RESEARCH ARTICLE

An amphipathic helix of vinexin α is necessary for a substratestiffness-dependent conformational change in vinculinNaoya Hino1,2, Takafumi Ichikawa1,*, Yasuhisa Kimura1, Michiyuki Matsuda2, Kazumitsu Ueda1,3

and Noriyuki Kioka1,3,‡

ABSTRACTExtracellular matrix (ECM) stiffness regulates various cell behaviors,including cell differentiation, proliferation and migration. Vinculinand vinexin α (an isoform encoded by the SORBS3 gene), both ofwhich localize to focal adhesions, cooperatively function asmechanosensors of ECM stiffness. On a rigid ECM, vinexin αinteracts with vinculin and induces a conformational change invinculin to give an ‘open’ form, which promotes nuclear localizationof Yes-associated protein (YAP, also known as YAP1) andtranscriptional coactivator with a PDZ-binding motif (TAZ, also knownas WWTR1) (hereafter YAP/TAZ). However, the detailed mechanismby which vinexin α induces the conformational change in vinculin hasnot been revealed.Here, we identify an amphipathic helix namedH2asa novel vinculin-binding site in vinexin α. The H2 helix interacts with thevinculin D1b subdomain and promotes the formation of a talin–vinculin–vinexin α ternary complex. Mutations in the H2 region not onlyimpair the ability of vinexin α to induce the ECM stiffness-dependentconformational change in vinculin but also to promote nuclearlocalization of YAP/TAZ on rigid ECM. Taken together, these resultsdemonstrate that the H2 helix in vinexin α plays a critical role in ECMstiffness-dependent regulation of vinculin and cell behaviors.

KEY WORDS: Vinexin, Vinculin, Focal adhesion, Matrix stiffness,Mechanotransduction

INTRODUCTIONExtracellular matrix (ECM) stiffness regulates various cellularbehaviors, including cell proliferation (Klein et al., 2009; Wanget al., 2000), differentiation (Engler et al., 2006) and migration (Loet al., 2000; Nagasato et al., 2017; Yamashita et al., 2014). Inaddition, some types of tumor cells generate stiffened ECM, whichpromotes cell proliferation and metastasis (Levental et al., 2009;Provenzano et al., 2009). Therefore, understanding how cells senseECM stiffness is important for tissue engineering and cancer therapy.Focal adhesions (FAs), integrin-based protein complexes formed

at cell–ECM contact sites, play critical roles in sensing ECMstiffness. FAs function as mechanical linkages between ECM and

force-generating actomyosin, and, thus, forces are exerted on FAs.Because stiff ECM is less deformable than soft ECM, greater forcesare thought to be loaded on FAs on rigid ECM (Sun et al., 2016).ECM stiffness-dependent force seems to cause a conformationalchange in mechanosensors localized at FAs, which translateinformation from physical cues into biochemical signals and geneexpression.

Vinculin, an FA scaffold protein, plays a crucial role in sensingECM stiffness due to its direct binding to the actin cytoskeleton(Hirata et al., 2014; Omachi et al., 2017; Thievessen et al., 2013;Yamashita et al., 2014). Vinculin is composed of an N-terminalhead region and a C-terminal tail region connected by a proline-richflexible linker region (Bakolitsa et al., 2004; Borgon et al., 2004)and interacts with various proteins, including talin (TLN1 andTLN2 in mammals), through the head region, and F-actin throughthe tail region (Gilmore et al., 1993; Johnson and Craig, 1995a). Thehead–tail intramolecular interaction keeps vinculin ‘closed’ andinhibits its interaction with F-actin. Once one of the bindingpartners, such as talin, α-catenin and α-actinin, binds to the vinculinhead domain through its amphipathic α-helix, the conformation ofvinculin changes into an ‘open’ form that has a higher affinity forF-actin (Bois et al., 2005; Izard et al., 2004; Peng et al., 2012). Inaddition, rigid ECM as well as cytoskeletal forces are considered tobe required for the vinculin conformational change (Carisey et al.,2013; Humphries et al., 2007; Liu et al., 2016; Omachi et al., 2017;Yamashita et al., 2014). A molecular dynamics simulation has alsoindicated that external forces promote the conformational change invinculin (Golji and Mofrad, 2013; Sun et al., 2017).

Vinexin α (an isoform encoded by the SORBS3 gene), anotherFA scaffold protein, cooperates with vinculin to work as amechanosensor of ECM stiffness (Ichikawa et al., 2017; Kurodaet al., 2017; Yamashita et al., 2014). On rigid ECM, vinexin αinteracts with vinculin in an actomyosin contractility-dependentmanner and induces the conformational change in vinculin to anopen form, promoting cell migration and the nuclear localization ofthe transcriptional co-activators, Yes-associated protein (YAP, alsoknown as YAP1) and transcriptional coactivator with a PDZ-binding motif (TAZ, also known as WWTR1) (hereafter YAP/TAZ). However, the detailed mechanism by which vinexin αinduces the conformational change in vinculin on rigid ECM is stillunclear. Interestingly, vinexin β, a splice variant that lacks theN-terminal half of vinexin α, interacts with vinculin through thesame Src homology 3 (SH3) domains as vinexin α (Kioka et al.,1999) but does not induce the ECM stiffness-dependent behaviorsof vinculin (Yamashita et al., 2014). This indicates that the SH3domains of vinexin α are not sufficient to induce the conformationalchange in vinculin and suggests that additional molecularmechanisms underlie the vinculin–vinexin α interaction.

Talin, an FA protein, has also been shown to play a crucial rolein mechanosensing (Austen et al., 2015; Elosegui-Artola et al.,Received 29 March 2018; Accepted 3 December 2018

1Division of Applied Life Sciences, Graduate School of Agriculture, KyotoUniversity, Sakyo-ku, Kyoto 606-8502, Japan. 2Laboratory of Bioimaging andCell Signaling, Graduate School of Biostudies, Kyoto University, Sakyo-ku,Kyoto 606-8501, Japan. 3Institute for Integrated Cell-Material Sciences (iCeMS),Kyoto University, Sakyo-ku, Kyoto 606-8507, Japan.*Present address: Developmental Biology Unit, European Molecular BiologyLaboratory (EMBL), Heidelberg 69117, Germany.

