The rapid development of CRISPR-Cas genome editing tools has greatly changed the way to conduct research and holds tremendous promise for clinical applications. During genome editing, CRISPR-Cas enzymes induce DNA breaks at the target sites and subsequently the DNA repair pathways are recruited to generate diverse editing outcomes. Besides off-target cleavage, unwanted editing outcomes including chromosomal structural variations and exogenous DNA integrations have recently raised concerns for clinical safety. To eliminate these unwanted editing byproducts, we need to explore the underlying mechanisms for the formation of diverse editing outcomes from the perspective of DNA repair. Here, we describe the involved DNA repair pathways in sealing Cas enzyme-induced DNA double-stranded breaks and discuss the origins and effects of unwanted editing byproducts on genome stability. Furthermore, we propose the potential risk of inhibiting DNA repair pathways to enhance gene editing. The recent combined studies of DNA repair and CRISPR-Cas editing provide a framework for further optimizing genome editing to enhance editing safety.

Efficient and precise gene editing is the gold standard of any reverse genetic study. The recently developed prime editing approach, a modified CRISPR/Cas9 [clustered regularly interspaced palindromic repeats (CRISPR)/CRISPR-associated protein] editing method, has reached the precision goal but its editing rate can be improved. We present an improved methodology that allows for routine prime editing in the model plant Physcomitrium patens, whilst exploring potential new prime editing improvements. Using a standardized protoplast transfection procedure, multiple prime editing guide RNA (pegRNA) structural and prime editor variants were evaluated targeting the APT reporter gene through direct plant selection. Together, enhancements of expression of the prime editor, modifications of the 3' extension of the pegRNA, and the addition of synonymous mutation in the reverse transcriptase template sequence of the pegRNA dramatically improve the editing rate without affecting the quality of the edits. Furthermore, we show that prime editing is amenable to edit a gene of interest through indirect selection, as demonstrated by the generation of a Ppdek10 mutant. Additionally, we determine that a plant retrotransposon reverse transcriptase enables prime editing. Finally, we show for the first time the possibility of performing prime editing with two independently coded peptides.

Gene editing has recently emerged as a promising technology to engineer genetic modifications precisely in the genome to achieve long-term relief from corneal disorders. Recent advances in the molecular biology leading to the development of Clustered Regularly Interspaced Short Palindromic Repeats (CRISPRs) and CRISPR-associated systems, zinc finger nucleases and transcription activator like effector nucleases have ushered in a new era for high throughput in vitro and in vivo genome engineering. Genome editing can be successfully used to decipher complex molecular mechanisms underlying disease pathophysiology, develop innovative next generation gene therapy, stem cell-based regenerative therapy, and personalized medicine for corneal and other ocular diseases. In this review we describe latest developments in the field of genome editing, current challenges, and future prospects for the development of personalized gene-based medicine for corneal diseases. The gene editing approach is expected to revolutionize current diagnostic and treatment practices for curing blindness.

To study the pathophysiology of human diseases, develop innovative treatments, and refine approaches for regenerative medicine require appropriate preclinical models. Pigs share physiologic and anatomic characteristics with humans and are genetically more similar to humans than are mice. Genetically modified pigs are essential where rodent models do not mimic the human disease phenotype. The male germline stem cell or spermatogonial stem cell (SSC) is unique; it is the only cell type in an adult male that divides and contributes genes to future generations, making it an ideal target for genetic modification. Here we report that CRISPR/Cas9 ribonucleoprotein (RNP)-mediated gene editing in porcine spermatogonia that include SSCs is significantly more efficient than previously reported editing with TALENs and allows precise gene editing by homology directed repair (HDR). We also established homology-mediated end joining (HMEJ) as a second approach to targeted gene editing to enable introduction of larger transgenes and/or humanizing parts of the pig genome for disease modeling or regenerative medicine. In summary, the approaches established in the current study result in efficient targeted genome editing in porcine germ cells for precise replication of human disease alleles.

