1. Cell Biology
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Molecular mechanism for direct actin force-sensing by α-catenin

  1. Lin Mei
  2. Santiago Espinosa de los Reyes
  3. Matthew J Reynolds
  4. Rachel Leicher
  5. Shixin Liu
  6. Gregory M Alushin  Is a corresponding author
  1. Laboratory of Structural Biophysics and Mechanobiology, The Rockefeller University, United States
  2. Tri-Institutional PhD Program in Chemical Biology, The Rockefeller University, United States
  3. Laboratory of Nanoscale Biophysics and Biochemistry, The Rockefeller University, United States
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Cite this article as: eLife 2020;9:e62514 doi: 10.7554/eLife.62514

Abstract

The actin cytoskeleton mediates mechanical coupling between cells and their tissue microenvironments. The architecture and composition of actin networks are modulated by force; however, it is unclear how interactions between actin filaments (F-actin) and associated proteins are mechanically regulated. Here we employ both optical trapping and biochemical reconstitution with myosin motor proteins to show single piconewton forces applied solely to F-actin enhance binding by the human version of the essential cell-cell adhesion protein αE-catenin but not its homolog vinculin. Cryo-electron microscopy structures of both proteins bound to F-actin reveal unique rearrangements that facilitate their flexible C-termini refolding to engage distinct interfaces. Truncating α-catenin’s C-terminus eliminates force-activated F-actin binding, and addition of this motif to vinculin confers force-activated binding, demonstrating that α-catenin’s C-terminus is a modular detector of F-actin tension. Our studies establish that piconewton force on F-actin can enhance partner binding, which we propose mechanically regulates cellular adhesion through α-catenin.

eLife digest

All of the cells in our bodies rely on cues from their surrounding environment to alter their behavior. As well sending each other chemical signals, such as hormones, cells can also detect pressure and physical forces applied by the cells around them. These physical interactions are coordinated by a network of proteins called the cytoskeleton, which provide the internal scaffold that maintains a cell’s shape. However, it is not well understood how forces transmitted through the cytoskeleton are converted into mechanical signals that control cell behavior.

The cytoskeleton is primarily made up protein filaments called actin, which are frequently under tension from external and internal forces that push and pull on the cell. Many proteins bind directly to actin, including adhesion proteins that allow the cell to ‘stick’ to its surroundings. One possibility is that when actin filaments feel tension, they convert this into a mechanical signal by altering how they bind to other proteins.

To test this theory, Mei et al. isolated and studied an adhesion protein called α-catenin which is known to interact with actin. This revealed that when tiny forces – similar to the amount cells experience in the body – were applied to actin filaments, this caused α-catenin and actin to bind together more strongly. However, applying the same level of physical force did not alter how well actin bound to a similar adhesion protein called vinculin. Further experiments showed that this was due to differences in a small, flexible region found on both proteins. Manipulating this region revealed that it helps α-catenin attach to actin when a force is present, and was thus named a ‘force detector’.

Proteins that bind to actin are essential in all animals, making it likely that force detectors are a common mechanism. Scientists can now use this discovery to identify and manipulate force detectors in other proteins across different cells and animals. This may help to develop drugs that target the mechanical signaling process, although this will require further understanding of how force detectors work at the molecular level.

Introduction

Cells probe and respond to the mechanical properties of their surroundings through cytoskeletal networks composed of actin filaments (F-actin), myosin motor proteins, and dozens of actin-binding proteins (ABPs). These networks transmit forces through cell-matrix focal adhesions (Humphrey et al., 2014) and cell-cell adherens junctions (Charras and Yap, 2018), plasma membrane-associated many-protein assemblies which serve as hubs for the conversion of mechanical cues and stimuli into biochemical signaling cascades (mechanotransduction). Defects in mechanotransduction are associated with numerous diseases (Jaalouk and Lammerding, 2009), including muscular dystrophies, cardiomyopathies, and metastatic cancer, yet therapeutics which specifically target these pathways are largely absent due to our ignorance of the mechanisms that transduce mechanical signals through the cytoskeleton.

Diverse force regimes modulate the polymerization dynamics, micron-scale architecture, and protein composition of actin assemblies by molecular mechanisms that remain unclear (Harris et al., 2018; Romet-Lemonne and Jégou, 2013). Cellular-scale pressures in the range of hundreds of pascal regulate the assembly and mechanical power of branched-actin networks generated by the ARP2/3 complex in vitro (Bieling et al., 2016) and in vivo (Mueller et al., 2017), tuning network geometric properties including filament length, density, and distribution of filament orientations. Live-cell imaging studies identified a subset of ABPs that preferentially localize to the cytoskeleton in vivo in response to this magnitude of mechanical stimulation (Schiffhauer et al., 2016), which were also postulated to recognize properties of network geometry. Molecular-scale forces in the piconewton range have also been shown to modulate the activity of F-actin polymerization (Courtemanche et al., 2013; Jégou et al., 2013; Risca et al., 2012; Zimmermann et al., 2017) and severing (Hayakawa et al., 2011) factors in vitro, suggesting that molecular components of actin networks could be mechanically regulated by physiological forces. While F-actin binding by the actin-depolymerization factor cofilin was reported to be directly regulated by tension across the filament to modulate its activity (Hayakawa et al., 2011), a recent study has suggested that cofilin is tension-insensitive, instead having its severing activity regulated by filament bending (Wioland et al., 2019). F-actin has been reported to adopt a structural landscape of co-existing conformations in cryo-electron microscopy (cryo-EM) studies (Galkin et al., 2010b), leading to speculation that actin filaments could themselves serve as tension sensors by presenting distinct binding interfaces to ABPs in the presence of load (Galkin et al., 2012). It nevertheless remains unclear to what extent mechanical modulation of functional interactions between ABPs and F-actin occurs through direct regulation of F-actin’s binding interactions by force. Furthermore, structural mechanisms enabling ABPs to detect force on F-actin, to our knowledge, are unknown.

Inspired by the report that the enhanced cytoskeletal localization response of ABP isoforms differed substantially when cells were mechanically stimulated (Schiffhauer et al., 2016), we reasoned that biophysical and structural analysis of closely-related ABPs would be a powerful approach for mechanistically dissecting mechanically regulated F-actin binding. Here we specifically investigate enhanced binding to F-actin when the load is applied solely to the filament, which we refer to as ‘force-activated binding’. Our studies focus on the homologous adhesion proteins α-catenin (Kobielak and Fuchs, 2004) and vinculin (Ziegler et al., 2006), which are major ABP components found in adherens junctions (α-catenin and vinculin) and focal adhesions (vinculin) that are critical for the force-dependent strengthening of cellular adhesion and mechanotransduction (Dumbauld et al., 2013; Yonemura et al., 2010). Vinculin is a strictly auto-inhibited globular protein that must engage with multiple adhesion partners to be activated and bind F-actin (Johnson and Craig, 1995), stabilizing adhesion through incompletely defined mechanisms. On the other hand, α-catenin exists in two distinct populations maintained in dynamic equilibrium in the cell (Drees et al., 2005). It serves as a central component of the membrane-anchored heterotrimeric α-catenin–β-catenin–cadherin complex (the ‘cadherin-catenin complex’) at adherens junctions, which lacks F-actin-binding activity in traditional assays when isolated (Yamada et al., 2005). It also forms a soluble homodimer with constitutive modest F-actin-binding activity, thought to play a role in the generation and maintenance of actin bundles by cross-linking filaments and inhibiting ARP2/3 binding, thereby suppressing branched-actin formation (Drees et al., 2005). The structural mechanisms by which forces transmitted through the actin cytoskeleton modulate the complex networks of binding interactions formed by α-catenin and vinculin during adhesion maturation remain unknown.

Both proteins are entirely α-helical (Bakolitsa et al., 2004; Rangarajan and Izard, 2013) and are composed of a large N-terminal ‘head’ domain, which engages in protein-protein interactions with other adhesion molecules (Kobielak and Fuchs, 2004; Ziegler et al., 2006) and a smaller C-terminal 5-helix bundle F-actin binding ‘tail’ domain (Bakolitsa et al., 1999; Ishiyama et al., 2013Figure 1—figure supplement 1, Helices H1–H5), connected by a flexible linker. The isolated actin-binding domains (ABDs), which we utilize in our study, retain their structures and actin-binding activities (Bakolitsa et al., 1999; Ishiyama et al., 2013), engaging a similar site on the filament surface (Hansen et al., 2013; Janssen et al., 2006; Kim et al., 2016; Thompson et al., 2014). Recent single-molecule force-spectroscopy studies reported that both α-catenin in the cadherin-catenin complex (Buckley et al., 2014) and vinculin (Huang et al., 2017) form catch bonds with F-actin, characterized by increased bond lifetime when moderate forces around 10 pN are applied across the ABP-actin interface. These studies provided one potential resolution to the apparent contradiction between biochemical studies demonstrating the cadherin-associated α-catenin population lacks F-actin binding activity (Drees et al., 2005) and cellular studies suggesting α-catenin plays a key role in linking adherens junctions to the cytoskeleton (Kobielak and Fuchs, 2004). However, in vivo both vinculin and α-catenin primarily engage contractile actomyosin bundles, whose component actin filaments are constitutively exposed to myosin-generated forces. This led us to hypothesize that force-activated binding to tensed F-actin could also be a key regulatory mechanism at adhesions, suitable for promoting the formation of initial attachments to actomyosin through membrane-anchored ABPs (e.g. the cadherin complex) before the ABP-F-actin interface itself coming under load. Furthermore, it could also serve as a mechanism to enrich soluble ABPs (e.g. the α-catenin dimer) whose binding geometry is fundamentally incompatible with catch-bond formation.

Here using simultaneous optical trapping and confocal fluorescence microscopy, we show that tension on the order of 1 pN across actin filaments directly enhances F-actin binding by human αE-catenin, but not vinculin. Utilizing a novel Total Internal Reflection Fluorescence (TIRF) microscopy in vitro reconstitution assay, we further show that physiological forces generated by myosin motor proteins activate α-catenin F-actin binding. Approximately 3 Å resolution cryo-EM structures of both proteins bound to F-actin (to our knowledge, the highest-resolution structures of ABP-F-actin complexes reported to date) reveal they share an overlapping major actin-binding site. However, they undergo markedly different rearrangements at their flexible N- and C-termini upon binding, resulting in distinct contacts mediated by their C-terminal extensions (CTEs) which re-fold on the actin surface. Truncating α-catenin’s CTE results in constitutive strong F-actin binding regardless of force, and a chimeric construct of vinculin featuring α-catenin’s CTE gains force-activated binding, demonstrating the α-catenin CTE to be a modular ‘force-detector’ which negatively regulates low-force binding. Together, our studies indicate piconewton force on F-actin can be discriminated by flexible elements in ABPs to mediate direct force-activated binding.

Results

Piconewton tension across individual actin filaments directly activates binding by α-catenin

To determine whether the actin-binding activity of vinculin or α-catenin could be regulated solely by force across F-actin, we performed correlative force and fluorescence measurements with a commercial instrument (Hashemi Shabestari et al., 2017) that combines dual-trap optical tweezers and confocal fluorescence microscopy (Figure 1A). In these experiments, Alexa-555 phalloidin-stabilized biotinylated actin filaments were captured from a laminar stream across a microfluidic flow-cell between two optically trapped streptavidin-coated beads. Tethered filaments were then transferred to a reservoir containing Halo-tagged ABD labeled with Alexa-488 and subjected to a constant velocity pulling protocol while simultaneously recording confocal movies of both the actin and ABD fluorescence signals.

Figure 1 with 2 supplements see all
Piconewton load across individual filaments activates α-catenin binding, while vinculin is force insensitive.

(A) Top: Geometry of the microfluidic chamber. Bottom: Cartoon of correlative single-filament force and fluorescence assay. (B and C) Representative single-filament constant velocity pulling experiments (0.1 µm/s) for vinculin ABD (B) and α-catenin ABD (C). Top: Force versus time plot. Red dots, raw data; black dots, force values binned to an interval of confocal frames. Middle: Montage of confocal frames of actin and ABD channels. Positions of beads are indicated with yellow circles (diameter, 4 μm). Bottom: IABD/Iactin ratio versus time plot. High-force: Average of five intensity ratios before actin breakage; low-force: average of five intensity ratios immediately before high-force average. Concentration of vinculin ABD or α-catenin ABD: 2 μM. (D and E) Paired analysis of low-force and high-force averages across trials for vinculin ABD (D) (N = 6, p=0.84) and α-catenin ABD (E) (N = 7, p=0.0156). Wilcoxon signed-rank test: *p<0.05; n.s. (not significant), p>0.05. (F and G) Scatter plots of normalized fluorescence ratios versus force. Statistical comparisons are between intensity ratios divided into two bins: force below and above 1 pN or 2.2 pN (dotted lines) for vinculin ABD (F) (1 pN: below: n = 62, above: n = 20, p=0.65; 2.2 pN: below: n = 72, above: n = 10, p=0.74), and force below and above 1 pN or 1.8 pN (dotted lines) for α-catenin ABD (G) (1pN: below: n = 53, above: n = 15, p<0.0001; 1.8 pN: below: n = 59, above: n = 9, p=0.0002). KS test: ****p<0.0001; ***p<0.001; n.s., p>0.05.

We frequently observed a monotonic increase in force once a threshold extension was reached, followed by an instantaneous return to baseline (Figure 1B,C; Figure 1—figure supplement 2A,B), consistent with resistance from a tether composed of one or more actin filaments extended beyond their resting length (visible as straightening in the actin fluorescence channel) followed by force-induced breakage. To correlate force with ABD binding, we calculated the background-subtracted ratio of the ABD fluorescence intensity to the actin intensity (which we refer to as ‘IABD/Iactin’) in each frame of the confocal movies (Materials and methods; Figure 1B,C, bottom), as well as the corresponding average force during frame acquisition (Figure 1B,C, top, black connected points). Apparent binding by the vinculin ABD fluctuated and did not change in response to load during individual pulling trajectories (Figure 1B), consistent with a previous report that mechanical stimulation does not promote cytoskeletal localization of vinculin in cells (Schiffhauer et al., 2016). However, we observed an apparent steady increase in binding along individual tethers by the α-catenin ABD (Figure 1C), consistent with force-activated binding. To quantify this phenomenon, due to the inherent fluctuations of individual IABD/Iactin traces in this assay (potentially due to dynamic changes in ABD binding density along individual tethers and instability in the focus during confocal imaging), we first focused our analysis on a subset of long-lived tethers which featured at least 10 frames. We examined the difference between a ‘high-force’ average, defined as the mean IABD/Iactin value from the five frames before the final tether rupture, and a ‘low-force’ average, defined as the mean IABD/Iactin value from the five preceding frames during each recording (Figure 1B,C). Paired analysis of low-force/high force averages (Materials and methods) showed a significant increase consistent with force-activated binding only by the α-catenin ABD (Figure 1D and E; Figure 1—figure supplement 2A).

Next, we sought to determine the impacts of force on ABD binding at the level of individual actin filaments. As our tether assembly procedure captures filaments from solution, in principle each tether could be composed of a single filament or multiple filaments. We thus elaborated the possible multi-filament tether configurations and designed a series of criteria to identify and exclude them from the analysis. The first plausible multi-filament tether configuration (1) is composed of multiple filaments of different lengths, all attached to the trapped beads at both ends (Figure 1—figure supplement 2B), which we identified by the presence of multiple breaking peaks in the force curve, generated as the shortest remaining filament in the tether reaches full extension then ruptures (Figure 1—figure supplement 2B). The next plausible configuration (2) is multiple filaments of different lengths, not all of which are attached at both ends (Figure 1—figure supplement 2C), which we identified by fluorescence line scans of the actin intensity, where we observe step-like reductions which we interpret to correspond to the ends of non-bridging filaments (Figure 1—figure supplement 2C). The third and final configuration (3), composed of multiple filaments, all of essentially identical length and attached at both ends (Figure 1—figure supplement 2D), is the most difficult to rigorously identify. We observed significant heterogeneity in the raw actin fluorescence intensity values between tethers (potentially due to slight differences in focus), with no significant difference between configuration one tethers and those featuring a single rupture force peak (data not shown), suggesting raw actin intensity cannot be used as a proxy for filament number. We thus analyzed the distribution of final breaking forces for all trials (pooling data collected in the presence of both the α-catenin and vinculin ABD), reasoning that configuration three tethers composed of multiple fully-extended actin filaments in parallel at the final break would generally rupture at higher forces. Consistent with this prediction, we observed a single major peak in the breaking force distribution centered at ~6 pN, which is likely to primarily consist of single filaments, as well as a long tail of ≥16.5 pN breaking forces, which we interpret to primarily encompass configuration three tethers. We thus additionally excluded all tethers with ≥16.5 pN breaking force from further analysis.

Based on these criteria, only 21 tethers (11 vinculin ABD and 10 α-catenin ABD) out of 183 total trials were included. This subset very likely predominantly contains tethers composed of single filaments. Consistent with the expected relative fragility of single-filament tethers with low breaking forces, most of these recordings only contained a limited number of frames, precluding quantification of force-dependent binding changes in individual tethers. We thus instead examined the relationship between force magnitude and ABD binding by pooling our recordings for each protein and plotting normalized IABD/Iactin versus force (Figure 1F,G). This analysis revealed no apparent correlation between vinculin ABD binding and force (Figure 1F). However, the α-catenin ABD plot showed an apparent step-like transition from force-uncorrelated binding to consistent strong binding above a threshold force of approximately 1 pN (Figure 1G).