‡Author for correspondence (nkioka@kais.kyoto-u.ac.jp)

N.H., 0000-0003-0190-3386; T.I., 0000-0002-0490-4176; N.K., 0000-0002-2708-537X

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2016). Talin interacts with the cytoplasmic tail of integrins throughits N-terminal head domain (Calderwood et al., 1999) and withF-actin through its C-terminal rod domain (Hemmings et al., 1996)and therefore is subject to mechanical forces. Mechanical forceloading on talin induces a conformational change in helix bundlesin the rod domains, resulting in exposure of cryptic vinculin-binding sites (VBSs) and recruitment of vinculin to FAs(Ciobanasu et al., 2014; del Rio et al., 2009; Hirata et al., 2014).Therefore, the talin–vinculin interaction also functions as a sensorfor ECM stiffness. However, the relationship between talin–vinculin and vinculin–vinexin α mechanosensing systems ispoorly understood.Here, we identify an amphipathic helix named H2 in the

N-terminal region of vinexin α as a novel vinculin-bindingregion. H2 helix interacts with the vinculin D1b subdomain andpromotes the formation of a talin–vinculin–vinexin α ternarycomplex. Mutations in the H2 region not only impair the ability ofvinexin α to induce the ECM stiffness-dependent conformationalchange in vinculin but also diminish nuclear localization of YAP/TAZ on rigid ECM. However, the H2 region alone does not inducethe conformational change in vinculin. Taken together, these resultsindicate that vinexin α interacts with vinculin through both its SH3

domains and H2 helix and induces the vinculin conformationalchange, which plays a pivotal role in sensing ECM stiffness.

RESULTSThe amphipathic helix H2 in vinexin α is necessary forhigh-affinity binding to vinculinVinexin α, but not vinexin β, is necessary for the ECM-stiffnessdependent behavior of vinculin (Yamashita et al., 2014). Thisobservation indicates that an N-terminal region of vinexin α, whichis not included in vinexin β, plays a critical role in regulation ofvinculin behavior (Fig. 1A). However, the molecular structure andthe function of the N-terminal region is poorly elucidated. To furtheranalyze the importance of this region, secondary structureprediction was performed using the program PORTER (Pollastriand McLysaght, 2005). Although most of the region was predictedto be random coils or disordered regions, a few regions werepredicted to form α-helices (Fig. S1A). Since vinculin bindingpartners, such as talin, α-actinin and IpaA, interact with vinculinthrough their amphipathic helices to induce the conformationalchange in vinculin (Bois et al., 2005; Izard et al., 2004, 2006), wefocused on two regions named predicted Helix1 (H1) (residues193–214) and predicted Helix2 (H2) (residues 329–355), which

Fig. 1. Amphipathic helix in vinexin α is necessary for high-affinity interaction with vinculin. (A) Schematic representations of the domain structure ofvinexin α and β. (B) Helical wheel models of predicted amphipathic helices H1 (amino acids 193–214) and H2 (amino acid 329–355). The models were createdusing Helical Wheel Projections (http://rzlab.ucr.edu/scripts/wheel/wheel.cgi) and modified. The filled and open diamonds indicate hydrophilic and hydrophobicresidues, respectively. The gray diamonds show intermediate residues. (C) Amino acid sequences of five vinexin α mutants (H1Pa, H1Pb, H2Pa, H2Pb andH2Pc) used in this study. The residues in the open squares were replaced with proline residues (shown as filled squares), and the name of the mutants are shownbelow. (D) Pulldown assays using purified GST–vinculin (0.14 µM). GST-tagged proteins and co-precipitated GFP–vinexin αΔN or vinexin β were analyzed byimmunoblotting using anti-GST and anti-GFP antibodies, respectively. The result is representative of three independent experiments. (E) Co-precipitated GFP-tagged proteins were quantified using ImageJ. The bars represent the mean+s.e.m. (n=3). *P<0.05 comparedwith GFP–vinexin αΔNWT (Tukey’s post-hoc test).

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were predicted to form amphipathic helices (Fig. 1A,B). To reveal arole for these amphipathic helices in the interaction with vinculin,we constructed five vinexin α mutants (H1Pa, H1Pb, H2Pa, H2Pb,and H2Pc) in which hydrophobic residues in the H1 region (H1Paand H1Pb) or the H2 region (H2Pa, H2Pb, and H2Pc) were replacedwith proline residues to disrupt the secondary structure (Fig. 1C).Because GST-tagged full-length vinexin α is highly degraded inE. coli, as described previously (Yamashita et al., 2014), truncatedmutant vinexin αΔN, which lacks residues 1–156 of vinexin α, andαΔN mutants having mutations in H1 or H2 helices were used toperform pulldown assays. Vinexin αΔN wild type (WT) co-precipitated with vinculin more efficiently than vinexin β (Fig. 1D,E), suggesting a possible binding region in the N-terminal region ofvinexin α. Interestingly, the H2Pa, H2Pb and H2Pc mutationsdramatically decreased the amount of co-precipitated vinculin to alevel comparable with that obtained with vinexin β. By contrast, theH1Pa and H1Pb mutations did not affect the co-precipitation. Theseresults clearly indicate that the H2 region is required for the high-affinity interaction between vinexin α and vinculin.To examine whether the H2 region can adopt an α-helical

conformation, the secondary structure of a chemically synthesizedH2 peptide (residues 329–355) was analyzed by circular dichroism(CD) spectroscopy. As shown in Fig. S1B, the H2 peptide exhibiteda spectrum pattern similar to that of synthesized talin VBS3 peptide(residues 1945–1970), which has already been revealed to form

an amphipathic helix (Izard et al., 2004). These peptides showedparticular negative peaks near 200 and 220 nm, which can beattributed to the superposition of negative peaks derived from atypical α-helix at 210 and 220 nm and from a random coil at 200 nm(Greenfield and Fasman, 1969). Therefore, this result suggests thatthe H2 region can adopt an α-helical conformation that is similar tothe talin VBS3 region.

Hydrophobic residues of H2 helix are necessary for directbinding to vinculinAlignment of vinculin-binding amphipathic helices among talin,α-actinin and α-catenin showed the conserved hydrophobic residuesin these helices, which are involved in binding to vinculin (Boiset al., 2005; Izard et al., 2004; Peng et al., 2012), and H2 helixincludes similar hydrophobic residues (Fig. 2A). Therefore, toinvestigate the role of the hydrophobic residues in H2 helix in theinteraction with vinculin, we constructed additional vinexin αmutants (H2Q1, H2Q2, H2Q3 and H2Q4) whose hydrophobicresidues were replaced with glutamine residues to impair thehydrophobic properties (Fig. 2B). GFP-tagged vinexin mutantsexpressed in 293T cells were pulled down using GST–vinculin. Asshown in Fig. 2C, GFP–vinexin α H2Q2 and H2Q3 mutants co-precipitated with GST–vinculin much less than GFP–vinexin αWT.The H2Q1 and H2Q4 mutant showed a modest effect on the co-precipitation. To further confirm the effect of these mutations on

