CRISPR-Cas9 genome editing induces megabase-scale chromosomal truncations

Hematopoietic stem and progenitors cells gene editing: Beyond blood disorders

Lessons learned from decades-long practice in the transplantation of hematopoietic stem and progenitor cells (HSPCs) to treat severe inherited disorders or cancer, have set the stage for the current ex vivo gene therapies using autologous gene-modified hematopoietic stem and progenitor cells that have treated so far, hundreds of patients with monogenic disorders. With increased knowledge of hematopoietic stem and progenitor cell biology, improved modalities for patient conditioning and with the emergence of new gene editing technologies, a new era of hematopoietic stem and progenitor cell-based gene therapies is poised to emerge. Gene editing has the potential to restore physiological expression of a mutated gene, or to insert a functional gene in a precise locus with reduced off-target activity and toxicity. Advances in patient conditioning has reduced treatment toxicities and may improve the engraftment of gene-modified cells and specific progeny. Thanks to these improvements, new potential treatments of various blood- or immune disorders as well as other inherited diseases will continue to emerge. In the present review, the most recent advances in hematopoietic stem and progenitor cell gene editing will be reported, with a focus on how this approach could be a promising solution to treat non-blood-related inherited disorders and the mechanisms behind the therapeutic actions discussed.

Targeted Integration of Transgenes at the Mouse Gt(ROSA)26Sor Locus

The targeting of transgenic constructs at single copy into neutral genomic loci avoids the unpredictable outcomes associated with conventional random integration approaches. The Gt(ROSA)26Sor locus on chromosome 6 has been used many times for the integration of transgenic constructs and is known to be permissive for transgene expression and disruption of the gene is not associated with a known phenotype. Furthermore, the transcript made from the Gt(ROSA)26Sor locus is ubiquitously expressed and subsequently the locus can be used to drive the ubiquitous expression of transgenes.Here we report a protocol for the generation of targeted transgenic alleles at Gt(ROSA)26Sor, taking as an example a conditional overexpression allele, by PhiC31 integrase/recombinase-mediated cassette exchange of an engineered Gt(ROSA)26Sor locus in mouse embryonic stem cells. The overexpression allele is initially silenced by the presence of a loxP flanked stop sequence but can be strongly activated through the action of Cre recombinase.

Genomic instability drives tumorigenesis and metastasis and its implications for cancer therapy

Genetic instability can be caused by external factors and may also be associated with intracellular damage. At the same time, there is a large body of research investigating the mechanisms by which genetic instability occurs and demonstrating the relationship between genomic stability and tumors. Nowadays, tumorigenesis development is one of the hottest research areas. It is a vital factor affecting tumor treatment. Mechanisms of genomic stability and tumorigenesis development are relatively complex. Researchers have been working on these aspects of research. To explore the research progress of genomic stability and tumorigenesis, development, and treatment, the authors searched PubMed with the keywords "genome instability" "chromosome instability" "DNA damage" "tumor spread" and "cancer treatment". This extracts the information relevant to this study. Results: This review introduces genomic stability, drivers of tumor development, tumor cell characteristics, tumor metastasis, and tumor treatment. Among them, immunotherapy is more important in tumor treatment, which can effectively inhibit tumor metastasis and kill tumor cells. Breakthroughs in tumorigenesis development studies and discoveries in tumor metastasis will provide new therapeutic techniques. New tumor treatment methods can effectively prevent tumor metastasis and improve the cure rate of tumors.

Current status and future prospects in cannabinoid production through in vitro culture and synthetic biology

For centuries, cannabis has been a rich source of fibrous, pharmaceutical, and recreational ingredients. Phytocannabinoids are the most important and well-known class of cannabis-derived secondary metabolites and display a broad range of health-promoting and psychoactive effects. The unique characteristics of phytocannabinoids (e.g., metabolite likeness, multi-target spectrum, and safety profile) have resulted in the development and approval of several cannabis-derived drugs. While most work has focused on the two main cannabinoids produced in the plant, over 150 unique cannabinoids have been identified. To meet the rapidly growing phytocannabinoid demand, particularly many of the minor cannabinoids found in low amounts in planta, biotechnology offers promising alternatives for biosynthesis through in vitro culture and heterologous systems. In recent years, the engineered production of phytocannabinoids has been obtained through synthetic biology both in vitro (cell suspension culture and hairy root culture) and heterologous systems. However, there are still several bottlenecks (e.g., the complexity of the cannabinoid biosynthetic pathway and optimizing the bioprocess), hampering biosynthesis and scaling up the biotechnological process. The current study reviews recent advances related to in vitro culture-mediated cannabinoid production. Additionally, an integrated overview of promising conventional approaches to cannabinoid production is presented. Progress toward cannabinoid production in heterologous systems and possible avenues for avoiding autotoxicity are also reviewed and highlighted. Machine learning is then introduced as a powerful tool to model, and optimize bioprocesses related to cannabinoid production. Finally, regulation and manipulation of the cannabinoid biosynthetic pathway using CRISPR- mediated metabolic engineering is discussed.

Safeguarding genome integrity during gene-editing therapy in a mouse model of age-related macular degeneration

Ensuring genome safety during gene editing is crucial for clinical translation of the high-efficient CRISPR-Cas9 toolbox. Therefore, the undesired events including chromosomal translocations, vector integrations, and large deletions arising during therapeutic gene editing remain to be adequately addressed or tackled in vivo. Here, we apply CRISPR-Cas9TX in comparison to CRISPR-Cas9 to target Vegfa for the treatment of age-related macular degeneration (AMD) disease in a mouse model. AAV delivery of both CRISPR-Cas9 and CRISPR-Cas9TX can efficiently inhibit laser-induced neovascularization. Importantly, Cas9TX almost eliminates chromosomal translocations that occur at a frequency of approximately 1% in Cas9-edited mouse retinal cells. Strikingly, the widely observed AAV integration at the target Vegfa site is also greatly reduced from nearly 50% of edited events to the background level during Cas9TX editing. Our findings reveal that chromosomal structural variations routinely occur during in vivo genome editing and highlight Cas9TX as a superior form of Cas9 for in vivo gene disruption.

