Optimized CRISPR/Cas9 Electroporation and Single Cell Cloning Protocol for Generating Pure Cellular Models in Human Immortalized Myoblasts

BackgroundGenome editing in human skeletal muscle research requires protocols that maximize delivery while preserving viability and clonal outgrowth. We sought to develop a reagent-free workflow for CRISPR/Cas9 editing in human immortalized myoblasts and to demonstrate its performance in two use cases, an IARS1 knockout and an MLIP homozygous knock-in. MethodsWe optimized electroporation parameters using a green fluorescent protein reporter to compare three electrical settings for transfection and survival in E6/E7 myoblasts, then applied ribonucleoprotein delivery for editing. We evaluated the effect of confluency at electroporation, performed single-cell cloning without antibiotics or fluorescence-activated sorting, and validated edits by high-resolution melting pre-screen followed by Sanger sequencing. ResultsElectroporation optimization identified one parameter set that maximized delivery while preserving viability. Performing electroporation at low confluency increased clonal outgrowth and editing rates. The workflow yielded an 84% success rate for IARS1 knockout and a 3.3% success rate for MLIP homozygous knock-in. High-resolution melting provided a very sensitive pre-screen, detecting 96% to 100% of actual edits, reducing the number of Sanger sequencing needed. Performance was reproducible across runs and myoblast lines and increasing single-cell seeding scaled yields without compromising purity. ConclusionsThis work provides a practical and reproducible selection-free protocol that couples electroporation optimization, low confluency editing, single-cell cloning, and high-resolution melting sorting to generate pure edited myoblast lines. The approach is applicable to disease modeling in neuromuscular research and clarifies feasibility boundaries for essential genes and homology-directed repair in these cells.