Fig. 2. H2 helix of vinexinα interactswith the vinculin head. (A) Alignment of amino acid sequences of H2helix with vinculin-binding sites in known vinculin head-binding proteins. Hydrophobic residues are indicated by filled squares. (B) Amino acid sequences of vinexin α mutants (H2Q1, H2Q2, H2Q3 and H2Q4). Thehydrophobic residues shown in open squares were replaced with glutamine residues (indicated by filled squares), and the name of the mutants are shown below.(C) Pulldown assay using purified GST–vinculin (0.34 µM). Lysates fromHEK293T cells transfected with GFP-tagged vinexin αmutants or vinexin βwere subjectedto pulldown assays. GST-tagged proteins and co-precipitated GFP-tagged proteins were analyzed by immunoblotting using anti-GST and anti-GFP antibodies,respectively. The result is representative of two independent experiments. (D) GFP depletion assay using purified GST–vinculin (0.34 µM). Lysates fromHEK293T cells transfectedwithGFP-tagged vinexin αmutants or vinexin βwere subjected to aGFPdepletion assay. The fraction ofGFP-tagged proteins that boundto vinculin was determined by calculating the reduction in GFP fluorescence of the supernatant incubated with GST–vinculin compared with that of the supernatantincubated with GST. The values represent the mean+s.e.m. from three independent experiments. *P<0.05 compared with GFP–vinexin α WT (Tukey’s post-hoctest). (E) Pulldown assay using purified GST–H2 (5 µM), GST–H2-H2Q3 mutant (5 µM), and GST–vinexin αΔN (0.2 µM). Lysates from HEK293T cells transfectedwith GFP-tagged vinculin were subjected to pulldown assays. GST-tagged proteins and co-precipitated GFP-tagged proteins were analyzed by immunoblottingusing anti-GST and anti-GFP antibodies, respectively. The result is representative of three independent experiments.

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vinculin binding, the depletion of GFP fluorescence in thesupernatant was determined to analyze the fraction of the GFP-tagged vinexin mutants that bound to vinculin (Fig. 2D). Consistentwith the pulldown assay, the H2Q2 and H2Q3 mutationssignificantly decreased the bound fraction to a level that wascomparable with that found for vinexin β. These results suggest thatthe hydrophobicity of H2 helix is necessary for the high-affinityinteraction between vinexin α and vinculin.To investigate whether the H2 helix directly interacts with vinculin

or allosterically modulates the binding of vinexin α to vinculin, wetested whether the H2 helix itself has a vinculin-binding ability. TheH2 helix (residues 329–355) was purified as a GST-tagged protein,and vinculin binding was testedwith a pulldown assay. GFP–vinculinwas co-precipitated with the GST–vinexin α H2 helix, whileintroduction of the H2Q3 mutation into GST–vinexin α H2 helixclearly impaired the co-precipitation (Fig. 2E). This result indicatesthat H2 helix directly interacts with vinculin.

H2 helix interacts with the vinculin D1b helix bundlePrevious studies have shown that the SH3 domains of vinexin αinteract with the vinculin linker region (Kioka et al., 1999). Toidentify the H2 helix-binding site in vinculin, we first split vinculininto two parts, the N-terminal half containing a head region(residues 1–850) (head) and the C-terminal half containing aproline-rich linker and a tail region (836–1066) (linker-tail), andthen performed pulldown assays using these constructs (Fig. 3A,B).GST–vinexin α H2 helix pulled down the head region (1–850) butnot the linker-tail region (836–1066). Introducing the H2Q3mutation into GST–vinexin α H2 helix specifically disrupted the

co-precipitation with the head region (1–850). In contrast, GST–vinexin αΔN WT, which includes the SH3 domains, was co-precipitated with both constructs (Fig. 3B). These results indicatethat the head region contains an H2 helix-binding site.

The vinculin head region is composed of four helix bundles,called D1 to D4 domains (Fig. 3A) (Bakolitsa et al., 2004). Thus, wenext investigated which domain in the head region interacts with H2helix. Both GST–vinexin αΔN WT and GST–vinexin α H2 helixwere co-precipitated with the vinculin D1 domain (1–258) moreefficiently than with the other domains (Fig. S1C). H2Q3 mutationof vinexin αΔNmarkedly decreased the co-precipitation with the D1domain (1–258). Taken together, these findings show that vinexin αinteracts with the vinculin D1 domain through the H2 helix, inaddition to the interaction with the vinculin linker region throughthe SH3 domains.

The D1 domain consists of two helix bundles, the D1a and D1bsubdomains, each of which contains four helices (Bakolitsa et al.,2004). Talin VBS3 and most other amphipathic helices that interactwith and activate vinculin bind to the D1a subdomain (Izard et al.,2004). Replacement of the alanine at residue 50 in the D1asubdomain with an isoleucine residue (A50I) disturbs theinteraction with talin VBS3 (Bakolitsa et al., 2004). To examinewhether H2 helix interacts with the D1 domain in a manner similarto talin VBS3, we performed pulldown assays using a GFP–vinculinA50I mutant (Fig. 3C,D). The A50I mutation decreased the co-precipitation with talin VBS3, as expected. In contrast, the A50Imutation did not affect the co-precipitation with H2 helix. GST–vinexin α H2 helix having H2Q3 mutation did not co-precipitatewith either wild-type vinculin or the A50I mutant. These results

Fig. 3. H2 helix of vinexin α interacts with the vinculin D1b subdomain. (A) Schematic representations of full-length (FL) and truncated mutants of vinculin.(B–E) Pulldown assays using purified GST–H2 (3 µM), GST–H2-H2Q3 (3 µM), GST–vinexin αΔN (0.3 µM) or GST–talin VBS3 (3 µM). Lysates from HEK293Tcells transfected with GFP-tagged vinculin head or linker-tail (B), GFP-tagged vinculinWTor A50I mutant (C,D), andGFP-taggedD1 or D1b (E) were subjected topulldown assays. GST-tagged proteins and co-precipitated GFP-tagged proteins were analyzed by immunoblotting using anti-GST and anti-GFP antibodies,respectively. The results are representative of two independent experiments. (D) The amount of co-precipitated GFP–vinculin from experiments in C wasquantified using ImageJ. The bars represent the mean+s.e.m. (n=3). The P-values were calculated with a Tukey’s post-hoc test (n.s., not significant).

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suggest that the H2-binding site of vinculin is different from thetalin VBS3-binding site. To verify this observation, GFP–D1a(1–130) and GFP–D1b (127–252) helix bundles were constructed.However, GFP–D1a was co-precipitated with GST alone non-specifically, possibly due to improper folding of D1a (data notshown). Therefore, we focused on the GFP–D1 and GFP–D1bconstructs. GST–talin VBS3 co-precipitated with GFP–D1 (1–258)but not GFP–D1b (127–252), while GST–vinexin α H2 helix, aswell as GST–vinexin αΔN, co-precipitated with both GFP–D1 (1–258) and GFP–D1b (127–252) (Fig. 3E). These results suggest thatthe H2 helix of vinexin α interacts with the D1b subdomain, whiletalin VBS3 binds to the D1a subdomain.

H2 helix and talin VBS3 cooperatively interact with thevinculin head regionThe above observation that vinculin binds to H2 helix of vinexin αand talin VBS3 through different subdomains raises the possibilitythat vinexin α and talin simultaneously interact with vinculin. Toinvestigate this possibility, the effect of the talin VBS on theinteraction of H2 helix with vinculin was examined. Interestingly,the addition of a chemically synthesized talin VBS3 peptideincreased the amount of full-length vinculin and the vinculin headregion (1–850) co-precipitated with GST–vinexin α H2 helix in adose-dependent manner (Fig. 4A). Triplicate experiments usingvarious concentrations of GST–vinexin α H2 helix furtherconfirmed the effect of talin VBS3 on the co-precipitation ofvinculin with the vinexin α H2 helix (Fig. S1D,E). These results

suggest that the interaction of talin with vinculin promotes thevinculin–vinexin α interaction.

