Abstract

Cardiovascular disease is the leading cause of death in the U.S. and the rest of the World. Physiological assays with iPSC-cardiomyocytes are powerful new tools for the development of new drugs, particularly assays for measuring action potential and calcium transient kinetics. However, high throughput methods to quantify cardiomyocyte contraction and relaxation kinetics are less prevalent. Furthermore, there are no optical high-throughput methods to quantify cardiomyocyte force generation and cell stiffness changes simultaneously during a cardiac cycle. Here, we present a method to measure the mechanical tension produced by a monolayer culture of cardiomyocytes as well as the elastic modulus. By utilizing the forces generated by the cells themselves to probe the material properties, our method is fully non-invasive. iPSC-cardiomyocytes were cultured on deformable substrates doped with fluorescent beads and were live stained with fluorescently-labeled wheat germ agglutinin (WGA). We acquired high framerate videos of the iPSC-cardiomyocytes labeled with WGA and of the fluorescently-labeled beads. From the WGA movies, we determined the strain field of the cell monolayer and its variations during the cardiac cycle. Similarly, the substrate deformations were measured by tracking the motion of the beads, and were used to compute the cell-substrate forces using Traction Force Microscopy. From these datasets, we computed the intracellular stresses using Monolayer Stress Microscopy, and estimated the elastic modulus of the cells by fitting the relation between the independently measured stress and strain maps to Hooke's law. Our method is comparable to existing data from invasive methods such as AFM. Additionally, we present a high-throughput application of our method by testing a series of benchmark compounds on monolayers of iPSC-cardiomyocytes in multi-well plates. This work represents the first high-throughput assay for simultaneous force and stiffness characterization in iPSC-cardiomyocytes.

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