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The intracellular force transmission is critical for various biological events. It has recently been suggested that the cell nucleus is exposed to the mechanical forces transmitted through the cytoskeleton from outside the cell, and the changes in nuclear morphology possibly affect the regulation of cell functions. However, it remains unclear at this stage how much force is applied to the intracellular nucleus and how the morphology of the nucleus changes during cell contraction or release of intracellular tension. In the present study, we investigated the changes in the traction forces of the vascular smooth muscle cells (SMCs) cultured on polydimethylsiloxane-based elastic micropillar array substrates in the state of the contraction with actomyosin activation or the release of tension with F-actin disruption. We found that the total tension of SMCs increased from ~100 nN to ~200 nN following cell contraction, and decreased to less than 1/5 following F-actin disruption. We also measured the three-dimensional morphology of the nucleus of SMCs in the state of contraction or tension releasing using confocal microscopy. The nuclear height decreased significantly by ~10% following cell contraction, and dramatically increased by ~50% following F-actin disruption, indicating that the intracellular nuclei were compressed physiologically. Based on these mechanical and morphological data, we analyzed the force applied to the nucleus using a simple two-dimensional cell model, and estimated the compressive stiffness of the nucleus for the first time. Even though the nucleus has been believed to be the stiffest organelle, the estimated nuclear stiffness was in the range of 10 nN/!m in this study, indicating that the intracellular nucleus has a relatively large deformability similar to that of cytoplasm.

Quantitative analysis of cellular traction forces using a micropillar substrate and estimation of the intracellular force applied to the nucleus