway. This scheme represents the molecular elements that transmit the force between the actomyosin cytoskeleton and the ECM. In addition to the kinetic parameters of the interactions (k+ and k− ), and the mechanical properties of the proteins, the activities of several ABPs are expected to control force transmission positively (green arrow) or negatively (red arrow). The capping of actin filament barbed ends by tensin (or vinculin) provides a stable link between ABPs and actin filaments. In contrast, the processive elongation of actin filaments by VASP and mDia1 (or mDia2) releases the actomyosin tension. In response to mechanical strain, a complex made of zyxin, VASP and ␣-actinin repairs strain-induced damage along stress fibres to maintain force transmission. The inside out activation of integrins by talin reinforces the connection between the actomyosin cytoskeleton and the ECM (see Fig. 3). Filamin A binding to integrin ␤-tail competes with talin and prevents the activation of integrins (see Fig. 3). Finally, the ECM properties influences force transmission. ECM rigidity and force-induced fibrillogenesis favour force transmission.

et al., 2002), suggested that the force generated by the actomyosin cytoskeleton could stretch talin to expose cryptic VBSs. Molecular dynamics simulations first proposed a model in which a force applied on a fragment of the talin rod domain, comprising 12 helices of the R1, R2 and R3 helix bundles, breaks the structure to expose VBSs, allowing vinculin head (Vh) binding (Hytonen and Vogel, 2008) (Fig. 3A). This model also suggested that talin can be overstretched, preventing vinculin association. In vitro, the use of magnetic tweezers and atomic force microscopy demonstrated that the stretching of the R1-R3 fragment of the talin rod domain induces Vh binding (del Rio et al., 2009). However, no reduction of the association could be observed at the highest force applied. In cells, the distance between fluorescent tags fused with the Nand C-termini of talin oscillates from 100 to 350 nm in a myosindependent manner (Margadant et al., 2011). These observations, together with the stabilisation of the extended conformation of talin by Vh in cells (Margadant et al., 2011), support a model in which the actomyosin-dependent stretching of talin is a prerequisite for vinculin association. The function of this mechanosensitive switch is still unclear. The mechanosensitive exposure of the actin binding tail of vinculin (Vt) could reinforce the coupling between FAs and the retrograde flow of actin or resist the actomyosin force (Fig. 2). Vinculin could also provide a secondary path to transmit force to talin and stretch its rod domain, leading to a positive feedback loop. A recent report showed that Vt is required to couple the actin retrograde flow to the ECM but dispensable for FA growth (Thievessen et al., 2013). In agreement with these findings, the overexpression

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Fig. 3. Actin binding proteins integrate actin dynamics and integrin regulation in a mechanosensitive manner. (A) Talin couples the activation of integrins to the mechanosensitive regulation of actin assembly. (Left panel) In its inactive state, talin dimer adopts a donut-shaped conformation in which the two heads fill the central hole. The binding of the head domain of talin to the plasma membrane induces the release of the rod, allowing the interaction of the THATCH domain (R13) with actin filaments. Talin is represented as a monomer to simplify this scheme but the active form of talin is a dimer. The binding of the F3 domain (PTB) of talin to integrin ␤-tail induces the inside-out activation of integrin characterised by the open-extended conformation. The inside-out activation of integrin involves the synergistic or sequential action of talin and kindlin (K). In response to the force generated by the acomyosin stress fibres (or the lamellar retrograde flow), talin is stretched and exposes cryptic vinculin binding sites. The activation of vinculin reinforces the anchoring of actin filaments with focal adhesions and blocks actin filament barbed end elongation. (Right panel) In its inactive state, several domains of talin are auto-inhibited. The scheme shows the contact between the F3 (PTB) domain of the head and the R9 helix bundle located in the rod domain. This contact prevents integrin ␤-tail to interact with F3. An inhibitory contact also exists between the VBS-containing region made of the R1 and R2 helix bundles and the F2-F3 domains of the head. The binding of F2 to PtdIns(4,5)P2 at the surface of the plasma membrane and F3 to integrin ␤-tail disrupts the auto-inhibitory contacts. Talin interacts with both the membrane proximal (MP) and the proximal NPXY motif (Y) of integrin ␤-tail, while kindlin interacts with the distal NPXY (Y) motif. In response to the force generated by the actomyosin cytoskeleton, the VBS-containing domain is stretched, leading to the exposure of VBSs. The binding of vinculin head (Vh) to the newly exposed VBS induces the release of the actin binding tail of vinculin (Vt). Vinculin tail interacts with both the side and the barbed end (+) of an actin filament. (B) Filamin A inhibits integrins in response to a mechanical strain. (Upper panel) Filamin A crosslinks actin filaments in an orthogonal manner. The mechanical strain applied on the actin network induces the binding of filamin A to integrin ␤-tail. Filamin A does not activate integrins and prevents talin and kindlin to activate integrins. Although integrins are represented in the closed conformation, the filamin-bound integrin conformation is not known. Adapted from Ehrlicher et al. (2011). (Lower panel) Filamin A responds to a mechanical strain by exposing a cryptic integrin binding site located in the 21st immunoglobulin domain (IgFLNa 21). Integrin ␤-tail replaces the ␤ strand of IgFLNa 20. Adapted from Rognoni et al. (2013).