Gene-editing technology is an emerging therapeutic modality for manipulating the eukaryotic genome by using target-sequence-specific engineered nucleases. Because of the exceptional advantages that gene-editing technology offers in facilitating the accurate correction of sequences in a genome, gene editing-based therapy is being aggressively developed as a next-generation therapeutic approach to treat a wide range of diseases. However, strategies for precise engineering and delivery of gene-editing nucleases, including zinc finger nucleases, transcription activator-like effector nuclease, and CRISPR/Cas9 (clustered regularly interspaced short palindromic repeats-associated nuclease Cas9), present major obstacles to the development of gene-editing therapies, as with other gene-targeting therapeutics. Currently, viral and non-viral vectors are being studied for the delivery of these nucleases into cells in the form of DNA, mRNA, or proteins. Clinical trials are already ongoing, and in vivo studies are actively investigating the applicability of CRISPR/Cas9 techniques. However, the concept of correcting the genome poses major concerns from a regulatory perspective, especially in terms of safety. This review addresses current research trends and delivery strategies for gene editing-based therapeutics in non-clinical and clinical settings and considers the associated regulatory issues.

Usher syndrome (USH) is a rare autosomal recessive disease and the most common inherited form of combined visual and hearing impairment. Up to 13 genes are associated with this disorder, with USH2A being the most prevalent, due partially to the recurrence rate of the c.2299delG mutation. Excluding hearing aids or cochlear implants for hearing impairment, there are no medical solutions available to treat USH patients. The repair of specific mutations by gene editing is, therefore, an interesting strategy that can be explored using the CRISPR/Cas9 system. In this study, this method of gene editing is used to target the c.2299delG mutation on fibroblasts from an USH patient carrying the mutation in homozygosis. Successful in vitro mutation repair was demonstrated using locus-specific RNA-Cas9 ribonucleoproteins with subsequent homologous recombination repair induced by an engineered template supply. Effects on predicted off-target sites in the CRISPR-treated cells were discarded after a targeted deep-sequencing screen. The proven effectiveness and specificity of these correction tools, applied to the c.2299delG pathogenic variant of USH2A, indicates that the CRISPR system should be considered to further explore a potential treatment of USH.

The sturgeon (Acipenseriformes) is an important farmed species because of its economical value. However, neither gene transfer nor gene editing techniques have been established in sturgeon for molecular breeding and gene functional study until now. In this study, we accomplished gene transfer and gene editing in sterlet (Acipenser ruthenus), which has the shortest sexual maturation period of sturgeons. The plasmid encoding enhanced green fluorescent protein (EGFP) was transferred into the embryos of sterlet at injection concentration of 100 ng/μL, under which condition high survival rate and gene transfer rate could be achieved. Subsequently, exogenous EGFP was efficiently disrupted by transcription activator-like effector nucleases (TALENs) or clustered regularly interspaced short palindromic repeats (CRISPR)/Cas9 nuclease/guide RNA (gRNA), with injection concentrations of 300 ng/μL TALENs, or 100 ng/μL Cas9 nuclease and 30 ng/μL gRNA, respectively, under which condition high survival rate and gene mutation rate could be achieved. Finally, the endogenous gene no tail in sterlet was successfully mutated by Cas9 nuclease/gRNA. We observed the CRISPR-induced no tail mutation, at a high efficiency with the mutant P0 embryos displaying the expected phenotype of bent spine and twisted tail.