CRISPR base editing of cis-regulatory elements enables the perturbation of neurodegeneration-linked genes

CRISPR technology has demonstrated broad utility for controlling target gene expression; however, there remains a need for strategies capable of modulating expression via the precise editing of non-coding regulatory elements. Here, we demonstrate that CRISPR base editors, a class of gene-modifying proteins capable of creating single-base substitutions in DNA, can be used to perturb gene expression via their targeted mutagenesis of cis-acting sequences. Using the promoter region of the human huntingtin (HTT) gene as an initial target, we show that editing of the binding site for the transcription factor NF-κB led to a marked reduction in HTT gene expression in base-edited cell populations. We found that these gene perturbations were persistent and specific, as a transcriptome-wide RNA analysis revealed minimal off-target effects resulting from the action of the base editor protein. We further demonstrate that this base-editing platform could influence gene expression in vivo as its delivery to a mouse model of Huntington's disease led to a potent decrease in HTT mRNA in striatal neurons. Finally, to illustrate the applicability of this concept, we target the amyloid precursor protein, showing that multiplex editing of its promoter region significantly perturbed its expression. These findings demonstrate the potential for base editors to regulate target gene expression.

Visual function restoration in a mouse model of Leber congenital amaurosis via therapeutic base editing

Leber congenital amaurosis (LCA), an inherited retinal degeneration, causes severe visual dysfunction in children and adolescents. In patients with LCA, pathogenic variants, such as RPE65, are evident in specific genes, related to the functions of retinal pigment epithelium and photoreceptors. In contrast to the original Cas9, base editing tools can correct pathogenic substitutions without generation of DNA double-stranded breaks (DSBs). In this study, dual adeno-associated virus (AAV) vectors containing split adenine base editors (ABEs) with trans-splicing intein were prepared for in vivo base editing in retinal degeneration of 12 (rd12) mice, an animal model of LCA, possessing a nonsense mutation of C to T transition in the Rpe65 gene (p.R44X). Subretinal injection of AAV-ABE in retinal pigment epithelial cells of rd12 mice resulted in an A to G transition. The on-target editing was sufficient for recovery of wild-type mRNA, RPE65 protein, and light-induced electrical responses from the retina. Compared with our previous therapeutic editing strategies using Cas9 and prime editing, or with the gene transfer strategy shown in the current study, our results suggest that, considering the editing efficacy and functional recovery, ABEs could be a strong, reliable method for correction of pathogenic variants in the treatment of LCA.

Gene editing strategies to treat lysosomal disorders: The example of mucopolysaccharidoses

Lysosomal storage disorders are a group of progressive multisystemic hereditary diseases with a combined incidence of 1:4,800. Here we review the clinical and molecular characteristics of these diseases, with a special focus on Mucopolysaccharidoses, caused primarily by the lysosomal storage of glycosaminoglycans. Different gene editing techniques can be used to ameliorate their symptoms, using both viral and nonviral delivery methods. Whereas these are still being tested in animal models, early results of phase I/II clinical trials of gene therapy show how this technology may impact the future treatment of these diseases. Hurdles related to specific hard-to-reach organs, such as the central nervous system, heart, joints, and the eye must be tackled. Finally, the regulatory framework necessary to advance into clinical practice is also discussed.

CRISPR/Cas9-based genome editing for the modification of multiple duplications that cause Duchenne muscular dystrophy

With the development of basic research, some genetic-based methods have been found to treat Duchenne muscular dystrophy (DMD) with large deletion mutations and nonsense mutations. Appropriate therapeutic approaches for repairing multiple duplications are limited. We used the CRISPR (clustered regularly interspaced short palindromic repeat)/Cas9 system with patient-derived primary myoblasts to correct multiple duplications of the dystrophin gene. Muscle tissues from a patient carrying duplications of dystrophin were obtained, and tissue-derived primary cells were cultured. Myoblasts were purified with an immunomagnetic sorting system using CD56 microbeads. After transduction by lentivirus with a designed single guide RNA (sgRNA) targeting a duplicated region, myoblasts were allowed to differentiate for 7 days. Copy number variations in the exons of the patient's myotubes were quantified by real-time PCR before and after genetic editing. Western blot analysis was performed to detect the full-length dystrophin protein before and after genetic editing. The ten sequences predicted to be the most likely off-targets were determined by Sanger sequencing. The patient carried duplications of exon 18-25, dystrophin protein expression was completely abrogated. Real-time PCR showed that the copy number of exon 25 in the patient's myotubes was 2.015 ± 0.079 compared with that of the healthy controls. After editing, the copy number of exon 25 in the patient's modified myotubes was 1.308 ± 0.083 compared with that of the healthy controls (P < 0.001). Western blot analysis revealed no expression of the dystrophin protein in the patient's myotubes before editing. After editing, the patient's myotubes expressed the full-length dystrophin protein at a level that was ~6.12% of that in the healthy control samples. Off-target analysis revealed no abnormal editing at the ten sites predicted to be the most likely off-target sites. The excision of multiple duplications by the CRISPR/Cas9 system restored the expression of full-length dystrophin. This study provides proof of evidence for future genome-editing therapy in patients with DMD caused by multiple duplication mutations.

Comparison of CRISPR-Cas9-mediated megabase-scale genome deletion methods in mouse embryonic stem cells

The genome contains large functional units ranging in size from hundreds of kilobases to megabases, such as gene clusters and topologically associating domains. To analyse these large functional units, the technique of deleting the entire functional unit is effective. However, deletion of such large regions is less efficient than conventional genome editing, especially in cultured cells, and a method that can ensure success is anticipated. Here, we compared methods to delete the 2.5-Mb Krüppel-associated box zinc finger protein (KRAB-ZFP) gene cluster in mouse embryonic stem cells using CRISPR-Cas9. Three methods were used: first, deletion by non-homologous end joining (NHEJ); second, homology-directed repair (HDR) using a single-stranded oligodeoxynucleotide (ssODN); and third, HDR employing targeting vectors with a selectable marker and 1-kb homology arms. NHEJ-mediated deletion was achieved in 9% of the transfected cells. Inversion was also detected at similar efficiency. The deletion frequency of NHEJ and HDR was found to be comparable when the ssODN was transfected. Deletion frequency was highest when targeting vectors were introduced, with deletions occurring in 31-63% of the drug-resistant clones. Biallelic deletion was observed when targeting vectors were used. This study will serve as a benchmark for the introduction of large deletions into the genome.

Gene Editing Technologies to Target HBV cccDNA

Hepatitis B virus (HBV) remains a significant cause of mortality and morbidity worldwide, since chronic HBV infection is associated with elevated risk of cirrhosis and hepatocellular carcinoma. Current licensed therapies against HBV efficiently suppress viral replication; however, they do not have significant effects on the intrahepatic covalently closed circular DNA (cccDNA) of the viral minichromosome responsible for viral persistence. Thus, life-long treatment is required to avoid viral rebound. There is a significant need for novel therapies that can reduce, silence or eradicate cccDNA, thus preventing HBV reemergence after treatment withdrawal. In this review, we discuss the latest developments and applications of gene editing and related approaches for directly targeting HBV DNA and, more specifically, cccDNA in infected hepatocytes.

