Using quantitative fluorescence microscopy and FRET imaging to measure spatiotemporal signaling events in single living cells.

The mechanisms that mediate how migratory eukaryotic cells amplify a shallow, extracellular chemoattractant gradient into a steep intracellular gradient of signaling components to guide chemotaxis remains unknown. To unravel these mechanisms, it is essential to quantitatively measure the spatiotemporal patterns of chemoattractant gradients, the dynamic movement of intracellular signaling pathway molecules, and the localized activation of these molecules in single living cells. Recent developments in live-cell fluorescence microscopy have permitted direct visualization and quantitative measurement of signal transduction events with high temporal and spatial resolution. Here, we outline fluorescence imaging methods to simultaneously visualize and quantitatively measure spatiotemporal changes in chemoattractant concentration by using the fluorescent tracer dye Alexa 594. Next, we provide a method to correlate the dynamic changes in ligand to the spatiotemporal changes in the second messenger phosphatidylinositol 3,4,5-triphosphate (PIP3) along the inner surface of the plasma membrane in live cells. Finally, we describe a fluorescence resonance energy transfer (FRET) method to determine the extent of heterotrimeric G protein activation in single living cells in response to various chemoattractant fields.