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Current preclinical models fail to capture the mechanics of oligodendrocyte myelination. Lasli et al. 1 now demonstrate that the mechanical compliance of the axonal niche is a key determinant of oligodendrocyte maturation. By developing a platform that mimics the extreme mechanical softness of the central nervous system, they reveal myelination as a mechanically gated process as much as a biochemically regulated one.

Communications Biology is inviting submissions on the topic of live microscopy – from new tools to emerging techniques, from conventional to advanced light microscopy - with the aim of publishing high-quality research devoted to advance our understanding of biology.

Many cellular processes rely on cells generating or responding to nanoscale mechanical forces. This protocol describes STED–traction force microscopy (STFM), which allows these forces to be measured with higher resolution and accuracy than standard TFM. Cells continuously exert or respond to mechanical force. Measurement of these nanoscale forces is a major challenge in cell biology; yet such measurement is essential to the understanding of cell regulation and function. Current methods for examining mechanical force generation either necessitate dedicated equipment or limit themselves to coarse-grained force measurements on the micron scale. In this protocol, we describe stimulated emission depletion traction force microscopy—STED-TFM (STFM), which allows higher sampling of the forces generated by the cell than conventional TFM, leading to a twofold increase in spatial resolution (of up to 500 nm). The procedure involves the preparation of functionalized polyacrylamide gels loaded with fluorescent beads, as well as the acquisition of STED images and their analysis. We illustrate the approach using the example of HeLa cells expressing paxillin-EGFP to visualize focal adhesions. Our protocol uses widely available laser-scanning confocal microscopes equipped with a conventional STED laser, open-source software and common molecular biology techniques. The entire STFM experiment preparation, data acquisition and analysis require 2–3 d and could be completed by someone with minimal experience in molecular biology or biophysics.

This protocol uses fluorescence recovery after photobleaching (FRAP) to dissect the reaction dynamics leading to protein turnover in macromolecular complexes in living cells. Proteins within most macromolecular complexes or organelles continuously turn over. This turnover results from association and dissociation reactions that are mediated by each of the protein's functional domains. Thus, studying organelle or macromolecular formation from the bottom up using theoretical and computational modeling approaches will necessitate the determination of all of these reaction rates in vivo . Yet current methods for examining protein dynamics either necessitate highly specialized equipment or limit themselves to basic measurements. In this protocol, we describe a broadly applicable method based on fluorescence recovery after photobleaching (FRAP) for determining how many reaction processes participate in the turnover of any given protein of interest, for characterizing their apparent association and dissociation rates, and for determining their relative importance in the turnover of the overall protein population. Experiments were performed in melanoma M2 cells expressing mutant forms of ezrin that provide a link between the plasma membrane and the cortical actin cytoskeleton. We also describe a general strategy for the identification of the protein domains that mediate each of the identified turnover processes. Our protocol uses widely available laser-scanning confocal microscopes, open-source software, graphing software and common molecular biology techniques. The entire FRAP experiment preparation, data acquisition and analysis require 3–4 d.

The cytoplasm is the largest part of the cell by volume and hence its rheology sets the rate at which cellular shape changes can occur. Recent experimental evidence suggests that cytoplasmic rheology can be described by a poroelastic model, in which the cytoplasm is treated as a biphasic material consisting of a porous elastic solid meshwork (cytoskeleton, organelles, macromolecules) bathed in an interstitial fluid (cytosol). In this picture, the rate of cellular deformation is limited by the rate at which intracellular water can redistribute within the cytoplasm. However, direct supporting evidence for the model is lacking. Here we directly validate the poroelastic model to explain cellular rheology at short timescales using microindentation tests in conjunction with mechanical, chemical and genetic treatments. Our results show that water redistribution through the solid phase of the cytoplasm (cytoskeleton and macromolecular crowders) plays a fundamental role in setting cellular rheology at short timescales. It has been suggested that the cytoplasm of living cells can be described as a porous elastic meshwork bathed in an interstitial fluid. Microindentation tests now show that intracellular water redistribution plays a fundamental role in cellular rheology and that at physiologically relevant timescales cellular responses to mechanical stresses are consistent with such a poroelastic model.

Quantifying small, rapidly progressing three-dimensional forces generated by cells remains a major challenge towards a more complete understanding of mechanobiology. Traction force microscopy is one of the most broadly applied force probing technologies but ascertaining three-dimensional information typically necessitates slow, multi-frame z-stack acquisition with limited sensitivity. Here, by performing traction force microscopy using fast single-frame astigmatic imaging coupled with total internal reflection fluorescence microscopy we improve the temporal resolution of three-dimensional mechanical force quantification up to 10-fold compared to its related super-resolution modalities. 2.5D astigmatic traction force microscopy (aTFM) thus enables live-cell force measurements approaching physiological sensitivity. Quantifying rapidly progressing three-dimensional forces generated by cells remains a major challenge in mechanobiology. Here, the authors show that combining traction force microscopy with astigmatic imaging permits sensitive out-of-plane force estimation on the second timescale.

Living systems rely, for biological function, on the spatiotemporal organization of their structures. Cellular order naturally emerges by dissipation of energy. Consequently, energy-consuming processes operating far from thermodynamic equilibrium are a necessary condition to enable biological systems to respond to environmental cues that allow their transitions between different steady-states. Such self-organization was predicted for the actin cytoskeleton in theoretical considerations and has repeatedly been observed in cell-free systems. We now demonstrate in our recent work how self-organizing actin patterns such as vortices, stars, and asters may allow cells to adjust their membrane architecture without affecting their cell mechanical properties.

Quantifying small, rapidly evolving forces generated by cells is a major challenge for the understanding of biomechanics and mechanobiology in health and disease. Traction force microscopy remains one of the most broadly applied force probing technologies but typically restricts itself to slow events over seconds and micron-scale displacements. Here, we improve >2-fold spatially and >10-fold temporally the resolution of planar cellular force probing compared to its related conventional modalities by combining fast two-dimensional total internal reflection fluorescence super-resolution structured illumination microscopy and traction force microscopy. This live-cell 2D TIRF-SIM-TFM methodology offers a combination of spatio-temporal resolution enhancement relevant to forces on the nano- and sub-second scales, opening up new aspects of mechanobiology to analysis. Quantifying rapid and small cellular forces is a major challenge in mechanobiology. Here, the authors show a >2-fold spatially and >10-fold temporally force sampling improvement combining traction force microscopy with total internal reflection fluorescence super-resolution structured illumination microscopy.