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Various strategies of effector accumulation to improve the efficiency of genome editing and derivative methodologies.
CRISPR-Cas9 is a sophisticated tool in which Cas9/sgRNA complexes bind to the programmed target sequences and induce DNA double-strand breaks (DSBs) enabling highly efficient genome editing. Moreover, when nuclease-inactive Cas9 (dCas9) is employed, its specific DNA-binding activity provides a variety of derivative technologies such as transcriptional activation/repression, epigenome editing, and chromosome visualization. In these derivative technologies, particular effector molecules are fused with dCas9 or recruited to the target site. However, there had been room for improvement, because both genome editing and derivative technologies require not only the DNA-binding tools but also the additional components for their efficient and flexible outcomes. For genome editing, DSB repair molecules and knock-in donor templates need to act at the DSB sites. Derivative technologies also require their various effector domains to be gathered onto the target sites. Recently, many groups have developed and utilized inventive platforms to accumulate these additional components to the target sequence by modifying Cas9 protein and/or sgRNA. Here, we summarize the strategies of CRISPR-based effector accumulation and the improved methodologies using these creative platforms.
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Strategies to produce T-DNA free CRISPRed fruit trees via Agrobacterium tumefaciens stable gene transfer.
Genome editing via CRISPR/Cas9 is a powerful technology, which has been widely applied to improve traits in cereals, vegetables and even fruit trees. For the delivery of CRISPR/Cas9 components into dicotyledonous plants, Agrobacterium tumefaciens mediated gene transfer is still the prevalent method, although editing is often accompanied by the integration of the bacterial T-DNA into the host genome. We assessed two approaches in order to achieve T-DNA excision from the plant genome, minimizing the extent of foreign DNA left behind. The first is based on the Flp/FRT system and the second on Cas9 and synthetic cleavage target sites (CTS) close to T-DNA borders, which are recognized by the sgRNA. Several grapevine and apple lines, transformed with a panel of CRISPR/SpCas9 binary vectors, were regenerated and characterized for T-DNA copy number and for the rate of targeted editing. As detected by an optimized NGS-based sequencing method, trimming at T-DNA borders occurred in 100% of the lines, impairing in most cases the excision. Another observation was the leakage activity of Cas9 which produced pierced and therefore non-functional CTS. Deletions of genomic DNA and presence of filler DNA were also noticed at the junctions between T-DNA and genomic DNA. This study proved that many factors must be considered for designing efficient binary vectors capable of minimizing the presence of exogenous DNA in CRISPRed fruit trees.
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Highly efficient heritable targeted deletions of gene clusters and non-coding regulatory regions in Arabidopsis using CRISPR/Cas9.
Genome editing using CRISPR/Cas9 is considered the best instrument for genome engineering in plants. This methodology is based on the nuclease activity of Cas9 that is guided to specific genome sequences by single guide RNAs (sgRNAs) thus enabling researchers to engineer simple mutations or large chromosomal deletions. Current methodologies for targeted genome editing in plants using CRISPR/Cas9 are however largely inefficient, mostly due to low Cas9 activity, variable sgRNA efficiency and low heritability of genetic lesions. Here, we describe a newly developed strategy to enhance CRISPR/Cas9 efficiency in Arabidopsis thaliana focusing on the design of novel binary vectors (pUbiCAS9-Red and pEciCAS9-Red), the selection of highly efficient sgRNAs, and the use of direct plant regeneration from induced cell cultures. Our work demonstrates that by combining these three independent developments, heritable targeted chromosomal deletions of large gene clusters and intergenic regulatory sequences can be engineered at a high efficiency. Our results demonstrate that this improved CRISPR/Cas9 methodology can provide a fast, efficient and cost-effective tool to engineer targeted heritable chromosomal deletions, which will be instrumental for future high-throughput functional genomics studies in plants.
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Wildtype heterogeneity contributes to clonal variability in genome edited cells.
