α-catenin is a key mechanosensor that forms force-dependent interactions with F-actin, thereby coupling the cadherin-catenin complex to the actin cytoskeleton at adherens junctions (AJs). However, the molecular mechanisms by which α-catenin engages F-actin under tension remained elusive. Here we show that the α1-helix of the α-catenin actin-binding domain (αcat-ABD) is a mechanosensing motif that regulates tension-dependent F-actin binding and bundling. αcat-ABD containing an α1-helix-unfolding mutation (H1) shows enhanced binding to F-actin in vitro. Although full-length α-catenin-H1 can generate epithelial monolayers that resist mechanical disruption, it fails to support normal AJ regulation in vivo. Structural and simulation analyses suggest that α1-helix allosterically controls the actin-binding residue V796 dynamics. Crystal structures of αcat-ABD-H1 homodimer suggest that α-catenin can facilitate actin bundling while it remains bound to E-cadherin. We propose that force-dependent allosteric regulation of αcat-ABD promotes dynamic interactions with F-actin involved in actin bundling, cadherin clustering, and AJ remodeling during tissue morphogenesis.
The mechanical coupling of intercellular adhesion proteins to the cytoskeleton plays a key role in balancing the integrity and plasticity of epithelial tissues. Mechanical tension generated by cortical actomyosin is transmitted through the epithelial sheet by adherens junctions (AJs), allowing contractile forces to change cell and tissue shape1,2. The cadherin-catenin cell adhesion complex is the major building block of AJs, and has a crucial function in the dynamic behaviors of epithelial cells, such as cell polarization and cell rearrangements3,4. The enormous versatility of cadherin-mediated cell adhesion in tissue morphogenesis and homeostasis requires catenin-dependent regulation of the dynamic cadherin-actin interface in response to variable tension.
α-catenin is an actin-binding and actin-bundling protein responsible for connecting the cadherin-catenin complex to filamentous actin (F-actin) at AJs5,6,7,8. It plays critical roles in development and tissue homeostasis across the metazoans9,10,11,12, and α-catenin gene mutations have been linked to a variety of physiological abnormalities13,14,15, including tumor metastasis16. The α-catenin family includes three paralogs expressed in amniotes, E (epithelial), N (neuronal), and T (testis and heart), as well as a single homolog expressed in invertebrates, such as Drosophila17. Monomeric α-catenin binds to cadherin-bound β-catenin and anchors the cell adhesion complex to the actin cytoskeleton7,18,19. α-catenin dissociated from β-catenin can homodimerize to promote actin bundling5, but the underlying mechanism and function of α-catenin dimers in cell adhesion have been controversial20,21 and remain to be clarified.
The structure of α-catenin (100 kDa) consists of three distinct domains. The N-terminal (N) domain (30 kDa) facilitates β-catenin binding and homodimerization in a mutually exclusive manner22,23. The central mechanosensitive modulatory (M) domain (40 kDa) contains a cryptic binding site for another F-actin-binding protein vinculin6,24,25,26,27. The C-terminal actin-binding domain (ABD) (28 kDa), which is connected to the rest by a flexible P-linker region28 (2 kDa), directly binds to F-actin, and closely resembles the vinculin ABD (vin-ABD)27,29. Unlike vinculin that forms an autoinhibitory head-to-tail interaction30, the unhindered αcat-ABD23,27 forms a catch bond with F-actin that stabilizes the interaction under tension8. However, the molecular basis of this catch bond is unknown, and the physiological significance of its distinctive mechanical properties has not yet been demonstrated.
Here we reveal that a force-dependent conformational change in the αcat-ABD allosterically regulates direct F-actin binding. Several lines of evidence suggest that α1-helix unfolding changes the conformational dynamics of the actin-binding site. Furthermore, the αcat-ABD in an activated state homodimerizes to facilitate actin bundling. Our data suggest that manipulation of the ABD-dependent mechanosensory function of α-catenin severely interferes with AJ remodeling in mammalian cells and Drosophila embryos. Surprisingly, not only loss but also gain of F-actin binding propensity dramatically compromises α-catenin function in morphogenesis. Based on these results, we propose a new mechanism of the force-dependent, dynamic cadherin-actin linkage regulated by the ABD of α-catenin.
Force-dependent unfolding of αcat-ABD enhances actin binding
The direct interaction between α-catenin and F-actin was demonstrated to be a catch bond8, an interaction that is stabilized by increased force31,32. Since the C-terminal tail (residues 865-906) of α-catenin is postulated to be part of the interface between the αcat-ABD and F-actin33,34,35, we hypothesized that a regulatory motif resides within or near the N terminus of ABD. We monitored the disassembly and reformation of AJs in α-catenin-deficient R2/7 epithelial cells36,37 expressing various αE-catenin deletion mutants (Supplementary Fig. 1a; Supplementary Table 1). We found that the deletion of residues 663-696 from the ABD was associated with an unusual accumulation of cadherin-catenin-F-actin complexes in the cytoplasm after trypsinization of cell monolayers (Supplementary Fig. 1b, c), and delayed reformation of AJs with a unique square wave-like arrangement (Supplementary Fig. 2a). Cells with these deformed junctions showed diminished tight junction barrier function compared to full-length αE-catenin (αEcatFL)-expressing cells (Supplementary Fig. 2b). In addition, the αEcat-ABD residues 663-906 expressed in R2/7 cells colocalized with actin-rich regions at the cell periphery (Fig. 1a), whereas an N-terminally truncated form of ABD (ABD*; residues 697-906) prominently accumulated along stress fibers and actin rods (Fig. 1a), consisting of tightly packed actin bundles (Supplementary Fig. 2c). These results suggest the αE-catenin residues 663-696 regulate the association of αcat-ABD with different actin assemblies (Fig. 1a), and are critical for the normal function of αcat-ABD in forming AJs and, consequently, epithelial differentiation.
Comparison of crystal structures of αcat-ABDs27,38 with the vin-ABD30 revealed several highly conserved motifs of α-catenin potentially involved in its unique actin-binding mechanism: an N-terminal α1-helix (αE-catenin residues 669-675), a β-hairpin (βH; residues 799-810), and a C-terminal tail (Fig. 1b, c and Supplementary Fig. 3). Considering that the α1-helix is part of the ABD truncation (residues 663-696) that resulted in abnormal F-actin association and a failure to form normal AJs in R2/7 cells (Fig. 1a and Supplementary Fig. 1b, 2a‒c), we sought to explore the potential role of α1-helix in the regulation of force-dependent αcat-ABD-F-actin interaction. We performed equilibrium and constant-force steered molecular dynamics (SMD) simulations of the αN-catenin ABD (αNcat-ABD) to gain insights into how α1-helix may respond to increasing mechanical tension at the cadherin-actin interface. To help discuss equivalent residues between αN-catenin and αE-catenin with different residue numbering (e.g., V795 of αN-catenin is equivalent to V796 of αE-catenin), henceforth the αN-catenin residues will be denoted by using the equivalent αE-catenin residue numbers accompanied by a subscripted ‘N’ (e.g., V795 as V796N) for clarity. The SMD simulations showed α1-helix unfolding after a constant pulling force was applied on αNcat-ABD for 60 ns (Fig. 1d, Supplementary Fig. 4a, b and Supplementary Movie 1). Interestingly, shortly before α1-helix unfolded (at ~45 ns), the side chain of V796N turned over from a cryptic position to an exposed position (Fig. 1e and Supplementary Movie 2). αN-catenin residues V796N and I792N are equivalent to the vinculin actin-binding site residues, V1001 and I99739 (Fig. 1c). These results suggest that the conformational flexibility of α1-helix and the dynamics of V796N are mechanically coupled within the αNcat-ABD. This mechanism would be consistent with catch bond formation, if the conformation change of α1-helix exposes V796N and enhances the bond strength between αcat-ABD and F-actin.
To assess whether the α1-helix affects the α-catenin-F-actin interaction, we performed in vitro actin cosedimentation assays with three ABD variants of αE- and αN-catenin (αE-catenin residue numbers are shown): a wild type form of ABD (ABD-WT; residues 652-906), an ABD with a structure-guided helix-1 mutation (H1) designed to unfold α1-helix (ABD-H1; RAIM670-673GSGS) (Fig. 1c), and an ABD with a partially deleted α1-helix (ABD-Δα1; residues 671-906)33. The structural integrity of αcat-ABD was not affected by these mutations (Supplementary Fig. 4c‒e). We observed a nearly two-fold increase in the cosedimented amount of either ABD-H1 or ABD-Δα1 compared to ABD-WT (Fig. 1f). These results indicate that the α1-helix attenuates the αcat-ABD-F-actin interaction, and alterations in α1-helix significantly enhance the F-actin-binding activity of both αEcat-ABD and αNcat-ABD.
α1-helix unfolding induces weak αcat-ABD homodimerization
To examine the structural details of αcat-ABD with enhanced F-actin binding, we determined crystal structures of αNcat-ABD-H1 (Fig. 2a, Supplementary Fig. 5, and Supplementary Table 2). The αNcat-ABD-H1 structure closely resembles the overall fold of αNcat-ABD-WT (PDB ID: 4K1O)27 (Supplementary Fig. 6a), except for the α1-helix residues. However, unlike the monomeric αNcat-ABD-WT structure, αNcat-ABD-H1 crystallized as a homodimer connected by two βH motifs (Fig. 2a and Supplementary Fig. 6b). The dimer interface involves L807N of the βH, which mimics M673N of α1-helix interacting with the hydrophobic patch in the αNcat-ABD-WT structure (Fig. 2b and Supplementary Fig. 6c). Moreover, the observation of αNcat-ABD-H1 dimerization, which occludes 3100 Å2 of solvent-accessible surface (Fig. 2a), in two distinct crystal forms (Supplementary Fig. 6b) provides a basis for further examining the physiological relevance of this ABD-dimer interface. Our NMR analysis of αNcat-ABD-H1 in solution showed that concentration-dependent chemical shift perturbations (CSPs) mostly occurred in the βH motif (Fig. 2c and Supplementary Fig. 7a, b). In addition, we observed the increased propensity for αEcat-ABD-H1 to dimerize, albeit very weakly, in a concentration-dependent manner compared to αEcat-ABD-WT by size-exclusion chromatography-coupled multiangle light scattering (SEC-MALS) (Fig. 2d and Supplementary Fig. 7c). These results further support that the unfolded α1-helix propagates the weak dimerization of αcat-ABD through the βH-dependent interface.