The Cas9 endonuclease can be programmed by guide RNA to introduce sequence-specific breaks in genomic DNA. Thus, Cas9-based approaches present a range of novel options for genome manipulation and precision editing. African trypanosomes are parasites that cause lethal human and animal diseases. They also serve as models for studies on eukaryotic biology, including 'divergent' biology. Genome modification, exploiting the native homologous recombination machinery, has been important for studies on trypanosomes but often requires multiple rounds of transfection using selectable markers that integrate at low efficiency. We report a system for delivering tetracycline inducible Cas9 and guide RNA to Trypanosoma brucei. In these cells, targeted DNA cleavage and gene disruption can be achieved at close to 100% efficiency without further selection. Disruption of aquaglyceroporin (AQP2) or amino acid transporter genes confers resistance to the clinical drugs pentamidine or eflornithine, respectively, providing simple and robust assays for editing efficiency. We also use the new system for homology-directed, precision base editing; a single-stranded oligodeoxyribonucleotide repair template was delivered to introduce a single AQP2 - T791G/L264R mutation in this case. The technology we describe now enables a range of novel programmed genome-editing approaches in T. brucei that would benefit from temporal control, high-efficiency and precision.

Zebrafish have practical features that make them a useful model for higher-throughput tests of gene function using CRISPR/Cas9 editing to create 'knockout' models. In particular, the use of G0 mosaic mutants has potential to increase throughput of functional studies significantly but may suffer from transient effects of introducing Cas9 via microinjection. Further, a large number of computational and empirical tools exist to design CRISPR assays but often produce varied predictions across methods leaving uncertainty in choosing an optimal approach for zebrafish studies.

Gene editing stands for the methods to precisely make changes to a specific nucleic acid sequence. With the recent development of the clustered regularly interspaced short palindromic repeats (CRISPR)/Cas9 system, gene editing has become efficient, convenient and programmable, leading to promising translational studies and clinical trials for both genetic and non-genetic diseases. A major concern in the applications of the CRISPR/Cas9 system is about its off-target effects, namely the deposition of unexpected, unwanted, or even adverse alterations to the genome. To date, many methods have been developed to nominate or detect the off-target sites of CRISPR/Cas9, which laid the basis for the successful upgrades of CRISPR/Cas9 derivatives with enhanced precision. In this review, we summarize these technological advancements and discuss about the current challenges in the management of off-target effects for future gene therapy.

Chimeric editing is often reported in gene editing. To assess how the general chimeric editing is, we created a transgenic tobacco line carrying a marker, beta-glucuronidase gene (gusA), introduced a CRISPR-Cas9 editing vector into the transgenic tobacco line for knocking out gusA, and then investigated the gusA editing efficiencies in T0 and subsequent generations. The editing vector carried a Cas9 gene, which was driven by the cauliflower mosaic virus 35S promoter, and two guide RNAs, gRNA1 and gRNA2, which were driven by Arabidopsis U6 (AtU6) and U3 (AtU3) promoter, respectively. The two gRNAs were designed to knock out a 42-nucleotide fragment of the coding region of gusA. The editing vector was transformed into gusA-containing tobacco leaves using Agrobacterium tumefaciens-mediated transformation and hygromycin selection. Hygromycin-resistant, independent T0 transgenic lines were used to evaluate gusA-editing efficiencies through histochemical GUS assays, polymerase chain reactions (PCR), and next-generation sequencing of PCR amplicons. Profiles of targeted sequences of 94 T0 transgenic lines revealed that these lines were regenerated from non-edited cells where subsequent editing occurred and created chimeric-edited cells in these lines during or after regeneration. Two of them had the target fragment of 42 bp pairs of nucleotides removed. Detail analysis showed that on-target mutations at the AtU6-gRNA1 site and the AtU3-gRNA2 site were found in 4.3% and 77.7% of T0 transgenic lines, respectively. To overcome the issue of extremely low editing efficiencies in T0 lines, we conducted a second round of shoot induction from the chimeric line(s) to enhance the success of obtaining lines with all or most cells edited. The mutation profiles in T0 transgenic lines provide valuable information to understand gene editing in plant cells with constitutively expressed CRISPR-Cas9 and gRNAs.

