1. Biochemistry and Chemical Biology
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Talin-activated vinculin interacts with branched actin networks to initiate bundles

  1. Rajaa Boujemaa-Paterski  Is a corresponding author
  2. Bruno Martins
  3. Matthias Eibauer
  4. Charlie T Beales
  5. Benjamin Geiger
  6. Ohad Medalia  Is a corresponding author
  1. Department of Biochemistry, University of Zurich, Switzerland
  2. Université Grenoble Alpes, France
  3. Department of Immunology, Weizmann Institute of Science, Israel
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Cite this article as: eLife 2020;9:e53990 doi: 10.7554/eLife.53990

Abstract

Vinculin plays a fundamental role in integrin-mediated cell adhesion. Activated by talin, it interacts with diverse adhesome components, enabling mechanical coupling between the actin cytoskeleton and the extracellular matrix. Here we studied the interactions of activated full-length vinculin with actin and the way it regulates the organization and dynamics of the Arp2/3 complex-mediated branched actin network. Through a combination of surface patterning and light microscopy experiments we show that vinculin can bundle dendritic actin networks through rapid binding and filament crosslinking. We show that vinculin promotes stable but flexible actin bundles having a mixed-polarity organization, as confirmed by cryo-electron tomography. Adhesion-like synthetic design of vinculin activation by surface-bound talin revealed that clustered vinculin can initiate and immobilize bundles from mobile Arp2/3-branched networks. Our results provide a molecular basis for coordinate actin bundle formation at nascent adhesions.

Introduction

Integrin-based cell adhesions mediate the interactions of the actin cytoskeleton with the extracellular matrix (ECM), enabling cell migration, proliferation and differentiation. Vinculin, a key adapter protein regulating cell adhesion signaling, contributes to tissue homeostasis, cell morphogenesis, immune processes, and wound healing (Carisey and Ballestrem, 2011). Composed of multiple ‘adhesome’ scaffolding and signaling components, integrin adhesions, for example focal adhesions (FAs), are associated with the termini of contractile actin and myosin-rich stress fibers (Geiger, 1979; Horton et al., 2016; Zaidel-Bar et al., 2007a). These assemblies develop in response to external or internal (mostly cytoskeletal) traction forces (Balaban et al., 2001; Geiger et al., 2009; Livne et al., 2014; Vicente-Manzanares et al., 2009). The mechanism whereby integrin activation leads to FA formation is rather complex, and involves a cascade of conformational transitions of key mechanoresponsive proteins such as talin, vinculin, and FA kinase (Burridge and Connell, 1983; Carisey et al., 2013; del Rio et al., 2009; Grashoff et al., 2010; Kong et al., 2009; Legate et al., 2011; Roberts and Critchley, 2009; Webb et al., 2004; Yang et al., 2016; Zamir and Geiger, 2001). During nascent adhesion formation, talin changes its conformation to enable integrin binding (Jiang et al., 2003; Wegener et al., 2007). Next, force-sensing proteins such as vinculin are recruited and activated through its tail (Burridge and Mangeat, 1984; del Rio et al., 2009; Hemmings et al., 1996; Hu et al., 2016), leading to a series of molecular interactions involving adhesome scaffolding and signaling components (Horton et al., 2016; Zaidel-Bar et al., 2007a). Furthermore, the association of vinculin with both talin and the actin cytoskeleton generates tension that, in turn, induces conformational changes in vinculin, further activating it (Grashoff et al., 2010). Thus, vinculin is thought to act as a molecular clutch that couples the actin network to nascent adhesions and FAs (Bachir et al., 2014; Case and Waterman, 2015; Choi et al., 2008; Thievessen et al., 2013).

Particularly intriguing is the mode of interaction of vinculin with the actin cytoskeleton. In motile cells, vinculin is believed to be involved in the coupling of the retrograde flow of the lamellipodial branched actin network to nascent integrin adhesions, which controls their initiation, maturation and disassembly dynamics (Case et al., 2015; Choi et al., 2008; Giannone et al., 2007; Oakes et al., 2012; Thievessen et al., 2013; Zaidel-Bar et al., 2007a). Vinculin was shown to be a mechanosensitive partner involved in FA maturation at the lamellipodium-lamellum border, where it responds to actomyosin-induced tension by increasing growth and robustness of FAs and their associated stress fibers (Balaban et al., 2001; Chrzanowska-Wodnicka and Burridge, 1996; Humphries et al., 2007). Vinculin is a component not only of nascent adhesions and FAs, but also of adherens-type cell-cell junctions, where it also acts as a mechanosensing component (Seddiki et al., 2018; Yonemura et al., 2010).

Vinculin is a flexible multidomain protein comprising a seven-helical head domain, a proline-rich linker, and a five-helical bundle tail domain (Bakolitsa et al., 2004; Borgon et al., 2004; Molony and Burridge, 1985; Winkler et al., 1996). In its inactive state, it exists in an auto-inhibitory conformation in which the head interacts with the tail domain of the protein, thus masking many of its functional binding sites (Borgon et al., 2004; Chorev et al., 2018). Interaction of vinculin with activated talin, coupled with mechanical perturbation, results in an open vinculin conformation, which enables its binding to various adhesome and cytoskeletal proteins (Bois et al., 2005; Brindle et al., 1996; Carisey and Ballestrem, 2011; Case et al., 2015; DeMali et al., 2002; Hüttelmaier et al., 1997; Jockusch and Isenberg, 1981; Johnson and Craig, 1994; Kim et al., 2016; Thompson et al., 2017; Zamir and Geiger, 2001). While a large body of evidence confirms the pivotal role of vinculin as a mechanosensor and a transducer of actin-generated tension to integrin adhesions throughout their functional states, little is known of how vinculin interacts with and organizes the architecture of the actin cytoskeleton at these sites.

Here we designed an experimental system to assess interactions of dynamic actin networks with full-length vinculin, in solution and upon vinculin interaction with immobilized talin fragment, resembling an adhesion site. We utilized in vitro reconstitutions of purified proteins to determine the molecular mechanisms underlying vinculin-mediated bundle initiation from branched actin networks. Next, we analyzed the effects of soluble activated vinculin on both dense actin organizations and sparse filament networks, to resolve the effect of actin crowding on vinculin crosslinking. Using reconstituted patterned actin organizations, we showed that activated full-length vinculin slows down lamellipodial branched networks growth and binds cellular-like actin organizations with comparable affinities. Through single filament total internal reflection fluorescence (TIRF) microscopy, we provide evidence that vinculin remodels Arp2/3 complex-mediated dendritic actin networks. These interactions are dominated by rapid binding of vinculin and fast crosslinking of actin filaments (F-actin) into stable and flexible bundles. Spatial restriction on vinculin activation by surface-bound talin, indicated its ability to interact with mobile Arp2/3 complex-mediated branched actin networks, modify their organization and to initiate a complex bundle formation. Therefore, our results highlight the ability of vinculin to connect filaments from various actin organizational states and suggest that talin-associated vinculin engages lamellipodial branched networks, from which it can initiate the formation of a stable network of actin bundles at the integrin nascent adhesion sites.

Results

Talin-activated vinculin alters the organization and dynamics of lamellipodium-like Arp2/3 complex-mediated branched actin networks

The lamellipodial actin network is a dynamic and dense mesh made of short and branched filaments (Svitkina and Borisy, 1999) capable of opposing the front-edge membrane tension and inducing protrusions (Pollard and Borisy, 2003). The Arp2/3 complex is a canonical actin nucleator of the leading edge (Mullins et al., 1998; Svitkina and Borisy, 1999), that together with the capping protein αβ (CP, a physiological marker of lamellipodia) (Iwasa and Mullins, 2007; Pollard and Borisy, 2003), organize actin into stiff and propulsive dendritic networks (Achard et al., 2010; Akin and Mullins, 2008; Loisel et al., 1999). The Arp2/3 complex is constitutively inactive. It is recruited to and activated at the plasma membrane by the WAVE signaling complex, where it assembles branched filaments that generate force and push the membrane forwards (Machesky et al., 1999; Svitkina and Borisy, 1999). The lamellipodial actin treadmills centripetally and was proposed to be coupled via vinculin to early stages of nascent adhesions (Case and Waterman, 2015; Choi et al., 2008; Thievessen et al., 2013; Zaidel-Bar et al., 2003). Yet, how vinculin-F-actin interactions modulate branched network dynamics remains unclear.

First, we expressed the full-length version of human vinculin in insect cells, since post-translational modifications might be relevant for its function (Golji et al., 2012). The activation of the full-length vinculin by the constitutively active vinculin-binding site 1 (VBS1, residues 482–636) of talin 1 (Cohen et al., 2005; Papagrigoriou et al., 2004) allowed its binding to F-actin. Next, we investigated the effect of vinculin-F-actin interactions with branched networks at a density and stiffness that resembles the cellular environment. Using two-dimensional patterns printed on a protein-repellent glass coverslip and functionalized with a nucleation promoting factor (NPF), a C-terminal fragment of the Wiskott-Aldrich syndrome protein (WASP-pWA), which activates Arp2/3 complex-mediated nucleation of dendritic actin networks. Due to its fast and stable interactions with the actin barbed end (Schafer et al., 1996), CP prevents the expansion of Arp2/3-mediated branches away from the nucleation patterns. Capped actin branches host de novo nucleation by Arp2/3 complex, thereby ensuring the formation of a rigid branched meshwork, the growth of which results in a backward motion of the entire network (Figure 1A, Figure 1—video 1Boujemaa-Paterski et al., 2017). Thus, the addition of VBS1-activated vinculin to lamellipodium-like structures significantly reduced their growth (Figure 1B, Figure 1—video 1), in line with in vivo studies showing that adhesion-bound vinculin engages and halts lamellipodial retrograde flow (Choi et al., 2008; Thievessen et al., 2013).

Figure 1 with 1 supplement see all
Vinculin alters the dynamics and the organization of lamellipodium-like branched actin networks.

(A) A schematic illustration of dendritic network growth from a WASP pWA-micropattern on a two-dimensional glass coverslip (left) in the presence of profilin–actin (in blue-yellow), CP (in white), and Arp2/3 (in purple). The three-dimensionally constrained growth of lamellipodium-like branched actin network is forced by the geometry (middle scheme). Median, orthogonal ZY and ZX views from confocal imaging of a representative network is shown on the right. The blue arrowhead indicates the position of the orthogonal ZX section. The purple arrowhead indicates the position of the Nucleation promoting factor (NPF) pattern. Scale bars, 4 µm. (B) Wide-field epifluorescence images of the growth of different actin networks on 3 × 15 μm2 GST-WASP pWA-coated patterned bar. Actin polymerized in a 10 µm high chamber in the presence of 4 μM G-actin Alexa-568 labeled, 8 μM Profilin, 120 nM Arp2/3 complex, 16 nM CP and talin VBS1-vinculin as indicated. (C) Analyses of longitudinal and transversal vinculin to actin fluorescence ratio along and across 25 networks, assembled as in B. Fluorescence intensities were measured along the line-scans (doted arrows) shown in the schematics. Plateau means are indicated. (D) Steady state saturation curve was fitted assuming mass-action binding at equilibrium. (E) Fluorescence quantification of the networks showed a stable vinculin to actin fluorescence ratio over the actin density. (C, D, E) Error bars represent the standard deviation from 25 networks per condition. (F) Dependence of networks length and actin fluorescence density on vinculin concentration. Statistical comparisons using Holm-Šídák test and a one-way analysis of variance (ANOVA) showed significant variations from 39, 83, 89 values for 0, 0.25, 0.5 µM vinculin, respectively, with p values < 0.0001. (B, C) Scale bars, 10 µm.

Steady state analyses showed that activated vinculin distributed homogenously throughout the entire network (Figure 1C) with an apparent dissociation constant of 0.2 mM (Figure 1D), independent of the network density (Figure 1E). Surprisingly, increasing the concentration of vinculin resulted in networks with higher actin density (Figure 1F). This may reflect (i) a mesoscale effect that results from a reorganization of the dendritic network by vinculin, or (ii) a microscale, biochemical effect of vinculin either on the dendritic nucleation activity of Arp2/3 complex, as vinculin was shown to bind Arp2/3 complex in vivo (Chorev et al., 2014; Chorev et al., 2018; DeMali et al., 2002), or on actin polymerization dynamics (Jannie et al., 2015; Le Clainche et al., 2010; Wen et al., 2009).

The influence of actin organization on vinculin-mediated bundling

Vinculin was shown to participate in bridging cell adhesions with lamellipodial and lamellar actin, of which architecture, dimensionality and density may vary greatly (Geiger et al., 2009; Thievessen et al., 2013; Xu et al., 2012). We therefore explored how local actin organization affects vinculin recruitment and its ability to bundle the filamentous networks. Patterns of dotted rings were produced for studying three main types of cellular-like actin organization; namely, branched actin networks (Svitkina and Borisy, 1999), filopodial-like uniform-polarity filament bundles (Svitkina and Borisy, 1999; Vignjevic et al., 2006), and mixed-polarity bundles resembling the contractile segments of stress fibers (Hotulainen and Lappalainen, 2006; Xu et al., 2012Figure 2, Figure 2—video 1). We used a similar setting as described above, but omitted CP. This led branched filaments to be nucleated on functionalized patterns and grow unrestricted with their fast-growing ends directed outwards (Figure 2A).

Figure 2 with 6 supplements see all
The affinity of vinculin to actin in variety of network organizations.

(A) An example of patterned dots used to reconstitute actin networks (upper left corner). Schematic view shows the organization of actin filaments on two adjacent patterned dots. An Arp 2/3 complex-mediated branched actin network organization assembled on the WASP pWA-coated micropatterns (in gray) with 0.8 µM actin, 2.4 µM profilin, and 130 nM Arp2/3 complex. The Arp2/3 complex is activated by pWA and actin branches grow with their barbed end oriented outwards. (B) Representative steady state fluorescence images of reconstituted networks showing actin and the corresponding vinculin to actin fluorescence ratio. The image set illustrates the analysis in D. Conditions as in A but supplemented with VBS1-activated vinculin. (C) Dotted and continuous patterned rings indicating the line-scans used for quantifications of actin and vinculin fluorescence intensities across mixed polarity (green), uniform polarity (blue) and branched networks (purple). Conditions as in B. (D) Vinculin to actin fluorescence intensity ratio as a function of actin network fluorescence density. Conditions as in B. Inset, Steady state saturation curve was fitted assuming mass-action binding at equilibrium. In all cases, the concentration of vinculin was much lower than the KD and that of actin was much higher. Apparent dissociation equilibrium constant had a confidence interval of 31 to 38 for uniform polarity, 35 to 39 for mixed polarity, and 31 to 36 for the branched networks. (D) Inset. Error bars represent standard deviation from 529, 348, 181 patterns, as indicated (details in Materials and methods section). Scale bars, 20 µm.