We next examined whether vinexin α, talin and vinculin form aternary complex. Vinexin αWT, its H2Q3 mutant or vinexin β wasexpressed in vinexin-knockout (KO) mouse embryonic fibroblasts(MEFs) together with GFP–vinculin. Although the expression levelof vinexin αWT was similar to that of the vinexin α H2Q3 mutant,the expression level of vinexin β was much higher than that ofvinexin α (Fig. 4B). Co-immunoprecipitation assays using anti-vinexin antibody showed that talin, as well as GFP–vinculin, co-precipitated with the vinexin α H2Q3 mutant significantly less thanwith vinexin α WT (Fig. 4C,E,F). Talin also co-precipitatedsignificantly less with vinexin β. To determine whether vinexin αinteracts with talin directly or indirectly through the interaction withvinculin, we performed co-immunoprecipitation assays using GFP-or GFP–vinculin-expressing vinculin-KO MEFs. Talin co-precipitated with vinexin α in GFP–vinculin-expressing cells butnot in GFP-expressing cells (Fig. 4D). Taken together, these resultssuggest that vinexin α indirectly forms a complex with talin throughinteraction with vinculin, resulting in the formation of a ternarycomplex.

The vinexin α H2 helix is necessary for inducing theconformational change in vinculinVinexin α interacts with vinculin and induces a conformationalchange in vinculin to an open form in vitro (Yamashita et al., 2014).To determine whether the H2 helix is involved in the

Fig. 4. H2 helix and talin cooperatively interact with vinculin. (A) The cell lysates of HEK293T cells transfected with GFP-tagged full-length vinculin or thevinculin head domain were incubated with purified GST–H2 (1 µM) in the presence of the indicated concentration of chemically synthesized talin VBS3 peptide.GST-tagged proteins and co-precipitated GFP-tagged proteins were analyzed by immunoblotting using anti-GST and anti-GFP antibodies, respectively. Theresult is representative of three independent experiments. (B) Vinexin-KO MEF cells re-expressing vinexins and expressing GFP–vinculin were lysed and thensubjected to immunoblotting using anti-vinculin, -vinexin and -β-tubulin antibodies. (C,D) Vinexin-KO MEF cells re-expressing vinexins (C) or vinculin-KO MEFcells re-expressing vinculin (D) were lysed and subjected to immunoprecipitation using anti-vinexin antibody. Vinexin and co-precipitated proteins were detectedby immunoblotting using the indicated antibodies. The results are representative of three independent experiments. (E,F) The levels of co-precipitated talin (E)and GFP–vinculin (F) from experiments as in C were quantified using ImageJ. The bars represent the mean+s.e.m. of three independent experiments. n=3. TheP-values were calculated with a Tukey’s post hoc test.

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conformational change, we assessed the effect of H2 helix onvinculin conformation using a purified vinculin fluorescenceresonance energy transfer (FRET) conformation probe that showsa high FRET ratio in a closed form compared with an open form(Chen et al., 2005). Consistent with a previous report (Yamashitaet al., 2014), the addition of GST–vinexin αΔN WT decreased theFRET ratio in a concentration-dependent manner (Fig. 5A). Thevinexin αΔN H2Q3 mutant did not decrease the FRET ratio, whichis comparable to what was seen with vinexin β, which lacks theN-terminal region of vinexin α, including H2 helix (Fig. 5A). Theseresults suggest that the H2 helix of vinexin α is necessary forinducing the conformational change in vinculin.In the open form, vinculin has a higher affinity for F-actin than in

the closed form, and thus, we also performed F-actin co-sedimentation assays to examine the effect of the H2 helix onvinculin affinity for F-actin. Consistent with the result from thein vitro FRET assay, GST–vinexin α WT increased the amount ofvinculin that co-sedimented with F-actin in a concentration-dependent manner (Fig. 5B,C). The H2Q3 mutant, as well asvinexin β, co-sedimented significantly less vinculin (Fig. 5B,C).These results support the notion that H2 helix of vinexin α plays acritical role in inducing the conformational change in vinculin.Next, we investigated whether the H2 helix is sufficient to induce

the conformational change in vinculin. Vinculin FRET analysisshowed that GST–vinexin αH2 helix did not change the FRET ratioeven at high concentrations, while GST–talin VBS3 decreased theFRET ratio in a concentration-dependent manner (Fig. 5D). TheF-actin co-sedimentation assay showed results consistent with this

observation (Fig. 5E,F). GST–talin VBS3 increased the amount ofvinculin that co-sedimented with F-actin in a concentration-dependent manner, whereas GST–vinexin α H2 helix did notaffect the amount of co-sedimented vinculin (Fig. 5E,F). Takentogether, these results indicate that H2 helix of vinexin α isnecessary but not sufficient for induction of the conformationalchange in vinculin.

The vinexin α H2 helix is necessary for the ECM stiffness-dependent conformational change in vinculin in MEFsTo examine the function of H2 helix in vivo, we first investigated thesubcellular localization of vinexin proteins and their effect onvinculin recruitment to FAs on glass cover slips. The vinexin αH2Q3mutant was predominantly localized to FAs, similar to vinexin αWT(Fig. 6A). Re-expression of vinexin αWT led to 1.8-fold and 1.4-foldincreases in the intensity of GFP–vinculin at FAs and the number ofGFP-vinculin-positive FAs, respectively, compared with mock cellsunder this experimental condition (Fig. 6B,C). Re-expression ofvinexin β showed a moderate effect. In contrast, expression of thevinexin α H2Q3 mutant did not increase either GFP–vinculinintensity in FAs or the number of FAs, suggesting that the H2 helixpromotes the recruitment of GFP–vinculin to FAs (Fig. 6B,C). Thedifference between vinexin α H2Q3 and vinexin β is possibly due tomuch higher expression of vinexin β than that of vinexin α WT orvinexin α H2Q3 (see Fig. 4B,C).

Next, to investigate the effect of the H2 helix on the ECM-stiffnessdependent localization of vinculin to FAs, vinexin-KO MEFs re-expressing different forms of vinexins were seeded on soft (3.8 kPa)

Fig. 5. H2 helix is necessary for the conformational change in vinculin induced by vinexin α. (A,D) Purified vinculin FRET probe (0.1 µM) was incubatedwith the indicated amount of purified GST-fused proteins in the presence of pre-polymerized F-actin (4.6 µM). The fluorescent intensity of CFP and FRET wereacquired with a SpectraMax M2e (A) or a Cytation 5 imaging plate reader (D). The ratio of cFRET to CFP signals was calculated. The values represent the mean±s.e.m. of three independent experiments. (B,E) F-actin co-sedimentation assay with the vinculin FRET probe. The purified vinculin FRET probe (0.1 µM)incubated as described in Awas ultracentrifuged at 150,000 g for 1 h. The pellets and the supernatants were subjected to SDS-PAGE, followed by CBB staining.The results are representative of three independent experiments. (C,F) Quantification of the amount of co-precipitated vinculin FRET probe in the F-actin co-sedimentation assay. The bars represent the mean+s.e.m. of three independent experiments. The P-values were calculated with a Tukey’s post-hoc test.

