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Highly Efficient Mouse Genome Editing by CRISPR Ribonucleoprotein Electroporation of Zygotes.
The CRISPR/Cas9 system has been employed to efficiently edit the genomes of diverse model organisms. CRISPR-mediated mouse genome editing is typically accomplished by microinjection of Cas9 DNA/RNA and single guide RNA (sgRNA) into zygotes to generate modified animals in one step. However, microinjection is a technically demanding, labor-intensive, and costly procedure with poor embryo viability. Here, we describe a simple and economic electroporation-based strategy to deliver Cas9/sgRNA ribonucleoproteins into mouse zygotes with 100% efficiency for in vivo genome editing. Our methodology, designated as CRISPR RNP Electroporation of Zygotes (CRISPR-EZ), enables highly efficient and high-throughput genome editing in vivo, with a significant improvement in embryo viability compared with microinjection. Using CRISPR-EZ, we generated a variety of editing schemes in mouse embryos, including indel (insertion/deletion) mutations, point mutations, large deletions, and small insertions. In a proof-of-principle experiment, we used CRISPR-EZ to target the tyrosinase (Tyr) gene, achieving 88% bi-allelic editing and 42% homology-directed repair-mediated precise sequence modification in live mice. Taken together, CRISPR-EZ is simple, economic, high throughput, and highly efficient with the potential to replace microinjection for in vivo genome editing in mice and possibly in other mammals.
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Limit on lab-grown human embryos dropped by stem-cell body.
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Cell-Specific Single Viral Vector CRISPR/Cas9 Editing and Genetically Encoded Tool Delivery in the Central and Peripheral Nervous Systems.
CRISPR/Cas9 gene editing represents an exciting avenue to study genes of unknown function and can be combined with genetically encoded tools such as fluorescent proteins, channelrhodopsins, DREADDs, and various biosensors to more deeply probe the function of these genes in different cell types. However, current strategies to also manipulate or visualize edited cells are challenging due to the large size of Cas9 proteins and the limited packaging capacity of adeno-associated viruses (AAVs). To overcome these constraints, we developed an alternative gene editing strategy using a single AAV vector and mouse lines that express Cre-dependent Cas9 to achieve efficient cell-type specific editing across the nervous system. Expressing Cre-dependent Cas9 from a genomic locus affords space to package guide RNAs for gene editing together with Cre-dependent, genetically encoded tools to manipulate, map, or monitor neurons using a single virus. We validated this strategy with three common tools in neuroscience: ChRonos, a channelrhodopsin, for studying synaptic transmission using optogenetics, GCaMP8f for recording Ca2+ transients using photometry, and mCherry for tracing axonal projections. We tested these tools in multiple brain regions and cell types, including GABAergic neurons in the nucleus accumbens, glutamatergic neurons projecting from the ventral pallidum to the lateral habenula, dopaminergic neurons in the ventral tegmental area, and proprioceptive neurons in the periphery. This flexible approach could help identify and test the function of novel genes affecting synaptic transmission, circuit activity, or morphology with a single viral injection.
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CRISPR/Cas9 gene editing clarifies the role of CD33 SNP rs12459419 in gemtuzumab ozogamicin-mediated cytotoxicity.
The single-nucleotide polymorphism (SNP) rs12459419 is located at the intron/exon junction of CD33 exon2. When exon2 is skipped by this CD33 SNP, the full-length CD33 (CD33FL) is converted to a short CD33 isoform (CD33D2). Since gemtuzumab ozogamicin (GO) only recognizes CD33FL, the CD33 SNP may affect the clinical efficacy of GO. To elucidate the significance of CD33 SNP on GO reactivity, we leveraged the CRISPR/Cas9 genome-editing system to create OCI-AML3 cell lines with specifically modified CD33 SNPs. Levels of CD33 D2 mRNA were significantly higher in the T/T clone (p < 0.001), but CD33D2 protein was not detectable in any clones. There was no significant difference in CD33FL mRNA expression across edited clones, and CD33FL protein expression was lowest in T/T clones, followed by T/C and C/C. Cytotoxicity assays revealed that the IC50 of GO was significantly lower in T/C and C/C clones than in the T/T clone (p < 0.001). Our study demonstrated a difference in GO-induced cytotoxicity in CD33 SNP-edited clones, clearly indicating that at least one CD33 SNP allele, rs12459419 C, is important for sensitivity to GO.