Recent developments in gene editing technology have enabled scientists to modify DNA sequence by using engineered endonucleases. These gene editing tools are promising candidates for clinical applications, especially for treatment of inherited disorders like sickle cell disease (SCD). SCD is caused by a point mutation in human β-globin gene (HBB). Clinical strategies have demonstrated substantial success, however there is not any permanent cure for SCD available. CRISPR/Cas9 platform uses a single endonuclease and a single guide RNA (gRNA) to induce sequence-specific DNA double strand break (DSB). When this accompanies a repair template, it allows repairing the mutated gene. In this study, it was aimed to target HBB gene via CRISPR/Cas9 genome editing tool to introduce nucleotide alterations for efficient genome editing and correction of point mutations causing SCD in human cell line, by Homology Directed Repair (HDR). We have achieved to induce target specific nucleotide changes on HBB gene in the locus of mutation causing SCD. The effect of on-target activity of bone fide standard gRNA and newly developed longer gRNA were examined. It is observed that longer gRNA has higher affinity to target DNA while having the same performance for targeting and Cas9 induced DSBs. HDR mechanism was triggered by co-delivery of donor DNA repair templates in circular plasmid form. In conclusion, we have suggested methodological pipeline for efficient targeting with higher affinity to target DNA and generating desired modifications on HBB gene.

Myostatin (MSTN) encodes a negative regulator of skeletal muscle mass that might have applications for promoting muscle growth in livestock. In this study, we aimed to test whether targeted MSTN editing, mediated by transcription activator-like effector nucleases (TALENs), is a viable approach to create myostatin-modified goats (Capra hircus).

This review will highlight some of the recent advances in genome engineering with applications for both clinical and basic science investigations of HIV-1.

Cystic fibrosis (CF) is a lethal autosomal recessive disease caused by mutations in the cystic fibrosis transmembrane conductance regulator (CFTR) gene. Nuclease-mediated precise gene editing (PGE) represents a promising therapy for CF, for which an efficient strategy that is free of viral vector, drug selection, and reporter enrichment (VDR free) is desirable. Here we compared different transfection methods (lipofectamine versus electroporation) and formats (plasmid DNA versus ribonucleoprotein) in delivering the CRISPR/Cas9 elements along with single-stranded oligodeoxynucleotides (ssODNs) to clinically relevant cells targeting major CFTR mutation loci. We demonstrate that, among different combinations, electroporation of CRISPR/Cas9 and guide RNA (gRNA) ribonucleoprotein (Cas9 RNP) is the most effective one. By using this VDR-free method, 4.8% to 27.2% efficiencies were achieved in creating dF508, G542X, and G551D mutations in a wild-type induced pluripotent stem cell (iPSC) line. When it is applied to a patient-derived iPSC line carrying the dF508 mutation, a greater than 20% precise correction rate was achieved. As expected, genetic correction leads to the restoration of CFTR function in iPSC-derived proximal lung organoids, as well as in a patient-derived adenocarcinoma cell line CFPAC-1. The present work demonstrates the feasibility of gene editing-based therapeutics toward monogenic diseases such as CF.

The prototypical genetic autoimmune disease is immune dysregulation polyendocrinopathy enteropathy X-linked (IPEX) syndrome, a severe pediatric disease with limited treatment options. IPEX syndrome is caused by mutations in the forkhead box protein 3 (FOXP3) gene, which plays a critical role in immune regulation. As a monogenic disease, IPEX is an ideal candidate for a therapeutic approach in which autologous hematopoietic stem and progenitor (HSPC) cells or T cells are gene edited ex vivo and reinfused. Here, we describe a CRISPR-based gene correction permitting regulated expression of FOXP3 protein. We demonstrate that gene editing preserves HSPC differentiation potential, and that edited regulatory and effector T cells maintain their in vitro phenotype and function. Additionally, we show that this strategy is suitable for IPEX patient cells with diverse mutations. These results demonstrate the feasibility of gene correction, which will be instrumental for the development of therapeutic approaches for other genetic autoimmune diseases.