VBS1-activated vinculin formed stable bundles composed of uniform- and mixed-polarity filaments, and bound branched F-actin networks (Figure 2B, Figure 2—video 2). We quantified the fluorescence intensity ratio (vinculin:actin) of the three different networks (Figure 2—figure supplement 1) assembled on >500 dotted patterned as well as on continuous patterned rings (Figure 2C,D). Here, high and low density actin networks, represented by high and low fluorescence intensity networks respectively, were exposed to identical vinculin concentrations and were analyzed. Control experiments showed that addition of VBS1 or vinculin alone has no effect on the actin organization (Figure 2—figure supplement 2). These experiments demonstrated that while vinculin binding to actin is independent of the actin organization (i.e. vinculin affinities are similar in branched, uniform, or mixed-polarity networks) it is sensitive to actin density. Namely, sparser actin networks showed a higher occupancy than the denser ones (Figure 2D) while dense lamellipodium-like networks that contain CP exhibit ~eightfold higher vinculin-actin dissociation constant (Figure 1D).

Since actin architectures polymerized by adhesion-associated actin nucleators can be attached to their nucleation sites (Balaban et al., 2001; Choi et al., 2008; Chorev et al., 2014; DeMali et al., 2002; Lavelin et al., 2013; Legerstee et al., 2019; Oakes et al., 2012), we tested how surface anchorage and filament density in the intermeshed, branched networks (on the patterns) affect the bundling. We utilized continuously patterned rings and polymerize an actin network of reduced branched density. In the absence of vinculin, isolated branched actin networks polymerized and intermeshed, resulting in smooth coverage of the patterns (Figure 2—figure supplement 3A,B,E; Figure 2—video 1). In the presence of VBS1-activated vinculin, dense networks polymerized on the rings were decorated by vinculin, and prominent bundles formed between the densely actin subnetworks (Figure 2—figure supplement 3C,D,E; Figure 2—video 3). These findings showed that vinculin not only binds branched and intermeshed actin filaments but reorganizes them into stable bundles.

Rapid reorganization of actin branched networks by vinculin crosslinking

To characterize the effects of vinculin on branched actin networks at the single filament level, we utilized dual-color total internal reflection fluorescence (TIRF) microscopy. Here, we monitored actin polymerization and vinculin-F-actin interactions simultaneously. Vinculin and Arp2/3 complex were mixed with actin monomers in the absence or in the presence of their activators, VBS1 and the C-terminal amino acids of the WASP-family verprolin-homologous protein (WAVE WA) (Machesky et al., 1999), respectively.

Dendritic actin networks were assembled by Arp2/3 complex and rapidly decorated by vinculin, with no detectable lag time between actin polymerization and decoration (Figure 3A). Actin branches were initiated on both vinculin-decorated actin filaments and bundles induced by vinculin (Figure 3A, Figure 3—videos 1 and 2). The branched actin filaments were cross-linked either to the mother filament or to neighboring branches, that is to form uniform- or mixed-polarity bundles, respectively. As polymerization proceeded, the Arp2/3 complex-mediated branched networks were reorganized by vinculin and interconnected via stable and dynamic vinculin-F-actin bundles (Figure 3—videos 1 and 2).

Figure 3 with 9 supplements see all
Vinculin induced uniform and mixed polarity filaments bundles.

(A) Actin polymerization and bundling by 0.3 µM full-length vinculin at, in the presence of 0.9 µM actin, 12 nM Arp2/3 complex, 80 nM WAVE WA fragment, and 1.4 µM talin VBS1. Crosslinking within and between independent networks are followed over time, NW1 and NW2. Dynamic actin branches (purple arrowheads), uniform polarity (UP, green arrowheads) and mixed polarity (MP) bundles are indicated. Schemes defining UP and MP with a pair of actin filaments crosslinked by vinculin (in green) are drawn below; barbed ends (B) are indicated. (B) Quantifications of branch density or vinculin to actin fluorescence ratio of networks polymerized with the mentioned proteins at concentrations indicated in A, are shown as histograms. Error bars represent the standard deviation from 15 to 20 values per condition. (C, D) Controls assays with the mentioned proteins at concentrations indicated in A. Scale bars, 5 µm.

Vinculin-decorated actin branches freely diffused or were recruited to existing bundles, with no obvious detectable debranching of the vinculin-decorated network. Quantitative measurements showed that the branch density of dendritic networks was not significantly affected by vinculin decoration, and vinculin binding was not significantly influenced by the presence of Arp2/3 complex (Figure 3B). Furthermore, control assays confirmed that the bundling of actin branches was solely due to the presence of activated vinculin (Figure 3C) and in the absence of Arp2/3 activator, no actin branching was detected (Figure 3D). This suggests that vinculin may directly interact only with a specific cellular sub-population of Arp2/3 complex or additional cellular factors are presumably required (Chorev et al., 2014).

To characterize the bundling activity of full-length vinculin protein in higher resolution and details, we applied hybrid methods combining TIRF microscopy and cryo-ET with an experimental set-up that allowed actin filaments to freely diffuse and adopt random orientations prior to bundling by vinculin. While actin polymerization was not affected by the presence of either VBS1 or vinculin alone and chemical labelling of vinculin did not affect its native auto-inhibited configuration (Figure 3—figure supplement 1), assembly kinetics confirmed that actin polymerization was not altered by vinculin binding (Figure 3—figure supplement 2D). Here, we calculated an on-rate constant of the actin filament’s fast-growing barbed end of 10 ± 1 µM−1 sec−1, similar to values obtained for free barbed end actin growth in solution. Our findings were in agreement with Leclainche and collaborators Le Clainche et al., 2010 who used bacterially expressed full-length vinculin. However, we measured no activation of actin polymerization by vinculin, as was shown by bacterially expressed chicken vinculin tail fragment (Jannie et al., 2015; Wen et al., 2009). Since we detected no apparent lag time between filament polymerization and vinculin binding (Figure 3—figure supplement 2A,B), an on-rate constant of vinculin can be approximated to 10 ± 1 µM−1 sec−1, as both proteins were used at similar concentrations. Interestingly, we found that the ‘zippering’ process was strikingly fast, reaching a velocity of 38 ± 17 µm/min, equivalent to 230 ± 110 actin subunits/sec, with zippering angles that exceed 60 degrees (Figure 3—figure supplement 2C). These crosslinks were mechanically robust enough to occasionally break existing actin filaments (Figure 3—video 3), and the bundles were flexible and could buckle with curvatures exceeding 0.2 ± 0.06 µm−1 for up to 9-filament bundles (Bathe et al., 2008Figure 3—figure supplement 2E).

Using cryo-electron tomography (cryo-ET) combined with image processing approaches we resolved the organization and polarity of vinculin-F-actin bundles. Bundles induced by the vinculin tail domain showed an inter-filament spacing, 10 ± 1 nm (Figure 3—figure supplement 3A), as previously reported (Janssen et al., 2006). However, bundles assembled with activated full-length vinculin showed a wider inter-filament spacing, 29 ± 5 nm, presumably due to the lack of vinculin heads anchorage through cellular integrin-bound talin (Case et al., 2015Figure 3—figure supplement 3). More importantly, we reconstructed actin filaments to a resolution of 14 Angstrom and unambiguously determined filaments’ polarity within these in vitro bundles. Next, we determined the polarity of the original filaments (Figure 3—figure supplement 4A–D and Materials and methods section, Martins et al., 2020). Evaluating the polarity of each actin filament with respect to its neighboring filaments in 3D revealed a mixed-polarity organization, wherein neighboring filaments could exhibit both uniform- and mixed-polarity orientation (Figure 3—figure supplement 4E).

Thus, our light microscopy and cryo-ET results suggested that, in a force-independent system, VBS1-activated vinculin mediates stable bundling through a rapid binding and fast crosslinking process. Moreover, this bundling does not favor uniform over a mixed actin polarity for bundled filaments.

Substrate-bound VBS1-activated vinculin initiates bundles out of branched networks

Cytosolic vinculin is recruited to early integrin adhesions (e.g. nascent adhesions and focal complexes) (Choi et al., 2008; Zaidel-Bar et al., 2003), and is activated at these adhesion sites by talin, where it is exposed to the shear flow generated by lamellipodial actin retrograde flow (Case et al., 2015; Thievessen et al., 2013). It is currently unclear how adhesion localized vinculin is involved in local actin bundle formation and how the centripetally flowing F-actin sheet is transformed into a FA-associated bundle.

To address these questions, we spotted talin VBS1 onto micropatterned surfaces before flowing a solution containing soluble, inactive vinculin, WA-activated Arp2/3 complex, and G-actin over the surfaces. We then followed actin dynamics using TIRF microscopy (Figure 4A). We observed the rapid recruitment of vinculin to the VBS1-anchored spots (Figure 4B, first two panels, and Figure 4—video 1). In parallel control experiments, micropatterns coated with bovine serum albumin protein (BSA) did not produce any significant vinculin recruitment (Figure 4—figure supplement 1, Figure 4—video 2). These results suggest that vinculin was specifically recruited and activated by the immobilized VBS1. Furthermore, control experiments combining microfluidics and TIRF microscopy confirmed that VBS1-activated vinculin immobilized on actin filaments is active and can engage flowing and naked actin filaments (Figure 4—figure supplements 2 and 3, Figure 4—video 3).

Figure 4 with 6 supplements see all
Vinculin activated by surface immobilized VBS1 initiate actin bundles only at coated spots.

(A) Scheme of the experimental setup shows that the passivated coverslip is patterned (1) and coated with talin-VBS1 (2). Next the patterned surface was mounted into a flow chamber in which a mix of 300 nM inactive full-length vinculin, 0.3 µM actin monomer, 18 nM Arp2/3 and its activator, 80 nM WAVE WA, was delivered (3). (B) Bundling of Arp2/3-branched actin networks by vinculin was observed only at the sites of patterned VBS1. Dotted arrows indicate the direction of movement of the branched networks, which are bundled. Dotted circle shows the position of the VBS1-coated pattern decorated by vinculin (green in the upper panel). Mixed polarity (MP) and uniform polarity (UP) bundles formed on the pattern. Scale bar, 2 µm. (C) Kymographs along the three bundles, shown in B, confirm the crosslinking of Arp2/3 complex-mediated branches into MP (highlighted by arrows) and UP (arrowheads highlight the fusion of 4 branches to a secondary network). (D) The schematic view depicts the bundles initiated from the two branched networks, colored in white and in red and imaged in B.

As actin polymerization proceeded, branched actin networks were formed in solution, flowing over the immobilized VBS1-vinculin complexes and eventually being engaged by them. Strikingly, at these functionalized spots, the branched networks were profoundly rearranged into growing uniform- and mixed-polarity bundles (Figure 4B–D, Figure 4—video 1). Moreover, the actin bundling process was strictly local, occurring only at the immobilized VBS1-vinculin spots, but not observed elsewhere. In the control experiments, BSA-coated patterns showed no apparent interactions and no network remodeling of actin was detected (Figure 4—figure supplement 1, Figure 4—video 2). The fact that vinculin, once selectively recruited and activated by spotted talin VBS1, is able to bind and bundle branched actin networks is highly reminiscent of actin bundling along focal complexes (Bachir et al., 2014; Case et al., 2015; Choi et al., 2008; Thievessen et al., 2013) and maturing FAs (Alexandrova et al., 2008; Gardel et al., 2008; Geiger et al., 2009; Tee et al., 2015).

Discussion

Vinculin is a canonical component of the integrin-mediated adhesion machinery (Geiger, 1979). It interacts with multiple adhesome proteins (Horton et al., 2016; Zaidel-Bar et al., 2007a; Zamir and Geiger, 2001) and therefore participates in interactions between the ECM and the actin cytoskeleton. Previously, it was shown that vinculin exists in an auto-inhibited state (Bakolitsa et al., 2004; Borgon et al., 2004; Izard et al., 2004). Vinculin is activated after binding to talin at nascent adhesion sites and subsequent exposure to mechanical force, driving focal adhesion assembly (Galbraith et al., 2002; Thievessen et al., 2013). Monitoring the early stages of adhesion formation using live cell microscopy demonstrated that the assembly of primordial adhesions (‘nascent adhesions’ and ‘focal complexes’) at the rear of the lamellipodium is followed by centripetal extension of actin filaments from these sites (Bachir et al., 2014; Choi et al., 2008; Humphries et al., 2007; Tee et al., 2015; Thievessen et al., 2013). Understanding the properties of proteins that participate in the initiation and maturation of these nascent adhesion-cytoskeleton interactions is of a major importance.

Inspired by the dynamic interplay between the branched actin network, generated at leading edge, that flows centripetally, and the subjacent, nascent integrin adhesions (Case and Waterman, 2015; Geiger et al., 2009), we searched for an in vitro ‘molecular model’ that may simulate early events of adhesion assembly (Choi et al., 2008; Thievessen et al., 2013). In pursuit of this goal, we focused on the interplay between relevant cytoskeletal network components and full-length vinculin. A combination of real-time fluorescence microscopy, microfluidics, advanced substrate micro-patterning and cryo-ET provided insights into the dynamic molecular processes whereby active, nascent adhesion-associated vinculin transforms Arp2/3-induced branched actin networks into mature adhesion-associated bundles.

Relatively little is known about how vinculin controls the organization of lamellipodial branched actin networks, despite its crucial role in bridging actin dynamics and adhesions turnover. In vivo high-resolution traction-force and fluorescence microscopy showed that the vinculin tail domain mediates interactions of F-actin retrograde flow to the mature focal adhesion (Alexandrova et al., 2008; Thievessen et al., 2013), and plays a critical role controlling nascent adhesion maturation (Choi et al., 2008; Thievessen et al., 2013) and organization of the lamellipodial-lamellar actin border (Alexandrova et al., 2008; Thievessen et al., 2013). However, the underlying molecular mechanisms remain unclear. Here we showed that the vinculin-binding domain of Talin1 (VBS1) activates full-length vinculin, enabling it to interact and control the dynamics and the organization of reconstituted lamellipodium-like Arp2/3 complex-mediated actin networks (Figure 1). Our reconstitution elucidates a fundamental standing question of how vinculin, activated in a force-independent regime, induces a mesoscale reorganization of lamellipodial branched actin networks.

Our results show that, while binding talin VBS1, vinculin readily decorated and reorganized Arp2/3 complex-mediated branched networks of variable density and stiffness (Figures 2 and 3). The interaction between vinculin and F-actin was fast and independent of actin polymerization resulting into the formation of mixed-polarity bundles. Yet, it is worth noting that in the in vitro experiments (Figure 3—figure supplement 3) the full-length vinculin protein may not fit within the tight actin bundle that is formed by vinculin tail domain alone (Janssen et al., 2006). In cells, vinculin binds to integrin-anchored talin, therefore 3D actin bundles crosslinked only by vinculin are unlikely.

Previous structural investigations (Bakolitsa et al., 2004; Borgon et al., 2004; Dedden et al., 2019; Goult et al., 2013; Izard et al., 2004), in vitro stretching assays (del Rio et al., 2009; Yao et al., 2015), live cell microscopy studies (Margadant et al., 2011; Thievessen et al., 2013), and Förster resonance energy-based tension sensors for both proteins (Case et al., 2015; Grashoff et al., 2010; Kumar et al., 2016) have revealed the mechanosensitive nature of talin and vinculin. These studies support the current model requiring actin-generated force to unveil key regulatory domains that potentiate their role in adhesion initiation and maturation. However, recent studies propose an alternative pathway to the force-dependent relief of auto-inhibition in talin and vinculin. Vinculin binding site R3 (Izard et al., 2004) was found to be available for vinculin binding in soluble closed talin (Dedden et al., 2019), while Paxillin was shown to promote efficient targeting of vinculin to the adhesion site prior to its activation by talin (Case et al., 2015). Recent in vivo reconstitutions confirmed the key role of R3 in the binding of vinculin to talin in a force-independent regime, as well as paxillin association to closed vinculin and talin (Atherton et al., 2020). These interactions may form precomplexes at an early-stage nascent adhesion (Choi et al., 2008). Furthermore, in vivo cross-correlated fluctuations analysis producing high-resolution spatial and temporal maps (Digman et al., 2009) showed that talin-vinculin association precedes their recruitment to nascent adhesions (Bachir et al., 2014). Finally, Kelley and colleagues showed that phosphoinositides enable talin-vinculin-actin association in a force independent-regime (Kelley et al., 2020). The work presented here supports these findings and shows that vinculin, activated in a force-independent regime, can reorganize Arp2/3 complex dendritic networks.

