A variety of methods are available for monitoring of cell forces. In this paper, a novel approach using an in-plane deformable microsystem is utilized in which displacements induced by cultured cells are measured via optical profilometry. The high resolution obtainable from profilometry gives an order of magnitude improvement in measurement resolution compared to conventional optical techniques and demonstrates a spatial measurement resolution of 12 nm (126 nN). The work focuses on both fixed and living fibroblasts and epithelial cells with estimates of forces exerted significantly higher using living cells compared to fixed cells. The methodology was developed to give no restriction to the cell environment, thereby allowing the potential for a broad range of experiments in the field.
The aim of this study was to compare uniaxial traction forces exerted by different cell types using a novel sensor design and to test the dependence of measured forces on cytoskeletal integrity. The sensor design detects forces generated between 2 contact points by cells spanning a gap. The magnitude of these forces varied according to cell type and were dependent on cytoskeletal integrity. The response time for drug-induced cytoskeletal disruption also varied between cell types: dermal fibroblasts exerted the greatest forces and had the slowest drug response times; EBV-transformed epithelial cells also had slow cytoskeletal depolymerisation times but exerted the lowest forces overall. Conversely, lung epithelial tumor cells exerted low forces but had the fastest depolymerisation drug response. These results provide proof of principle for a new design of force-measurement sensor based on optical interferometry, an approach that can be used to study cytoskeletal dynamics in real time.
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