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Integrase-Deficient Lentiviral Vector as a Platform for Efficient CRISPR/Cas9-Mediated Gene Editing for Mucopolysaccharidosis IVA.
Mucopolysaccharidosis IVA (MPS IVA) is a lysosomal storage disorder causing systemic skeletal dysplasia due to a deficiency of N-acetyl-galactosamine-6-sulfate sulfatase (GALNS) enzyme activity, leading to the impaired degradation and accumulation of glycosaminoglycans (GAGs), keratan sulfate (KS) and chondroitin-6-sulfate. While treatments such as enzyme replacement therapy (ERT) and hematopoietic stem cell transplantation (HSCT) are available, they have significant limitations regarding efficacy in skeletal tissues and long-term safety, highlighting the need for more effective therapies. We evaluated a novel gene therapy approach using a dual Integrase-deficient lentiviral vector (IDLV) to deliver an expression cassette that includes human GALNS cDNA and Cas9 sgRNA, targeting the upstream region of the mouse Galns initial codon. This approach leverages the endogenous promoter to drive transgene expression. We assessed in vitro transduction, editing, and functional correction in NIH3T3 and MPS IVA mouse fibroblasts. In vivo efficacy was successfully evaluated via the facial vein injection in MPS IVA newborn mice. In vitro, this IDLV platform demonstrated supraphysiological GALNS activity in cell lysate, resulting in the normalization of KS levels. In vivo direct IDLV platform in newborn MPS IVA mice led to sustained plasma GALNS activity, reduced plasma KS, and favorable biodistribution. Partial correction of heart and bone pathology was observed, with no vector toxicity and minimal antibody responses. This dual IDLV-CRISPR/Cas9 approach effectively mediated targeted GALNS knock-in, yielding sustained enzyme activity, reduced KS storage, and partial pathological amelioration in MPS IVA mice. In conclusion, IDLVs represent an efficient, safe platform for delivering the CRISPR/Cas9 gene editing system for MPS IVA.
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Manipulation of the Tyrosinase gene permits improved CRISPR/Cas editing and neural imaging in cichlid fish.
Direct tests of gene function have historically been performed in a limited number of model organisms. The CRISPR/Cas system is species-agnostic, offering the ability to manipulate genes in a range of models, enabling insights into evolution, development, and physiology. Astatotilapia burtoni, a cichlid fish from the rivers and shoreline around Lake Tanganyika, has been extensively studied in the laboratory to understand evolution and the neural control of behavior. Here we develop protocols for the creation of CRISPR-edited cichlids and create a broadly useful mutant line. By manipulating the Tyrosinase gene, which is necessary for eumelanin pigment production, we describe a fast and reliable approach to quantify and optimize gene editing efficiency. Tyrosinase mutants also remove a major obstruction to imaging, enabling visualization of subdermal structures and fluorophores in situ. These protocols will facilitate broad application of CRISPR/Cas9 to studies of cichlids as well as other non-traditional model aquatic species.
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Applications of CRISPR as a potential therapeutic.
Genetic disorders and congenital abnormalities are present in 2-5% of births all over the world and can cause up to 50% of all early childhood deaths. The establishment of sophisticated and controlled techniques for customizing DNA manipulation is significant for the therapeutic role in such disorders and further research on them. One such technique is CRISPR that is significant towards optimizing genome editing and therapies, metabolic fluxes as well as artificial genetic systems. CRISPR-Cas9 is a molecular appliance that is applied in the areas of genetic and protein engineering. The CRISPR-CAS system is an integral element of prokaryotic adaptive immunity that allows prokaryotic cells to identify and kill any foreign DNA. The Gene editing property of CRISPR finds various applications like diagnostics and therapeutics in cancer, neurodegenerative disorders, genetic diseases, blindness, etc. This review discusses applications of CRISPR as a therapeutic in various disorders including several genetic diseases (including sickle cell anemia, blindness, thalassemia, cystic fibrosis, hereditary tyrosinemia type I, duchenne muscular dystrophy, mitochondrial disorders), Cancer, Huntington's disease and viral infections (like HIV, COVID, etc.) along with the prospects concerning them. CRISPR-based therapy is also being researched and defined for COVID-19. The related mechanism of CRISPR has been discussed alongside highlighting challenges involved in therapeutic applications of CRISPR.
