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Pre-complexation of talin and vinculin without tension is required for efficient nascent adhesion maturation

  1. Sangyoon J Han  Is a corresponding author
  2. Evgenia V Azarova
  3. Austin J Whitewood
  4. Alexia Bachir
  5. Edgar Guttierrez
  6. Alex Groisman
  7. Alan R Horwitz
  8. Benjamin T Goult
  9. Kevin M Dean  Is a corresponding author
  10. Gaudenz Danuser  Is a corresponding author
  1. Lyda Hill Department of Bioinformatics, University of Texas Southwestern Medical Center, United States
  2. Department of Biomedical Engineering, Michigan Technological University, United States
  3. Department of Cell Biology, University of Texas Southwestern Medical Center, United States
  4. School of Biosciences, University of Kent, United Kingdom
  5. Department of Cell Biology, University of Virginia, United States
  6. Department of Physics, University of California San Diego, United States
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Cite this article as: eLife 2021;10:e66151 doi: 10.7554/eLife.66151

Abstract

Talin and vinculin are mechanosensitive proteins that are recruited early to integrin-based nascent adhesions (NAs). In two epithelial cell systems with well-delineated NA formation, we find these molecules concurrently recruited to the subclass of NAs maturing to focal adhesions. After the initial recruitment under minimal load, vinculin accumulates in maturing NAs at a ~ fivefold higher rate than in non-maturing NAs, and is accompanied by a faster traction force increase. We identify the R8 domain in talin, which exposes a vinculin-binding-site (VBS) in the absence of load, as required for NA maturation. Disruption of R8 domain function reduces load-free vinculin binding to talin, and reduces the rate of additional vinculin recruitment. Taken together, these data show that the concurrent recruitment of talin and vinculin prior to mechanical engagement with integrins is essential for the traction-mediated unfolding of talin, exposure of additional VBSs, further recruitment of vinculin, and ultimately, NA maturation.

Introduction

Cell-matrix adhesions are macromolecular complexes that link the extracellular matrix (ECM), typically via integrin transmembrane receptors, to the actin cytoskeleton. Being both a force-transmitter and a force-sensor, cell-matrix adhesions are critical to cell morphogenesis and mechanosensation (Parsons et al., 2010; Discher et al., 2005). Indeed, in response to ECM alterations, adhesions undergo constant changes in morphology and motion that involve the recruitment and recycling of a large number of adhesion molecules. For example, nascent adhesions (NAs) emerge within the actin-dense lamellipodia and then slide in the direction opposite of the protrusion as a result of polymerization-driven flow of the actin network (Parsons et al., 2010). Many of these NAs, which are less than 0.5 µm long (and thus appear as diffraction or near-diffraction limited spots in fluorescence microscopy) are rapidly turned over and disassembled. A subset of NAs matures into longer focal complexes (FCs, >0.5 µm in length) and focal adhesions (FAs,>2 µm in length) at the lamellipodia-lamella interface (Parsons et al., 2010; Gardel et al., 2010). During this progression, NAs go through multiple decision processes regarding fate and morphology. Compared to the well-studied FAs, for which the interconnection between structure, signaling, and force transmission is largely understood (Kanchanawong et al., 2010; Plotnikov et al., 2012; Thievessen et al., 2013; Geiger et al., 2009; Chrzanowska-Wodnicka and Burridge, 1996; Han et al., 2012; Riveline et al., 2001; Balaban et al., 2001; Stricker et al., 2011), much less is known about the molecular and mechanical factors that determine NA assembly, turnover, and maturation. In part, this is because until recently, it has not been technically feasible to measure whether individual NAs transduce traction forces. By combining high refractive-index and mechanically tuned substrates that are compatible with total internal reflection microscopy (Gutierrez et al., 2011) and numerical methods for the computational reconstruction of cell-substrate traction at the single micron length-scale, we demonstrated that force transmission is essential for the stabilization and maturation of NAs (Han et al., 2015). However, it remains unknown which molecular factors determine whether a NA begins to bear forces and thus continues to assemble.

One possible factor that may influence NA maturation is the stoichiometry of the earliest molecular components recruited to the site of its formation (Zaidel-Bar et al., 2004; Digman et al., 2009). In particular, the recruitment of talin, vinculin, and paxillin could play a critical role as they all are known to be mechanosensitive (Austen et al., 2015; Humphrey et al., 2014; del Rio et al., 2009; Kumar et al., 2016; Carisey et al., 2013; Humphries et al., 2007; Schiller et al., 2011; Pasapera et al., 2010). Talin is an integrin activator (Moser et al., 2009; Tadokoro et al., 2003) that directly links integrins to the actin cytoskeleton (Calderwood et al., 2013). Under force, the helix bundle domains in talin’s rod-like region unfold (del Rio et al., 2009), which both disrupts ligand binding and exposes cryptic-binding sites for vinculin and other proteins (del Rio et al., 2009; Yao et al., 2016; Yan et al., 2015; Goult et al., 2013; Goult et al., 2018). Vinculin, when bound to talin’s exposed binding sites, can indirectly strengthen the connection between actin and integrins by (1) forming a catch bond with F-actin (Case et al., 2015; Huang et al., 2017), (2) establishing multivalent linkages between talin and F-actin (Yao et al., 2016; Yan et al., 2015; Atherton et al., 2015), and (3) stabilizing talin’s unfolded state (Yao et al., 2014). In this scenario, talin first binds to integrins and F-actin, is unfolded under the initial load, and serves as a scaffold for the recruitment of vinculin. Indeed, at the level of FAs, direct evidence for catch-bonds (Bell, 1978; Thomas, 2008; Thomas et al., 2008), and the exposure of hidden binding sites under load (Vogel and Sheetz, 2006; Zhu et al., 2008), has established the idea of force-assisted adhesion growth. Further evidence for this model indicates that downregulation of actomyosin contractility reduces the recruitment of vinculin (Pasapera et al., 2010) and other adhesion proteins (Kuo et al., 2011), as well as the association between talin and integrins (Bachir et al., 2014).

In contrast to a model of hierarchical FA growth and stabilization where talin arrives first and recruits vinculin, fluorescence fluctuation analyses (Bachir et al., 2014) and co-immunoprecipitation experiments (Pasapera et al., 2010) have suggested that talin and vinculin might form a complex before talin associates with integrins. While talin-vinculin pre-association implies vinculin’s force-independent binding to talin, it is not clear whether this pre-association is required for NA assembly, and if so, whether pre-association affects the decision processes for NA maturation. Moreover, paxillin, a scaffolding protein that works in close relationship with focal adhesion kinase (FAK) (Pasapera et al., 2010; Parsons, 2003; Schlaepfer and Mitra, 2004; Mitra and Schlaepfer, 2006), is thought to be recruited and stabilized by force at an early phase of NA assembly (Plotnikov et al., 2012; Schiller et al., 2011; Choi et al., 2008; Deakin and Turner, 2008). However, vinculin’s recruitment and role in establishing tension across NAs remains poorly understood (Laukaitis et al., 2001; Webb et al., 2004; Wiseman et al., 2004).

Here, we investigated the integration of molecular recruitment and mechanical forces in determining the fate of NAs. Specifically, we combined high-resolution traction force microscopy with highly sensitive particle detection and tracking software to evaluate the time courses of force transmission and the molecular composition of individual NAs. A comprehensive inventory of these traces revealed broad heterogeneity in NA behavior. Thus, we applied machine-learning approaches to divide NAs into subgroups with distinct characteristics and identified that five subgroups are necessary to account for the different kinematic, kinetic, and mechanical properties of NAs. By focusing on the NA subgroup that matures into stable FAs, we found that the pre-complexation of talin and vinculin under force-free conditions was mediated by talin’s R8 domain. These complexes strengthen the link between talin and F-actin, aid the unfolding of talin upon initial weak force transmission in order to expose additional vinculin-binding sites, and ultimately support the transition of spontaneously formed nascent adhesions into stable focal adhesions.

Results

Nine adhesion classes can be distinguished based on different kinetic and kinematic behaviors

To investigate the time courses of traction forces and protein recruitment in NAs, we performed two-channel time-lapse, total internal reflection fluorescence (TIRF) imaging of Chinese Hamster Ovary epithelial cells (ChoK1, Figure 1—figure supplement 1a). We chose these cells because they form a stable footprint in close apposition to a flat substrate, which is ideal for TIRF microscopy of individual adhesions (Digman et al., 2009; Bachir et al., 2014; Bate et al., 2012; Macdonald et al., 2008). In addition, these cells display a relatively slow progression of adhesion assembly, offering the opportunity to track the fate of NAs and to identify the requirements for their maturation (Choi et al., 2008; Laukaitis et al., 2001; Webb et al., 2004; Choi et al., 2011). ChoK1 cells were transiently transfected with low-levels of either talin-GFP, vinculin-GFP, or paxillin-GFP, and plated and allowed to spread and migrate on high-refractive index silicone gels. To measure traction forces, deformable silicone gels were densely coated with 40 nm fluorescent beads (1.54 ± 0.22 beads/μm, 0.42 ± 0.17 μm bead-to-bead spacing, Figure 1—figure supplement 1b,c). After each experiment, cells were removed from the substrate and the beads were imaged in the relaxed, undeformed configuration of the silicone gel, which permits the quantitative reconstruction of traction forces at sub-micron scales (Figure 1—figure supplement 1e,g,h; Han et al., 2015). As expected, all traction force vectors pointed from the cell periphery to the cell center, independent of which adhesion protein was imaged (Figure 1a–c). Regardless of which protein was ectopically expressed, ChoK1 cells spread and generated traction forces indistinguishably (Figure 1—figure supplement 2).

Figure 1 with 7 supplements see all
Experimental/computational pipeline to analyze heterogeneous adhesion dynamics in ChoK1 cells.

(a–c) High-resolution traction maps co-imaged with mGFP-tagged adhesion protein, talin (d), vinculin (e), and paxillin (f). 5 kPa silicone gel coated with high density beads was used as a TFM substrate. (g) Trajectories of individual nascent and focal adhesions overlaid on a region of interest cropped from (e). Tracking is based on all detected point sources, (red circles). Big segmented focal contacts/adhesions (orange, closed freeform overlays) were used as additional information for feature selection. (h) Some of the key features used for supervised classification, tabulated in Supplementary file 1A. (i) Classification of adhesion trajectories into nine different groups, overlaid on the adhesion image. Five different NA groups, three FA groups and one noise group were distinguished by the support vector machine classifier. (j–m) Comparison of feature values among the five NA groups, G1, G2, G3, G4, and G5: edge protrusion speed (j), adhesion movement speed, positive when sliding toward protruding edge (k), mean intensity (l), and lifetime (m), extracted from six vinculin-tagged cells. All features show a significant shift in value for at least one subgroup. (n) Average traction magnitude, read from traction map, at individual NA trajectories per each group. The number of samples per each group is shown in the lower right corner of the figure.

Fluorescently tagged adhesion proteins (Figure 1d–f) were detected and tracked, and their intensity time courses extracted from their trajectories (Figure 1g, Figure 1—figure supplement 1f). To account for the heterogeneity of adhesions, we collected 22 features from each trajectory (Figure 1h, Supplementary file 1A) and classified the adhesions into nine groups (see Supplementary file 1B for a summary of each group) with a supervised machine learning pipeline (see Methods, Software Availability). To generate training data, a human operator used a dedicated graphical user interface for labeling ~120 adhesion tracks (~10 tracks per group, out of ~10,000 tracks per movie). Based on these data we trained a support vector machine (SVM) classifier (validation accuracy: 70–80%, Figure 1—figure supplement 3a). All features were inspected for redundancy and similarity (Figure 1—figure supplement 3b–c), and each group was distinct in terms of its Euclidean distance to the closest group in feature space (Figure 1—figure supplement 3d). SVM-based classification of trajectories that were excluded from the training data assigned each adhesion to one of nine different classes, G1, G2, …, G9 (Figure 1i). Five of the nine classes (G1-G5) identified NAs, three (G6-G8) identified FAs, and one group (G9) contained insignificant, noise-like trajectories (Video 1). The five NA classes significantly differed in terms of features such as ‘edge protrusion speed’ (Figure 1j), ‘adhesion movement speed’ (Figure 1k), ‘average fluorescence intensity’ (Figure 1l), and ‘lifetime’ (Figure 1m). For example, NAs classified as G3 form at the tip of the protruding edge and move forward with the protrusion. Of all NA classes, their fluorescence intensity is the lowest (Figure 1l). NAs classified as G2 form at the protruding edge but slide rearward relative to the substrate, mature to form larger FCs or FAs, and have the highest intensity and longest lifetime (Figure 1l–m). Indeed, all G2 NAs (~100%) mature to FCs, and ~32% mature into FAs (Figure 1—figure supplement 4). NAs classified as G1 also form at the protruding edge, but they are relatively stationary (Figure 1k) and are characterized by a weak fluorescence intensity and a short lifetime (Figure 1l–m).

Video 1
Time-lapse images of GFP-tagged vinculin in a ChoK1 cell, overlaid with adhesion trajectories and classification state from the support vector machine (SVM)-based machine-learning.

Different colors represent different classes: G1 (light green), G2 (dark blue), G3 (red), G4 (light blue), G5 (yellow), G6 (cyan), G7 (pink), G8 (dark green), and G9 (brown). The time interval per frame: 2 s. Playing speed: 25 frames/s. Duration of the movie: 12 min. Scale bar: 5 μm.

Nine adhesion classes exhibit distinct mechanical behaviors

Next, we tested the hypothesis that these spatially and kinetically distinct classes of NAs generated differential traction forces. Indeed, we found that the subgroup of maturing NAs, G2, showed the highest traction magnitude shortly after their initial assembly (Figure 1n). This is consistent with previous findings that demonstrated the tension-mediated and myosin-dependent maturation of FAs (Schiller et al., 2011; Choi et al., 2008; Schiller et al., 2013). Interestingly, NAs in G3 exhibited an insignificant amount of traction that did not increase throughout their lifetime (Figure 1n, Figure 1—figure supplement 5), suggesting that this population of anterogradely moving NAs might not interact with the retrogradely flowing F-actin cytoskeleton. NAs in G1 had higher traction than those in G3, implicating that short-lived, non-maturing NAs can transmit significant amounts of traction forces, which is consistent with our previous findings (Han et al., 2015). Importantly, these trends were consistent regardless of which adhesion protein was used for tracking (talin, vinculin, or paxillin; Figure 1—figure supplement 6). Furthermore, we observed shifts in the relative proportion of adhesion classes when cells were cultured on stiffer substrates (Figure 1—figure supplement 7). For example, large FAs (G8) were more abundant in ChoK1 cells cultured on 18 kPa substrates when compared to ChoK1 cells on 5 kPa substrates, which is a stereotypical output for stiffness sensing across adhesions (Han et al., 2012; Elosegui-Artola et al., 2016). Likewise, many populations of NAs (i.e. those in G1, G2, and G4) decreased for cells on 18 kPa relative to their 5 kPa counterparts. Altogether, these results confirm the reliability of the SVM classifier and suggest that kinetically unique NAs also show differences in terms of force transduction.

Talin, vinculin, and paxillin are recruited sequentially in non-maturing NAs, but concurrently in maturing NAs with traction development

To evaluate the relationship between molecular recruitment and traction force in NAs, we performed high-resolution traction force microscopy on cells labeled with either talin, vinculin, or paxillin, and processed these data using the aforementioned SVM classifier (Videos 19). We focused our analysis on the differences between NAs in G1 and G2 (Figure 2), which are for simplicity henceforth referred to as non-maturing and maturing NAs, respectively. For all proteins evaluated, fluorescence intensity traces for non-maturing NAs had a lifetime of ~6–7 min, with distinct phases of rise and fall (Figure 2a–f, top). The associated traction traces exhibited intermittent rises and falls, but with an overall magnitude that was much smaller than the traction traces in maturing NAs (Figure 2a–f, bottom). As expected, maturing NAs showed a steady increase in both fluorescence intensity and traction, and had a lifetime greater than 15 min (Figure 2g–l). The fluorescence intensity and traction of individual non-maturing and maturing NAs reflected this stereotypical behavior, and so did the average behavior, that is a slight increase and fall for non-maturing NAs, and more steady increase for maturing NAs (Figure 2—figure supplement 1a–i). A further analysis with cohort plots, where traces with similar lifetimes are grouped and displayed separately, revealed that average traces of many cohorts follow the stereotypical behavior (Figure 2—figure supplement 1j–o).

Figure 2 with 2 supplements see all
Talin and vinculin in non-maturing NAs are recruited in a sequential manner before traction development whereas in maturing NAs they are recruited concurrently, along with paxillin, briefly before the initial traction rise.