To quantify this phenomenon, we initially examined the distribution of intensity ratios above and below the subjectively identified 1 pN threshold, finding a significant difference only for the α-catenin ABD (Figure 1F,G). To objectively define the threshold force, we performed K-means clustering analysis (Materials and methods) which revealed that the α-catenin ABD force-fluorescence distribution could be optimally divided into two clusters with a threshold force of 1.8 pN. We found that the intensity ratios were also significantly different between these clusters (Figure 1G). Although there was no obvious correlation between force and binding by the vinculin ABD, as a control we nevertheless used K-means to divide the data into two clusters, which separated at a threshold force of 2.2 pN. Consistent with the lack of a force-binding correlation for the vinculin ABD, we find no significant difference between the distribution of intensity ratios in these two clusters (Figure 1F). This analysis, which is insensitive to the exact force threshold employed, supports a force-dependent increase in F-actin binding only for the α-catenin ABD. Collectively, these data suggest that piconewton-level tensile force along individual actin filaments is sufficient to activate α-catenin’s F-actin binding.

Physiological forces generated by myosin motors activate F-actin binding by α-catenin

As ~1 pN is the magnitude of force generated by individual myosin motor domains (Finer et al., 1994), we hypothesized α-catenin’s F-actin binding would also be enhanced by physiological forces generated by myosins. To test this hypothesis, we developed a novel adaptation of the gliding filament assay (Kron and Spudich, 1986) to apply force to filaments mimicking actomyosin contractility in vivo (Figure 2A). In our preparation, plus-end directed myosin V motor proteins and minus-end directed myosin VI motor proteins are randomly surface-immobilized inside a flow chamber assembled on a cover glass for TIRF, resulting in a configuration where the motors are poised to engage in tug-of-war along non-stabilized, rhodamine-labeled actin filaments. A Halo-tagged, JF-646 (Grimm et al., 2015) labeled ABD is then flowed into the chamber in the absence of ATP, and a 2-color TIRF movie is recorded to visualize the basal level of actin binding when filaments are anchored by the rigor-state motors. The ABD is then re-introduced into the same chamber in the presence of ATP to activate the motors, and a second movie is recorded to visualize binding in the presence of force generation. Visual inspection of -ATP/+ATP TIRF movie pairs for the vinculin and α-catenin ABDs suggest they respond to motor-generated forces on F-actin distinctly. Actin localization of the vinculin ABD did not change in response to motor activity (Figure 2B; Figure 2—video 1); however, α-catenin ABD’s actin localization was enhanced upon motor activation, indicative of force-activated actin binding (Figure 2C; Figure 2—video 2).

Figure 2 with 9 supplements see all
Forces generated by myosin motors activate α-catenin binding to F-actin.

(A) Cartoon of TIRF force reconstitution assay utilizing surface-anchored myosin motor proteins with opposing directionality. (B and C) TIRF assays of vinculin ABD (B) and α-catenin ABD (C). Left: Representative movie frames of actin and ABD channels in the absence (top) and presence (bottom) of ATP to activate motors. Scale bar, 20 μm. Right: Montages of individual filament regions. Scale bar, 5 μm. Concentration of vinculin ABD or α-catenin ABD: 2 μM. (D and E) Time-averaged IABD/Iactin intensity ratio distributions of filament regions before and after ATP addition for vinculin ABD (D, quantification of (B)) and α-catenin ABD (E, quantification of (C)). (F and G) Paired analysis of the overall average intensity ratio change before and after ATP addition for vinculin ABD (F) (N = 9, p=0.50) and α-catenin ABD (G) (N = 6, p=0.031). Wilcoxon signed-rank test: *p<0.05; n.s. (not significant), p>0.05.

As we expected the random deposition and inherent stochasticity of molecular motors in our assay to give rise to a distribution of forces and ABD binding states, we implemented an image analysis procedure to quantify ABD association by automatically identifying and tracking tens to hundreds of ‘filament regions’ through time (Materials and methods, Figure 2—figure supplement 1A). For each region, IABD/Iactin was calculated in each frame, then averaged over all frames in which the region was detected. Consistent with our qualitative interpretation, histograms of IABD/Iactin distributions before and after ATP addition to individual flow chambers showed no shift for the vinculin ABD (Figure 2D, Figure 2—figure supplement 1B). However, this analysis demonstrated a clear shift toward higher values upon ATP addition for the α-catenin ABD, supporting force-activated binding (Figure 1E, Figure 2—figure supplement 1B). While the reported trends were consistent across experiments for both ABDs, we nevertheless observed variability between trials (Figure 2—figure supplement 1B), potentially due to differences in the background intensities in both channels resulting from inconsistencies in cover-glass surface preparation (Materials and methods). We thus performed paired analysis of the mean IABD/Iactin between the -ATP/+ATP conditions for each chamber (Figure 2F and G), which demonstrated a significant increase only for the α-catenin ABD (Figure 2G).

Our optical-trapping experiments suggest that force along individual filaments is sufficient to activate α-catenin binding. However, in the cellular context, both α-catenin and vinculin primarily engage actin-myosin bundles. In our TIRF assay, visual inspection supports increased F-actin bundling by both vinculin and α-catenin upon motor activation (Figure 2B and C), presumably due to motility facilitating encounters between filaments. Although IABD/Iactin measurements are internally normalized for the local density of F-actin in each region, we are aware that inter-filament ABD bundling contacts could in principle enhance apparent binding. Additionally, while the ATP-dependence of this enhanced F-actin binding strongly suggests it is activated by force, allosteric remodeling of actin filament structure due to local deformations imposed by motor binding could also potentially contribute (Gurel et al., 2017). To decouple these effects, we performed assays in the presence of individual motors. In the presence of both myosin V alone (Figure 2—figure supplement 2, Figure 2—video 3 and Figure 2—video 4) and myosin VI alone (Figure 2—figure supplement 2, Figure 2—video 5 and Figure 2—video 6) we observe ATP-dependent formation of bundles. However, we observe no significant increase in apparent vinculin ABD or α-catenin ABD binding in either condition. This strongly suggests that force-activated α-catenin ABD binding is dependent upon the tug-of-war between motors of opposed directionality, mimicking the forces generated by bi-polar myosin II filaments in vivo.

We note that forces generated by the randomly distributed force generators in the dual-motor assay are complex, and can in principle include tension, compression, and torsional forces (Beausang et al., 2008; Sun et al., 2007). While our optical trapping studies suggest tensile forces are sufficient to activate α-catenin ABD binding (Figure 1), future studies will be required to explicitly dissect the contribution of compression and torsion in the presence of myosin motors.

Additionally, we note that there are caveats associated with our TIRF experiments. First, the relatively low apparent affinity of our α-catenin ABD construct in this assay necessitates utilizing a high working concentration. The commensurate high fluorescence background is refractory to definitively measuring the intensity level constituting F-actin binding saturation in the absence or presence of force generation. This limits our ability to establish the binding stoichiometry range in which force-activated actin binding occurs, which has the potential to constrain or discriminate between plausible molecular mechanisms (see Discussion). In contemporary studies, Xu and colleagues have reported a series of α-catenin ABD N-terminal deletion constructs with substantially increased F-actin affinity in the absence of force, notably a construct consisting of residues 698–906, which boosts affinity ~18-fold (Xu et al., 2020). Presuming this construct maintains force-activated binding activity, it may be a useful tool to dissect the impact of α-catenin’s binding stoichiometry. Second, due to the strong propensity of the α-catenin ABD to bundle actin filaments in our TIRF experiments, it remains to be conclusively determined if myosin motor activity can also activate α-catenin binding along single filaments. It may be possible to achieve a sufficiently low surface density of actin filaments to avoid bundling in the dual-motor assay in presence of the actin stabilizing drug phalloidin, which would enable probing α-catenin’s association with individual actin filaments in the presence of active force generation. We will pursue these studies, and their outcome will be presented in a follow-up report.

Regardless, the data we present here collectively suggest physiological forces generated by myosin motor proteins in an appropriate configuration can directly activate α-catenin binding to F-actin, and that force-activated α-catenin binding also occurs in the context of actin bundles, the primary cytoskeletal architecture engaged by the protein in vivo.

Cryo-EM structures of the metavinculin ABD-F-actin and α-catenin ABD-F-actin complexes

Hypothesizing that differences in the F-actin-binding interfaces of vinculin and α-catenin could underlie their differential force-activated actin binding, we pursued structural studies of both ABDs bound to F-actin with cryo-EM (Figure 3; Figure 3—figure supplement 1; Table 1). As optimizing the density of fully-decorated, well-separated individual filaments in cryo-EM images is a major bottleneck for single-particle analysis of F-actin-ABP complexes, we chose to use the ABD of the vinculin splice variant metavinculin for these studies, where a 68 amino-acid insert displaces the H1 helix and replaces it with helix H1’, producing a protein which retains actin binding but completely loses actin bundling activity (Janssen et al., 2012; Kim et al., 2016; Oztug Durer et al., 2015Figure 1—figure supplement 1B). Previous studies have suggested these isoforms engage an essentially identical site on the F-actin surface with equivalent affinity (Janssen et al., 2012; Kim et al., 2016), and we further found the metavinculin ABD lacks force-activated actin-binding activity in our TIRF assay, validating its use in these studies (Figure 2—figure supplement 1C,D). We were able to obtain fields of individual decorated filaments using this construct (Figure 3A, top). After careful optimization (Materials and methods), we were also able to acquire cryo-EM images of filaments decorated with the α-catenin ABD (Figure 3A, bottom), although persistent bundling by this construct necessitated the collection of substantially more images to obtain a sufficient dataset of individual segments to obtain a high-resolution reconstruction (Table 1; Figure 3—figure supplement 1).

Figure 3 with 1 supplement see all
Cryo-EM structures of metavinculin and α-catenin–F-actin complexes.

(A) Representative cryo-EM micrographs of F-actin decorated with metavinculin ABD (top) and α-catenin ABD (bottom). Scale bars, 100 nm. (B) Segmented regions of reconstructed density maps of F-actin decorated with metavinculin ABD (left) and α-catenin ABD (right), colored as indicated. (C) Overlay of the metavinculin ABD-actin complex and α-catenin ABD-actin complex atomic models, superimposed on actin and colored as in (B). Actin subunits from the α-catenin structure are displayed in varying shades.

Table 1
Cryo-EM data collection, refinement, and validation statistics.
Metavinculin ABD‒F-actin
(EMDB-20844, PDB 6UPW)
α-catenin ABD‒F-actin
(EMDB-20843, PDB 6UPV)
Data collection and processing
MicroscopeTitan KriosTitan Krios
Voltage (kV)300300
DetectorK2 SummitK2 Summit
Magnification29,00029,000
Electron exposure (e2)6060
Exposure rate (e/pixel/ s)66
Calibrated pixel size (Å)1.031.03
Defocus range (µm)‒1.5 to ‒3.5‒1.5 to ‒3.5
Helical symmetryC1C1
27.06 Å rise27.03 Å rise
−167.07° twist−166.88° twist
Initial filament segments (no.)237,503540,533
Final filament segments (no.)215,369414,486
Map resolution (Å)2.943.24
FSC threshold0.1430.143
Refinement
Initial model (PDB ID)3JBK, 3J8A4IGG, 3J8A
Model resolution (Å)2.983.28
FSC threshold0.50.5
Map sharpening B factor (Å2)‒58‒85
Model composition5 actin protomers, 2 actin-binding domains
Non-hydrogen atoms16,89617,233
Protein residues21522201
Ligands5 Mg.ADP
B factors (Å2)
Protein53.4171.59
Ligand47.2956.65
R.M.S. deviations
Bond lengths (Å)0.0120.012
Bond angles (°)0.8600.833
Validation
MolProbity score1.231.33
Clash score3.324.63
Poor rotamers (%)0.060.05
Ramachandran plot
Favored (%)97.4697.56
Allowed (%)2.542.44
Disallowed (%)0.000.00
EMRinger Score4.403.87

Using the Iterative Helical Real Space Reconstruction (IHRSR) approach (Egelman, 2007) as implemented in Relion 3.0 (He and Scheres, 2017; Zivanov et al., 2018; Materials and methods, Figure 3—figure supplement 1), we obtained reconstructions of the metavinculin ABD (residues 879–1134)–F-actin complex (Figure 3B, left) at 2.9 Å overall resolution (Figure 3—figure supplement 1) and the α-catenin ABD (residues 664–906)–F-actin complex (Figure 3B, right) at 3.2 Å overall resolution (Figure 3—figure supplement 1). As local resolutions ranged from 2.7 Å to 3.6 Å, radially decaying outward from the helical axis (Figure 3—figure supplement 1C,D) for both reconstructions, atomic models for the complete sequence of Mg-ADP α-actin and continuous segments of ABD residues 981–1131 for metavinculin (Figure 3C) and 699–871 for α-catenin (Figure 3C) were built and refined into the maps (Figure 3—figure supplement 1). In their contemporary work, Xu and colleagues also reported a 3.6 Å resolution cryo-EM reconstruction of the α-catenin ABD bound to F-actin (Xu et al., 2020), which shows a highly similar conformation of the complex to that presented here (actin, 0.5 Å RMSD; α-catenin residues 711–842, corresponding to helices H2–H5, 1.0 Å RMSD).

Superposition of the actin-bound metavinculin ABD with the full-length vinculin crystal structure (Bakolitsa et al., 2004) confirms that actin binding by both vinculin isoforms is auto-inhibited by intramolecular interactions between the N-terminal head and C-terminal ABD tail domains (Johnson and Craig, 1995Figure 4—figure supplement 1A,B), as the head domain clearly clashes with F-actin in the crystallized conformation. Full-length α-catenin was crystallized as an asymmetric ‘left-handshake dimer’, characterized by differential relative orientations between the head and tail domains of each protomer (Rangarajan and Izard, 2013). Comparing the actin-bound α-catenin ABD with both conformers in the asymmetric dimer crystal structure also reveals severe clashes between the α-catenin head and actin (Figure 4—figure supplement 1C–F), suggesting that full-length α-catenin must also undergo substantial conformational rearrangements to bind F-actin.

Next, we compared the metavinculin ABD-F-actin and α-catenin ABD-F-actin structures (Figure 3C), confirming previous low- and moderate-resolution studies (Janssen et al., 2006; Janssen et al., 2012; Kim et al., 2016; Thompson et al., 2014) that both ABDs engage a major site spanning the longitudinal interface of 2 actin protomers, which we term Actin I and Actin II (numbered from the plus end of the filament). In turn, each actin protomer also contacts 2 ABDs, leading to a 1:1 binding stoichiometry at saturation. Our structures establish this region is almost identical between the two ABDs, comprising 2040 Å2 of buried surface area for the metavinculin ABD and 1920 Å2 for the α-catenin ABD. However, our high-resolution models also reveal a previously unobserved minor interface for each ABD (confirming a recent computational prediction in the case of metavinculin [Krokhotin et al., 2019]), mediated by residues in their flexible C-terminal extensions (CTEs), which are entirely distinct between the two proteins (Figure 1—figure supplement 1A).

Differential conformational remodeling is linked to unique features of actin binding

Consistent with our previous medium-resolution structural studies (Kim et al., 2016), we find the metavinculin ABD undergoes substantial conformational remodeling upon F-actin engagement, characterized by displacement of helix H1’ from the 5-helix bundle to license a rearrangement of helices H2–H5 to relieve clashes with F-actin (Figure 4A,B; Figure 4—video 1 and Figure 4—video 2). N-terminal residues 879–980 are not visible in the map (Figure 4A, transparent brown), and are presumably disordered in the actin-bound state, while residues 981–985 (Figure 4A, brown) undergo a slight rearrangement, extending helix H2 by 1.5 turns (five residues). Our high-resolution map reveals this contact to be mediated by the (meta)vinculin CTE (Figure 4). The CTE is released from its pre-bound position, extending helix H5 by two turns (six residues), then undergoing an approximately 60° swing to engage a site along actin subdomain one proximal to H5 (Figure 4A and B; Figure 5—video 1). Coupled to this transition, helices H2–H5 slightly rearrange to accommodate actin binding and avoid steric clashes (Figure 4B and C; Figure 4—video 2).

Figure 4 with 3 supplements see all
Major ABD conformational rearrangements upon actin binding.

(A) Cartoons and crystal structures of pre-bound (meta)vinculin ABD (PDB 1st6, dark orange, top) and α-catenin ABD (PDB 4igg chain B, dark pink, bottom). Regions that undergo major rearrangements are highlighted in brown (metavinculin) or magenta (α-catenin); regions not resolved in the actin-bound cryo-EM structures are transparent. (B) Cryo-EM structures of metavinculin ABD (orange, top) and α-catenin ABD (pink, bottom). Flexible regions colored as in (A); actin, shades of blue. (C) Superimposed pre-bound and post-bound structures of metavinculin ABD (top) and α-catenin ABD (bottom). Green arrows indicate displacement of helices; rotation angles indicate repositioning of C-terminal extensions (CTEs).

We find that the α-catenin ABD also undergoes an order-to-disorder transition at its N-terminus upon actin binding (Figure 4A,B; Figure 4—video 1), as no density for residues 664–698, the majority of H0-H1, is present in our map (Figure 3D; Figure 4A,B; Figure 4—video 1), confirming recent reports that this region is important for activating α-catenin’s actin engagement (Ishiyama et al., 2018; Xu et al., 2020). This is accompanied by a twisting rearrangement of helices H2–H4 reminiscent of that found in (meta)vinculin (Figure 4B; Figure 4—video 2), as well as the extension of H4 by three turns (nine residues) through folding of the H3-H4 loop, to sculpt a major actin-binding interface sterically compatible with the filament (Figure 4B; Figure 4—video 1). This suggests that N-terminal helix release allosterically coupled to ABD helical-bundle rearrangement is a fundamentally conserved mechanistic feature of actin binding by members of the vinculin/α-catenin family. However, we observe distinct rearrangements in the α-catenin CTE, which undergoes a slight lateral shift and helical unfurling, rather than a swing, to engage a site spanning a different surface of actin subdomain 1 (Figure 4A and B; Figure 5—video 1).