One functional implication for α-catenin dimerization is actin bundling5, which has been presumed to occur through N-domain dimerization of αcatFL20,27,38. However, the ability of α-catenin to homodimerize through the ABD suggests an alternative actin-bundling mechanism. Indeed, actin bundling assays showed that both isolated αEcat-ABD-WT and αEcat-ABD-H1 proteins are capable of actin bundling (Fig. 2e). We next examined the involvement of α1-helix and βH motifs in ABD-dependent actin bundling. A βH-deletion mutant (αEcat-ABD-ΔβH) and a construct carrying both the H1 and βH-deletion mutations (αEcat-ABD-H1ΔβH) were well folded (Supplementary Fig. 4d, e), but cosedimented markedly less with F-actin at a high centrifugal force (100,000×g), indicating that the ΔβH mutation inadvertently affected F-actin binding of αEcat-ABD (Fig. 2e). Nevertheless, αEcat-ABD-ΔβH displayed residual actin bundling, whereas αEcat-ABD-H1ΔβH was unable to bundle F-actin (Fig. 2e). These results suggest that actin bundling can be facilitated by ABD dimerization through the βH-dependent interface, as well as through an unknown mechanism involving the α1-helix in our assays. In addition, our NMR transferred cross saturation (TCS) experiments with 15N/2H-labeled αNcat-ABD-WT and unlabeled F-actin indicated that the ABD directly interacts with F-actin through α5- and α6-helices, likely involving I792N and V796N, and, unexpectedly, through α3- and α4-helices on the opposite side of ABD (Fig. 2f and Supplementary Fig. 7d). This finding may point to a secondary contact site involved in actin bundling (Fig. 2f). Collectively, these results support the view that α-catenin facilitates actin bundling through ABD homodimerization.
ABD mutations compromise AJ remodeling in cells and embryos
Our finding that α-catenin can dimerize and mediate actin bundling independent of the N domain implicates the AJ-associated pool of α-catenin in reorganization of the actin cytoskeleton. To determine how alterations of α1-helix and βH would affect cadherin-mediated cell-cell adhesion, we tested the function of α-catenin mutants in R2/7 cells and Drosophila embryos. First, we examined R2/7 cells stably expressing αEcatFL fused with monomeric GFP (Supplementary Fig. 8a). Cells expressing αEcatFL or αEcat-H1 showed the typical cobblestone appearance of well-adhered epithelial cells with consistent colocalization of α-catenin and actin at AJs (Supplementary Fig. 8b). In contrast, cells expressing αEcat-ΔβH, αEcat-H1ΔβH, or a construct that lacks ABD entirely (αEcat-ΔABD) did not form cohesive cell monolayers and showed increased presence of α-catenin in protrusions (Supplementary Fig. 8b). Similarly, both αEcatFL or αEcat-H1 cells formed three-dimensional spheroids on ultra-low-attachment plates, whereas cells expressing other mutants remained in a semi-aggregated state (Supplementary Fig. 8c).
To find out how the H1 and ΔβH mutations affect the cell-cell adhesive strength, we performed an epithelial sheet disruption assay40. αEcatFL or αEcat-H1 cell monolayers lifted as a continuous sheet from the culture plate upon dispase treatment prior to mechanical disruption (Fig. 3a), but αEcat-ΔβH, αEcat-H1ΔβH, and αEcat-ΔABD cell sheets disintegrated into numerous pieces (Supplementary Fig. 8d). Subsequent mechanical disruption of cell monolayers caused αEcatFL monolayers to fragment, whereas αEcat-H1 monolayers remained mostly intact (Fig. 3a, b). These observations indicate that monolayers formed by αEcat-H1 cells have increased resistance towards mechanical stress compared to αEcatFL cells.
Next, we challenged R2/7 cells in scratch wound assays. In contrast to unchallenged cells, αEcatFL cells at the wound front displayed punctate AJs connected to actin cables aligned along the wound edge, whereas αEcat-H1 cells formed less organized punctate AJs and actin assemblies (Fig. 3c and Supplementary Fig. 9a). High-resolution live-imaging revealed that αEcat-H1 AJs were less organized towards the wound front, resulting in unproductive cell-cell tugging events that appeared to interfere with forward sheet migration (Supplementary Movie 3). In fact, αEcat-H1, αEcat-ΔβH, and αEcat-H1ΔβH cells were all inferior to αEcatFL cells in wound closure, and no better than αEcat-ΔABD cells (Fig. 3d, e and Supplementary Fig. 9b). By tracking cells individually, we found that αEcat mutant cells moved with similar speeds as αEcatFL cells, but less persistently, contributing to overall reduced epithelial sheet migration (Fig. 3f, Supplementary Fig. 9b and Supplementary Movie 4). In addition, differences between αEcatFL and αEcat-H1 cell trajectories were independently validated using a particle image velocimetry (PIV)-based tracking method (Supplementary 9c). These observations suggest that α-catenin with a defective α1-helix can support AJs in static epithelia but fails to support dynamic AJ rearrangements and cell movements.
To further assess α1-helix and βH functions in tissue organization we generated mutants in Drosophila α-Catenin (α-Cat) and tested their function in transgenic animals. Both α1-helix and βH regions are conserved in α-Cat (Fig. 1c), and previous work showed that the ABD of Drosophila α-Cat (αCat-ABD) is essential for cell adhesion7. Moreover, αCat-ABD-H1 showed enhanced actin binding and bundling activity compared to αCat-ABD (Supplementary Fig. 10a) similar as mammalian proteins (Fig. 2e). Zygotic null mutants for α-Cat (αCat−/−) show embryonic lethality and severe defects in head morphogenesis9 (Fig. 4a, b). Expression of full-length α-Cat (αCatFL) did rescue αCat−/− mutants to adulthood. In contrast, αCat-H1, αCat-Δα1, αCat-ΔβH, and αCat-H1ΔβH did not rescue the embryonic lethality of αCat−/− mutants similar to αCat-ΔABD (Fig. 4a). Expression of αCat-H1, αCat-Δα1, and αCat-H1ΔβH led to some improvements in head morphogenesis, and a small number of animals expressing αCat-H1 or αCat-H1ΔβH survived to larval stages (Fig. 4a). Immunoblot analysis (Supplementary Fig. 10b) and tissue staining in an αCat−/− mutant background (Fig. 4c) showed that our constructs are expressed at levels similar to endogenous α-Cat and are effectively recruited to AJs. Efficient recruitment of α-Cat proteins to AJs in a wildtype background indicated that mutant proteins are not outcompeted by endogenous α-Cat (Supplementary Fig. 10c). We noted that overexpression of αCat-H1 or αCat-Δα1 had a toxic effect on survival with most animals dying as larvae, whereas overexpression of other α-Cat constructs led to pupal lethality or adult survival (Fig. 4a). The failure of α-Cat proteins with a compromised α1-helix to substantively rescue αCat−/− mutants was surprising as those variants are likely capable of coupling cadherin to the actin cytoskeleton to promote intercellular adhesion. On the other hand, enhanced F-actin binding of αCat-ABD-H1 (Supplementary Fig. 10a) could explain the observed toxicity upon overexpression of these constructs. Our findings indicate that the function of the α1-helix in attenuating interactions between α-catenin and F-actin is instrumental for AJ function in developing epithelia.
We further examined the role of α1-helix in wound repair, which is driven by the polarized assembly of actin at the interface between wounded and adjacent cells in the Drosophila embryonic epidermis41. Polarization of actin (and the non-muscle myosin II) in the cells adjacent to the wound results in the assembly of a supracellular contractile cable around the perimeter of the wound that drives tissue repair42. The quantified wound closure dynamics revealed that αCat−/− embryos expressing αCatFL repaired damage to their epidermis faster than αCat−/− embryos, whereas αCat-H1 expression did not significantly accelerate wound closure in an αCat−/− epidermis. (Fig. 4d, e). These results are consistent with our whole animal rescue experiments, as well as our scratch wound healing assays (Fig. 3d‒f), and collectively suggest that a compromised α1-helix severely interferes with α-catenin function in tissue morphogenesis.
Actin-binding site residues are essential for α-cat function
Considerable in vitro evidence suggests that α-catenin can directly interact with F-actin5,7,8,20,27,29,33,35,43,44. A previously determined low resolution (18 Å) cryo-EM map of an αcat-ABD-F-actin complex precluded any detailed analysis of the complex interface29. Nonetheless, it suggested that the αcat-ABD interacts with two actin monomers adjacently aligned on the long axis of F-actin. A similar arrangement was observed in a recently determined 8.5-Å cryo-EM structure of a vin-ABD-F-actin complex, which revealed that the last two α-helices of vin-ABD interact with F-actin45. Considering the relatively high sequence identity shared between αcat- and vin-ABDs (~30%)27, we generated an atomic model of the αNcat-ABD-H1-F-actin complex based on the vin-ABD-F-actin structure. In this model, α5- and α6-helices of the αNcat-ABD-H1 interact with two axially arranged actin monomers of F-actin (Fig. 5a). In particular, the α5-helix contains the highly conserved residues, I792N and V796N (Fig. 1c). I792N of αN-catenin assumes an exposed position closely resembling the vinculin actin-binding site residue I99730. In contrast, the conformation of V796 remains ambiguous, partly due to poorly defined electron density of this region in the 3.7-Å crystal structure of human αE-catenin38, and a cryptic position of V796N in the αNcat-ABD structure27 (Supplementary Fig. 11a) compared to the fully exposed V1001 of vinculin30.
