Current technologies for cellular metabolomics are complex, costly, and lack single-cell resolution. We propose optical metabolic imaging (OMI) as a tool to noninvasively monitor cellular metabolism on a single-cell level. OMI uses two-photon microscopy and time correlated single photon counting to measure the fluorescence lifetime and intensity of endogenous metabolic coenzymes NAD(P)H and FAD. However, the biochemical basis of changes in OMI parameters such as redox ratio (NAD(P)H intensity divided by FAD intensity), NAD(P)H lifetime, and FAD lifetime remains unclear. A thorough understanding of these biochemical sources of contrast is needed to interpret OMI studies of drug efficacy, and thus leverage OMI as a tool for drug screening and metabolic research. To address this issue, we profiled 11 CRISPR-mediated knockout human cell lines with single gene deletions corresponding to mitochondrial proteins involved in characterized pathways, and complexes ranging from oxidative phosphorylation to mitochondrial structure. The wild-type and knockout cells were plated at an equal density on 35mm imaging dishes in parallel with additional plates for other analyses. After 48hrs, cells were imaged or collected for mass spectrometry-based metabolomic, proteomic, and lipidomic analyses. OMI of the wild-type and knockout haploid cells yielded statistically significant differences in redox ratio, NAD(P)H and FAD mean lifetimes (ANOVA, p<0.005) that were specific to the knockout. Comparisons between OMI and parallel multi-omic analysis of each knockout are currently underway. This comprehensive dataset will provide a better understanding of the biochemical basis for OMI measurements, which could enable robust, single-cell monitoring of metabolic activity.