(a–l) Representative traces of protein recruitment and traction generation at non-maturing (a,b,c), or maturing (g,h,i) NAs. Each panel contains three views at different time points of mGFP-tagged talin (a,g, top), vinculin (b,h, top), and paxillin (c,i, top) and associated traction maps (bottom). Yellow boxes indicate positions of example adhesions whose fluorescent signals and traction levels are shown in time lapse montages with finer resolution underneath. Scale bar: 1 μm. Green circle represents the time point of initial talin/vinculin/paxillin signal rise, yellow circles show the time point of initial traction rise, and white circles shows the time of the peak amplitude, while red circles show regular detections in between these events. Scale bar: 1 µm. (d–l) Traces of fluorescence intensity amplitude (top) and traction (bottom). Blue and red segments indicate periods of significant fluorescence intensity amplitude and of traction, respectively, illustrated as ‘detected signal’ in (m). The black segments indicate the remaining background-subtracted fluorescence intensity and traction levels outside the detected signal period, that is, pre-signal and post-signal illustrated in (m). Whereas colored segments are read at positions moving with the adhesion center, pre- and post-signal traces are read at the first and last position of detected signal. An inset in (l) indicates that also in this trace the traction is gradually increasing. Blue and red arrows with the numbers on top mark the time points in seconds of the first intensity increase (FII) and the first traction increase (FTI), respectively, which are defined in (m). (m–o) Analysis of time-shifts between protein recruitment and FTI. (m) Traces of fluorescence intensity (top) and traction (bottom). Illustrated is the detection of the first significant value in both series. The gray signal represents the local background around the detected NA. Distinct distributions of time lags between FII and FTI in non-maturing (n) and maturing (o) NAs. Sample numbers, extracted from six cells for talin, five cells for vinculin and four cells for paxillin, are shown with each y-axis label. *p<1×10−2, ****p<1×10−30 by Mann-Whitney U test.

Video 2
Time-lapse images of GFP-tagged talin in a ChoK1 cell.

The time interval per frame: 1.644 s. Playing speed: 25 frames/s. Duration of the movie: 8 min 13 s. Scale bar: 5 μm.

Video 3
Time-lapse images of traction force magnitude for a ChoK1 cell expressing GFP-talin.

Traction forces are reconstructed from the bead images using high-resolution, L1-regularized, TFM software (Han et al., 2015). The color scale is the same as one at Figure 1a, that is, 0 (blue) – 300 Pa (red). The time interval per frame: 1.644 s. Playing speed: 25 frames/s. Duration of the movie: 8 min 13 s. Scale bar: 5 μm.

Video 4
Time-lapse images of GFP-tagged talin in a ChoK1 cell, overlaid with adhesion trajectories and classification state from the support vector machine (SVM)-based machine-learning.

The color coding is the same as in the legend of Video 1. The time interval per frame: 1.644 s. Playing speed: 25 frames/s. Duration of the movie: 8 min 13 s. Scale bar: 5 μm.

Video 5
Time-lapse images of GFP-tagged vinculin in a ChoK1 cell.

The time interval per frame: 2 s. Playing speed: 25 frames/s. Duration of the movie: 12 min. Scale bar: 5 μm.

Video 6
Time-lapse images of traction force magnitude for a ChoK1 cell expressing GFP-vinculin.

Traction forces are reconstructed from the bead images using high-resolution, L1-regularized, TFM software (Han et al., 2015). The color scale is the same as one at Figure 1b, that is, 0 (blue) – 1800 Pa (red). The time interval per frame: 2 s. Playing speed: 25 frames/s. Duration of the movie: 12 min. Scale bar: 5 μm.

Video 7
Time-lapse images of GFP-tagged paxillin in a ChoK1 cell.

The time interval per frame: 1.644 s. Playing speed: 25 frames/s. Duration of the movie: 8 min 13 s. Scale bar: 5 μm.

Video 8
Time-lapse images of traction force magnitudes generated by a ChoK1 cell expressing GFP-paxillin.

Traction forces are reconstructed from the bead images using high-resolution, L1-regularized, TFM software (Han et al., 2015). The color scale is the same as one at Figure 1c, that is, 0 (blue) – 1000 Pa (red). The time interval per frame: 1.644 s. Playing speed: 25 frames/s. Duration of the movie: 8 min 13 s. Scale bar: 5 μm.

Video 9
Time-lapse images of GFP-tagged paxillin in a ChoK1 cell, overlaid with adhesion trajectories and classification state from the support vector machine (SVM)-based machine-learning.

The color coding is the same as in the legend of Video 1. The time interval per frame: 1.644 s. Playing speed: 25 frames/s. Duration of the movie: 8 min 13 s. Scale bar: 5 μm.

Next, we developed an event-based time-series analysis method that identifies the first time point of significant fluorescence and force increase, respectively, and then measures the time delay between the two (Figure 2m). The blue and red arrows in Figure 2d–f,j–l show, in two example traces, the time points identified statistically as the initial time point of intensity and traction increase, respectively. Using this approach, we first determined the fraction of NAs per group with a significant traction increase at any point throughout their lifetime (Figure 2—figure supplement 2). Interestingly, both non-maturing and maturing NAs showed such force increases, that is, they were engaging and clutching against F-actin’s retrograde flow at one point with the substrate. In contrast, NAs classified as groups G3-5 exhibited lower fractions of force increase, suggesting that a very large number of adhesion protein complexes, detectable through either talin, vinculin, or paxillin recruitment, do not transduce measurable traction forces (e.g. less than 20 Pa). Indeed, these complexes could result from an incomplete molecular clutch and by being engaged with the substrate but not F-actin, or vice versa.

To evaluate if maturing and non-maturing NAs were differentially assembled, we next analyzed protein recruitment using the initial traction force increase as a time fiduciary. In non-maturing NAs, talin, vinculin and paxillin were recruited ~18 s, ~8 s and ~4 s before the onset of force transmission, respectively (Figure 2n). In contrast, in maturing NAs, talin, vinculin and paxillin were recruited concurrently ~4 s before the onset of force transmission (Figure 2o). We also noted that the temporal distribution of talin recruitment was significantly wider in non-maturing NAs than in maturing NAs. These findings suggest that in maturing NAs talin localizes with vinculin prior to its association with integrins, which more efficiently leads to force transmission and maturation to a FC. These data also suggest that NAs that sense force at the earliest stages of assembly are more likely to successfully mature. In contrast, although non-maturing NAs also support some lower level of force transmission eventually (Figure 1n), talin and vinculin assemble sequentially and remain under force-free conditions for a much longer duration of time.

In maturing NAs, vinculin assembles faster than in non-maturing NAs, but talin and paxillin show no difference

The rod domain of talin contains 13 helical bundles, 9 of which include cryptic vinculin binding sites (VBSs) that are exposed upon tension-mediated unfolding (Geiger et al., 2009; del Rio et al., 2009; Goult et al., 2013). As such, we hypothesized that the simultaneous recruitment of talin and vinculin in maturing NAs could further accelerate vinculin binding compared to non-maturing NAs. To test this, we quantified the assembly rate of each protein by obtaining the slope of the fluorescence intensity over the first 10 s after initial appearance (Figure 3a). Interestingly, only vinculin showed a significant difference in the assembly rate between non-maturing vs. maturing vinculin complexes, while talin and paxillin showed no such differences (Figure 3a). Thus, the concurrent arrival time of talin and vinculin in maturing NAs could prime talin to expose additional VBS domains, which in turn would further reinforce vinculin recruitment and adhesion maturation. We also quantified the traction force growth in those NAs with an expectation that there would be an immediate rise in force with faster vinculin binding. However, the traction force growth rate for the first 10 s showed no significant difference between non-maturing and maturing NAs (Figure 3b). This observation could arise because the forces were below our limits of detection for traction, or because traction does not develop during the earliest stage of molecular recruitment. N, a difference was observed when the force growth rate was quantified over a longer period, that is, 2 min (Figure 3c), which is consistent with our previous observations (Han et al., 2015). These findings imply that increased vinculin recruitment in maturing NAs supports the rise in traction force by increasing binding to F-actin, albeit with some time delay.

Vinculin, but not talin and paxillin, is recruited significantly faster in maturing NAs than in non-maturing NAs.

(a) Assembly rate of talin, vinculin, and paxillin, to G1 (non-maturing) or to G2 (maturing) NAs, quantified by the slope of fluorescence intensity over the initial 10 s after detection. (b) Traction growth rate at the NAs in (a) for the initial 10 s after detection. (c) Traction growth rate quantified over the first 2 min after detection. *p<1×10−2, ***p<1×10−10, ****p<1×10−30 by Mann-Whitney U test.

Vinculin can bind to talin without force through a ‘threonine belt’ in talin R8 domain

Previous work established that under tension the R3 domain in talin unfolds first, as it contains a destabilized hydrophobic core due to the presence of a ‘threonine belt’. By mutating the threonine residues to isoleucines and valines (the so called ‘IVVI mutant’) it was possible to stabilize the core and prevent talin from unfolding, which significantly reduces the exposure of the two cryptic VBS (Goult et al., 2013; Yao et al., 2014; Elosegui-Artola et al., 2016). Moreover, we showed that the VBS in R8 was able to bind vinculin readily in the absence of force (Gingras et al., 2010). Like R3, R8 also contains a threonine belt, consisting of T1502, T1542, and T1562 (Figure 4a). Thus, we hypothesized that a similar mutational strategy, using a T1502V, T1542V, T1562V ‘R8vvv mutant’, could stabilize the R8 domain and reduce access to the VBS. To test this hypothesis, we made a 'R7R8vvv' construct and compared its unfolding characteristics to the wild-type (WT) R7R8 fragment (R7R8wt) with circular dichroism (CD; Figure 4b). We included the R7 domain to improve the stability of the fragment and maintain R8 in its native conformation. In the R7R8wt the two domains unfolded cooperatively with a single unfolding step at a melting temperature (Tm) of 55°C. In contrast, stabilization of the R8 domain in the R7R8vvv mutant resulted in the domains unfolding independently, with R7 unfolding at a similar temperature to the WT (Tm56°C), but the melting temperature of the R8 domain increased from 56°C to 82°C. Strikingly, the two unfolding steps indicate that in the R7R8vvv mutant the R7 and R8 behave independently with regard to thermal stability. Together, these results show that the R7R8vvv mutant stabilizes R8.

Figure 4 with 1 supplement see all
Stabilizing the ‘threonine belt’ in the R8 domain of talin inhibits talin-vinculin interactions under tension-free conditions.

(a) Cartoon representation of talin R7R8 (pdb id 2X0C) showing the ‘threonine belt’, comprised of residues T1502, T1542, and T1562, labeled and shown as sticks (cyan), the VBS helix is colored red. (a.i) side on view (N.B. helix 31 transparent), (a.ii) top down view. (b) Denaturation profiles for WT R7R8wt (red) and R7R8vvv (black) measured by monitoring the change in circular dichroism at 208 nm with increasing temperature. R7R8wt has a melting temperature of 55°C, whereas R7R8vvv unfolds in two steps, one (R7) with a melting temperature of 56°C and R8 unfolding at 82°C. (c) Chromatograms showing binding of talin R7R8 to the vinculin head (Vd1). R7R8wt (red) and R7R8vvv (black) binding to Vd1. Complex peaks and unbound peaks are indicated.

To test whether stabilization of R8 would affect its interaction with vinculin, we used analytical gel filtration to look at protein complex formation. Preincubation of R7R8wt or R7R8vvv with vinculin Vd1 showed that both constructs were able to form complexes. However, the R7R8vvv:Vd1 complex peak was substantially smaller than the WT:Vd1 peak, confirming that the R7R8vvv bound less Vd1 (Figure 4c). Thus, by stabilizing the threonine belt with the R8 valine mutations, vinculin’s accessibility to the cryptic VBS was reduced. Interestingly, the R8 domain is also a Leucine-Aspartic acid (LD) motif binding site, that is it binds to multiple proteins that act downstream of adhesion, including RIAM and DLC1 (Goult et al., 2013; Goult et al., 2018; Zacharchenko et al., 2016). Using a fluorescence polarization assay described previously (Whitewood et al., 2018), we measured the binding affinities of R8 ligands RIAM TBS1 and DLC1 peptides with R7R8wt and R7R8vvv (Figure 4—figure supplement 1). Both peptides showed comparable binding affinities toward WT R7R8 and the R8R7vvv mutant, confirming that R7R8vvv was still able to bind LD-motif proteins. Altogether, these biochemical characterizations suggest that the threonine belt in talin R8 exposes the R8 cryptic VBS and binds vinculin under force-free conditions. The mutant also provided a tool for us to probe the functional implications of talin-vinculin force-free complex formation on NA assembly and maturation in vivo without interfering with binding of other binding partners.

Cells with R8vvv mutant talin show less maturing NAs and sparser and smaller FAs

To investigate whether talin’s ability to form a complex with vinculin under force free conditions promotes adhesion maturation, we introduced the R8vvv mutation into full-length talin1 and tagged it with mNeonGreen (‘talin1 R8vvv-mNG’). For imaging, we knocked down endogenous talin1 expression with an shRNA, and rescued ChoK1 cells with shRNA-resistant forms of WT or R8vvv talin. The expression of talin1 R8vvv-mNG was slightly less than that of endogenous talin1 in WT ChoK1 cells, but more than the remaining endogenous talin1 expression in knockdown cells (Figure 5—figure supplement 1). Interestingly, cells expressing the talin1 R8vvv-mNG mutant contained many more NAs (Figure 5a,b,f,g,k) and less and smaller FCs and FAs (Figure 5a,b,f,g,l,m) than control cells expressing WT talin1. Furthermore, cells expressing the R8vvv variant of talin1 also showed less traction than cells rescued with WT talin1 (Figure 5c,h,n). Consequently, with less traction and more NAs, the edge protrusion and retraction velocity was also faster in cells expressing the talin1 R8vvv-mNG mutant (Figure 5d,i). Moreover, a lower fraction of NAs and FCs in R8vvv mutant cells grew in size to FAs than NAs in cells with WT talin1 rescue (Figure 5e,j,o,p). Together, these results demonstrate that talin R8vvv mutation restricts NAs from maturing into focal adhesions.

Figure 5 with 4 supplements see all
Expression of the talin1 R8vvv mutant in ChoK1 cells with endogenous talin1 knocked down results in the formation of denser NAs, but lesser and smaller FAs, lower traction, more active protrusions, and less maturing adhesions compared to WT talin.

(a–j) Adhesion, traction, and protrusion phenotype of a representative ChoK1 cell on 5 kPa substrate expressing (a–e) talin R8vvv mutant or (f–j) WT talin. (a and f) inverted talin-mNeonGreen images. (b and g) detection of NAs, FCs and FAs. (c and h) traction force maps. (d and i) snapshots of computer vision-extracted cell boundaries at 0 and 10 min of a movie. (e and j) overlay of NAs and FCs that mature to FAs. (k–p) Box plots of (k) NA density, (l) FA density, (m) FA area, (n) total traction integrated over cell area, (o) fraction of NAs maturing to FAs relative to all NAs, (p) and of the fraction of FCs maturing to FAs (relative to all FCs). Number of adhesions imaged are listed under each box plot. Number of independently imaged cells for talin1 R8vvv-mNG and WT talin1-mNG rescue were 5 and 7, respectively. Scale bar: 10 µm. ****: p<1×10-30 by Mann-Whitney U test.

Despite the reduced NA maturation, ChoK1 cells expressing the talin1 R8vvv mutant exhibited some large FAs (Figure 5a,b and m). Accordingly, we asked if these FAs could reflect a population of NAs that mature under recruitment of residual endogenous talin1 (Figure 5—figure supplement 1), or perhaps talin2, which we did not specifically knockdown. Western Blot analysis showed that talin2 expression was minimal in ChoK1 cells compared to talin1 and knocking down talin1 did not result in compensatory expression of talin2 (Figure 5—figure supplement 2). The traction magnitude of ChoK1 talin1 knockdown cells was significantly lower than of WT cells ectopically expressing talin1 or knockdown cells rescued with talin1 (Figure 5—figure supplement 3). Together, these data suggest that influence of a residual talin1 pool or of compensatory expression of talin2 on the behavior of ChoK1 cells with a talin1 knockdown and rescue by either R8vvv- or WT talin1 is minimal.

To ensure that the observed R8vvv NA maturation defect is independent of endogenous talin1 and talin2 expression, we lentivirally introduced the WT and the R8vvv variants of talin1 into talin1/2 null inner medullary collecting duct (IMCD) cells (see Materials and methods) (Mathew et al., 2017). Demonstrating the importance of talin for cell adhesion, talin1/2-null IMCD cells grew in suspension, and only adhered to the substrate when rescued with talin1 constructs. IMCD cells rescued with R8vvv-talin1 showed an adhesion phenotype that is very similar to ChoK1 rescued with R8vvv-talin1. Specifically, cells rescued with talin1 R8vvv had more NAs, smaller FCs and FAs, less traction, less NAs and FCs maturing to FAs than IMCD cells rescued with WT talin1 (Figure 5—figure supplement 4). Thus, we concluded that the remaining large FAs likely result from talin1-vinculin complexes, which still form in the presence of the R8vvv mutation, albeit to a lesser extent (see Figure 4C).

