Double-stranded DNA donors and CRISPR-Cas9 for universal correction of mutations causing cystic fibrosis in human airway cells.

Cystic fibrosis (CF) is a devastating genetic disease caused by mutations in the cystic fibrosis transmembrane conductance regulator (CFTR) gene. As morbidity and mortality from CF results from a lack of mucus clearance that leads to chronic bacterial infections and progressive loss of lung function, site-specific insertion of a CFTR cDNA into the endogenous CFTR locus in airway basal stem cells (ABSCs) could prove curative for all disease-causing mutations. This study describes the development of genome-editing approach utilizing nonviral reagents, designed to be packaged into nonviral delivery systems. An sgRNA targeting the 5' untranslated region (UTR) of CFTR was characterized as directing high on-target cutting and displaying a safe off-target profile. Airway cell lines electroporated with chemically modified (1-Aminohexane, AmC6), linear double-stranded DNA (ldsDNA) constructs were utilized as a homology directed repair (HDR) donor, initially optimized with an mCitrine reporter. Expectedly, when the 780 bp mCitrine cDNA was replaced with the 4.4 kb CFTR cDNA, integration efficiency dropped significantly. However, 1%-2% integration of codon-optimized donors was sufficient to restore CFTR expression in the bulk-edited population of a human bronchial epithelial cell line, 16HBE14o- (16HBE), to levels reaching 50% of wild-type expression as measured by western blot. Electrophysiological validation of CFTR ion channel function measured via Ussing chamber assay revealed that these bulk-edited populations exhibit greater than 40% restoration of the chloride ion currents of the measured wild-type controls. These results demonstrate that low levels of CFTR integration can be made therapeutically relevant by optimizing the designs of gene editing reagents. Importantly, this work utilizes nonviral-editing reagents, an essential step toward in vivo gene therapy for CF.