of Vh is sufficient to induce the growth of FAs (Humphries et al., 2007). Assuming that vinculin binding to stretched-talin triggers FA growth, these observations indicate that force transmission through vinculin does not contribute to talin stretching.

Interestingly, the dissociation of a variety of FA proteins, induced by myosin inhibition, is dramatically reduced in cells expressing Vh (Carisey et al., 2013). These data suggest that the stabilisation of the stretched conformation of talin by Vh could play the role of

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a general switch that governs directly or indirectly the recruitment of many FA proteins in response to the actomyosin force. Filamin stretching provokes the exposure of an integrin binding site Structural studies of the rod domain of filamin A revealed that the integrin binding site located in the 21st immunoglobulin domain (IgFLNa 21) is masked by an autoinhibitory contact with a ␤ strand of the 20th adjacent domain (IgFLNa 20) (Lad et al., 2007) (Fig. 3B). In vitro, the application of a mechanical strain to a filamincrosslinked actin network provokes the concomitant dissociation of the signalling protein FilGAP (FLNa-binding RhoGTPase-Activating Protein) and the association of integrin ␤7 (Ehrlicher et al., 2011). The stretching of single molecules of IgFLNa 20–21 by optical traps showed that a force of 2–5 pN is sufficient to dissociate the two domains (Rognoni et al., 2013). Another study used magnetic tweezers to show that IgFLNa 20 and IgFLNa 21 are individually unfolded at higher forces. Interestingly, IgFLNa 20 can be unfolded in different conformations that refold at different rates. The authors predict that, at low force, this refolding rate is the limiting parameter for the reassociation of IgFLNa 20 and IgFLNa 21 that dictates the time during which the integrin binding site is exposed (Chen et al., 2013). This non-equilibrium mechanism exemplifies how the intensity of a mechanical stimulus controls the lifetime of a mechanosensitive complex. Reinforcement and maintenance of stress fibres Several mechanosensitive machineries contribute to the formation, reinforcement and maintenance of SFs. In cells submitted to a cyclic mechanical stress, zyxin accumulates along SFs and drives the recruitment of ␣-actinin and VASP in a ternary complex that induces actin polymerisation to reinforce SFs or repair straininduced damage (Smith et al., 2010) (Fig. 2). The formin mDia1 was first described to play a major role in the mechanosensitive growth of FAs downstream of RhoA (Riveline et al., 2001). Two recent reports showed that, in vitro, the processive elongation of actin filaments mediated by the mammalian formin mDia1, or the yeast Bni1p, is accelerated by the force generated by a micro-fluidic flow (Courtemanche et al., 2013; Jegou et al., 2013) (see the review by Romet-Lemonne and Jegou in this issue). Whether the assembly of SFs, that accompanies the maturation of FAs, involves the forceinduced acceleration of actin filament elongation by mDia1 remains to be determined. Other formins are good candidate for such a function. It would be interesting to determine if the formin FHOD1, that is localised along transverse arcs and SFs, stimulates actin assembly in a force-dependent manner (Schonichen et al., 2013). Actin binding proteins control integrin activation Integrins do not constitutively interact with the ECM. The activation of integrins requires a conformational change from a bent or closed conformation to an extended-open conformation (WehrleHaller, 2012) (Fig. 3). The binding of integrins to the ECM induces a conformational change leading to the extended conformation. In this conformation, integrin ␣5␤1 behaves as a catch-bond that interacts with a higher affinity with the ECM when a drag force is applied on the cytoplasmic tail of the ␤ subunit (Kong et al., 2009). This mechanism implies that the kinetic parameters of the interaction between integrins and ABPs and ABPs and actin filaments, that control force transmission, also influence the activation of integrins. In addition to the outside-in activation of integrins, an inside-out pathway involves the activation of integrins by cytoplasmic components such as talin (Tadokoro et al., 2003). The cytoplasmic tail of the ␤ subunit of integrins contains several sites