To better characterize the αE-catenin actin-binding site, we elucidated a crystal structure of αEcat-ABD-WT at 2.2-Å resolution (Fig. 5b, Supplementary Fig. 5, and Supplementary Table 2). The electron density map of α5-helix clearly shows that V796 adopts a conformation that exposes its side chain on the ABD surface, along with two additional hydrophobic residues L785 and I792 (Supplementary Fig. 5g and 11b). Our site-directed mutagenesis and actin cosedimentation assays with the αEcat-ABD variants support critical roles of these hydrophobic residues in F-actin-binding: Ala substitutions of L785, I792 and V796, individually or together as 3A, led to a range of reduction (75, 36, 47, and 78%, respectively) in the amount of ABD cosedimenting with F-actin compared to αEcat-ABD-WT (Fig. 5c). The effects of I792A and V796A were greater in the H1 background (reduction of 70% and 73%, respectively), confirming that alterations of these residues significantly reduce F-actin binding by αEcat-ABD-H1 (Fig. 5c). In contrast, Ala substitution of V714, which is located on the α3-helix surface, resulted in no reduction (Fig. 5c). Also, none of the above mutations appear to interfere with the ability of αEcat-ABD to bundle F-actin (Supplementary Fig. 12). The equally significant reduction observed with either the L785A mutation alone or 3A suggests that L785 plays a central role in establishing the critical hydrophobic interface between F-actin and αcat-ABD. The measurable reduction in F-actin binding with I792A or V796A suggests that I792 and V796 are likely involved in further stabilizing this interface, and any changes to these residues could modulate the F-actin-binding activity of αcat-ABD. These results confirm that the hydrophobic residues on the α5-helix surface constitute an important binding surface for F-actin interaction.
Next we tested the in vivo importance of this interaction by expressing an αCat-3A (L798A + I805A + V809A) mutant in Drosophila. All three key hydrophobic residues identified in mammalian αE-catenin or αN-catenin are conserved in Drosophila α-Cat (Fig. 1c). αCat-3A was recruited normally to the cadherin-catenin complex (Fig. 4c and Supplementary Fig. 10c) but failed to show a rescue of the αCat−/− mutant phenotype; a fraction of embryos showed more severe defects than αCat−/− mutants, consistent with a mild dominant-negative effect of αCat-3A expression (Fig. 4a, b). We conclude that direct interaction between α-catenin and F-actin is essential for AJ assembly and function during development.
Allosteric coupling between α1-helix and V796 dynamics
Our observations of the cryptic (attenuated) and exposed (activated) conformations of V796 (Supplementary Fig. 11a, b), despite the nearly identical primary sequences of αEcat- and αNcat-ABDs (87% identity; Supplementary Fig. 3), indicate that this residue resides within a conformationally dynamic region. Consistent with this idea, the αEcat-ABD-WT structure contains an internal cavity that could accommodate V796 in the cryptic state similar to V796N in the αNcat-ABD-WT structure (Fig. 5b, Supplementary Fig. 11c). This internal cavity is partly formed by the side chain of M673 from α1-helix (Fig. 1e), hence raising the possibility that α1-helix unfolding allosterically affects the F-actin-binding site by changing the conformational dynamics of V796.
To determine the influence of α1-helix on the actin-binding site of α-catenin, we developed a new bio-layer interferometry (BLI) approach to measure the kinetics of the αcat-ABD-F-actin interaction. We immobilized F-actin onto the streptavidin-coated optical sensor with biotinylated LifeAct actin-binding peptides (LAbio)46, and measured subsequent association and dissociation of αcat-ABD (Fig. 6a and Supplementary Fig. 13a). We determined that concentration-dependent F-actin binding curves of αEcat-ABD-WT fit well with a 2:1 hetero-ligand:receptor model with two KD values, KD1 = 2.0 μM and KD2 = 0.3 μM (Fig. 6b, Table 1, and Supplementary Fig. 13b). This model supports αEcat-ABD-WT in an equilibrium between the attenuated and activated actin-binding states, respectively. The lower KD2 value is consistent with the positive cooperativity of F-actin binding by αEcat-ABD as previously reported8,29. In contrast, the BLI data of αEcat-ABD-H1 fit well with a 1:1 ligand:receptor model with the single KD value of 0.58 μM (Fig. 6c, Table 1 and Supplementary Fig. 13b), reflecting a predominantly activated state of αEcat-ABD-H1. The effects of mutations in the α-catenin actin-binding site, as well as ΔβH mutation, resulted in decreased affinity (Table 1 and Supplementary Fig. 14a, b) that are consistent with our actin cosedimentation assay results (Figs. 2e and 5c). Our data support the conclusion that the unfolded α1-helix contributed to an apparent equilibrium shift towards an activated state of αcat-ABD.
Comparison of the 15N/1H TROSY NMR spectra of αNcat-ABD-WT47 and αNcat-ABD-H1 showed that a region (αN-catenin residues 794–814) containing V796N and βH was one of three regions affected by the H1 mutation (Supplementary Fig. 15a), likely indicating an altered conformation in this region (Supplementary Fig. 15b). We further confirmed by NMR relaxation and MD simulations studies that the unfolded α1-helix increased molecular motions in the V796N/βH region (Supplementary Fig. 15c, d). In addition, we performed chemical shift (CS)-based Rosetta comparative modeling (CM)48 to show that V796N of αNcat-ABD-WT remained in the cryptic state, whereas αNcat-ABD-H1 displayed a large conformational change that exposed V796N on the surface (Fig. 6d), resembling V796 in the crystal structure of αEcat-ABD-WT (Fig. 5b). Our extended equilibrium MD calculations of αcat-ABDs support that the unfolded α1-helix accelerates the conformational change to favor the exposed state of V796 (Fig. 6e, f). As the exposure of V796N precedes complete unwinding of α1-helix during the constant-force simulation (Supplementary Movie 2), we expect that α1-helix unfolding ‘locks’ V796N in the activated state. Taken together, our observations indicate that allosteric coupling between α1-helix and the actin-binding residue V796 is central to the force-induced association of α-catenin with F-actin.
We show that the unique molecular features of αcat-ABD, α1-helix, V796, and βH, confer mechanosensitivity to α-catenin and its ability to dynamically regulate and reorganize actin filaments directly associated with cadherin-catenin complexes at intercellular junctions. The importance of α-catenin to directly associate with F-actin in a mechanosensitive manner is underscored by experiments showing that αcat-H1 with enhanced F-actin binding was equally inferior to αcatFL function as mutants with diminished F-actin binding (e.g., 3A) during mammalian and Drosophila wound healing, and Drosophila development (Figs. 3 and 4). Although a high-resolution structure of the αcat-ABD-F-actin complex remains to be solved, we have shown that the critical actin-binding site residues, L785, I792, and V796, are located away from the αcat-ABD mechanosensory motif, α1-helix, thus raising the possibility that the N-terminal region of ABD acts allosterically to regulate F-actin binding. Based on these observations, we propose that the coupled conformational states of α1-helix and V796 provide the structural basis of force-dependent allosteric regulation of the α-catenin-F-actin interaction.
In the proposed mechanism, the ABD of α-catenin in the attenuated state can weakly associate with F-actin, whereas its interaction with F-actin under force would trigger α1-helix unfolding and the exposure of V796 to form a catch bond interaction between the cadherin-catenin complex and F-actin at nascent contacts8 (Fig. 7). As nascent contacts grow, multiple α-catenin molecules will bind to F-actin in a cooperative manner8,29 to promote the formation of cadherin-catenin complex clusters (Fig. 7). Although αcat-H1 or other constructs without the α1-helix can support AJ formation in R2/7 cells (Supplementary Fig. 1b and 8b), these do not restore normal α-catenin function (Supplementary Fig. 2a, b) and may reflect a lack of extensive junctional remodeling in these cells. A similar discrepancy between confluent R2/7 cells, wound-healing assays, and in vivo performance was noted for αEcat-NMI (residues 1-402): this ABD-deficient construct forms AJs in R2/7 cells through the recruitment of vinculin6, but does not support normal α-catenin function during wound closure6, and a corresponding Drosophila construct (αCat-NM1) showed no rescue of αCat−/− embryos (R.S. and U.T., unpublished).