Metavinculin H5 extension is facilitated by binding interactions with Actin I (Figure 5A,B; Figure 5—video 1), notably a hydrophobic interaction between metavinculin I1114 and Actin Y91, and a salt bridge between metavinculin R1117 and actin E100. This positions the CTE to form an extended interface with actin subdomain 1, contiguous with that mediated by H5, with metavinculin W1126 buried in a proximal hydrophobic pocket formed by actin residues A7, P102, P130, A131, and W356, bolstered by a distal salt bridge between metavinculin R1128 and actin E361 (Figure 5B). By contrast, the α-catenin CTE retains an overall conformation similar to its pre-bound state (Figure 5A and C; Figure 5—video 1). An extensive hydrophobic network we term a ‘tryptophan latch’ embraces CTE residue W859 in both pre-bound and post-bound states, preventing α-catenin CTE unfurling (Figure 5C, right; Figure 5—video 1). A single turn of helix H1 on the N-terminal side of the ABD remains folded, with H1 residue W705 packing against CTE residue M861, encircling W859 along with CTE residues L852 and L854, as well as residues W705, I712, I763, L776, P768, V833, and Y837 from neighboring regions of the helical bundle, facilitating coordinated conformational transitions between the N- and C-terminal flexible regions of the α-catenin ABD upon actin binding. A putative hydrogen bond is also maintained between S840 and the single nitrogen atom in W859’s indole ring, maximizing the binding potential of this residue.

Figure 5 with 2 supplements see all
The actin-binding interfaces of metavinculin and α-catenin.

(A) Overlay of the metavinculin ABD-actin complex and α-catenin ABD-actin complex atomic models highlighting C-terminal extensions (CTEs) of metavinculin ABD and α-catenin ABD, superimposed on Actin I and colored as in Figure 4. Actins from α-catenin structure are displayed. (B and C) Detailed views of key contacts at minor interfaces: metavinculin CTE and actin (B); within the α-catenin tryptophan latch (right) and between its CTE and actin (left) (C). (D) Overlay of the metavinculin ABD-actin complex and α-catenin ABD-actin complex atomic models highlighting helical binding interfaces of metavinculin ABD and α-catenin ABD, superimposed and colored as in (A). Actins from α-catenin structure are displayed. (E, F, and G) Detailed views of key contacts at the major interface between metavinculin/α-catenin helices H4–H5 and Actin I (E); Actin II (F); Actin II D-loop (G). (H) Actin Cα traces colored by per-residue RMSD from the indicated comparisons. For superposition, segmented actin density from the cryo-EM maps was first aligned, followed by fitting the atomic models into their corresponding maps.

The latch positions the neighboring region of the CTE to bind a distinct site on Actin I’s subdomain 1 (Figure 5C, left; Figure 5—video 1) mediated by proximal salt bridges (α-catenin E865 – actin R28, α-catenin K866 – actin D24/D25, α-catenin K867 – actin E93) and distal hydrophobic interactions (α-catenin L869/V870 – actin P333/Y337). Consistent with a key role for the latch in coordinating conformational transitions that enable F-actin binding, Xu et al. report that mutating W859 to alanine reduces α-catenin’s F-actin-binding affinity 10-fold (Xu et al., 2020). Superposition of the actin-bound conformation of the α-catenin ABD with the pre-bound conformation of the (meta)vinculin ABD (Figure 4—figure supplement 1G) reveals a striking positional overlap between α-catenin W859 and metavinculin W1126 (vinculin W1058 is identically positioned, not shown), which is also engaged by a sparser latch in the pre-bound conformation (Figure 5—video 1). We thus speculate the extensive latch of α-catenin prevents W859 release and the extension of its CTE to engage the same site as metavinculin W1126.

Major filament binding interfaces discriminate actin D-loop conformations

The complete non-overlap of the (meta)vinculin and α-catenin minor actin-binding interfaces mediated by their CTEs lead us to hypothesize that the CTEs could be involved in differential force-activated actin binding. To identify other potential contributing structural elements, we undertook a detailed comparison of their major actin-binding interfaces mediated by helices H4–H5 in both proteins (Figure 5D). The Actin II binding interface is almost identical between the two ABDs (Figure 5E), mediated by an extensive series of conserved hydrophobic contacts: α-catenin I792/metavinculin I1065 – actin I345, α-catenin V796/metavinculin V1069 – actin I341, and α-catenin V800/metavinculin M1073 – actin P333/E334. The Actin I interface, on the other hand, is more variable and characterized by few clear residue-level binding interactions (Figure 5F), notably likely weak long-distance salt bridges (metavinculin R1044 – actin E83 and α-catenin K845 – actin E99) specific to each protein, despite the overall shape complementarity across the interface. Each ABD also features a unique hydrophobic interaction with actin Y91 (metavinculin I1114/α-catenin V838).

The Actin I interface extends into contacts with the actin D-loop (Figure 5G), a flexible region of actin which mediates structurally polymorphic longitudinal interactions between protomers (Galkin et al., 2010b) reported to be modulated by actin nucleotide state (Chou and Pollard, 2019; Merino et al., 2018) and ABPs (Dominguez and Holmes, 2011; Oda et al., 2019). Both ABDs form a potential weak long-distance interaction with actin D-loop residue K50: in metavinculin, a hydrogen bond through N1048, and in α-catenin, a salt bridge through D775. The D-loop then adopts subtly different conformations between the two interfaces centered at M47. Although clear residue-level binding interactions are not readily apparent, the conformation of M47 at the metavinculin interface would clash with α-catenin Y786, a position occupied by the smaller residue I1059 in metavinculin, suggesting local sterics unique to each ABD determine compatibility with a distinct D-loop conformation. Comparison of the actin conformation observed in a similar-resolution structure of ADP F-actin in isolation (‘F-actin alone’, M.S. in preparation) versus when bound to metavinculin or α-catenin, as well as comparison of the metavinculin-bound and α-catenin-bound conformations reveals minimal rearrangements throughout the majority of the structure (Figure 5H). This contrasts with a previous report of α-catenin-induced structural changes in F-actin inferred from low-resolution cryo-EM analysis (Hansen et al., 2013), but it is consistent with the high-resolution studies of Xu et al., 2020. The sole region featuring rearrangements greater than 1 Å RMSD is a 3–4 residue stretch of the D-loop centered on M47. As force across the filament could feasibly modulate D-loop structure to regulate ABP binding, we hypothesized ABD residues mediating D-loop interactions could also mediate differential force-activated actin binding.

The α-catenin C-terminal extension is a force-detector element

To investigate whether D-loop interactions contribute to α-catenin force-activated binding, we designed a triple point-mutant α-catenin ABD construct where three residues in close proximity to the D-loop were replaced by those in vinculin: α-cat ABDA778Q Y779V Y786I. In force reconstitution assays, this construct did not visibly associate with actin in either the –ATP or +ATP condition in the concentration regime accessible by TIRF (Figure 5—figure supplement 1A). While these data suggest that the α-catenin D-loop interactions contribute to overall affinity for F-actin, the complete lack of binding is refractory to determining whether this interface has a separable role in force-activated actin binding.

We thus returned to our initial hypothesis that differential force-activated binding could be mediated by the CTEs. Although we were unable to accurately model the final three residues of the metavinculin CTE, weak density is clearly present (Figure 5—figure supplement 1B, red), suggesting the entire CTE engages F-actin. By contrast, density for the α-catenin CTE is only present until K871 (Figure 5C; Figure 5—figure supplement 1C). Notably, all three human α-catenin isoforms have highly homologous CTEs that extend an additional 35 amino acids (Figure 1—figure supplement 1A), diverging in sequence and length from the vinculin CTE. Consistent with previous primary-structure-function analysis (Pokutta et al., 2002) showing that residues after P864, which bear no homology to vinculin, are necessary for actin binding, our structure shows that residues 865–871 are in direct contact with actin (Figure 5C; Figure 5—video 1), forming an extensive interface. We therefore hypothesized that distal residues 872–906, a 35-residue element unique to α-catenin (Figure 1—figure supplement 1A) that was not resolved in our cryo-EM analysis and is thus presumably conformationally flexible, could uniquely contribute to force-activated actin binding.

To test whether distal residues 872–906 have a separable role in force-activated binding, that is as a ‘force detector’, we first truncated them from the α-catenin ABD (α-cat ABDΔC). Consistent with a regulatory role for this segment, α-cat ABDΔC constitutively associated with F-actin in TIRF assays (Figure 6A; Figure 6—figure supplement 1A; Figure 6—video 1), with no significant increase in binding upon ATP addition, suggesting that this region is necessary for force-activated binding by negatively-regulating low-force binding. This contrasts with the observations of Xu et al., who report a modest (~2-fold) reduction in F-actin-binding affinity when this region is truncated in solution co-sedimentation assays, notably in the background of a construct where H0 has also been truncated in order to boost affinity overall (Xu et al., 2020). Possible sources of this discrepancy include differences in α-catenin binding behavior between solution assays and our TIRF assays, where filaments are immobilized. Additionally, coordination between rearrangements in the CTE and H0–H1 (potentially through allosteric mechanisms) may be necessary to mediate the force-detector’s negative regulatory effects.

Figure 6 with 4 supplements see all
The distal tip of α-catenin’s C-terminus is a force detector.

(A, B, and C) TIRF force reconstitution assays. Left: Representative movie frames in the presence and absence of ATP. Scale bar, 30 μm. Right: Cartoon of ABD construct (left) and paired analysis of the overall intensity ratio change upon ATP addition (right). Wilcoxon signed-rank test: *p<0.05; n.s. (not significant), p>0.05. Constructs assayed were: α-cat ABDΔC (A) (αE-catenin664-871, N = 5, p=0.81); Vinc ABD-NCSwap (B) (αE-catenin664-708-vinculin916-1041-αE-catenin837-906, N = 6, p=0.031); Vinc ABD-NCSwapΔC (C) (αE-catenin664-708-vinculin916-1041-αE-catenin837-871, N = 5, p=0.63). Concentration of ABD constructs: 2 μM.

To investigate the sufficiency of the force-detector, we generated chimeric ABDs featuring the H2–H5 bundle region of vinculin and the flexible termini of α-catenin. A vinculin ABD construct where only the CTE was substituted was non-functional (data not shown). However, consistent with structural coordination between the α-catenin N-terminal segment and the CTE through the tryptophan latch (Figure 5C; Figure 5—video 1), a construct featuring both the α-catenin N-terminal segment and the CTE (vinc ABD-NCSwap, Materials and methods) gained force-activated binding activity, with diminished low-force binding observed (Figure 6B; Figure 6—figure supplement 1B; Figure 6—video 2) in contrast to the wild-type vinculin ABD (Figure 1B). A C-terminal truncation of this construct (vinc ABD-NCSwapΔC) equivalent to α-cat ABDΔC reverted to constitutive binding regardless of force (Figure 6C; Figure 6—figure supplement 1C; Figure 6—video 3), supporting the α-catenin CTE as the key determinant of force-activated binding, in which 872–906 serves as the force-detector. We thus conclude that the distal C-terminus (residues 872–906) of α-catenin is necessary and sufficient for force-activated actin binding through negative regulation of low-force binding.

Discussion

Our studies reveal that α-catenin preferentially binds tensed F-actin, using its flexible C-terminus to detect piconewton load that can be generated by myosin motors. We propose this force-activated F-actin binding could play an important role in the formation and reinforcement of cell-cell adhesion complexes, facilitating mechanically regulated interactions between α-catenin and actomyosin cables tuned for high sensitivity to motor-generated forces.

We further provide evidence that actin filaments are themselves tension sensors that can transduce mechanical force into regulated partner binding. Our finding that the force-detector element of α-catenin can be functionally transplanted to vinculin demonstrates that force-activated F-actin binding is a modular activity that can be conferred by short sequence motifs. We thus speculate there may be other actin-binding proteins with functionally analogous force-detectors, which could broadly mediate mechanotransduction through the cytoskeleton.

Molecular mechanism for detecting force on actin through a flexible structural element

While our studies pinpoint the final 35 amino acids of α-catenin as a force-detector, the exact molecular mechanism by which this segment negatively regulates low-force binding to F-actin remains to be elucidated. Here we propose two potential, non-exclusive conceptual models for this modulation. As we observe the distal tip of the ordered region of the CTE to be in close apposition to the next ABD binding site along the actin filament, with potential contacts between CTE residues V870 and K871 with the H4–H5 loop and the N-terminal tip of H5 in longitudinally adjacent ABD (Figure 5—figure supplement 1C), the first model invokes steric exclusion (Figure 7A). In the absence of force, the force-detector (Figure 6; Figure 7) physically blocks the adjacent binding site through steric hindrance, which can be relieved by an increase in protomer axial spacing in the presence of tension, consistent with prior truncation studies suggesting residues 884–906 may inhibit α-catenin’s actin binding (Pappas and Rimm, 2006). As the force-detector likely represents a flexibly tethered conformational ensemble, in the presence of thermal fluctuations we envision this would manifest as a tension-dependent increase of the binding on-rate at the site due to its increased fractional availability. Although saturating the filament for structural studies could lead to non-physiological inter-ABD interactions, cooperative F-actin binding by the α-catenin ABD has previously been reported under non-saturating conditions (Hansen et al., 2013), and supplemental soluble ABD enhanced catch-bonding by the cadherin complex (Buckley et al., 2014), suggesting communication between actin-bound ABDs is likely to be physiologically relevant. We note that cooperative and inhibitory inter-ABD communication are not a priori mutually exclusive, and the interplay of these opposing effects could produce differential outcomes as a function of ABD concentration and filament load, an important subject for future studies.

Molecular and cellular mechanisms for force-activated binding by α-catenin.

(A and B) Cartoons of two conceptual mechanistic models for force-activated actin binding by the α-catenin ABD. Orange box represents the force detector (residues 872–906) at the distal tip of the CTE. Blue gradient represents a force-induced conformational transition in the actin protomer. (C) Cartoon of proposed biological functions for force-activated actin binding by the two cellular populations of α-catenin. Membrane-associated cadherin-catenin complexes are illustrated in purple.

The second model invokes a conformational change in the actin protomer that specifically occurs in the presence of mechanical load, which is recognized and preferentially bound by the force-detector, relieving inhibition (Figure 7B). Although our studies suggest only minor actin conformational changes when the binding is driven by mass action (Figure 5H), they do not rule out as yet unobserved actin conformations specifically evoked by force. Furthermore, while for simplicity we have framed both models in terms of discrete transitions between structural states, low piconewton forces could also modulate the intrinsic structural fluctuations of F-actin to control α-catenin engagement through either mechanism, as has previously been speculated for cofilin (Galkin et al., 2012; Hayakawa et al., 2011; Wioland et al., 2019). Although currently technically prohibitive, structural studies of the α-catenin ABD–F-actin interface in the presence of active force generation, as well as supporting functional experiments, will be necessary to dissect the interplay of these models.

Mechanically-regulated adhesion through α-catenin and vinculin

Our finding that approximately 1 pN of tension along individual filaments is sufficient for force-activated binding suggests that the actin-binding interface of α-catenin has been evolutionarily optimized to sense contractile forces generated by myosin motors (Finer et al., 1994). We speculate force-activated binding enables the cadherin-catenin complex to recognize and preferentially engage pre-stressed actomyosin cables adjacent to the plasma membrane at adherens junctions (Figure 7C), providing a mechanism for initial engagement between actin and the cadherin-associated population of α-catenin, as well as rapidly strengthening adhesion after preliminary attachments are formed. This could facilitate the transition from nascent cell-cell contacts to mature adherens junctions, as punctate nascent adhesions associated with radial actin cables coalesce and spread along a developing circumferential band of tensed actomyosin bundles (Vaezi et al., 2002), as well as support dynamic adherens junction remodeling during epithelial morphodynamics (Lecuit and Yap, 2015). It also provides a mechanism for concentrating the soluble, cytoplasmic α-catenin population at sites of cytoskeletal tension (Figure 7C). Previous studies have demonstrated this population is enriched on actomyosin bundles linked to adherens junctions, where it suppresses lamellipodium activity and promotes adhesion maturation (Drees et al., 2005). While enrichment has been speculated to occur via local release from the cadherin complex, this model is difficult to reconcile with its low cellular concentration and the high concentration of soluble α-catenin required to inhibit Arp2/3-mediated actin branching in vitro (Drees et al., 2005). Force-activated binding provides a feasible cellular mechanism for this enrichment of the homodimeric cytoplasmic α-catenin population, driven by enhanced affinity for tensed F-actin rather than mass action effects due to local concentration. Our identification of a functionally separable force-detector in α-catenin with amino-acid level precision will facilitate a detailed examination of these models in cell lines and in vivo.

We note that force-activated binding is an additional, rather than alternative, mechanism to catch-bond formation (Buckley et al., 2014) for mechanical regulation of F-actin binding. Our finding that vinculin lacks this activity despite forming catch-bonds with F-actin (Huang et al., 2017) strongly suggests that these two modes of mechanical regulation operate by unique structural mechanisms, likely to fulfill distinct biological functions. Vinculin’s lack of force-activated actin-binding activity is consistent with the ordered sequence of mechanically-regulated binding events underlying its coordination with α-catenin at adherens junctions, where only after preliminary attachments form through the cadherin complex and come under load is vinculin recruited and activated to bind F-actin (Yonemura et al., 2010). The lower force threshold for force-activated binding than catch-bond formation (~1 pN vs. ~10 pN) supports a model in which the formation of initial attachments to the cytoskeleton through the cadherin complex is stimulated through force-activated binding, which is subsequently strengthened by catch-bonding through both α-catenin and vinculin during adhesion maturation.