Range of DNA repair in response to double-strand breaks induced in human preimplantation embryos remains uncertain due to the complexity of analyzing single- or few-cell samples. Sequencing of such minute DNA input requires a whole genome amplification that can introduce artifacts, including coverage nonuniformity, amplification biases, and allelic dropouts at the target site. We show here that, on average, 26.6% of preexisting heterozygous loci in control single blastomere samples appear as homozygous after whole genome amplification indicative of allelic dropouts. To overcome these limitations, we validate on-target modifications seen in gene edited human embryos in embryonic stem cells. We show that, in addition to frequent indel mutations, biallelic double-strand breaks can also produce large deletions at the target site. Moreover, some embryonic stem cells show copy-neutral loss of heterozygosity at the cleavage site which is likely caused by interallelic gene conversion. However, the frequency of loss of heterozygosity in embryonic stem cells is lower than in blastomeres, suggesting that allelic dropouts is a common whole genome amplification outcome limiting genotyping accuracy in human preimplantation embryos.

Genome editing tools such as the clustered regularly interspaced short palindromic repeat (CRISPR)-associated system (Cas) have been widely used to modify genes in model systems including animal zygotes and human cells, and hold tremendous promise for both basic research and clinical applications. To date, a serious knowledge gap remains in our understanding of DNA repair mechanisms in human early embryos, and in the efficiency and potential off-target effects of using technologies such as CRISPR/Cas9 in human pre-implantation embryos. In this report, we used tripronuclear (3PN) zygotes to further investigate CRISPR/Cas9-mediated gene editing in human cells. We found that CRISPR/Cas9 could effectively cleave the endogenous β-globin gene (HBB). However, the efficiency of homologous recombination directed repair (HDR) of HBB was low and the edited embryos were mosaic. Off-target cleavage was also apparent in these 3PN zygotes as revealed by the T7E1 assay and whole-exome sequencing. Furthermore, the endogenous delta-globin gene (HBD), which is homologous to HBB, competed with exogenous donor oligos to act as the repair template, leading to untoward mutations. Our data also indicated that repair of the HBB locus in these embryos occurred preferentially through the non-crossover HDR pathway. Taken together, our work highlights the pressing need to further improve the fidelity and specificity of the CRISPR/Cas9 platform, a prerequisite for any clinical applications of CRSIPR/Cas9-mediated editing.

RNA editing is a post-transcriptional process that can act upon transcripts from mitochondrial, nuclear, and chloroplast genomes. In chloroplasts, single-nucleotide conversions in mRNAs via RNA editing occur at different frequencies across the plant kingdom. These range from several hundred edited sites in some mosses and ferns to lower frequencies in seed plants and the complete lack of RNA editing in the liverwort Marchantia polymorpha. Here, we report the sequence and edited sites of the chloroplast genome from the liverwort Pellia endiviifolia. The type and frequency of chloroplast RNA editing display a pattern highly similar to that in seed plants. Analyses of the C to U conversions and the genomic context in which the editing sites are embedded provide evidence in favor of the hypothesis that chloroplast RNA editing evolved to compensate mutations in the first land plants.

Plant gene editing is typically performed by delivering reagents such as Cas9 and single guide RNAs to explants in culture. Edited cells are then induced to differentiate into whole plants by exposure to various hormones. The creation of edited plants through tissue culture is often inefficient, time-consuming, works for only limited species and genotypes, and causes unintended changes to the genome and epigenome. Here we report two methods to generate gene-edited dicotyledonous plants through de novo meristem induction. Developmental regulators and gene-editing reagents are delivered to somatic cells of whole plants. This induces meristems that produce shoots with targeted DNA modifications, and gene edits are transmitted to the next generation. The de novo induction of gene-edited meristems sidesteps the need for tissue culture and promises to overcome a bottleneck in plant gene editing.