Genome editing tools such as CRISPR/Cas9 enable the rapid and precise manipulation of genomes. CRISPR-based genome editing has greatly simplified the study of gene function in cell lines, but its widespread use has also highlighted challenges of reproducibility. Phenotypic variability among different knockout clones of the same gene is a common problem confounding the establishment of robust genotype-phenotype correlations. Optimized genome editing protocols to enhance reproducibility include measures to reduce off-target effects. However, even if current state-of-the-art protocols are applied phenotypic variability is frequently observed. Here we identify heterogeneity of wild-type cells as an important and often neglected confounding factor in genome-editing experiments. We demonstrate that isolation of individual wild-type clones from an apparently homogenous stable cell line uncovers significant phenotypic differences between clones. Strikingly, we observe hundreds of differentially regulated transcripts (477 up- and 306 downregulated) when comparing two populations of wild-type cells. Furthermore, we show a variety of cellular and biochemical alterations in different wild-type clones in a range that is commonly interpreted as biologically relevant in genome-edited cells. Heterogeneity of wild-type cells thus contributes to variability in genome-edited cells when these are generated through isolation of clones. We show that the generation of monoclonal isogenic wild-type cells prior to genomic manipulation reduces phenotypic variability. We therefore propose to generate matched isogenic control cells prior to genome editing to increase reproducibility.
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Dissecting ELANE neutropenia pathogenicity by human HSC gene editing.
Severe congenital neutropenia (SCN) is a life-threatening disorder most often caused by dominant mutations of ELANE that interfere with neutrophil maturation. We conducted a pooled CRISPR screen in human hematopoietic stem and progenitor cells (HSPCs) that correlated ELANE mutations with neutrophil maturation potential. Highly efficient gene editing of early exons elicited nonsense-mediated decay (NMD), overcame neutrophil maturation arrest in HSPCs from ELANE-mutant SCN patients, and produced normal hematopoietic engraftment function. Conversely, terminal exon frameshift alleles that mimic SCN-associated mutations escaped NMD, recapitulated neutrophil maturation arrest, and established an animal model of ELANE-mutant SCN. Surprisingly, only -1 frame insertions or deletions (indels) impeded neutrophil maturation, whereas -2 frame late exon indels repressed translation and supported neutrophil maturation. Gene editing of primary HSPCs allowed faithful identification of variant pathogenicity to clarify molecular mechanisms of disease and encourage a universal therapeutic approach to ELANE-mutant neutropenia, returning normal neutrophil production and preserving HSPC function.
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CRISPR-based metabolic pathway engineering.
A highly effective metabolic pathway is the key for an efficient cell factory. However, the engineered homologous or heterologous multi-gene pathway may be unbalanced, inefficient and causing the accumulation of potentially toxic intermediates. Therefore, pathways must be constructed optimally to minimize these negative effects and maximize catalytic efficiency. With the development of CRISPR technology, some of the problems of previous pathway engineering and genome editing techniques were resolved, providing higher efficiency, lower cost, and easily customizable targets. Moreover, CRISPR was demonstrated as robust and effective in various organisms including both prokaryotes and eukaryotes. In recent years, researchers in the field of metabolic engineering and synthetic biology have exploited various CRISPR-based pathway engineering approaches, which are both effective and convenient, as well as valuable from a theoretical standpoint. In this review, we systematically summarize novel pathway engineering techniques and strategies based on CRISPR nucleases system, CRISPR interference (CRISPRi), and CRISPR activation (CRISPRa), including figures and descriptions for easy understanding, with the aim to facilitate their broader application among fellow researchers.
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Advancing fish disease research through CRISPR-Cas genome editing: Recent developments and future perspectives.