R8vvv mutation promotes talin recruitment overall but impedes traction growth rate

To investigate whether force-free complex formation of talin1 with vinculin affects talin recruitment itself, we compared the time of talin1 recruitment in R8vvv and WT talin1 rescue cells for non-maturing (G1) and maturing (G2) NAs. Consistent with the data in Figure 2, in both cases non-maturing NAs showed talin recruitment, on average, ~18–14 s prior to the initial rise in traction (Figure 6a,c,e,g,i), while maturing NAs showed a near-immediate, that is, with ~6 s before traction, talin recruitment (Figure 6b,d,f,h,i). This indicates that the ability of talin to bind vinculin in the absence of the force does not affect talin recruitment. For both WT and R8vvv mutant talin, the assembly rates were statistically indistinguishable between non-maturing and maturing NAs (Figure 6j). The rate of traction development in NAs, however, was significantly affected in talin1 R8vvv-mNG mutant cells. Overall, the traction growth rate was reduced in R8vvv cells, both for non-maturing and maturing NAs (Figure 6k). Moreover, whereas maturing NAs in WT talin1 rescue cells showed faster traction growth than non-maturing NAs, consistent with the data in Figure 3c, maturing adhesions in talin1 R8vvv-mNG mutant cells exhibited a force growth similar to that of non-maturing NAs (Figure 6k). Interestingly, the talin1 assembly rate of both NA types in R8vvv expressing cells was higher than WT talin1 rescue (Figure 6j). Inspection of individual NA trajectories (one example in Figure 6—figure supplement 1a,b) showed that R8vvv-talin is recruited fast but also disassembles fast. We speculate that this fast assembly relates to the high NA density shown in R8vvv-expressing cells (Figure 5k). These transient R8vvv-talin NAs also showed insignificant vinculin association (Figure 6—figure supplement 1c,d). These results suggest that the R8vvv mutation elevates talin’s own recruitment rate in both maturing and non-maturing NAs, but interferes with force transduction in maturing NAs.

Figure 6 with 1 supplement see all
Expression of talin1 R8vvv-mNG mutant does not change the recruitment timing of talin to NAs, but reduces the force growth rate in NAs.

(a–h) Representative talin (top) and traction force (bottom) images of talin1 R8vvv-mNG expressing cells (a–d) and WT talin-mNG rescue cells (e–h) within non-maturing (a,c,e,g) and maturing (b,d,f,h) NAs. (a,b,e,f) talin-mNG images (top) and traction images (bottom) of three different time points, that is at initial nucleation, at maximum fluorescence intensity, and at the end of the NA portion of the track. Yellow boxes indicate positions of example adhesions whose fluorescent signals and traction levels are shown in time lapse montages with finer resolution underneath. Green circles indicate the time points of initial talin signal rise, yellow circles show the time points of initial traction rise, and white circles show the time of the peak amplitude. Red circles show normal default detections without special events. Scale bar: 1 µm. (c–d, g–h) Traces of talin-mNeonGreen fluorescence intensity (top) and traction (bottom). Phases of the traces with significant fluorescence above background are indicated in blue and red, respectively. The black time series outside the colored signal are the background-subtracted intensities read at the first or last position detected by the particle tracker. Blue and red arrows mark the time points of the first intensity increase and the first traction increase, respectively (i–k) Distributions of time lags of fluorescence intensity onset relative to traction onset (i), talin assembly rates (j), and traction growth rates (k) of non-maturing (G1) and maturing (G2) NAs in talin1 R8vvv-mNG mutant and WT talin1-mNG rescue cells. Time integration time for calculating slopes in j and k was 20 and 60 s, respectively. *p<1×10−2, **p<1×10−3, ***p<1×10−10, ****p<1×10−30 by Mann-Whitney U test.

Differential vinculin recruitment between non-maturing vs. maturing NAs vanishes with talin1 R8vvv mutation

Vinculin recruitment to the NA is critical for both force growth and adhesion maturation (Figure 3; Thievessen et al., 2013). To examine whether the assembly rate of vinculin is affected by talin’s ability to form a complex with vinculin without initial tension, we performed two-channel imaging of vinculin-SnapTag-TMR-Star and WT- or R8vvv-talin-mNeonGreen (see Materials and methods). As performed previously, we captured and analyzed time-series of each pair of talin-vinculin signals in non-maturing vs. maturing NAs (Figure 7a–h) and quantified the vinculin assembly rate within 30 s after first detection (Figure 7i). In talin1 R8vvv-mNG mutant cells, vinculin assembly rates were statistically indistinguishable between non-maturing and maturing NAs, whereas in wild-type talin1-rescue cells vinculin rates were significantly higher in maturing NAs, consistent with the data acquired in control cells (Figure 3a). This result suggests that early vinculin binding to talin R8 domain indeed contributes to faster recruitment of additional vinculin. The insignificant difference in vinculin recruitment in R8vvv mutant cells for non-maturing vs. maturing NAs might be related to the reverted traction growth rates between the two NA groups observed in these mutant cells (Figure 6k). It is also worth noting that the vinculin signal in maturing NAs of cells with WT talin-rescue tended to keep increasing while talin intensity was relatively flat (t = 200 ~ 600 s in Figure 7h,d), suggesting that the number of exposed talin VBSs is increasing, and thus the number of bound vinculin proteins, over time under tension. The same trend was observed in talin1 R8vvv mutant cells (Figure 7b,f), but the vinculin recruitment rate was again much less than those found in WT talin1-rescue. Altogether, this data strongly suggests that vinculin recruitment is significantly reduced in the absence of tension-free vinculin-talin pre-complexation.

Vinculin recruitment is reduced in talin1 R8vvv mutant cells.

(a,d,g,j) Representative two-channel time-lapse images of talin-mNeonGreen (top) and vinculin-SnapTag-TMR-Star (bottom) of G1 NA in a Talin1 R8vvv mutant cell (a), G1 NA in WT talin1 rescue cell (d), G2 adhesion in a Talin1 R8vvv mutant cell (g), and G2 adhesion in WT talin1 rescue cell (j). NAs of interest are indicated with a yellow box. Scale bar: 1 μm. (b–k) Time series of talin-mNeonGreen amplitude (top) and vinculin-SnapTag-TMR-Star amplitude (bottom) of G1 non-maturing (b,e) and G2 maturing (h,k) NAs in cells expressing the talin1 R8vvv mutant (b,h) and WT talin (e,k) constructs. Colored time periods (blue for talin, magenta for vinculin) indicate the phases where the adhesion is detected as a significant particle of robust trackability. The black time series outside the colored signal are the background-subtracted intensities read at the first or last position detected by the particle tracker. Blue and magenta arrows and the text around them indicate the time of talin and vinculin recruitment onset, respectively. (c,f,k,l) Time lapse montages of individual NAs shown in a, d, g, and j, respectively, overlaid with colored circles as detected centers of NAs of interest. Green circle represents the time point of initial talin signal rise, yellow the time point of initial vinculin signal onset, white the time of the peak amplitude, while red circles show normal default detections without special events. Talin and vinculin’s initial recruitments are indicated with green and yellow arrows to highlight the time delay occurring between talin and vinculin in G1 adhesions and the concurrent recruitment in G2 adhesions, regardless of R8vvv mutations. (m) Vinculin assembly rates at non-maturing and maturing NAs in R8vvv mutant and WT talin rescue cells, quantified by the slope of vinculin-SnapTag-TMR-Star fluorescence intensity over the initial 20 s after the first detection in the talin-mNeonGreen channel. (n) Time delays of vinculin recruitment onset relative to talin recruitment onset of non-maturing vs. maturing NAs in talin1 R8vvv-mNG mutant and WT talin1 mNG cells. Vinculin recruitment onsets in non-maturing NAs are positive, that is, vinculin recruitment starts after talin. In contrast, vinculin recruitment onsets in maturing NAs are nearly coincidental with talin. See the text for further description. (o) Lifetimes of maturing NAs classified in talin1 R8vvv mutant and WT talin1 mNG rescue cells. ****p<1×10−15, **p<0.05 by Mann-Whitney U test. The numbers of adhesions (N), extracted from seven cells each for cells with talin1 R8vvv-mNG and WT talin1-mNG, are shown per each condition name at each panel.

Simultaneous talin-vinculin imaging confirms vinculin’s recruitment after talin for non-maturing NAs and concurrent recruitment for maturing NAs

To confirm the recruitment order of talin and vinculin with respect to traction force development (Figure 2), we quantified the time difference between the first significant increase in talin fluorescence intensity and the first significant increase in vinculin fluorescence intensity (blue and magenta arrows in Figure 7a–l,n). For non-maturing NAs, both in talin1 R8vvv mutant and WT talin1-rescue cells, vinculin was delayed to talin (Figure 7n), consistent with the delay we inferred indirectly based on alignment of the fluorescent intensity increases with the first significant traction force increase (Figure 2n). In maturing NAs, vinculin and talin recruitment coincided (Figure 7i,l,n), also consistent with the indirect inference presented in Figure 2o. This shows directly that tension-free talin-vinculin pre-complexation enhances the probability of NA maturation. In more detail, vinculin recruitment in non-maturing NAs of talin1 R8vvv mutant cells was on average ~17 s after talin recruitment, whereas vinculin recruitment in the wild-type talin rescue condition was preceded by the talin recruitment on average by ~14 s (Figure 7n). We interpret the small but significant difference between the two recruitment time distributions as the result of the mutation in talin’s R8 domain, which reduces the ability of vinculin to bind talin prior to mechanical unfolding. Moreover, even though some maturing NAs eventually grow also in talin1 R8vvv mutant cells, the absence of efficient vinculin binding to the VBS in R8 propagates into an overall less efficient vinculin recruitment. In agreement with this interpretation, we found that the lifetimes of maturing NAs in the mutant cells were much shorter than those in cells with WT talin1 rescue (Figure 7o). In summary, our data establishes that early association of talin with vinculin via the talin R8 domain is critical for accelerated vinculin binding, which in turn contributes to the development of the level of force transmission required for NA maturation.

Discussion

Our experiments show that the maturation of NAs depends upon the concurrent early recruitment of talin and vinculin without tension. Previous models have inferred that tension across talin, which can establish direct bridges between integrins and actin filaments, is sufficient to trigger a conformational change that exposes several vinculin binding sites. These binding sites were thought to promote the recruitment of vinculin to further strengthen the link between the integrin-talin complex and F-actin (Goult et al., 2018; Sun et al., 2016). However, these models were derived primarily from observations in focal adhesions, that is, at a late stage of the maturation process (Thievessen et al., 2013; Atherton et al., 2015). Here, we exploit our ability to simultaneously measure nanonewton-scale traction forces and molecular associations within individual NAs using total internal reflection microscopy on high-refractive index soft substrates (Gutierrez et al., 2011; Han et al., 2015). Our data suggests that the tension born by an individual talin bridge between an integrin and actin filaments is insufficient to maximize the number of VBSs available for reinforcing the link with F-actin. This further lowers the lifetimes of catch-bond-like molecular associations (Huang et al., 2017; Sun et al., 2016; Hákonardóttir et al., 2015) between talin and vinculin, vinculin and actin, and talin and actin, resulting in turnover of the NAs (Figure 8, top). In contrast, the concurrent recruitment of talin and vinculin, perhaps as a preexisting cytosolic complex, immediately establishes a strong link between integrins and F-actin, as indicated by the immediate onset of traction forces. The fast loading rate promotes an efficient unfolding of the talin rod domain, which exposes several additional VBSs for further recruitment of vinculin and strengthening of the talin/F-actin interaction. This results in robust increase of traction force transmission and stabilization of catch-bond-like molecular associations that contribute to the maturation of the NA (Figure 8, bottom).

A suggested mechanism of differential recruitment of talin and vinculin determining maturation of nascent adhesions.

(Top) For non-maturing NAs, talin binds to integrin before vinculin recruitment. Talin stretching might be limited to a shorter level, which limits the exposure of vinculin-binding-sites. Inefficient vinculin binding, in turn, limits the number of F-actin that can connect to the adhesion complex, allowing for only a low amount of tension across the complex. Insufficient loading level reduces the lifetime of catch-bond like associations between molecules, resulting in turnover of the NA complex. (Bottom) For maturing NAs, talin and vinculin interact before engagement with integrin. Upon concurrent recruitment to the NA traction force builds immediately. Talin might be stretched in a faster manner by pre-associated vinculin and talin’s own binding to F-actin accommodate faster, efficient recruitment of additional vinculin. High loading levels across the complex stabilizes molecular bonds, which facilitates the maturation of the NA. The sites for paxillin binding, for example, to vinculin or β-integrin via FAK, are inferred from the literature (Humphries et al., 2007; Turner et al., 1990; Hu et al., 2014; Lawson et al., 2012).

Our data also show that the colocalization of talin and vinculin is promoted by talin’s R8 domain, which contains a VBS that is exposed for vinculin recruitment without tension-mediated unfolding of talin. We generated a talin mutant with a more stable R8 domain that reduces the spontaneous association with vinculin. Cells expressing this mutant have a large fraction of NAs that cannot mature into FAs and transmit only low-level forces. Ultimately, our data suggests that maturing NAs are likely to be initiated by an R8-mediated talin-vinculin association. Intriguingly, complex formation prior to binding integrin receptors requires spontaneous encounters of mobile talin and vinculin at the plasma membrane or even within the cytosol. These are likely rare events, which may explain the surprising finding that the number of maturing (G2) NAs (3.5 ± 1.6%, Mean ± Standard Error of the Mean, N = 20 movies) is low compared to G1 NAs (28.8 ± 3.5%, Mean ± S.E.M., N = 20 movies) among all NAs.

Despite the general phenotype of less NA maturation, talin1-R8vvv-expressing cells also exhibited a small number of large FAs (Figure 5j). These FAs are not due to residual endogenous talin1 or talin2 expression, as they are also evident in talin1/2 double knockout cells. Instead, these FAs in R8vvv-expressing cells likely result from one or more mechanisms. For example, (1) talin-R8vvv binding to vinculin is not abolished, only reduced (Figure 4c), (2) talin-R8vvv’s association with integrins and F-actin could be quick enough to expose other VBSs, or perhaps (3) arise from one of the other NA classes (we focused largely on G1 and G2 here). Nonetheless, by performing three-color imaging of talin, vinculin, and traction forces (Figure 7), we show that R8vvv cells do have maturing NAs, but less of them, with reduced lifetimes, and importantly, with impaired vinculin association rates (Figure 7m). Thus, it appears that the second possibility might be less likely. Alternatively, we speculate that there may be a compensation effect at play where the fewer maturing NAs accumulate excess talin (Figure 6j).

Our findings regarding the talin R8 domain offer an alternative perspective on talin-vinculin association, including our own report describing talin’s R3 domain as the weakest region that can unfold under tension (Yao et al., 2016; Atherton et al., 2015). Likewise, there are recent results that seem conflicting with one another. Using truncated talin fragments, it would appear that the talin R4-R8 fragment does not bind the vinculin head domain, whereas the talin R1-R3 fragment does (Dedden et al., 2019). However, another recent study has shown that a talin lacking R2R3 is able to interact with both inactive and active vinculin (Atherton et al., 2020). Moreover, a few studies have supported an idea that R3 requires force, albeit small, for engagement with vinculin (Austen et al., 2015; Yao et al., 2016; Elosegui-Artola et al., 2016): First, a study with a talin tension sensor has shown that talin1’s engagement with the cytoskeleton must precede vinculin’s binding to the N-terminal VBS on talin1 (Austen et al., 2015). Second, IVVI mutant in talin R3 has been reported to prevent mechanotransduction, as assessed by traction exertion and YAP nuclear localization (Elosegui-Artola et al., 2016). Additionally, based on published data (Goult et al., 2013; Lee et al., 2013), the R3 domain is likely to be bound to the Rap1-interacting adaptor molecule (RIAM) prior to the force (Vigouroux et al., 2020). Thus, early, tension-independent interactions between talin and vinculin is more likely mediated by a different site, for which our data suggests the talin R8 domain.

Our finding of a role for R8-mediated talin-vinculin complex formation in the earliest stages of adhesion assembly is also somewhat unexpected in view of the paradigm that both full-length talin and vinculin reside largely in an auto-inhibited conformation that prevents mutual interaction. Indeed, a recent study showed that full-length talin1 in a closed conformation does not form a complex with vinculin regardless of vinculin’s conformation (Dedden et al., 2019). In contrast, another study reported that only the ‘activated’ conformation of vinculin can form a complex with talin, or vice versa (i.e. an activated talin can form a complex with vinculin in its closed form) (Atherton et al., 2020). We speculate that both talin and vinculin are highly dynamic proteins that exist in an equilibrium that transitions between closed, open, and intermediate states in living cells (Dedden et al., 2019), and the potential activation energy necessary for activation could easily arise from F-actin-driven forces. Additionally, although biochemically not yet confirmed, a talin-vinculin precomplex appears feasible as the structure of full-length talin shows an exposed R8 VBS. These considerations collectively align with the observation that the occurrence of maturing G2 adhesions is less than the occurrence of non-maturing G1 adhesions. Thus, the conditions for efficient maturation, including the formation of a talin-vinculin complex without tension, are rarely fulfilled. Nonetheless, a rare but significant number of talin-vinculin precomplexes are sufficiently available to nucleate a population of G2 adhesions, which ultimately drives adhesion maturation.

