Twin-prime editing enables endogenous protein visualization and mutant allele tracking

Abstract

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.