Vinculin bundling properties

Quantitative and structural analyses of talin VBS1-activated vinculin-induced reorganization of patterned actin structures (Figure 2) and single filaments (Figure 3) revealed fundamental features of vinculin-actin interactions: (i) activated vinculin binds to uniform, mixed-polarity and intermeshed branched actin organizations with a similar affinity; (ii) three-dimensional bundles assembled were characterized by a random polarity neighborhood; (iii) activated vinculin displayed an association rate constant resembling that of actin polymerization and filaments zippering rate of ~0.6 µm/sec. This rate is considerably faster than the lamellipodial actin polymerization rate, which falls in the range of 1–11 µm/min (Geraldo et al., 2008; Medeiros et al., 2006; Watanabe and Mitchison, 2002). It infers that the zippering is independent of actin polymerization, suggesting that vinculin-mediated crosslinking is not a rate-limiting step at the leading edge of migrating cells, where nascent adhesions emerge under the highly dynamic lamellipodial actin network (Choi et al., 2008; Ponti et al., 2004; Zaidel-Bar et al., 2003). Altogether, our data describes talin-activated vinculin as an efficient candidate which (in the absence of any tensile force) can efficiently bind randomly-oriented actin branches of mobile lamellipodial meshwork (Svitkina and Borisy, 1999), forming mixed-polarity bundles, modifying the meshwork organization and halting its dynamics (Alexandrova et al., 2008; Choi et al., 2008; Hu et al., 2007; Shemesh et al., 2009). We speculate that in the absence of tensile forces, talin-vinculin precomplexes (Atherton et al., 2020; Bachir et al., 2014; Kelley et al., 2020) efficiently reorganize the mobile lamellipodial actin branches into random polarity bundles, which present at a random orientation (Svitkina and Borisy, 1999). This may trigger the formation of a self-sustained system whereby actin retrograde flow exerts force through preformed bundles on talin-vinculin complexes, completing their activation and association to integrins (Bachir et al., 2014). This would represent a selective process whereby maturation of surviving nascent adhesions (Choi et al., 2008) is regulated by an elegant feedback mechanosensitive mechanism that may reinforce uniform polarity bundles (Case and Waterman, 2015; Huang et al., 2017).

Bundled, dendritic networks at talin-bound spots, in absence of tension

Nascent adhesions emerge at the basal aspect of the lamellipodium where they are linked to the extracellular matrix, attenuating the retrograde lamellipodial actin flow without the presence of a local actin bundle (Gardel et al., 2008; Geiger et al., 2009; Zaidel-Bar et al., 2007b; Zhang et al., 2008).

Our experiment showed that immobilized talin VBS1 recruits and activates vinculin which initiates linear bundle formation by reorganizing flowing Arp2/3-actin networks (Figure 4—video 1). This implies that surface-bound talin-vinculin complex can initiate actin bundles from flowing actin networks and connects them at initial stages of adhesion formation (Choi et al., 2008; Thievessen et al., 2013). At adhesion sites, such crosslinked actin bundles may be formed by few vinculin proteins occupying several vinculin bindings sites on a talin protein or by vinculin dimers. These hypotheses are summarized in a dynamic ‘core mechanism’ model for vinculin-actin filament interactions throughout adhesion initiation and maturation (Figure 5).

A model depicting the initiation of actin bundles from branched networks by membrane bound talin-activated vinculin at nascent adhesions, independent of myosin-mediated force.

Schematic representation of a nascent integrin-based adhesion site, localized in proximity to the leading edge of a cell (left). (1) The Arp2/3 branched actin network treadmills and flows centripetally. Vinculin is activated by interacting with talin (Atherton et al., 2020; Kelley et al., 2020). (2) At nascent adhesion sites, talin-activated vinculin stably binds and bundles mobile branched networks likely affecting actin dynamics. (3) Actin retrograde flow generates tension that further activates talin and vinculin, reinforcing the link to integrins. Adhesion-anchored vinculin interacting with the flowing, branched actin networks initiates bundles at the nascent adhesion site. Vinculin engages the actin retrograde flow, applying tension to the mechanoresponsive components of the adhesion, enabling adhesion self-sustained assembly dynamics and recruitment of additional FA components such as myosin II, α-actinin, and zyxin (Case et al., 2015; Choi et al., 2008; Thievessen et al., 2013).

Beside vinculin, other actin crosslinkers, for example α-actinin 1, could concomitantly be recruited and function during the formation of nascent adhesion (Bachir et al., 2014; Otey et al., 1990). Since α-actinin harbors a cryptic VBS (Le et al., 2017) that is likely to be exposed due lamellipodial-generated tension, it may contribute to activating vinculin during adhesion formation. However, α-actinin one may also contribute to crosslinking actin filament bundles anchored by integrin-bound talin-vinculin complexes (Case and Waterman, 2015; Ciobanasu et al., 2014).

The work described here presents mechanistic insights into the activation-dependent interactions of full-length vinculin with actin networks, crucial for initiation and maturation of nascent adhesions. It provides a molecular view on the mechanical interplay between talin-activated vinculin and the retrograde flow of branched actin networks, independent of myosin-mediated tension. These findings provide mechanistic insights on the force dependent molecular regulation of cell-matrix interactions and cell migration.

Materials and methods

Key resources table
Reagent type
(species) or resource
DesignationSource or referenceIdentifiersAdditional information
Gene
(Homo sapiens)
VCLHuman Genome Nomenclature DatabaseHGNC:12665
Cell line used for cloning
(human skin)
Human A431Sigma-AldrichCat# 85090402Cell line for cloning VCL
Strain, strain background
(S. frugiperda)
SF9 cellsThermoFisher ScientificCat# 12659017Cell line for baculovirus generation and protein expression
Strain, strain background
(E. coli)
DH5αThermoFisher ScientificCat# 18265017Chemically competent cells
Strain, strain background
(E. coli)
DB3.1Gift from Raymond DutzlerChemically competent cells
Strain, strain background
(E. coli)
BL21(DE3)pLysSSigma-AldrichCat# 69451Chemically competent cells
Recombinant DNA reagentpFBXC3GH-VCLThis paper,
Subcloned in vector gifted by R. Dutzler
For recombinant baculovirus generation and full-length human VCL expression in SF9 cells
Recombinant DNA reagentpBXNHG3-TLN1-VBS1This paper,
Subcloned from gift from Christophe LeClainche, in vector gifted by R. Dutzler
Human Talin1-VBS1 (482–636)
Recombinant DNA reagentpBXNHG3-VCL-TailThis paper,
Subcloned from pFBXC3GH-VCL
Human vinculin tail (879–1066)
Commercial assay, kitBac-to-Bac Vector KitThermoFisher ScientificCat# 10359016For cloning
Commercial assay, kitRNeasy mini KitQiagenCat# 74104For cloning
Commercial assay, kitAffinityScript qPCR cDNA Synthesis KitAgilentCat# 600559For cloning
Chemical compound, drugAlexa Fluor 488 C5 MaleimideThermoFisher ScientificCat# A10254For protein labeling
Chemical compound, drugAlexa Fluor 647 C2 MaleimideThermoFisher ScientificCat# A20347For protein labeling
Chemical compound, drugAlexa Fluor 568 C5 MaleimideThermoFisher ScientificCat# A20341For protein labeling
Chemical compound, drugmPEG-Silane, MW 5 kCreative PEGWorksCat# PLS-2011Surface passivation
Chemical compound, drugPLL(20)-g(3.5)-PEG(2)SuSoS AGSurface passivation
Chemical compound, drugMethyl CelluloseSigma-AldrichCat# M0387Reconstitution assays
Peptide, recombinant proteinGlucose Oxidase Type IISigma-AldrichCat# G6125Reconstitution assays
Peptide, recombinant proteinCatalaseSigma-AldrichCat# C9322Reconstitution assays
OtherAdhesive TapeNittoCat# 5601Microscopy chamber assembly
Othersticky-Slide VI 0.4IbidiCat# 80608Reconstitution assays
Software, algorithmFIJIhttps://imagej.net/Fiji/DownloadsFor data analysis
Software, algorithmPrismGraphPadhttps://www.graphpad.com/scientific-software/prism/For data analysis
Software, algorithmMATLABMathWorkshttps://www.mathworks.com/products/matlab.htmlFor data analysis
Software, algorithmSerialEMUniversity of Colorado
Open source under an MIT license
https://bio3d.colorado.edu/SerialEM/download.htmlFor data acquisition
Software, algorithmUCSF ChimeraUniversity of Californiahttps://www.cgl.ucsf.edu/chimera/download.htmlFor structure visualization

Human full-length vinculin cloning

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Full-length vinculin cDNA was isolated from a human A431 cell line by extracting mRNAs using the RNeasy Kit (QIAGEN), followed by a reverse transcriptase reaction with a specific primer (5’-GGTGCCTACTGGTACCAGGGAG-3’) using the AffinityScript kit (Agilent). The PCR product (using the forward 5’-ATATATGCTCTTCTAGTATGCCAGTGTTTC-3’, and the reverse 5’-TATATAGCTCTTCATGCCTGGTACCAGGG-3’ primers) of 1066 amino acids long, was cloned into a pFBXC3GH insect cell expression vector containing a C-terminal 3C protease cleavage site followed by a GFP and His tag, using the fragment exchange (FX) cloning strategy (Geertsma and Dutzler, 2011). Protein expression was performed by generating a recombinant baculovirus for Sf9 insect cell infection at a density of 2.0 × 106 mL−1 using the Bac-to-Bac system (Invitrogen). Briefly, the recombinant vinculin-GFP encoding plasmid pFBMS was transformed into DH10Bac E. coli cells, which enabled the transposition of the recombinant gene into the bacmid genome. The recombinant bacmid DNA was then isolated and transfected into Sf9 cells to generate the full-length vinculin-expressing baculovirus.

Human full-length vinculin tail (879–1066) cloning

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Human vinculin tail (879–1066) was cloned from pFBXC3GH-human FL vinculin into a pBXNHG3 bacterial expression vector, containing an N-terminal His tag followed a GFP and 3C protease cleavage site, using the fragment exchange (FX) cloning strategy (Geertsma and Dutzler, 2011).

Protein expression and purification

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Sf9 cells were infected with the full-length vinculin-expressing baculovirus for recombinant protein expression. Expressing cells were harvested by gentle centrifugation and lysed by sonication in 20 mM Tris-HCl pH 7.5, 0.4 M NaCl, 5% Glycerol, 1 mM DTT, 0.1% Triton, and protease inhibitors. Full-length vinculin or vinculin tail fragment 879–1066 were purified from clarified cell extract on Profinity Ni-charged Immobilized metal affinity chromatography, IMAC resin (Bio-Rad Laboratories, Inc). The C-terminal GFP followed by a 10xHis tag was cleaved by incubating the resin-bound protein with 3C protease, and full-length vinculin was eluted with 20 mM Tris Ph 7.5, 150 mM NaCl, 5% glycerol, 3 mM beta mercaptoethanol, 5 mM Imidazole. Proteins were further purified on a Superdex 200 size exclusion column (GE Healthcare) and eluted with 20 mM Tris-HCl pH 7.8, 0.15 M KCl, 1 mM MgCl2 and 5% Glycerol. Protein aliquots supplemented with 15–20% glycerol were flash-frozen in liquid nitrogen and stored at −80°C.

Actin was purified from rabbit skeletal muscle acetone powder, purified according to the method of Spudich and Watt, 1971, and stored in G-buffer (5 mM Tris-HCl pH 7.8, 0.2 mM CaCl2, 0.5 mM DTT, 0.2 mM ATP). The Arp2/3 complex was purified for bovine thymus by WAVE WA affinity chromatography as described (Egile et al., 1999). Human profilin, mouse capping protein (α1/β2), human WAVE WA and human WASP pWA constructs were obtained from L. Blanchoin. Human profilin and His-tagged mouse capping protein constructs were expressed in BL21 (DE3)pLysS strain and purified using ion-exchange and size exclusion chromatography as described (Lu and Pollard, 2001; Palmgren et al., 2001). GST-tagged human WAVE WA or human WASP pWA fragment constructs, and GST-tagged S. cerevisiae. Human Talin 1 VBS1 residues 482–636 and human α-actinin one constructs were obtained from C. Le Clainche, expressed in BL21 (DE3)pLysS strain and purified by glutathione-Sepharose affinity chromatography as described (Ciobanasu et al., 2015). Purified polypeptides were supplemented with 10–20% glycerol, aliquoted, flash-frozen in liquid nitrogen and stored at −80°C. Ca-monomeric actin was stored on ice and used within 2 weeks for TIRFM and micropatterning assays.

Fluorescence labeling of protein

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Proteins were labeled through cysteines with Alexa Fluor maleimide dyes (Invitrogen) or Cy3B maleimide dye (GE Healthcare). Actin was labeled with Alexa647, for TIRFM and with Alexa568 for epifluorescence microscopy, vinculin was conjugated to Alexa488, α-actinin one to Alexa568. Labeling reactions of vinculin or α-actinin one were performed in 50 mM K-Phosphate buffer pH 7, and actin labeling in 2 mM Tris-HCl pH 7, 0.2 mM CaCl2, 0.2 mM ATP, for 16 hr. To limit excessive labeling, protein/dye molar ratio was kept ≤1:3. The reactions were stopped by adding 10 mM DTT. The labeled vinculin was purified using Superdex 200 size exclusion column (GE Healthcare), in 20 mM Tris-HCl pH 7.8, 0.15 M KCl, 1 mM MgCl2 and 5% Glycerol. Labeled actin was polymerized, depolymerized, and gel-filtered using Superdex 200 size exclusion column (GE Healthcare), in 2 mM Tris-HCl pH 7, 0.2 mM CaCl2, 0.2 mM ATP, 0.5 mM DTT. The degree of labeling (DOL) was determined by absorption spectroscopy, using the known extinction coefficients for the dyes and proteins, or using coefficients predicted from the amino acid sequences using PredictProtein software (ExPASy, Swiss Institute Bioinformatics Resource portal). Actin, vinculin, and α-actinin 1 DOLs ranged between 25–35%. Purified polypeptides were supplemented with 10–20% glycerol, aliquoted, flash-frozen in liquid nitrogen and stored at −80°C. Ca-monomeric actin was stored on ice and used within 2 weeks for TIRFM and micropatterning assays.

