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A pipeline for rapidly generating genetically engineered mouse models of pancreatic cancer using in vivo CRISPR-Cas9-mediated somatic recombination.
Genetically engineered mouse models (GEMMs) that recapitulate the major genetic drivers in pancreatic ductal adenocarcinoma (PDAC) have provided unprecedented insights into the pathogenesis of this lethal neoplasm. Nonetheless, generating an autochthonous model is an expensive, time consuming and labor intensive process, particularly when tissue specific expression or deletion of compound alleles are involved. In addition, many of the current PDAC GEMMs cause embryonic, pancreas-wide activation or loss of driver alleles, neither of which reflects the cognate human disease scenario. The advent of CRISPR/Cas9 based gene editing can potentially circumvent many of the aforementioned shortcomings of conventional breeding schema, but ensuring the efficiency of gene editing in vivo remains a challenge. Here we have developed a pipeline for generating PDAC GEMMs of complex genotypes with high efficiency using a single "workhorse" mouse strain expressing Cas9 in the adult pancreas under a p48 promoter. Using adeno-associated virus (AAV) mediated delivery of multiplexed guide RNAs (sgRNAs) to the adult murine pancreas of p48-Cre; LSL-Cas9 mice, we confirm our ability to express an oncogenic Kras G12D allele through homology-directed repair (HDR), in conjunction with CRISPR-induced disruption of cooperating alleles (Trp53, Lkb1 and Arid1A). The resulting GEMMs demonstrate a spectrum of precursor lesions (pancreatic intraepithelial neoplasia [PanIN] or Intraductal papillary mucinous neoplasm [IPMN] with eventual progression to PDAC. Next generation sequencing of the resulting murine PDAC confirms HDR of oncogenic KrasG12D allele at the endogenous locus, and insertion deletion ("indel") and frameshift mutations of targeted tumor suppressor alleles. By using a single "workhorse" mouse strain and optimal AAV serotype for in vivo gene editing with combination of driver alleles, we present a facile autochthonous platform for interrogation of the PDAC genome.
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Amount of Cas9 protein introduced into mouse embryos via electroporation affects the genome-editing rate.
Genetically engineered animals can be produced quickly using genome editing technology. A new electroporation technique, technique for animal knockout system by electroporation (TAKE), aids in the production of genome-edited animals by introducing nucleases into intact embryos using electroporation instead of microinjection. It is difficult to confirm nuclease delivery into embryos after electroporation using the conventional TAKE method. We previously reported the successful visualization of fluorescently-labeled tracrRNA in embryos after electroporation Cas9 paired with the crRNA:tracrRNA-ATTO550 duplex. However, the amount of fluorescence signal from labeled tracrRNA in embryos did not correlate with the genome editing rate of the offspring. This study examined the visualization of Cas9 protein in embryos after electroporation and its correlation with the genome editing rate of the offspring using a fluorescent Cas9 fusion protein. The fluorescent Cas9 protein was observed in all embryos that survived following electroporation. We found that the efficiency of Cas9 protein delivery into embryos via electroporation depended on the pulse length. Furthermore, we demonstrated that the amount of fluorescent Cas9 protein detected in the embryos correlated with the genome editing efficiency of the embryos. These data indicate that the TAKE method using fluorescently-labeled nucleases can be used to optimize the delivery conditions and verify nuclease delivery into individual embryos prior to embryo transfer for the efficient production of genome-edited animals.
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Epigenetic editing of marine medaka ( Oryzias melastigma) fgf2 using CRISPR/dCas9-Tet1CD.
