Summary The cytosolic protein α–catenin is a postulated force-transducer at cadherin complexes [1]. The demonstration of force activation, identification of consequent downstream events in live cells, and development of tools to study these dynamic processes in living cells are central to elucidating the role of α–catenin in cellular mechanics and tissue function [2–10]. Here we demonstrate that α–catenin is a force-activatable mechano-transducer at cell-cell junctions, using an engineered α-catenin conformation sensor, based on fluorescence resonance energy transfer (FRET). This sensor reconstitutes α-catenin-dependent functions in α-catenin depleted cells, and recapitulates the behavior of the endogenous protein. Dynamic imaging of cells expressing the sensor demonstrated that α-catenin undergoes immediate, reversible conformational switching, in direct response to different mechanical perturbations of cadherin adhesions. Combined magnetic twisting cytometry with dynamic FRET imaging [11] revealed rapid, local conformational switching, upon the mechanical stimulation of specific cadherin bonds. At acutely stretched cell-cell junctions, the immediate, reversible conformational change further reveals that α-catenin behaves like an elastic spring in series with cadherin and actin. The force-dependent recruitment of vinculin—a principal α-catenin effector—to junctions requires the vinculin-binding-site of the α–catenin sensor [1, 12–16]. In cells, the relative rates of force-dependent α–catenin conformation switching and vinculin recruitment reveal that α–catenin activation and vinculin recruitment occur sequentially rather than in a concerted process, with vinculin accumulation being significantly slower. This engineered α-catenin sensor revealed that α–catenin is a reversible, stretch-activatable sensor that mechanically links cadherin complexes and actin, and is an indispensable player in cadherin-specific mechano-transduction at intercellular junctions.

This report elucidates an E-cadherin-based force-transduction pathway that triggers changes in cell mechanics through a mechanism requiring epidermal growth factor receptor (EGFR), phosphoinositide 3-kinase (PI3K), and the downstream formation of new integrin adhesions. This mechanism operates in addition to local cytoskeletal remodeling triggered by conformational changes in the E-cadherin-associated protein α-catenin, at sites of mechanical perturbation. Studies using magnetic twisting cytometry (MTC), together with traction force microscopy (TFM) and confocal imaging identified force-activated E-cadherin-specific signals that integrate cadherin force transduction, integrin activation and cell contractility. EGFR is required for the downstream activation of PI3K and myosin-II-dependent cell stiffening. Our findings also demonstrated that α-catenin-dependent cytoskeletal remodeling at perturbed E-cadherin adhesions does not require cell stiffening. These results broaden the repertoire of E-cadherin-based force transduction mechanisms, and define the force-sensitive signaling network underlying the mechano-chemical integration of spatially segregated adhesion receptors.

Imaging the location and dynamics of individual interacting protein pairs is essential but often difficult because of the fluorescent background from other paired and non-paired molecules, particularly in the sub-diffraction cellular space. Here we develop a new method combining bimolecular fluorescence complementation and photoactivated localization microscopy for super-resolution imaging and single-molecule tracking of specific protein–protein interactions. The method is used to study the interaction of two abundant proteins, MreB and EF-Tu, in Escherichia coli cells. The super-resolution imaging shows interesting distribution and domain sizes of interacting MreB–EF-Tu pairs as a subpopulation of total EF-Tu. The single-molecule tracking of MreB, EF-Tu and MreB–EF-Tu pairs reveals intriguing localization-dependent heterogonous dynamics and provides valuable insights to understanding the roles of MreB–EF-Tu interactions.

We apply fast relaxation imaging (FReI) as a novel technique for investigating the folding stability and dynamics of proteins within polyacrylamide hydrogels, which have diverse and widespread uses in biotechnology. FReI detects protein unfolding in situ by imaging changes in fluorescence resonance energy transfer (FRET) after temperature jump perturbations. Unlike bulk measurements, diffraction-limited epifluorescence imaging combined with fast temperature perturbations reveals the impact of local environment effects on protein-biomaterial compatibility. Our experiments investigated a crowding sensor protein (CrH2) and phosphoglycerate kinase (PGK), which undergoes cooperative unfolding. The crowding sensor quantifies the confinement effect of the cross-linked hydrogel: the 4% polyacrylamide hydrogel is similar to aqueous solution (no confinement), while the 10% hydrogel is strongly confining. FRAP measurements and protein concentration gradients in the 4% and 10% hydrogels further support this observation. PGK reveals that noncovalent interactions of the protein with the polymer surface are more important than confinement for determining protein properties in the gel: the mere presence of hydrogel increases protein stability, speeds up folding relaxation, and promotes irreversible binding to the polymer even at the solution-gel interface, whereas the difference between the 4% and the 10% hydrogels is negligible despite their large difference in confinement. The imaging capabilities of FReI, demonstrated to be diffraction limited, further revealed spatially homogeneous protein unfolding across the hydrogels at 500 nm length scales and revealed differences in protein properties at the gel-solution boundary.