Glass surface passivation and deep-ultraviolet (UV) micropatterning

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Using passivated flow cells was a prerequisite for preventing non-specific protein/substrate interactions and improve signal-to-noise ratio during TIRFM image acquisition. Thus, slides and coverslips (CVs) were used to assemble the reaction chambers were drastically cleaned by successive chemical treatments (adapted from Boujemaa-Paterski et al., 2014): 2 hr in 2% Hellmanex III, rising in ultrapure water, 30 min in acetone, 30 min ethanol 96%, rinsing in ultrapure water. Slides and CVs were dried using a filtered nitrogen gas flow and oxidized with oxygen plasma (3 min, 30 Watt, Femto low-pressure plasma system Type A, Diener electronic GmbH, Germany), just before an overnight incubation in a solution containing tri-ethoxy-silane-PEG (5 kDa, PLS-2011, Creative PEGWorks, USA) 1 mg/mL in ethanol 96% and 0.02% of HCl, with gentle stirring for TIRFM assays. mPEG-silane coated slides and CVs were then rinsed in ethanol and extensively in ultrapure water. Passivated slides and CVs were then stored in a clean container and used within a week time. Alternatively, for deep UV-assisted micropatterning, plasma cleaned slides and CVs were incubated in PLL(20)-g(3.5)-PEG(2) (SuSoS AG, Switzerland) at 1 mg/mL in Hepes 10 mM pH 7.4. PLL-PEG-coated CVs were immediately patterned, as described below (adapted from Boujemaa-Paterski et al., 2014), and mounted onto a PEGylated slide using a double-sided tape (3M electronics or Nitto).

Likewise, passivated flow cells were mounted by gluing a PEG-silane CV onto a PEG-silane slide and used for TIRFM imaging.

TIRFM imaging

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Reconstitution assays were performed using fresh actin polymerization buffer, containing 20 mM Hepes pH 7.0, 40 mM KCl, 1 mM MgCl2, 1 mM EGTA, 100 mM β-mercaptoethanol, 1.2 mM ATP, 20 mM glucose, 40 μg/mL catalase, 100 μg/mL glucose oxidase, and 0.4% methylcellulose. The final actin concentrations were 0.7 to 1.1 µM, with 13–16% Alexa monomer labels. The polymerization medium was supplemented with vinculin, talin VBS1, Arp2/3 complex, WAVE WA, and α-actinin one as indicated in the figure legends and methods (below). The reaction medium was rapidly injected into a passivated flow cell at the onset of actin assembly, imaging started after 2 min. Time-lapse TIRFM was recorded every 5 or 15 s. TIRF images were acquired using a Widefield/TIRF – Leica SR GSD 3D microscope, consisting of an inverted widefield microscope (Leica DMI6000B/AM TIRF MC) equipped with a 160x objective (HCX PL APO for GSD/TIRF, NA 1.43), a Leica SuMo Stage, a PIFOC piezo nanofocusing system (Physik Instrumente, Germany) to minimize the drift for an accurate imaging, and combined with an Andor iXon Ultra 897 EMCCD camera (Andor, Oxford Instruments). Fluorescent proteins were excited using three solid-state diode lasers, 488 nm (300 mW), 532 nm (500 mW), 642 nm (500 mW). Laser power was set to 5% for Alexa-labeled proteins, and dyes were excited for 50 ms. Image acquisition was performed with 25 degrees-equilibrated samples and microscope stage. The microscope and devices were driven by Leica LAS X software (Leica Microsystems, GmbH, Germany).

Deep UV-assisted micropatterning and widefield fluorescence microscopy

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To direct actin filament polymerization to predefined positions on glass CVs, we used the micropatterning strategy (Boujemaa-Paterski et al., 2014; Reymann et al., 2010) by printing adhesive patterns on a protein-repellent surface. PLL-PEG-coated CVs were exposed to short-wavelength UV radiation (184.9 nm and 253.7 nm, Jelight, USA) for 2 min through 24 × 24 transparent micropatterns printed on a photomask (Compugraphics, Germany). To ensure high-resolution printing of micropatterns on the PLL-PEG-coated CVs, during UV exposure CVs and the photomask were mounted onto a custom-made vacuum-compatible holder. Immediately after UV exposure, micropatterned CVs were incubated in 0.2–0.3 µM GST-WASp pWA solution for 15 min at room temperature under gentle agitation, then washed in a buffer containing 2 mM Tris-HCl pH 7, 0.2 mM CaCl2, 0.2 mM ATP, 0.5 mM DTT, mounted onto a PLL-PEG coated slide, and injected with a defined reaction medium.

To restrict actin filament polymerization on pWA-functionalization micropatterns, reconstitution media contained both the Arp2/3 complex, recruited and activated by pWA attached to the patterned surfaces, and profilin that inhibits spontaneous nucleation of actin filaments in solution. However, Arp2/3 complex-mediated actin nucleation on the micropatterns is allowed. Thus, to polymerize patterned actin networks, we injected micropatterned flow cells with an actin polymerization medium, containing 20 mM Hepes pH 7.4, 30 mM KCl, 1 mM MgCl2, 1.7 mM EGTA, 40 mM DTT, 1.2 mM ATP, 20 mM glucose, 40 μg/mL catalase, 100 μg/mL glucose oxidase, and 0.2% methylcellulose. In all tests, final actin concentration was 0.8 µM, with 18% Alexa568-labeled monomers, profilin was 2.4 µM, the Arp2/3 complex was 130 µM, vinculin and/or talin VBS1 as indicated in the figure legends. Imaging started after 15 min. Time-lapse microscopy was recorded every 1 or 2 min. To polymerize stiff, dense and motile branched networks, the polymerization medium was supplemented with 16 nM Capping protein, 4 µM actin, with 5% Alexa568-labeled monomers, 8 µM profilin, 120 nM Arp2/3 complex, and vinculin, talin-VBS1 and/or α-actinin 1, as indicated in the figure legends. Fluorescence images were acquired on an inverted widefield Leica DMI4000B microscope equipped with a 63x oil objective (HCX PL APO 63x; NA 1.40–0.60) and combined to Leica DFC 365 FX camera (Leica Microsystems, GmbH, Germany). Illumination was set to 17% of halogen lamp power and fluorescent dyes were excited for 80 ms. Image acquisition was performed with 25 degrees Celsius-equilibrated samples and microscope stage. The microscope and devices were driven by Leica LAS X software (Leica Microsystems, GmbH, Germany). Samples were also explored using an Olympus IXplore SpinSR10 spinning disk confocal imaging system (Olympus Scientific Solutions, USA), equipped with a 60x Silicon oil objective (UPLSAPO UPlan S Apo 60x, NA 1.3) and with two Prime BSI scientific CMOS cameras (Teledyne Photometrics, USA).

Deep UV-assisted micropatterning and TIRF microscopy

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Micropatterns were also coated with GST-talin VBS1 to address vinculin activation to the patterned surfaces. Briefly, immediately after UV exposure, micropatterned CVs were incubated in 0.4 µM GST-talin VBS1 solution for 15 min at room temperature under gentle agitation, then washed in a buffer containing 2 mM Tris-HCl pH 7, 0.2 mM CaCl2, 0.2 mM ATP, 0.5 mM DTT, passivated with a solution of 4.5 µM BSA, washed again, mounted onto a PLL-PEG coated slide, and injected with a defined reaction medium. Control patterns were similarly treated, except for the coating step where GST-talin VBS1 was replaced by a BSA.

To monitor the recruitment of vinculin to the GST-talin VBS1-coated patterns, and subsequent remodeling of Arp2/3-branched actin network to these adhesion-like spots, we injected micropatterned flow cells with an actin polymerization medium, containing 20 mM Hepes pH 7.0, 40 mM KCl, 1 mM MgCl2, 1 mM EGTA, 100 mM β-mercaptoethanol, 1.2 mM ATP, 20 mM glucose, 40 μg/mL catalase, 100 μg/mL glucose oxidase, 0.4% methylcellulose, 0.2% BSA. Vinculin was 300 nM with 30% Alexa488-labeled monomers, actin was 0.3 µM with 18% Alexa568-labeled monomers, the Arp2/3 complex was 15 nM, WAVE WA was 80 nM. Imaging started when actin filaments enter the evanescent field. Time-lapse microscopy was recorded every 15 s. Control flow chambers assembled with BSA-coated patterns were injected with the same solution. The reaction was monitored with TIRFM using the Widefield/TIRF – Leica SR GSD 3D microscope described above.

Microfluidics and TIRF microscopy

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Microfluidic channels were assembled as described above using a sticky-Slide VI 0.4 (ibidi GmbH, Germany). The microfluidic channel inlet was connected to a PHD 2000Infusion/Withdraw pump (Harvard Apparatus, USA). The flow was set to 25 µL per min. Typically, (1) 0.8–1 µM actin with 18% Alexa568-labeled monomers, 0.5–1 µM vinculin with 30% Alexa488-labeled monomers, 2–4 µM talin VBS1, were supplied in the above-mentioned polymerization buffer from the inlet. (2) At steady state of actin assembly, all soluble components are washed away with five channel volumes, and (3) a solution of a pre-assembled branched actin networks is injected. The branched network was assembled with 0.5 µM actin Alexa647-labeled monomers, 15 nM Arp 2/3 complex and 40 nM WAVE WA, and diluted twofold. Time-lapse imaging of vinculin-actin bundles and branched network started as soon as step (3) was completed. Images were recorded every 15 s with TIRFM using the Widefield/TIRF – Leica SR GSD 3D microscope described above.

Image processing and data analysis of fluorescence images

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Time-lapse videos of filament growth were processed with Fiji software (NIH). For each sample, we measured the rates of barbed end elongation of at least ~10 filaments, typically from at least 20–40 frames of imaging over a span of ~200–400 s, using TIRFM. The growth rate of actin filaments was calculated using Multikymograph Fiji software plugins or a matrix-to-8-bit-image conversion program written in MATLAB, Kymographs were generated over the time course of actin assembly along the actin filament or bundle trace. For filaments or bundles that were stable in the observation field, traces were automatically picked using Multikymograph Fiji software plugins. For mobile structures, traces were manually picked. The alignment of filament pointed-end and internal fluorescence speckles of actin filaments in MATLAB-generated kymograghs accounted for the accuracy of the method.

To measure the zippering velocity of vinculin-induced crosslinking, the length of the zippered region of single filaments was measured over time for in three different experiments where 0.7 to 1.1 µM actin was polymerized in the presence of 600 nM vinculin and 2 µM Talin VBS1, or 300 nM vinculin and 1 µM Talin VBS1, or 300 nM vinculin, 1 µM Talin VBS1, 12 nM Arp2/3 complex and 80 nM WAVE WA. In all three experiments, measured velocities varied within the same range. Determining the number of filaments per vinculin-mediated bundle was conducted by analyzing the fluorescence intensity of bundles using the line-scan tool in Fiji software, and subsequently normalized to the fluorescence intensity of a single actin filament from the same experiment and that was as many times illuminated as the bundle. Additionally, in most cases, time-lapse videos provided bundle formation history hence an evidence from the calculated number of filaments.

To determine the fluorescence ratio of vinculin to actin within patterned actin networks, we analyzed two color widefield fluorescence images taken after 60 min of actin assembly. (1), we piled, separately, the images for each channel (568 for actin or 488 for vinculin signals) and for each patterned geometry, namely continuous circles and dotted circles where the dot-to-dot distance was 6, 11, or 18 µm. (2), Piled images were then aligned according to the patterned geometry using ‘Template Matching’ plugins of Fiji software. (3), By thresholding the vinculin channel images, we created binary masks to isolate the vinculin-decorated network and filter out the background fluorescence for each channel (568 for actin or 488 for vinculin signals). (4) We then generated ratiometric images (488 fluorescence intensity/568 fluorescence intensity, as shown in Figure 2B), using the default plugins in Fiji. (5) Since we kept constant all illumination parameters and all biochemical parameters of the reconstituted assays (except vinculin concentration that was 0.25 or 0.5 µM), we were able to perform quantitative analyses. For all patterns (>1000), the fluorescence density of actin (from actin fluorescence images) and related vinculin to actin fluorescence ratio (from ratiometric images) were measured for each network along the pixel selections presented in Figure 2C. For data analyses, we wrote a MATLAB program to correlate, for each pixel, the vinculin/actin fluorescence ratio to the actin fluorescence density. For each series of pattern geometry (repeated 29 to 70 times), we computed the distribution and the mean value of each network (branched, uniform or mixed polarity) separately. In final statistics, we characterized each type of network by pooling the mean ratio values related to the same actin density value, and calculated their mean and standard deviation, which were plotted as a function of actin fluorescence density (Figure 2D). To estimate the apparent equilibrium dissociation constant of vinculin to actin networks, we calibrated the fluorescence intensities to the protein concentration using labelled actin or labelled vinculin. Mean fluorescence intensities for 3–5 known protein concentrations were measured and averaged. Calibrations were carried out using pegylated glass and coverslips using the same reaction chamber, volumes and composition as for the experiments. The mean fluorescence intensities were converted into protein concentration in the networks. This allowed to determine a steady state saturation curve showing local concentration of bound vinculin, which represents the local concentration vinculin-actin complex, as a function of local actin concentration. The apparent equilibrium dissociation constant was then obtained by fitting the steady state saturation curve using Graphpad software and assuming mass-action binding at equilibrium.

Synthesis of tiopronin Monolayer-Protected AuNPs

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Water-soluble tiopronin-protected gold nanoparticles (AuNP-TP) were synthesized as described previously (Dahan et al., 2018). Briefly, 0.1 g of Gold(III) chloride trihydrate (Sigma–Aldrich) and 0.11 g of N-(2-mercaptopropionyl) glycine (Tiopronin, Cayman Chemical) were dissolved in 13 mL methanol:acetic acid solution (6:1 v/v) and gently stirred at room temperature for 1 hr. 1 mL of 1M sodium borohydride (Acros Organics) in deionized water was prepared and immediately added to the solution. The solution was vigorously stirred for 2 hr at room temperature. The solution was dialyzed against deionized water for 72 hr with a 8–10 kDa Float-a-Lyzer G2 (Spectrum). Dialyzed solution was stored at 4°C.

Functionalization of tiopronin Monolayer-Protected AuNPs

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AuNP-TPs were coupled to GST-VBS1 in a two-step reaction. First, a concentrated solution of AuNP-TPs (OD = 1 at 520 nm) in 0.1M, pH6 MES buffer were ‘activated’ by incubating with 40 mM EDC (1-ethyl-3-[3-dimethylaminopropyl] carbodiimide hydrochloride, Sigma–Aldrich) and 80 mM sulfo-NHS (N-hydroxysulfosuccinimide, Thermo) in 0.1M pH6 MES buffer for 15 min at room temperature. The volume was increased to 1 mL with 0.1M pH6 MES buffer and concentrated using a 30 kDA MWCO centricon (Millipore). This was repeated twice with 0.1M pH6 MES and twice with 2 mM pH 7.4 sodium phosphate buffer. The concentration of the AuNPs was adjusted to an OD = 0.1 at 520 nm and reacted with 1.5 nmole VBS1-GST in a 50 µL final volume for 2 hr at room temperature. The reaction mixture was stored on ice at 4°C for later use. The conjugation reaction was analyzed agarose gel electrophoresis. 16 µL of sample was mixed with 4 µL 30% glycerol and loaded into a 0.75% agarose gel. The gel was run at 90V for 20 min.