Cadherin clustering and AJ maturation likely require trans-interactions and cis-interactions of cadherin ectodomains, as well as an active process involving intracellular coupling of the cadherin-catenin complex to actin networks49. Our αNcat-ABD-H1 crystal structures revealed an unexpected ABD homodimerization (Fig. 2a), which can facilitate F-actin bundling in vitro (Fig. 2e). It involves the βH motif forming an extensive dimer interface with the hydrophobic patch uncovered by α1-helix unfolding (Fig. 2b and Supplementary Fig. 6c). Considering the very weak αcat-ABD dimerization (Fig. 2c and Supplementary Fig. 7c), which is marginally increased by the H1 mutation in solution (Fig. 2d), it is possible that tension-induced unfolding of α1-helix allosterically changes the conformational dynamics of the actin-binding site without affecting dimerization. Nonetheless, AJ-localized α-catenin cooperatively binding to F-actin would likely increase the propensity of αcat-ABD to dimerize and promote F-actin bundling. Uncovering this ABD dimerization interface motivated us to propose a new monomer-dimer model for α-catenin at the cadherin-actin interface (Fig. 7). Although both in vitro8 and in vivo7 studies consistently concluded that monomeric α-catenin forms the essential link between the cadherin-β-catenin complex and F-actin, the current model fails to account for the capacity of α-catenin to bundle F-actin5 at AJs. The ABD-dependent dimerization as demonstrated here allows actin filaments to be tightly bundled in an antiparallel fashion (Fig. 8) and places the α3-α4 surface of ABD in close proximity with F-actin, which is consistent with our NMR saturation transfer data (Fig. 2f). In addition, the ABD dimerization allows F-actin bundling to occur while the N domain of α-catenin remains associated with cadherin-bound β-catenin (Fig. 7). Hence, our proposed model differs from the previous monomer-dimer model of α-catenin by arguing that (i) the E-cadherin/β-catenin/α-catenin/F-actin complex regulates the cadherin-actin linkage without disrupting the β-catenin-α-catenin interaction; (ii) that α-catenin as a component of the complex can bundle F-actin, and (iii) that α-catenin controls actin binding through force-dependent allosteric regulation of the actin-binding site within the ABD. The versatility of α-catenin to modulate the attachment of the cadherin-catenin complex to F-actin from transient interaction to stable actin bundling, and to the dynamic cortical actin network50, will likely involve additional dynamic connections provided by the recruitment of other F-actin-binding proteins, such as vinculin, afadin, ZO-1 and EPLIN, to intercellular junctions6,27,51,52,53.
We have employed an integrative structure/function approach to show that the structural motifs of αcat-ABD involved in the regulation of tension-sensitive actin binding are essential for normal tissue morphogenesis and wound healing. Although the occurrence of actin bundling involving ABD-linked α-catenin dimers at intercellular junctions remains to be tested, our model reconciles previous observations of α-catenin as a critical mechanosensor engaged in reorganization of AJs by facilitating dynamic F-actin association8,21, and actin bundling through homodimerization5,20. Moreover, the significance of this mechanism lies in the ability of α-catenin to modulate cadherin-mediated cell adhesion through force-dependent F-actin binding and actin remodeling without dissociation from the cadherin-β-catenin complex.
Protein expression and purification
The cDNA corresponding to the actin-binding domain (ABD) of mouse αE-catenin (652-906), mouse αN-catenin (651-905) and all related mutants (e.g., the αE-catenin H1 mutation RAIM670-673GSGS) were amplified by PCR and individually subcloned into the pGEX4T1 vector (GE Healthcare). The fly αcat-ABD (659-917) cDNA was amplified by PCR and subcloned into a modified pET-SUMO vector. Site-directed mutagenesis was performed using the Quikchange protocol (Stratagene) to produce all single-/multiple-residue and deletion mutants. Recombinant proteins were expressed as N-terminal glutathione S-transferase (GST) fusion proteins in Escherichia coli BL21-CodonPlus cells. Cells were grown to an O.D.600 of 0.8 at 37 °C and the recombinant protein expression was induced with 0.5 mM isopropyl β-D-1-thiogalactopyranoside for 16 h at 16 °C. Cells harvested by centrifugation were resuspended in the lysis buffer (50 mM Tris-HCl, pH 8.0, 300 mM NaCl, 10 mM β-mercaptoethanol, 1 mM Tris(2-carboxyethyl)phosphine (TCEP)), sonicated on ice, and subjected to centrifugation to isolate soluble proteins. GST-fusion proteins were isolated using the glutathione-sepharose resin (GE Healthcare). His-SUMO fusion proteins were isolated using the Ni2+-NTA resin (ThermoFisher Scientific). GST-fusion and His-SUMO proteins were cleaved by thrombin or SUMO protease (Ulp-1), respectively. The cleaved proteins were further purified by size-exclusion chromatography using Superdex 75 (GE Healthcare) in the running buffer (50 mM Tris-HCl, pH 8.0, 300 mM NaCl, 1 mM TCEP). The purified proteins were exchanged into protein storage buffer (50 mM Tris-HCl, pH 8.0, 100 mM NaCl, 1 mM TCEP).
Size-exclusion chromatography-multiangle light scattering
Purified protein (5 mg/mL, 100 μL injection volume) was subjected to size-exclusion chromatography (SEC) using a Superdex-200 Increase 10/300 GL column (GE Healthcare) equilibrated in SEC-MALS buffer (20 mM Tris-HCl pH 7.0, 100 mM NaCl) at a flow rate of 0.5 mL/min. Multi-angle light scattering (MALS) measurements were performed in-line with SEC by using a three-angle (45°, 90°, and 135°) miniDawn light-scattering instrument and an Optilab rEX differential refractometer (Wyatt Technologies). Molecular weight was calculated by using the ASTRA software (Wyatt Technologies). Dimer peak area integration was performed by using ImageJ54. Statistical analysis was performed by Two-way ANOVA followed by Bonferroni’s comparison test.
Actin cosedimentation assay
Monomeric rabbit skeletal muscle actin was purified from rabbit muscle acetone powder55 (Pel-Freez Biologicals). Purified globular actin (G-actin) was diluted to 20 μM in a fresh Buffer-G (2 mM Tris-HCl, pH 8.0, 0.2 mM ATP, 0.5 mM DTT, 0.1 mM CaCl2), and subsequently polymerized in Buffer-F (5 mM Tris-HCl, pH 8.0, 50 mM KCl, 2 mM MgCl2, 1 mM ATP, 0.2 mM CaCl2, 0.5 mM DTT) for 1 h at RT. The αcat-ABD samples were subjected to buffer exchange into Buffer-F. Samples of F-actin and ABD were mixed (the protein mixture contains 5 μM ABD and 5 μM actin in 50 μL) in Ultra-Clear Centrifuge Tubes (Beckman Coulter) and incubated for 1 h at RT. F-actin with bound protein samples were cosedimented by centrifugation using a Beckman Coulter Airfuge with a chilled A-100/30 rotor at 28 psi (≥100,000×g) for 20 min at RT. To assess actin bundling, F-actin with bound protein samples were cosedimented by centrifugation using a benchtop microcentrifuge at low relative centrifugal force (RCF; 10,000×g) for 30 m at 4 °C. Supernatant and pellet fractions were analyzed by SDS-PAGE with coomassie blue stain. Gel band intensity was measured by using ImageJ54. Statistical analysis of three or more groups was performed by One-way ANOVA followed by Tukey’s multiple comparison test. Statistical analysis of two groups was performed by Two-way ANOVA followed by Bonferroni’s comparison test.
Crystallization and data collection
Crystals of the αN-catenin ABD-H1 were grown at 277 K by vapor diffusion. For crystallization the αNcat-ABD-H1 sample was exchanged into Buffer-P (20 mM K/Na phosphate, pH 6.0, 150 mM NaCl, 1 mM TCEP), and the protein solution (30 mg/mL) was mixed with an equal volume of the reservoir solution, which consists of either solution A (2.0 M (NH4)2SO4, 10 mM CoCl2) for form A crystals, or solution B (100 mM Na acetate/acetic acid, pH 4.5, 0.8 M NaH2PO4, 1.2 M K2HPO4) for form B crystals. Similarly, crystals of the αEcat-ABD-WT in Buffer-P (30 mg/mL) were grown at 277 K by vapor diffusion with the reservoir solution consisting of 0.2 M KBr, 2.2 M (NH4)2SO4 and 3% (w/v) D-Galactose. Crystallization of αEcat-ABD-H1 was unfruitful. Crystals were briefly soaked in crystallization solution containing 25% glycerol for data collection at 100 K. Diffraction data were collected at the Canadian Light Source-Canadian Macromolecular Crystallography Facility (CMCF) beamline 08ID-1 (Saskatoon, Canada) and processed with HKL200056. Br-SAD data were collected with αEcat-ABD crystals. Statistics pertaining to the diffraction data are presented in Supplementary Table 2.
Crystal structure determination and refinement
Crystal structures of the αNcat-ABD-H1 in forms A and B were determined at 2.2 and 2.8 Å resolution, respectively. The structure solution was solved by molecular replacement using PHASER57 with the αNcat-ABD-WT crystal structure (PDB ID: 4K1O) as a search model. Successive rounds of manual model building and refinement were performed by using Coot58 and PHENIX59 to refine the models of αNcat-ABD-H1. The crystal structure of the αEcat-ABD-WT was initially determined at 2.3 Å resolution by the single-wavelength anomalous dispersion method, and further refined at 2.2 Å by using PHENIX. Refinement statistics are presented in Supplementary Table 2. Molecular graphics representations were prepared using PyMOL (http://www.pymol.org/).
The NMR experiments of 15N/13C labeled αNcat-ABD-WT and αNcat-ABD-H1 were performed on Bruker AVANCE II 800 MHz (Bruker Biospin) spectrometer equipped with a cryogenic triple-resonance z-gradient probe. Labeled proteins were expressed in E. coli BL21-CodonPlus with M9 minimal media containing 15N-ammonium chloride and 13C-glucose for 15 h at 288 K. The purification of labeled ABD proteins was performed in a similar manner as described above. The backbone assignment of αNcat-ABD-H1 was processed using standard 1H-15N experiments. 15N relaxation data were acquired at 288 K in the presence and absence of a 3 s 1H saturation period prior to 15N excitation using the 15N-1H heteronuclear NOE pulse sequence60. NMR spectra were processed using NMRPipe61 and resonance assignment was carried out using NMRView62. Errors in peak intensity values were estimated from the signal-to-noise ratio of each spectrum.
The transferred cross saturation (TCS) experiments were performed at 293 K to detect the resonances of αNcat-ABD-WT in the free state after binding to F-actin in solution. The 15N/2H-labeled αNcat-ABD-WT was mixed with unlabeled F-actin at the molar ratio of 1:0.1 (ABD:G-actin) in the modified actin polymerization buffer (20 mM Tris-HCl, pH 7.0, 150 mM NaCl, 0.2 mM ATP, 0.1 mM CaCl2, 1 mM TCEP, 50 mM KCl, 2 mM MgCl2) containing 12% H2O to avoid the dipole coupling between the amides63. Control TCS experiments were carried out without F-actin to assess the effects of the residual aliphatic protons in the ABD.
