Precise analysis of endogenous protein behavior in mosaic tissues requires strategies that both modify native alleles and identify successfully edited cells at single-cell resolution. Here, we establish a CRISPR-based genome-editing platform that combines endogenous protein visualization with allele-specific mutant tracking in vivo. Using the chick neural tube and {beta}-catenin as a model system, we validate cytosine base editing, prime editing, and twin-prime editing by introducing stabilizing mutations that reproduce their expected cellular and morphological phenotypes. We then develop a twin-prime editing strategy that couples installation of a defined oncogenic mutation to simultaneous insertion of minimal peptide tags, thereby making productive editing directly observable at single-cell resolution without clonal selection. Comparative analysis of ALFA, V5, and split-GFP tags identifies split-GFP as the most reliable strategy for endogenous protein visualization in vivo, and fluorescence-based cell selection strongly enriches for the intended twin-prime editing product. Finally, we extend the approach to endogenous wild-type {beta}-catenin, identifying a functionally neutral insertion site that enables visualization at physiological, non-stabilized levels while preserving normal protein behavior. Together, these results establish twin-prime editing as a versatile platform for directly linking precise endogenous genome modification to protein visualization and allele-specific mutation analysis in intact vertebrate tissues.
Menendez, A., Ramos, C., PONS, S.
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