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Rapid, Efficient, and Cost-Effective Gene Editing of Enterococcus faecium with CRISPR-Cas12a.
Considered a serious threat by the Centers for Disease Control and Prevention, multidrug-resistant Enterococcus faecium is an increasing cause of hospital-acquired infection. Here, we provide details on a single-plasmid CRISPR-Cas12a system for generating clean deletions and insertions. Single manipulations were carried out in under 2 weeks, with successful deletions/insertions present in >80% of the clones tested. Using this method, we generated three individual clean deletion mutations in the acpH, treA, and lacL genes and inserted codon-optimized unaG, enabling green fluorescent protein (GFP)-like fluorescence under the control of the trehalase operon. The use of in vivo recombination for plasmid construction kept costs to a minimum. IMPORTANCE Enterococcus faecium is increasingly associated with hard-to-treat antibiotic-resistant infections. The ability to generate clean genomic alterations is the first step in generating a complete mechanistic understanding of how E. faecium acquires pathogenic traits and causes disease. Here, we show that CRISPR-Cas12a can be used to quickly (under 2 weeks) and cheaply delete or insert genes into the E. faecium genome. This substantial improvement over current methods should speed up research on this important opportunistic pathogen.
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Highly efficient gene editing via targeted Cas9 insertion into chicken housekeeping gene.
Achieving stable and efficient transgene expression is a key challenge in advancing avian genome engineering. Although viral vector-based and piggyBac-mediated transgenesis have been widely used in chickens, both approaches are prone to epigenetic silencing, leading to inconsistent, tissue-specific, and often diminished expression over time. This variability limits used of transgenes requiring robust and long-term expression across multiple tissues. In mammals, site-specific integration into genomic safe harbor loci, such as Rosa26, has enabled stable and predictable transgene expression without disrupting endogenous gene function; however, such strategy has not been established in birds. In this research, we hypothesized that integrating Cas9 into endogenous housekeeping genes (the ACTB and GAPDH) could achieve efficient gene editing in chickens through stable and ubiquitous transgene expression. Using two different approaches, 3'-targeted gene insertion and gene tagging, we inserted Cas9 and GFP cassettes into defined genomic loci in chicken DF-1 cells. Both approaches exhibited stable expression of transgenes in the cells, and functional assays confirmed that Cas9 showed highly efficient nuclease activity following guide RNA delivery. Additionally, we derived single-cell clones stably expressing Cas9, enabling uniform and reproducible genome editing in downstream applications. Targeted insertion of transgenes into active housekeeping genes as candidate safe harbor loci mitigates the limitations of random integration and promoter silencing, offering a robust platform for consistent transgene expression in poultry biotechnology and genome engineering.
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Exosome-mediated delivery of CRISPR-Cas9: A revolutionary approach to cancer gene editing.
Several molecular strategies based on targeted gene delivery systems have been developed in recent years; however, the CRISPR-Cas9 technology introduced a new era of targeted gene editing, precisely modifying oncogenes, tumor suppressor genes, and other regulatory genes involved in carcinogenesis. However, efficiently and safely delivering CRISPR-Cas9 to cancer cells across the cell membrane and the nucleus is still challenging. Using viral vectors and nanoparticles presents issues of immunogenicity, off-target effects, and low targeting affinity. Naturally, extracellular vesicles called exosomes have garnered the most attention as delivery vehicles in oncology-related CRISPR-Cas9 calls due to their biocompatibility, loading capacity, and inherent targeting features. The following review discusses the current progress in using exosomes to deliver CRISPR-Cas9 components, the approaches to load the CRISPR components into exosomes, and the modification of exosomes to increase stability and tumor-targeted delivery. We discuss the latest strategies in targeting recently accomplished in the exosome field, including modifying the surface of exosomes to enhance their internalization by cancer cells, as well as the measures taken to overcome the impacts of TME on delivery efficiency. Focusing on in vitro and in vivo experimentation, this review shows that exosome-mediated CRISPR-Cas9 can potentially treat cancer types, including pancreatic, lymphoma, and leukemia, for given gene targets. This paper compares exosome-mediated delivery and conventional vectors regarding safety, immune response, and targeting ability. Last but not least, we present the major drawbacks and potential development of the seemingly promising field of exosome engineering in gene editing, with references to CRISPR technologies and applications that may help make the target exosomes therapeutic in oncology.
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