Sample preparation for cryo-electron tomography

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In order to favor a random orientation of filaments in solution, and prevent a biased orientation due to excessive filament length, we incubated short preformed actin filaments with vinculin in the presence of talin VBS1. In a buffer containing 20 mM Hepes, pH 7.0, 50 mM CaCl2, 1 mM MgCl2, 1 mM EGTA, 0.5 mM DTT, 0.2 mM ATP, 10 µM Mg-ATP actin monomers were preassembled in the presence 50 nM capping protein α1β2. Thus, the ratio CP:actin monomers was 1:200 yielding filaments of ~0.7 µm in average, unable of end-to-end annealing. Subsequently, 0.5 µM capped actin filaments were incubated for 30 min at 25°C with 1.2 µM full-length vinculin in the presence of 3 µM talin VBS1. When GFP-VBS1 was used, 0.3 µM capped actin filaments were incubated for 1 hr at 25°C with 2 µM vinculin and 8 µM GFP-VBS1. Gold nanoparticles labeling was conducted as followed: 0.3 µM capped actin filaments were incubated for 1 hr at 25°C with 2 µM vinculin, 0.8 µM AuNP-GST-VBS1, and 7.2 µM GST-VBS1. For the experiments using vinculin tail fragment, 0.3 µM capped actin filaments were incubated for 30 min at 25°C with 2.2 µM tail fragment. Next, 4 µL of the reconstituted solution and 1 µL of a 10 nm fiducial gold marker solution (Aurion, Netherlands) was applied onto glow-discharged EM grids coated with a holey carbon mesh (Cu R2/1, 200 mesh, Quantifoil). Finally, the EM grids were manually blotted and plunge frozen in liquid ethane. The EM grids were stored in liquid N2 until used for cryo-ET data acquisition.

Cryo-ET data collection

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Cryo-ET data was acquired on a FEI Titan Krios transmission electron microscope (FEI, Hillsboro, USA) equipped with a GIF Quantum energy filter and a K2-Summit direct electron detector (Gatan, Pleasanton, USA). The microscope was operated at 300 keV in zero-loss mode with the slit width of the energy filter set to 20 eV.

Using SerialEM (Mastronarde, 2005), image stacks were recorded at each tilt angle in super-resolution mode with an electron flux of ~8 electrons per pixel per second.

The image stacks were acquired at a dose-fractionated frame rate of 1 frame per 0.2 s at a magnification of 42,000×, resulting in a pixel size of 0.17 nm. The tilt-series covered an angular range of − 60° to + 60° and were recorded at increments of 2°, at a defocus of −4 μm and a total accumulated electron dose of ~70 electrons per Å2.

The image stacks were 2 × 2 down-sampled and subjected to motion-correction using MotionCorr (Li et al., 2013), resulting in a final pixel size of 3.4 Å. Next, the tilt-series were contrast transfer function corrected (Eibauer et al., 2012) and finally reconstructed in volumes with a size of 1024 × 1024×512 voxel (total down-sampling is 8 × 8, voxel size of 13.6 Å) using the TOM Toolbox (Nickell et al., 2005).

Data acquisition for samples containGFP-VBS1 and AuNP-TPs-GST-VBS1 vinculin bundles were acquired using a dose-fractionated frame rate of 6 frames per 1.2 s with a magnification of 64,000 × resulting in a pixel size of 0.11 nm. The tilt-series covered an angular range of − 60° to + 60° and were recorded at increments of 3°, at a defocus of −4 μm and a total accumulated electron dose of ~85 electrons per Å2. Reconstruction of the tomograms was done with IMOD (Kremer et al., 1996), using gold fiducials for tilt-series alignment and weighted back-projection reconstructions. Cryo-tomograms were 2 × 2 down-sampled for reconstruction resulting in a pixel size of 2.22 Å.

Structural analysis of actin filaments from bundles by cryo-ET

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Actin filaments were automatically segmented, as we described in Martins et al., 2020, from 16 tomograms using a convolutional neural network algorithm (Chen et al., 2017). Based on the segmentations, 3D coordinates of actin segments were chosen along the filaments with a spacing of 16.5 nm. Next, actin segments were reconstructed into sub-tomograms of 144 × 144×144 voxels, corresponding to a box size of ~50 nm and pixel size of 3.4 Å. A total number of 39,231 actin segments were extracted. The 3D coordinate extraction from segmentations was performed using MATLAB (Mathworks, Natick) and the sub-tomogram reconstruction was performed using the TOM Toolbox.

Next, the sub-tomograms were aligned parallel to the x-axis. A 22 nm thick central section of the aligned sub-tomograms was projected and an 11 nm thick filamentous mask along the x-axis was applied.

Subsequently, we performed one round of 2D classification with the pre-aligned using RELION (Scheres, 2012). Thereby, we removed low-quality particles and false positive particles (e.g. gold markers, background), resulting in a cleaned particle set of 29,643 actin segments. Representative 2D classes are shown in Figure 3—figure supplement 4B.

Next, we subjected these particles to a helical 3D-refinement using RELION, using a cylinder as an initial template (He and Scheres, 2017), a helical rise of 27.6 Å and a helical twist of 166.7 degrees (Galkin et al., 2015). The obtained 3D reconstruction of actin (Figure 3—figure supplement 4C) resolved to ~14 Å, as indicated by the RELION software. Actin structures were visualized with UCSF Chimera (Pettersen et al., 2004).

Mapping the polarity in actin bundles

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Using the segments that contributed to the final actin structure, all transformations that were applied to the segments during reconstruction (that are pre-alignment, projection, and helical reconstruction), were inverted. The inverted transformations describe how an actin segment is oriented at the position of its extraction in the tomograms. In particular, the first Euler rotation of a segment around the z-axis (termed psi angle) resolves the segments polarity, given that the barbed- and pointed ends can be identified in the reconstructed structure. As expected, histograms of the psi angles show two peaks per bundle, 180° apart (Figure 3—figure supplement 5).

In the next step, we connected the segments back to filaments. In principle, this information is contained in the initial segmentations. However, the automatic segmentation procedure tends to fuse filaments, if they run parallel or cross each other with small distances. Therefore, we manually combined the segments into filaments. When filament continuity within the tomogram was questionable, we kept the smaller pieces, so that an unambiguous assignment was possible. Therefore, this procedure underestimates the length of certain filaments. However, it has no influence on the parameter we investigated in this work, namely the polarity and distances between actin filaments.

In order to determine the polarity of filaments we plot the segments belonging to each filament as columns of circles (Figure 3—figure supplement 5). The color was assigned according to segment transformation, left (blue) or right (red) psi angle. The columns show a clear tendency to be either blue or red, confirming that we determined the direction of actin segments belonging to the same filaments in a consistent manner. Since the procedure did not consider which segment belongs to which filament, this result serves as an internal control. Since an actin filament can only have a single polarity, we decided the final direction of a filament based on the majority of its segments and mapped the direction of a filament accordingly (Figure 3—figure supplement 4D, Figure 3—figure supplement 5).

A confidence score measurement providing an additional confirmation to the reliability of determining the polarity of each filament (Figure 3—figure supplement 6A). We termed this score the ‘majority confidence score’. It is defined as the maximum number of segments pointing in a similar direction divided by the total number of segments of the respective filament. In this study we extracted 606 filaments from 16 tomograms. Around 20% of the filaments had a majority confidence score <2/3 (Figure 3—figure supplement 6A). These filaments were excluded from further analysis, for example the black filaments in Figure 3—figure supplement 4D. In order to measure the distance between the filaments and to quantify the distribution of uniform polarity and mixed polarity regions within the bundles, we performed a 3D neighborhood analysis that described in details in Martins et al., 2020.

In brief, for every actin segment we extracted the polarities of the (up to) three nearest segments of neighboring filaments, within a 3D distance of 40 nm, defining its neighborhood. Based on that we determined for each segment the degree of uniform polarity (UP) and mixed polarity (MP) of its neighborhood (Figure 3—figure supplement 6C). Next, we defined a score (MPs-UPs)/(MPs+UPs) to characterize the overall polarity of each bundle.

The calculated scores range from −1 to 1, for example if all filaments in a bundle point in the same direction the score is −1 (Figure 3—figure supplement 4E). The 3D neighborhood analysis was programmed in MATLAB.

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Decision letter

  1. Edward H Egelman
    Reviewing Editor; University of Virginia, United States
  2. Suzanne R Pfeffer
    Senior Editor; Stanford University School of Medicine, United States
  3. Christoph Ballestrem
    Reviewer; University of Manchester, United Kingdom

In the interests of transparency, eLife publishes the most substantive revision requests and the accompanying author responses.

[Editors’ note: the authors submitted for reconsideration following the decision after peer review. What follows is the decision letter after the first round of review.]

Thank you for submitting your work entitled "Talin-activated vinculin interacts with branched actin networks to initiate bundles" for consideration by eLife. Your article has been reviewed by four peer reviewers, one of whom is a member of our Board of Reviewing Editors, and the evaluation has been overseen by a Senior Editor. The following individual involved in review of your submission has agreed to reveal their identity: Christoph Ballestrem (Reviewer #4).

Our decision has been reached after consultation between the reviewers. Based on these discussions and the individual reviews below, we regret to inform you that your work will not be considered further for publication in eLife.

Overall, there were concerns about the novelty of the work and the new insights that might be gained into function. While some of the points raised below might be addressed with new experiments or even more detailed explanations, it was felt that the scope of the revisions needed were outside of the bounds of what is required for an eLife resubmission. We realize that this decision will be disappointing but hope that you will find these reviews helpful in planning your next experiments.

Reviewer #1:

This is a rather complicated study that aims to provide new insights into how talin and vinculin interact with actin to form bundles in cells. The work done is purely in vitro, which is necessary to simplify and control the system. While some aspects appear novel, I will need to defer to other reviewers in terms of the biological significance of the results. I am not bothered at all about the seeming conflicts with Janssen et al., 2006, since Kim et al., 2016, came to very different conclusions. The work on actin polarity appears to have been done well and is convincing.

Reviewer #2:

The finding that vinculin does not decorate actin without vinculin activation is not novel and thus not surprising. The finding that the addition of talin VBS1, which activates vinculin, results in vinculin binding to polymerizing actin filaments is also neither new and thus not surprising. The quantification of the speed of active vinculin binding to the cytoskeleton is a new detail in the field and perhaps of importance from a mechanistic point of view of nascent adhesion formation.

Likewise, the observations about the robustness of these vinculin-actin interactions is interesting while again the idea that filament elongation and vinculin binding occurring simultaneously have also already been known.

The α-actinin observations with regards to simultaneous binding with vinculin to actin are interesting but perhaps not fully explored and characterized.

How were "rather short filaments" generated?

The analysis of the bundled actin seems problematic where "bridging densities" might be over interpreted as vinculin dimers. Given that the vinculin tail domain binds to actin, I assume the authors are suggesting that the vinculin head domain dimerizes which has not been demonstrated biochemically or structurally. What would cause the vinculin head domain to dimerize? Thompson et al., 2013, suggested that the vinculin tail domain dimerizes but the "bridging densities" do not seem to correlate with such a dimer.

Figure 1A, why 0.6 and 9.3 micromoles of vinculin instead no vinculin at all?

Figure 2A-C is actin alone but 2E analyzes vinculin, where are those actin-vinculin images please?

Figure 2F leaves room for several interpretations.

Figure 3, there seems to be a crucial control missing, actin plus Arp2/3 without vinculin to exclude the possibility that the observed effects are solely due to Arp2/3.

Figure 6, the dimerization is debatable and without any integrin experiments this is also speculative while much of the rest of the model has been published by Waterman's lab although some of her key findings are ignored.

As a side note, the supplementary figures are not labeled and take an effort to track

Figure 1—figure supplement 2, (G) and (H) pull-downs – are all lanes in (G) pellet and are lanes in (H) supernatant and its suggested that only lanes 1-8 are but then what are the other lanes – if they all are pellet (G) and all supernatant (H) then with some actin in the supernatant and vinculin in the pellet, any interpretation would be tricky

I am not certain if Arp2/3 and vinculin binding are sterically hindering each other, surely this depends on the affinity. Arp2/3 is supposed to (biochemical data are missing in the literature) bind the proline-rich region connecting the vinculin head and tail of severed vinculin while the vinculin tail domain binds actin, i.e. two different regions on vinculin harbor the Arp2/3 and actin binding sites.

This study does, however, have some interesting novel aspects. For example, the authors use vinculin that they generated in insect cells which is a plus and new for the field that used to use E. coli expressed vinculin. I wonder if there are any remaining residues on this C-terminus after cleavage of the tag and if the C-terminal tag would prevent the closed vinculin conformer.

Videos – what are the arrows pointing to? Without a legend they are difficult to follow.

I am left with the feeling that this manuscript is hard to follow because of too many tangential experiments while a lot has already been published which dilutes the novelty here, that I mentioned above.

Reviewer #3:

The manuscript entitled "Talin-activated vinculin interacts with branched actin networks to initiate bundles" by Paterski et al., describes studies characterizing the underlying mechanism of actin network reorganization and bundling by talin-activated vinculin. Interactions between full-length vinculin and F-actin are monitored, using a simplified in vitro experimental setup to recapitulate actin reorganization in the early events of adhesion assembly. The investigators observe that talin-activated vinculin crosslinks actin filaments into stable and flexible bundles with uniform and mixed polarity using cryo-electron tomography and micro-patterned surface approaches. Further, talin-bound active vinculin is also observed to promote Arp2/3 complex-induced branched actin network organization and dynamics.

Vinculin is a scaffolding protein that localizes to focal adhesions (FA) and adherens junctions where it links the actin cytoskeleton to the adhesive super-structure. It is an abundant and ubiquitously expressed cell adhesion protein that plays a key role in mediating cell adhesion, motility, and cellular response to force. Understanding how talin-mediated activation of full length vinculin promotes actin cytoskeletal rearrangements and how other interacting proteins such as Arp2/3 and actinin modulate these actin networks, will aid in deciphering how vinculin couples the extracellular matrix with the actin cytoskeleton to regulate morphology, motility and mechano-transduction. However, concerns exist with the manuscript, as key controls are missing and some of the data presented does not agree with the existing data/literature. Some of the discrepancies are not acknowledged and discussed.

Detailed comments are listed below:

1) The investigators find that the mean distance between F-actin filaments within the F-actin bundle is 21 +/- 6 nm. This is twice as large as inter-filament distance previously measured by Cryo-electron tomography experiments using isolated Vt domain (10.5 nm) (Janssen, et al., 2006). In the Discussion, authors cite this previous paper without bringing the actual numbers for comparison, and do not elaborate on this difference. However, this difference is crucial for the mechanism of vinculin-mediated F-actin bundling. If the distance between F-actin filaments is increased 2-fold, two Vt domains cannot interact. This would imply that vinculin in the assays should mediate bundling through interaction with linker region or head domain. Indeed, the authors speculate: "Thus, the ~21 nm interspace within vinculin-mediated bundles may represent the length of an open vinculin protein suggesting that vinculin's head domain may play a role in the dimerization process to produce an activated full-length vinculin dimers." However, this is counter to an abundance of evidence in the literature which support actin binding and dimerization primarily through the tail domain.