In the case of non-maturing NAs, talin and vinculin were recruited significantly before our measurements could detect a significant traction onset. This finding implies that there are sub-populations of interacting talin and vinculin that transduce little force. Additionally, talin is present for a longer time than vinculin before traction onset in non-maturing adhesions. While this is conceptually consistent with a previous finding that vinculin binding to talin requires talin’s actin binding for tension development (Austen et al., 2015), the time lag between talin and vinculin recruitment suggests that talin’s sole engagement with F-actin without vinculin potentially impedes talin’s own role as an integrin activator (Shattil et al., 2010) and promoter of integrin clustering (Saltel et al., 2009). Additionally, before vinculin binding, talin may be bound to RIAM (Lee et al., 2009), which is replaced by vinculin only after the R2R3 domain unfolds (Goult et al., 2013).

How a potential talin-vinculin precomplex promotes faster tension development and talin unfolding in maturing adhesions remains to be determined. Potential mechanisms imply that (1) the complex is also pre-bound to F-actin through the vinculin tail (as vinculin bound to talin is almost certainly in an open conformation) (Humphries et al., 2007; Golji and Mofrad, 2013) and (2) that the talin-vinculin interactions via talin’s R8-domain do not interfere with talin’s direct binding to F-actin, thus accelerating talin’s actin-binding rate.

In maturing adhesions, paxillin is recruited concurrently with talin and vinculin. Which molecular partners in NAs and binding sites within those molecules are responsible for paxillin’s concurrent binding needs further investigation. A previous study has shown that talin-vinculin interaction can facilitate efficient paxillin recruitment regardless of the paxillin-binding-site in vinculin’s tail domain (Humphries et al., 2007). This finding suggests that paxillin’s concurrent recruitment in maturing nascent adhesions is associated with the talin-vinculin precomplex but not necessarily via paxillin’s direct binding to vinculin (Carisey and Ballestrem, 2011).

How non-maturing (G1) NAs switch to disassembly also necessitates further investigation. One potential scenario includes that talin is competitively bound by RIAM or DLC1 before it can associate with vinculin. Alternatively, vinculin binding to other VBSs within talin could interfere with force transduction and adhesion maturation (e.g. vinculin binding to R3, after partial unfolding by F-actin-mediated forces at ABS3). Whether vinculin’s binding to such VBSs, that is, other than R8, interfere or synergize vinculin binding remains to be determined. However, our data suggests that when talin binds F-actin in G1 NAs – directly or indirectly via vinculin – the further recruitment of vinculin is much slower than that of G2 NAs where talin and vinculin arrive simultaneously in what we hypothesize is a precomplex.

Our data also indicates that the onset of traction force is accompanied by paxillin recruitment, regardless of the fate of the NA (Figure 2). This suggests that paxillin is recruited after vinculin, and this is particularly true in non-maturing NAs. Indeed, the tension dependency of paxillin recruitment is well-established (Schiller et al., 2011). Our data suggests that tension-dependent recruitment of paxillin is through vinculin, which is consistent with previous findings that suggest paxillin recruitment can be induced by vinculin (Humphries et al., 2007) as it binds to the tail-domain of vinculin (Turner et al., 1990). Alternatively, paxillin has been reported to be recruited after focal adhesion kinase (FAK) in endothelial cells (Hu et al., 2014). Given evidence that talin can also be recruited by FAK (Lawson et al., 2012), paxillin’s recruitment after talin and vinculin might be coincident with vinculin-paxillin binding mediated by FAK. In line with our measurements, a FRET-based tension sensor study showed that of the three molecules (paxillin, talin, and vinculin), paxillin levels correlate strongest with traction force levels (Morimatsu et al., 2015). Altogether, our findings agree with previous evidence that paxillin levels are an accurate reporter of traction levels, but not NA nucleation.

In summary, our work establishes an unexpected role for an early association of talin and vinculin in a tension-free state as a mechanical prerequisite to the further recruitment of vinculin to talin, which is the foundation of adhesion maturation. While the possibility of talin-vinculin pre-complexation has been discussed in previous studies (Pasapera et al., 2010; Bachir et al., 2014; Khan and Goult, 2019), their function has remained obscure until now. Here, we show that these interactions are an essential step in adhesion assembly. How the pre-complexations are promoted, and whether they are regulated by cellular signals, are two of the critical questions to be addressed in future studies.

Materials and methods

Key resources table
Reagent type (species)
or resource
DesignationSource or referenceIdentifiersAdditional information
E. coliBL21(DE3)Sigma-AldrichCMC0016Electrocompetent cells
Cell line (Cricetulus griseus)CHO-K1ATCCCCL-61Courtesy of Dr.Horwitz, Allen Institute
Cell line
Mus musculus
IMCD Talin1/2 KOThis paperCourtesy of Dr. Zent, Vanderbilt University
Lentiviral expression vectorpLVX-shRNA1ClontechLentiviral construct for stable expression of shRNA.
Lentiviral expression vectorpLVX-CMV-100AddgeneCatalog # 110718Lentiviral construct for stable expression of ectopic proteins.
Transient expression vectorpCDNA3.1(+)ThermoFisher ScientificCatalog # V79020DNA construct for transient expression of ectopic proteins.
Primary antibodyAnti-talin 1AbcamCatalog # 71333WB 1:1000
Primary antibodyAnti-talin 2AbcamCatalog # 105458WB 1:1000
Secondary antibodyGoat anti-Mouse IgG (H+L) Cross-Adsorbed- Perosidaxe AntibodyThermoFisher ScientificCatalog # G-21040WB 1:1000
Secondary antibodyGoat anti-Rabbig IgG (H+L) Cross-Adsorbed-Peroxidase AntibodyThermoFisher ScientificCatalog # G-21234WB 1:1000
Transient expression vectorpCDNA-mNG-Talin-shRNA-R8R8 mutant of Talin1 resistant to shRNA.
Transient expression vectorpCDNA-mNG-shRNA-TalinWT Talin1 resistant to shRNA
Transient expression vectorpCDNA-mNG-TalinWT Talin1
Lentiviral expression vectorpLVXCMV100-mNG-Talin-shRNA-R8R8 mutant of Talin1 resistant to shRNA.
Lentiviral expression vectorpLVXCMV100-mNG-Talin-shRNAWT Talin1 resistant to shRNA
Lentiviral expression vectorpLVXCMV100-mNG-TalinWT Talin1
Transient expression vectorPaxillin-GFPOriginal construct from Horwitz Lab
Transient expression vectorVinculin-GFPOriginal construct from Horwitz Lab
Transient expression vectorTalin-GFPOriginal construct from Horwitz Lab
Recombinant expression vectorpet151-Talin1-R7R8-WT (murine)GeneArtOriginal construct from Goult Lab
Recombinant expression vectorpet151-Talin1-R7R8-VVV (murine)GeneArtOriginal construct from Goult Lab
Recombinant expression vectorpet151-Vinculin-Vd1 (murine)Original construct from Goult Lab
Synthetic peptideRIAM-TBS1 peptide
(RIAM_6_30-C) DIDQMFSTLLGEMDLLTQSLGVDT-C
GLBiochem
Synthetic peptideDLC1 peptide (DLC1_465_489-C)
IFPELDDILYHVKGMQRIVNQWSEK-C
GLBiochem

Cell line

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ChoK1 cells were received from the Horwitz lab (UVA). The ChoK1 cell line was regularly tested for mycoplasma infection in the Horwitz lab. Upon transfer to the Danuser lab, they were again tested for mycoplasma infection, which were negative. IMCD talin1/2 cells were obtained from Roy Zent's lab at Vanderbilt University, where they were tested for authenticity and mycoplasma infection before shipment.

Cell culture

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ChoK1 cells were cultured in Dulbecco’s Modified Eagle Medium (DMEM) with 4.5 g/L D-Glucose, L-Glutamine, and Sodium Pyruvate (Gibco, 11995–065) supplemented with 10% Fetal Bovine Serum (Equitech-Bio, Inc, SFBU30), 1% Anti-Anti (Gibco, 15240112), and 1% Non-Essential Amino Acids (Gibco, 11140076). For transfection, cells were plated in a 6-well plate at ~30% confluency and transfected the next day with 350 ng of fluorescent protein-, or SNAP-tagged adhesion marker, 1 μg of pBluescript (Stratagene) as non-specific DNA, 10 μL of Lipofectamine LTX (Gibco, 15338030), and 2 mL of reduced serum Opti-MEM (Gibco, 31985088) according to the manufacturer’s directions. Four hours after adding the DNA-lipid mixture to the cells, the media was replaced with full DMEM media. Twenty-four hr later, cells were trypsinized, and enriched with flow cytometry for low-level GFP-positive cells. Of this pool, 50,000 cells were seeded on fibronectin-coated (see below) traction-force microscopy substrates in pH 7.4 HyQ-CCM1 media (GE Lifesciences, SH30058.03), supplemented with 1.2 g/L of sodium bicarbonate and 25 mM HEPES. mGFP-talin1 (provided by N. Bate and D. Critchley), paxillin-eGFP (provided by I. Schneider), and mGFP-vinculin (provided by M. Humphries) were used for adhesion-TFM two-channel experiments.

Knock-out and knock-down experiments

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For knock-out experiments, Murine Inner Medullary Collecting Duct (IMCD) Talin1/2 knockout cells (the kind gift of Dr. Roy Zent, Vanderbilt University [Mathew et al., 2017]) were grown in DMEM/F-12 Dulbecco's Modified Eagle Medium/Nutrient Mixture F-12 with L-Glutamine, 15 mM HEPES (Gibco, 11330–032) 10% Fetal Bovine Serum (Equitech-Bio, Inc, SFBU30), and 1% Anti-Anti (Gibco, 15240112) under standard cell culture conditions (5% CO2, 37°C) and passaged regularly at ~70% confluency. Owing to the complete knockout of Talin1/2, these cells are nonadherent and propagate in suspension. To rescue Talin1, IMCD cells were separately transduced using lentivirus with mNeonGreen-tagged WT and R8 mutant Talin1 (T1502V, T1542V, and T1562V) at low-levels using a truncated cytomegalovirus promoter (pLVX-CMV-100, https://www.addgene.org/110718/). To generate lentivirus, human embryonic kidney cells were transiently transfected with transfer vector (pLVX-CMV-100-mNG-Talin-18/pLVX-CMV-100-mNG-Talin-18-R8), viral packaging (psPAX2, https://www.addgene.org/12260/) vector, and viral envelope (pMD2.G, https://www.addgene.org/12259/) vector, at a 1:1:1 ratio (5 mg each) using 250 ml of Opti-MEM (Gibco, 31985088) and 45 ml (1 mg/ml) of polyethyleneimine. The final mixture was incubated for 15 min at room temperature and then transferred dropwise to human embryonic kidney cells. After 48 hr, the viral supernatant was gently removed from the human embryonic kidney cells, filtered with low protein binding 0.45 mm syringe filters, and added to IMCD Talin1/2 knockout cells at ~70% confluency with 12 micrograms/mL of polybrene. Twenty-four hr later the media was replaced with full DMEM/F-12 media, and changes in the adhesion behavior were observed 48 hr post-transduction. Approximately one-week post-infection, non-adherent cells were washed from the dish, and the remaining knockout cells were trypsinized and evaluated for traction force microscopy.

For knock-down experiments, a previously validated shRNA hairpin against talin (GGAAAGCTTTGGACTACTA), located on the N-terminus of talin1, was stably introduced into ChoK1 cells with a pLVX-shRNA1 lentiviral system (Clontech) and selected for with 5 μg/mL of puromycin. Western blot analysis indicated decreased levels of talin expression (Figure 5—figure supplement 1). For rescue experiments, talin1 from mouse Mus musculus was subcloned into the pCDNA3.1(+) mammalian expression vector (ThermoFisher Scientific, V79020). To make the reconstitution vectors insensitive to the shRNA, silent mutations were introduced into the corresponding shRNA target sequence. The new sequence of the reconstitution vectors is now GGAAGGCCCTAGACTACTA. This silent mutation was introduced into mNeonGreen talin1 pcDNA3.1 vector, which was used for WT reconstitution. The pCDNA3.1-mNG-Talin1-shRNA vector was mutated for talin1 R8vvv mutant with mutation sites at T1502V, T1542V, and T1562V. The R8 mutations (T1502V, T1542V, and T1562V, according to mouse numbering), alter the stability of the talin R8 domain. For vinculin imaging, mNeonGreen (Allele Biotechnology) was replaced with SNAP-Tag using seamless cloning, and labeled with SNAP-Cell TMR-Star (New England Biolabs, S9105S) or SNAP-Cell 647-SiR (New England Biolabs, S9102S) according to manufacturer’s recommendations. All protein-coding regions of expression constructs were verified with traditional primer walking and Sanger sequencing.

Expression of recombinant polypeptides

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For in vitro analyses, murine vinculin Vd1 (residues 1–258), murine talin R7R8wt (residues 1357–1653), and R7R8vvv (residues 1357–1653; T1502V, T1542V and T1562V) were cloned into a pET151 vector (Invitrogen) and expressed in E. coli BL21(DE3) cells cultured in LB. Standard nickel-affinity chromatography was used to purify the His-tagged recombinant proteins as described previously (Whitewood et al., 2018). The proteins were further purified using anion exchange chromatography following cleavage of the 6xHis-tag with TEV protease. Protein concentrations were determined using their respective extinction coefficients at 280 nm.

Circular dichroism (CD)

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Spectroscopy was performed using a JASCO J-715 spectropolarimeter equipped with a PTC-423S temperature control unit. Denaturation profiles were measured from 20 to 80°C at 0.2°C intervals by monitoring the unfolding of α-helices at 208 nm. A total of 0.1 mg/mL of protein was dissolved in phosphate buffered saline (PBS). Measurements were made in a quartz cell of 0.1 cm path length.

Fluorescence polarization assays

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To determine if other binding partners of talin R8 domain except for vinculin can still bind to R7R8vvv fragment, the relative binding affinities were measured using an in vitro fluorescence polarization assay. The R8 interacting, LD-motif containing peptides from DLC1 and RIAM, that is, DLC1_465–489-C (IFPELDDILYHVKGMQRIVNQWSEK-C) and RIAM_6–30-C (DIDQMFSTL LGEMDLLTQSLGVDT-C), were coupled to a thiol-reactive fluorescein dye via the terminal cysteine. Peptides with a C-terminal cysteine were synthesized by GLBiochem (China). Stock solutions (i.e. peptide + fluorescein) were made in phosphate-buffered saline (PBS; 137 mM NaCl, 27 mM KCl, 100 mM Na2HPO4, 18 mM KH2PO4, pH 7.4), 1 mM TCEP and 0.05% Triton X-100. Excess dye was removed using a PD-10 desalting column (GE Healthcare, Chicago, IL, USA). Titrations were performed in PBS using a constant 1 μM concentration of fluorescein-coupled peptide with increasing concentration of R7R8 fragment (either WT or vvv mutant); final volume 100 μM in a black 96-well plate. Fluorescence polarization (FP) measurements, in which the binding between the two polypeptides results in an increase in the fluorescence polarization signal, were recorded on a BMGLabTech CLARIOstar plate reader at room temperature and analyzed using GraphPad Prism. Kd values were calculated with nonlinear curve fitting using a one-site total binding model.

Analytical gel filtration

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Gel filtration was performed using a Superdex-75 size exclusion chromatography column (GE Healthcare) at a flow rate of 0.7 mL/min at room temperature in 50 mM Tris pH 7.5, 150 mM NaCl, 2 mM DTT. A sample of 100 μL consisting of 100 μM of each protein was incubated at a 1:1 ratio at 25°C for 10 min. The elution was monitored by a Malvern Viscotek SEC-MALS-9 (Malvern Panalytical, Malvern, UK).

Western blot

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Cells were transfected under identical conditions as they were for imaging experiments but with a 10 cm dish and sorted with a flow cytometer (FACS Aria II SORP) for low expression. Cells were lysed by adding 2x laemmli + 10% b-ME, vortexing, and heating at 95°C for 10 min. Protein concentration was measured, and the same amount was loaded for each lane. The gel was semi-dry transferred with a turbo blot, and then incubated overnight in 5% milk in tris-buffered saline with 0.1% Tween 20 (TBST) at four degrees. Protein was visualized with an anti-talin antibody at 1:1000 and the loading control was visualized with anti-b-actin at 1:5000, each in 0.5% milk/TBST overnight at 4°C. Gels were then rinsed with TBST, and probed with IgG:horseradish peroxidase in 0.5% milk/TBST at 4°C for 1 hr and then at room temperature for another 30 min. Gels were rinsed three times for 20 min in TBST and then detected with enhanced chemiluminescence.