In their parallel work, Xu et al. speculate that the displacement of H1 from the α-catenin ABD upon actin binding could be stabilized or induced by force (Xu et al., 2020), providing a possible structural mechanism for catch-bond formation which we have previously proposed could also be employed by vinculin (Kim et al., 2016; Swaminathan et al., 2017). This model predicts that the force required to dissociate H1 from the α-catenin ABD should be lower than the force required to displace the ABD in the mechanically-reinforced, strongly bound state from F-actin. It furthermore predicts that the constitutive high F-actin affinity α-catenin ABD construct (residues 711–842) lacking H0-H1 reported by Xu et al., which they propose mimics the strongly-bound state (Xu et al., 2020), should no longer form catch bonds with F-actin due to its anticipated inability to switch between weakly and strongly bound states. Thus, this proposed structural mechanistic framework is now well-positioned to be subjected to explicit experimental scrutiny. We believe dissecting the interplay between force-activated binding and catch-bond formation in vitro and in vivo, as well as the structural basis for coordinated actin catch-bonding by α-catenin and vinculin, are important subjects for future studies.

Implications for cytoskeletal mechanotransduction

Our direct observation of force-activated binding to F-actin mediated by a short, flexible sequence element suggests this mechanism could feasibly be employed by other ABPs. Proteins from the large Calponin Homology (CH) domain ABP superfamily have diverse functions as cytoskeletal cross-linkers and plasma membrane/organelle tethers (Liem, 2016; Razinia et al., 2012). Their ABDs have also been reported to have sequence elements that undergo folding transitions associated with filament engagement (Avery et al., 2017; Galkin et al., 2010a; Iwamoto et al., 2018) which can sterically regulate their actin binding (Harris et al., 2019). This suggests members of this family could plausibly employ force-activated binding mechanisms similar to α-catenin to coordinate diverse mechanotransduction pathways throughout the cell, as could other ABPs with similar properties. The experimental strategy established here should be broadly useful for identifying force-sensitive ABPs and defining their force-detector-F-actin interfaces, such as the α-catenin CTE-F-actin interaction, in atomistic detail. This will facilitate elucidating the molecular and cellular mechanisms of cytoskeletal mechanotransduction with sufficient precision for guiding inhibitor development.

Materials and methods

Key resources table
Reagent type
(species) or resource
DesignationSource or referenceIdentifiersAdditional information
Recombinant DNA ReagentpET His-Strep-TEV and pET His-Strep-TEV-Halo vectorsThis paperAmpicillin resistance; expression vectors for E. coli
Contact Alushin Lab for distribution
Recombinant DNA ReagentpFastBac Dual Expression VectorThermoFisher10712024Expression vectors for sf9 insect cells
Gene (H. sapiens)Vinculin; HsVCL isoform 1Synthesized by GenewizUniProt:P18206-2
Gene (H. sapiens)Metavinculin; HsVCL isoform 2Synthesized by GenewizUniProt:P18206-1
Gene (H. sapiens)αE-catenin; HsCTNNA1Addgene # 24194UniProt:P35221
Strain, strain background (E. coli)Rosetta2 (DE3)Novagen71400–4E. coli strain for protein expression
Cell line (S. frugiperda)Gibco Sf9 insect cellThermoFisher11496015Insect cell line for protein expression
Transfected construct (S. frugiperda)Gibco Bac-to-Bac Baculovirus Expression SystemThermoFisher10359016Insect cell line transfection kit for protein expression
Chemical compound, drugHalo-tag ligand amine O4PromegaP6741Synthetic precursor for the fluorescent dye JF-646-Halo
Chemical compound, drugJF-646 NHS-ester building blockTOCRIS6148Synthetic precursor for the fluorescent dye JF-646-Halo
Chemical compound, drugHaloTag Alexa Fluor 488 LigandPromegaG1001
Chemical compound, drugAlexa Fluor 555-PhalloidinThermoFisherA34055
Biological sample
(O. cuniculus)
Actin Protein (Rhodamine): Rabbit Skeletal MuscleCytoskeleton, IncAR05-C
Biological sample
(O. cuniculus)
Actin Protein (Biotin): Rabbit Skeletal MuscleCytoskeleton, IncAB07-C
OtherStreptavidin Coated Polystyrene ParticlesSpherotechSVP-40–5Used in optical trapping experiments
OtherC-flat Holey Carbon Grids for TEM - Gold OnlyEMSCF313-100-AuUsed in cryo-EM experiments
Software, algorithmC-trap viewerLUMICKShttp://www.nat.vu.nl/~iheller/download.html
Software, algorithmTWOM viewerLUMICKShttp://www.nat.vu.nl/~iheller/download.html
Software, algorithmImageJ 1.5Schneider et al., 2012
https://doi.org/10.1038/nmeth.2089
RRID:SCR_003070ImageJ plugin for TIRF image analysis:https://github.com/alushinlab/ActinEnrichment
Software, algorithmRELION 3.0He and Scheres, 2017
https://doi.org/10.1016/j.jsb.2017.02.003
Schneider et al., 2012
https://doi.org/10.1016/j.jmb.2011.11.010
Zivanov et al., 2018
https://doi.org/10.7554/eLife.42166
RRID:SCR_016274
Software, algorithmCTFFIND 4.1.5Rohou and Grigorieff, 2015
https://doi.org/10.1016/j.jsb.2015.08.008
RRID:SCR_016732
Software, algorithmPhenix 1.17.1Afonine et al., 2018
https://doi.org/10.1107/S2059798318006551
RRID:SCR_014224
Software, algorithmCoot 0.8.9Emsley et al., 2010
https://doi.org/10.1107/S0907444910007493
RRID:SCR_014222
Software, algorithmUCSF Chimera 1.14Pettersen et al., 2004
https://doi.org/10.1002/jcc.20084
RRID:SCR_004097
Software, algorithmUCSF ChimeraX 1.0Goddard et al., 2018
https://doi.org/10.1002/pro.3235
RRID:SCR_015872
Software, algorithmGraphPad Prism 8GraphPad Software, IncRRID:SCR_002798

Contact and materials availability

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Further information and requests for resources and reagents should be directed to the corresponding author, Gregory M. Alushin (galushin@rockefeller.edu). All reagents generated in this study are available from the corresponding author without restriction.

Experimental model and subject details

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Globular actin (G-actin) monomers were purified from chicken skeletal muscle as described previously (Pardee and Spudich, 1982) and maintained in G-Ca buffer: G buffer (2 mM Tris-Cl pH 8, 0.5 mM DTT, 0.2M ATP, 0.01% NaN3) supplemented with 0.1 mM CaCl2, at 4°C before use. C-terminally GFP-tagged mouse myosin V HMM and myosin VI S1 were purified from SF9 insect cells using published protocols (Wang et al., 2000). All other proteins were heterologously expressed in Rosetta2(DE3) E. coli cells (Novagen) grown in LB media as described in Method details.

Method details

F-actin preparation

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Filamentous actin (F-actin) was polymerized fresh for each experiment from actin monomers in G-Mg (G buffer supplemented with 0.1 mM MgCl2) and KMEI (50 mM KCl, 1 mM MgCl2, 1 mM EGTA, 10 mM imidazole pH 7.0, 1 mM DTT) buffers as described previously (Gurel et al., 2017); 30% rhodamine-labeled actin filaments used in TIRF microscopy assays were prepared by copolymerizing unlabeled actin monomers and rhodamine-labeled actin monomers (Cytoskeleton catalog # AR05) at a 7:3 molar ratio (total actin concentration 1 μM) at room temperature for 2 hr in the dark. For optical trapping/confocal microscopy assays, 10% biotinylated actin filaments were prepared by copolymerizing purified unlabeled actin monomers and biotinylated actin monomers (Cytoskeleton catalog # AB07) at a 9:1 molar ratio (total actin concentration 1 μM) at room temperature for 2 hr. The biotinylated F-actin was subsequently stabilized and fluorescently labeled by adding 1.2 molar equivalents of Alexa Fluor 555 Phalloidin (ThermoFisher catalog # A34055).

Expression cloning

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Vectors for expression of WT H. sapiens vinculin ABD (879–1066), H. sapiens metavinculin ABD (879–1134), H. sapiens α-catenin ABD (αE-catenin664-906), α-catenin ABDΔC (αE-catenin664-871), vinculin ABD-NCSwap (αE-catenin664-708-vinculin916-1041-αE-catenin837-906), and vinculin ABD-NCSwapΔC (αE-catenin664-708-vinculin916-1041-αE-catenin837-871) were constructed by inserting the corresponding cDNA sequence into a pET bacterial expression vector with an N-terminal His6-tag, Strep-tag, TEV cleavage site, and, for proteins used in fluorescence imaging experiments, a subsequent C-terminal Halo-tag by Gibson assembly (Gibson et al., 2009). Constructs for proteins used in Cryo-EM structural studies are not Halo-tagged. Three WT α-catenin TIRF trials were performed with a previously reported Halo-tagged α-catenin ABD construct (αE-catenin671-906), and no difference was observed.

Expression and purification of actin-binding proteins

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C-terminally GFP-tagged mouse myosin V HMM and myosin VI S1 were purified from SF9 cells using published protocols (Wang et al., 2000). Human calmodulin (CaM) was purified from Rosetta2(DE3) E. coli cells using a published protocol (Putkey et al., 1985) and stored in gel filtration buffer (20 mM Tris-Cl pH 8.0, 100 mM NaCl, 2 mM β-mercaptoethanol) supplemented with 5% v/v glycerol.

All other ABPs were expressed in Rosetta2(DE3) E. coli cells (Novagen) grown in LB media at 37°C to an optical density of 0.8–1.0 and induced with 0.7 mM IPTG. After induction, the cells were grown for 16 hr at 16°C, then cell pellets were collected and stored at −80°C until use. To purify Halo-tagged ABPs for fluorescent labeling, cells pellets were resuspended in lysis buffer (50 mM Tris-Cl pH 8.0, 150 mM NaCl, 5% v/v glycerol, 2 mM β-mercaptoethanol, 20 mM imidazole) and lysed with an Avestin Emulsiflex C5 homogenizer, after which the lysate was clarified at 15,000 g for 30 min. Cleared lysate was incubated with Ni-NTA resin (Qiagen) for 1 hr on a rotator at 4°C, after which the flow-through was discarded and the resin was washed with five bed volumes of lysis buffer. Proteins were subsequently eluted in elution buffer (50 mM Tris-Cl pH 8.0, 150 mM NaCl, 5% v/v glycerol, 2 mM β-mercaptoethanol, 300 mM imidazole). Purified His-tagged TEV protease (prepared according to a published protocol [Tropea et al., 2009]) was added at 0.05 mg/mL working concentration, then the eluate was dialyzed against dialysis buffer (20 mM Tris-Cl pH 8.0, 300 mM NaCl, 5% v/v glycerol, 2 mM β-mercaptoethanol) for 16 hr. The protein solution was then reapplied to Ni-NTA resin, and the flow-through was collected.

Protein was then sequentially purified by a HiTrapQ HP anion exchange column (GE Healthcare) followed by size exclusion chromatography on a Superdex 200 Increase column in gel filtration buffer supplemented with 10% v/v glycerol, and then snap-frozen in liquid nitrogen and stored at −80°C. Non-Halo-tagged proteins prepared for cryo-EM studies were purified identically, except glycerol was omitted from all buffers.

After rapid thawing before use, proteins were clarified by ultracentrifugation at 45,000 rpm in a TLA100 rotor for 10 min at 4°C. All protein concentrations were estimated using the Bradford colorimetric assay (Pierce), calibrated with BSA.

Synthesis of Halo-JF-646 and labeling of ABPs

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Fluorescent dye JF-646 (Grimm et al., 2015) NHS-ester building block (TOCRIS) was conjugated with Halo-tag ligand amine O4 (Promega) by synthetic chemistry according to published protocols (Grimm et al., 2017). Briefly, 1.5 equivalents of amine O4 ligand were added to one equivalent of the JF-646 NHS-ester in DMF followed by adding 5% triethylamine. The reaction was vigorously stirred for 16 hr at room temperature and the product was purified by silica gel chromatography, dried by SpeedVac (ThermoFisher), and reconstituted in DMSO.

For optical trapping/confocal microscopy assays, HaloTag Alexa Fluor 488 Ligand (Promega) was utilized as described above, followed by desalting through a PD SpinTrap G-25 column (GE Healthcare) according to the manufacturer’s protocol to remove unreacted dye before use. To label the Halo-tagged actin-binding proteins with Halo-JF-646 for TIRF microscopy assays, two equivalents of synthesized Halo-JF-646 dye was added to the protein solution, followed by incubation for at least 2 hr in the dark at 4°C before use. Subsequent removal of excess dye was not required, as JF-646 is a fluorogenic dye (Grimm et al., 2015).

Correlative force spectroscopy and confocal microscopy assays

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Experiments were performed at room temperature (approximately 25°C) on a LUMICKS C-Trap instrument combining confocal fluorescence with dual-trap optical tweezers (Hashemi Shabestari et al., 2017; Wasserman et al., 2019). The optical traps were cycled through pre-set positions in the five channels of a microfluidic flow cell by an automated stage (Figure 1A). Channels 1–3 were separated from each other by laminar flow, which we utilized to form actin filament tethers between two 4 μm-diameter streptavidin-coated polystyrene beads (Spherotech) held in optical traps with a stiffness of 0.3 pN/nm. We first captured a single bead in each trap in channel 1. The traps were then transferred to channel 2, containing 5–20 nM Alexa 555 phalloidin-stabilized, 10% biotinylated F-actin in motility buffer (‘MB’: 20 mM MOPS pH7.4, 5 mM MgCl2, 0.1 mM EGTA, 1 mM DTT) supplemented with 1 µM dark phalloidin, where tethers were formed by briefly moving 1 of the two traps toward the other trap against the direction of flow, followed by rapidly moving the traps to channel 3, which contained only buffer (MB + 1 μM dark phalloidin). The presence of a tether was verified by carefully separating the traps and observing an associated increase in force when monitoring the force-extension curve, applying the minimum extension feasible to avoid prematurely rupturing the tether.

The traps were then moved to orthogonal channel 4 or 5, which contained 2 μM fluorescently labeled vinculin ABD or α-catenin ABD (diluted in MB supplemented with 1 μM dark phalloidin), respectively, and flow was ceased during data acquisition. Force data were acquired at 200 Hz during constant velocity (0.1 μm/s) pulling experiments while simultaneously acquiring 2-color confocal fluorescence scans at 33 ms line scan time, exciting Alexa Fluor 488 HaloTag ligand and Alexa Fluor 555 phalloidin fluorophores with laser lines at 488 nm and 532 nm, respectively.

Data analysis was performed using ImageJ and custom software provided by LUMICKS. Force data from the two traps were averaged and binned to the confocal frame interval. The intensity values are measured by drawing a box in ImageJ to measure the fluorescence intensities of actin and ABP in both channels with background subtraction, calculating the background from an equal-sized box from that frame in an area devoid of filaments or beads. For paired analysis, the ‘high-force’ and ‘low-force’ averages were calculated only from the final tether to rupture, as long as the entire force trace has at least 10 quantifiable confocal image frames. For fluorescence-force correlation plots, only single-filament tethers selected based on Figure 1—figure supplement 2A–C were used. IABP/IActin values were normalized by dividing the values in each recording by the largest value observed during that recording. K-means clustering analysis was performed to identify the cutoff force in the correlation plots for each ABP. Briefly, the force-fluorescence data were grouped into two clusters using scikit-learn (Pedregosa et al., 2011). For both α-catenin and vinculin, the data separated along the force axis, providing a threshold force for each ABP. Furthermore, a silhouette analysis for both ABPs confirmed that the data should not be clustered into more than two clusters (Kaufman and Rousseeuw, 2009).

TIRF force reconstitution assays

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Glass coverslips (Rectangular: Corning 22 × 50 mm #1½ Cover Glass; Square: Fisherbrand 22 × 22 mm #1½ Microscope Cover Glass) were cleaned by 30 min 100% acetone wash, 10 min 100% ethanol wash, and 2 hr 2% Hellmanex III liquid cleaning concentrate (HellmaAnalytics) wash in a bath sonicator followed by rinsing with water. The cleaned glass coverslips were coated with 1 mg/mL mPEG5K-Silane (Sigma) in a 96% ethanol, 10 mM HCl solution for at least 16 hr. After coating, the coverslips were rinsed with 96% ethanol and water, then air-dried and stored at 4°C until use. Flow cells were prepared with one square and one rectangular coverslip, both coated with mPEG-Silane. Double-sided adhesive tape (3M) was used to make ~4 mm-wide flow chambers between the coverslips, which were open on both sides to facilitate buffer exchange when adding components for imaging.

For each assay, 6 mg/mL anti-GFP antibody (Sigma #G1546) solution reconstituted in water was first introduced into the flow chamber and incubated for 2 min. Subsequently, MB containing 0.075 μM GFP-myosin V S1 and 0.15 μM GFP-myosin VI S1 were flowed into the chamber and incubated for another 2 min. A solution of 1 mg/mL bovine serum albumin (BSA) in MB was then flowed into the flow chamber and incubated for at least 2 min, after which 1 μM rhodamine-labeled F-actin in MB was flowed into the chamber and incubated for 20–30 s. The flow chamber was rinsed with MB buffer to remove F-actin not bound to the rigor-state motors, then imaging buffer (MB without ATP, supplemented with 0.01% Nonidet P-40 substitute [Roche], 1 μM calmodulin, 15 mM glucose [Sigma], 1 μg/mL glucose oxidase [Sigma], and 0.05 μg/mL catalase [Sigma]) containing 2 μM fluorescently labeled ABP was flowed into the chamber. The first movie (-ATP, no force) was then recorded. A second imaging buffer, identical to the first but now including 100 μM ATP, was then introduced into the same chamber, and a second movie (+ATP, with force) was recorded. For each solution that was introduced, complete buffer exchange was facilitated by applying a filter paper at the other end of the flow chamber while pipetting.