The disconnect with published literature raises the following concerns:

i) Does Alexa88-modified full length vinculin alter actin bundling and/or other vinculin interactions? Actin binding but surprisingly not bundling controls are reported for baculovirus expressed full length vinculin but not the fluorescent-tagged protein. A comparison of actin binding and bundling is needed for both non-modified baculovirus expressed and fluorescently-tagged vinculin, in the absence and presence of the talin VBS1 peptide.

ii) Alternatively, could the actin "bundling" observed be due to non-specific association of vinculin head domains (Vh) and linkers connecting Vh to Vt? In cells, the position of Vh is strongly restrained by its binding to cell membrane through talin. The experiments herein are performed on a 2D coverslip, where Vh is free to move and potentially create gel-like structures, bridging F-actin filaments together. As a result, Vh domains may be forced between actin filaments (since they cannot escape in 3D) and may prevent F-actin fibers from coming close enough for Vt domains to interact?

2) The authors observe that vinculin and a-actinin colocalize within F-actin bundles. In support of this observation the authors argue that a-actinin and vinculin supposedly have compatible inter-filaments spacing (~20 nm). This conclusion contradicts published literature (P. Kanchanawong, et al., Nanoscale architecture of integrin-based cell adhesions. Nature 468, 580-4 (2010); L. B. Case, et al., Molecular mechanism of vinculin activation and nanoscale spatial organization in focal adhesions. Nat. Cell Biol. 17, 880-892 (2015)), in which a-actinin and vinculin mostly occupy distinct layers, consisting of a membrane-associated integrin signaling layer, a force transduction layer containing talin and vinculin, and an actin-regulatory layer containing a-actinin as well as zyxin and VASP.

3) Known vinculin activation, actin binding and bundling mechanisms should be discussed, to put the data into context. For example, upon vinculin activation (which is still not completely understood), actin binding to Vt promotes a conformational change in the tail domain that facilitates actin dimerization and bundling. Moreover, actinin engages the head domain and can promote Vt activation in a manner similar to talin (Bois et al., JBC, 2006), which may be why these activating ligands localize to different “layers” within the focal adhesion. Arp2/3 engages the proline rich domain and may not compete with direct actin binding interactions through the tail domain. As mentioned above, in cells, vinculin has been shown to localize to different pools, which are “lost” in the simplified in vitro system used here. Thus, the investigators need to extrapolate their findings to a physiologically relevant system whereby vinculin localization occurs with distinct ligands.

4) The authors state that vinculin-mediated F-actin bundles have mixed polarity. The quantity plotted at Figure 2E is (MPs-UPs)/(MPs+UPs), where MPs describes degree of mixed polarity of neighboring fibers, and UPs describe degree of uniform polarity. However, the investigators do not adequately provide a definition of MPs and UPs. Figure 2F depicts crosslinking bridge between the actin filaments. Can the authors elaborate on the orientation of vinculin at these crossing junctions? If bundling is mediated by the tail domain, it is unclear how Vt can dimerize to form a mixed bundle. Moreover, while the mixed polarity observation is intriguing, it is unclear how this fits with published findings that vinculin forms directionally asymmetric catch bonds with F-actin (Huang et al, 2017). Such "mixed polarity bundles" should be very unstable in vivo and shear apart under force.

5) It is apparent from Figure 2D that the actin bundles, except bundle-5, show mixed polarities. However, Figure 2E shows bundles 4,5, and 7 that exhibit uniform polarity. It would be helpful to clarify the apparent discrepancy between Figure 2D and 2E. Moreover, bundles 5 and 7 in Figure 2D depict long and uniform filaments of actin in sharp contrast to bundles 4 and 16. Some discussion is needed to explain these differences.

6) The authors observe that vinculin doesn't enhance F-actin polymerization rate. This observation is also in contradiction with previously published results (Wen et al, 2009; Jannie et al., 2015) which showed that vinculin promotes F-actin polymerization.

7) The vinculin-actin interaction model presented in the Figure 6 seems a bit misleading as it gives the impression that vinculin forms a dimer at the talin interface through its head domain. Actin binding to vinculin has been shown to be mediated primarily through the tail domain. Some discussion is needed here.

8) All proteins are conjugated with covalent fluorophores with varying degree of labeling. And since various parameters noted in the paper are calculated based on the fluorescence quantification coming from these fluorophores, it would be helpful if the authors tabulate the protein with respective fluorophore and degree of labelling and their effective percentages used in fluorescence ratio calculation for each experiment.

Reviewer #4:

Boujemaa-Paterski and colleagues use a minimal in vitro system to mimic situations in cells where initial adhesion complexes become linked to the branched actin network in protruding lamellipodia of migrating cells.

Major findings are that only vinculin activated by talin is able to bind and bundle actin filaments of mixed population; another focal adhesion protein, a-actinin1, is attracted at later stages. When immobilised on micropatterns, activated vinculin traps and organises (bundles) Arp2/3 induced branched actin filaments. At first, many of these individual aspects may not seem entirely novel but the comprehensive way of combining cutting edge technology reveals in a very clear manner how individual components, following specific activation steps, act together to tether initial adhesion complexes to the actin network.

Points to consider:

1) Authors speculate that vinculin dimerization involving its head domain may be important for the bundling function but they do not show this. A potential experiment to clarify could be to use the actin binding vinculin tail only for their experiments.

2) Vinculin itself binds to the Arp2/3 complex. Does this interaction have a role in trapping the Arp2/3 induced branched network?

3) A limitation of the study may be it has not included talin itself, which binds and organises actin prior to vinculin. The presence of talin could change protein binding kinetics and aspects of actin organisation. New experiments may be beyond the scope of this study but this matter requires discussion in the relevant section (see comment point 5).

4) There is no statistics to the histogram in Figure 3A.

5) The Discussion remains superficial in relevant aspects:

a) In a somewhat similar study, Ciobanasu et al., 2014, have shown how the talin-vinculin complex leads to actin anchoring and a-actinin contributes to the crosslinking (bundling) of the initial network. Questions arising are, whether for example kinetics of a-actinin (and vinculin) binding to filaments or the distance between filaments etc. would change in presence of full-length talin?

b) A recent study (Atherton et al. JCB) shows that the release of an autoinhibition motif in talin unmasks VBSs that recruit and activate vinculin. The finding seems in line with the present study showing that an isolated VBS can activate vinculin to bind to actin and should be discussed.

c) There is little discussion about the potential role of Arp2/3 binding to vinculin (DeMali et al., 2002 and other manuscripts). Could the Arp2/3 binding site in vinculin contribute to the connection to the actin meshwork?

[Editors’ note: further revisions were suggested prior to acceptance, as described below.]

Thank you for submitting your article "Talin-activated vinculin interacts with branched actin networks to initiate bundles" for consideration by eLife. Your article has been reviewed by four peer reviewers, one of whom is a member of our Board of Reviewing Editors, and the evaluation has been overseen by Suzanne Pfeffer as the Senior Editor. The following individual involved in review of your submission has agreed to reveal their identity: Christoph Ballestrem (Reviewer #4).

The reviewers have discussed the reviews with one another and the Reviewing Editor has drafted this decision to help you prepare a revised submission.

We would like to draw your attention to changes in our revision policy that we have made in response to COVID-19 (https://elifesciences.org/articles/57162). Specifically, we are asking editors to accept without delay manuscripts, like yours, that they judge can stand as eLife papers without additional data, even if they feel that they would make the manuscript stronger. Thus the revisions requested below only address clarity and presentation.

The reviewers have agreed that the revised paper is a substantial improvement over the original submission, and is close to being ready for acceptance in eLife. However, in a somewhat unusual move, the reviewers thought that the final paper should contain less data than what is currently shown (rather than more), due to the major concerns about Figure 3—figure supplement 2. Since it was felt that the biochemistry shown here is not essential to the paper, the easiest path is simply to have you remove it rather than asking for a major revision where this work would be redone.

One reviewer stated:

Panels F and H cannot be unambiguously interpreted:

Their pull-down shows that their vinculin aggregates and thus pellets so one cannot say anything about vinculin binding to F-actin if the readout is its pelleting, as it pellets in the absence of F-actin also. More complications are caused by F-actin not completely being polymerized as they also have actin in the supernatant. To me, these data are thus uninterpretable. However, they get the assay to work in panel G where vinculin is not aggregating, and actin does not seem to be unpolymerized.

Since the other reviewers are positive about this manuscript and since these panels are not even mentioned in the manuscript, one way to reconcile the three reviewers would be to take those panels out.

Also please correct this issue and clarify the text as needed:

With regard to Figure 3—figure supplement 2, it is concerning that full length vinculin pellets in the presence of VBS1 but in the absence of actin (vinculin oligomerization, stability issues?), and that actin is present in the soluble fraction upon polymerization. This raises additional concerns on my end regarding use of purified vinculin for their assays. Also, in Materials and methods, it is stated that the pelleting speed is 380,000, but in the figure legend, the pelleting speed is stated as 80,000.

https://doi.org/10.7554/eLife.53990.sa1

Author response

[Editors’ note: The authors appealed the original decision. What follows is the authors’ response to the first round of review.]

Reviewer #1:

This is a rather complicated study that aims to provide new insights into how talin and vinculin interact with actin to form bundles in cells. The work done is purely in vitro, which is necessary to simplify and control the system. While some aspects appear novel, I will need to defer to other reviewers in terms of the biological significance of the results. I am not bothered at all about the seeming conflicts with Janssen et al., 2006, since Kim et al., 2016, came to very different conclusions. The work on actin polarity appears to have been done well and is convincing.

We thank the reviewer for his insightful comments and for his appreciation of the work.

We reorganized the manuscript in order to focus the manuscript around the central claim, namely the ability of activated vinculin to reorganize lamellipodial-like branched actin networks.

We hope that displaying vinculin’s ability to affect such networks from the mesoscale, to the macroscale, down to the filament scale will make our study less complicated and easier to follow.

Our results obtained with single-filament TIRF microscopy as well as the structural analysis of our in vitro vinculin-F-actin bundles using cryo-ET and image processing are now used as supplementary information to the reorganization of Arp2/3 complex-branched followed by TIRF microscopy.

Reviewer #2:

The finding that vinculin does not decorate actin without vinculin activation is not novel and thus not surprising. The finding that the addition of talin VBS1, which activates vinculin, results in vinculin binding to polymerizing actin filaments is also neither new and thus not surprising. The quantification of the speed of active vinculin binding to the cytoskeleton is a new detail in the field and perhaps of importance from a mechanistic point of view of nascent adhesion formation.

We agree with the reviewer’s concerns raised concerning the novelty of the ability of talin-VBS1 to relieve vinculin autoinhibition (Papagrigoriou et al., 2004). Our study focuses on the ability of vinculin to bind and bundle branched actin network. For that purpose, we used the published knowledge and activated vinculin in order to study its intrinsic properties with regard to actin binding and bundling.

To the best of our knowledge the quantifications (using full length protein) are novel and may explain the efficacity at which talin-vinculin complexes can engage and reorganize lamellipodial branched-actin networks at the nascent adhesions.

Likewise, the observations about the robustness of these vinculin-actin interactions is interesting while again the idea that filament elongation and vinculin binding occurring simultaneously have also already been known.

We would like to emphasize here that there was a discrepancy around the question of how vinculin affect actin polymerization.

Using spectroscopic measurements, the isolated vinculin tail domain (Vt, only 18% of the full length protein) was first shown to activate actin polymerization (Wen et al, 2009). This was contradicted by LeClainche et al., who proposed that Vt specifically blocks actin filaments barbed end elongation via its C-terminal amino-acids. Namely, it leads actin filaments to elongate only from their pointed ends. In addition, the study also showed that full length vinculin expressed in bacteria (and activated by VBS1) fails to block the barbed end of actin filament (Le Clainche et al., 2010).

We acknowledged the discrepancy in our revised manuscript. Yet, the dynamics of vinculin-F-actin bundles formation, the dynamics and the orientation of filaments within the bundles, the flexibility of bundles were not yet reported. Therefore, these are novel and important basic characteristics of vinculin.

The α-actinin observations with regards to simultaneous binding with vinculin to actin are interesting but perhaps not fully explored and characterized.

We agree with the reviewer’s comment. We fell that an in-depth comparison of the effects of vinculin and α-actinin is beyond the scope of our study, therefore, the α-actinin observations were removed.

How were "rather short filaments" generated?

To preform short actin filaments, we first polymerized actin monomers in the presence of the capping protein α1β2, CP, with a 1:200 molar ratio CP:actin. Given that CP binds actin filament barbed ends fast and stably (Kd ≈ 0.1 nM, Schafer et al., 1996), we expected CP-capped actin filaments to measure 0.7 µm in average.

This information is now provided in the revised manuscript, Figure 3—figure supplement 4 legend and in the revised Material and methods section.

The analysis of the bundled actin seems problematic where "bridging densities" might be over interpreted as vinculin dimers. Given that the vinculin tail domain binds to actin, I assume the authors are suggesting that the vinculin head domain dimerizes which has not been demonstrated biochemically or structurally. What would cause the vinculin head domain to dimerize? Thompson et al., 2013 suggested that the vinculin tail domain dimerizes but the "bridging densities" do not seem to correlate with such a dimer.

We agree with the reviewer that only structural determination would allow an unambiguous statement about the vinculin bridges, seen in our tomograms. However, since vinculin does not seem to form homogeneous coating of the actin, we could not identify the structure of vinculin within the averaged structures. Nevertheless, we preformed additional experiments to identify vinculin structures and locations within tomograms of vinculin-induced actin bundles.

First, we increased the density of bridges by activating vinculin with GFP-talin-VBS1 construct. Second, we used ~2.4 nm gold-nanoparticles (AuNPs) that were synthetized, coupled to talin-VBS1 and used to activated vinculin and induced bundles. These experiments were added as a Figure 3—figure supplement figure 3 and the technical details can be found in the Material and methods section.

As expected, the densities observed between the filaments (“bridges”) are irregularly distributed along the actin as well as the gold-labelled VBS1.

As a comparison, we reconstructed actin bundles using the Vt (Figure 3—figure supplement 3A and Material and methods section).

Figure 1A, why 0.6 and 9.3 micromoles of vinculin instead no vinculin at all?

Control experiments are now provided in Figure 3—figure supplement 1.

Figure 2A-C is actin alone but 2E analyzes vinculin, where are those actin-vinculin images please?

The figure indeed showed bundles which were assembled in the presence of talin-VBS1-activated vinculin.

In the new experiments, the occurrence of these densities was increased. This information is provided in Figure 3—figure supplement 3.

Figure 2F leaves room for several interpretations.

We agree that in a context where vinculin is not attached to a surface, the inter-filament spacing we measured may reflect interactions within vinculin dimer, as previously depicted by Molony and Burridge, 1985, or of a different nature, and that may not be seen in vivo. However, since this is not the major aim in this study, we avoid this analysis.

In this analysis, we rather focused on determining the polarity of filaments within vinculin-F-actin bundles, which to our understanding is a key intrinsic property of vinculin that explains its ability to bundle the intermeshed, branched lamellipodial actin network. However, it is plausible that previous experiments using Vt (187 aa) may not represent the precise functional mechanism of full-length vinculin (1066 aa).

Figure 3, there seems to be a crucial control missing, actin plus Arp2/3 without vinculin to exclude the possibility that the observed effects are solely due to Arp2/3.

We have added the requested controls and quantifications. No actin bundling in the absence of activated-vinculin was detected. The information is provided in Figure 3.