CRISPR/dCas9-mediated epigenetic editing offers a versatile approach for transcriptional regulation without introducing DNA strand breaks. Although this strategy has been explored in a limited number of species, its application in aquatic vertebrates remains largely uncharacterized. In this study, ten-eleven translocation methylcytosine dioxygenase 1 ( tet1) was cloned and molecularly characterized in marine medaka ( Oryzias melastigma). Decitabine treatment identified fibroblast growth factor 2 ( fgf2) as a methylation-sensitive gene, with a regulatory CpG island located within its promoter region. Subsequently, a CRISPR/dCas9-Tet1CD activation system was constructed by fusing the catalytic domain of Tet1 (Tet1CD, Ala1352-Thr2034) to dCas9, enabling locus-specific DNA demethylation. Targeting fgf2, this CRISPR/dCas9-Tet1CD system induced efficient and selective demethylation of the CpG island, resulting in a maximal 2.41-fold increase in fgf2 transcript levels. Whole-genome bisulfite sequencing and transcriptomic analysis confirmed high on-target precision with minimal off-target effects. Epigenetic activation of fgf2 further modulated downstream gene networks associated with growth, promoting durable transcriptional enhancement and increased cellular proliferation. Collectively, these results establish a robust and highly specific epigenetic editing platform in marine medaka, providing a powerful tool for functional genomics studies and regulatory element analysis in aquatic models.
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New frontiers in the therapy of primary immunodeficiency: From gene addition to gene editing.
The most severe primary immune deficiency diseases (PIDs) have been successfully treated with allogeneic hematopoietic stem cell transplantation for more than 4 decades. However, such transplantations have the best outcomes when there is a well-matched donor available because immune complications, such as graft-versus-host disease, are greater without a matched sibling donor. Gene therapy has been developed as a method to perform autologous transplantations of a patient's own stem cells that are genetically corrected. Through an iterative bench-to-bedside-and-back process, methods to efficiently add new copies of the relevant gene to hematopoietic stem cells have led to safe and effective treatments for several PIDs, including forms of severe combined immune deficiency, Wiskott-Aldrich syndrome, and chronic granulomatous disease. New methods for gene editing might allow additional PIDs to be treated by gene therapy because they will allow the endogenous gene to be repaired and expressed under its native regulatory elements, which are essential for genes involved in cell processes of signaling, activation, and proliferation. Gene therapy is providing exciting new treatment options for patients with PIDs, and advances are sure to continue.
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Dynamically covalent lipid nanoparticles mediate CRISPR-Cas9 genome editing against choroidal neovascularization in mice.
As an important modality for choroidal neovascularization (CNV) treatment, intravitreal injection of vascular endothelial growth factor A (VEGFA) inhibitors suffers from undesired response rate, low patient compliance, and ocular damage. Here, dynamically covalent lipid nanoparticles (LNPs) were engineered to mediate VEGFA gene editing and CNV treatment by codelivering Cas9 mRNA (mCas9) and single guide RNA (sgRNA) targeting VEGFA (sgVEGFA). A library of lipidoids bearing iminoboronate ester linkage was developed via facile "one-pot" synthesis, and the top-performing lipidoid-A4B3C7 was formulated into LNP-A4B3C7 with the highest mRNA transfection efficiency. Inside the diseased retinal pigment epithelial cells, LNPs were dissociated upon H2O2-triggered lipidoid degradation, facilitating mRNA/sgRNA release to potentiate the gene editing efficiency. In laser-induced CNV mice, mCas9/sgVEGFA@LNP-A4B3C7 after single intravitreal injection led to pronounced VEGFA disruption and CNV area reduction, outperforming the clinical anti-VEGF drug in eliciting sustained therapeutic effect. This study establishes a robust nonviral platform for mRNA delivery and genome editing and renders a promising strategy for CNV treatment.
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The BBSome assembly is spatially controlled by BBS1 and BBS4 in human cells.
