Measurement of cellular traction forces during confined migration.

To migrate efficiently through tissues, cells must transit through small constrictions within the extracellular matrix. However, in vivo environments are geometrically, mechanically, and chemically complex, and it has been difficult to understand how each of these parameters contribute to the propulsive strategy utilized by cells in different confining environments. To address this, we employed a sacrificial micromolding approach to generate polymer substrates with tunable stiffness, controlled adhesivity, and user-defined microscale geometries. We combined this together with live-cell imaging and three-dimensional traction force microscopy to quantify the forces that cells use to transit through constricting channels. Surprisingly, rather than enlarging the constriction via pushing forces, we observe that mesenchymal cells migrating through compliant constrictions generate inwardly directed contractile forces that decrease the size of the opening and pull the channel walls closed around the nucleus. This had the effect of increasing nuclear deformation compared to cells migrating through comparably sized rigid confinements. Additionally, the nucleus took longer to transit through compliant constrictions compared to similarly sized rigid constrictions. These findings show that nuclear deformation during confined migration can be accomplished by internal cytoskeletal machinery rather than by reactive forces from the substrate, and our approach provides a mechanism to test between different models for how cells translocate their nucleus through narrow constrictions. The methods, analysis, and results presented here will be useful to understand how cells choose between propulsive strategies in different physical environments.