Figure 6, the dimerization is debatable and without any integrin experiments this is also speculative while much of the rest of the model has been published by Waterman's lab although some of her key findings are ignored.

We would like to mention that the work of Waterman’s lab was of course cited also in the previous version. However, our revised manuscript stressed the fact that Thievessen et al., 2013 have shown that vinculin tail interacts with lamellipodial actin and affects nascent adhesions turnover. We also highlighted that due to actin-generated tension vinculin adopts an extended conformation (Case et al., 2015). This can be found in the revised manuscript. Here, we show direct interactions between the proteins, while inside cells it is much more challenging to identify such direct interactions.

Moreover, Thievessen et al. paper stated that “…As the resolution of our TFM was not sufficient to analyze nascent FA, we restricted our analysis to mature FA…”. In addition, Xu et al. 2012 Nat. Methods, reported that investigation of the lamellipodial actin was hindered due to actin density and diffraction-limited scale of nascent adhesions.

While cellular study is of a major importance, biochemical and dynamics properties in vitro are fundamental to understand cellular behavior in details. In our study, vinculin interactions with actin are investigated in in vitro systems that may be relevant for the early-stage of nascent adhesion formation. As discussed in the revised manuscript, several lines of evidence (Bachir et al., 2014 ; Atherton et al., 2020; Dedden et al., 2020 ; Kelley et al., 2020) show that talin-vinculin precomplex can form before associating with integrins. On the basis of our reconstitution experiments, we propose that these precomplexes may stably interact with branched lamellipodial actin, and bundles can be initiated in a force independent manner. The formation of bundles from lamellipodial actin may stabilize nascent adhesions, which is necessary for their maturation into focal adhesions. Additionally, our in vitro reconstitution experiments unambiguously demonstrate that the reorganization of the lamellipodial network induced by vinculin slow down its progression (Figure1, in agreement with in vivo reports by Choi et al., 2008 and Thievessen et al., 2013) which may serve as a key step for the formation of a stable connection between actin – talin-vinculin, at nascent adhesions.

As a side note, the supplementary figures are not labeled and take an effort to track

Figure 1—figure supplement 2, (G) and (H) pull-downs – are all lanes in (G) pellet and are lanes in (H) supernatant and its suggested that only lanes 1-8 are but then what are the other lanes – if they all are pellet (G) and all supernatant (H) then with some actin in the supernatant and vinculin in the pellet, any interpretation would be tricky.

We apologize for the inconvenience. All figures are labelled. The figure legend has been rephrased and the revised figure is now Figure 3—figure supplement 2.

I am not certain if Arp2/3 and vinculin binding are sterically hindering each other, surely this depends on the affinity. Arp2/3 is supposed to (biochemical data are missing in the literature) bind the proline-rich region connecting the vinculin head and tail of severed vinculin while the vinculin tail domain binds actin, i.e. two different regions on vinculin harbor the Arp2/3 and actin binding sites.

Our quantifications of the binding of Arp2/3 and vinculin alone or together indicated that the differences fall within the standard deviation of measurements. Thus, we do not see significant hindering of one in the presence of the other.

We also did not record any activation of Arp2/3 complex by vinculin. This suggest that vinculin may interact with a cellular subpopulation of Arp2/3 (Chorev et al., 2014) or an additional factor is needed for the two to form a complex.

The information is provided in revised Figure 3.

This study does, however, have some interesting novel aspects. For example, the authors use vinculin that they generated in insect cells which is a plus and new for the field that used to use E. coli expressed vinculin. I wonder if there are any remaining residues on this C-terminus after cleavage of the tag and if the C-terminal tag would prevent the closed vinculin conformer.

We thank the reviewer for the supportive comment.

Indeed, the vinculin was expressed and purified from insect cells is in a close conformation, as we previously determined its structure by X-ray crystallography (Chorev et al., 2018).

Videos – what are the arrows pointing to? Without a legend they are difficult to follow.

We apologize for missing the legends for these videos. Legend are now provided.

I am left with the feeling that this manuscript is hard to follow because of too many tangential experiments while a lot has already been published which dilutes the novelty here, that I mentioned above.

We agree with the reviewer and focused the manuscript in order to make its message clearer.

Reviewer #3:

The manuscript entitled "Talin-activated vinculin interacts with branched actin networks to initiate bundles" by Paterski et al., describes studies characterizing the underlying mechanism of actin network reorganization and bundling by talin-activated vinculin. Interactions between full-length vinculin and F-actin are monitored, using a simplified in vitro experimental setup to recapitulate actin reorganization in the early events of adhesion assembly. The investigators observe that talin-activated vinculin crosslinks actin filaments into stable and flexible bundles with uniform and mixed polarity using cryo-electron tomography and micro-patterned surface approaches. Further, talin-bound active vinculin is also observed to promote Arp2/3 complex-induced branched actin network organization and dynamics.

Vinculin is a scaffolding protein that localizes to focal adhesions (FA) and adherens junctions where it links the actin cytoskeleton to the adhesive super-structure. It is an abundant and ubiquitously expressed cell adhesion protein that plays a key role in mediating cell adhesion, motility, and cellular response to force. Understanding how talin-mediated activation of full length vinculin promotes actin cytoskeletal rearrangements and how other interacting proteins such as Arp2/3 and actinin modulate these actin networks, will aid in deciphering how vinculin couples the extracellular matrix with the actin cytoskeleton to regulate morphology, motility and mechano-transduction. However, concerns exist with the manuscript, as key controls are missing and some of the data presented does not agree with the existing data/literature. Some of the discrepancies are not acknowledged and discussed.

Detailed comments are listed below:

1) The investigators find that the mean distance between F-actin filaments within the F-actin bundle is 21 +/- 6 nm. This is twice as large as inter-filament distance previously measured by Cryo-electron tomography experiments using isolated Vt domain (10.5 nm) (Janssen et al., 2006). In the Discussion, authors cite this previous paper without bringing the actual numbers for comparison, and do not elaborate on this difference. However, this difference is crucial for the mechanism of vinculin-mediated F-actin bundling. If the distance between F-actin filaments is increased 2-fold, two Vt domains cannot interact. This would imply that vinculin in the assays should mediate bundling through interaction with linker region or head domain. Indeed, the authors speculate: "Thus, the ~21 nm interspace within vinculin-mediated bundles may represent the length of an open vinculin protein suggesting that vinculin's head domain may play a role in the dimerization process to produce an activated full-length vinculin dimers." However, this is counter to an abundance of evidence in the literature which support actin binding and dimerization primarily through the tail domain.

We agree with the reviewer comments, although one should take into consideration that a small part/domain of a protein (Vt,187aa) may not fully reflect the structure and dynamics of a large protein (Vinculin, 1066aa). Therefore, studying the full-length protein, ~117 kDa, is of major importance and relevance in comparison to a ~21 kDa protein segment, the Vt. Moreover, we would like to remind the reviewer here that the ability of vinculin to interact and bundle a branched actin network is the major focus of our work and therefore the actin bundles characterization is moved into the supplementary information.

Nevertheless, we have now reconstituted bundles with Vt and obtained an inter-filament distance of 10 nm, matching the value reported by Janssen et al. We have also developed two additional strategies: increasing the density of vinculin density by activating vinculin with GFP-talin-VBS1 construct, and by using 2-3 nm Gold-nanoparticles that were coupled to talin-VBS1 and used to activate vinculin. This information is added as Figure 3—figure supplement 3 and Material and methods section.

In the context of reconstitution using F-actin and soluble, activated VBS1-vinculin, the inter-filament spacing was 29 nm, and depicted an intrinsic property of activated vinculin. Similarly, the 10 nm inter-filament spacing obtained with the tail is an intrinsic property of the isolated Vt.

We agree that a wealth of evidence extensively characterized in vitro the dimerization of isolated Vt (Johnson and Craig, 1995, Kim et al., 2016, and as reviewed in Thompson et al., 2013). in vivo characterizations measured the conformational change in vinculin (Johnson and Craig, 2000, Case et al., 2015), showing that the open, extended state of vinculin correlates with focal adhesion maturation state. Yet, the dynamics of vinculin’s opening upon its activation by talin as well as its stretching remains unclarified.

Therefore, the inter-filament spacing we reported here might be of interest as it describes an intrinsic property of the full-length protein. Yet, our result does not determine whether the spacing represents a transient intermediate state of vinculin-mediated bundles at the onset of the adhesion formation, nor describe the structure and dynamics of the reconstituted bridges. That is beyond the scope of our study and needs further investigation, although in a recent cryo-ET of analysis (Martins et al., 2020, https://www.biorxiv.org/content/10.1101/2020.03.11.987263v2) we measure a mean distance of 28 nm between filaments at FAs in intact cells.

We clarified our statement as mentioned in the revised manuscript.

We have also moved this information in the supplementary information of Figure 3, which describes the branched network reorganization at the single-filament scale. The revised figure label is Figure 3—figure supplement 4.

The disconnect with published literature raises the following concerns:

i) Does Alexa88-modified full length vinculin alter actin bundling and/or other vinculin interactions? Actin binding but surprisingly not bundling controls are reported for baculovirus expressed full length vinculin but not the fluorescent-tagged protein. A comparison of actin binding and bundling is needed for both non-modified baculovirus expressed and fluorescently-tagged vinculin, in the absence and presence of the talin VBS1 peptide.

We have added a time-lapse sequence showing that unlabeled vinculin expressed insect was unable to bundle actin filaments in the absence of the talin VBS1 domain, while addition of talin-VBS1 activated its bundling activity.

As we reported bundling activity also with unlabeled proteins, we concluded that the bundling observed with labeled vinculin was not due to any perturbation resulting from chemical labeling by Alexa-488.

This information is provided in Figure 3—figure supplement 1.

The reviewer would agree that expressing vinculin (a eukaryotic protein) in eukaryotic expression system is much more reliable than the expression of mammalian protein in prokaryotes, as previously done.

ii) Alternatively, could the actin "bundling" observed be due to non-specific association of vinculin head domains (Vh) and linkers connecting Vh to Vt? In cells, the position of Vh is strongly restrained by its binding to cell membrane through talin. The experiments herein are performed on a 2D coverslip, where Vh is free to move and potentially create gel-like structures, bridging F-actin filaments together. As a result, Vh domains may be forced between actin filaments (since they cannot escape in 3D) and may prevent F-actin fibers from coming close enough for Vt domains to interact?

The experiments in solution are designed to characterize the basic properties of full-length vinculin. In physiological conditions, vinculin binds activated talin, which is attached to integrin (Case and Waterman, 2015) or membrane-associated PIP2 (Kelley et al., 2020), and therefore it is only activated close to the membrane. This property is demonstrated in Figure 4, which shows that bundling of actin occurs when vinculin is indirectly attached to the surface (via talin VBS1). At FAs, the close proximity of vinculin binding domain in talin may force the head domains to be very close to each other, as well as the localization of many talin proteins in close proximity. Therefore, the spatial proximity of the Vh domain at adhesion sites is dictated, while the bulk experiments may force such a proximity in another manner. In any case, in the revised manuscript we removed the claim that Vh-Vh form a dimer, despite the electron microscopy images provided by Molony and Burridge, 1985

We agree with the reviewer that the interactions and affinity between Vh-Vh domains is of prime interest but it is beyond the scope of our study.

2) The authors observe that vinculin and a-actinin colocalize within F-actin bundles. In support of this observation the authors argue that a-actinin and vinculin supposedly have compatible inter-filaments spacing (~20 nm). This conclusion contradicts published literature (Kanchanawong et al., 2010; Case, et al., 2015), in which a-actinin and vinculin mostly occupy distinct layers, consisting of a membrane-associated integrin signaling layer, a force transduction layer containing talin and vinculin, and an actin-regulatory layer containing a-actinin as well as zyxin and VASP.

Due to the comments from reviewer #1 we removed the a-actinin experiments. However, our data does not contradict Dr. Waterman’s findings. In fact, several studies have detected α-actinin at the onset of adhesion assembly, at the stage of nascent ones. Choi et al., 2008 and Bachir et al., 2014, show that α-actinin, concomitantly to vinculin and talin, transiently associates to nascent adhesions during their stabilization and growth phases. These findings do not exclude nor contradict a later recruitment of α-actinin to the acto-myosin associated fibers.

3) Known vinculin activation, actin binding and bundling mechanisms should be discussed, to put the data into context. For example, upon vinculin activation (which is still not completely understood), actin binding to Vt promotes a conformational change in the tail domain that facilitates actin dimerization and bundling. Moreover, actinin engages the head domain and can promote Vt activation in a manner similar to talin (Bois et al., 2006), which may be why these activating ligands localize to different “layers” within the focal adhesion. Arp2/3 engages the proline rich domain and may not compete with direct actin binding interactions through the tail domain. As mentioned above, in cells, vinculin has been shown to localize to different pools, which are “lost” in the simplified in vitro system used here. Thus, the investigators need to extrapolate their findings to a physiologically relevant system whereby vinculin localization occurs with distinct ligands.

We thank the reviewer for this remark. Following these suggestions, we extrapolated our findings to physiologically relevant scenarios and extended our discussion, as mentioned in the revised manuscript, Discussion section.

4) The authors state that vinculin-mediated F-actin bundles have mixed polarity. The quantity plotted at Figure 2E is (MPs-UPs)/(MPs+UPs), where MPs describes degree of mixed polarity of neighboring fibers, and UPs describe degree of uniform polarity. However, the investigators do not adequately provide a definition of MPs and UPs.

We apologize for the unclear term. We have properly defined it in the revised manuscript. Uniform-polarity score (UPs) and mixed-polarity score (MPs) are now provided in the Materials and methods section and in the figure legend Figure 3—figure supplement 4.

Figure 2F depicts crosslinking bridge between the actin filaments. Can the authors elaborate on the orientation of vinculin at these crossing junctions?

In order to reveal the orientation of vinculin along actin, a uniform interaction (binding) along actin filaments is needed. However, it seems that full-length vinculin cannot saturate all sites on actin presumably because of the size of the protein, therefore our trials to determine the structure were unsuccessful.

If bundling is mediated by the tail domain, it is unclear how Vt can dimerize to form a mixed bundle. Moreover, while the mixed polarity observation is intriguing, it is unclear how this fits with published findings that vinculin forms directionally asymmetric catch bonds with F-actin (Huang et al, 2017). Such "mixed polarity bundles" should be very unstable in vivo and shear apart under force.

We do agree with the reviewer’s comment and with this respect we have expanded our Discussion, as mentioned in our revised manuscript.

We speculate that vinculin’s early interactions with the branched network (which comprises branches randomly oriented to each other) are fast and enable the protein to assemble random-polarity bundles. This may be a key mechanism that potentiate the connection to mobile lamellipodial network, which in turn applies a selection pressure reinforcing only uniform polarity bundles and further stabilization and maturation of the nascent adhesions.

Moreover, we have found that in intact cells focal adhesion-associated actin has non-uniform polarity (Martins et al., 2020)

5) It is apparent from Figure 2D that the actin bundles, except bundle-5, show mixed polarities. However, Figure 2E shows bundles 4,5, and 7 that exhibit uniform polarity. It would be helpful to clarify the apparent discrepancy between Figure 2D and 2E. Moreover, bundles 5 and 7 in Figure 2D depict long and uniform filaments of actin in sharp contrast to bundles 4 and 16. Some discussion is needed to explain these differences.