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and rigid (25 kPa) polyacrylamide (PAA) gel substrates and thensubjected to confocal microscopy (Fig. 7A,B; Fig. S2A). In vinexin αWT-re-expressing cells, both the intensity and the number of GFP–vinculin-positive FAs were increased on the rigid substratescompared with on the soft substrates under this experimentalcondition. Although mock and vinexin α H2Q3-re-expressing cellssomewhat retained the ECM stiffness dependency, both the intensityand the number of GFP–vinculin-positive FAs on the rigid substrateswere significantly decreased compared with vinexin α WT-re-expressing cells. Vinexin β-re-expressing cells did not show anyECM stiffness-dependency. Therefore, these results show that the H2helix promotes accumulation of GFP–vinculin to FAs on rigid ECM.A closed form of vinculin localized to FAs is readily washed out

by treatment with CSK buffer containing Triton X-100, whereas anopen form of vinculin is resistant to CSK treatment, mainly becauseof its tight interaction with F-actin (Omachi et al., 2017; Yamashitaet al., 2014). Our previous study showed that vinexin α, but notvinexin β, increases the amount of CSK-resistant vinculin in cells onrigid ECM. To investigate whether H2 helix is involved in thisincrease, vinexin-KO MEFs re-expressing different forms ofvinexins were seeded on PAA gel substrates with different levelsof stiffness, and subjected to CSK treatment. As expected, invinexin α WT-re-expressing cells, the GFP–vinculin intensity atFAs and the number of CSK-resistant vinculin-positive FAs were

8.4-fold and 4.7-fold higher, respectively, on the rigid substrates(25 kPa) compared with on the soft substrates (3.8 kPa) (Fig. 7C,D;Fig. S2B). By contrast, CSK-resistant vinculin-positive FAs inmock, vinexin α H2Q3 and vinexin β-re-expressing cells culturedon either the soft or rigid substrate were barely visible, and there wasno significant difference between cells cultured on soft and rigidsubstrates (Fig. 7C,D; Fig. S2B). Combined with our previousreport showing that rigid ECM promotes the interaction of vinexin αwith vinculin compared to soft ECM, these results suggest that H2helix of vinexin α is necessary for the ECM stiffness-dependentconformational change in vinculin in MEFs.

The vinexin α H2 helix is involved in stiffness-dependentregulation of cell behaviorsNuclear localization of YAP/TAZ is promoted on rigid ECM inmesenchymal stem cells in a manner dependent on vinexin andvinculin expression (Kuroda et al., 2018, 2017). To explore the roleof H2 helix in the nuclear localization of YAP/TAZ, vinexin-KOMEFs re-expressing vinexin αWTor H2Q3 cultured on soft or rigidsubstrates were immunostained with anti-YAP/TAZ antibody(Fig. 8A,B). On soft substrates, there was no significant differencein nuclear localization of YAP/TAZ between mock cells andvinexin-KO MEFs re-expressing vinexin α WT, H2Q3 mutant orvinexin β. In contrast, on rigid substrates, vinexin α WT-re-

Fig. 6. H2 helix promotes the formation of vinculin-rich focal adhesions. (A) Immunofluorescence detected using an anti-vinexin antibody in vinexin-KOMEFcells re-expressing vinexins and GFP–vinculin. The FA-containing areas indicated by open boxes are shown at a higher magnification in the lower rightof each panel. Scale bar: 30 µm. (B,C) Quantification of total signal intensity of GFP–vinculin (B) and the number of GFP–vinculin-positive FAs (C). Black linesrepresent the mean values for n=30. The P-values were calculated with a Tukey’s post-hoc test.

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expressing cells showed more prominent nuclear localization ofYAP/TAZ than mock, vinexin α H2Q3- or vinexin β-re-expressingcells. These results indicate that vinexin α H2 helix promotes YAP/TAZ nuclear localization on rigid substrates.

DISCUSSIONIn this study, we identified a novel vinculin-binding site in vinexinα, the H2 helix, which forms an amphipathic helix structure. The H2helix interacts with the vinculin D1b subdomain, which is includedin the vinculin head region, and the disruption of the H2 helix resultsin impairment of the conformational change in vinculin and thenuclear localization of YAP/TAZ on rigid ECM. However, the H2helix alone does not induce the conformational change in vinculin.Similarly, the SH3 domains of vinexin α are required but notsufficient to induce the conformational change in vinculin(Yamashita et al., 2014). Taken together, these observationssuggest that, on rigid ECM, vinexin α interacts with vinculinthrough both the H2 helix and SH3 domains to induce theconformational change of vinculin and nuclear translocation ofYAP/TAZ (Fig. 8C), and that the interaction regulates behaviors ofvinculin that act as a sensor for ECM stiffness.

Amphipathic helices are known to associate with vinculin in twodifferent ways, via a ‘helical bundle conversion’ and ‘a helixaddition mechanism’. Amphipathic helices present in talin, α-actinin, or IpaA bind the vinculin D1a subdomain and inducedynamic remodeling of the structure of the vinculin D1 domain,leading to the conformational change in vinculin to an open form(Bois et al., 2005; Izard et al., 2004, 2006), a process called helicalbundle conversion. A few amphipathic helices, including IpaAVBS2, are known to interact with the vinculin D1b subdomain(Nhieu and Izard, 2007). These interactions induce neither thedynamic remodeling of the structure of the vinculin D1 domain northe conformational change in vinculin, a process called a helixaddition mechanism. In this study, we showed that H2 helix ofvinexin α interacts with the vinculin D1b subdomain and does notinduce the conformational change in vinculin by itself.Interestingly, the H2 helix shows significant similarity to IpaAVBS2 (563–587) (Fig. S1F). Thus, it can be speculated that H2helix interacts with vinculin through a helix addition mechanism. Itwill be of interest to examine this possibility in the future.

Because both the H2 helix and the SH3 domains of vinexin α arenecessary for induction of the conformational change in vinculin,

Fig. 7. H2 helix is involved in ECMstiffness-dependent vinculin behaviors. (A) Immunostaining of vinexin-KOMEF cells re-expressing vinexin seeded on soft(3.8 kPa) and rigid (25 kPa) PAA gels. To improve visibility, the gray images were inverted. Scale bar: 30 µm. (B) Quantification of total GFP–vinculin intensity.Black lines represent the mean values; n=45 in all samples except vinexin β on rigid PAA gels (n=35). The P-values were calculated with a Tukey’s post-hoc test.(C) Immunostaining of CSK buffer-treated vinexin-KO MEF cells re-expressing vinexin seeded on soft (3.8 kPa) and rigid (25 kPa) PAA gels. To improvevisibility, the gray imageswere inverted. Scale bar: 30 µm. (D) Quantification of CSK-resistant GFP-vinculin intensity. Black lines represent themean values; n=60(mock; soft), n=52 (mock; rigid), n=60 (vinexin αWT; soft), n=45 (vinexin αWT; rigid), n=60 (vinexin α H2Q3; soft), n=53 (vinexin α H2Q3; rigid), n=49 (vinexin β;soft), and n=60 (vinexin β; rigid). The P-values were calculated with a Tukey’s post-hoc test.