NMR chemical shift (CS)-guided structure modeling of the αNcat-ABD-WT and αNcat-ABD-H1 (28 kDa) was performed by employing the CS-Rosetta-CM approach48 with NMR chemical shift data (αNcat-ABD-WT and αNcat-ABD-H1) and the αNcat-ABD-WT crystal structure (PDB ID: 4K1O) as the template. This approach enables CS-Rosetta modeling to be effective for proteins larger than 15 kDa. The POMONA server (https://spin.niddk.nih.gov/bax/nmrserver/pomona/) was used to prepare the Rosetta input files.
To determine a dissociation constant for the αcat-ABD-F-actin interaction, we devised a biolayer interferometry approach which uses label/modification-free F-actin and minimizes any occurrence of actin bundling. All BLI experiments were performed at 26 °C using Octet384 (Fortebio). All proteins used in BLI experiments were buffer exchanged into the assay buffer (2 mM Tris-HCl, pH8.0, 0.2 mM ATP, 0.5 mM DTT, 0.1 mM CaCl2, 50 mM KCl, 2 mM MgCl2, 0.1% BSA, 0.02% Tween-20). F-actin was polymerized for 1 h at RT, and subsequently diluted to 1 μM for the assay. We first load the optical surface of the Streptavidin (SA) biosensors with the widely-used F-actin-binding peptide LifeAct46 containing a C-terminal biotinylation (LAbio) at the concentration of 2 μg/mL. The SA sensors coated with the N-terminally biotinylated LA (bioLA) did not produce any response signals in the presence of F-actin (Supplementary Fig. 13a). The SA-LAbio sensor was used to immobilize F-actin through the LifeAct-F-actin interaction with high avidity. To measure the sub-micromolar affinity binding of αEcat-ABD-H1, the Super-Streptavidin (SSA) biosensors with increased Streptavidin density were used. The SA-LAbio-F-actin sensors were incubated with αcat-ABD at various concentrations for the association step, and transferred to a buffer to monitor its dissociation. Octet analysis software (Fortebio) was used to perform the kinetic analysis of ABD-F-actin interaction.
Circular dichroism spectroscopy data for 40 μM αcat-ABD samples were collected on a Jasco J-815 CD spectrometer (Jasco, Tokyo, Japan) at 293 K using a 0.1 cm path length cuvette with a scanning speed of 20 nm/min (1 nm increments). Thermal denaturation data were acquired at 220 nm with a scan rate of 1 °C/min.
Multiple sequence alignment of human αE-catenin (accession P35221), mouse αE-catenin (accession NP_033948.1), mouse αN-catenin (accession NP_663785.2), mouse αT-catenin (accession Q65CL1.2), Drosophila α-Catenin (P35220.2), and mouse vinculin (accession Q64727) was performed by using T-Coffee (http://tcoffee.crg.cat)64 and ESPript (http://espript.ibcp.fr)65.
Mammalian cell culture
The αE-catenin cDNA was amplified by PCR and subcloned into a modified pCAH vector66 to express αE-catenin-mGFP (monomeric EGFP with the dimerization-disrupting A206K mutation67). The full-length or deletion variant αE-catenin cDNA was subcloned into pCA vector to express αE-catenin-FLAG. Site-directed mutagenesis was performed using the Quikchange protocol (Stratagene) to produce αE-catenin deletion mutants, NMIABD, NMIABD* and NMI. with the constitutively active MI/vinculin-binding site (VBS; residues 305-352). R2/7 cells were cultured in Dulbecco’s modified Eagle’s medium (Thermo Fisher Scientific) supplemented with 10% v/v fetal bovine serum (Invitrogen), 100 U/ml penicillin, and 100 mg/ml streptomycin (Sigma-Aldrich). Cells were transfected with expression vectors by using Lipofectamine 3000 (ThermoFisher Scientific), and cells with stable protein expression was selected based on antibiotic resistance (300 μg/mL hygromycin for pCAH vectors) and subsequently isolated by flow cytometry. For live imaging analysis of αE-catenin-mGFP-expressing R2/7 monolayers post-scratch wounding, cells were also infected with a lentiviral LifeAct-Ruby purchased from Addgene (pLenti.PGK.LifeAct-Ruby, Addgene Plasmid #51009).
The following primary antibodies were used: hybridoma mouse anti-α-catenin (5B11; undiluted), polyclonal rabbit anti-phospho-serine 641 αCat (21330; 1:300, Signalway Antibody), monoclonal mouse anti-GAPDH (9484; 1:5000, Abcam). Secondary antibodies for Western blotting included fluorescently labeled donkey anti-mouse and anti-rabbit antibodies (680RD or 800RD; 1:5000, LiCor Biosciences). Secondary antibodies for immunofluorescence included IgG Alexa Fluor 488 or 568-conjugated goat anti-mouse or anti-rabbit antibodies (1:200, Invitrogen). For R2/7 cells expressing αE-catenin-FLAG, the following antibodies were used: anti-DDDDK tag rabbit polyclonal antibody, which recognizes the FLAG tag (1:200, MBL), anti-ZO-1 mouse monoclonal antibody (T8-754; undiluted, a gift from Sa. Tsukita, Osaka University, Japan), anti-E-cadherin mouse monoclonal antibody (clone36/E-cadherin; 1:100, BD) and anti-E-cadherin rat monoclonal antibody (ECCD2; 1:200, a gift from M. Takeichi, Riken, Japan). Alexa Fluor 488-conjugated or 555-conjugated secondary antibodies (1:200) were purchased from Invitrogen. Alexa Fluor 647-conjugated phalloidin (1:50, Invitrogen) was used for staining actin filaments.
Fluorescence microscopy and confocal imaging
Cells were grown on coverslips, fixed in 2% paraformaldehyde (Electron Microscopy Services, Hatfield, PA) for 15 m, quenched with glycine, permeabilized with 0.3% Triton X-100 (Sigma), and blocked with normal goat serum (Sigma). Primary and secondary antibody incubations were performed at RT for 1 h, interspaced by multiple washes in PBS, and followed by mounting coverslips in ProLong Gold fixative (Life Technologies). Confocal images were acquired with an Olympus IX81 inverted confocal microscope (Olympus) through a Plan-Apochromat x60/1.42 NA oil-immersion objective lens (Olympus). Images were processed by Olympus FV10-ASW.
For R2/7 cells expressing αE-catenin-FLAG, images were taken using Olympus BX51 microscope with a UPlanFLN 40 × /0.75 Ph2 lens or a PlanApo 60 × /1.40 Oil Ph3 lens.
The images of R2/7 cells expressing αE-catenin variants at the scratch wound fronts were collected on a Nikon Ti Eclipse inverted microscope equipped with a Yokogawa CSU-X1 spinning disk head, Perfect Focus system (Nikon), a 100 × 1.49 NA APO TIRF objective, and an Andor xION EMCCD camera (Andor Technology), controlled by MetaMorph 188.8.131.52 software (Molecular Devices). Cells were maintained at 37 °C plus 5% CO2 during imaging using a Tokai-Hit stage-top incubator (Tokai-Hit) or an Okolab gas mixer (Okolab).
For imaging z-stacks of cell piling, slides were imaged at RT with 60× oil Apo TIRF NA 1.49 objective on the Nikon A1R laser scanning confocal inverted microscope equipped with two standard PMTs (408, and 640) and two high sensitivity GaAsP detectors (488, 561).
Scratch wound assay
250,000 R2/7 cells were plated for 24 h on LabTek #1 4-well chamber slide (43300-776, Thermo Fisher Scientific), wounded with a P200 micropipette tip and cells were allowed to recover for 2 h. Prior to imaging, DMEM media was replaced with FluoroBright DMEM (Life Technologies) and 10 g/mL Mitomycin C (Sigma) to limit cell proliferation. Cells in Fig. 3d were imaged with the 20× objective every 10 min (both phase contrast and fluorescent channels) on the Nikon Biostation IM-Q with the slide holder module (located in Northwestern University Nikon Imaging Facility) at 37 °C, 5% CO2, for 15 h. 10–12 fields of view (FOV) were captured along the wound edge. Instrument controlled by Biostation IM software, version 2.21 build 144. To quantify change in wound area, the resulting .ids file was imported to ImageJ54, and the wound edge of the phase-contrast image was traced with the polygon tool at time = 0 and time = 15 h. The area of the resulting polygon was measured in pixels² and the resulting data (35 FOV; 5BR) were compared by performing One-way ANOVA statistical analysis followed by Tukey’s multiple comparison test. Images presented in the paper were adjusted for brightness/contrast but were otherwise unprocessed.
To determine the velocity (μm/sec) and persistence (total path length (μm)/net displacement (μm)) of cells at the wound front, tracked individual cells at each interval of the time lapse sequence using the MTrackJ plug-in (FIJI) (see colored tracks in Supplementary Fig. 9b). The center of the nucleus (centroid) was hand-marked at each interval. Data points for each of the 5 constructs were compiled from scratch wounds carried out on three different days to ensure biological reproducibility and significance by ANOVA.
To validate cell trajectories for the WT versus H1 αE-catenin mutant comparison using a fully unbiased method, a particle image velocimetry (PIV) analysis (i.e., template matching by cross-correlation) was carried out using the PIV_jar plug-in (FIJI). Briefly, each acquired image frame was divided into an interrogation window size that approximated single cells or a small cell group depending on degree of cell packing (100 μm2). Vectors were averaged from all 91 frames/movie for 3 αEcat-WT and 4 αEcat-H1 wounds (two biological replicates from separate days).