Bardet-Biedl syndrome (BBS) is a pleiotropic ciliopathy caused by dysfunction of primary cilia. More than half of BBS patients carry mutations in one of eight genes encoding for subunits of a protein complex, the BBSome, which mediates trafficking of ciliary cargoes. In this study, we elucidated the mechanisms of the BBSome assembly in living cells and how this process is spatially regulated. We generated a large library of human cell lines deficient in a particular BBSome subunit and expressing another subunit tagged with a fluorescent protein. We analyzed these cell lines utilizing biochemical assays, conventional and expansion microscopy, and quantitative fluorescence microscopy techniques: fluorescence recovery after photobleaching and fluorescence correlation spectroscopy. Our data revealed that the BBSome formation is a sequential process. We show that the pre-BBSome is nucleated by BBS4 and assembled at pericentriolar satellites, followed by the translocation of the BBSome into the ciliary base mediated by BBS1. Our results provide a framework for elucidating how BBS-causative mutations interfere with the biogenesis of the BBSome.
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Comparative Analysis of Promoter Activity in Crassostrea gigas Embryos: Implications for Bivalve Gene Editing.
In recent years, CRISPR/Cas9 gene editing technology has emerged as a powerful genetic tool with potential application in aquaculture. Crassostrea gigas, as a valuable species in aquaculture, holds promising potential for genetic enhancement and breeding through gene editing. However, the lack of efficient promoters for driving exogenous gene expression poses a major obstacle in bivalve gene editing. In this study, we isolated the promoter sequences of the β-tub and histone H3.3A genes from C. gigas. DNA expression constructs were generated by linking the promoters with the enhanced green fluorescent protein (EGFP) reporter and compared with the promoter activity of the endogenous EF-1α gene and an exogenous OsHV-1 promoter in C. gigas embryos. All four promoters effectively drive the expression of EGFP during early embryonic development in C. gigas. Among these four promoters, the β-tub promoter is the most potent promoter in driving EGFP expression in C. gigas embryos as early as 4.5 h after fertilization. The OsHV-1 promoter showed similar activity as β-tub promoter and appeared to be more active than the EF-1α and histone H3.3A promoters in C. gigas embryos. Furthermore, we assessed their performance in other three C. gigas relatives (Crassostrea ariakensis, Crassostrea nippona, and Crassostrea sikamea) and similar results were found. Collectively, these data suggest that the β-tub promoter is an effective promoter in directing gene expression in directing gene expression in oyster embryos, thus offering a potential application for gene editing in bivalves.
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Metarhizium anisopliae engineering mediated by a CRISPR/Cas9 recyclable system.
The advent of CRISPR/Cas technology has revolutionized genome editing, offering simplicity, precision, and cost-effectiveness. While its application in biological control fungi has been limited, including the cosmopolitan fungus Metarhizium anisopliae, recent advancements show promise. However, integrating cas9 and selection-marker genes into fungal genomes poses challenges, including reduced efficiency, toxicity, and off-target effects. Besides, marker-free genetic engineering through a CRISPR recyclable system presents a viable solution, enabling efficient mutant generation without compromising fitness and virulence. This study pioneers the construction of marker-free strains of M. anisopliae using a CRISPR/Cas9 recyclable system. Precise deletion of albA and ku70, alongside gfp cassette insertion, confirms the system efficiency. This innovative approach holds significant potential for facilitating in-depth molecular studies, understanding their ecological roles in agricultural systems, and enhancing biocontrol efficacy against insect pests through genetic improvement.
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DNAJC14 gene-edited pigs are resistant to classical pestiviruses.
Infectious diseases remain a major impediment to livestock production, negatively impacting both productivity and welfare. Where key interactions between viruses and host proteins have been identified, it is possible to rationally devise intervention strategies. In vitro studies have identified the host protein DNAJC14 as a core component of the replicative cycle of classical pestiviruses. Outbreaks caused by this group of viruses cause enormous losses in stock farming due to culling and export restrictions. Using CRISPR/Cas9 gene editing, we produced a cohort of pigs with altered DNAJC14. Primary cells from these animals did not support replication of either classical swine fever virus (CSFV) or bovine viral diarrhoea virus (BVDV) in vitro. In vivo challenge with CSFV revealed that the edited pigs displayed complete resistance to infection. This establishes gene editing as an additional strategy that can contribute to the control of classical pestiviruses.
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Empowering chimeric antigen receptor T-cell therapy with CRISPR.
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