Despite the utility of CRISPR-Cas9 nucleases for genome editing, the potential for off-target activity limits their application, especially for therapeutic purposes1,2. We developed a yeast-based assay to identify optimized Streptococcus pyogenes Cas9 (SpCas9) variants that enables simultaneous evaluation of on- and off-target activity. We screened a library of SpCas9 variants carrying random mutations in the REC3 domain and identified mutations that increased editing accuracy whilst maintaining editing efficiency. We combined four beneficial mutations to generate evoCas9, a variant that has fidelity exceeding both wild-type (79-fold improvement) and rationally designed Cas9 variants3,4 (4-fold average improvement), while maintaining near wild-type on-target editing efficiency (90% median residual activity). Evaluating evoCas9 on endogenous genomic loci, we demonstrated a substantial improved specificity and observed no off-target sites for 4 of the 8 sgRNAs tested. Finally, we showed that following long-term expression (40 days), evoCas9 strongly limited the unspecific cleavage of a difficult-to-discriminate off-target site and fully abrogated the cleavage of two additional off-targets.

The method of delivery of CRISPR-Cas9 into target cells is a strong determinant of efficacy and specificity in genome editing. Even though high efficiency of Cas9 delivery is necessary for optimal editing, its long-term and high levels of expression correlate with increased off-target activity. We developed vesicles (VEsiCas) carrying CRISPR-SpCas9 ribonucleoprotein complexes (RNPs) that are efficiently delivered into target cells through the fusogenic glycoprotein of the vesicular stomatitis virus (VSV-G). A crucial step for VEsiCas production is the synthesis of the single guide RNA (sgRNA) mediated by the T7 RNA polymerase in the cytoplasm of producing cells as opposed to canonical U6-driven Pol III nuclear transcription. In VEsiCas, the absence of DNA encoding SpCas9 and sgRNA allows rapid clearance of the nuclease components in target cells, which correlates with reduced genome-wide off-target cleavages. Compared with SpCas9 RNPs electroporation, which is currently the method of choice to obtain transient SpCas9 activity, VEsiCas deliver the nuclease with higher efficiency and lower toxicity. We show that a wide variety of cells can be edited through VEsiCas, including a variety of transformed cells, induced pluripotent stem cells (iPSCs), and cardiomyocytes, in vivo. VEsiCas is a traceless CRISPR-Cas9 delivery tool for efficient and safe genome editing that represents a further advancement toward the therapeutic use of the CRISPR-Cas9 technology.

In vivo application of the CRISPR-Cas9 technology is still limited by unwanted Cas9 genomic cleavages. Long-term expression of Cas9 increases the number of genomic loci non-specifically cleaved by the nuclease. Here we develop a Self-Limiting Cas9 circuit for Enhanced Safety and specificity (SLiCES) which consists of an expression unit for Streptococcus pyogenes Cas9 (SpCas9), a self-targeting sgRNA and a second sgRNA targeting a chosen genomic locus. The self-limiting circuit results in increased genome editing specificity by controlling Cas9 levels. For its in vivo utilization, we next integrate SLiCES into a lentiviral delivery system (lentiSLiCES) via circuit inhibition to achieve viral particle production. Upon delivery into target cells, the lentiSLiCES circuit switches on to edit the intended genomic locus while simultaneously stepping up its own neutralization through SpCas9 inactivation. By preserving target cells from residual nuclease activity, our hit and go system increases safety margins for genome editing.

Cystic fibrosis (CF) is an autosomal recessive disease caused by mutations in the CFTR gene. The 3272–26A>G and 3849+10kbC>T CFTR mutations alter the correct splicing of the CFTR gene, generating new acceptor and donor splice sites respectively. Here we develop a genome editing approach to permanently correct these genetic defects, using a single crRNA and the Acidaminococcus sp. BV3L6, AsCas12a. This genetic repair strategy is highly precise, showing very strong discrimination between the wild-type and mutant sequence and a complete absence of detectable off-targets. The efficacy of this gene correction strategy is verified in intestinal organoids and airway epithelial cells derived from CF patients carrying the 3272–26A>G or 3849+10kbC>T mutations, showing efficient repair and complete functional recovery of the CFTR channel. These results demonstrate that allele-specific genome editing with AsCas12a can correct aberrant CFTR splicing mutations, paving the way for a permanent splicing correction in genetic diseases.

Major mutations associated with HIV-I integrase inhibitors (INI) resistance are rare in INI-naive patients. However, polymorphisms at positions that may influence the genetic barrier and/or drive the selection of specific INI resistance pathways are common in HIV non-B subtypes. The aim was to evaluate the presence of natural polymorphisms and/or INI resistance mutations in HIV-1 non-B subtype samples obtained from INI-naive patients living in rural west Cameroon. Thirty-three HIV-1 non-B samples were obtained from INI-naive African women and, as controls, 15 samples of HIV-1 subtype B were obtained from antiretroviral-naive Italian patients. The integrase gene was amplified and sequenced using Trugene Core Reagents. Several amino acid positions in B and non-B subtypes were found to be polymorphic. Interestingly, two patients infected with the CRF02_AG subtype had the resistance mutations N155H and E157Q/E and 12% of African samples had an amino acid substitution at position 143. Silent mutations leading to a higher increment of genetic barriers were detected at 140 and 151 positions in non B-subtypes. Although most polymorphisms may have little effect on INI susceptibility, the IN gene variations found in the present study should be taken into consideration as they may facilitate or delay the emergence of variants fully resistant to INIs.