BACKGROUND: Single cell gene profiling has been successfully applied to cultured cells. However, isolation and preservation of a cell's native gene expression state from an intact embryo remain problematic. RESULTS: Here, we present a strategy for in vivo single cell profiling that optimizes cell identification, isolation and amplification of nucleic acids with nominal bias and sufficient material detection. We first tested several photoconvertible fluorescent proteins to selectively mark a cell(s) of interest in living chick embryos then accurately identify and isolate the same cell(s) in fixed tissue slices. We determined that the dual color mDendra2 provided the optimal signal/noise ratio for this purpose. We developed proper procedures to minimize cell death and preserve gene expression, and suggest nucleic acid amplification strategies for downstream analysis by microfluidic reverse transcriptase quantitative polymerase chain reaction or RNAseq. Lastly, we compared methods for single cell isolation and found that our fluorescence-activated cell sorting (FACS) protocol was able to preserve native transcripts and generate expression profiles with much higher efficiency than laser capture microdissection (LCM). CONCLUSIONS: Quantitative single cell gene expression profiling may be accurately applied to interrogate complex cell dynamics events during embryonic development by combining photoconversion cell labeling, FACS, proper handling of isolated cells, and amplification strategies.