Fluctuations in osmotic pressure represent a critical challenge of the cellular environment. Differences in solution composition across plasma membranes cause bulk water movement in the direction of decreasing water activity, driving cell shrinkage or swelling. Cells dynamically respond to such stresses with osmo-regulatory mechanisms aimed at maintaining volume (tonic) control. Depending on the cell?s tolerance for mechanical stress, the adapted state may only partially correct the underlying osmotic imbalance. As a result, variations in intracellular water activity also perturb osmotically sensitive interactions that involve changes in molecular hydration. Osmotic stress arises from exposure to non-isotonic environments or rapid metabolic turnover in proliferating cells, and an increasing number of human diseases are connected to persistent osmotic stress. Osmotic pressure is therefore a parameter of interest to many areas of biomedical research. Current technologies cannot directly access osmotic pressure inside the cell. They infer osmotic pressure from functional or other correlates such as cell volume, gas vesicles, gene expression or macromolecular crowding. These indirect metrics, which are particular to different cell types but not specific to osmotic disturbances, limit their general utility. Direct access to intracellular osmotic pressure would enable investigators to establish a standard metric for evaluating osmotic responses, and compare different cellular systems or stress conditions. To address this unmet need, this proposal is aimed at validating a novel solution to directly report intracellular osmotic pressure using common imaging and flow cytometric instrumentation. Our approach is based on osmotically sensitive transcription factors, which bind high- and low-affinity DNA target sequences with distinct dependence on osmotic pressure. We postulate that differential transactivation of reporter genes by osmotically sensitive transcription factors at high- and low-affinity DNA enhancers could yield a direct ratiometric readout of the intracellular osmotic pressure. To validate this concept, we will use as initial design the transcription factor PU.1, whose osmotic sensitivities are characterized. We will 1) construct fluorescent protein reporter systems that are differentially responsive to osmotic pressure. 2) We will validate their operational basis using osmotically impaired mutant factors and calibrate the osmotic pressure readout in live cells. 3) To maximize the addressable range of organisms, we will generalize our design to remove the requirement for factor-specific transcriptional machinery. 4) Finally, we will integrate a time-sensitive feature into the sensor by controlling metabolic reporter turnover. An emphasis in our approach is a modular design that will accept a wide range of alternate transcription factors, promoters, and reporter moieties. This feature greatly enhances risk management. If successful, these innovations will lead to a direct and non-invasive approach for directly determining the latency, rate, and completeness of hypo- and hyperosmotic stress response by cells from all kingdoms of life.
Living cells are highly sensitive to stress caused by imbalances in their internal water content, known as osmotic pressure. Currently, osmotic pressure within living cells is not directly measurable by experiment, a shortcoming that limits our understanding of how cells respond to various environmental stresses. This proposal will investigate a novel approach to directly measure osmotic pressure in living cells from all the kingdoms of life.