In Vivo Hematopoietic Stem Cell Genome Editing: Perspectives and Limitations

The tremendous evolution of genome-editing tools in the last two decades has provided innovative and effective approaches for gene therapy of congenital and acquired diseases. Zinc-finger nucleases (ZFNs), transcription activator- like effector nucleases (TALENs) and CRISPR-Cas9 have been already applied by ex vivo hematopoietic stem cell (HSC) gene therapy in genetic diseases (i.e., Hemoglobinopathies, Fanconi anemia and hereditary Immunodeficiencies) as well as infectious diseases (i.e., HIV), and the recent development of CRISPR-Cas9-based systems using base and prime editors as well as epigenome editors has provided safer tools for gene therapy. The ex vivo approach for gene addition or editing of HSCs, however, is complex, invasive, technically challenging, costly and not free of toxicity. In vivo gene addition or editing promise to transform gene therapy from a highly sophisticated strategy to a "user-friendly' approach to eventually become a broadly available, highly accessible and potentially affordable treatment modality. In the present review article, based on the lessons gained by more than 3 decades of ex vivo HSC gene therapy, we discuss the concept, the tools, the progress made and the challenges to clinical translation of in vivo HSC gene editing.

Clinical trials and promising preclinical applications of CRISPR/Cas gene editing

Treatment of genetic disorders by genomic manipulation has been the unreachable goal of researchers for many decades. Although our understanding of the genetic basis of genetic diseases has advanced tremendously in the last few decades, the tools developed for genomic editing were not efficient and practical for their use in the clinical setting until now. The recent advancements in the research of Clustered Regularly Interspaced Short Palindromic Repeats (CRISPR) and CRISPR-associated protein (Cas) systems offered an easy and efficient way to edit the genome and accelerated the research on their potential use in the treatment of genetic disorders. In this review, we summarize the clinical trials that evaluate the CRISPR/Cas systems for treating different genetic diseases and highlight promising preclinical research on CRISPR/Cas mediated treatment of a great diversity of genetic disorders. Ultimately, we discuss the future of CRISPR/Cas mediated genome editing in genetic diseases.

CRISPR nuclease off-target activity and mitigation strategies

The discovery of CRISPR has allowed site-specific genomic modification to become a reality and this technology is now being applied in a number of human clinical trials. While this technology has demonstrated impressive efficacy in the clinic to date, there remains the potential for unintended on- and off-target effects of CRISPR nuclease activity. A variety of in silico-based prediction tools and empirically derived experimental methods have been developed to identify the most common unintended effect-small insertions and deletions at genomic sites with homology to the guide RNA. However, large-scale aberrations have recently been reported such as translocations, inversions, deletions, and even chromothripsis. These are more difficult to detect using current workflows indicating a major unmet need in the field. In this review we summarize potential sequencing-based solutions that may be able to detect these large-scale effects even at low frequencies of occurrence. In addition, many of the current clinical trials using CRISPR involve ex vivo isolation of a patient's own stem cells, modification, and re-transplantation. However, there is growing interest in direct, in vivo delivery of genome editing tools. While this strategy has the potential to address disease in cell types that are not amenable to ex vivo manipulation, in vivo editing has only one desired outcome-on-target editing in the cell type of interest. CRISPR activity in unintended cell types (both on- and off-target) is therefore a major safety as well as ethical concern in tissues that could enable germline transmission. In this review, we have summarized the strengths and weaknesses of current editing and delivery tools and potential improvements to off-target and off-tissue CRISPR activity detection. We have also outlined potential mitigation strategies that will ensure that the safety of CRISPR keeps pace with efficacy, a necessary requirement if this technology is to realize its full translational potential.

T Cells Directed against the Metastatic Driver Chondromodulin-1 in Ewing Sarcoma: Comparative Engineering with CRISPR/Cas9 vs. Retroviral Gene Transfer for Adoptive Transfer

Ewing sarcoma (EwS) is a highly malignant sarcoma of bone and soft tissue with early metastatic spread and an age peak in early puberty. The prognosis in advanced stages is still dismal, and the long-term effects of established therapies are severe. Efficacious targeted therapies are urgently needed. Our previous work has provided preliminary safety and efficacy data utilizing T cell receptor (TCR) transgenic T cells, generated by retroviral gene transfer, targeting HLA-restricted peptides on the tumor cell derived from metastatic drivers. Here, we compared T cells engineered with either CRISPR/Cas9 or retroviral gene transfer. Firstly, we confirmed the feasibility of the orthotopic replacement of the endogenous TCR by CRISPR/Cas9 with a TCR targeting our canonical metastatic driver chondromodulin-1 (CHM1). CRISPR/Cas9-engineered T cell products specifically recognized and killed HLA-A*02:01+ EwS cell lines. The efficiency of retroviral transduction was higher compared to CRISPR/Cas9 gene editing. Both engineered T cell products specifically recognized tumor cells and elicited cytotoxicity, with CRISPR/Cas9 engineered T cells providing prolonged cytotoxic activity. In conclusion, T cells engineered with CRISPR/Cas9 could be feasible for immunotherapy of EwS and may have the advantage of more prolonged cytotoxic activity, as compared to T cells engineered with retroviral gene transfer.

In vivo application of base and prime editing to treat inherited retinal diseases

Inherited retinal diseases (IRDs) are vision-threatening retinal disorders caused by pathogenic variants of genes related to visual functions. Genomic analyses in patients with IRDs have revealed pathogenic variants which affect vision. However, treatment options for IRDs are limited to nutritional supplements regardless of genetic variants or gene-targeting approaches based on antisense oligonucleotides and adeno-associated virus vectors limited to targeting few genes. Genome editing, particularly that involving clustered regularly interspaced short palindromic repeat (CRISPR)-Cas9 technologies, can correct pathogenic variants and provide additional treatment opportunities. Recently developed base and prime editing platforms based on CRISPR-Cas9 technologies are promising for therapeutic genome editing because they do not employ double-stranded breaks (DSBs), which are associated with P53 activation, large deletions, and chromosomal translocations. Instead, using attached deaminases and reverse transcriptases, base and prime editing efficiently induces specific base substitutions and intended genetic changes (substitutions, deletions, or insertions), respectively, without DSBs. In this review, we will discuss the recent in vivo application of CRISPR-Cas9 technologies, focusing on base and prime editing, in animal models of IRDs.