Dual-color TIRF image sequences (movies) were recorded at room temperature (approximately 25°C) using a Nikon TiE inverted microscope equipped with an H-TIRF module and an Agilent laser launch, driven by Nikon Elements software. Images were taken every 2 s with an Apo TIRF 60 × 1.49 NA objective (Nikon) on an Andor iXon EMCCD camera; Rhodamine and JF646 fluorophores were excited by laser lines at 561 nm and 640 nm, respectively.

Filament region quantification

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To quantify ABP association with individual ‘filament regions’ of TIRF movies, we developed a custom ImageJ (Schneider et al., 2012) plugin (Figure 2—figure supplement 1A) which features a graphical user interface (GUI). The plugin takes as input two movie files, the actin channel and the ABP channel from a dual-color TIRF experiment, as well as an adjustable set of parameters (set by default in the GUI to the optimized values used in this study).

To identify regions of interest (ROIs) in each frame, the actin channel image series was preprocessed (unsharp mask, median filter, rolling ball subtraction), binarized, then segmented into contiguous regions of pixels using the built-in ImageJ plugin ‘Analyze Particles’. ROIs whose centroids were fewer than 30 pixels from the edge of the field-of-view were excluded from further analysis due to incompatibility with downstream background subtraction procedures. The ROIs were then tracked through the image series and sorted into filament regions (representing individual filaments or small groups of filaments) by shortest Euclidean distance between ROI centroids in neighboring frames, with a maximum distance cutoff of 24 pixels. ROIs that were not matched with a pre-existing filament region by this criterion (i.e. whose centroid was greater than 24 pixels away from any ROI in the previous frame) were considered to represent a newly appeared filament region. Although this may result in overcounting the absolute number of filaments (should this be of interest; we do not believe this caveat impacts the conclusions of the present study), we find this procedure reliably handles events such as filament breakages and desultory motion in a completely automated fashion. To account for poorly tethered filaments fluctuating in and out of the evanescent field, only filament regions detected in at least 10 consecutive frames were included in the analysis.

Intensity in both the actin channel and the ABP channel were then quantified for each region. For each channel, the local background for each filament region in each frame was calculated as the mean gray value of the pixels from a 60 by 60 pixel box centered on the region’s centroid, excluding pixels belonging to the region itself or any other filament region detected in the frame. Background-subtracted mean gray values were then calculated, followed by the ratio of these values across all frames in which the filament region was detected and their average, which we here report as the overall IABP/IActin for that filament region. The program then outputs all frame and average IABP/IActin values of the tracked filaments sorted by filament number for analysis, as well as a file containing all tracked regions.

Cryo-EM sample preparation and data collection

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F-actin was polymerized in G-Mg and KMEI from 5 µM unlabeled actin monomers at room temperature for 1 hr and then diluted to 0.6 μM in KMEI before use. Purified metavinculin ABD (879–1134) was diluted in KMEI to 10 μM before use. After screening grids prepared with finely sampled ABP concentrations, we found diluting purified α-catenin ABD (664-906) to 20 μM in KMEI before use gave an optimal balance between filament decoration and bundling.

Immediately before sample preparation, CF-1.2/1.3-3Au 300-mesh gold C-flat holey carbon cryo-TEM grids (Protochips) were plasma cleaned with a Hydrogen/Oxygen mixture for 5 s in a Gatan Solarus. Actin (3 μL) was first applied to the grid in the humidified chamber of a Leica EM GP plunge freezer and incubated for 60 s at 25°C. Actin-binding protein (3 μL) was then applied and incubated for 30 s. Solution (3 μL) was then removed and an additional 3 μL of the same actin-binding protein solution was applied. After an additional 30 s, 3 μL of solution was removed, then the grid was back-blotted for 5 s, plunge-frozen in ethane slush, and stored in liquid nitrogen until imaging.

Cryo-EM data were recorded on a Titan Krios (ThermoFisher/FEI) operated at 300 kV equipped with a Gatan K2 Summit camera. SerialEM (Mastronarde, 2005) was used for automated data collection. Movies were collected at a nominal magnification of 29,000X in super-resolution mode resulting in a calibrated pixel size of 1.03 Å/pixel (superresolution pixel size of 0.515 Å/pixel), over a defocus range of −1.5 to −3.5 μm; 40 frames were recorded over 10 s of exposure at a dose rate of 6 electrons per pixel per second (1.5 electrons per Å2 per second) for a cumulative dose of 60 electrons per Å2.

Cryo-EM image processing

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Unless otherwise noted, all image processing was performed within the RELION-3.0 package (Zivanov et al., 2018). Movie frames were aligned and summed with 2 × 2 binning using the MotionCor2 algorithm (Zheng et al., 2017) as implemented in RELION (Zivanov et al., 2019), utilizing subframe motion correction with 5 × 5 patches. Contrast transfer function (CTF) parameters were estimated from non-doseweighted summed images with CTFFIND4 (Rohou and Grigorieff, 2015). Bimodal angular searches around psi angle priors were utilized in all subsequent 2D and 3D alignment/classification procedures. Around 2000 segments were initially manually picked, extracted, and subjected to 2D classification to generate templates for auto-picking. Helical auto-picking was then performed utilizing a step-size of 3 asymmetric units with a 27 Å helical rise. Segments were extracted from dose-weighted (Grant and Grigorieff, 2015) sum images in 512 × 512 pixel boxes which were not further down-sampled, then a second round of 2D classification followed by auto-picking with featureful class averages was performed.

A total of 237,503 particles from 1708 images (for the metavinculin ABD-actin dataset) and 540,553 particles from 7317 images (for the α-catenin ABD-actin dataset) were then extracted and subjected to whole-dataset 2D classification (Figure 3—figure supplement 1) using a 200 Å tube diameter and 300 Å mask diameter. 234,703 segments from the metavinculin ABD-actin dataset and 428,335 particles from the α-catenin ABD-actin dataset contributed to featureful class averages and were selected for 3D analysis.

All subsequent 3D classification and 3D auto-refine steps were primed with estimates of helical rise and twist of 27.0 Å and −167.0°, respectively, utilizing an initial reference low-pass filtered to 35 Å resolution, with the outer tube diameter set to 200 Å, inner tube diameter set to −1, and the mask diameter set to 300 Å. The first round of 3D classification into three classes was performed utilizing a reconstruction of a bare actin filament (EMBD-7115) as the initial reference. A second iteration of 3D classification was then performed as above, utilizing a featureful class with clear ABP density produced by the first round as the initial reference. For both datasets, this second round of 3D classification yielded two classes with helical parameters similar to the initial estimates and well-resolved 3D features, and one junk class with aberrant helical parameters and distorted features (Figure 3—figure supplement 1). Segments contributing to the two good classes were then pooled (215,369 particles for the metavinculin ABD-actin dataset and 414,486 particles for the α-catenin ABD-actin dataset) for 3D auto-refinement.

The first round of auto-refinement was then performed using one of the two good 3D averages as an initial reference. All masks for subsequent post-processing steps were calculated with 0 pixel extension and a six pixel soft edge from the converged reconstruction produced by that round of refinement, low-pass filtered to 15 Å and thresholded to fully encompass the density map. The first-round post-processing was performed with a 50% z length mask, followed by CTF refinement without beam-tilt estimation and Bayesian polishing (Zivanov et al., 2019). A second round of auto-refinement was then performed using the converged reconstruction from the first round as the initial reference. The second-round post-processing was performed with a 30% z length mask, followed by a second round of CTF refinement with beam-tilt estimation and Bayesian polishing. A final round of auto-refinement was then performed using the converged reconstruction from the second round as the initial reference. We found that this iterative procedure of tightening the mask for polishing resulted in substantial resolution improvements, potentially by mitigating the effects of medium-range disorder in F-actin previously speculated to limit the resolution of reconstructions of this filament (Galkin et al., 2012; Merino et al., 2018).

The final reconstructions converged with helical rise of 27.1 Å and twist of −167.1° for the metavinculin ABD–F-actin complex, and a helical rise of 27.0 Å and twist of −166.9° for the α-catenin ABD-F-actin complex, consistent with our finding that actin rearrangements evoked by these ABPs are minimal (Figure 5H). Final post-processing was performed with a 30% z length mask, leading to global resolution estimates of 2.9 Å for the metavinculin ABD–F-actin complex and 3.2 Å for the α-catenin ABD–F-actin complex by the gold-standard Fourier shell correlation (FSC) 0.143 criterion (Figure 3—figure supplement 1). B-factors of both datasets estimated during post-processing were then used to generate sharpened, local-resolution filtered maps with RELION. The key statistics summarizing cryo-EM image processing are reported in Table 1.

Asymmetric focused classification (without alignment) utilizing masks isolating the ABD region showed no evidence of segments with unoccupied binding sites (data not shown), suggesting that decoration of actin filaments by both 10 μM metavinculin ABD and 20 μM α-catenin ABD was essentially complete, with 100% occupancy at the limit of detection of current methods.

Model building and refinement

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Sharpened, local-resolution filtered maps as described above were used for model building. The 2.9 Å and 3.2 Å density maps were of sufficient quality for de novo atomic model building. As structures of components were available, initial models of actin (PDB 3j8a), metavinculin ABD (PDB 3jbk) truncated to residues 981–1131 and α-catenin ABD (PDB 4igg chain B) truncated to residues 699–871 were fit into the density map using Rosetta (Wang et al., 2016). Models were subsequently inspected and adjusted with Coot (Brown et al., 2015; Emsley et al., 2010), and regions that underwent significant conformational rearrangements were manually rebuilt. The models were then subjected to several rounds of simulated annealing followed by real-space refinement in Phenix (Adams et al., 2010; Afonine et al., 2018) alternating with manual adjustment in Coot. A final round of real-space refinement was performed without simulated annealing. The key statistics summarizing model building, refinement, and validation are reported in Table 1.

Molecular graphics and structure analysis

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Structural figures and movies were prepared with ChimeraX (Goddard et al., 2018). Per-residue RMSD analysis was performed with UCSF Chimera (Pettersen et al., 2004) as previously described (Zhang et al., 2015). The surface area of actin-binding interfaces was calculated with PDBePISA (Krissinel and Henrick, 2007; (EMBL-EBI). Model quality was assessed with EMRinger (Barad et al., 2015) and MolProbity (Chen et al., 2010) as implemented in Phenix.

Sequence alignments

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Protein sequences of human vinculin (UniProt Accession Code P18206-2), human metavinculin (P18206-1), human αE-catenin (P35221), human αN-catenin (P26232), and human αT-catenin (Q9UI47) were aligned with ClustalOmega (Sievers and Higgins, 2014; EMBL-EBI).

Quantification and statistical analysis

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Plotting and statistical analysis of data from TIRF force reconstitution assays and force-spectroscopy/confocal microscopy assays was performed with GraphPad Prism 8. All the details can be found in the figure legends of these figures and in the Method details. The data collection and refinement statistics of the cryo-EM structures can be found in Table 1. Resolution estimations of cryo-EM density maps and statistical validation performed on the deposited models are described in the Method details.

Data and code availability

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The atomic coordinates for the metavinculin ABD–F-actin complex and α-catenin ABD–F-actin complex have been deposited in the Protein Data Bank (PDB) with accession codes 6UPW and 6UPV, and the corresponding cryo-EM density maps in the Electron Microscopy Data Bank (EMDB) with accession codes EMD-20844 and EMD-20843.

The code for analyzing TIRF movies is freely available as an ImageJ plugin with a graphical user interface at https://github.com/alushinlab/ActinEnrichment. (copy archived at https://github.com/elifesciences-publications/ActinEnrichmentAlushinlab, 2020).

All other data are available in the manuscript or supplementary materials.

Data availability

The atomic coordinates for the metavinculin ABD-F-actin complex and α-catenin ABD-F-actin complex have been deposited in the Protein Data Bank (PDB) with accession codes 6UPW and 6UPV, and the corresponding cryo-EM density maps in the Electron Microscopy Data Bank (EMDB) with accession codes EMD-20844 and EMD-20843. The code for analyzing TIRF movies is freely available as an ImageJ plugin with a graphical user interface at https://github.com/alushinlab/ActinEnrichment (copy archived at https://github.com/elifesciences-publications/ActinEnrichment). All other data are available in the manuscript or supplementary materials.

The following data sets were generated
    1. Mei L
    2. Alushin GM
    (2020) RCSB Protein Data Bank
    ID 6UPW. Metavinculin ABD-F-actin complex.
    1. Mei L
    2. Alushin GM
    (2020) RCSB Protein Data Bank
    ID 6UPV. Alpha-E-catenin ABD-F-actin complex.

References

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    Acta Crystallographica. Section D, Biological Crystallography 66:486–501.
    https://doi.org/10.1107/S0907444910007493
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    2. Rousseeuw PJ
    (2009)
    Finding Groups in Data: An Introduction to Cluster Analysis, 344
    John Wiley & Sons.
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    2. Varoquaux G
    3. Gramfort A
    4. Michel V
    5. Thirion B
    6. Grisel O
    7. Blondel M
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    9. Weiss R
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    11. Vanderplas J
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    15. Perrot M
    16. Duchesnay E
    (2011)
    Scikit-learn: machine learning in Python
    The Journal of Machine Learning Research 12:2825–2830.
    1. Putkey JA
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    The Journal of Biological Chemistry 260:4704–4712.

Decision letter

  1. Andrew P Carter
    Reviewing Editor; MRC Laboratory of Molecular Biology, United Kingdom
  2. Suzanne R Pfeffer
    Senior Editor; Stanford University School of Medicine, United States

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

Acceptance summary:

The actin cytoskeleton has the unique ability to propagate mechanical stimuli at cell-cell and cell-matrix interactions, resulting in the maturation of adhesion complexes. The actin filaments that are placed under tension in these regions create unique binding sites for actin-binding proteins. Vinculin and α-catenin have been shown differentially bind to tensed and relaxed actin filaments, but the mechanism remains elusive. Here, Mei et al. address this important question. They combine beautiful cryo-EM structures of actin decorated by metavinculin or α-catenin with optical trap and TIRF motility assays to explore the how actin binding proteins interact with actin under tension.

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 "Molecular mechanism for direct actin force-sensing by α-catenin" for consideration by eLife. Your article has been reviewed by three peer reviewers, one of whom is a member of our Board of Reviewing Editors, and the evaluation has been overseen by a Senior Editor. The reviewers have opted to remain anonymous.

Our decision has been reached after consultation between the reviewers. Based on these discussions and the individual reviews below, we regret to inform you that your work will not be considered further for publication in eLife at this stage. As you will see from the comments below the reviewers were impressed by your cryoEM structures and felt the work was of general interest and potentially suitable for eLife. However they had a number of concerns particularly concerning the force experiments.

The reviewers all agree that your work has the potential to make a stronger paper. You will need to make the decision what you want to do with the manuscript, especially given the current situation. However, if you think you can solve most of the issues raised then we encourage you to resubmit to eLife later in the year.

Reviewer #1:

The actin cytoskeleton has the unique ability to propagate mechanical stimuli at cell-cell and cell-matrix interactions, resulting in the maturation of adhesion complexes. The actin filaments that are placed under tension in these regions create unique binding sites for actin-binding proteins. Vinculin and α-catenin have been shown differentially to bind to tensed and relaxed actin filaments, but the mechanism remains elusive. Here, Mei et al. explore the binding of vinculin and α-catenin to actin under tension to address this important question.

The authors first develop an assay to pull single actin filaments and assess the binding intensity of vinculin and α-catenin. Vinculin did not show differential binding to filaments under high or low pulling force, but α-catenin showed a mild preference for filaments pulled beyond 1 pN. To validate this result, the authors developed another assay where opposite polarity myosin motors were used to place actin filaments under tension. This showed that α-catenin binding was enhanced when ATP was added, whereas vinculin binding was unchanged. The authors obtain beautiful cryo-EM structures of actin decorated by metavinculin or α-catenin at high resolution. The major binding site for the two proteins on actin was largely similar, whereas the minor binding site was distinct. In particular, the C-terminal extensions followed different paths on actin, and the extreme 35 amino acids on α-catenin were unresolved. The authors speculate that these unresolved residues mediated the force-sensitivity. Truncating these residues abolished force sensitivity, but switching the major binding region of α-catenin with vinculin did not, suggesting that the C-terminal extension of α-catenin is necessary and sufficient to mediate force-sensitivity. While this work provides important insight into how forces can be propagated at tensed cellular environments, I have a few major concerns that should be addressed by the authors.

1) In Figure 1, high force was taken to be the 5 frames before breakage and low force to be 5 frames before that. Therefore, a specific amount of time is taken to represent high versus low force, which is not appropriate. Furthermore, the low number of replicates raises questions on the robustness of the result. Therefore, I find that 1D and E weaken the author's claims, which are better represented in 1F and G, where the data is plotted against force. In the latter, however, it is not clear why 1 pN is chosen to divide the data. A regression analysis is more appropriate, with R2 values reported.

2) A necessary assumption in the myosin assay is that myosins behave similarly in all experiments. However, this is not validated and may not be the case. For example, Video 7 (ABD-NCSwap) clearly has more bundled actin filaments after ATP addition than Video 6 (ABD-ΔC) and Video 8 (ABD-NCSwapΔC). The authors even hint that myosin inhibition can occur in certain cases. Additional experiments to assess myosin activity is required for these data to be reliable. A suitable assay would be myosin gliding in the presence of the different constructs, for example.