Epithelial sheet disruption assay
R2/7 cells expressing α-catenin variants were plated in a 12-well culture plates (Corning) and allowed to reach confluency. After 36 h, the monolayer was washed 2x in DPBS supplemented with 0.5 mM Mg2+ and 1 mM Ca2+ (HyClone) and then incubated for 30 m at 37 °C in 1 mg/ml Dispase (Roche) diluted in PBS and supplemented with the indicated amount of Ca2+. Subsequently monolayers were disrupted by subjecting the plate to a shaking force of 1400 rpm for 15 s. An image of each well was captured before and after shaking, and monolayer fragments were counted. Seventy-five epithelial fragments were established as the upper limit for counting. The assay results were collected from 3 technical replicates. Statistical analysis was performed by One-way ANOVA followed by Tukey’s multiple comparison test.
Measurement of transepithelial electric resistance
Confluent monolayers of R2/7 cell lines expressing various α-catenin mutants grown in TranswellTM chamber were used. Transepithelial electric resistance (TER) was measured using a Millicell-ERS epithelial volt-ohmmeter (Millipore) and normalized by the area of the monolayer. The background TER of blank TranswellTM filters was subtracted from the TER of cell monolayers. Statistical analysis was performed by One-way ANOVA followed by Tukey’s multiple comparison test.
Cells were fixed with 2% paraformaldehyde, 2.5% glutaraldehyde in 0.1 M sodium cacodylate buffer, pH 7.5 for 2 h at RT. Samples were conventionally dehydrated and embedded in resin (Polybed 812; Polyscience). Ultrathin sections were cut and stained with uranyl acetate and lead citrate before observation with an electron microscope (JEM-1010; JEOL).
Molecular dynamics simulations
The structure of the αNcat-ABD-WT was obtained from the Protein Data Bank (PDB ID 4K1O27). The structure of αNcat-ABD-H1 was obtained from the form A crystal structure of αN-catenin ABD-H1. The structure of αEcat-ABD-WT was obtained from the crystal structure of αEcat-ABD-WT chain A. All-atom MD simulations were performed using NAMD268, the CHARMM27 force field for proteins and ions69,70, and the TIP3P model for explicit water71. The Solvate and Autoionize plugins of VMD were used to solvate the system in a water box with at least 15 Å between the protein and the boundary of the box, and to add 150 mM NaCl and neutralize the net charge of the system, respectively72. All simulations were performed using periodic boundary conditions and a time step of 2 fs. A constant temperature of 310 K was maintained using Langevin dynamics with a damping coefficient of 0.5 ps-1(refs. 73,74). Short-range, non-bonded interactions were calculated using a cutoff distance of 12 Å, and long-range electrostatic forces were described with the Particle Mesh Ewald method75. Throughout the simulations, bond distances involving hydrogen atoms were fixed using the SHAKE algorithm76. After an initial 10,000 steps of energy minimization with all Cα atoms fixed, the system was equilibrated in an NVT (constant Number, Volume, and Temperature) ensemble for 500 ps, during which all protein Cα atoms were fixed to allow relaxation of the side chains and water. Subsequent equilibrium simulations were performed in an NVT ensemble. In constant-force SMD simulations of the αcat-ABD, a 100-pN pulling force was applied to the Cα atom of N-terminal residue 668/669 (αN-/αE-catenin). It should be noted that the force-dependent binding between α-catenin and F-actin was previously detected in vitro at ~5 pN8. We chose a 100-pN pulling force for the SMD simulations by considering the trade-off between the improved accuracy and longer time required to observe conformational changes under low force. Assuming the SMD simulation successfully captures the physiological unfolding pathway, we expect unfolding of α1-helix to occur at lower force (<100 pN) over a longer timescale (>60 ns). The center of mass (the sulfur atom of M722/723) was fixed to prevent the overall translation of the system in response to the applied external force. The force was directed along the x-axis between the two anchoring points, and the vector pointing from the center of mass to the N-terminal residue was defined as the positive direction. Simulation outputs were analyzed using VMD72 and plotted using the matplotlib plotting library77. The portion of each equilibrium simulation during which the RMSD monotonically increased (the first 7–13 ns) was excluded from analysis to avoid biasing the results toward the initial coordinates.
3D cell culture
We observed for the ability of R2/7 cells expressing αE-catenin variants to form spheroid structures by using Non-adherent Corning 96-well round bottom ultra-low attachment microplate (Corning). In each well, 25–200 cells were suspended in 100–200 μL of DMEM containing 10% FBS. The microplate was placed in a CO2 incubator, and occasionally taken out of the incubator to acquire phase contrast images with a stereomicroscope equipped with a digital camera at various time points.
Drosophila stocks, overexpression, and rescue experiments
Paired-GAL4, Act5c-GAL4, and da-GAL4 were obtained from the Bloomington Drosophila stock center (https://bdsc.indiana.edu). Transgenic constructs inserted at attP2 were recombined with α-Cat1 and balanced over TM3, Ser, twi-Gal4, UAS-GFP. These flies were crossed to act-Gal4 da-Gal4 α-Cat1/TM3, Ser twi-Gal4 UAS-GFP and eggs were collected on apple juice agar plates at 25 °C.
For quantification of whole-animal rescue experiments, a total of 100–300 fertilized non-GFP embryos were collected, allowed to develop at 25 °C, and monitored daily. Dead embryos were mounted in Hoyer’s medium and lactic acid (1:1 ratio) for examination of the embryonic cuticle. Lethality counts were performed on larvae that hatched, at each of the stages of Drosophila development indicated below. A specific score from −2 to 8 was given to each recued and control animal with a score of 0 denoting the phenotype most frequently observed in α-Cat1 zygotic null mutant embryos9. The following scoring criteria were applied to measuring the extent of enhancement or rescue of the α-Cat mutant phenotype: (−2) embryonic lethal with both a severe head defect (‘head open’) and a dorsal open phenotype indicating a failure of dorsal closure; (−1) embryonic lethal with both the head open defect and a hole in the dorsal epidermis indicating incomplete closure; (0) embryonic lethal with a head open defect; (1) embryonic lethal with weak head defects (‘abnormal head’); (2) embryonic lethal with normal head; (3) lethal at first larval instar; (4) lethal at second larval instar; (5) lethal at third larval instar; (6) early pupa lethal; (7) late pupa lethal; (8) adult. Statistical significance was assessed with a non-parametric Kolmogorov-Smironov test. Data are presented as mean plus SD.
Immunohistochemistry and histology of Drosophila embryos
To prepare embryonic cuticle, embryos were de-chorionated in 50% bleach for 5 min, washed with 0.1% TritonX-100, mounted in 1:1 solution of Hoyer’s and lactic acid and incubated overnight at 60 °C. Embryos were examined under phase contrast using a Zeiss Axiophot microscope.
The heat fixation method78 was used for antibody staining of α-Cat1 embryos expressing UAS-driven transgenes with da-Gal4. Embryos over-expressing transgenes using Paired-Gal4 were fixed in a 1:1 mixture of 5% formaldehyde in 1× PBS and Heptane for 20 m. In both cases, embryos were devitellinized using a 1:3 mixture of Heptane and Methanol. Primary antibodies used were: rat mAb anti-HA (3F10, 1:500, Sigma), guinea pig pAb anti-α-Catenin (p121; 1:1000, Sarpal et al., 2012), mouse mAb anti-Arm (N2-7A1, 1:50; Developmental Studies Hybridoma Bank [DSHB]). Fluorescent secondary antibodies were used at a dilution of 1:400 (Jackson ImmunoResearch Laboratories and Invitrogen). Samples were analyzed on a Leica SP8 scanning laser confocal microscope using a 40x oil immersion lens (NA 1.3). Images were prepared and assembled in ImageJ, Adobe Photoshop, and Adobe Illustrator.
Generation of Drosophila transgenes
To generate the Drosophila UASp-αCat constructs, full-length αCat (2751 nucleotides) was cloned into Gateway pENTRTM/D-TOPO entry vector (Invitrogen) digested with Not1 and Asc1, using 3-part Gibson assembly reaction (NEB). αCat cDNAs carrying various mutations were cloned using 2-part or 3-part Gibson assembly reactions using UASp-αCat in pENTRTM/D-TOPO as the backbone digested with following restriction enzymes: Not1 and Blp1 to generate αCat-H1 and αCat-Δα1; Blp1 and Asc1 to generate αCat-ΔβH, Not1 and Asc1 to generate αCat-H1ΔβH; Blp1 and Asc1 to generate αCat-3A. The Gateway® LR® Clonase Enzyme mix was used to clone all entry vector constructs into the pPWH (pUASP-Gateway Cassette with C-terminal 3x HA) vector containing an attB recombination site that was added using the NSi1 restriction site. Transgenic animals were produced by Best Gene Inc., by using flies carrying the attP2 recombination site on the left arm of the third chromosome. Amino acids of α-Cat proteins are: αCatFL (1-917), αCat-H1 (683REAM > 683GSGS), αCat-Δα1 (1-658/691-917), αCat-ΔβH (1- 811/824-917), αCat-H1ΔβH (1-811/824-917 + 683REAM > 683GSGS), αCat-3A (L798A + I805A + V809A), and αCat-ΔABD (1-708)7.