Delivering the CRISPR/Cas9 system for engineering gene therapies: Recent cargo and delivery approaches for clinical translation

Clustered Regularly Interspaced Short Palindromic Repeats associated protein 9 (CRISPR/Cas9) has transformed our ability to edit the human genome selectively. This technology has quickly become the most standardized and reproducible gene editing tool available. Catalyzing rapid advances in biomedical research and genetic engineering, the CRISPR/Cas9 system offers great potential to provide diagnostic and therapeutic options for the prevention and treatment of currently incurable single-gene and more complex human diseases. However, significant barriers to the clinical application of CRISPR/Cas9 remain. While in vitro, ex vivo, and in vivo gene editing has been demonstrated extensively in a laboratory setting, the translation to clinical studies is currently limited by shortfalls in the precision, scalability, and efficiency of delivering CRISPR/Cas9-associated reagents to their intended therapeutic targets. To overcome these challenges, recent advancements manipulate both the delivery cargo and vehicles used to transport CRISPR/Cas9 reagents. With the choice of cargo informing the delivery vehicle, both must be optimized for precision and efficiency. This review aims to summarize current bioengineering approaches to applying CRISPR/Cas9 gene editing tools towards the development of emerging cellular therapeutics, focusing on its two main engineerable components: the delivery vehicle and the gene editing cargo it carries. The contemporary barriers to biomedical applications are discussed within the context of key considerations to be made in the optimization of CRISPR/Cas9 for widespread clinical translation.

Comprehensive assessment of miniature CRISPR-Cas12f nucleases for gene disruption

Because of their small size, the recently developed CRISPR-Cas12f nucleases can be effectively packaged into adeno-associated viruses for gene therapy. However, a systematic evaluation of the editing outcomes of CRISPR-Cas12f is lacking. In this study, we apply a high-throughput sequencing method to comprehensively assess the editing efficiency, specificity, and safety of four Cas12f proteins in parallel with that of Cas9 and two Cas12a proteins at multiple genomic sites. Cas12f nucleases achieve robust cleavage at most of the tested sites and mainly produce deletional fragments. In contrast, Cas9 and Cas12a show relatively higher editing efficiency at the vast majority of the tested sites. However, the off-target hotspots identified in the Cas9- and Cas12a-edited cells are negligibly detected in the Cas12f-edited cells. Moreover, compared to Cas9 and Cas12a nucleases, Cas12f nucleases reduce the levels of chromosomal translocations, large deletions, and integrated vectors by 2- to 3-fold. Therefore, our findings confirm the editing capacity of Cas12f and reveal the ability of this nuclease family to preserve genome integrity during genome editing.

Zebrafish Models of Paediatric Brain Tumours

Paediatric brain cancer is the second most common childhood cancer and is the leading cause of cancer-related deaths in children. Despite significant advancements in the treatment modalities and improvements in the 5-year survival rate, it leaves long-term therapy-associated side effects in paediatric patients. Addressing these impairments demands further understanding of the molecularity and heterogeneity of these brain tumours, which can be demonstrated using different animal models of paediatric brain cancer. Here we review the use of zebrafish as potential in vivo models for paediatric brain tumour modelling, as well as catalogue the currently available zebrafish models used to study paediatric brain cancer pathophysiology, and discuss key findings, the unique attributes that these models add, current challenges and therapeutic significance.

Ultra-deep sequencing validates safety of CRISPR/Cas9 genome editing in human hematopoietic stem and progenitor cells

As CRISPR-based therapies enter the clinic, evaluation of safety remains a critical and active area of study. Here, we employ a clinical next generation sequencing (NGS) workflow to achieve high sequencing depth and detect ultra-low frequency variants across exons of genes associated with cancer, all exons, and genome wide. In three separate primary human hematopoietic stem and progenitor cell (HSPC) donors assessed in technical triplicates, we electroporated high-fidelity Cas9 protein targeted to three loci (AAVS1, HBB, and ZFPM2) and harvested genomic DNA at days 4 and 10. Our results demonstrate that clinically relevant delivery of high-fidelity Cas9 to primary HSPCs and ex vivo culture up to 10 days does not introduce or enrich for tumorigenic variants and that even a single SNP in a gRNA spacer sequence is sufficient to eliminate Cas9 off-target activity in primary, repair-competent human HSPCs.

Recursive Editing improves homology-directed repair through retargeting of undesired outcomes

CRISPR-Cas induced homology-directed repair (HDR) enables the installation of a broad range of precise genomic modifications from an exogenous donor template. However, applications of HDR in human cells are often hampered by poor efficiency, stemming from a preference for error-prone end joining pathways that yield short insertions and deletions. Here, we describe Recursive Editing, an HDR improvement strategy that selectively retargets undesired indel outcomes to create additional opportunities to produce the desired HDR allele. We introduce a software tool, named REtarget, that enables the rational design of Recursive Editing experiments. Using REtarget-designed guide RNAs in single editing reactions, Recursive Editing can simultaneously boost HDR efficiencies and reduce undesired indels. We also harness REtarget to generate databases for particularly effective Recursive Editing sites across the genome, to endogenously tag proteins, and to target pathogenic mutations. Recursive Editing constitutes an easy-to-use approach without potentially deleterious cell manipulations and little added experimental burden.

Paired nicking-mediated COL17A1 reframing for junctional epidermolysis bullosa

Junctional epidermolysis bullosa (JEB) is a debilitating hereditary skin disorder caused by mutations in genes encoding laminin-332, type XVII collagen (C17), and integrin-α6β4, which maintain stability between the dermis and epidermis. We designed patient-specific Cas9-nuclease- and -nickase-based targeting strategies for reframing a common homozygous deletion in exon 52 of COL17A1 associated with a lack of full-length C17 expression. Subsequent characterization of protein restoration, indel composition, and divergence of DNA and mRNA outcomes after treatment revealed auspicious efficiency, safety, and precision profiles for paired nicking-based COL17A1 editing. Almost 46% of treated primary JEB keratinocytes expressed reframed C17. Reframed COL17A1 transcripts predominantly featured 25- and 37-nt deletions, accounting for >42% of all edits and encoding C17 protein variants that localized accurately to the cell membrane. Furthermore, corrected cells showed accurate shedding of the extracellular 120-kDa C17 domain and improved adhesion capabilities to laminin-332 compared with untreated JEB cells. Three-dimensional (3D) skin equivalents demonstrated accurate and continuous deposition of C17 within the basal membrane zone between epidermis and dermis. Our findings constitute, for the first time, gene-editing-based correction of a COL17A1 mutation and demonstrate the superiority of proximal paired nicking strategies based on Cas9 D10A nickase over wild-type Cas9-based strategies for gene reframing in a clinical context.

