Abstract
Protein phosphatases undo the post-translational modifications of kinase-signaling networks, but phosphatase activation in cells is difficult to measure and interpret. Here, we report the design of a quantitative and high-throughput assay platform for monitoring cellular phosphatase activity toward specific phosphoprotein targets. Protein substrates of interest are purified recombinantly, phosphorylated in vitro using the upstream kinase, and adsorbed to 96-well plates. Total phosphatase extracts from cells are then added to trigger a solid-phase dephosphorylation reaction. After stopping the reaction, phosphoprotein levels are quantified by ELISA with a phospho-specific antibody, and the loss of phospho-specific immunoreactivity is used as the readout of phosphatase activity. We illustrate the generality of the method by developing specific phosphatase-activity assays for the three canonical mitogen-activated protein phospho-kinases: ERK, JNK, and p38. The assays capture changes in activity with a dynamic range of 25-100-fold and are sensitive to a limit of detection below 25,000 cells. When applied to cytokine-induced signaling, the assays revealed complex and dynamic regulation of phosphatases suggesting cross-communication and a means for cellular memory. Our assay platform should be beneficial for phosphoproteomic surveys and computational-systems models of signaling, where phosphatases are known to be important but their activities are rarely measured.