Immunoblotting with fly samples
Transgenic flies were crossed to da-Gal4. An overnight collection of embryos was dechorionated and homogenized in cold lysis buffer (RIPA-25 mM Tris-HCl, pH 7.6, 150 mM NaCl, 1% TritonX-100, 1% sodium deoxycholate, 0.1% SDS) containing protease inhibitors (Roche). The protein concentrations were measure using the Bradford assay. 25 μg of total protein lysates were resolved by SDS-PAGE. Immunoblotting was performed according to recommended protocols provided by Life Technologies iBlot 2® Dry Blotting system. Primary antibodies used were anti-HA (rat monoclonal 3F10, 1:500; Abcam), anti-β-tubulin (mouse monoclonal, E7-1:1000; DSHB) and anti-α-Cat (guinea pig polyclonal, p121; 1:1000; Sarpal et al., 2012). Secondary antibodies used were: anti-rat 680LT (Goat-1:5000; LI-COR), anti-mouse 800CW (Donkey-1:5000, LI-COR) and anti-guinea pig 800LT (1:5000, LI-COR). Protein bands were visualized using the LI-COR Odyssey® Fc Dual-Mode Imaging system.
Wound healing assay
For time-lapse imaging stage 15 Drosophila embryos were dechorionated in 50% bleach for 2 min, rinsed with water, and mounted ventrolateral-side-down onto a coverslip using heptane glue. Embryos were covered with a 1:1 mix of halocarbon oil 27:700 (Sigma-Aldrich), and imaged using a Revolution XD spinning disk confocal (Andor) with a 60x oil-immersion lens (NA 1.35; Olympus). Images were captured with an iXon Ultra 897 camera (Andor) and Metamorph (Molecular Devices) as the image acquisition software. 16-bit Z-stacks were acquired in 0.2 μm steps (21 slices per stack) every 30 s. Maximum intensity projections were used for analysis. Wounds were made using a pulsed Micropoint N2 laser (Andor) tuned to 365 nm. The laser produced 120 μJ pulses at the source with a duration of 2–6 ns. To wound embryos, ten laser pulses were delivered at each of seven spots along a 14 μm line.
To quantify wound closure dynamics, wounds were delineated using the Livewire algorithm, a semi-automated optimal path search method for image segmentation, in which the user traces the wound margin with the mouse, and the algorithm automatically identifies the brightest pixels that follow the trajectory of the mouse. We used the Livewire implementation in SIESTA, an image analysis platform that we develop79,80. The wound closure rate was calculated by subtracting the wound area 30 min after wounding from the maximum wound area for each wound, divided by the elapsed time.
We compared sample variances using the F-test for statistical analysis. To compare mean sample values, we used Student’s t test for populations with equal or unequal variances (depending on the outcome of the F-test), applying Holm’s correction to account for the comparison between three groups. For time series, error bars indicate SEM. For box plots, error bars show SD, the box indicates the SEM, and gray lines denote the mean.
Crystal structure coordinates and structure factors of αNcat-ABD-H1 form A, αNcat-ABD-H1 form B and αEcat-ABD-WT are deposited in the Protein Data Bank under accession codes: 6DUW, 6DUY, and 6DV1, respectively. Backbone chemical shift assignments of αNcat-ABD-H1 are deposited in the Biological Magnetic Resonance Data Bank under accession code 27526. All reagents and experimental data are available from the authors upon request. A Reporting Summary for this Article is available as a Supplementary Information file. Source Data are provided for Figs. 1f, 2d, 2f, 3b, 3e, 3f, 4a, 4e, 5c, 6b, 6c, 6f, and Supplementary Figs. 2b, 2c, 4b‒d, 7a, 7c, 8a, 10a‒b, 14a‒b, 15a‒c as a Source Data file.
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Heisenberg, C.-P. & Bellaïche, Y. Forces in tissue morphogenesis and patterning. Cell 153, 948–962 (2013).
Heller, E. & Fuchs, E. Tissue patterning and cellular mechanics. J. Cell Biol. 211, 219–231 (2015).
Takeichi, M. Dynamic contacts: rearranging adherens junctions to drive epithelial remodelling. Nat. Rev. Mol. Cell Biol. 15, 397–410 (2014).
Harris, T. J. C. & Tepass, U. Adherens junctions: from molecules to morphogenesis. Nat. Rev. Mol. Cell Biol. 11, 502–514 (2010).
Rimm, D. L., Koslov, E. R., Kebriaei, P., Cianci, C. D. & Morrow, J. S. α1(E)-catenin is an actin-binding and -bundling protein mediating the attachment of F-actin to the membrane adhesion complex. Proc. Natl Acad. Sci. USA 92, 8813–8817 (1995).
Yonemura, S., Wada, Y., Watanabe, T., Nagafuchi, A. & Shibata, M. α-Catenin as a tension transducer that induces adherens junction development. Nat. Cell Biol. 12, 533–542 (2010).
Desai, R. et al. Monomeric α-catenin links cadherin to the actin cytoskeleton. Nat. Cell Biol. 15, 261–273 (2013).
Buckley, C. D. et al. Cell adhesion. The minimal cadherin-catenin complex binds to actin filaments under force. Science 346, 1254211 (2014).
Sarpal, R. et al. Mutational analysis supports a core role for Drosophila α-catenin in adherens junction function. J. Cell Sci. 125, 233–245 (2012).
Kofron, M., Spagnuolo, A., Klymkowsky, M., Wylie, C. & Heasman, J. The roles of maternal α-catenin and plakoglobin in the early Xenopus embryo. Development 124, 1553–1560 (1997).
Torres, M. et al. An α-E-catenin gene trap mutation defines its function in preimplantation development. Proc. Natl Acad. Sci. USA 94, 901–906 (1997).
Costa, M. et al. A putative catenin-cadherin system mediates morphogenesis of the Caenorhabditis elegans embryo. J. Cell Biol. 141, 297–308 (1998).
Park, C., Falls, W., Finger, J. H., Longo-Guess, C. M. & Ackerman, S. L. Deletion in Catna2, encoding αN-catenin, causes cerebellar and hippocampal lamination defects and impaired startle modulation. Nat. Genet. 31, 279–284 (2002).
Li, J. et al. Loss of αT-catenin alters the hybrid adhering junctions in the heart and leads to dilated cardiomyopathy and ventricular arrhythmia following acute ischemia. J. Cell Sci. 125, 1058–1067 (2012).
Saksens, N. T. M. et al. Mutations in CTNNA1 cause butterfly-shaped pigment dystrophy and perturbed retinal pigment epithelium integrity. Nat. Genet. 48, 144–151 (2016).
Ding, L. et al. Genome remodelling in a basal-like breast cancer metastasis and xenograft. Nature 464, 999–1005 (2010).
Zhao, Z.-M., Reynolds, A. B. & Gaucher, E. A. The evolutionary history of the catenin gene family during metazoan evolution. BMC Evol. Biol. 11, 198 (2011).
Aberle, H. et al. Assembly of the cadherin-catenin complex in vitro with recombinant proteins. J. Cell Sci. 107, 3655–3663 (1994).
McCrea, P. D. & Gumbiner, B. M. Purification of a 92-kDa cytoplasmic protein tightly associated with the cell-cell adhesion molecule E-cadherin (uvomorulin). Characterization and extractability of the protein complex from the cell cytostructure. J. Biol. Chem. 266, 4514–20 (1991).
Drees, F., Pokutta, S., Yamada, S., Nelson, W. J. & Weis, W. I. α-catenin is a molecular switch that binds E-cadherin-β-catenin and regulates actin-filament assembly. Cell 123, 903–915 (2005).
Yamada, S., Pokutta, S., Drees, F., Weis, W. I. & Nelson, W. J. Deconstructing the cadherin-catenin-actin complex. Cell 123, 889–901 (2005).
Pokutta, S. & Weis, W. I. Structure of the dimerization and β-catenin-binding region of α-catenin. Mol. Cell 5, 533–543 (2000).
Pokutta, S., Choi, H.-J., Ahlsen, G., Hansen, S. D. & Weis, W. I. Structural and thermodynamic characterization of cadherin·β-catenin·α-catenin complex formation. J. Biol. Chem. 289, 13589–13601 (2014).
Yao, M. et al. Force-dependent conformational switch of α-catenin controls vinculin binding. Nat. Commun. 5, 4525 (2014).
Rangarajan, E. S. & Izard, T. The cytoskeletal protein α-catenin unfurls upon binding to vinculin. J. Biol. Chem. 287, 18492–18499 (2012).
Choi, H.-J. et al. αE-catenin is an autoinhibited molecule that coactivates vinculin. Proc. Natl Acad. Sci. USA 109, 8576–8581 (2012).
Ishiyama, N. et al. An autoinhibited structure of α-catenin and its implications for vinculin recruitment to adherens junctions. J. Biol. Chem. 288, 15913–15925 (2013).
Escobar, D. J. et al. α-Catenin phosphorylation promotes intercellular adhesion through a dual-kinase mechanism. J. Cell Sci. 128, 1150–1165 (2015).
Hansen, S. D. et al. αE-catenin actin-binding domain alters actin filament conformation and regulates binding of nucleation and disassembly factors. Mol. Biol. Cell 24, 3710–3720 (2013).
Bakolitsa, C. et al. Structural basis for vinculin activation at sites of cell adhesion. Nature 430, 583–586 (2004).
Dembo, M., Torney, D. C., Saxman, K. & Hammer, D. The reaction-limited kinetics of membrane-to-surface adhesion and detachment. Proc. R. Soc. Lond. Biol. Sci. 234, 55–83 (1988).
Marshall, B. T. et al. Direct observation of catch bonds involving cell-adhesion molecules. Nature 423, 190–193 (2003).
Pokutta, S., Drees, F., Takai, Y., Nelson, W. J. & Weis, W. I. Biochemical and structural definition of the l-afadin- and actin-binding sites of α-catenin. J. Biol. Chem. 277, 18868–18874 (2002).
Pappas, D. J. & Rimm, D. L. Direct interaction of the C-terminal domain of α-catenin and F-actin is necessary for stabilized cell-cell adhesion. Cell Commun. Adhes. 13, 151–170 (2006).