3) Metavinculin is used for structural studies to prevent bundling, and the authors validate its use by stating that it is similarly force-insensitive in the myosin assay. However, this is represented by a single experiment in Figure 2—figure supplement 1D, while it is clear from Figure 2—figure supplement 1B that there is significant variability in this type of data. Therefore, the use of metavinculin as a proxy for vinculin in structural analyses is not properly justified. Additional replicates will be required.

4) In testing whether the D-loop interactions contribute to force-sensitivity, mutations are made in α-catenin to resemble metavinculin at the D-loop interface. The mutations abolish binding to actin altogether, and an interpretation provided by the authors is that "this interface does not have a separable role in force-activated actin binding". This interpretation cannot be correct if no binding was detected altogether, and the authors should remove the statement.

5) The motivation to truncate the disordered region of the C-terminal extension of α-catenin is not entirely clear. The residues of the C-terminal extension preceding this disordered stretch are clearly different between α-catenin and vinculin in the two cryo-EM structures. This region is therefore also a good candidate force sensor. The authors need to explain why they didn't switch this region between the two proteins.

Reviewer #2:

Using distinct biophysical assays, Mei et al. investigate the effect of pN scale forces on the interaction between actin filaments and two adhesion-associated actin-binding proteins (either α-catenin or its homolog vinculin). They show that forces moderately promote the binding of α-catenin but not vinculin. Extending their previous structural work on (meta) vinculin, they then determine high-resolution structures of the ABDs of either protein on actin filaments in the absence of force, which reveal differences in the C-terminal tail regions. Using chimeric constructs, they provide additional biochemical support for the C-terminus of α-catenin conferring force sensitivity.

Overall, the paper clearly contains several strong elements that merit publication. It focuses on a pertinent question of broad interest (the molecular mechanisms of force sensing in the actin cytoskeleton) and the data quality is high throughout. The TIRFM motility assay is a useful addition to the field and the observation of force-sensitive binding of α-catenin is in itself novel and interesting.

The central weakness of the paper is that it falls rather short in providing answers to the most pressing questions: The nature of the force-induced conformational change in actin filaments remains ultimately unclear. Similarly, the "force-sensing" C-terminal element of α-catenin (residues 872-906) is not resolved in the cryoEM structure. Hence, the detailed mechanism of force sensing remains to be elucidated.

That being said, I still feel that the paper deserves to be published even in the absence of a comprehensive structural mechanism. My main points rather address some of the immediate technical and conceptual issues that require improvement.

1) Both vinculin and α-catenin ABDs are known to bundle actin filaments. I was therefore surprised to see that the authors seem to neglect potential confounding effects that bundling might cause in their reconstitutions. Both TIRFM and optical trapping experiments are done in a configuration in which filaments appear to be clearly bundled. The authors even mention that filaments become more strongly bundled in the TIRFM assay upon ATP addition. I am concerned that some of their observations might be equally explainable by differential effects of bundling/filament overlap configurations on vinculin and α-catenin. The authors need to perform control experiments under conditions where they clearly observe and analyze only individual actin filaments to demonstrate that the effect of force is indeed based on a conformational change in the filament itself.

2) The authors should be more explicit and conscious about the ABD concentrations employed in their in vitro assays. This information can currently only be found in the Materials and methods section (should be part of the figures/captions!). Why did they chose to work at 2uM for either protein in buffers of low ionic strength? Previous measurements show that both α-catenin and vinculin ABDs binds with sub-micromolar affinity to filaments (Hansen et al., 2013 and Le Clainche et al., 2010), which means that the authors work at or close to saturation. This should mask or at least severely diminish the effects that force might exert on ABD binding. In my opinion, the authors need to take much greater care of the fractional occupancy of filaments by ABDs (by measuring the ABD affinities) under their experimental conditions to ensure that they work still in a sensitive concentration regime. Determining exact affinities in the presence and absence of force in the TIRFM assay also ought to give them additional information concerning the relative number of binding sites at saturation (maximal signal), which might allow them to discriminate between the proposed force sensing mechanisms.

3) The "tug of war" motility assay employed by the authors, while ingenious, suffers from a central weakness. The opposite polarity motors are equally likely to produce tensile AND compressive forces on the surface-bound filament. This important limitation is not discussed by the authors at all. Admittedly, the optical trapping assay shows that the effect can also be observed for tensile forces only. However, the magnitude in the TIRFM assay is much greater, which is why one cannot help but wonder whether compression is the true binding-promoting cue. I am fully aware that this cannot easily be addressed experimentally. However, the authors should be more open in discussing this alternative interpretation.

4) The "steric exclusion" model put forward by the authors posits that adjacent α-catenin ABDs interfere with synchronous binding. How can this be reconciled with the observation of cooperative α-catenin ABD binding (Hansen et al., 2013), indicating stabilizing effects? The authors cite and superficially discuss this point, but ultimately do not provide a satisfactory explanation. Is there any indication of partial binding in their cryoEM data? Along the same vein, have they compared the binding stoichiometry of α-catenin (either wt or C-terminal deletion mutants) at saturation via TIRFM or co-sedimentation?

Reviewer #3:

Mei et al. present a very interesting study of force-dependent binding of actin-binding domains to filamentous actin. Using optical trapping, they demonstrate forces > 1pN are sufficient to show enhanced actin binding of a-catenin, a process they call force-activated binding, but not vinculin. The authors identify a C-terminal motif of a-catenin that is required for force-activated binding and that confers similar behavior to vinculin when added. Using cryoEM, they obtain structures of both a-catenin and vinculin bound to actin, showing where the C-terminal motif binds. This work provides interesting new information about how forces on filaments can influence actin-binding proteins. However, the mechanistic insight is limited by the lack of data on how a 1pN force alters actin and how a-catenin might sense that change. In addition, I have concerns about interpretation of the optical trapping and TIRF data and am unclear why constructs and labeling schemes vary across the different experimental methods used.

1) The optical trapping results appear to pool data on single filaments and bundled filaments (as noted in the Materials and methods), which makes interpretation of the data very difficult and raises questions about the conclusions.

a) How many single filaments vs bundled filaments are included in the data set? The caption of Figure 1 indicates that it shows data on individual filaments, but the images of actin do not look like single filaments. For example, in the series of images of actin filaments in Figure 1C, the 4th actin filament looks like it could be several filaments, and the 5th image seems to show a broken filament. I'm not convinced that a single rupture force conclusively shows that only one filament is being imaged, as all filaments need not be load bearing and may only be laterally associated with a filament that is attached to the bead. Could a calibrated measurement of filament fluorescence intensity be used to quantify the number of filaments in a bundle?

b) It seems that the amount of force felt by each filament will depend on precisely how each is coupled to the beads and to each other, which is hard to know. Won't the inclusion of actin bundles lead to an error in estimating the force at which binding of a-catenin increases?

c) How do the force-extension curves obtained in this study compare with previous measurements of actin force-extension, such as Liu and Pollack (Biophys J, 2002)? This might help to identify single vs bundled filaments.

d). How do the authors correct for filament length in their analysis? The effect of 1pN on a fluctuating filament will depend on the length of the filament.

e) In the vinculin ABD binding experiment in Figure 1B, why is the relative ABD intensity varying so much (~30%) if binding of the ABD has equilibrated on the filament? And how is the intensity ratio normalized in Figure 1F and 1G?

2) The myosin TIRF experiments also raise several questions:

a) Are the filaments imaged in the TIRF experiments single filaments or bundles (or both)? As discussed above, can calibrated fluorescence intensity be used to identify single vs bundled filaments?

b) Won't some filaments in the myosin TIRF assay experience compression while others experience tension? What about twist? How does this affect data interpretation and the force that is responsible for a-catenin force-activated binding?

c) The authors note the variability in the assay. Any thoughts on the source of this variability? Would a lower density of filaments in Figure 2B and 2C enable easier data collection? Also, the variability in the TIRF data shown in Figure 6—figure supplement 1 is puzzling. Why are some values of normalized intensity negative?

3) The cryoEM data is very impressive. While the structures of a-catenin and vinculin ABDs on actin are well done, they don't provide insight into how force-activated binding might be happening.

a) While imaging filaments under a load in cryoEM would be very difficult, can the authors identify two different classes of filaments in their data, analogous to their "low force" and "high force" grouping of trapping data in Figure 1? If a force of only 1pN is necessary to drive enhanced binding, perhaps filaments under forces that are above and below that may be captured during freezing, which could provide valuable mechanistic insight.

b) Was a structure of the vinculin ABD with the a-catenin C-terminal motif solved to confirm that it interacts with the filament in the same way as it does on a-catenin?

4) The choice of filament labeling strategies and ABD used appears to vary from experiment and would benefit from more explanation and/or experiments with the same constructs.

a) In the cryoEM structures, metavinculin was used rather than vinculin to avoid bundling. Why wasn't metavinculin used in the optical trapping and TIRF experiments, given that bundling complicates analysis and interpretation of the data?

b) The optical trapping experiments used phalloidin-labeled actin filaments, while the TIRF experiments used rhodamine-labeled actin filaments. Why weren't the experiments done with the same actin labeling method?

5) The proposed models seem reasonable but would benefit from more consideration of fluctuations and comparison with other existing models.

a) It is unclear how the thermal fluctuations of a filament under physiological conditions would affect the proposed a-catenin steric exclusion and actin filament conformational change models. Won't the small changes associated with a force of 1pN already be sampled by thermal fluctuations? While detailed modeling is beyond the scope of the manuscript, some discussion of thermal fluctuations and the proposed models would be helpful.

b) The authors cite Ishiyama et al., 2018, which presents a different mechanism for a-catenin force dependence based on conformational unfolding of the ABD under force, but there is little discussion of it. Are the proposed models compatible with this?

c) As a minor point, the authors say that they report the first direct observation of enhanced actin binding by an actin-binding protein. However, Hayakawa et al., 2011, which the authors cite, previously reported tension-dependent binding of cofilin to actin.

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

Thank you for submitting your article "Molecular mechanism for direct actin force-sensing by α-catenin" for consideration by eLife. Your article has been reviewed by two peer reviewers, one of whom is a member of our Board of Reviewing Editors, and the evaluation has been overseen by Suzanne Pfeffer as the Senior Editor. The reviewers have opted to remain anonymous.

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

We would like to draw your attention to changes in our revision policy that we have made in response to COVID-19 (https://elifesciences.org/articles/57162). Specifically, when editors judge that a submitted work as a whole belongs in eLife but that some conclusions require a modest amount of additional new data, as they do with your paper, we are asking that the manuscript be revised to either limit claims to those supported by data in hand, or to explicitly state that the relevant conclusions require additional supporting data.

Our expectation is that the authors will eventually carry out the additional experiments and report on how they affect the relevant conclusions either in a preprint on bioRxiv or medRxiv, or if appropriate, as a Research Advance in eLife, either of which would be linked to the original paper.

Summary:

The actin cytoskeleton has the unique ability to propagate mechanical stimuli at cell-cell and cell-matrix interactions, resulting in the maturation of adhesion complexes. The actin filaments that are placed under tension in these regions create unique binding sites for actin-binding proteins. Vinculin and α-catenin have been shown differentially bind to tensed and relaxed actin filaments, but the mechanism remains elusive. Here, Mei et al. address this important question. They combine beautiful cryo-EM structures of actin decorated by metavinculin or α-catenin with optical trap and TIRF motility assays to explore the how actin binding proteins interact with actin under tension.

Revisions for this paper:

The manuscript is much improved and the additional replicate experiments and modified analysis of optical tweezer data strengthen the results. The varied experimental approaches used by the authors are impressive.

The reviewers still have concerns about the TIRF motility data (listed below). Their suggestion is to either address these directly now, or to discuss in full the caveats to the TIRF experiments and explain how they will address them in future work.

1) The ABD concentrations employed in the TIRFM experiments remain a source of concern. Instead of resorting to speculations about whether they might not be at saturation, the reviewers feels the authors should try to perform experiments to find out. A recent preprint by the Weis and Hanein labs shows that the presence/absence of the N-terminal H0 can drastically affect a-catenin affinity. Given that a-catenin ABD intensities are close to background levels in TIRFM, the use of a shorter and high-affinity construct might be helpful.

2) The authors opted to not repeat the TIRF motility experiments using single filament densities as requested. While the single motor control experiments (Figure 2—figure supplement 2) are in principle suitable controls, it is concerning to see so little specific filament binding of a-catenin in the raw data. As detailed also in the previous comments by reviewer 3, the authors are encouraged to invest some time in optimizing their TIRFM assay to perform these important controls.

3) The authors should comment about their findings in relation to the study from Weis' lab.

https://doi.org/10.7554/eLife.62514.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 actin cytoskeleton has the unique ability to propagate mechanical stimuli at cell-cell and cell-matrix interactions, resulting in the maturation of adhesion complexes. The actin filaments that are placed under tension in these regions create unique binding sites for actin-binding proteins. Vinculin and α-catenin have been shown differentially to bind to tensed and relaxed actin filaments, but the mechanism remains elusive. Here, Mei et al. explore the binding of vinculin and α-catenin to actin under tension to address this important question.

The authors first develop an assay to pull single actin filaments and assess the binding intensity of vinculin and α-catenin. Vinculin did not show differential binding to filaments under high or low pulling force, but α-catenin showed a mild preference for filaments pulled beyond 1 pN. To validate this result, the authors developed another assay where opposite polarity myosin motors were used to place actin filaments under tension. This showed that α-catenin binding was enhanced when ATP was added, whereas vinculin binding was unchanged. The authors obtain beautiful cryo-EM structures of actin decorated by metavinculin or α-catenin at high resolution. The major binding site for the two proteins on actin was largely similar, whereas the minor binding site was distinct. In particular, the C-terminal extensions followed different paths on actin, and the extreme 35 amino acids on α-catenin were unresolved. The authors speculate that these unresolved residues mediated the force-sensitivity. Truncating these residues abolished force sensitivity, but switching the major binding region of α-catenin with vinculin did not, suggesting that the C-terminal extension of α-catenin is necessary and sufficient to mediate force-sensitivity. While this work provides important insight into how forces can be propagated at tensed cellular environments, I have a few major concerns that should be addressed by the authors.

1) In Figure 1, high force was taken to be the 5 frames before breakage and low force to be 5 frames before that. Therefore, a specific amount of time is taken to represent high versus low force, which is not appropriate. Furthermore, the low number of replicates raises questions on the robustness of the result. Therefore, I find that 1D and E weaken the author's claims, which are better represented in 1F and G, where the data is plotted against force. In the latter, however, it is not clear why 1 pN is chosen to divide the data. A regression analysis is more appropriate, with R2 values reported.

We appreciate the reviewer’s concerns with the low-force / high-force analysis. We have significantly revised the optical trapping portion of our manuscript, notably including substantially more trials. Out of 183 optical trapping traces, only 13 fit our criteria for low-force / high-force analysis, and only 21 fit our criteria for single-filament analysis.

As the reviewer surely appreciates, tether-to-tether variability is a major limitation of these experiments. We thus respectively disagree that the single tether analysis weakens our claims; rather, we believe it is important to show that we do see an increase in a-catenin binding along individual tethers when force is applied. We do believe the 5 frame window is necessary to see this effect, based on the noise level of the data (Figure 1B, C). We believe our findings from this analysis justifies the subsequent analysis based on pooling multiple tethers.

Based on the comments of the other two reviewers, we have implemented a detailed procedure for ensuring that the force vs. binding analysis focuses on tethers composed of individual actin filaments. Thus, while our overall dataset contains 183 recordings, only 21 tethers were included in this analysis based on the stringent criteria we imposed.

Although it is possible that determining a suitable fit function would be possible if we had many more points, to our eyes binding transitions from force-uncorrelated to strong binding in an essentially step-like fashion, within the noise level of the data (see Figure 1B and 1C: we now state this explicitly in the text). Therefore, regression analysis is not a feasible choice. Regardless, the reviewer’s point that 1 pN is a subjectively determined threshold is well-taken. We have performed K-means clustering analysis to more objectively identify the cutoff force value directly from the data in both plots, which we describe in the Results and the Materials and methods section. We have since used these force values, as well as the original 1 pN criteria, to divide the data in Figure 1F and Figure 1G. We observe a significant difference for a-catenin, but not vinculin, in both cases, suggesting this analysis is not sensitive to chosen threshold force.

Finally, we hope the reviewer can appreciate the low-throughput nature and low efficiency of these experiments. Out of 183 recordings, only 21 tethers could be included in the stringent single-filament subset, a “yield” of 11%. Under the current working conditions imposed by the ongoing COVID-19 pandemic with social distancing restrictions, our additional data was acquired with substantial difficulty. It is currently not feasible to obtain the massive datasets that would likely be required for a meaningful, well justified regression analysis as proposed by the reviewer.

2) A necessary assumption in the myosin assay is that myosins behave similarly in all experiments. However, this is not validated and may not be the case. For example, Video 7 (ABD-NCSwap) clearly has more bundled actin filaments after ATP addition than Video 6 (ABD-ΔC) and Video 8 (ABD-NCSwapΔC). The authors even hint that myosin inhibition can occur in certain cases. Additional experiments to assess myosin activity is required for these data to be reliable. A suitable assay would be myosin gliding in the presence of the different constructs, for example.

We thank the reviewer for this suggestion. We have performed single-motor experiments with the wild-type constructs. As noted by the all the reviewers, significant actin bundling occurs with both constructs in the dual motor experiments. We find that significant bundling also occurs in the presence of either the a-catenin ABD or the vinculin ABD with either myosin V alone or myosin VI alone, which unsurprisingly disrupts processive motility. This result is not surprising as any kind of actin movement, driven by either one myosin motor or two myosin motors, will help actin filaments sample larger space and cause bundling in the presence of an actin-bundling protein. Note that we are confident the motors are active (we always perform control gliding filament assays with both individual motors in the absence of ABPs where we do observe processive motility, data not shown). Therefore, a detailed quantification of the impacts of these constructs on myosin activity (e.g. processive gliding) is not possible, and we did not extend this analysis to all of the constructs employed in our work, which would be a substantial effort with limited potential for meaningful interpretation.

