Abstract
The multistep process of cell migration requires cells to dynamically couple to extracellular interfaces and generate traction force or friction for displacement of the cell body. When deformed, biopolymer networks, including fibrillar collagen and fibrin, undergo a nonlinear elasticity change that is termed strain stiffening and is commonly measured by bulk rheology. It remains poorly characterized, however, whether forces generated by moving cells suffice to induce strain stiffening. To detect strain stiffening at the leading edge of normal and tumor cells moving across fibrillar type I collagen, we combined AFM nanoindentation and differential field probing with confocal reflection microscopy. In different cell models, gradient-like fiber realignment, densification, and elevation of Young's modulus ahead of the leading edge were observed, with peak increases of up to 1.15 kPa near the leading edge. Moving fibroblasts generated a larger anterograde strain field with a higher amplitude and up to 6-fold increased cumulative strain stiffening (52 kPa) compared with mesenchymal HT1080 fibrosarcoma cells (8.8 kPa) and epithelial SCC38 cancer cells (9.8 kPa). Collectively moving SCC38 cells produced 4-fold increased cumulative strain stiffening (38 kPa) compared with individually moving SCC38 cells in a beta1 integrin- and actomyosin-dependent manner. This indicates that the extent of strain stiffening by the leading edge of moving cells scales with cell type, multicellular cooperativity, integrin availability, and contractility. By straining, migrating cells realign and densify fibrillar extracellular matrix and thus adopt an autonomous strategy to move on a "traveling wave" of stiffened substrate, which reaches levels sufficient for mechanosensory activation and self-steering of migration.