CRISPR-Cas genome editing technology has transformed genetic research, by enabling unprecedented precision in modifying DNA sequences across various organisms, including fish. This review explores the significant advancements and potential uses of CRISPR-Cas technology in the study and management of fish diseases, which pose serious challenges to aquaculture and wild fish populations. Fish diseases cause significant economic losses and environmental impacts, therefore effective disease control a top priority. The review highlights the pivotal role of CRISPR-Cas in identifying disease-associated genes, which is critical to comprehending the genetic causes of disease susceptibility and resistance. Some studies have reported key genetic factors that influence disease outcomes, using targeted gene knockouts and modifications to pave the way for the development of disease-resistant fish strains. The creation of such genetically engineered fish holds great promise for enhancing aquaculture sustainability by reducing the reliance on antibiotics and other conventional disease control measures. In addition, CRISPR-Cas has facilitated in-depth studies of pathogen-host interactions, offering new insights into the mechanisms by which pathogens infect and proliferate within their hosts. By manipulating both host and pathogen genes, this technology provides a powerful tool for uncovering the molecular underpinnings of these interactions, leading to the development of more effective treatment strategies. While CRISPR-Cas has shown great promise in fish research, its application remains limited to a few species, primarily model organisms and some freshwater fish. In addition, challenges such as off-target effects, ecological risks, and ethical concerns regarding the release of genetically modified organisms into the environment must be carefully addressed. This review also discusses these challenges and emphasizes the need for robust regulatory frameworks and ongoing research to mitigate risks. Looking forward, the integration of CRISPR-Cas with other emerging technologies, such as multi-omics approaches, promises to further advance our understanding and management of fish diseases. This review concludes by envisioning the future directions of CRISPR-Cas applications in fish health, underscoring its potential to its growing in the field.
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P.F508del editing in cells from cystic fibrosis patients.
Development of genome editing methods created new opportunities for the development of etiology-based therapies of hereditary diseases. Here, we demonstrate that CRISPR/Cas9 can correct p.F508del mutation in the CFTR gene in the CFTE29o- cells and induced pluripotent stem cells (iPSCs) derived from patients with cystic fibrosis (CF). We used several combinations of Cas9, sgRNA and ssODN and measured editing efficiency in the endogenous CFTR gene and in the co-transfected plasmid containing the CFTR locus with the p.F508del mutation. The non-homologous end joining (NHEJ) frequency in the CFTR gene in the CFTE29o- cells varied from 1.25% to 2.54% of alleles. The best homology-directed repair (HDR) frequency in the endogenous CFTR locus was 1.42% of alleles. In iPSCs, the NHEJ frequency in the CFTR gene varied from 5.5% to 12.13% of alleles. The best HDR efficacy was 2.38% of alleles. Our results show that p.F508del mutation editing using CRISPR/Cas9 in CF patient-derived iPSCs is a relatively rare event and subsequent cell selection and cultivation should be carried out.
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Protocol for genomic recombineering in Yersinia ruckeri using CRISPR Cas12a coupled with the λ Red system.
Genomic manipulation of Yersinia ruckeri, a pathogen of salmonid fish species, is essential for understanding bacterial physiology and virulence. Here, we present a protocol for genomic recombineering in Y. ruckeri, a species reluctant to standard genomic engineering, using CRISPR Cas12a coupled with the λ Red system. We describe steps for identifying protospacer guides, preparing repair template plasmids, and electroporating Yersinia cells with Cpf1 and protospacer plasmids with homologous arms. We then detail procedures for genome editing and plasmid curing.
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Generation of Cas9 transgenic zebrafish and their application in establishing an ERV-deficient animal model.
OBJECTIVES: To investigate the effect of endogenous Cas9 on genome editing efficiency in transgenic zebrafish. RESULTS: Here we have constructed a transgenic zebrafish strain that can be screened by pigment deficiency. Compared with the traditional CRISPR injection method, the transgenic zebrafish can improve the efficiency of genome editing significantly. At the same time, we first observed that the phenotype of vertebral malformation in early embryonic development of zebrafish after ZFERV knockout. CONCLUSIONS: The transgenic zebrafish with expressed Cas9, is more efficient in genome editing. And the results of ZFERV knockout indicated that ERV may affect the vertebral development by Notch1/Delta D signal pathway.
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