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to which synergistic activators or competitive inhibitors interact (Bouvard et al., 2013; Calderwood et al., 2013; Kim et al., 2011; Meves et al., 2009; Wehrle-Haller, 2012). Interestingly, several regulators of integrins are ABPs. Since a large part of our knowledge on integrins is based on the study of the ␤1 and ␤3-containing integrins the next paragraphs describe mainly the mechanisms by which ABP regulate these integrins. Inside-out activation of integrins by talin The PTB (Phospho Tyrosine Binding)-like F3 subdomain of talin FERM domain interacts with both the membrane proximal domain and the proximal NPXY motif of integrin ␤-tails (Fig. 3A). In vitro, the binding of the talin head domain to integrin is sufficient to induce the extended conformation of the heterodimer (Ye et al., 2010). However, in cells, the binding of kindlins (fermitins) to the distal NPXY motif of integrin ␤-tail is required for talin-induced integrin activation (Montanez et al., 2008; Moser et al., 2008) (Fig. 3A). The three kindlin isoforms (kindlin-1, kindlin-2 and kindlin-3) share a similar domain organisation characterised by a FERM domain and a plekstrin homology domain (PH) but differ in their subcellular localisation and affinity for integrin ␤-tail. The mechanism by which talin and kindlins cooperate to activate integrins is still debated (Calderwood et al., 2013). In addition to the activation mechanism described previously, the mechanosensitive binding of vinculin to R1-R3 (del Rio et al., 2009), could also facilitate the interaction between the head domain and integrin ␤-tail by preventing the re-association of the head to the rod (Fig. 3A). In agreement with this hypothesis, the expression of vinculin head increases the force needed to detach cells bound to a fibronectin-coated surface (Carisey et al., 2013). ˛-Actinin, tensin and filamin regulate talin-mediated integrin activation According to a recent report, talin and ␣-actinin activate integrin ␤3 in a competitive manner whereas they activate integrin ␤1 in a synergistic manner. In mature FAs, ␣-actinin would replace talin to maintain integrin activation and provide a strong link with the actin cytoskeleton (Roca-Cusachs et al., 2013). Structural studies are needed to determine why the structurally distinct PTB domain of talin and the R1-R2 spectrin repeats of ␣-actinin can form a ternary complex with integrin ␤3 while they compete for the closely related integrin ␤1. The hypothesis that talin and ␣-actinin transmit force at different steps of FA maturation suggests that the two proteins have different mechanical properties. ␣-Actinin is a rigid dimer in which the integrin binding sites are spaced by only 10 nm while the activated dimer of talin is composed of extended flexible protomers at the end of which the integrin binding sites are spaced by more than 100 nm. Whether these differences influence the clustering of integrins and ultimately force transmission remains to be determined. Like the PTB domain of talin, the PTB domain of tensin1 is thought to activate integrins. In contrast with talin, tensin1 binding to integrin is not reduced by the tyrosine phosphorylation of the NPXY motif (McCleverty et al., 2007). Because the NPXY is a substrate for Src, these authors proposed that the phosphorylation of integrins by Src acts as a switch to replace talin by tensin1 and direct integrins towards fibrillar adhesions. As discussed earlier, the IgFLNa 21 domain of filamin A interacts directly with a sequence of the integrin ␤-tail in a force-dependent manner. The filamin A binding site overlaps with talin and kindlin binding sites (Kiema et al., 2006), making filamin A an inhibitor of the inside-out activation of integrins (Nieves et al., 2010) (Fig. 3B). Migfilin (filamin-binding LIM protein 1) binding to filamin A has been proposed to release filamin A from integrins in order for