However, fortuitously we find that neither the a-catenin ABD nor the vinculin ABD displays ATP-enhanced binding in the presence of either motor alone, despite the presence of bundling. This strongly suggests that the increased a-catenin binding is due to the tug-of-war between motors of opposing directionality, rather than simply being an indirect effect of bundling. We have thus included this data in the revised manuscript (Figure 2—figure supplement 2) which we believe substantially strengthens our conclusions.

3) Metavinculin is used for structural studies to prevent bundling, and the authors validate its use by stating that it is similarly force-insensitive in the myosin assay. However, this is represented by a single experiment in Figure 2—figure supplement 1D, while it is clear from Figure 2—figure supplement 1B that there is significant variability in this type of data. Therefore, the use of metavinculin as a proxy for vinculin in structural analyses is not properly justified. Additional replicates will be required.

As requested, we have performed additional replicates, which support our conclusion that metavinculin is force insensitive. This data is included in Figure 2—figure supplement 1D, supporting our use of metavinculin in the structural experiments. We believe use of metavinculin is also supported by the predictive power of the structures in delineating the force detector of a-catenin.

4) In testing whether the D-loop interactions contribute to force-sensitivity, mutations are made in α-catenin to resemble metavinculin at the D-loop interface. The mutations abolish binding to actin altogether, and an interpretation provided by the authors is that "this interface does not have a separable role in force-activated actin binding". This interpretation cannot be correct if no binding was detected altogether, and the authors should remove the statement.

The reviewer’s point is well-taken. We have adjusted this statement to: “While these data suggest that the α-catenin D-loop interactions contribute to overall affinity for F-actin, the complete lack of binding is refractory to determining whether this interface has a separable role in force-activated actin binding.”

5) The motivation to truncate the disordered region of the C-terminal extension of α-catenin is not entirely clear. The residues of the C-terminal extension preceding this disordered stretch are clearly different between α-catenin and vinculin in the two cryo-EM structures. This region is therefore also a good candidate force sensor. The authors need to explain why they didn't switch this region between the two proteins.

We thank the reviewer’s comment on this issue. The protein residues we have switched in order to make the chimera, a-catenin residues 837-871, does include this region. The reason why we did not truncate this region is because a previous study has clearly shown that truncating this region of a-catenin will abolish its actin binding (Pokutta et al., 2002). As we have discussed in the main text this is consistent with our structure in which we have found this region directly forms a binding interface with actin.

Reviewer #2:

Using distinct biophysical assays, Mei et al. investigate the effect of pN scale forces on the interaction between actin filaments and two adhesion-associated actin-binding proteins (either α-catenin or its homolog vinculin). They show that forces moderately promote the binding of α-catenin but not vinculin. Extending their previous structural work on (meta) vinculin, they then determine high-resolution structures of the ABDs of either protein on actin filaments in the absence of force, which reveal differences in the C-terminal tail regions. Using chimeric constructs, they provide additional biochemical support for the C-terminus of α-catenin conferring force sensitivity.

Overall, the paper clearly contains several strong elements that merit publication. It focuses on a pertinent question of broad interest (the molecular mechanisms of force sensing in the actin cytoskeleton) and the data quality is high throughout. The TIRFM motility assay is a useful addition to the field and the observation of force-sensitive binding of α-catenin is in itself novel and interesting.

The central weakness of the paper is that it falls rather short in providing answers to the most pressing questions: The nature of the force-induced conformational change in actin filaments remains ultimately unclear. Similarly, the "force-sensing" C-terminal element of α-catenin (residues 872-906) is not resolved in the cryoEM structure. Hence, the detailed mechanism of force sensing remains to be elucidated.

That being said, I still feel that the paper deserves to be published even in the absence of a comprehensive structural mechanism. My main points rather address some of the immediate technical and conceptual issues that require improvement.

We of course agree with the reviewer that visualizing force-evoked structural rearrangements in F-actin is the critical next step in dissecting the mechanism of force-activated binding, as we clearly state in the Discussion. However, as this effectively requires developing a new structural biology method, we reaffirm the reviewer’s point that this is not a reasonable request for revision of our current paper. The current manuscript is not data-light, and we believe it provides a substantial advance that sets the foundation for pursuing a structure of a-catenin bound to F-actin in the presence of force as a follow-up study.

1) Both vinculin and α-catenin ABDs are known to bundle actin filaments. I was therefore surprised to see that the authors seem to neglect potential confounding effects that bundling might cause in their reconstitutions. Both TIRFM and optical trapping experiments are done in a configuration in which filaments appear to be clearly bundled. The authors even mention that filaments become more strongly bundled in the TIRFM assay upon ATP addition. I am concerned that some of their observations might be equally explainable by differential effects of bundling/filament overlap configurations on vinculin and α-catenin. The authors need to perform control experiments under conditions where they clearly observe and analyze only individual actin filaments to demonstrate that the effect of force is indeed based on a conformational change in the filament itself.

The reviewer’s point is well-taken. We have acquired substantially larger optical trapping datasets for both vinculin and a-catenin and implemented a detailed procedure to focus our force vs. binding analysis on tethers composed of individual actin filaments (Figure 1F, 1G). This analysis still clearly shows a force-dependent binding increase only for a-catenin.

Additionally, at the recommendation of reviewer 1, we have performed TIRFM reconstitution experiments in the presence of individual myosin motor proteins (myosin V alone and myosin VI alone). Fortuitously, these experiments also show increased bundling in the presence of ATP, but no evidence of increased a-catenin binding (Figure 2—figure supplement 2, Video 9-12). This strongly suggests that increased binding in the dual motor TIRFM experiments is not simply due to bundling.

We believe that collectively these new experiments convincingly make the case that force activated a-catenin binding occurs at the level individual filaments, and have substantially strengthened our paper. Furthermore, we believe demonstrating that force-activated binding also occurs in context of actin bundles is a strength, not a weakness, since this is the cytoskeletal architecture recognized by a-catenin in vivo, a point which we have emphasized in the text.

2) The authors should be more explicit and conscious about the ABD concentrations employed in their in vitro assays. This information can currently only be found in the Materials and methods section (should be part of the figures/captions!). Why did they chose to work at 2uM for either protein in buffers of low ionic strength? Previous measurements show that both α-catenin and vinculin ABDs binds with sub-micromolar affinity to filaments (Hansen et al., 2013 and Le Clainche et al., 2010), which means that the authors work at or close to saturation. This should mask or at least severely diminish the effects that force might exert on ABD binding. In my opinion, the authors need to take much greater care of the fractional occupancy of filaments by ABDs (by measuring the ABD affinities) under their experimental conditions to ensure that they work still in a sensitive concentration regime. Determining exact affinities in the presence and absence of force in the TIRFM assay also ought to give them additional information concerning the relative number of binding sites at saturation (maximal signal), which might allow them to discriminate between the proposed force sensing mechanisms.

We have now included the ABD concentrations in the figure legends. However, due to substantial variability in reported F-actin binding KDs in the literature, we believe the order of magnitude of the concentration is far more relevant than the exact value. For example, Rimm et al. reported in their 1995 PNAS paper that the a-catenin ABD has a 3μM affinity to F-actin. We chose 2 μm because based on numerous previous reports, including the ones by Hansen et al. and Rimm et al., both proteins have essentially μM affinity for F-actin. 2 μm, in our hands, is the lowest concentration we can see significant effects for the force-sensitive actin binding for a-catenin.

Hansen et al. reported approximately 0.5 μM F-actin affinity for the a-catenin ABD. Note that we observe very little a-catenin binding in the absence of force in the TIRFM experiments (Figure 2C), explicitly contradicting the reviewer’s assertion that we are working close to saturation. Hansen et al. noted distinct behaviors of N-terminally labelled GFP a-catenin ABD and unlabeled a-catenin ABD, so a potential source of the apparent lower affinity of a-catenin in our hands is the presence of a C-terminal halo tag. It is also plausible that there are discrepancies in our measurements of protein concentration. We used a Bradford assay calibrated with BSA (as noted in the Materials and methods), while Hansen et al. did not report their method of protein concentration measurement.

Le Clainche et al. performed co-sedimentation studies to examine the affinity of the vinculin ABD for F-actin (Figure 4D of their paper). Although it is difficult to exactly compare cosedimentation assays and our TIRFM studies due to the many differences in the conditions (immobilization of F-actin, etc.), at the point in their curves most closely related to our assays (2 μM vinculin ABD and 1 μM actin), they report that ~25 % of the vinculin ABD is bound, i.e. 0.5 μM. As the maximum stoichiometry of vinculin ABD to actin protomers is 1:1, this would correspond to 50% of sites being occupied, precisely the range where we would expect to see sensitivity to additional modulating factors of binding such as force.

Although it is once again difficult to compare assays with different modalities, we found it required an order of magnitude higher concentrations of both ABDs (10 μM for metavinculin and 20 μM for a-catenin) to achieve saturation in our cryo-EM studies. While we find it is common that higher concentrations are required to achieve saturation in cryo-EM of ABP-Factin complexes, this nevertheless suggests that the concentrations employed in our biophysical experiments are not nearing saturation.

Finally, we appreciate the reviewer’s suggestion that it would be useful to explicitly determine the effects of force on binding affinity by performing our TIRFM experiments at different concentrations. As noted in the text, there is considerable variability between replicates even when the assay is performed with identical concentrations. We thus do not believe it would be feasible to perform these experiments unless the TIRFM assay is systematically improved, which we believe is an appropriate subject for a subsequent follow-up study, which we now note in the Discussion.

In summary, we believe that we are not working close to saturation, and that performing systematic studies of the quantitative impact of force on ABD affinity are not currently feasible as a revision to our present manuscript. We were very careful in our writing to make clear that, although we quantitate our data to support our conclusions, the fundamental phenomenon we report is simply an “increase” in F-actin binding by the a-catenin ABD in the presence of force. A quantitative dissection of Kd vs. force would constitute a study in itself, which would require substantial refinement of the methods we introduce in this paper.

3) The "tug of war" motility assay employed by the authors, while ingenious, suffers from a central weakness. The opposite polarity motors are equally likely to produce tensile AND compressive forces on the surface-bound filament. This important limitation is not discussed by the authors at all. Admittedly, the optical trapping assay shows that the effect can also be observed for tensile forces only. However, the magnitude in the TIRFM assay is much greater, which is why one cannot help but wonder whether compression is the true binding-promoting cue. I am fully aware that this cannot easily be addressed experimentally. However, the authors should be more open in discussing this alternative interpretation.

We are certainly aware of the multiple types of forces that can in principle be generated in our dual-motor assay. We believe it is unlikely that compression is the dominant binding promoting cue, since many tethers in our optical trapping experiments are bent at the onset of the experiment (e.g. Figure 1B), suggesting they are compressed beyond their resting contour length when captured in the microfluidic flow cell, but do not show increased a-catenin binding. Regardless, we have included a sentence about the different motor forces generated in the TIRFM assay, including torsional forces, to the Results section. Dissecting these different forces through elaborations and improvements to the assay will be an important subject for future studies.

4) The "steric exclusion" model put forward by the authors posits that adjacent α-catenin ABDs interfere with synchronous binding. How can this be reconciled with the observation of cooperative α-catenin ABD binding (Hansen et al., 2013), indicating stabilizing effects? The authors cite and superficially discuss this point, but ultimately do not provide a satisfactory explanation. Is there any indication of partial binding in their cryoEM data? Along the same vein, have they compared the binding stoichiometry of α-catenin (either wt or C-terminal deletion mutants) at saturation via TIRFM or co-sedimentation?

We thank the reviewer for the insightful comment. We believe these two theories, our “steric exclusion” model and the cooperative binding of α-catenin, are not mutually exclusive. It is plausible that an α-catenin molecule can have both cooperative and inhibitory effects for a neighboring α-catenin molecule to bind actin: the cooperativity can facilitate its engaging, while the steric exclusion can reduce it. For α-catenin’s actin binding interaction, having binding cooperativity does not rule out the possibility that there may still be steric hindrance between neighboring molecules globally reducing binding, which can be alleviated by force on actin. Indeed, these opposing interaction modes provide multiple axes for regulation which could be harnessed by the cell. We have added an expanded discussion about this issue in the main text (subsection “Molecular mechanism for detecting force on actin through a flexible structural element”).

We are confident that actin filaments are fully decorated in our Cryo-EM data. Our lab has extensive experience in analyzing heterogeneity in cytoskeletal complexes, including occupancy (e.g. Kim et al., 2016). As indicated in the Materials and methods, we do not see any evidence of partial occupancy in the cryo-EM data through classification methods, and we believe if there was partial binding we would have found it. Our structure clearly shows that, at saturation, the binding stoichiometry is 1:1, which we now explicitly emphasize. There may be some confusion as the a-catenin binding site spans two actin protomers, but keep in mind that each actin protomer also contacts 2 a-catenin molecules.

As the binding stoichiometry of the wild-type is 1:1 with the actin protomer (e.g. the maximum possible), it is not plausible that the C-terminal deletion mutant would have a higher binding stoichiometry. We thus do not believe it is meaningful to examine binding stoichiometry through other methods, since it is clearly maximal for the wild-type as indicated from the structural data.

Reviewer #3:

Mei et al. present a very interesting study of force-dependent binding of actin-binding domains to filamentous actin. Using optical trapping, they demonstrate forces > 1pN are sufficient to show enhanced actin binding of a-catenin, a process they call force-activated binding, but not vinculin. The authors identify a C-terminal motif of a-catenin that is required for force-activated binding and that confers similar behavior to vinculin when added. Using cryoEM, they obtain structures of both a-catenin and vinculin bound to actin, showing where the C-terminal motif binds. This work provides interesting new information about how forces on filaments can influence actin-binding proteins. However, the mechanistic insight is limited by the lack of data on how a 1pN force alters actin and how a-catenin might sense that change. In addition, I have concerns about interpretation of the optical trapping and TIRF data and am unclear why constructs and labeling schemes vary across the different experimental methods used.

We thank the reviewer for their overall appreciation of the significance of our findings. While we of course agree that a structure of a-catenin bound to F-actin in the presence of force will be very insightful, this requires development of a novel structural biology method, and thus goes significantly beyond the scope of a revision. Beyond identifying a-catenin as a force activated F-actin binding protein (itself a significant advance), we nevertheless believe that our current work provides significant insights into the mechanism, notably the identification of the force-detector through our combined cryo-EM structural studies and functional biophysics approach. Beyond constraining plausible biophysical mechanisms for future studies, this finding immediately enables cell biological experiments to investigate the role of a-catenin’s force activated binding in vivo by examination of the DC mutant. Thus, we believe the significance of our work will be broadly appreciated by the readership of eLife.

1) The optical trapping results appear to pool data on single filaments and bundled filaments (as noted in the Materials and methods), which makes interpretation of the data very difficult and raises questions about the conclusions.

a) How many single filaments vs bundled filaments are included in the data set? The caption of Figure 1 indicates that it shows data on individual filaments, but the images of actin do not look like single filaments. For example, in the series of images of actin filaments in Figure 1C, the 4th actin filament looks like it could be several filaments, and the 5th image seems to show a broken filament. I'm not convinced that a single rupture force conclusively shows that only one filament is being imaged, as all filaments need not be load bearing and may only be laterally associated with a filament that is attached to the bead. Could a calibrated measurement of filament fluorescence intensity be used to quantify the number of filaments in a bundle?

We very much appreciate this point, which is similar to ones brought up by reviewers 1 and 2. Broadly, we have collected substantially expanded optical trapping datasets, which enabled us to institute a strenuous series of criteria for excluding multi-filament tethers, described in the text and Figure 1—figure supplement 2, for the force vs. fluorescence analysis displayed in Figure 1F-G. To specifically address the case of laterally associated filaments which are not loadbearing: Motivated by the fact that such filaments are unlikely to be exactly the same length as the load-bearing filament, we examined fluorescence line-scans of the actin intensity. These show a clear step-like drop in intensity when such filaments are present (Figure 1—figure supplement 2C), enabling us to exclude them from further analysis.

b) It seems that the amount of force felt by each filament will depend on precisely how each is coupled to the beads and to each other, which is hard to know. Won't the inclusion of actin bundles lead to an error in estimating the force at which binding of a-catenin increases?

The reviewer’s point is well-taken, and we now only include single-filament tethers in the force vs. binding analysis in Figure 1F-G. Additionally, we have implemented an objective clustering based analysis to identify the threshold force for the binding increase, as discussed above in our answer to the reviewer #1.

c) How do the force-extension curves obtained in this study compare with previous measurements of actin force-extension, such as Liu and Pollack (Biophys J, 2002)? This might help to identify single vs bundled filaments.

We appreciate this suggestion. The rupture forces reported by Liu and Pollack with cantilevers for single filaments were generally > 200 pN, i.e. 2 orders of magnitude greater than the modal rupture force for single filaments that we now report (6 pN), with tethers surviving several cycles of stretch. While it is possible that the different apparatuses employed (including our dual-color imaging), and differences in loading rate and/or tethering chemistry, can give rise to this discrepancy, and we do not want to cast aspersions on the pioneering work of Liu and Pollack, it is unclear how these authors ensured they were pulling on single filaments. We believe that the very large discrepancy makes it difficult to directly compare the force extension curves.

Two of our co-authors (M. Reynolds and S. Liu) recently developed a novel analysis procedure for analyzing DNA force-extension curves (Leicher…Liu. Biorxiv, 2019) to estimate persistence length. We attempted to apply this analysis to our data, but did not observe a consistent difference between clear multi-filament tethers and single filament tethers, likely due to deviations from the WLC model for actin, the low rupture forces, as well as the generally short length of our recordings for single filaments, limiting power of the fits.