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the simultaneous interaction of H2 helix and the SH3 domains withvinculin is required for induction of the conformational change.Owing to the flexibility of the proline-rich linker region to which thefirst and second SH3 domains of vinexin bind, the distance betweenthe linker region and D1b subdomain is roughly estimated to be50 Å, as determined from the crystal structures of vinculin(Bakolitsa et al., 2004; Borgon et al., 2004; Nhieu and Izard,2007). The H2 helix and the first SH3 are 95 amino acids apart, mostof which are predicted to form disordered or random coil regions.The maximum length of one amino acid in an extended peptide is3.8 Å long (Erickson, 1994); thus 95 amino acids are enough lengthfor connecting a 50 Å distance, suggesting that vinexin α could bindvinculin through the H2 helix and SH3 domains simultaneously.Previous studies have indicated that molecules that induce theconformational change in vinculin interact with a single domain ofvinculin; talin, α-actinin, and α-catenin interact with the vinculinhead region, and phosphatidylinositol 4,5-bisphosphate (PIP2)interacts with the vinculin tail region (Bois et al., 2005; Izardet al., 2004; Johnson and Craig., 1995b; Peng et al., 2012). Thus, itis plausible that vinexin α induces a conformational change invinculin through a novel mechanism, in which interactions at twodifferent sites of vinculin are required.Mechanical forces induce stretching of the talin rod, leading to

exposure of cryptic VBSs and resulting in force-dependentrecruitment of vinculin to FAs (Ciobanasu et al., 2014; del Rio

et al., 2009; Hirata et al., 2014). This talin–vinculin interactionseems to play a crucial role in ECM stiffness sensing (Austen et al.,2015; Elosegui-Artola et al., 2016). Indeed, tension applied to talinis regulated by ECM stiffness (Austen et al., 2015; Kumar et al.,2016), especially in soft ranges (less than 4 kPa) in fibroblasts(Austen et al., 2015). The tensions detected in the Austen et al. studyare enough for unfolding the most unstable talin rod domain (R3)and exposing VBSs (Yao et al., 2014). However, cell responses toECM stiffness have a wide dynamic range, and traction forcegeneration increases as the ECM becomes stiffer from 3 kPa to52 kPa (Austen et al., 2015); thus, cells seem to have other systemsfor sensing ECM stiffness. In addition to the possibility that othertalin rod domains or the number of talin molecules subjected toforce contribute to sensing this range of ECM stiffness, our previousobservation indicates that amount of open-form vinculin isincreased depending on the interaction with vinexin α and theECM stiffness in a stiffness range of 3.8–43 kPa (Yamashita et al.,2014), implicating that the vinculin–vinexin α machinery is alsoinvolved in sensing of ECM stiffness. However, the interplaybetween the talin–vinculin and vinculin–vinexin αmechanosensorysystems has been never addressed. In the present study, we showedthat talin VBS3 promotes the interaction between vinculin and H2helix of vinexin α in a concentration-dependent manner, and talinand vinexin α can bind to vinculin simultaneously to form a ternarycomplex. Therefore, our observations suggest a collaboration

Fig. 8. H2 helix in vinexin α promotes YAP/TAZ nuclear localization on rigid ECM. (A) Immunostaining of vinexin-KOMEF cells re-expressing vinexin seededon soft (2.2 kPa) and rigid (43 kPa) PAA gels using antibody against YAP/TAZ and Hoechst 33342. Scale bar: 30 µm. (B) Quantification of the nuclear localizationratio of YAP/TAZ. Black lines represent the mean values; n=101 (mock; soft), n=102 (mock; rigid), n=106 (vinexin α WT; soft), n=95 (vinexin α WT; rigid),n=87 (vinexin αH2Q3; soft), n=127 (vinexin αH2Q3; rigid), n=84 (vinexin β; soft), and n=90 (vinexin β; rigid). TheP-valueswere calculatedwith a Tukey’s post-hoctest. (C) Model for the function of H2 helix in vinexin α.

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between the talin–vinculin and vinculin–vinexin αmachinery and amulti-tiered architecture of mechanosensory systems that allowscells to respond to a wide range of ECM stiffnesses.Vinexin, c-Cbl associated protein (CAP) and Arg-binding

protein2 (ArgBP2) constitute an adaptor protein family called thevinexin (SORBS) family and contain one sorbin homology (SoHo)domain and three SH3 domains in common (Kioka et al., 2002).Since our recent study indicated that CAP, as well as vinexin α,interacts with vinculin and induces the ECM stiffness-dependentconformational change in vinculin, it apparently seems that vinexinα and CAP are functionally redundant (Ichikawa et al., 2017).However, an alignment of amino acid sequences and secondarystructure predictions implies that an amphipathic helix, such as H2helix in vinexin α, is not conserved in the other vinexin familyproteins (data not shown). Therefore, it is likely that vinexin α andCAP interact with vinculin through different mechanisms and showdifferent responses to ECM stiffness. Indeed, vinexin α is involvedin differentiation of mesenchymal stem cells into myoblasts on 15kPa substrates, while CAP is involved in the differentiation intoosteoblasts on 42 kPa substrates (Holle et al., 2016). Thus, acomparative study of the molecular mechanism of the vinexin α–vinculin and CAP–vinculin interactions is needed.Collectively, we conclude that the H2 helix in the N-terminal

region of vinexin α plays a crucial role in the ECM stiffness-dependent regulation of vinculin and cell behaviors. Given that theexpression levels of vinexin in various tissues are correlated withthose of vinculin (Kioka et al., 1999), regulation of vinculinfunction by vinexin α would be likely to play critical roles. Becauseit has been revealed that vinculin is involved in a variety of cellfunctions that are dependent on ECM stiffness, such as stem celldifferentiation and tumor progression (Kuroda et al., 2017;Rubashkin et al., 2014), further studies concerning the role ofvinexin in cancer progression and cell differentiation might providenovel insights into cancer therapy and tissue engineering.