TFM substrate preparation

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All silicone substrates had a diameter of 35 mm, a stiffness of 5 kPa (with the exception of Figure 1—figure supplement 7), were embedded with 580/605 or 640/647 (λEXEM) 40 nm-diameter beads, and were compatible with total internal reflection fluorescence illumination. Substrates were coated with fibronectin (Sigma Aldridge, F1141) the same day as imaging experiments were conducted by mixing 20 μL of a 10 mg/mL 1-ethyl-3-(3-dimethylaminopropyl) carbodiimide hydrochloride (EDC) solution, 30 μL of a 5 mg/mL fibronectin solution, and 2 mL of Ca2+ and Mg2+ containing Dulbecco’s Phosphate Buffered Saline (DPBS, Gibco, 14040117) for 30 min at room temperature. Thereafter, the substrate was rinsed two times with DPBS, and incubated with 2 mL of 0.1% (w/v) bovine serum albumin in DPBS for another 30 min at room temperature, and rinsed several times with PBS prior to seeding with 50,000 transiently transfected cells.

Live-cell TIRF imaging for TFM and adhesion proteins

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After being seeded, the cells were allowed to adhere to the substrate for ~1 hr prior to imaging. This was one of the more effective ways to make sure that cells had active protrusions. Cells were imaged with a DeltaVision OMX SR (General Electric) equipped with ring-TIRF, which mitigates laser coherence effects and provides a more homogeneous illumination field. This microscope is equipped with a 60x, NA = 1.49, objective, and 3 sCMOS cameras, configured at a 95 MHz readout speed to further decrease readout noise. The acquired images were in 1024 × 1024 pixel format with an effective pixel size of 80 nm. Imaging was performed at 37°C, 5% carbon dioxide, and 70% humidity. To maintain the optimal focus, laser-based identification of the bottom of the substrate was performed prior to image acquisition, with a maximum number of iterations set to 10. Laser powers were decreased as much as possible, and the integration time set at 200 ms, to avoid phototoxicity. At the back pupil of the illumination objective, the laser power for both 488 and 568 nm lasers was ~44 μW. Imaging was performed at a frequency of 1 Hz for 5–10 min, and deviations between the alignment for each camera were corrected in a post-processing step that provides sub-pixel accuracy. After imaging, cells were removed from the substrate with a 30% bleach solution, and the beads on the relaxed gel substrate were imaged for each cell position. Rapid imaging was necessary to mitigate swelling effects in the silicone substrate and to resolve traction forces in nascent adhesions.

TFM force reconstruction

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Bead images of the deformed gel – acquired when a cell was on the substrate – and a ‘reference bead image’ of the relaxed gel acquired after cell removal – were processed for traction reconstruction as described previously (Han et al., 2015). Briefly, the bead images of the deformed gel were compared with the reference image using particle image velocimetry. A template size of 17–21 pixels, and a maximum displacement of 10–80 pixels, depending on the bead density and overall deformation, were used for cross-correlation-based tracking of the individual beads. The displacement field, after outlier removal, was used for traction field estimation over an area of interest. The area of interest on the reference bead image was meshed with square grids of the same width, which depends on the average area per bead. The forward map, which defines the expected deformation of the gel at all bead locations given a unit force at a particular mesh of the force grid, was created by solving Boussinesq Eq. under the assumption of infinite gel depth. This forward map was then used to solve the inverse problem, that is given the measured field of bead displacements, the underlying traction field is determined. The solution to this inverse problem is ill-conditioned in that small perturbations in the measured displacement field can yield strong variation in the reconstructed traction field. To circumvent this problem, the traction field was estimated subject to L1-norm regularization. As discussed in detail in Han et al., 2015, L1-norm regularization preserved the sparsity and magnitude of the estimated traction field. Also as discussed and validated in Han et al., 2015, the application of L1-norm regularization over the L2-norm regularization most traction force microscopy studies employ is essential to resolve force variation at the length scale of the distances between individual nascent adhesions. The level of regularization is determined by a control parameter. We chose the parameter based on L-curve analysis, which guaranteed a fully automated and unbiased estimate of the traction field (Han et al., 2015). Strain energy, which represents the mechanical work a cell has put into the gel, was quantified as 1/2 × displacement × traction, integrated over a segmented cell area. The unit of this integral is femto-Joule.

Adhesion segmentation, detection, and tracking

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Focal adhesions (FAs) and diffraction-limited NAs were detected and segmented as previously described (Han et al., 2015). Briefly, FAs from images of either labeled paxillin, talin, or vinculin were segmented with a threshold determined by a combination of Otsu’s and Rosin’s algorithms after image pre-processing with noise removal and background subtraction. Segmented areas larger than 0.2 μm2 were considered for focal contacts (FCs) or FAs, based on the criteria described by Gardel et al., 2010. Individual segmentations were assessed for the area and the length, which is estimated by the length of major axis in an ellipse that fit in each FA segmentation. FA density was calculated as the number of all segmentations divided by the cell area. Nascent adhesions were detected using the point source detection described in Aguet et al., 2013. Briefly, fluorescence images were filtered using the Laplacian of Gaussian filter and then local maxima were detected. Each local maximum was then fitted with an isotropic Gaussian function (standard deviation: 2.1 pixels, i.e. ~180 nm) and outliers were removed using a goodness of fit test (p=0.05). NA density was defined as the number of NAs divided by the entire cell area.

Adhesion classification

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From the adhesion tracks, features 1–9 in Supplementary file 1A were captured from the individual fluorescence intensity traces, and features 10–21 in Supplementary file 1A from the corresponding spatial properties, some in reference to the position and movement of the proximal cell edge and to the overlap with segmentations of focal adhesions and focal complexes. A total of nine class outputs were used to train the adhesion tracks. We believe that nine classes are a minimum number that represents a heterogeneity of the cell-ECM adhesions in a cell because the five were dedicated to classify NAs. The three classes assigned for FA classes can be viewed as insufficient in terms of representing a whole FA population and can be expanded further into a larger number of classes, but they were sufficient in this study because NAs were its main scope.

The classification was accomplished using a cubic support vector machine (SVM), which was proven to be the most accurate among linear classifiers. The classifier was evolved in a human-in-the-loop fashion, i.e. the user iteratively adjusted machine-generated classifications. The initial training data was labeled with qualitative criteria described in Supplementary file 1B. To facilitate the labeling process, an automatic, filtering-based, labeling was also employed (see Supplementary file 1C).

Both manual labeling and automatic labeling have advantages and drawbacks in terms of classification accuracy: while the manual labeling is less consistent due to subjectivity and human error, the automatic labeling has deficiencies in terms of incompleteness of the filtering criteria in capturing all essential properties of different adhesion classes. To overcome these drawbacks, both methods were employed in a way that the automatic labeling was performed first, and then manual labeling was added for insufficiently-labeled classes. During the manual labeling, adhesion classifications were immediately updated and presented to the user to allow class reassignments of selected adhesions. The labeling process was regarded to be completed once at least 10 adhesions were labeled for each class. To remove classification bias due to potential imbalance in the number of labels across the classes, the minority classes were oversampled, and the majority classes were undersampled, based on the mean number of labels (Krawczyk, 2016). After training a classifier on one movie, for a new movie, another iteration of automatic-and-manual labeling was executed to update the classifier, which was applied to predict the classes of adhesions in the movie. Separate classifiers were built for talin-, vinculin-, and paxillin-tagged adhesions.

Statistical methods

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Processed data such as traction, assembly rate, time delays, lifetimes, adhesion densities and areas in different conditions were compared using Mann-Whitney non-parametric test since all individual distributions do not follow a normal distribution. Testing for normal distribution was done by one-sample Kolmogorov-Smirnov test using a ‘kstest’ function in Matlab.

Software availability

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A GUI-based Matlab software is shared via GitHub at https://github.com/HanLab-BME-MTU/focalAdhesionPackage.gitHan, 2021; copy archived at swh:1:rev:6aeb3593a5fd3ace9b0663d1bf0334decfb99835.

References

Decision letter

  1. Jonathan A Cooper
    Senior Editor; Fred Hutchinson Cancer Research Center, United States
  2. Reinhard Fässler
    Reviewing Editor; Max Planck Institute of Biochemistry, Germany
  3. Reinhard Fässler
    Reviewer; Max Planck Institute of Biochemistry, Germany
  4. Nico Strohmeyer
    Reviewer
  5. Thomas Weidemann
    Reviewer; Max Planck Institute of Biochemistry, Germany

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

Acceptance summary:

The manuscript nicely demonstrates how talin, vinculin and paxilin coordinate maturation of nascent cell-substrate adhesions and mechanotransduction to control motility and shape of cells.

Decision letter after peer review:

[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 "Formation of talin-vinculin pre-complexes dictates maturation of nascent adhesions by accelerated vinculin recruitment" for consideration by eLife. Your article has been reviewed by three peer reviewers, including Reinhard Fässler as the Reviewing Editor and Reviewer #1, and the evaluation has been overseen by a Senior Editor. The following individuals involved in review of your submission have agreed to reveal their identity: Nico Strohmeyer (Reviewer #2); Thomas Weidemann (Reviewer #3).

The reviewers spent quite a bit of time reviewing and then discussing this paper, in part because of the numerous concerns that were raised in the reviews. Although the reviewers found the manuscript very interesting and considered the possibility of having you resubmit a revised manuscript after clarifying the concerns/comments, it was ultimately decided that the experiments that needed to be done were too many for the time frame of 2-3 months. Based on these discussions, we regret to inform you that your work will not be considered further for publication in eLife. We strongly encourage you, however, to submit a new version of your manuscript in case you address our concerns and comments.

Reviewer #1:

The paper reports five classes of NAs based on their different mechanical behavior and recruitment dynamics of talin, vinculin and paxillin. The authors focus their study on two NA types, non-maturing and maturing NAs, and demonstrate that they greatly differ with respect to their force levels and talin/vincluin recruitment. Whereas maturing NAs recruit talin preloaded with vinculin, non-maturing NAs a non-vinculin bound talin, which prevents them to transduce high forces to the integrin/ligand bond and strengthen the bond.

The described methodology (combination of machine learning, high-resolution traction force microscopy, single-particle-tracking and fluorescence fluctuation time-series analysis) and findings (division of NAs into different subgroups based on their kinetic and mechanical properties and the importance of a pre-formed talin-vinculin complex for NA maturation to FAs) are of interest for the readership of eLife. However, publication in eLife requires additional analysis:

1) To validate their results the authors should test with simple manipulations whether the balance between the different adhesion subclasses can be shifted (e.g. Mn2+-treatment, PLL, different substrate stiffness) to validate their findings.

2) It is not clear why the authors used ChoK1 cells as their "model system" and expressed fluorescently tagged talin, vinculin or paxillin on top of the endogenous proteins or after knock-down of talin1 instead of a clean knockout background. How can the authors be sure that the fluorescently tagged-proteins are incorporated into NAs and FAs similar to untagged proteins?

Regarding the talin1 knockdown: is it know if talin2 is expressed in ChoK1 cells? Did the authors observe differences in the expression levels of talin1 wt-mNG or talin1 R8vvv-mNG? Why is the mNG-tagged talin1 running at a seemingly similar MW then the endogenous talin1 in Figure 5—figure supplement 1?

3) Their data demonstrate that the talin1 R8vvv mutation, which prevents talin-vinculin pre-complex formation, restricts NAs from maturing into focal adhesions. Is there a way to increase talin-vinculin pre-complex formation (e.g. by expressing an activated form of vinculin) to increase the number of maturing (G2) NAs? To support their hypothesis of the importance of pre-assembled talin-vinculin complexes it would be interesting to test the effect of talin1 variants impaired in actin binding to determine the importance of these independent actin linkages. What has a stronger effect?

4) The authors write that their analysis suggests, "that a very large number of adhesion protein aggregates, detectable through either talin, vinculin, or paxillin recruitment, never engage with the substrate." Is it not an overstatement to say that the failure to detect traction forces shows that these adhesions are not engaged with the substrate? Could they engage with the substrate but not link to actin?

Reviewer #2:

Han et al. describe a pre-complex of talin and vinculin formed in the cytosol, in which the vinculin vd1 domain interacts with the talin R8 domain. The authors demonstrated this complex with recombinant protein fragments of talin and kindlin. An R8vvv mutation of talin mechanically stabilizes R8, reduces the formation of the pre-complex and the number of maturing NAs in cells. The authors used high resolution traction force microscopy and TIRF to conclude that the talin-vinculin pre-complex is essential to mature NAs and sustain traction forces applied to them characterized. While the data and the model potentially provide a functional role of R8 in NA maturation, several questions arise.

1) To better understand the experimental setup, a figure explaining the method as well as an image of the flexible substrate with embedded beads measuring traction forces (including the processing steps) would be helpful. Additionally, the bead density and average distance between beads in these substrates should be reported. The mechanical properties of the substrates should be evaluated, as they might differ from batch to batch. Technically, I am wondering where the different background traction levels (see for example Figure 2, different traction force traces) or the traction measured outside a cell (see Figure 5A-C) arise from.

2) Talin and vinculin are known to exhibit auto-inhibited conformations, in which an interaction between R8 and Vd1 may not be possible. Can the authors show in a more physiological setting (preferably full-length proteins) that talin and vinculin form a complex in solution? Additionally, in Figure 4C it seems that talin1 R8vvv can still bind vinculin, although not as proficient. Can the authors show the importance of vinculin binding to R8 for the maturation of NAs into FCs more directly by mutating only the vinculin binding site in R8?

3) The authors state that in maturing (G2) NAs vinculin and talin are recruited together. However, in Figure 2O it appears that in about 25% (and in Figure 7J ~50%) of cases, vinculin is recruited before talin. In Figure 7H it appears that vinculin is recruited to NAs 14 s earlier than talin. Could the authors elaborate how vinculin is recruited before talin is visible?

4) How do the authors exclude the possibility that talin is quickly unfolded (as described in the legend of Figure 8) by forces from the cytoskeleton, which induces a fast recruitment of vinculin to talin in maturing NAs (which may be dependent on R8)? This process could be faster than the sampling rate and hence a pre-complex is not required to form in the cytosol.

5) The authors show multiple traction force maps and images acquired for cells that show very different phenotypes although being the same cell line. In Figure 1A-F ChoK1 cells are shown that overexpress mGFP-tagged talin, vinculin or paxillin but have very different morphologies, sizes (d-f) and exhibit very different traction forces (a-c) where the scale is 0-300 Pa for talin, 0-1800 Pa for vinculin and 0-1000 Pa for paxillin. In Figure 2 it is similarly diverse where traction force scales range from 0-800 Pa (in h) to 0-4000 Pa (in a). What is the reason for this heterogeneity in morphologies and traction forces within the same cell line?

6) In maturing (G2) NAs, does the constantly increasing traction force arise from the accumulating more vinculin at the adhesion site (i.e. stronger connection to the actomyosin), or from the growth of the adhesion site (i.e. accumulating more integrins)? In Figure 2G, a continuous recruitment of talin is observed, which would argue for the increase in traction forces due to the recruitment of additional integrins. How would the authors explain the pre-formation of talin-vinculin complexes to be important for the recruitment of integrins to the adhesion site (in maturing NAs)?

7) For testing the effect of the R8vvv talin mutant in cells, the authors used shRNA to knock-down talin1 levels before rescuing cells with either WT or R8vvv talin1. How is the talin expression level of WT talin1 rescued cells compared to R8vvv rescued cells? Further, an assessment of the morphology (whole cells and possible adhesion sites) of talin1 KD cells would be beneficial to appreciate the difference between R8vvv and WT talin 1. This would exclude the possibility that apparent differences between R8vvv and WT talin1 re-expressing cells arise from potentially different talin1 expression levels, a potential expression of talin2 in response to reduced talin1 levels or residual expression of WT talin1 in KD cells.

8) The authors state that the talin-vinculin pre-complex formation is reduced in talin1-R8vvv expressing cells. However, talin1-R8vvv expressing cells show adhesion sites larger than NAs (Figure 5A). How do larger adhesion sites form in cells expressing talin1 R8vvv, if the vinculin binding to R8 is essential for NAs maturation? Representative image of labeled vinculin and talin within one cell (as analysed for Figure 7) would be beneficial to understand how the vinculin recruitment is affected by the R8vvv mutation.

9) In their very clear conclusive Figure 8 the authors depict different sites for paxillin in non-maturing vs. maturing NAs. Do the authors have experimental evidence for this? Further, the legend describes hypotheses, which can explain experimental data but experimental evidence from the reported data is limited. For example, that talin stretching is limited to a shorter level in non-maturing NAs and talin stretches faster in maturing NAs has, to my point of view, not been shown in the presented data.

Reviewer #3:

Han et al. describe in great detail the molecular decision process that leads from transient nascent adhesions (NAs) forming constantly at the plasma membrane to focal adhesions (FAs) stably connecting the ECM with the actin cytoskeleton. The assembly kinetics of three GFP-tagged NA components (talin, vinculin and paxilin) is tracked down by TIRF microscopy and their behavior is correlated with local traction force determined by imaging the displacement of fluorescent beads in the hydrogel support, on which the cells grow. The authors apply machine learning algorithms to identify two classes of events both linked to force-assisted growth but with opposite fates. Using a new talin mutant, it is shown that binding of vinculin at a specific talin domain (R8) is required to establish the feed forward control for further vinculin recruitment and the steady build-up of local traction. In contrast, sequential binding of talin, vinculin and paxilin leads to maturation abort. I consider the study to be compelling, the technology pioneering and the results significant. Given that the topic is quite complex, the manuscript is of remarkable clarity. A direct comparison with R3 talin-mutants would add further evidence, however, such experiments are probably not within the scope of a revision. My listed points represent mainly comprehension questions that could encourage improvements of the text. Overall, I am in full support of publication.

