Mechanical forces are central to developmental, physiological and pathological processes1. However, limited understanding of force transmission within sub-cellular structures is a major obstacle to unravelling molecular mechanisms. Here we describe the development of a calibrated biosensor that measures forces across specific proteins in cells with pico-Newton (pN) sensitivity, as demonstrated by single molecule fluorescence force spectroscopy2. The method is applied to vinculin, a protein that connects integrins to actin filaments and whose recruitment to focal adhesions (FAs) is force-dependent3. We show that tension across vinculin in stable FAs is ~2.5 pN and that vinculin recruitment to FAs and force transmission across vinculin are regulated separately. Highest tension across vinculin is associated with adhesion assembly and enlargement. Conversely, vinculin is under low force in disassembling or sliding FAs at the trailing edge of migrating cells. Furthermore, vinculin is required for stabilizing adhesions under force. Together, these data reveal that FA stabilization under force requires both vinculin recruitment and force transmission, and that, surprisingly, these processes can be controlled independently.

In the absence of perfusable vascular networks, three-dimensional (3D) engineered tissues densely populated with cells quickly develop a necrotic core [1]. Yet the lack of a general approach to rapidly construct such networks remains a major challenge for 3D tissue culture [2–4]. Here, we 3D printed rigid filament networks of carbohydrate glass, and used them as a cytocompatible sacrificial template in engineered tissues containing living cells to generate cylindrical networks which could be lined with endothelial cells and perfused with blood under high-pressure pulsatile flow. Because this simple vascular casting approach allows independent control of network geometry, endothelialization, and extravascular tissue, it is compatible with a wide variety of cell types, synthetic and natural extracellular matrices (ECMs), and crosslinking strategies. We also demonstrated that the perfused vascular channels sustained the metabolic function of primary rat hepatocytes in engineered tissue constructs that otherwise exhibited suppressed function in their core.

We report the establishment of a library of micromolded elastomeric micropost arrays to modulate substrate rigidity independently of effects on adhesive and other material surface properties. We demonstrate that micropost rigidity impacts cell morphology, focal adhesions, cytoskeletal contractility, and stem cell differentiation. Furthermore, early changes in cytoskeletal contractility predicted later stem cell fate decisions at the single cell level.Cell function is regulated primarily by extracellular stimuli, including soluble and adhesive factors that bind to cell surface receptors. Recent evidence suggests that mechanical properties of the extracellular matrix (ECM), particularly rigidity, can also mediate cell signaling, proliferation, differentiation, and migration 1,2 . Culturing cells on hydrogels derived from natural ECM proteins at different densities has dramatic effects on cell adhesion, morphology, and function 3 . However, changing densities of the gels impacts not only mechanical rigidity, but also the amount of ligand, leaving uncertainty as to the relevant contribution of these two matrix properties on the observed cellular response. Synthetic ECM analogs such as polyacrylamide or polyethylene glycol gels, which vary rigidity by modulating the amount of cross-linker, has revealed that substrate rigidity alone can modulate many cellular functions, including stem cell differentiation 4-6 . However, altered cross-linker amount impacts not only bulk mechanics, but also molecular-scale material properties including porosity, surface chemistry, backbone flexibility, and binding properties of immobilized adhesive ligands 7,8 . Consequently, whether cells transduce substrate rigidity at the microscopic scale (eg sensing the rigidity between adhesion sites) or the nanoscopic scale (eg sensing local alterations in receptor-ligand binding characteristics) remains an open question 7,8 . While hydrogels will continue to play a major role in characterizing and controlling cell-material interactions, alternative approaches are necessary to further elucidate the basis by which cells sense changes in substrate rigidity.

Actomyosin contractility affects cellular organization within tissues in part through the generation of mechanical forces at sites of cell-matrix and cell-cell contact. While increased mechanical loading at cell-matrix adhesions results in focal adhesion growth, whether forces drive changes in the size of cell-cell adhesions remains an open question. To investigate the responsiveness of adherens junctions (AJ) to force, we adapted a system of microfabricated force sensors to quantitatively report cell-cell tugging force and AJ size. We observed that AJ size was modulated by endothelial cell-cell tugging forces: AJs and tugging force grew or decayed with myosin activation or inhibition, respectively. Myosin-dependent regulation of AJs operated in concert with a Rac1, and this coordinated regulation was illustrated by showing that the effects of vascular permeability agents (S1P, thrombin) on junctional stability were reversed by changing the extent to which these agents coupled to the Rac and myosin-dependent pathways. Furthermore, direct application of mechanical tugging force, rather than myosin activity per se, was sufficient to trigger AJ growth. These findings demonstrate that the dynamic coordination of mechanical forces and cell-cell adhesive interactions likely is critical to the maintenance of multicellular integrity and highlight the need for new approaches to study tugging forces.adherens junction | mechanotransduction | myosin | PDMS | traction force

Physical forces generated by cells drive morphologic changes during development and can feedback to regulate cellular phenotypes. Because these phenomena typically occur within a 3-dimensional (3D) matrix in vivo, we used microelectromechanical systems (MEMS) technology to generate arrays of microtissues consisting of cells encapsulated within 3D micropatterned matrices. Microcantilevers were used to simultaneously constrain the remodeling of a collagen gel and to report forces generated during this process. By concurrently measuring forces and observing matrix remodeling at cellular length scales, we report an initial correlation and later decoupling between cellular contractile forces and changes in tissue morphology. Independently varying the mechanical stiffness of the cantilevers and collagen matrix revealed that cellular forces increased with boundary or matrix rigidity whereas levels of cytoskeletal and extracellular matrix (ECM) proteins correlated with levels of mechanical stress. By mapping these relationships between cellular and matrix mechanics, cellular forces, and protein expression onto a bio-chemo-mechanical model of microtissue contractility, we demonstrate how intratissue gradients of mechanical stress can emerge from collective cellular contractility and finally, how such gradients can be used to engineer protein composition and organization within a 3D tissue. Together, these findings highlight a complex and dynamic relationship between cellular forces, ECM remodeling, and cellular phenotype and describe a system to study and apply this relationship within engineered 3D microtissues.biomechanics ͉ cell mechanics ͉ collagen ͉ morphogenesis ͉ PDMS