MATERIALS AND METHODSPlasmid constructionThe lentiviral transfer vector pCDH-EF1-IRES-Puro from SystemBiosciences (Mountain View, CA) was modified into pCDH-EF1-IRES-Blast as described previously (Ichikawa et al., 2017). The full-length vinexinα and β, as well as monomeric GFP-tagged full-length vinexin α, vinexinαΔN (157–733), and vinexin β were subcloned into pCDH-EF1-IRES-Blast. Monomeric GFP-tagged full-length chicken vinculin was insertedinto pCDH-EF1-IRES-Puro. Vinculin head (1–850), linker-tail (836–1066),D1–D4 (1–836), D1 (1–258), D2 (253–487), D3 (491–717), D4 (718–836),D1a (1–130) and D1b (127–252) were generated by PCR and thensubcloned into pCDH-EF1-IRES-Puro vector as a monomeric GFP-taggedform. The vinculin FRET conformation probe (Chen et al., 2005) wassubcloned into pColdI vector (Takara Bio, Ohtsu, Japan). Vinexin αΔN(157–733), H2 (328–355) and talin1 VBS3 (1945–1970) were generated byPCR and then subcloned into pColdI-ΔHis-GST vector. Vinculin was alsosubcloned into pColdI-ΔHis-GST vector. Vinexin α H1Pa (M199P; F200P;I203P), H1Pb (L210P; L212P), H2Pa (I329P; L332P; L333P), H2Pb (L337P;L340P; S341P; L344P), H2Pc (L348P; I351P; L355P), H2Q1 (I329Q;L332Q), H2Q2 (L333Q; L337Q), H2Q3 (L340Q; L344Q) and H2Q4(L348Q; I351Q; L355Q) were generated by site-directed mutagenesis usingan In-Fusion® HD cloning kit (Clontech, Mountain View, CA).

Antibodies and reagentsMouse monoclonal anti-vinculin (hVIN-1, cat. no. V9131, 1:20,000 dilutionfor immunoblotting, 1:400 dilution for immunofluorescence) and anti-talin(8d4, cat. no. T3287, 1:10,000 dilution for immunoblotting) antibodies werepurchased from Sigma-Aldrich (St Louis,MO).Mouse monoclonal anti-GSTantibody (5A7, cat. no. 013-21851, 1:5000 dilution for immunoblotting) was

purchased fromWako (Osaka, Japan). Mouse monoclonal anti-GFP (B-2, cat.no. sc-9996, 1:2000 dilution for immunoblotting) and anti-YAP (63.7, cat.no. sc-101199, 1:100 dilution for immunofluorescence) antibodies werepurchased from Santa Cruz Biotechnology (Dallas, TX). Rabbit polyclonalanti-vinexin antibody (1:10,000 dilution for immunoblotting, 1:400 dilutionfor immunofluorescence) was as described previously (Kioka et al., 1999).Goat anti-mouse IgGHRP conjugate (cat. no. 172-1011, 1:10,000 dilution forimmunoblotting) and goat anti-rabbit IgG-HRP conjugate (cat. no. 172-1019,1:10,000 dilution for immunoblotting) antibodies were from Bio-Rad(Hercules, CA). Alexa Fluor 546-conjugated goat anti-rabbit IgG antibody(1:1000 dilution for immunofluorescence) was fromThermo Fisher Scientific(Waltham, MA). Hoechst 33342 was from Invitrogen (Carlsbad, CA, 1:2000dilution for immunofluorescence). Type I collagen was from Nitta Gelatin(Osaka, Japan). Puromycin and Blasticidin S were from Sigma-Aldrich andKaken Pharmaceutical (Tokyo, Japan), respectively. Actin protein (>99%pure) purified from rabbit skeletal muscle was purchased from Cytoskeleton,Inc. (Denver, CO). Peptides of vinexin α H2 (329–355) and talin1 VBS3(1945–1970) were synthesized and purified by Toray Research Center (Otsu,Japan). Full size blots for appropriate figures are presented in Fig. S3.

Cell culture and lentiviral transductionSpontaneously immortalized WT and vinexin-KO MEFs were as describedpreviously (Kioka et al., 2010). Vinculin-KO MEFs were gifted from DrEileen Adamson (Burnham Institute, La Jolla, CA). Lentiviruses for proteinexpression were generated as described using lentiviral third-generationvectors [pCDH, pMD2.G (#12259), pRSV-Rev (#12253) and pMDLg/pRRE (#12251)] (Addgene, Cambridge, MA). The target cells wereincubated with lentivirus, followed by the incubation with mediumcontaining 1 µg/ml Puromycin or 3 µg/ml Blasticidin S for more than7 days. All cells were grown in Dulbecco’s modified Eagle’s medium(DMEM; Nacalai tesque, Kyoto, Japan) supplemented with 10% fetalbovine serum (FBS; Gibco/Thermo Fisher Scientific) at 37°C in ahumidified atmosphere containing 5% CO2. PAA gel substrates wereprepared as previously described (Yamashita et al., 2014), with slightmodifications; PAA gels were coated with 100 µg/ml type I collagen. 2.2and 3.8 kPa PAA gels were used as soft substrates and 25 and 43 kPa wereused as rigid substrates.

Immunofluorescence and confocal microscopyImmunofluorescence of FA proteins was performed as previously described(Ichikawa et al., 2017; Yamashita et al., 2014), with slight modifications.Briefly, vinexin-KO cells re-expressing vinexin together with GFP–vinculinwere cultured on 10 µg/ml type I collagen-coated No. 1 coverslips or 100 µg/ml type I collagen-coated PAA gels, and subjected to immunostaining. Tovisualize the total amount of vinculin or vinexin in cells on coverslips or PAAgels, cells were fixed with 2% (w/v) paraformaldehyde in PBS for 30 min atroom temperature and then permeabilized with 0.2% (v/v) Triton X-100 inPBS for 5 min at room temperature. To visualize CSK-resistant GFP–vinculinin cells on PAA gels, cells were treated with CSK buffer (0.5% Triton X-100,10 mMPIPES, pH 6.8, 50 mMNaCl, 3 mMMgCl2, 300 mM sucrose) on icefor 2 min, followed by fixation with 4% (w/v) paraformaldehyde in PBS for30 min at room temperature. For immunostaining, cells were incubated with10% goat serum (Sigma-Aldrich) and 0.01% (v/v) Triton X-100 in PBS for30 min at room temperature, followed by sequential incubation with primaryand secondary antibodies diluted with PBS containing 10% goat serum and0.01% (v/v) Triton X-100 for 1 h at room temperature or overnight at 4°C.Images were acquired using an LSM700 laser scanning confocal microscopewith a Plan-Apochromat 40x/1.3NAoil immersion objective lens (Carl Zeiss,Oberkochen, Germany). Image quantifications were performed using ImageJsoftware. Focal adhesions were classed as structures of 1–20 µm2 in the‘Analyze Particles’ command. Gray images were treated with ‘EnhanceContrast’ command and then inverted to increase their visibility.Immunostaining of YAP/TAZ was performed as described previously(Kuroda et al., 2018), with slight modifications. Briefly, vinexin-KO cellsre-expressing vinexin together with GFP–vinculin were cultured on 100 µg/ml type I collagen-coated PAA gels, and subjected to immunostaining. Thecells were fixed with 4% (w/v) paraformaldehyde in PBS for 15 min at roomtemperature and then permeabilized with 0.2% (v/v) Triton X-100 in PBS for

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5 min. The cells were further incubated with 10% goat serum in PBS for30 min, followed by sequential incubation with the anti-YAP antibody thatcrossreacts with TAZ (Kuroda et al., 2017) and second antibody diluted withPBS containing 10% goat serum for 1 h at room temperature or overnight at4°C. Images were acquired using an FLUOVIEW FV1000 laser scanningconfocal microscope (Olympus, Tokyo, Japan) equipped with an UPlanSApo 20×/0.75 NA dry objective lens (Olympus) with 3× zoom. To evaluateYAP/TAZnuclear localization, themean intensity of YAP/TAZ in the regionsof the nucleus and cytosol ( just outside the nucleus) with the same area wasanalyzed, followed by calculation of nucleus:cytosol intensity ratio.