We thus believe that, while imperfect, examination of the breaking force distribution is the best analysis for distinguishing individual filaments, when coupled with our other criteria for excluding multi-filament tethers.

d) How do the authors correct for filament length in their analysis? The effect of 1pN on a fluctuating filament will depend on the length of the filament.

We observe no systematic relationship between the force dependent increase in binding (the difference between high-force and low-force averages) and filament length, or breaking force and filament length (see Author response image 1). We thus have not corrected for filament length in our analysis.

Author response image 1

e) In the vinculin ABD binding experiment in Figure 1B, why is the relative ABD intensity varying so much (~30%) if binding of the ABD has equilibrated on the filament? And how is the intensity ratio normalized in Figure 1F and 1G?

We believe the intensity is fluctuating due to the variations inherent to a single-molecule pulling experiment. Sources of variation include fluctuations in focus, fluctuations in binding density along the tether, performance of the fluorophores, and performance of the detector. We believe the IABD/Iactin analysis is the best possible correction for these factors, yet variations do remain.

The intensity ratio in Figure 1F-G is normalized on a tether-by-tether basis by the largest value in the image sequence for each tether, which is stated in the Materials and methods.

2) The myosin TIRF experiments also raise several questions:

a) Are the filaments imaged in the TIRF experiments single filaments or bundles (or both)? As discussed above, can calibrated fluorescence intensity be used to identify single vs bundled filaments?

It is likely that both single filaments and bundles are present in the TIRF data, which we explicitly note in the text. At the reviewer’s suggestion, we analyzed the distribution of raw actin intensity values of all filament regions in a representative video. If were able to distinguish bundles from individual filaments, we would anticipate observing a multi-modal distribution, where each mode represents the number of filaments in a bundle. Instead, we observe a unimodal distribution (see the plot in Author response image 2), suggesting that we cannot discriminate the number of filaments in each region from intensity alone. We nevertheless believe the bundles are physiologically relevant: indeed, actomyosin bundles are the primary substrate of a-catenin and vinculin in vivo. We thus believe force-activated binding to bundles is an important finding, which we emphasize in the text.

Author response image 2

A point of concern raised by the other reviewers is that bundling increases upon activating the myosins, which is correct. At the suggestion of reviewer 1, we have now conducted singlemotor experiments (Figure 2—figure supplement 2), which, while still showing an ATP-dependent increase in bundling, show no increase in actin binding by either ABD. We believe these experiments convincingly demonstrate that force-activated binding is not simply a product of increased bundling in the myosin-motor reconstitution experiments.

b) Won't some filaments in the myosin TIRF assay experience compression while others experience tension? What about twist? How does this affect data interpretation and the force that is responsible for a-catenin force-activated binding?

We agree with the reviewer that tension, compression, and torque may all be present in the TIRF assays. We have thus carefully stated that “myosin generated forces” are sufficient to activate a-catenin binding. While the optical-trapping experiments suggest tension is sufficient to activate binding, we have now included an explicit discussion of this issue in the text.

c) The authors note the variability in the assay. Any thoughts on the source of this variability? Would a lower density of filaments in Figure 2B and 2C enable easier data collection? Also, the variability in the TIRF data shown in Figure 6—figure supplement 1 is puzzling. Why are some values of normalized intensity negative?

We would be delighted to conclusively know the source of variability in these experiments, as we would then hopefully be able to eliminate it. Our current best guess is batch-to-batch variation in our cover-glass preparation procedure, which strongly affects the amount of background present in our experiments. While we do perform a background subtraction step for quantification, systematic variations may nevertheless affect the IABD / Iactin distribution. We now explicitly include a discussion of this issue in the Results section.

Obtaining an appropriate filament density is a delicate balance, as too low of a filament density tends to result in the filaments being quickly shattered by the motors, precluding analysis. Too many filaments prevent the reliable identification of individual regions. The actin concentration we employed was optimized in light of these considerations.

Negative values are expected to occasionally arise due to Poisson distributed shot-noise on the camera. In filament regions featuring very low binding (essentially zero), it is formally possible that the local background will randomly contain slightly more counts than the region itself, producing a negative value for the IABD upon background subtraction. The Iactin will always have a positive value, since actin intensity is used for thresh-holding and region identification. Therefore, IABD/Iactin can occasionally have a (small) negative value, which is indeed what we observe.

3) The cryoEM data is very impressive. While the structures of a-catenin and vinculin ABDs on actin are well done, they don't provide insight into how force-activated binding might be happening.

Respectfully, we strongly disagree with this statement. The cryo-EM structures explicitly enabled us to precisely identify the force detector in a-catenin. Furthermore, our design of the vinculin-a-catenin chimera which gains force-activated binding was enabled by the structures, which provides perhaps the strongest evidence in our paper that the a-catenin C-terminus is the force detector.

a) While imaging filaments under a load in cryoEM would be very difficult, can the authors identify two different classes of filaments in their data, analogous to their "low force" and "high force" grouping of trapping data in Figure 1? If a force of only 1pN is necessary to drive enhanced binding, perhaps filaments under forces that are above and below that may be captured during freezing, which could provide valuable mechanistic insight.

We observe no evidence of this type of variability in our dataset, as described in extensive detail in the Materials and methods, as well as Figure 3—figure supplement 1. As our lab has extensive expertise in analyzing heterogenous filament datasets (e.g. Kim et al., 2016), we believe if it was there, we would have found it.

We realize that there is literature suggesting that the conditions of cryo-EM sample preparation expose actin filaments to tensile forces, which we cite in the Introduction. It is our opinion that the nature of these forces remains speculative, and we are not comfortable building upon this speculation in our work without well-controlled experiments. We are currently pursuing such studies, which are of sufficient scope to be the subject of an independent follow-up paper.

b) Was a structure of the vinculin ABD with the a-catenin C-terminal motif solved to confirm that it interacts with the filament in the same way as it does on a-catenin?

We did not obtain such a structure, which would be a massive investment of time and resources which are currently quite limited during the ongoing COVID 19 pandemic. We believe such a request is outside the scope of what is normally considered appropriate for a revision. While there is formally a very small chance that this chimera adopts a different binding mode, this is far from the most parsimonious explanation for our data.

4) The choice of filament labeling strategies and ABD used appears to vary from experiment and would benefit from more explanation and/or experiments with the same constructs.

a) In the cryoEM structures, metavinculin was used rather than vinculin to avoid bundling. Why wasn't metavinculin used in the optical trapping and TIRF experiments, given that bundling complicates analysis and interpretation of the data?

As the reviewer might expect, the studies described in our paper evolved over the course of several years. As vinculin is the canonical isoform expressed in most cells, we chose to employ this for our initial studies. We then turned to structural studies to interrogate the mechanism, where we chose to focus on metavinculin to avoid bundling. As this structure had substantial predictive power when we engineered vinculin mutants, we believe this choice was well-justified.

b) The optical trapping experiments used phalloidin-labeled actin filaments, while the TIRF experiments used rhodamine-labeled actin filaments. Why weren't the experiments done with the same actin labeling method?

Although presented in the manuscript with the optical trapping experiments first, we actually performed the dual-motor TIRF experiments with the wild-type constructs first during the course of our studies. At the time, we were concerned that phalloidin (which stabilizes F-actin) would prevent force-dependent conformational changes in F-actin recognized by mechanosensitive ABPs. We thus chose to employ rhodamine-labeled actin. Once we discovered a-catenin’s force-activated actin binding activity, we then pursued the optical trapping studies to validate this observation and dissect the contribution of tension from other motor-dependent phenomena (compression and torsion, as noted by the reviewer).

As the optical trapping experiments had to be performed at very low F-actin concentrations in order to have a chance at capturing individual filaments, they necessitated the use of phalloidin. Fortuitously, we found that phalloidin did not inhibit force-activated binding. We have also performed control experiments where we visually confirmed that un-labelled phalloidin does not impact force-activated binding to rhodamine-labelled filaments in myosin TIRF reconstitution experiments (data not shown).

We believe that demonstrating a-catenin’s force-activated binding to be independent of the actin-labelling strategy employed is a strength, not a weakness, of our manuscript.

5) The proposed models seem reasonable but would benefit from more consideration of fluctuations and comparison with other existing models.

a) It is unclear how the thermal fluctuations of a filament under physiological conditions would affect the proposed a-catenin steric exclusion and actin filament conformational change models. Won't the small changes associated with a force of 1pN already be sampled by thermal fluctuations? While detailed modeling is beyond the scope of the manuscript, some discussion of thermal fluctuations and the proposed models would be helpful.

We thank the reviewer for this suggestion. We agree that low piconewton forces affecting the distribution of actin conformations is a reasonable and attractive model. We believe either mechanism we propose could operate in this framework, where discrete conformations are substituted with conformational ensembles. We have added an explicit note to this regard in the revised Discussion, including referencing previous work suggesting this regulatory mechanism for cofilin.

b) The authors cite Ishiyama et al., 2018, which presents a different mechanism for a-catenin force dependence based on conformational unfolding of the ABD under force, but there is little discussion of it. Are the proposed models compatible with this?

The mechanism proposed by Ishiyama et al. concerns force-dependent rearrangements in the N-terminus of the ABD (H0-H1), specifically force-dependent unfolding. Our actin-bound structure does indeed show this region unfolds upon actin-binding, which could reasonably be stabilized by force. This is a mechanism that could explain catch-bond formation (we have previously proposed a similar mechanism to explain vinculin catch-bond formation with F-actin in a commentary: Swaminathan, Alushin and Waterman, 2017), but it cannot explain force-activated binding. This is because the entire ABD would have to be bearing tension in order for this region, which does not directly contact actin, to “feel” force. As our paper focuses on force-activated binding rather than catch-bond formation, and already features a lengthy discussion, we did not delve into this in detail. However, this mechanism (which we emphasize to be quite speculative) is discussed at length in the competing pre-print from Volkman, Hanein, Weis, and colleagues which also reports a (somewhat lower resolution) structure of the a-catenin ABD bound to F-actin, that has emerged while our paper is under review (and that we now cite). We believe substantial further work (e.g. visualizing the entire cadherin complex bound to F-actin in the presence of tension) will be required to establish the mechanism of a-catenin catch-bonding to F-actin in detail.

c) As a minor point, the authors say that they report the first direct observation of enhanced actin binding by an actin-binding protein. However, Hayakawa et al., 2011, which the authors cite, previously reported tension-dependent binding of cofilin to actin.

We thank the reviewer for pointing out this oversight. We have included a statement that tension dependent binding of cofilin to F-actin was previously reported by Hayakawa et al., and also called into question by a more recent paper from Wioland, Jegou and Romet-Lemonne, 2019. We have also systematically pruned our paper of excessive claims of novelty, which we do not believe impacts the significance of our findings.

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

Revisions for this paper:

The manuscript is much improved and the additional replicate experiments and modified analysis of optical tweezer data strengthen the results. The varied experimental approaches used by the authors are impressive.

The reviewers still have concerns about the TIRF motility data (listed below). Their suggestion is to either address these directly now, or to discuss in full the caveats to the TIRF experiments and explain how they will address them in future work.

Due to the related work which has very recently appeared from Hanein, Volkman, Weis, and colleagues (Xu et al., 10.7554/eLife.60878), also in eLife, we have elected to discuss these caveats of the TIRF studies and propose specific experiments to address them in future work. This has been included in the revised manuscript.

1) The ABD concentrations employed in the TIRFM experiments remain a source of concern. Instead of resorting to speculations about whether they might not be at saturation, the reviewers feels the authors should try to perform experiments to find out. A recent preprint by the Weis and Hanein labs shows that the presence/absence of the N-terminal H0 can drastically affect a-catenin affinity. Given that a-catenin ABD intensities are close to background levels in TIRFM (see below), the use of a shorter and high-affinity construct might be helpful.

We have emphasized that Xu et al. report a series of high-affinity N-terminal deletion constructs in the text, including a construct which lacks both H0 and H1, which has ~18 fold higher affinity for actin. We agree with the reviewer that examining a higher affinity construct, should it retain force-activated binding to actin, will likely be very useful for making accurate estimates of binding stoichiometry and their impact on force-activated binding. We will pursue these experiments as an immediate follow-up. The discussion of this caveat and our plan for addressing it are included in the revised manuscript.

2) The authors opted to not repeat the TIRF motility experiments using single filament densities as requested. While the single motor control experiments (Figure 2—figure supplement 2) are in principle suitable controls, it is concerning to see so little specific filament binding of a-catenin in the raw data. As detailed also in the previous comments by reviewer 3, the authors are encouraged to invest some time in optimizing their TIRFM assay to perform these important controls.

While our optical trapping studies support force-activated binding to individual filaments, we agree that it will be interesting to establish whether myosin motor activity is also sufficient to activate binding to individual filaments in addition to bundles. It may be feasible to achieve sufficiently low filament densities to avoid bundling in the presence of phalloidin, and we will pursue these experiments as an immediate follow-up. The discussion of this caveat and our plan for addressing it are included in the revised manuscript.

The outcome of these experiments (both points 1 and 2) will be posted as a bioRxiv pre-print and, if suitable, submitted as an eLife Research Advance. A statement to this effect has also been included in the revised manuscript.

3) The authors should comment about their findings in relation to the study from Weis' lab.

We have added detailed comparisons to the study of Xu et al., noting specific points of concordance and discordance between our papers in the Results section. This includes:

1) A quantitative comparison of the structural models and comments about their similarity.

2) Referencing the W859A mutant of Xu et al., which reduces actin affinity, in support of the importance of the tryptophan latch.

3) Noting the apparently conflicting results of a CTE deletion (in our hands, increased overall affinity and loss of force-activated binding in TIRF assays; in their hands, slight reduction in affinity in solution co-sedimentation assays), and potential reasons for this discrepancy.

We have furthermore included an additional paragraph in the Discussion regarding the catch-bonding model proposed by Xu et al., in relation to our present and previous work.

https://doi.org/10.7554/eLife.62514.sa2

Article and author information

Author details

  1. Lin Mei

    1. Laboratory of Structural Biophysics and Mechanobiology, The Rockefeller University, New York, United States
    2. Tri-Institutional PhD Program in Chemical Biology, The Rockefeller University, New York, United States
    Contribution
    Conceptualization, Resources, Data curation, Formal analysis, Validation, Investigation, Visualization, Methodology, Writing - original draft, Writing - review and editing
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0002-5056-4547
  2. Santiago Espinosa de los Reyes

    Laboratory of Structural Biophysics and Mechanobiology, The Rockefeller University, New York, United States
    Contribution
    Software, Formal analysis, Methodology, Writing - review and editing
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0003-4510-8296
  3. Matthew J Reynolds

    Laboratory of Structural Biophysics and Mechanobiology, The Rockefeller University, New York, United States
    Contribution
    Software, Formal analysis, Methodology, Writing - review and editing
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0002-2501-9280
  4. Rachel Leicher

    1. Tri-Institutional PhD Program in Chemical Biology, The Rockefeller University, New York, United States
    2. Laboratory of Nanoscale Biophysics and Biochemistry, The Rockefeller University, New York, United States
    Contribution
    Investigation, Methodology, Writing - review and editing
    Competing interests
    No competing interests declared
  5. Shixin Liu

    Laboratory of Nanoscale Biophysics and Biochemistry, The Rockefeller University, New York, United States
    Contribution
    Formal analysis, Supervision, Funding acquisition, Methodology, Project administration, Writing - review and editing
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0003-4238-7066
  6. Gregory M Alushin

    Laboratory of Structural Biophysics and Mechanobiology, The Rockefeller University, New York, United States
    Contribution
    Conceptualization, Data curation, Formal analysis, Supervision, Funding acquisition, Visualization, Methodology, Writing - original draft, Project administration, Writing - review and editing
    For correspondence
    galushin@rockefeller.edu
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0001-7250-4484

Funding

Irma T. Hirschl Trust (Research Award)

  • Gregory M Alushin

Pew Charitable Trusts (Pew Scholar Award)

  • Gregory M Alushin

National Institutes of Health (5DP5OD017885)

  • Gregory M Alushin

National Institutes of Health (1DP2HG010510)

  • Shixin Liu

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

Acknowledgements

We gratefully acknowledge Pinar Gurel (RU) for initial efforts in establishing the force reconstitution TIRF assay, Luke Lavis (HHMI Janelia) for the gift of JF-646 for pilot studies, and Yasuharu Takagi and James Sellers (NHLBI DIR) for the gift of myosin proteins and training in their purification. We also thank Sara Tafoya and Jordi Cabanas-Danés (LUMICKS) for assistance with C-trap experiments and data analysis, Muzaddid Sarker and Sharon Campbell (UNC) for the gift of metavinculin ABD used in cryo-EM studies, Rui Gong (RU) for assistance with cryo-EM data collection and analysis, and Mark Ebrahim and Johanna Sotiris (RU Cryo-EM Resource Center) for assistance with cryo-EM data collection. Template cDNA for α-catenin constructs was obtained from David Rimm via AddGene (#24194). This work was funded by grants from the Irma T Hirschl/Monique Weill-Caulier Trust and Pew Charitable Trusts to GMA, and NIH High-Risk High-Reward grants to GMA (5DP5OD017885, Early Independence Award) and SL (1DP2HG010510, New Innovator Award).

Senior Editor

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

Reviewing Editor

  1. Andrew P Carter, MRC Laboratory of Molecular Biology, United Kingdom

Publication history

  1. Received: August 27, 2020
  2. Accepted: September 21, 2020
  3. Accepted Manuscript published: September 24, 2020 (version 1)
  4. Version of Record published: October 26, 2020 (version 2)

Copyright

© 2020, Mei 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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