1) Is there statistics on how many G2-class NAs actually mature into FAs?

2) "Event-based time series analysis" measures the appearance of fluorescently tagged proteins. What is the sensitivity of the method? How many mGFPs have to accumulate to detect a spot?

3) At the stage of Figure 2, is the existence of an unstrained talin-vinculin pre-complex really a stringent conclusion? The time resolution of imaging is one second. During this time bin, proteins with a Kd in the μM range may undergo several binding and unbinding cycles. Likewise, the forces may exhibit fast fluctuations especially during the onset when only a small number of force transmitting proteins have been accumulated (see Figure 3B). Thus, there is plenty of "temporal" space for different scenarios of "what is first" that is technically inaccessible. Please comment.

4) Figure 3B, the average force growth in the first 10 seconds may simply be zero due to the sensitivity limit of TFM. Or does this represent a finding?

5) Are G1-NAs programmed to terminate, because talin has too much time for erroneous interactions? Explicitly, do traction-dependent vinculin binding site (VBS), other than R8, represent "wrong sites" in the sense that cooperative vinculin assembly is even prevented?

6) At which stage of NA maturation then must force be sensed?

7) Is mutant R8vvv-talin totally impaired in vinculin binding (Figure 4D) or is there residual probability for exposing the VBS (Figure 4C)? The biochemical result is contradictory.

8) The difference between vinculin epitopes in domain R3 and R8: As a naïve reader one would assume that a VBS of the relaxed state locates at the surface of the protein. I consider it conceptually difficult to acknowledge that the VBS in domain R8 is active in an unstrained state but nevertheless requires unlocking of the threonine belt and partial unfolding of the α-helical bundle; maybe this warrants a sentence or two.

9) Cells expressing the R8vvv mutant are clearly impaired in NA maturations as documented by the cellular analysis (Figure 5). However, there are mature FAs. This may either relate to residual wildtype talin that could not be suppressed by shRNA (as the WB shows, Figure 5—figure supplement 1), residual binding of the mutant R8vvv-talin or even an undefined pathway into the cooperative vinculin assembly. These possibilities are not mentioned.

10) The traction growth rates in Figure 6K reach much more into the negative range than those in Figure 3B and 3C. Do negative values signify initial forces that disappear? In addition, there is quite a large difference in the scale of y-axis in Figure 6K (100-200 Pa/sec) as compared to Figure 3B and C (1-2 Pa/sec).

11) The body text referring to Figure 6 at the end of the paragraph says "These results suggest that the vinculin-talin pre-complex is essential for the development of force across NAs…". In my opinion this statement still remains an assumption, since I could not find direct evidence in Figure 6. The behavior of the mutant R8vvv-talin, if G2 characteristics are fulfilled, appear to be indistinguishable from the mGFP-tagged version described in Figure 2. The fact that there is a problem with force growth can only be explained in the light of Figure 7. Re-ordering these arguments may be considered.

12) Again, the two expression systems lead to a different range of values: The absolute vinculin assembly rates in Figure 7I as well as the rate increase between G2 vs G1 are much smaller than in Figure 3A. Is this due to different protein expression levels? Why were different integration times used (30 seconds in Figure 7I in contrast to 10 seconds in Figure 3A)?

13) Is actin always engaged first and then the integrin tails? In other words, is there support for a "search mode" of a small number of competent actin-talin-vinculin-heads for integrin tails linked to the ECM (Figure 8, G2, first panel)?

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

Thank you for submitting your article "Talin-vinculin precomplex drives adhesion maturation by accelerated force transmission and vinculin recruitment" for consideration by eLife. Your article has been reviewed by three peer reviewers, including Reinhard Fässler as the Reviewing Editor and Reviewer #1, and the evaluation has been overseen by Jonathan Cooper as the Senior Editor. The following individuals involved in review of your submission have agreed to reveal their identity: Nico Strohmeyer (Reviewer #2); Thomas Weidemann (Reviewer #3).

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

Summary:

Cell adhesion to the extracellular matrix is mediated by integrins which require an activation step and exposure to tension for ligand binding, adhesion site formation and maturation. Han and co-workers elegantly demonstrate that the maturation of small into large adhesions depends on the isochronic association of talin (integrin activator) and vinculin (force transmitter) with integrins and the generation of increasing forces. The study represents an inspiring example for the power of microscopy and machine learning to approach the emergence of cellular functions that involve molecular decision making.

Essential revisions:

1) The conclusion that the concurrent recruitment of talin and vinculin to maturing NAs corresponds with the existence of a cytoplasmic precomplex of the two proteins. Although it is likely that such precomplexes form prior to NA recruitment, it has not been demonstrated in the current paper. Hence, it is necessary to tone this claim down in the title, Abstract, Results section and Discussion of the manuscript.

2) Use same assembly rates (Figure 3A, Figure 6J, and Figure 7M) in y-axis. It is not clear what is meant by 1/min in Figures 3A/7M, and what is the rational for the different time frames analysed (Figure 3A: 2 min; Figure 6: 1 min; Figure 7: 0.5 min).

3) Although the authors state that the R8vvv mutation does not affect the talin recruitment rate, Figure 6J shows that in both G1 and G2 NAs talin R8vvv is recruited at higher rates than WT talin1. Test the rates for statistical significance (for G1 and G2 R8vvv vs. WT). In case talin1 R8vvv is indeed recruited faster to adhesion sites, the authors should try finding an explanation.

4) The authors state that traction increase for talin1 R8vvv is reduced compared to WT talin1. The statistical test in Figure 6K to corroborate this statement is missing.

5) Subsection “Simultaneous talin-vinculin imaging confirms vinculin’s recruitment after talin for non-maturing NAs and concurrent recruitment for maturing NAs”: should probably reference Figure 7N (instead of Figure 7J) and Figure 7O (instead of Figure 7K).

6) "Fluorescence fluctuation analysis" may be misleading as it usually refers to FCS, PCH, i.e., methods that involve a statistical correlation analysis. Here the authors directly analyze the time traces by thresholds and slopes.

7) The density 2.2 beads/µm2 cannot be reproduced from average spacing 0.42 µm via the relation: density = 1/sqrt(spacing).

8) "Both peptides bound…" This sentence sounds strange and may be changed to "Both peptides showed comparable affinity towards wildtype R7R8 and the R8R7vvv mutant, confirming.…"

9) "The threonine belt in talin R8 is responsible…" The wording is not clear since only the binding epitope itself can be responsible for binding and the threonine belt is, rather indirectly, regulating accessibility and hence the on-rate. Do these data suggest that RIAM and DLC1 use binding epitopes spatially distinct from the VBS within the R8 bundle? Or is RIAM and vinculin binding indeed competitive?

https://doi.org/10.7554/eLife.66151.sa1

Author response

[Editors’ note: the authors resubmitted a revised version of the paper for consideration. What follows is the authors’ response to the first round of review.]

Reviewer #1:

The paper reports five classes of NAs based on their different mechanical behavior and recruitment dynamics of talin, vinculin and paxillin. The authors focus their study on two NA types, non-maturing and maturing NAs, and demonstrate that they greatly differ with respect to their force levels and talin/vincluin recruitment. Whereas maturing NAs recruit talin preloaded with vinculin, non-maturing NAs a non-vinculin bound talin, which prevents them to transduce high forces to the integrin/ligand bond and strengthen the bond.

The described methodology (combination of machine learning, high-resolution traction force microscopy, single-particle-tracking and fluorescence fluctuation time-series analysis) and findings (division of NAs into different subgroups based on their kinetic and mechanical properties and the importance of a pre-formed talin-vinculin complex for NA maturation to FAs) are of interest for the readership of eLife. However, publication in eLife requires additional analysis:

1) To validate their results the authors should test with simple manipulations whether the balance between the different adhesion subclasses can be shifted (e.g. Mn2+-treatment, PLL, different substrate stiffness) to validate their findings.

We thank the reviewer for this suggestion. To evaluate if a simple manipulation could shift the balance between adhesion classes, we compared adhesion classification for ChoK1 cells transfected with vinculin GFP on both 5 kPa and 18 kPa stiff substrates. As anticipated, the subpopulation for each group showed significant and meaningful differences between the two stiffness conditions. For example, we found major reduction in G1, G2, G3 and G5 groups – all NAs – and an increase in G7 and G8 groups – major FAs – for cells cultured on the stiffer 18 kPa substrates. We believe that these differences can be attributed to the increased stability of FAs in stiff environments.

2) It is not clear why the authors used ChoK1 cells as their "model system" and expressed fluorescently tagged talin, vinculin or paxillin on top of the endogenous proteins or after knock-down of talin1 instead of a clean knockout background. How can the authors be sure that the fluorescently tagged-proteins are incorporated into NAs and FAs similar to untagged proteins?

While there are pros and cons with any cell type, this study is a follow up on a series of papers by Rick Horwitz and colleagues, which had provided extensive (and widely accepted) insights into adhesion formation and maturation. Historically, Rick selected the CHO epithelial cell because its adhesion assembly process was slower than seen in highly contractile cells, and therefore the individual molecular steps of adhesion formation could be readily resolved by live cell imaging. Importantly, the literature suggests that the adhesion assembly and maturation process in CHO cells has been largely recapitulated in other cells types.

As requested, we now reproduce our key findings in talin1/2 double knockout IMCD cells. Of note, our concern with these cells is their potential for long-term genetic adaptation and phenotypic drift owing to the disruption of a central adhesion element. Indeed, these cells propagate in suspension, which is likely accompanied by significant shifts in gene expression and cell signaling. Technically, the IMCD cells were challenging to transiently transfect, which led us to stably transduce them with lentiviruses. Virus production was also very inefficient because full-length GFP-tagged talin1 is well-beyond the viral packaging capacity of most lentiviral systems. Nonetheless, after months of troubleshooting, we succeeded in the expression of both WT and R8vvv mutant talin in knockout cells. As in ChoK1 cells, rescue of IMCD cells expressing with R8vvv mutant showed significantly less adhesion maturation (See Figure 5).

GFP-tagged talin, paxillin, and vinculin have also been used productively for decades in the adhesion community. While one must be cautious tagging any protein, we use constructs that have been employed extensively in other labs and have been accepted as faithful reporters of adhesion formation or maturation. We also agree that overexpression is a valid concern. However, as described in the original manuscript, we transfected cells with a minimal amount of DNA (200 ng), or transduced cells with lentiviral vectors with a truncated CMV promoter that reduces expression 100-fold. In both cases, all cells were sorted for minimal expression (e.g., just greater than autofluorescence) with a flow cytometer just prior to imaging. Importantly, we observed quantitatively similar results for adhesion classification and maturation under talin knockdown and knockout conditions, for rescue experiments involving GFP- and mNeonGreen-tagged talin. Thus, our results are independent of the fusion protein, or whether or not endogenous talin is present.

Regarding the talin1 knockdown: is it know if talin2 is expressed in ChoK1 cells?

To evaluate the expression of different talin isoforms in ChoK1 cells, we performed a western blot for talin1 and talin2 in both WT and talin1 knockdown cells. Here, we find that talin2 is expressed at very low levels in WT and talin1 knockdown cells. While we cannot exclude some compensation between isoforms, the fact that we observe robust defects and proper rescue upon expression of talin1 variants suggests that interference by talin2, if any, is minor. Likewise, experiments in IMCD cells, where talin1 was rescued, but not talin2, also suggest that crosstalk between isoforms is negligible. These western blots are now provided in Figure 5—figure supplement 2 and discussed in the text.

Did the authors observe differences in the expression levels of talin1 wt-mNG or talin1 R8vvv-mNG?

As previously mentioned, all cells were evaluated and sorted with a flow cytometer just prior to imaging. Here, the intensity distributions of cells expressing wt-mNG talin1 and R8vvv-mNG talin1 were indistinguishable. Thus, we have no reason to believe that these variants are differentially expressed or degraded.

Why is the mNG-tagged talin1 running at a seemingly similar MW then the endogenous talin1 in Figure 5—figure supplement 1?

The molecular weight of talin1 and mNG are 270 and 26.6 kDa, respectively. As such, tagging talin1 with mNG only yields a 10% increase in molecular weight. Furthermore, in SDS-PAGE, electrophoretic migration scales logarithmically with the molecular weight of the protein. For these reasons, we anticipate that the tagged protein will have a very similar electrophoretic mobility. Nonetheless, we now include in Figure 5—figure supplement 1 the western blot results from WT cells overexpressing mNG-tagged talin 1 or mNG-talin R8vvv, which clearly show that a minor separation exists between endogenous talin 1 and mNG-tagged talin 1.

3) Their data demonstrate that the talin1 R8vvv mutation, which prevents talin-vinculin pre-complex formation, restricts NAs from maturing into focal adhesions. Is there a way to increase talin-vinculin pre-complex formation (e.g. by expressing an activated form of vinculin) to increase the number of maturing (G2) NAs?

We thank the reviewer for this suggestion. To increase the formation of a talin-vinculin pre-complex, we would need to identify a vinculin mutant with a higher affinity for R8. Unfortunately, we are not aware of such a mutant. There is a constitutively active vinculin mutant (T12) that promotes mature, over-sized adhesions, but it interacts with talin1 vinculin binding sites with its head domain (Cohen, D.M., Chen, H., Johnson, R.P., Choudhury, B., and Craig, S.W. (2005), J Biol Chem), which is not necessarily specific to R8. Thus, a potential experiment with the mutant will obviously induce more maturing NAs, but it would not test whether the pre-complex formation is R8-specific.

To support their hypothesis of the importance of pre-assembled talin-vinculin complexes it would be interesting to test the effect of talin1 variants impaired in actin binding to determine the importance of these independent actin linkages. What has a stronger effect?

We agree with the reviewer; it is a fascinating question whether talin R8 – vinculin – actin bridges could functionally replace the direct talin – actin bridges. However, we want to clarify, this is not our suggested model. As the cartoon in Figure 8 illustrates, we envision that both bridges have to work in parallel to support the formation of a maturing adhesion. Although we cannot exclude that in the very earliest stages talin R8 – vinculin – actin bridges might be sufficient to raise the traction above the levels seen measured in talin R8vvv expressing cells (i.e. the R8 – vinculin – actin bridges are more important than the talin – actin bridges), it is well established that the absence of the talin ABS causes a significant defect in adhesion formation and maturation that will likely make a G1 / G2 distinction very challenging. We point out to the reviewer that these experiments are tedious and given the reduced work capacity with the COVID pandemic we had to choose between the “necessary” and the “interesting”. We hope the reviewers and editor understand that we focused the continuation of the originally reviewed version of the study on bolstering the key novelty of our data: A talin – actin bridge alone is not sufficient to trigger mechanical adhesion maturation but the process requires additional early support from vinculin – actin bridges that bind to a still folded talin configuration.

4) The authors write that their analysis suggests, "that a very large number of adhesion protein aggregates, detectable through either talin, vinculin, or paxillin recruitment, never engage with the substrate." Is it not an overstatement to say that the failure to detect traction forces shows that these adhesions are not engaged with the substrate? Could they engage with the substrate but not link to actin?

We agree with the reviewer and have changed the wording appropriately. It now states that “a very large number of adhesion protein complexes, detectable through either talin, vinculin, or paxillin recruitment, do not transduce measurable traction forces (e.g., less than 20 Pa). Indeed, these complexes could result from an incomplete molecular clutch and be engaged with the substrate but not actin, or vice versa.”

Reviewer #2:

Han et al. describe a pre-complex of talin and vinculin formed in the cytosol, in which the vinculin vd1 domain interacts with the talin R8 domain. The authors demonstrated this complex with recombinant protein fragments of talin and kindlin. An R8vvv mutation of talin mechanically stabilizes R8, reduces the formation of the pre-complex and the number of maturing NAs in cells. The authors used high resolution traction force microscopy and TIRF to conclude that the talin-vinculin pre-complex is essential to mature NAs and sustain traction forces applied to them characterized. While the data and the model potentially provide a functional role of R8 in NA maturation, several questions arise.

1) To better understand the experimental setup, a figure explaining the method as well as an image of the flexible substrate with embedded beads measuring traction forces (including the processing steps) would be helpful. Additionally, the bead density and average distance between beads in these substrates should be reported. The mechanical properties of the substrates should be evaluated, as they might differ from batch to batch. Technically, I am wondering where the different background traction levels (see for example Figure 2, different traction force traces) or the traction measured outside a cell (see Figure 5A-C) arise from.

We now provide an explanatory figure that describes the traction force microscopy procedure (Figure 1—figure supplement 1A, E-H), as well as a detailed analysis of our bead density (~2.2 beads/µm2 or 0.42 ± 0.17 µm bead-to-bead spacing). Please see the new Figure 1—figure supplement 1B-D.

Every batch of substrates is evaluated prior to use, and substrates are only accepted if the stiffness falls within 10% of the nominal value.