Circular dichroism spectrometrySecondary structures of vinexin αH2 (329–355) and talin VBS3 (1945–1970)peptides were analyzed by circular dichroism spectrometry (J-805; JASCO,Tokyo, Japan) using a 0.1-mm quartz cell (121.027-QS, Q 10 mm; JASCO) asdescribed previously (Sato et al., 2013; Yamashita et al., 2014). Talin VBS3 orvinexin α H2 peptides (80 µM) dissolved in PBS containing 8% ethanol weresubjected to circular dichroism analysis. The obtained values were corrected bysubtracting the spectrum of PBS containing 8% ethanol alone.

Protein purificationPurifications of GST-tagged proteins were performed as describedpreviously (Takahashi et al., 2005; Yamashita et al., 2014). The His-tagged vinculin FRET conformation probe was expressed in E. coliRosetta™ (DE3) strain (Novagen/Merck Millipore, Darmstadt, Germany),and then E. coli were lysed in PBS with a XL2020 Sonicator (MISONIX,Farmingdale, NY). The protein was then purified with Profinity™ IMACNi-Charged Resin (Bio-rad), followed by anion exchange chromatographywith MonoQ 5/50 GL (GE Healthcare).

In vitro FRET assay and actin co-sedimentation assayThe in vitro FRET and actin co-sedimentation assays were performed asdescribed previously (Yamashita et al., 2014) with slight modifications.Briefly, the purified vinculin FRET conformation probe (final 0.1 µM) andGST-tagged protein were incubated with pre-polymerized F-actin (final4.6 µM) for 30 min at room temperature. The fluorescent intensity of CFPand FRET were acquired with a SpectraMax M2e (Molecular Devices, Inc.,Sunnyvale, CA) for the experiment using GST-tagged vinexin, and with aCytation 5™ imaging plate reader (BioTek Japan, Tokyo, Japan) for theexperiment using GST-tagged amphipathic helices (H2 and talin VBS3).CFP emission was detected at 478 nm with excitation at 436 nm, and FRETemission was detected at 526 nm with excitation at 436 nm. Spectralcrosstalk was corrected as described previously (Chen et al., 2005). In thisstudy, the values RY and RC were 0.048 and 0.402 in the measurement withSpectraMax M2e, and 0.059 and 0.494 with Cytation 5™ imaging platereader. The corrected FRET ratio (cFRET:CFP) was obtained as thecorrected FRET emission divided by the CFP emission. After themeasurement of fluorescence, the samples were subjected toultracentrifugation at 150,000 g for 1 h at 4°C for an actin co-sedimentation assay. The pellet and half of the supernatant were subjectedto SDS-PAGE, followed by Coomassie Brilliant Blue (CBB) staining.

ImmunoprecipitationThe immunoprecipitation assay was performed as described previously(Yamashita et al., 2014). Briefly, vinexin-KO cells re-expressing vinexintogether with GFP–vinculin, or vinculin-KOMEFs expressing GFP–vinculintogether with vinexin α were lysed with 1% (v/v) Triton X-100 in PBS. Thecell lysates (300 µg) were incubated with 1.5 µg anti-vinexin antibody at 4°Cfor 1 h, followed by further incubation with Protein G–Sepharose 4B beads(Sigma-Aldrich). The beads were collected and washed with 0.5% (v/v)Triton X-100 in PBS three times. Co-precipitated proteins were subjected toSDS-PAGE, followed by immunoblotting with specific antibodies.

Pulldown assay and GFP depletion assayThe pulldown and GFP depletion assays were performed as describedpreviously (Ichikawa et al., 2017) with slight modification. 293T cells weretransiently transfected with pCDH plasmids. At 24 h after transfection, the

cells were lysed with 1% (v/v) Triton X-100 in PBS. Cell lysates wereincubated with purified GST-tagged proteins at 4°C for 1 h followed byfurther incubation with glutathione–Sepharose 4B beads (GE Healthcare)for 1 h. The beads and the supernatant were separated by centrifugation (380g for 2 min). For the GFP depletion assay, the GFP fluorescence of thesupernatants was measured with a Cytation 5™ imaging plate reader.The fraction of bound GFP-tagged proteins were determined by calculatingthe reduction in GFP fluorescence of the supernatant incubated withGST–vinculin compared with that of the supernatant incubated with GST.For pulldown assay, the collected beads were washed with 0.5% (v/v) TritonX-100 in PBS three times. Co-precipitated proteins were subjected toSDS-PAGE, followed by immunoblotting with specific antibodies.

Statistical analysisData are presented as mean±s.e.m. Differences among compared groupswere assessed by one-way ANOVA followed by multiple Tukey’s post hoctests. P<0.05 was considered statistically significant. Statistical analysis wasperformed using Origin 8.6J or GraphPad Prism 7 software.

AcknowledgementsWe thank Dr K. Murakami and Dr K. Irie (Kyoto University) for advising us on CDspectroscopic analysis.

Competing interestsThe authors declare no competing or financial interests.

Author contributionsConceptualization: N.K.; Formal analysis: N.H., T.I., Y.K., M.M., K.U., N.K.;Investigation: N.H., T.I.; Writing - original draft: N.H.; Writing - review & editing: N.K.;Supervision: N.K.; Funding acquisition: K.U., N.K.

FundingThis work was supported in part by the Japan Society for the Promotion of ScienceGrants-in-Aid for Scientific Research (B) (JSPS KAKENHI grant numbers24380185, 18H02167 to N.K.), Grant-in-Aid for Challenging Exploratory Research(JSPS KAKENHI grant numbers 24658094, 26660291, 16K15090 to N.K.) and byGrants-in-Aid for JSPS Fellows [JSPS KAKENHI grant number 12J03633 (to T.I.)and 17J02107 (to N.H.)], the Ministry of Education, Culture, Sports, Science, andTechnology of Japan, Grant-in-Aid for Scientific Research (S) (25221203 to K.U.)and by a Grant-in-Aid for Scientific Research on Innovative Areas (MEXT KAKENHIgrant number 26112707 to N.K.), and by the Advanced Research and DevelopmentPrograms for Medical Innovation (N.K.) of the Japan Agency for Medical Researchand Development.

Supplementary informationSupplementary information available online athttp://jcs.biologists.org/lookup/doi/10.1242/jcs.217349.supplemental

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RESEARCH ARTICLE Journal of Cell Science (2019) 132, jcs217349. doi:10.1242/jcs.217349

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