Chen, C.-S. et al. α-Catenin-mediated cadherin clustering couples cadherin and actin dynamics. J. Cell Biol. 210, 647–661 (2015).
Watabe-Uchida, M. et al. α-Catenin-vinculin interaction functions to organize the apical junctional complex in epithelial cells. J. Cell Biol. 142, 847–857 (1998).
van Hengel, J. et al. Protein kinase C activation upregulates intercellular adhesion of α-catenin-negative human colon cancer cell variants via induction of desmosomes. J. Cell Biol. 137, 1103–1116 (1997).
Rangarajan, E. S. & Izard, T. Dimer asymmetry defines α-catenin interactions. Nat. Struct. Mol. Biol. 20, 188–193 (2013).
Thompson, P. M. et al. Identification of an actin binding surface on vinculin that mediates mechanical cell and focal adhesion properties. Structure 22, 697–706 (2014).
Wood, M. N. et al. α-Catenin homodimers are recruited to phosphoinositide-activated membranes to promote adhesion. J. Cell Biol. 216, 3767–3783 (2017).
Martin, P. & Lewis, J. Actin cables and epidermal movement in embryonic wound healing. Nature 360, 179–183 (1992).
Fernandez-Gonzalez, R. & Zallen, J. A. Wounded cells drive rapid epidermal repair in the early Drosophila embryo. Mol. Biol. Cell 24, 3227–3237 (2013).
Kwiatkowski, A. V. et al. In vitro and in vivo reconstitution of the cadherin-catenin-actin complex from Caenorhabditis elegans. Proc. Natl Acad. Sci. USA 107, 14591–14596 (2010).
Wickline, E. D. et al. αT-Catenin is a constitutive actin-binding α-catenin that directly couples the cadherin·catenin complex to actin filaments. J. Biol. Chem. 291, 15687–15699 (2016).
Kim, L. Y. et al. The structural basis of actin organization by vinculin and metavinculin. J. Mol. Biol. 428, 10–25 (2016).
Riedl, J. et al. Lifeact: a versatile marker to visualize F-actin. Nat. Methods 5, 605–607 (2008).
Nishikawa, T., Ishiyama, N., Wang, F. & Ikura, M. Backbone resonance assignments of the F-actin binding domain of mouse αN-catenin. Biomol. NMR Assign. 11, 21–24 (2016).
Shen, Y. & Bax, A. Homology modeling of larger proteins guided by chemical shifts. Nat. Methods 12, 747–750 (2015).
Mège, R.-M. & Ishiyama, N. Integration of cadherin adhesion and cytoskeleton at adherens junctions. Cold Spring Harb. Perspect. Biol. 9, a028738 (2017).
Zhang, J. et al. Actin at cell-cell junctions is composed of two dynamic and functional populations. J. Cell Sci. 118, 5549–5562 (2005).
Abe, K. & Takeichi, M. EPLIN mediates linkage of the cadherin-catenin complex to F-actin and stabilizes the circumferential actin belt. Proc. Natl Acad. Sci. USA 105, 13–19 (2008).
Imamura, Y., Itoh, M., Maeno, Y., Tsukita, S. & Nagafuchi, A. Functional domains of α-catenin required for the strong state of cadherin-based cell adhesion. J. Cell Biol. 144, 1311–1322 (1999).
Tachibana, K. et al. Two cell adhesion molecules, nectin and cadherin, interact through their cytoplasmic domain-associated proteins. J. Cell Biol. 150, 1161–1176 (2000).
Schneider, C. A., Rasband, W. S. & Eliceiri, K. W. NIH Image to ImageJ: 25 years of image analysis. Nat. Methods 9, 671–675 (2012).
Pardee, J. D. & Spudich, J. A. Purification of muscle actin. Methods Cell Biol. 24, 271–289 (1982).
Otwinowski, Z. & Minor, W. Processing of X-ray diffraction data collected in oscillation mode. Meth. Enzymol. 276, 307–326 (1997).
McCoy, A. J. et al. Phaser crystallographic software. J. Appl. Cryst. 40, 658–674 (2007).
Emsley, P. & Cowtan, K. Coot: model-building tools for molecular graphics. Acta Crystallogr. D. Biol. Crystallogr. 60, 2126–2132 (2004).
Adams, P. D. et al. PHENIX: a comprehensive Python-based system for macromolecular structure solution. Acta Crystallogr. D Biol. Crystallogr 66, (213–221 (2010).
Kay, L. E., Torchia, D. A. & Bax, A. Backbone dynamics of proteins as studied by 15N inverse detected heteronuclear NMR spectroscopy: application to staphylococcal nuclease. Biochemistry 28, 8972–8979 (1989).
Delaglio, F. et al. NMRPipe: a multidimensional spectral processing system based on UNIX pipes. J. Biomol. NMR 6, 277–293 (1995).
Johnson, B. A. & Blevins, R. A. NMR View: A computer program for the visualization and analysis of NMR data. J. Biomol. NMR 4, 603–614 (1994).
Shimada, I. NMR techniques for identifying the interface of a larger protein-protein complex: cross-saturation and transferred cross-saturation experiments. Meth. Enzymol. 394, 483–506 (2005).
Notredame, C., Higgins, D. G. & Heringa, J. T-Coffee: A novel method for fast and accurate multiple sequence alignment. J. Mol. Biol. 302, 205–217 (2000).
Robert, X. & Gouet, P. Deciphering key features in protein structures with the new ENDscript server. Nucleic Acids Res 42, W320–W324 (2014).
Ichii, T. & Takeichi, M. p120-catenin regulates microtubule dynamics and cell migration in a cadherin-independent manner. Genes Cells 12, 827–839 (2007).
Zacharias, D. A., Violin, J. D., Newton, A. C. & Tsien, R. Y. Partitioning of lipid-modified monomeric GFPs into membrane microdomains of live cells. Science 296, 913–916 (2002).
Phillips, J. C. et al. Scalable molecular dynamics with NAMD. J. Comput. Chem. 26, 1781–1802 (2005).
MacKerell, A. D. et al. All-atom empirical potential for molecular modeling and dynamics studies of proteins. J. Phys. Chem. B 102, 3586–3616 (1998).
Foloppe, N. & MacKerell, A. D. Jr All-atom empirical force field for nucleic acids: I. Parameter optimization based on small molecule and condensed phase macromolecular target data. J. Comput. Chem. 21, 86–104 (2000).
Jorgensen, W. L., Chandrasekhar, J., Madura, J. D., Impey, R. W. & Klein, M. L. Comparison of simple potential functions for simulating liquid water. J. Chem. Phys. 79, 926–935 (1983).
Humphrey, W., Dalke, A. & Schulten, K. VMD: visual molecular dynamics. J. Mol. Graph. 14, 33–38 (1996).
Martyna, G. J., Tobias, D. J. & Klein, M. L. Constant pressure molecular dynamics algorithms. J. Chem. Phys. 101, 4177–4189 (1994).
Feller, S. E., Zhang, Y., Pastor, R. W. & Brooks, B. R. Constant pressure molecular dynamics simulation: The Langevin piston method. J. Chem. Phys. 103, 4613–4621 (1995).
Darden, T., York, D. & Pedersen, L. Particle mesh Ewald: An N⋅log(N) method for Ewald sums in large systems. J. Chem. Phys. 98, 10089–10092 (1993).
Ryckaert, J.-P., Ciccotti, G. & Berendsen, H. J. C. Numerical integration of the cartesian equations of motion of a system with constraints: molecular dynamics of n-alkanes. J. Comput. Phys. 23, 327–341 (1977).
Hunter, J. D. Matplotlib: A 2D graphics environment. Comput. Sci. Eng. 9, 99–104 (2007).
Tepass, U. Crumbs, a component of the apical membrane, is required for zonula adherens formation in primary epithelia of Drosophila. Dev. Biol. 177, 217–225 (1996).
Fernandez-Gonzalez, R. & Zallen, J. A. Oscillatory behaviors and hierarchical assembly of contractile structures in intercalating cells. Phys. Biol. 8, 045005 (2011).
Leung, C. Y. B. & Fernandez-Gonzalez, R. Quantitative image analysis of cell behavior and molecular dynamics during tissue morphogenesis. Methods Mol. Biol. 1189, 99–113 (2015).
Mammalian expression vector pCA-αEcat-GFP was a gift from M. Takeichi. R2/7 cells were kindly provided by F. van Roy. We thank the CMCF beamline at the Canadian Light Source for assisting data collection. We thank G. Seabrook for help with NMR. We are grateful to K. Kakiguchi and H. Endoh for their help with electron microscopy. We thank the CSB Imaging Facility (U. of Toronto) and the Center for Advanced Microscopy (Northwestern U.) for support. We thank D. Kirchenbuechler for help with PIV analysis. We thank T. Hakoshima and T.Q.P. Uyeda for technical advice. Funding: Foundation Theme grant from the Canadian Institutes for Health Research (CIHR) (to M.I.), the Princess Margaret Cancer Foundation (to M.I.), the Canadian Foundation for Innovation (to M.I., U.T. and R.F.-G.), a project grant from the CIHR (to U.T.), a grant from the Canada First Research Excellence Fund (to U.T. and R.F.-G.), National Institutes of Health (NIH) grants GM076561 (to C.J.G.), P01HL071643 (to C.J.G.), HL134800 (to C.J.G.) and GM129312 (to C.J.G., N.I., M.I., U.T. and D.E.L.), National Science Foundation (NSF) grant CHE 12-3755 (to D.E.L.), a grant supported by CREST from JST, Japan (to S.Y.), and NSF Graduate Research Program DGE-1324585 (to M.N.W.). M.I. is the Canadian Research Chair (CRC) in Cancer Structural Biology, U.T. is the CRC in Epithelial Polarity and Development, and R.F.-G. is the CRC in Quantitative Cell Biology and Morphogenesis.