We also discuss possible sources of the background traction. Conceptually, traction outside the cell may originate from non-transfected, invisible cells. However, this is rather unlikely, as cells were sorted with a flow cytometer prior to imaging. Alternatively, differences in background traction levels and traction forces outside of the cell are much more likely to result from imperfections in the image registration after removal of the cells: Per experiment ~15 cells were imaged for 10 minutes each. Once imaging was complete, all cells were removed by gently adding a dilute bleach solution that was preheated to 37 degrees Celsius, and the relaxed substrate was measured as rapidly as possible to avoid swelling of the substrate. Consequently, the entire experiment took ~2.5 hours, required careful fluid handling, and accurate positioning of the sample by the microscope. It is quite possible that certain parts of the substrate exhibit local deformations that are not captured by the registration procedure. Importantly, these spatial variations in traction force do not affect our local, temporal fluctuation analysis results.

2) Talin and vinculin are known to exhibit auto-inhibited conformations, in which an interaction between R8 and Vd1 may not be possible. Can the authors show in a more physiological setting (preferably full-length proteins) that talin and vinculin form a complex in solution?

We understand the reviewer’s concern that the R8-Vd1 interaction might not be possible in full-length talin and vinculin, especially in their autoinhibited state. However, it is not feasible to test the full length proteins in this experiment as there are 11 VBSs, and of these, the R3 VBSs are easy to activate (if we use Vd1 it can break open R3 as well). In an effort to support our argument, we now cite a recent study from the Ballestrem group. In this manuscript, they show that full-length talin does not associate with full-length vinculin unless the proteins are activated (Atherton et al., 2020). This, as well as thermodynamic principles, implies that a small population of talin or vinculin exists in an active conformation that is capable of forming a complex. In our manuscript, we speculate that such activation could potentially result from F-actin-driven force or vinculin’s PIP2 binding. While this possibility would be difficult to test experimentally, we have included this discussion in the text.

Additionally, in Figure 4C it seems that talin1 R8vvv can still bind vinculin, although not as proficient. Can the authors show the importance of vinculin binding to R8 for the maturation of NAs into FCs more directly by mutating only the vinculin binding site in R8?

While we agree that the R8vvv mutant can still bind vinculin, the equilibrium is shifted towards the unbound state (Figure 4C). Mutating the VBS alone in R8 is a good suggestion. Unfortunately, our attempts at making a mutant that prevents the VBS binding were not successful, i.e., all mutants tested did not abolish vinculin binding. The main challenge we faced in generating such a mutant was maintaining the structural integrity of the folded R8 domain, which is integral to ligand binding and must remain unperturbed. Indeed, these vinculin binding epitopes are buried in the folded form, and mutations appeared to easily ruin the fold. We attempted to alter the sizes of the hydrophobic epitope (swapping the F-A) but were not successful here either. We hope the reviewer understands these challenges.

3) The authors state that in maturing (G2) NAs vinculin and talin are recruited together. However, in Figure 2O it appears that in about 25% (and in Figure 7J ~50%) of cases, vinculin is recruited before talin. In Figure 7H it appears that vinculin is recruited to NAs 14 s earlier than talin. Could the authors elaborate how vinculin is recruited before talin is visible?

We appreciate the reviewer’s observation, and we revisited our data and looked closely at the trajectories where vinculin recruitment was identified earlier than talin. In these adhesions, we found that the talin signal was present but spatially diffuse and was thus not detected as a distinctive spot with our particle detection software. In contrast, although the vinculin signal was dim, it was spatially concentrated and detected by the software. Consequently, the detection of the early rise in talin was delayed relative to vinculin our analyses.

To evaluate if we could more accurately detect the onset of the diffuse talin signal, we relaxed the statistical criteria for particle detection from a P-value of.05 to.1 (e.g., 1 in 10 chance of it being observed due to chance). After reanalyzing the data, the number of events where vinculin arrived before talin was markedly decreased (see Figure 7N). In an effort to better communicate this observation, we updated Figure 7J and replaced Figure 7H with a new time series pair that represents a time-lag of 0 seconds, as now provide montages that follow adhesion assembly.

With regard to Figure 2O, we find that the larger pre-traction rise time for vinculin (than talin) is due to heterogeneity in traction background, which happens depending on multiple factors (e.g., image quality, bead density, and overall deformation, which can be all heterogeneous). In an effort to improve the quality of these data, we collected more samples from each condition and have analyzed the time of recruitment (updated in the new Figure 2O). It is worth noting that the negative portion of the 25% percentile is similar between talin and paxillin, and that the overall trend didn’t change with the new analysis.

4) How do the authors exclude the possibility that talin is quickly unfolded (as described in the legend of Figure 8) by forces from the cytoskeleton, which induces a fast recruitment of vinculin to talin in maturing NAs (which may be dependent on R8)? This process could be faster than the sampling rate and hence a pre-complex is not required to form in the cytosol.

We appreciate the alternative scenario that the reviewer has proposed. Indeed, we cannot formally exclude the possibility that vinculin recruitment is faster than our sampling rate. In part, this is due to the limitations of microscopy. Here, we had to image fast enough to properly detect and track adhesion complex formation, at low enough probe concentrations as not to perturb the specimen, with laser powers that did not noticeably introduce photobleaching or phototoxicity, all the while imaging long enough to watch the relatively slow process of adhesion maturation. Nonetheless, our observation is in agreement with previous work that used fluorescence fluctuation analysis to evaluate adhesion assembly, which is a method that provides a faster temporal sampling (see Bachir et al., 2014).

In contrast, testing the possibility by molecular perturbation might be informative if the effects of adhesion maturation are approximately equal. For example, such an experiment could include a double mutant R8vvv and ABS3 KO. Nonetheless, our data suggests this is not the case and gives overwhelming evidence that the key factor is R8 domain. Indeed, it is the only thing that can explain the differences observed in the IMCD talin null rescue experiments. Given that these experiments would require a significant amount of time and resources we chose not to pursue them (see also response 3-2 to Rev #1).

5) The authors show multiple traction force maps and images acquired for cells that show very different phenotypes although being the same cell line. In Figure 1A-F ChoK1 cells are shown that overexpress mGFP-tagged talin, vinculin or paxillin but have very different morphologies, sizes (d-f) and exhibit very different traction forces (a-c) where the scale is 0-300 Pa for talin, 0-1800 Pa for vinculin and 0-1000 Pa for paxillin. In Figure 2 it is similarly diverse where traction force scales range from 0-800 Pa (in h) to 0-4000 Pa (in a). What is the reason for this heterogeneity in morphologies and traction forces within the same cell line?

We understand the reviewer’s concern. However, we believe that these images reflect the natural heterogeneity of this cell line. Indeed, in an effort not to bias our results and to maximize the statistical power of our subsequent analyses, every cell encountered in the dish was imaged regardless of their morphology.

With regard to overall traction and cell area, we show these distributions in (Figure 1—figure supplement 2). It is worth noting that while there was slight variation in each protein-overexpression condition, no statistical difference in the spread area and overall traction was observed between the protein groups.

6) In maturing (G2) NAs, does the constantly increasing traction force arise from the accumulating more vinculin at the adhesion site (i.e. stronger connection to the actomyosin), or from the growth of the adhesion site (i.e. accumulating more integrins)? In Figure 2G, a continuous recruitment of talin is observed, which would argue for the increase in traction forces due to the recruitment of additional integrins. How would the authors explain the pre-formation of talin-vinculin complexes to be important for the recruitment of integrins to the adhesion site (in maturing NAs)?

We appreciate the reviewer’s suggestion of another potential mechanism for traction increase within G2 adhesions. We believe that both scenarios are plausible. As the reviewer noticed, the time series of talin in WT G2 adhesions steadily increase during traction development (shown in both Figures 2G and 6H), which can be proportional to the number of clustered integrins. If we compare the (original) Figure 7B,D to 7F,H, where talin and vinculin are imaged simultaneously, we observe the following:

1). For wild-type talin – talin’s intensity remains relatively constant in G2 adhesions (t=200-600 sec in Figure 7D), whereas vinculin’s signal increases significantly (t=200-600 sec in Figure 7H).

2). For the R8vvv mutant of talin – talin’s intensity shows a large increase in G2 adhesions (t=82-280 sec in Figure 7B), whereas vinculin’s signal increases marginally (t=82-280 sec in Figure 7F).

In Figure 6B, the traction of R8vvv G2 adhesion shows a plateau after 400 sec while talin signal was exponentially increasing, implying that inability to incorporate further vinculin could result in failure of further traction development. This gives more weight to the evidence that the precomplex formation has a major effect on vinculin recruitment.

7) For testing the effect of the R8vvv talin mutant in cells, the authors used shRNA to knock-down talin1 levels before rescuing cells with either WT or R8vvv talin1. How is the talin expression level of WT talin1 rescued cells compared to R8vvv rescued cells?

We sorted cells with a flow cytometer prior to imaging, and both the R8vvv and wild-type cells had similar expression profiles before and after sorting (see also response to Rev #1 comment 2).

We also evaluated talin1/2 double knockout IMCD cells (from Dr. Roy Zent) to investigate R8vvv’s effect in a cleaner manner (e.g., in the absence of residual talin1 expression, as well as talin2 expression). Indeed, the adhesion phenotype in these cells was in agreement with the ChoK1 cells. In particular, the R8vvv mutant showed significantly less adhesion maturation (Figure 5—figure supplement 4).

Further, an assessment of the morphology (whole cells and possible adhesion sites) of talin1 KD cells would be beneficial to appreciate the difference between R8vvv and WT talin 1. This would exclude the possibility that apparent differences between R8vvv and WT talin1 re-expressing cells arise from potentially different talin1 expression levels, a potential expression of talin2 in response to reduced talin1 levels or residual expression of WT talin1 in KD cells.

We reproduced our key findings from ChoK1 cells in talin1/2 double KO IMCD cells. Consequently, this should eliminate the concern that the differences observed between R8vvv and WT talin1 rescue cells are due to variable talin1 or talin2 backgrounds. As replied to reviewer #1, the expression levels of R8vvv and WT talin1 were controlled via judicious transfection/transduction conditions and by sorting cells with a flow cytometer prior to imaging. Likewise, as we previously stated, we believe that the morphology of the cells reflects the natural heterogeneity of the cell line, and that we intentionally chose to image all cells regardless of their morphology as to not bias our analysis.

8) The authors state that the talin-vinculin pre-complex formation is reduced in talin1-R8vvv expressing cells. However, talin1-R8vvv expressing cells show adhesion sites larger than NAs (Figure 5A). How do larger adhesion sites form in cells expressing talin1 R8vvv, if the vinculin binding to R8 is essential for NAs maturation? Representative image of labeled vinculin and talin within one cell (as analysed for Figure 7) would be beneficial to understand how the vinculin recruitment is affected by the R8vvv mutation.

This is a valid observation. To investigate the possibility that the large adhesions in talin1-R8vvvexpressing cells are driven by residual endogenous talin1, we investigated the adhesion pattern in talin1/2 double knock-out IMCD cells with talin-R8vvv expression. Consistent with the ChoK1 cells, these cells also show fewer maturing NAs than control cells, but also some large adhesions. There are two non-mutually exclusive explanations for this: (1) talin-R8vvv still can bind vinculin or (2) that vinculin-talin precomplex formation is not the only path for adhesion maturation. Indeed, in support of the former, we show evidence in Figure 4C that binding is reduced, but not abolished, for talin-R8vvv and vinculin. Nonetheless, given the complexity of cell adhesions, it seems reasonable to assume that secondary mechanisms for adhesion maturation also exist.

In order to better communicate how vinculin recruitment is affected by the R8vvv mutation, we now include representative images of vinculin and talin for both R8vvv and WT talin1 expressing cells (new Figure 7). In agreement with reduced talin-vinculin binding, R8vvv cells do have G2 adhesions, albeit less of them. Possibly, there is a compensation effect at play where less abundant G2 adhesions accumulate more talin. We also show that vinculin recruitment in those G2 adhesions is indistinguishable compared to one in G1 adhesions in R8vvv expressing cells.

9) In their very clear conclusive Figure 8 the authors depict different sites for paxillin in non-maturing vs. maturing NAs. Do the authors have experimental evidence for this?

The sites depicted are based on findings in the literature, not from our experimental findings. We now added question marks on every paxillin’s appearance in the figure and added our explanation in the figure legend with appropriate citations.

Further, the legend describes hypotheses, which can explain experimental data but experimental evidence from the reported data is limited. For example, that talin stretching is limited to a shorter level in non-maturing NAs and talin stretches faster in maturing NAs has, to my point of view, not been shown in the presented data.

We agree with this critique and have toned down our interpretation.

Reviewer #3:

Han et al. describe in great detail the molecular decision process that leads from transient nascent adhesions (NAs) forming constantly at the plasma membrane to focal adhesions (FAs) stably connecting the ECM with the actin cytoskeleton. The assembly kinetics of three GFP-tagged NA components (talin, vinculin and paxilin) is tracked down by TIRF microscopy and their behavior is correlated with local traction force determined by imaging the displacement of fluorescent beads in the hydrogel support, on which the cells grow. The authors apply machine learning algorithms to identify two classes of events both linked to force-assisted growth but with opposite fates. Using a new talin mutant, it is shown that binding of vinculin at a specific talin domain (R8) is required to establish the feed forward control for further vinculin recruitment and the steady build-up of local traction. In contrast, sequential binding of talin, vinculin and paxilin leads to maturation abort. I consider the study to be compelling, the technology pioneering and the results significant. Given that the topic is quite complex, the manuscript is of remarkable clarity. A direct comparison with R3 talin-mutants would add further evidence, however, such experiments are probably not within the scope of a revision. My listed points represent mainly comprehension questions that could encourage improvements of the text. Overall, I am in full support of publication.

1) Is there statistics on how many G2-class NAs actually mature into FAs?

We thank the reviewer for the support. Indeed, a direct comparison with the talin1 R3 IVVI mutant would be of significant interest. As the reviewer noted, this requires a substantial investment in time and resources. Especially with our reduced lab capacity, we believe that this may take ~1 year to achieve. As such, we believe it is beyond the scope of this publication.

2) "Event-based time series analysis" measures the appearance of fluorescently tagged proteins. What is the sensitivity of the method? How many mGFPs have to accumulate to detect a spot?

We appreciate the reviewer’s insightful questions. To evaluate the number of GFPs necessary for detection, we performed protein number and brightness quantification (Digman et al., 2008 Biophys J) on a single movie (i.e., paxillin-mGFP). Using such an approach, we find that the minimum number of mGFPs necessary to detect NAs is ~2 (see Author response image 1).

Author response image 1
Histogram of mGFP molecules in one paxillin-mGFP time lapse images.

Mean value was 1.7 with 1.08 standard deviation. Up to 16 mGFP molecules were able to be resolved.

3) At the stage of Figure 2, is the existence of an unstrained talin-vinculin pre-complex really a stringent conclusion? The time resolution of imaging is one second. During this time bin, proteins with a Kd in the μM range may undergo several binding and unbinding cycles. Likewise, the forces may exhibit fast fluctuations especially during the onset when only a small number of force transmitting proteins have been accumulated (see Figure 3B). Thus, there is plenty of "temporal" space for different scenarios of "what is first" that is technically inaccessible. Please comment.

We agree with the reviewer’s thoughts. During our imaging experiments, we carefully balanced our imaging duration, image acquisition rate, and illumination conditions, such that we could simultaneously resolve both fast (molecular recruitment) and slow (e.g., maturation) adhesion processes. At our purposefully minimal expression levels, a frame rate of 1 Hz was the fastest we could achieve without introducing photodamage. Consequently, we would not be able to resolve phenomena that occur on sub-second timescales. Nonetheless, our results are consistent with previous work that used very fast fluctuation-based imaging methods (see Bachir et al., 2014).

In order to communicate this limitation to the readers, we have reworded the text. For example, we use “concurrent localization” instead of “pre-complex formation” in the paragraphs up to Figure 3. Only after Figure 4, where we show a complex between talin R7R8 fragment and vinculin head, do we begin using the term “pre-complex”.

4) Figure 3B, the average force growth in the first 10 seconds may simply be zero due to the sensitivity limit of TFM. Or does this represent a finding?

We appreciate the reviewer’s insightful point. We have analyzed the background fluctuation of the traction and found that the traction growth in the first 10 seconds is not significantly different from the traction background fluctuation. We have updated this point.

5) Are G1-NAs programmed to terminate, because talin has too much time for erroneous interactions? Explicitly, do traction-dependent vinculin binding site (VBS), other than R8, represent "wrong sites" in the sense that cooperative vinculin assembly is even prevented?

We are grateful for this interesting interpretation regarding G1 NAs. We haven’t explicitly tested this hypothesis because of the complexity involved in the experimental design. Nonetheless, we now add a paragraph that outlines potential scenarios for G1 NAs in the Discussion.

6) At which stage of NA maturation then must force be sensed?

We suggest that the force should be sensed at the earliest stage of the NA assembly. We have made this point clear in the text describing the results in Figure 2O.