The contribution of DNA repair pathways to genome editing and evolution in filamentous pathogens

DNA double-strand breaks require repair or risk corrupting the language of life. To ensure genome integrity and viability, multiple DNA double-strand break repair pathways function in eukaryotes. Two such repair pathways, canonical non-homologous end joining and homologous recombination, have been extensively studied, while other pathways such as microhomology-mediated end joint and single-strand annealing, once thought to serve as back-ups, now appear to play a fundamental role in DNA repair. Here, we review the molecular details and hierarchy of these four DNA repair pathways, and where possible, a comparison for what is known between animal and fungal models. We address the factors contributing to break repair pathway choice, and aim to explore our understanding and knowledge gaps regarding mechanisms and regulation in filamentous pathogens. We additionally discuss how DNA double-strand break repair pathways influence genome engineering results, including unexpected mutation outcomes. Finally, we review the concept of biased genome evolution in filamentous pathogens, and provide a model, termed Biased Variation, that links DNA double-strand break repair pathways with properties of genome evolution. Despite our extensive knowledge for this universal process, there remain many unanswered questions, for which the answers may improve genome engineering and our understanding of genome evolution.

Cas9/Nickase-induced allelic conversion by homologous chromosome-templated repair in Drosophila somatic cells

Repair of double-strand breaks (DSBs) in somatic cells is primarily accomplished by error-prone nonhomologous end joining and less frequently by precise homology-directed repair preferentially using the sister chromatid as a template. Here, a Drosophila system performs efficient somatic repair of both DSBs and single-strand breaks (SSBs) using intact sequences from the homologous chromosome in a process we refer to as homologous chromosome-templated repair (HTR). Unexpectedly, HTR-mediated allelic conversion at the white locus was more efficient (40 to 65%) in response to SSBs induced by Cas9-derived nickases D10A or H840A than to DSBs induced by fully active Cas9 (20 to 30%). Repair phenotypes elicited by Nickase versus Cas9 differ in both developmental timing (late versus early stages, respectively) and the production of undesired mutagenic events (rare versus frequent). Nickase-mediated HTR represents an efficient and unanticipated mechanism for allelic correction, with far-reaching potential applications in the field of gene editing.

Frequency and mechanisms of LINE-1 retrotransposon insertions at CRISPR/Cas9 sites

CRISPR/Cas9-based genome editing has revolutionized experimental molecular biology and entered the clinical world for targeted gene therapy. Identifying DNA modifications occurring at CRISPR/Cas9 target sites is critical to determine efficiency and safety of editing tools. Here we show that insertions of LINE-1 (L1) retrotransposons can occur frequently at CRISPR/Cas9 editing sites. Together with PolyA-seq and an improved amplicon sequencing, we characterize more than 2500 de novo L1 insertions at multiple CRISPR/Cas9 editing sites in HEK293T, HeLa and U2OS cells. These L1 retrotransposition events exploit CRISPR/Cas9-induced DSB formation and require L1 RT activity. Importantly, de novo L1 insertions are rare during genome editing by prime editors (PE), cytidine or adenine base editors (CBE or ABE), consistent with their reduced DSB formation. These data demonstrate that insertions of retrotransposons might be a potential outcome of CRISPR/Cas9 genome editing and provide further evidence on the safety of different CRISPR-based editing tools.

Cas9-induced large deletions and small indels are controlled in a convergent fashion

Repair of Cas9-induced double-stranded breaks results primarily in formation of small insertions and deletions (indels), but can also cause potentially harmful large deletions. While mechanisms leading to the creation of small indels are relatively well understood, very little is known about the origins of large deletions. Using a library of clonal NGS-validated mouse embryonic stem cells deficient for 32 DNA repair genes, we have shown that large deletion frequency increases in cells impaired for non-homologous end joining and decreases in cells deficient for the central resection gene Nbn and the microhomology-mediated end joining gene Polq. Across deficient clones, increase in large deletion frequency was closely correlated with the increase in the extent of microhomology and the size of small indels, implying a continuity of repair processes across different genomic scales. Furthermore, by targeting diverse genomic sites, we identified examples of repair processes that were highly locus-specific, discovering a role for exonuclease Trex1. Finally, we present evidence that indel sizes increase with the overall efficiency of Cas9 mutagenesis. These findings may have impact on both basic research and clinical use of CRISPR-Cas9, in particular in conjunction with repair pathway modulation.

CRISPR/Cas9 system has revolutionized science and therapy, but DNA damage it causes often goes beyond the desired ’precision editing’. Here, the authors identify general and target specific DNA repair pathways responsible for unwanted mutagenesis.

Understanding on CRISPR/Cas9 mediated cutting-edge approaches for cancer therapeutics

The research focus on CRISPR/Cas9 has gained substantial concentration since the discovery of 'an unusual repeat sequence' reported by Ishino et al. (J Bacteriol 169:5429-5433, 1987) and the journey comprises the recent Nobel Prize award (2020), conferred to Emmanuelle Charpentier and Jennifer Doudna. Cumulatively, the CRISPR has a short, compact, and most discussed success of its application in becoming one of the most versatile and paradigm shifting technologies of Biological Research. Today, the CRISPR/Cas9 genome editing system is almost ubiquitously utilized in many facets of biological research where its tremendous gene manipulation capability has been harnessed to create miracles. From 2012, the CRISPR/Cas 9 system has been showcased in almost 15,000 research articles in the PubMed database, till date. Backed by some strong molecular evidence, the CRISPR system has been utilized in a few clinical trials targeted towards various pathologies. While the area covered by CRISPR is cosmic, this review will focus mostly on the utilization of CRISPR/Cas9 technology in the field of cancer therapy.

Modern therapeutic approaches to liver-related disorders

The liver is a key production and processing site that is essential for health. Liver dysfunction can result in both systemic and liver-specific diseases. To combat these diseases, genetic approaches have been developed that have high liver tropism and are based on gene addition/editing or gene silencing. The gene addition/editing approach has yielded encouraging clinical data on the use of viral vectors in patients with haemophilia, as well as neuromuscular diseases, and has led to trials for liver-related disorders. However, the immune response and the long-term stability of exogenous expression remain important challenges. Gene editing and mRNA therapy have yielded first in-human proof-of-concept therapeutics and vaccines, but the road to the treatment of liver-related disorders remains long. Gene silencing is accomplished primarily via antisense oligonucleotides and small-interfering RNAs (siRNAs). siRNA modification with N-acetyl galactosamine results in hepatocellular-specific targeting and catapulted the liver to the centre of siRNA research. Several siRNA drugs for liver-related disorders have recently been approved, and the pipeline of drugs under investigation is crowded. Loss-of-function mutations might also be treated with enzyme substitution therapy. This review summarises current genetic approaches as well as key enzyme substitution therapies, focusing on recently approved compounds, potential adverse effects, and future challenges. Collectively, these recent advances place the liver at the forefront of precision medicine for metabolic and genetic diseases and are expected to transform the care and treatment of patients with both liver-specific and systemic diseases.

