For a quantitative analysis of intracellular mechanotransduction, it is crucial to know the mechanical properties of actin stress fibers in situ. Here we measured tensile properties of cultured aortic smooth muscle cells (SMCs) in a quasi-in situ tensile test in relaxed and activated states to estimate stiffness of their single stress fibers (SFs). An SMC cultured on substrates was held using a pair of micropipettes and detached from the substrate while maintaining its in situ cell shape and cytoskeletal integrity. Stretching up to approximately 15% followed by unloading was repeated three times to stabilize their tension-strain curves in the untreated (relaxed) and 10 microM-serotonin-treated (activated) condition. Cell stiffness defined as the average slope of the loading limb of the stable loops was approximately 25 and approximately 40 nN/% in relaxed and activated states, respectively. It decreased to approximately 10 nN/% following SF disruption with cytochalasin D in both states. The number of SFs in each cell measured with confocal microscopy decreased significantly upon serotonin activation from 21.5+/-3.8 (mean+/-SD, n=80) to 17.5+/-3.9 (n=77). The dynamics of focal adhesions (FAs) were observed in adherent cells using surface reflective interference contrast microscopy. FAs aligned and elongated along the cell major axis following activation and then merged with each other, suggesting that the decrease in SFs was caused by their fusion. Average stiffness of single SFs estimated by the average decrease in whole-cell stiffness following SF disruption divided by the average number of SFs in each cell was approximately 0.7 and approximately 1.6 nN/% in the relaxed and activated states, respectively. Stiffening of single SFs following SF activation was remarkably higher than stiffening at the whole-cell level. Results indicate that SFs stiffen not only due to activation of the actomyosin interaction, but also due to their fusion, a finding which would not be obtained from analysis of isolated SFs.
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