High-throughput quantitation of pathogen-induced calcium signals captured through live-cell fluorescence microscopy

Many intracellular pathogens manipulate host cell calcium to facilitate their survival and replication. Live-cell microscopy using fluorescent calcium indicators has become an indispensable tool for characterizing the mechanisms underlying both homeostatic and pathogen-induced cellular calcium dynamics, but such imaging must be coupled with robust quantitative analysis. Further, calcium imaging is most powerful when paired with reductive studies targeting calcium-modulating proteins. The lack of specific inhibitors or agonists to directly target most pathogen-induced calcium signals precludes many of the approaches that have allowed for robust characterization of major eukaryotic cell calcium signaling mechanisms, such as ER Ca²⁺ release by inositol triphosphate receptors. Given this, we sought to develop quantitative imaging pipelines tailored for the characterization of pathogen-induced calcium signals. Using rotavirus as a prototypical calcium-modulating pathogen, we developed and optimized a suite of computational tools for automated quantitation of both intra- and inter-cellular calcium signals detected via live-cell imaging of infected epithelial monolayers expressing genetically encoded calcium indicators. Using recombinant strains of rotavirus that express fluorescent markers, we developed a system that allows for automated detection of rotavirus-infected cells and normalization of signals to infectivity. All tools were built in ImageJ, making them freely available and adaptable across operating systems and microscope setups. These tools required minimal active time from the user and allowed for the extraction of signal parameters previously unquantifiable, increasing the speed and breadth of characterization.