The origin of unwanted editing byproducts in gene editing

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.

Mechanics of CRISPR-Cas12a and engineered variants on λ-DNA

Cas12a is an RNA-guided endonuclease that is emerging as a powerful genome-editing tool. Here, we selected a target site on bacteriophage λ-DNA and used optical tweezers combined with fluorescence to provide mechanistic insight into wild type Cas12a and three engineered variants, where the specific dsDNA and the unspecific ssDNA cleavage are dissociated (M1 and M2) and a third one which nicks the target DNA (M3). At low forces wtCas12a and the variants display two main off-target binding sites, while on stretched dsDNA at higher forces numerous binding events appear driven by the mechanical distortion of the DNA and partial matches to the crRNA. The multiple binding events onto dsDNA at high tension do not lead to cleavage, which is observed on the target site at low forces when the DNA is flexible. In addition, activity assays also show that the preferential off-target sites for this crRNA are not cleaved by wtCas12a, indicating that λ-DNA is only severed at the target site. Our single molecule data indicate that the Cas12a scaffold presents singular mechanical properties, which could be used to generate new endonucleases with biomedical and biotechnological applications.

Efficient Simultaneous Introduction of Premature Stop Codons in Three Tumor Suppressor Genes in PFFs via a Cytosine Base Editor

Base editing is an efficient and precise gene-editing technique, by which a single base can be changed without introducing double-strand breaks, and it is currently widely used in studies of various species. In this study, we used hA3A-BE3-Y130F to simultaneously introduce premature stop codons (TAG, TGA, and TAA) into three tumor suppressor genes, TP53, PTEN, and APC, in large white porcine fetal fibroblasts (PFFs). Among the isolated 290 single-cell colonies, 232 (80%) had premature stop codons in all the three genes. C−to−T conversion was found in 98.6%, 92.8%, and 87.2% of these cell colonies for TP53, PTEN, and APC, respectively. High frequencies of bystander C−to−T edits were observed within the editing window (positions 3−8), and there were nine (3.01%) clones with the designed simultaneous three-gene C−to−T conversion without bystander conversion. C−to−T conversion outside the editing window was found in 9.0%, 14.1%, and 26.2% of the 290 cell colonies for TP53, PTEN, and APC, respectively. Low-frequency C−to−G or C−to−A transversion occurred in APC. The mRNA levels of the three genes showed significant declines in triple-gene-mutant (Tri-Mut) cells as expected. No PTEN and a significantly lower (p < 0.05) APC protein expression were detected in Tri-Mut cells. Interestingly, the premature stop codon introduced into the TP53 gene did not eliminate the expression of its full-length protein in the Tri-Mut cells, suggesting that stop codon read-through occurred. Tri-Mut cells showed a significantly higher (p < 0.05) proliferation rate than WT cells. Furthermore, we identified 1418 differentially expressed genes (DEGs) between the Tri-Mut and WT groups, which were mainly involved in functions such as tumor progression, cell cycle, and DNA repair. This study indicates that hA3A-BE3-Y130F can be a powerful tool to create diverse knockout cell models without double-strand breaks (DSBs), with further possibilities to produce porcine models with various purposes.

Development and Application of CRISPR-Cas Based Tools

Abundant CRISPR-Cas systems in nature provide us with unlimited valuable resources to develop a variety of versatile tools, which are powerful weapons in biological discovery and disease treatment. Here, we systematically review the development of CRISPR-Cas based tools from DNA nuclease to RNA nuclease, from nuclease dependent-tools to nucleic acid recognition dependent-tools. Also, considering the limitations and challenges of current CRISPR-Cas based tools, we discuss the potential directions for development of novel CRISPR toolkits in the future.

Molecular Therapies for Myotonic Dystrophy Type 1: From Small Drugs to Gene Editing

Myotonic dystrophy type 1 (DM1) is the most common muscular dystrophy affecting many different body tissues, predominantly skeletal and cardiac muscles and the central nervous system. The expansion of CTG repeats in the DM1 protein-kinase (DMPK) gene is the genetic cause of the disease. The pathogenetic mechanisms are mainly mediated by the production of a toxic expanded CUG transcript from the DMPK gene. With the availability of new knowledge, disease models, and technical tools, much progress has been made in the discovery of altered pathways and in the potential of therapeutic intervention, making the path to the clinic a closer reality. In this review, we describe and discuss the molecular therapeutic strategies for DM1, which are designed to directly target the CTG genomic tract, the expanded CUG transcript or downstream signaling molecules.

CRISPRi for specific inhibition of miRNA clusters and miRNAs with high sequence homology

miRNAs form a class of noncoding RNAs, involved in post-transcriptional regulation of gene expression, broadly studied for their involvement in physiological and pathological context. Inhibition of mature miRNA transcripts, commonly used in miRNA loss-of-function experiments, may not be specific in case of miRNAs with high sequence homology, e.g. miRNAs from the same seed family. Phenotypic effects of miRNA repression might be biased by the repression of highly similar miRNAs. Another challenge is simultaneous inhibition of multiple miRNAs encoded within policistronic clusters, potentially co-regulating common biological processes. To elucidate roles of miRNA clusters and miRNAs with high sequence homology, it is of key importance to selectively repress only the miRNAs of interest. Targeting miRNAs on genomic level with CRISPR/dCas9-based methods is an attractive alternative to blocking mature miRNAs. Yet, so far no clear guidelines on the design of CRISPR inhibition (CRISPRi) experiments, specifically for miRNA repression, have been proposed. To address this need, here we propose a strategy for effective inhibition of miRNAs and miRNA clusters using CRISPRi. We provide clues on how to approach the challenges in using CRISPR/dCas in miRNA studies, which include prediction of miRNA transcription start sites (TSSs) and the design of single guide RNAs (sgRNAs). The strategy implements three TSS prediction online tools, dedicated specifically for miRNAs: miRStart, FANTOM 5 miRNA atlas, DIANA-miRGen, and CRISPOR tool for sgRNAs design; it includes testing and selection of optimal sgRNAs. We demonstrate that compared to siRNA/shRNA-based miRNA silencing, CRISPRi improves the repression specificity for miRNAs with highly similar sequence and contribute to higher uniformity of the effects of silencing the whole miRNA clusters. This strategy may be adapted for CRISPR-mediated activation (CRISPRa) of miRNA expression.