We apologize for being unclear. We now better defined MPs and UPs scores in our revised manuscript. A clearer information is provided in the Materials and methods section and in the figure legend Figure 3—figure supplement 4.

The histogram initially Figure 2E, in the revised manuscript Figure 3—figure supplement 4E, shows a polarity score of filament’s neighborhood. While in most bundles non-uniform polarity was found, the score between the different polarities was variable, covering all possibilities.

In brief, for every actin segment we extracted the polarities of the (up to) three nearest segments of neighboring filaments, within a 3D distance of 40 nm, defining its neighborhood. Based on that we determined for each segment the degree of uniform polarity (UP) and mixed polarity (MP) of its neighborhood (Figure 3—figure supplement 5C). This describes the microenvironment of an actin segment. Next, we defined a score (MPs-UPs)/(MPs+UPs) to characterize the overall polarity of each bundle.

The histogram shows that bundles 4 and 5 have a polarity ratio ≈ -0.2 meaning that ~20% of the segments display an opposite polarity with their closest neighbors (alternance blue/red filaments) and ~80% of them have a uniform polarity (packing blue/blue or red/red filaments), as depicted the directionality maps of these bundles in panel D. Thus, both polarities were detected in most bundles.

6) The authors observe that vinculin doesn't enhance F-actin polymerization rate. This observation is also in contradiction with previously published results (Wen et al., 2009); Jannie et al., 2015) which showed that vinculin promotes F-actin polymerization.

We thank the reviewer for this comment. The two papers mentioned used only the tail (187aa out of 1066aa). Therefore, it may not be not surprising that using a full-length protein, expressed in eukaryotic system represent a more realistic system.

Moreover, Leclainche et al., 2010 already contradicted the above-mentioned literature.

We acknowledged the discrepancy in our revised manuscript.

7) The vinculin-actin interaction model presented in the Figure 6 seems a bit misleading as it gives the impression that vinculin forms a dimer at the talin interface through its head domain. Actin binding to vinculin has been shown to be mediated primarily through the tail domain. Some discussion is needed here.

We thank the reviewer for this comment. We modified the model accordingly and expanded our Discussion.

8) All proteins are conjugated with covalent fluorophores with varying degree of labeling. And since various parameters noted in the paper are calculated based on the fluorescence quantification coming from these fluorophores, it would be helpful if the authors tabulate the protein with respective fluorophore and degree of labelling and their effective percentages used in fluorescence ratio calculation for each experiment.

We thank the reviewer for this valuable comment. Accordingly, we calibrated the fluorescence intensities and we are now able to provide apparent equilibrium dissociation constant that better characterize the affinity of vinculin to the different actin organizations. This information is provided in revised Figures 1 and 2, and the Materials and methods section.

Moreover, the cryo-ET measurements were conducted with unlabeled proteins.

Reviewer #4:

Boujemaa-Paterski and colleagues use a minimal in vitro system to mimic situations in cells where initial adhesion complexes become linked to the branched actin network in protruding lamellipodia of migrating cells.

Major findings are that only vinculin activated by talin is able to bind and bundle actin filaments of mixed population; another focal adhesion protein, a-actinin1, is attracted at later stages. When immobilised on micropatterns, activated vinculin traps and organises (bundles) Arp2/3 induced branched actin filaments. At first, many of these individual aspects may not seem entirely novel but the comprehensive way of combining cutting edge technology reveals in a very clear manner how individual components, following specific activation steps, act together to tether initial adhesion complexes to the actin network.

Points to consider:

1) Authors speculate that vinculin dimerization involving its head domain may be important for the bundling function but they do not show this. A potential experiment to clarify could be to use the actin binding vinculin tail only for their experiments.

We thank the reviewer with this comment.

Although we tuned down this part of the manuscript and discussed in greater details the bundling of branched actin network, we cloned the tail (879-1066) from our human vinculin construct. Reconstitution of actin bundles and structural analysis by cryo-ET showed bundles with an inter-filament spacing of 10 ± 1 nm (Figure 3—figure supplement 3 and 4), in agreement with previous reports (Janssen et al., 2006; Kim et al., 2016).

These experiments clearly demonstrated that Vt (187aa) behaves differently than the full-length vinculin (1066aa). There are several vinculin binding domains in talin that would anyhow force Vh domains to be very adjacent to each other. This would result in actin bundling, and is demonstrated in Figure 4.

Additional experiments revealed increased densities, mediated by talin-VBS1-activated full-length vinculin as well as gold labeling, indicating that bundling by vinculin does not form homogeneous structures as the Vt does. As we now discuss in the revised manuscript these observed bridge densities may also reflect an early, transient state of activated vinculin prior to its extension between the integrin and actin layers, and maybe that actin-generated force is needed to fully extent the protein.

2) Vinculin itself binds to the Arp2/3 complex. Does this interaction have a role in trapping the Arp2/3 induced branched network?

Using TIRFM assays we did not record any activation of Arp2/3 complex by vinculin. This information is added in revised Figure 3.

We speculate that either vinculin interacts with a cellular subpopulation of Arp2/3, as Chorev et al. have identified using mass-spectroscopy analysis (Chorev et al., 2014), or additional factors are needed to mediate the interaction. In Chorev et al., the Arp2/3 complex that contained vinculin was missing the p41-ARC protein. The Arp2/3 complex used in our assays comprises 7 subunits and was purified using a WAVE-WA affinity column chromatography.

3) A limitation of the study may be it has not included talin itself, which binds and organises actin prior to vinculin. The presence of talin could change protein binding kinetics and aspects of actin organisation. New experiments may be beyond the scope of this study but this matter requires discussion in the relevant section (see comment point 5).

We agree with the reviewer and we have expanded our discussed accordingly, as mentioned the revised manuscript. In Figure 4, we synthetically mimic a situation in which several VBS domains are in close proximity to each other. This better resembles the full-length talin and its increased concentration at FAs.

4) There is no statistics to the histogram in Figure 3A.

We indeed implemented our quantifications and the differences falls now within the standard deviation of measurements. The information is provided in revised Figure 3.

5) The Discussion remains superficial in relevant aspects:

a) In a somewhat similar study, Ciobanasu et al., 2014 have shown how the talin-vinculin complex leads to actin anchoring and a-actinin contributes to the crosslinking (bundling) of the initial network. Questions arising are, whether for example kinetics of a-actinin (and vinculin) binding to filaments or the distance between filaments etc. would change in presence of full-length talin?

We thank the reviewer for his comment and have expanded our Discussion accordingly, as mentioned in the revised manuscript. Interestingly, the distances measured between actin filament in FAs, by means of cryo-ET (Martins et al., 2020 , doi.org/10.1101/2020.03.11.987263), resemble the measure distances in this work.

However, we tuned down our initial work of α-actinin 1, since it was felt insufficient by the other reviewers. We do agree that an in-depth comparison of the dynamics of vinculin and α-actinin 1 is of prime interest, but beyond the scope of the present study.

b) A recent study (Atherton et al. JCB) shows that the release of an autoinhibition motif in talin unmasks VBSs that recruit and activate vinculin. The finding seems in line with the present study showing that an isolated VBS can activate vinculin to bind to actin and should be discussed.

We thank the reviewer for this comment and we have expanded our Discussion accordingly, as mentioned in the revised manuscript.

Additionally, during the revision of our manuscript, Kelley and colleagues (Kelley et al., 2020) used phosphoinositide to assemble in vitro activated talin-vinculin precomplexes in absence of tensile forces. These data are in line with our results and were discussed too.

c) There is little discussion about the potential role of Arp2/3 binding to vinculin (DeMali et al., 2002 and other manuscripts). Could the Arp2/3 binding site in vinculin contribute to the connection to the actin meshwork?

There might be several paths that would allow the interaction between Arp2/3 and vilnculin, however, such interactions were not detected in our experimental setup.

The results shown here and in previous studies by Chorev et al. suggest that lamellipodial actin- associated Arp2/3 and adhesion-associated Arp may be two distinct species.

[Editors’ note: what follows is the authors’ response to the second round of review.]

The reviewers have agreed that the revised paper is a substantial improvement over the original submission, and is close to being ready for acceptance in eLife. However, in a somewhat unusual move, the reviewers thought that the final paper should contain less data than what is currently shown (rather than more), due to the major concerns about Figure 3—figure supplement 2. Since it was felt that the biochemistry shown here is not essential to the paper, the easiest path is simply to have you remove it rather than asking for a major revision where this work would be redone.

We agree with the reviewer assessment that the biochemical experiments aimed at determining the affinity of vinculin to actin filaments are not central to this study.

Additionally, we thank the reviewers for their suggestion and believe that it is fully justified.

Therefore, we have removed panels F to H from Figure 3—figure supplement 2 and have revised the manuscript accordingly.

One reviewer stated:

Panels F and H cannot be unambiguously interpreted:

Their pull-down shows that their vinculin aggregates and thus pellets so one cannot say anything about vinculin binding to F-actin if the readout is its pelleting, as it pellets in the absence of F-actin also.

We removed these panels and agree with the reviewers that the experiments could have been improved, however, we would like to clarify the following.

When vinculin was incubated with VBS1 in the absence of actin, a fraction of vinculin pelleted using high-speed centrifugation (380,000 g), while vinculin alone cannot be pelleted.

These observations may represent oligomerization of activated vinculin, as previously reported by Molony and Burridge, 1985.

During data analysis we considered these activated vinculin polymers as our background and subtracted it for the purpose of Kd calculations. In any case it was removed from the manuscript.

More complications are caused by F-actin not completely being polymerized as they also have actin in the supernatant. To me, these data are thus uninterpretable. However, they get the assay to work in panel G where vinculin is not aggregating, and actin does not seem to be unpolymerized.

Unpolymerized actin that remained in the supernatants is a fundamental property of actin and will always be observed due to the equilibrium between F-actin and the critical concentration of actin (G-actin). Densitometric measurements showed that the unpolymerized actin ranged between 0.08 to 0.1 µM (as was shown in the first eight lanes where actin was incubated with increasing amount of VBS1-activated vinculin). This is the critical concentration of actin (Pollard, Analytical Biochemistry, 1983) that remains unpolymerized and is in equilibrium with polymerized filaments.

Similarly, sedimentation of filamentous actin in panel G showed unpolymerized actin corresponding to the critical concentration (as was shown in the last two lanes of “Pellets” and “Supernatants” gels). Most of the actin (filaments) went to the pellet except the critical concentration that remained unpolymerized, therefore staying in the supernatant. Thus, the intrinsic property of actin to polymerize above its critical concentration was not affected by the presence of vinculin in the absence or in the presence of VBS1.

Since the other reviewers are positive about this manuscript and since these panels are not even mentioned in the manuscript, one way to reconcile the three reviewers would be to take those panels out.

We thank the reviewer for the suggestion. We have removed all biochemistry data in panels G to H.

Also please correct this issue and clarify the text as needed:

With regard to Figure 3—figure supplement 2, it is concerning that full length vinculin pellets in the presence of VBS1 but in the absence of actin (vinculin oligomerization, stability issues?), and that actin is present in the soluble fraction upon polymerization. This raises additional concerns on my end regarding use of purified vinculin for their assays.

We hope the explanation that actin can only polymerize above its critical concentration (an intrinsic property of actin defined by the ratio of its dissociation over association rate constants, k-/k+) and therefore will also be found in the supernatant is convincing. Moreover, VBS1-activated vinculin can be found in the pellet due to some oligomerization states as observed by Molony and Burridge, 1985.

Also, in Materials and methods, it is stated that the pelleting speed is 380,000, but in the figure legend, the pelleting speed is stated as 80,000.

We apologize for the typo mistake and the confusion it brought.

We used high-speed sedimentation assays that were all carried out at 380,000x g.

https://doi.org/10.7554/eLife.53990.sa2

Article and author information

Author details

  1. Rajaa Boujemaa-Paterski

    1. Department of Biochemistry, University of Zurich, Zurich, Switzerland
    2. Université Grenoble Alpes, Grenoble, France
    Contribution
    Conceptualization, Formal analysis, Validation, Investigation, Visualization, Writing - original draft, Writing - review and editing
    For correspondence
    r.boujemaa@bioc.uzh.ch
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0001-9645-387X
  2. Bruno Martins

    Department of Biochemistry, University of Zurich, Zurich, Switzerland
    Contribution
    Investigation, Methodology
    Competing interests
    No competing interests declared
  3. Matthias Eibauer

    Department of Biochemistry, University of Zurich, Zurich, Switzerland
    Contribution
    Formal analysis, Methodology, Writing - review and editing
    Competing interests
    No competing interests declared
  4. Charlie T Beales

    Department of Biochemistry, University of Zurich, Zurich, Switzerland
    Contribution
    Investigation, Writing - review and editing
    Competing interests
    No competing interests declared
  5. Benjamin Geiger

    Department of Immunology, Weizmann Institute of Science, Rehovot, Israel
    Contribution
    Conceptualization, Supervision, Writing - review and editing
    Competing interests
    No competing interests declared
  6. Ohad Medalia

    Department of Biochemistry, University of Zurich, Zurich, Switzerland
    Contribution
    Conceptualization, Supervision, Funding acquisition, Writing - original draft, Project administration, Writing - review and editing
    For correspondence
    omedalia@bioc.uzh.ch
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0003-0994-2937

Funding

Schweizerischer Nationalfonds zur Förderung der Wissenschaftlichen Forschung (31003A_179418)

  • Ohad Medalia

Maxi Foundation (LMAN)

  • Ohad Medalia

Israel Science Foundation

  • Benjamin Geiger

Klaus Tschira Foundation

  • Benjamin Geiger

The funders had no role in study design, data collection and interpretation, or the decision to submit the work for publication.

Acknowledgements

This study was supported by the Swiss National Foundation, grant no. 31003A_179418, and the Mäxi Foundation (to OM), the Israel Science Foundation (to BG) and the Klaus Tschira Foundation (to BG). The authors would like to thank the Center of Microscopy and Image Analysis of the University of Zürich, and Sascha Weidner from the Workshop of the Department of Biochemistry, the University of Zürich, for the design and production of the mask holder used for the micropatterning. The authors are grateful to Prof. Hans-Werner Fink, Prof. Jürg Osterwalder and Prof. Marta Gibert from the Physik-Institut of the University of Zürich for their help with the UVO cleaner. We thank Ed Egelman (University of Virginia) for fruitful discussions during the initial stages of the project. We thank Julien Berro (Yale University) for insightful discussions. BG is the incumbent of the Erwin Neter Professorial Chair in Cell and Tumor Biology. We thank Dr Agnieszka Kawska (IllusScientia) for the artwork in Figure 5.

Senior Editor

  1. Suzanne R Pfeffer, Stanford University School of Medicine, United States

Reviewing Editor

  1. Edward H Egelman, University of Virginia, United States

Reviewer

  1. Christoph Ballestrem, University of Manchester, United Kingdom

Publication history

  1. Received: November 26, 2019
  2. Accepted: November 12, 2020
  3. Accepted Manuscript published: November 13, 2020 (version 1)
  4. Version of Record published: November 23, 2020 (version 2)

Copyright

© 2020, Boujemaa-Paterski et al.

This article is distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use and redistribution provided that the original author and source are credited.

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