Frameshift Mutations in Exon 1 of Fgf10 Frequently Yield Morphologically Normal Embryos

Prime Editing Mediated Generation and Correction of the mdx5cv Mutation Restores Dystrophin Expression in Myoblasts

Duchenne muscular dystrophy (DMD) is caused by mutations in the DMD gene that abolish dystrophin expression. Prime editing enables precise genome modification without generating double-strand DNA breaks or requiring donor DNA templates. We established an in vitro prime editing workflow to generate and subsequently correct the mdx5cv mutation in mouse C2C12 myoblasts. Following optimization of engineered prime editing guide RNAs (epegRNAs) and PAM-flexible prime editors, wild-type cells were edited, clonally isolated, and genotyped. Mutation correction was then evaluated using optimized epegRNA designs. Two rounds of prime editing introduced the mdx5cv mutation into approximately 20% of alleles in C2C12 cells creating the mdx5cv C2C12 cell line. Clonal isolation yielded five homozygous mutant clones among 59 expanded colonies. Optimization studies identified an epegRNA containing a 16 nucleotide reverse transcription template and a 10 nucleotide primer binding site (RTT16/PBS10) as the most efficient design. Correction of the pathogenic allele reached approximately 26%, whereas longer PBS lengths reduced editing efficiency. In silico off-target analysis using Cas-OFFinder identified no candidate genomic loci with fewer than three mismatches for the spacer sequences used in either mutation generation or correction, suggesting a favorable predicted specificity profile. Corrected mdx5cv C2C12 myoblasts retained their capacity to differentiate into multinucleated myotubes. Representative Western blot analysis detected dystrophin protein expression in differentiated corrected mdx5cv myotubes, consistent with successful correction of the pathogenic mutation. These findings establish a robust prime editing platform for both the generation and correction of the mdx5cv mutation and provide proof of concept that precise correction of the pathogenic mutation is associated with restoration of dystrophin expression following myogenic differentiation.

Evaluation of germline transmission of electroporation-mediated double gene-edited cattle lines.

Gene editing in livestock using clustered regularly interspaced short palindromic repeats/CRISPR-associated protein 9 (CRISPR/Cas9) offers a promising approach for genetic improvement in cattle. This study evaluated germline transmission and mutation stability of double-knockout cattle generated by zygote electroporation. Previously reported myostatin/beta-lactoglobulin (MSTN/BLG) and newly generated α-1,3-galactosyltransferase (GGTA1/BLG) double-knockout cattle were produced using CRISPR/Cas9-mediated genome editing. Targeted deep sequencing demonstrated extensive somatic mosaicism across multiple tissues. Computer-assisted sperm analysis (CASA) demonstrated normal sperm motility in MSTN/BLG double-knockout males. Fertilization of wild-type oocytes produced heterozygous embryos, with mutation frequencies of 37.76 ± 10.74% at the MSTN locus and 54.80 ± 7.73% at the BLG locus, as assessed by T7 endonuclease I (T7E1) assay. MSTN/BLG double-knockout sperm were subsequently used for embryo production and for artificial insemination of GGTA1/BLG double-knockout females. Healthy offspring were successfully obtained (n = 3), alongside one stillborn calf. Targeted deep sequencing of all four progenies revealed highly variable allele frequencies that deviated substantially from the approximately 50% expected for heterozygous germline transmission. In contrast, whole-genome sequencing (WGS) results were consistent with Mendelian expectations, underscoring the limitations of PCR-based targeted sequencing for assessing germline transmission in mosaic founders. These results show that CRISPR/Cas9-edited embryos generated by electroporation can develop into healthy, sexually mature cattle capable of germline transmission. While variable transmission rates were observed owing to founder mosaicism, non-mosaic F1 offspring were successfully generated. However, direct, embryo-mediated gene-editing strategies remain technically and economically challenging for large-scale commercial calf production, and reports in cattle are limited. This study provides a reference for future applications of gene-edited embryos and their germline propagation.

CRISPR-Mediated Gene Editing: Generating Gene Knockouts or Precision Base Changes in F0 Xenopus.

CRISPR/Cas9-based repair of a heterozygous HNF1A mutation in patient-derived hiPSCs

Human induced pluripotent stem cells (hiPSCs) represent a powerful platform for disease modeling, especially in monogenic diseases as they preserve the donor’s genetic background while enabling directed differentiation into disease-relevant cell types. This makes them highly suitable for studying disease mechanisms in a patient-specific and physiologically relevant context. Although CRISPR/Cas9 is widely applied for genome editing, precise correction of pathogenic variants in hiPSCs remains challenging due to the lack of standardized CRISPR component selection and experimental design. Here, we describe an optimized CRISPR-based strategy for correcting a heterozygous HNF1A frameshift mutation (c.235_236insG; p.Glu79Glyfs*16) in HNF1A-MODY patient-derived hiPSCs. Using electroporation, we efficiently delivered CRISPR components, including a ribonucleoprotein complex of Cas9 and single-guide RNA, along with a single-stranded oligodeoxynucleotide repair template. Corrected hiPSC lines were validated for pluripotency, absence of exogenous reprogramming factors, and off-target effects. Additionally, we discuss key technical challenges encountered during the editing process and provide practical recommendations that may improve the generation of mutation-corrected hiPSC lines. These guidelines could serve as a useful reference for researchers employing CRISPR-based strategies for generation of reliable disease modelling tools.

Genome-wide CRISPR screening identifies cellular factors controlling nonviral genome editing efficiency

To systematically map cellular factors constraining nonviral genome editing, influencing uptake and intracellular trafficking, we develop a genome-wide CRISPR screening platform linking perturbation of 19,114 genes to editing outcomes in human cells. We identify six negative regulators of delivery whose depletion increases editing efficiency by up to six-fold across diverse payloads, loci, and cell types. We test the top two factors, GJB2 and BET1L, in two distinct human models: correction of a pathogenic adenine base mutation in KCNJ13 and introduction of a cytosine base mutation in the GABAA receptor gene. Depletion of either improves base-editing outcomes by 6-fold, potentially through effects on delivery. In a patient-derived model of retinal channelopathy, knockdown of either gene improves lipid nanoparticle base editing efficiency by over 3.5-fold. This enables functional restoration of Kir7.1 ion channels in a subset of edited cells, highlighting cellular barriers as actionable targets to enhance the potency of genetic therapies. Low editing efficiency of nonviral delivery in post mitotic tissues presents a challenge to the field of gene therapy. Here, authors dissect the genetic regulators of nonviral delivery in post mitotic retinal epithelial cells describe strategies for improved base editor delivery and editing.

Genome-Wide CRISPR Screening Identifies Cellular Factors Controlling Nonviral Genome Editing Efficiency

A novel genome-wide CRISPR screening strategy that will facilitate the systematic engineering of novel nonviral genome editing delivery methods, where the identified novel gene hits can be further used to increase editing efficiency for other therapeutically relevant cell types.