A kinetic model predicts SpCas9 activity, improves off-target classification, and reveals the physical basis of targeting fidelity

The S. pyogenes (Sp) Cas9 endonuclease is an important gene-editing tool. SpCas9 is directed to target sites based on complementarity to a complexed single-guide RNA (sgRNA). However, SpCas9-sgRNA also binds and cleaves genomic off-targets with only partial complementarity. To date, we lack the ability to predict cleavage and binding activity quantitatively, and rely on binary classification schemes to identify strong off-targets. We report a quantitative kinetic model that captures the SpCas9-mediated strand-replacement reaction in free-energy terms. The model predicts binding and cleavage activity as a function of time, target, and experimental conditions. Trained and validated on high-throughput bulk-biochemical data, our model predicts the intermediate R-loop state recently observed in single-molecule experiments, as well as the associated conversion rates. Finally, we show that our quantitative activity predictor can be reduced to a binary off-target classifier that outperforms the established state-of-the-art. Our approach is extensible, and can characterize any CRISPR-Cas nuclease - benchmarking natural and future high-fidelity variants against SpCas9; elucidating determinants of CRISPR fidelity; and revealing pathways to increased specificity and efficiency in engineered systems.

Cas9 exo-endonuclease eliminates chromosomal translocations during genome editing

The mechanism underlying unwanted structural variations induced by CRISPR-Cas9 remains poorly understood, and no effective strategy is available to inhibit the generation of these byproducts. Here we find that the generation of a high level of translocations is dependent on repeated cleavage at the Cas9-targeting sites. Therefore, we employ a strategy in which Cas9 is fused with optimized TREX2 to generate Cas9TX, a Cas9 exo-endonuclease, which prevents perfect DNA repair and thereby avoids repeated cleavage. In comparison with CRISPR-Cas9, CRISPR-Cas9TX greatly suppressed translocation levels and enhanced the editing efficiency of single-site editing. The number of large deletions associated with Cas9TX was also reduced to very low level. The application of CRISPR-Cas9TX for multiplex gene editing in chimeric antigen receptor T cells nearly eliminated deleterious chromosomal translocations. We report the mechanism underlying translocations induced by Cas9, and propose a general strategy for reducing chromosomal abnormalities induced by CRISPR-RNA-guided endonucleases.

Time-controlled and muscle-specific CRISPR/Cas9-mediated deletion of CTG-repeat expansion in the DMPK gene

CRISPR/Cas9-mediated therapeutic gene editing is a promising technology for durable treatment of incurable monogenic diseases such as myotonic dystrophies. Gene-editing approaches have been recently applied to in vitro and in vivo models of myotonic dystrophy type 1 (DM1) to delete the pathogenic CTG-repeat expansion located in the 3′ untranslated region of the DMPK gene. In DM1-patient-derived cells removal of the expanded repeats induced beneficial effects on major hallmarks of the disease with reduction in DMPK transcript-containing ribonuclear foci and reversal of aberrant splicing patterns. Here, we set out to excise the triplet expansion in a time-restricted and cell-specific fashion to minimize the potential occurrence of unintended events in off-target genomic loci and select for the target cell type. To this aim, we employed either a ubiquitous promoter-driven or a muscle-specific promoter-driven Cas9 nuclease and tetracycline repressor-based guide RNAs. A dual-vector approach was used to deliver the CRISPR/Cas9 components into DM1 patient-derived cells and in skeletal muscle of a DM1 mouse model. In this way, we obtained efficient and inducible gene editing both in proliferating cells and differentiated post-mitotic myocytes in vitro as well as in skeletal muscle tissue in vivo.

Current developments in gene therapy for epidermolysis bullosa

INTRODUCTION

The genodermatosis epidermolysis bullosa (EB) is a monogenetic disease, characterized by severe blister formation on the skin and mucous membranes upon minimal mechanical trauma. Causes for the disease are mutations in genes encoding proteins that are essential for skin integrity. In EB, one of these proteins is either functionally impaired or completely absent. Therefore, the development and improvement of DNA and RNA-based therapeutic approaches for this severe blistering skin disease is mandatory to achieve a treatment option for the patients.

AREAS COVERED

Currently, there are several forms of DNA/RNA therapies potentially feasible for EB. Whereas some of them are still at the preclinical stage, others are clinically advanced and have already been applied to patients. In particular, this is the case for a cDNA replacement approach successfully applied for a small number of patients with junctional EB.

EXPERT OPINION

The heterogeneity of EB justifies the development of therapeutic options with distinct modes of action at a DNA or RNA level. Besides, splicing-modulating therapies, based on RNA trans-splicing or short antisense oligonucleotides, especially designer nucleases, have steadily improved in efficiency and safety and thus likely represent the most promising gene therapy tool in the near future.

Engineered pegRNAs improve prime editing efficiency

Prime editing enables the installation of virtually any combination of point mutations, small insertions or small deletions in the DNA of living cells. A prime editing guide RNA (pegRNA) directs the prime editor protein to the targeted locus and also encodes the desired edit. Here we show that degradation of the 3' region of the pegRNA that contains the reverse transcriptase template and the primer binding site can poison the activity of prime editing systems, impeding editing efficiency. We incorporated structured RNA motifs to the 3' terminus of pegRNAs that enhance their stability and prevent degradation of the 3' extension. The resulting engineered pegRNAs (epegRNAs) improve prime editing efficiency 3-4-fold in HeLa, U2OS and K562 cells and in primary human fibroblasts without increasing off-target editing activity. We optimized the choice of 3' structural motif and developed pegLIT, a computational tool to identify non-interfering nucleotide linkers between pegRNAs and 3' motifs. Finally, we showed that epegRNAs enhance the efficiency of the installation or correction of disease-relevant mutations.

Current Status of Genetically Modified Pigs That Are Resistant to Virus Infection

Pigs play an important role in agriculture and biomedicine. The globally developing swine industry must address the challenges presented by swine-origin viruses, including ASFV (African swine fever virus), PRRSV (porcine reproductive and respiratory syndrome virus), PEDV (porcine epidemic diarrhea virus), PRV (pseudorabies virus), CSFV (classical swine fever virus), TGEV (transmissible gastroenteritis virus), et al. Despite sustained efforts by many government authorities, these viruses are still widespread. Currently, gene-editing technology has been successfully used to generate antiviral pigs, which offers the possibility for increasing animal disease tolerance and improving animal economic traits in the future. Here, we summarized the current advance in knowledge regarding the host factors in virus infection and the current status of genetically modified pigs that are resistant to virus infection in the world. There has not been any report on PEDV-resistant pigs, ASFV-resistant pigs, and PRV-resistant pigs owing to the poor understanding of the key host factors in virus infection. Furthermore, we summarized the remaining problems in producing virus-resistant pigs, and proposed several potential methods to solve them. Using genome-wide CRISPR/Cas9 library screening to explore the key host receptors in virus infection may be a feasible method. At the same time, exploring the key amino acids of host factors in virus infection with library screening based on ABEs and CBEs (Bes) may provide creative insight into producing antiviral pigs in the future.