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kindlin and talin to activate integrins (Das et al., 2011). In addition to Filamin A, several proteins interact with integrin ␤-tails to prevent the talin-mediated activation of integrins. The spatiotemporal balance between activation and inactivation of integrins determines the adhesion strength and allows cells to respond rapidly to changes in ECM properties (Bouvard et al., 2013). Perspectives Although our understanding of the mechanisms by which ABPs coordinate actin dynamics and integrin activation to control force transmission in FAs has made rapid progress, several questions remain largely unanswered. The force transmission pathways are still poorly described. The systematic insertion of biosensors between protein domains is needed to draw a comprehensive map of the path followed by the force through the protein network formed at the cytoplasmic face of FAs. The mechanisms by which ABPs and their complexes regulate actin polymerisation in FAs remain poorly understood. In particular, whether the activities of vinculin, VASP and tensin, mentioned in this review act together in a synergistic manner to track and orientate actin filaments barbed ends or whether they act in a sequential manner during the maturation of FAs is not known. A possible role of the IPP complex in the regulation of actin assembly should be tested. Whether the structural and dynamic properties of the actin networks associated with FAs influence the spatial segregation of integrins remains to be established. Because FAs are heterogeneous structures in which a variety of integrins are connected to different actin networks under the control of complex signalling pathways, it becomes necessary to develop simplified in vitro assays that reconstitute isolated mechanosenstive circuits. The development of a minimal system made of purified components in which ABPs such as talin, ␣-actinin, filamin A or tensin connect a dynamic actomyosin cytoskeleton to a lipid bilayer containing integrins would be extremely useful. Acknowledgement CLC is supported by the Agence Nationale pour la Recherche (ANR-09-JCJC-0111, ADERACTIN). References Banno, A., Goult, B.T., Lee, H., Bate, N., Critchley, D.R., Ginsberg, M.H., 2012. Subcellular localization of talin is regulated by inter-domain interactions. J. Biol. Chem. 287, 13799–13812. Bouvard, D., Pouwels, J., De Franceschi, N., Ivaska, J., 2013. Integrin inactivators: balancing cellular functions in vitro and in vivo. Nat. Rev. Mol. Cell Biol. 14, 430–442. Breitsprecher, D., Kiesewetter, A.K., Linkner, J., Urbanke, C., Resch, G.P., Small, J.V., Faix, J., 2008. Clustering of VASP actively drives processive, WH2 domainmediated actin filament elongation. EMBO J. 27, 2943–2954. Burridge, K., Wittchen, E.S., 2013. The tension mounts: stress fibers as forcegenerating mechanotransducers. J. Cell Biol. 200, 9–19. Byfield, F.J., Wen, Q., Levental, I., Nordstrom, K., Arratia, P.E., Miller, R.T., Janmey, P.A., 2009. Absence of filamin A prevents cells from responding to stiffness gradients on gels coated with collagen but not fibronectin. Biophys. J. 96, 5095–5102. Calderwood, D.A., Campbell, I.D., Critchley, D.R., 2013. Talins and kindlins: partners in integrin-mediated adhesion. Nat. Rev. Mol. Cell Biol. 14, 503–517. Carisey, A., Tsang, R., Greiner, A.M., Nijenhuis, N., Heath, N., Nazgiewicz, A., Kemkemer, R., Derby, B., Spatz, J., Ballestrem, C., 2013. Vinculin regulates the recruitment and release of core focal adhesion proteins in a force-dependent manner. Curr. Biol. 23, 271–281. Chandrasekar, I., Stradal, T.E., Holt, M.R., Entschladen, F., Jockusch, B.M., Ziegler, W.H., 2005. Vinculin acts as a sensor in lipid regulation of adhesion-site turnover. J. Cell Sci. 118, 1461–1472. Chen, H., Chandrasekar, S., Sheetz, M.P., Stossel, T.P., Nakamura, F., Yan, J., 2013. Mechanical perturbation of filamin A immunoglobulin repeats 20–21 reveals potential non-equilibrium mechanochemical partner binding function. Sci. Rep. 3, 1642.

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