7) Is mutant R8vvv-talin totally impaired in vinculin binding (Figure 4D) or is there residual probability for exposing the VBS (Figure 4C)? The biochemical result is contradictory.

To resolve this contradiction, we repeated binding experiments with multiple variants of vinculin, including:

1) Vd1

2) A vinculin mutant (A50I) deficient for talin binding

3) A constitutively active vinculin mutant (T12)

4) Wild-type full-length vinculin.

Unfortunately, these experiments suffered from a high level of variation between repeats, and we were not able to identify the source of this variability. As such, we have chosen not to include them. Nonetheless, as we show in Figure 4C, the Vd1 domain of vinculin can bind the R8vvv mutant of talin, albeit to a lesser extent. For clarity, we have removed Figure 4D.

8) The difference between vinculin epitopes in domain R3 and R8: As a naïve reader one would assume that a VBS of the relaxed state locates at the surface of the protein. I consider it conceptually difficult to acknowledge that the VBS in domain R8 is active in an unstrained state but nevertheless requires unlocking of the threonine belt and partial unfolding of the α-helical bundle; maybe this warrants a sentence or two.

Both R3 and R8 have similar mechanical responses. For example, both domains are folded, but destabilized by their threonine belts. Consequently, although the VBS is not always accessible, the presence of the threonine belt shifts the equilibrium towards an accessible state.

From the literature, we know that full-length vinculin cannot bind to full-length talin unless one of the proteins is in a relaxed state (see Atherton et al.). As such, we hypothesize that a spontaneous conformational change (perhaps to an intermediate or semi-relaxed form) could release talin autoinhibition, expose the R8 VBS, and permit vinculin binding.

With regard to R3 in particular, the same study showed that talin lacking R2R3 is able to interact with both inactive and active forms of vinculin. Nonetheless, why vinculin cannot bind to the R3 domain in the absence of force is an open question in the field. One possibility is that RIAM, which binds to the R2R3 domain with a high affinity, stabilizes R2R3 in a conformation where the VBS is in a less accessible state. From our data, we have no reason to believe that such a masking interaction occurs on R8. In an effort to communicate these nuances to the reader, we added a paragraph in the Discussion on the role of R8 in VBS accessibility.

9) Cells expressing the R8vvv mutant are clearly impaired in NA maturations as documented by the cellular analysis (Figure 5). However, there are mature FAs. This may either relate to residual wildtype talin that could not be suppressed by shRNA (as the WB shows, Figure 5—figure supplement 1), residual binding of the mutant R8vvv-talin or even an undefined pathway into the cooperative vinculin assembly. These possibilities are not mentioned.

We appreciate the reviewer’s observation and potential interpretations, which were also raised by Reviewer 2. To investigate the possibility that the large adhesions in talin1-R8vvv-expressing cells result from an incomplete knock down of talin1, we performed rescue experiments in talin1/2 double knock-out IMCD cells. Importantly, rescue experiments in the IMCD cells phenocopied the experiments performed in ChoK1 cells. Specifically, cells rescued with the R8vvv mutant of talin1 have fewer maturing adhesions than cells rescued with WT talin1, but continue to display some large FAs (e.g., Figure 5J).

Another possibility is that these large FAs could be promoted by talin2. However, because the IMCD cells lacked talin2 expression but still created large FAs, this can be ruled out.

Thus, we think that residual binding on the mutant R8vvv-talin to vinculin is likely to explain these large FAs. As we show in Figure 4C, binding is reduced, but not abrogated, for talinR8vvv and vinculin. Nonetheless, given the complexity of cell adhesions, it seems reasonable to assume that secondary mechanisms for adhesion maturation may also exist. We now discuss these possibilities in the Discussion.

10) The traction growth rates in Figure 6K reach much more into the negative range than those in Figure 3B and 3C. Do negative values signify initial forces that disappear? In addition, there is quite a large difference in the scale of y-axis in Figure 6K (100-200 Pa/sec) as compared to Figure 3B and C (1-2 Pa/sec).

We have verified the value of the data and parameters that were used to quantify those data.

First, there was a trivial error in the y-axis labeling: The unit in Figure 6K was meant to be “Pa/min”, “Pa/sec” as indicated. The labels in Figure 6K and 3B,C are now consistent.

The differences in force growth rate between Figure 6K and Figure 3B-C can be easily explained. Specifically, in Figure 6K, we evaluated all adhesions (i.e., those in both G1 and G2). In contrast, in Figure 3B-C, we evaluated only “force-transmitting” adhesions. To avoid this confusion, we now present only the “force-transmitting” NAs in Figure 6k as well.

Of note, we chose to analyze the force-transmitting adhesions since this served as the best mechanism to identify de novo NA nucleation events. Furthermore, because we could align NA assembly to force onset, this allowed us to perform the time-delay analysis as shown in Figure 2M.

11) The body text referring to Figure 6 at the end of the paragraph says "These results suggest that the vinculin-talin pre-complex is essential for the development of force across NAs…". In my opinion this statement still remains an assumption, since I could not find direct evidence in Figure 6. The behavior of the mutant R8vvv-talin, if G2 characteristics are fulfilled, appear to be indistinguishable from the mGFP-tagged version described in Figure 2. The fact that there is a problem with force growth can only be explained in the light of Figure 7. Re-ordering these arguments may be considered.

We agree that the problem with force growth in G2 adhesions of R8vvv mutant cells can only be explained with evidence from Figure 7. As we observe the different force development in the G2 adhesions, we now ended the paragraph with a more direct summary of the observation.

12) Again, the two expression systems lead to a different range of values: The absolute vinculin assembly rates in Figure 7I as well as the rate increase between G2 vs G1 are much smaller than in Figure 3A. Is this due to different protein expression levels? Why were different integration times used (30 seconds in Figure 7I in contrast to 10 seconds in Figure 3A)?

In Figure 7, we used the amplitude of a Gaussian fit to approximate the experimental point spread function, which provided a noise-suppressed “signal” and allowed us to compare the two fluorescence images.

In Figures 2 and 3, where we compare fluorescence vs traction, we used the absolute intensity since it was enough to compare the relative difference.

To avoid the confusion, we now also report the amplitude of the Gaussian fits in Figures 2 and 3. We also apply the same “first-rate constant method” for Figure 7 as in Figure 3.

13) Is actin always engaged first and then the integrin tails? In other words, is there support for a "search mode" of a small number of competent actin-talin-vinculin-heads for integrin tails linked to the ECM (Figure 8, G2, first panel)?

No, we have no evidence that actin is always engaged first. Furthermore, we only have indirect evidence in the literature of actin engaging with the talin-vinculin complex. In an effort to communicate this graphically, we have added question marks on the assembly of the “actin” part in the cartoon Figure 8. We hope this clearly indicates that this part of the illustration is not supported by our data.

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

Essential revisions:

1) The conclusion that the concurrent recruitment of talin and vinculin to maturing NAs corresponds with the existence of a cytoplasmic precomplex of the two proteins. Although it is likely that such precomplexes form prior to NA recruitment, it has not been demonstrated in the current paper. Hence, it is necessary to tone this claim down in the title, Abstract, Result sections and Discussion of the manuscript.

We agree that our data does not provide direct evidence of a cytoplasmic precomplex, and we have sought to tone down this claim throughout the manuscript. In particular, we have replaced the word “precomplex” with “concurrent recruitment” or “force-free complex formation” in Abstract, Results, and Discussion. To generate a succinct summary of the main result in the title, we use the phrase “pre-complexation of talin and vinculin without tension”. This wording avoids the unproven notion that talin and vinculin come into the adhesion as precomplexes but emphasizes that the two molecules are recruited and interact, at least in adhesion sites, before tension develops. We hope that this is an appropriate tone-down.

2) Use same assembly rates (Figure 3A, Figure 6J, and Figure 7M) in y-axis. It is not clear what is meant by 1/min in Figures 3A/7M, and what is the rational for the different time frames analysed (Figure 3A: 2 min; Figure 6: 1 min; Figure 7: 0.5 min).

We thank the reviewer for catching the differences. The unit “1/min” was meant to be a first-order rate constant. However, to be consistent with our calculation of force growth rate, we decided to use the slope of the intensities. Accordingly, we changed the unit in Figures 3A and 7M to be “a.u./min” with correctly converted values. Importantly, these values are now comparable with Figure 6J.

For our analysis, we used a 10 second integration time for Figure 3A and a 20 second integration time for Figures 6J and 7M. In Figure 3A, we used a shorter integration time because it nicely demonstrated that vinculin had a substantially different association rate for non-maturing and maturing nascent adhesions. However, these results would not change markedly when analyzed with a 20 second integration time. In Figure 6J and 7M, we found that the longer integration time produced less variance in the calculated slopes. To better communicate these details, we have fixed the time information in the figure legends of figures 6J and 7M accordingly.

3) Although the authors state that the R8vvv mutation does not affect the talin recruitment rate, Figure 6J shows that in both G1 and G2 NAs talin R8vvv is recruited at higher rates than WT talin1. Test the rates for statistical significance (for G1 and G2 R8vvv vs. WT). In case talin1 R8vvv is indeed recruited faster to adhesion sites, the authors should try finding an explanation.

We thank the reviewer for noticing the potential difference in talin recruitment between R8vvv and WT talin. Indeed, there were statistical differences between the two conditions, in every combination of G1 and G2, which we now show in Figure 6J. Presumably the higher rate of R8vvv recruitment is consistent with the overall increased rate of NA nucleation. But we have no molecular explanation for this phenomenon. We tend to avoid grandiose speculations and thus would like to leave the presentation as is with the facts statistically pinned down. However, as can be seen in the adhesion trajectories, G1 adhesions under the R8vvv condition assemble fast in the first minute but also disassemble fast. Under these conditions, vinculin is not recruited significantly. To explain this to the readers, we added this explanation to the manuscript and now include an example (Figure 6—figure supplement 1).

4) The authors state that traction increase for talin1 R8vvv is reduced compared to WT talin1. The statistical test in Figure 6K to corroborate this statement is missing.

We added bars in Figure 6K that show statistical differences with asterisks.

5) Subsection “Simultaneous talin-vinculin imaging confirms vinculin’s recruitment after talin for non-maturing NAs and concurrent recruitment for maturing NAs”: should probably reference Figure 7N (instead of Figure 7J) and Figure 7O (instead of Figure 7K).

We corrected the figure panel and references accordingly. Thank you.

6) "Fluorescence fluctuation analysis" may be misleading as it usually refers to FCS, PCH, i.e., methods that involve a statistical correlation analysis. Here the authors directly analyze the time traces by thresholds and slopes.

We believe that this comment is valid, and have changed the wording to “event-detection time-series analysis”.

7) The density 2.2 beads/µm2 cannot be reproduced from average spacing 0.42 µm via the relation: density = 1/sqrt(spacing).

We used a formula, i.e., density = detected beads/area, to quantify the density, which was different from the relation given. However, we noticed that the given formula is more widely accepted. Thus, we changed the density accordingly and reflected in the manuscript and corresponding Figure 1—figure supplement 1.

8) "Both peptides bound…" This sentence sounds strange and may be changed to "Both peptides showed comparable affinity towards wildtype R7R8 and the R8R7vvv mutant, confirming.…"

We have adopted throughout the manuscript. Thank you.

9) "The threonine belt in talin R8 is responsible…" The wording is not clear since only the binding epitope itself can be responsible for binding and the threonine belt is, rather indirectly, regulating accessibility and hence the on-rate. Do these data suggest that RIAM and DLC1 use binding epitopes spatially distinct from the VBS within the R8 bundle? Or is RIAM and vinculin binding indeed competitive?

Thank you for pointing this out. We agree that the sentence was not sufficiently clear. Indeed, given that both RIAM and vinculin are capable of binding the VVV mutant form, the binding epitope remains unchanged. Instead, the VVV mutant has an altered bundle stability. In order to better communicate this, we have changed the wording of the corresponding sentence to: “Altogether, these biochemical characterizations suggested that the threonine belt in talin R8 is responsible for the increased accessibility of the VBS in R8 enhancing vinculin binding without force.” We hope this change clarifies the meaning.

https://doi.org/10.7554/eLife.66151.sa2

Article and author information

Author details

  1. Sangyoon J Han

    1. Lyda Hill Department of Bioinformatics, University of Texas Southwestern Medical Center, Dallas, United States
    2. Department of Biomedical Engineering, Michigan Technological University, Houghton, United States
    Contribution
    Conceptualization, Data curation, Software, Formal analysis, Funding acquisition, Validation, Investigation, Visualization, Methodology, Writing - original draft, Project administration, designed the experiments and performed TFM reconstruction, nascent adhesion analysis, and machine learning from the images
    For correspondence
    sjhan@mtu.edu
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0002-1384-665X
  2. Evgenia V Azarova

    Department of Cell Biology, University of Texas Southwestern Medical Center, Dallas, United States
    Contribution
    Investigation, Methodology, cultured IMCD talin1/2 KO cells and expressed WT talin1 and talin1 R8vvv vectors and performed imaging experiments with talin construct and its variations (e.g. R8vvv mutant)
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0002-3846-9176
  3. Austin J Whitewood

    School of Biosciences, University of Kent, Canterbury, United Kingdom
    Contribution
    Data curation, Validation, Investigation, Methodology, performed biochemical experiments and analyses
    Competing interests
    No competing interests declared
  4. Alexia Bachir

    Department of Cell Biology, University of Virginia, Charlottesville, United States
    Contribution
    Conceptualization, Investigation, performed imaging experiments for WT GFP-tagged vinculin and paxillin
    Competing interests
    No competing interests declared
  5. Edgar Guttierrez

    Department of Physics, University of California San Diego, San Diego, United States
    Contribution
    Resources
    Competing interests
    No competing interests declared
  6. Alex Groisman

    Department of Physics, University of California San Diego, San Diego, United States
    Contribution
    Resources, Provided high-refractive-index silicone substrates for traction force microscopy
    Competing interests
    No competing interests declared
  7. Alan R Horwitz

    Department of Cell Biology, University of Virginia, Charlottesville, United States
    Present address
    Allen Institute for Cell Science, Seattle, United States
    Contribution
    Conceptualization, Data curation, Funding acquisition, Methodology, Project administration
    Competing interests
    No competing interests declared
  8. Benjamin T Goult

    School of Biosciences, University of Kent, Canterbury, United Kingdom
    Contribution
    Conceptualization, Resources, Data curation, Formal analysis, Investigation, Methodology, Writing - review and editing, provided suggestions for talin structure and mutation and performed biochemical experiments and analyses
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0002-3438-2807
  9. Kevin M Dean

    Department of Cell Biology, University of Texas Southwestern Medical Center, Dallas, United States
    Contribution
    Data curation, Investigation, Methodology, Writing - review and editing, created ChoK1 cells with talin1 shRNA and talin1 R8vvv expression and performed imaging experiments with the cells
    For correspondence
    Kevin.Dean@UTsouthwestern.edu
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0003-0839-2320
  10. Gaudenz Danuser

    1. Lyda Hill Department of Bioinformatics, University of Texas Southwestern Medical Center, Dallas, United States
    2. Department of Cell Biology, University of Texas Southwestern Medical Center, Dallas, United States
    Contribution
    Conceptualization, Supervision, Funding acquisition, Investigation, Methodology, Project administration, Writing - review and editing
    For correspondence
    gaudenz.Danuser@utsouthwestern.edu
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0001-8583-2014

Funding

National Institutes of Health (R15GM135806)

  • Sangyoon J Han

National Institutes of Health (F32GM117793)

  • Kevin M Dean

National Institutes of Health (P01GM098412)

  • Alex Groisman
  • Alan Rick Horwitz
  • Gaudenz Danuser

National Institutes of Health (R35GM136428)

  • Gaudenz Danuser

Biotechnology and Biological Sciences Research Council (BB/N007336/1)

  • Ben Goult

Human Frontier Science Program (RGP00001/2016)

  • Ben Goult

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

Acknowledgements

We thank Roy Zent (Vanderbilt) and Olga Martha Viquez (Vanderbilt) for providing IMCD talin 1/2 double KO cells. We also thank Joseph Chi and Dana Reed for preparing DNA constructs and assisting with western blots, and Assaf Zaritsky (Ben Gurion University) for providing helpful comments about machine learning. This work was funded by the following grants: NIH R15GM135806 (SJH) NIH F32GM117793 (KMD), NIH P01GM098412 (AG, ARH, and GD), NIH R35GM136428 (GD), Biotechnology and Biological Sciences Research Council grant BB/N007336/1 (BTG) and a Human Frontier Science Program grant RGP00001/2016 (BTG).

Senior Editor

  1. Jonathan A Cooper, Fred Hutchinson Cancer Research Center, United States

Reviewing Editor

  1. Reinhard Fässler, Max Planck Institute of Biochemistry, Germany

Reviewers

  1. Reinhard Fässler, Max Planck Institute of Biochemistry, Germany
  2. Nico Strohmeyer
  3. Thomas Weidemann, Max Planck Institute of Biochemistry, Germany

Publication history

  1. Received: December 30, 2020
  2. Accepted: March 11, 2021
  3. Version of Record published: March 30, 2021 (version 1)

Copyright

© 2021, Han 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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