Considerations and practical implications of performing a phenotypic CRISPR/Cas survival screen

Genome-wide screens that have viability as a readout have been instrumental to identify essential genes. The development of gene knockout screens with the use of CRISPR-Cas has provided a more sensitive method to identify these genes. Here, we performed an exhaustive genome-wide CRISPR/Cas9 phenotypic rescue screen to identify modulators of cytotoxicity induced by the pioneer transcription factor, DUX4. Misexpression of DUX4 due to a failure in epigenetic repressive mechanisms underlies facioscapulohumeral muscular dystrophy (FHSD), a complex muscle disorder that thus far remains untreatable. As the name implies, FSHD generally starts in the muscles of the face and shoulder girdle. Our CRISPR/Cas9 screen revealed no key effectors other than DUX4 itself that could modulate DUX4 cytotoxicity, suggesting that treatment efforts in FSHD should be directed towards direct modulation of DUX4 itself. Our screen did however reveal some rare and unexpected genomic events, that had an important impact on the interpretation of our data. Our findings may provide important considerations for planning future CRISPR/Cas9 phenotypic survival screens.

Identification of novel HPFH-like mutations by CRISPR base editing that elevate the expression of fetal hemoglobin

Naturally occurring point mutations in the HBG promoter switch hemoglobin synthesis from defective adult beta-globin to fetal gamma-globin in sickle cell patients with hereditary persistence of fetal hemoglobin (HPFH) and ameliorate the clinical severity. Inspired by this natural phenomenon, we tiled the highly homologous HBG proximal promoters using adenine and cytosine base editors that avoid the generation of large deletions and identified novel regulatory regions including a cluster at the –123 region. Base editing at –123 and –124 bp of HBG promoter induced fetal hemoglobin (HbF) to a higher level than disruption of well-known BCL11A binding site in erythroblasts derived from human CD34+ hematopoietic stem and progenitor cells (HSPC). We further demonstrated in vitro that the introduction of –123T > C and –124T > C HPFH-like mutations drives gamma-globin expression by creating a de novo binding site for KLF1. Overall, our findings shed light on so far unknown regulatory elements within the HBG promoter and identified additional targets for therapeutic upregulation of fetal hemoglobin.

CRISPR-Cas9 induces large structural variants at on-target and off-target sites in vivo that segregate across generations

CRISPR-Cas9 genome editing has potential to cure diseases without current treatments, but therapies must be safe. Here we show that CRISPR-Cas9 editing can introduce unintended mutations in vivo, which are passed on to the next generation. By editing fertilized zebrafish eggs using four guide RNAs selected for off-target activity in vitro, followed by long-read sequencing of DNA from >1100 larvae, juvenile and adult fish across two generations, we find that structural variants (SVs), i.e., insertions and deletions ≥50 bp, represent 6% of editing outcomes in founder larvae. These SVs occur both at on-target and off-target sites. Our results also illustrate that adult founder zebrafish are mosaic in their germ cells, and that 26% of their offspring carries an off-target mutation and 9% an SV. Hence, pre-testing for off-target activity and SVs using patient material is advisable in clinical applications, to reduce the risk of unanticipated effects with potentially large implications.

Sob gene is critical to wing development in Bombyx mori and Tribolium castaneum

The development of insect appendages requires the expression of multiple genes in a strict spatial and temporal order. The odd-skipped family genes are vital transcriptional factors involved in embryonic development. The development and morphogenesis of the insect wing requires multiple transcription factors to regulate the expression of wing patterning genes at the transcriptional level. However, the function of odd-related genes in insect wing morphogenesis and development during postembryonic stages is unclear. We focused on the roles of the sister of odd and bowl (sob) gene, a member of odd-skipped family genes, during the wing morphopoiesis in Bombyx mori using the clustered regularly interspaced palindromic repeats (CRISPR)/CRISPR-associated protein 9 system and in Tribolium castaneum by RNA interference. The results showed that the wings were significantly smaller and degenerated, and wing veins were indistinct in the sob gene loss-of-function group in both B. mori and T. castaneum. Quantitative real-time polymerase chain reaction revealed that the Tcsob gene regulated the expression of wing development genes, such as the cht 7 and the vg gene. The findings suggest the importance of sob gene in insect wing morphology formation during postembryonic stages.

Base editing and prime editing in laboratory animals

Genome editing by programmable RNA-dependent Cas endonucleases has revolutionised the field of genome engineering, achieving targeted genomic change at unprecedented efficiencies with considerable application in laboratory animal research. Despite its ease of use and wide application, there remain concerns about the precision of this technology and a number of unpredictable consequences have been reported, mostly resulting from the DNA double-strand break (DSB) that conventional CRISPR editing induces. In order to improve editing precision, several iterations of the technology been developed over the years. Base editing is one of most successful developments, allowing for single base conversions but without the need for a DSB. Cytosine and adenine base editing are now established as reliable methods to achieve precise genome editing in animal research studies. Both cytosine and adenine base editors have been applied successfully to the editing of zygotes, resulting in the generation of animal models. Similarly, both base editors have achieved precise editing of point mutations in somatic cells, facilitating the development of gene therapy approaches. Despite rapid progress in optimising these tools, base editing can address only a subset of possible base conversions within a relatively narrow window and larger genomic manipulations are not possible. The recent development of prime editing, originally defined as a simple 'search and replace' editing tool, may help address these limitations and could widen the range of genome manipulations possible. Preliminary reports of prime editing in animals are being published, and this new technology may allow significant advancements for laboratory animal research.

CRISPR-based genome editing through the lens of DNA repair

Genome editing technologies operate by inducing site-specific DNA perturbations that are resolved by cellular DNA repair pathways. Products of genome editors include DNA breaks generated by CRISPR-associated nucleases, base modifications induced by base editors, DNA flaps created by prime editors, and integration intermediates formed by site-specific recombinases and transposases associated with CRISPR systems. Here, we discuss the cellular processes that repair CRISPR-generated DNA lesions and describe strategies to obtain desirable genomic changes through modulation of DNA repair pathways. Advances in our understanding of the DNA repair circuitry, in conjunction with the rapid development of innovative genome editing technologies, promise to greatly enhance our ability to improve food production, combat environmental pollution, develop cell-based therapies, and cure genetic and infectious diseases.