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Protocol for non-invasive delivery of CRISPR RNPs via virus-like particles for mouse model generation.
CRISPR-virus-like particle (VLP)-induced targeted mutagenesis (CRISPR-VIM) enables genome editing in mouse embryos through non-invasive delivery of CRISPR ribonucleoproteins (RNPs) via VLPs, eliminating the need for physical manipulation and specialized expertise. We detail protocols for VLP production, titration, and treatment for diverse genome edits. This protocol is compatible with zygotes and in vitro fertilization (IVF)-derived embryos via simple co-culture, facilitating high-efficiency and heritable mutations with minimized off-target effects, independent of specialized equipment and conducive to reduced animal use. For complete details on the use and execution of this protocol, please refer to Jeong et al.1.
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CRISPR-Cas9 genome editing in human cells occurs via the Fanconi anemia pathway.
CRISPR-Cas genome editing creates targeted DNA double-strand breaks (DSBs) that are processed by cellular repair pathways, including the incorporation of exogenous DNA via single-strand template repair (SSTR). To determine the genetic basis of SSTR in human cells, we developed a coupled inhibition-cutting system capable of interrogating multiple editing outcomes in the context of thousands of individual gene knockdowns. We found that human Cas9-induced SSTR requires the Fanconi anemia (FA) pathway, which is normally implicated in interstrand cross-link repair. The FA pathway does not directly impact error-prone, non-homologous end joining, but instead diverts repair toward SSTR. Furthermore, FANCD2 protein localizes to Cas9-induced DSBs, indicating a direct role in regulating genome editing. Since FA is itself a genetic disease, these data imply that patient genotype and/or transcriptome may impact the effectiveness of gene editing treatments and that treatments biased toward FA repair pathways could have therapeutic value.
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CRISPR/Cas9 Ribonucleoprotein-mediated Precise Gene Editing by Tube Electroporation.
Gene editing nucleases, represented by CRISPR-associated protein 9 (Cas9), are becoming mainstream tools in biomedical research. Successful delivery of CRISPR/Cas9 elements into the target cells by transfection is a prerequisite for efficient gene editing. This protocol demonstrates that tube electroporation (TE) machine-mediated delivery of CRISPR/Cas9 ribonucleoprotein (RNP), along with single-stranded oligodeoxynucleotide (ssODN) donor templates to different types of mammalian cells, leads to robust precise gene editing events. First, TE was applied to deliver CRISPR/Cas9 RNP and ssODNs to induce disease-causing mutations in the interleukin 2 receptor subunit gamma (IL2RG) gene and sepiapterin reductase (SPR) gene in rabbit fibroblast cells. Precise mutation rates of 3.57%-20% were achieved as determined by bacterial TA cloning sequencing. The same strategy was then used in human iPSCs on several clinically relevant genes including epidermal growth factor receptor (EGFR), myosin binding protein C, cardiac (Mybpc3), and hemoglobin subunit beta (HBB). Consistently, highly precise mutation rates were achieved (11.65%-37.92%) as determined by deep sequencing (DeepSeq). The present work demonstrates that tube electroporation of CRISPR/Cas9 RNP represents an efficient transfection protocol for gene editing in mammalian cells.
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Multiplex Genome Editing and Regulation in Bacillus subtilis with CRISPR-MAD7.
With the advent of MAD7, a Cpf1-like nuclease, there has been a renewed focus on the development of CRISPR-based genome engineering tools in recent years. To improve genome engineering methodologies in B. subtilis, we revisited the potential of MAD7 for gene modification and expression interference. A key challenge in these endeavors is the limited transformation efficiency. To overcome this, we developed an efficient transformation protocol using strains overexpressing competence genes. Our results showed that although MAD7 together with a B. subtilis chromosome-targeting gRNA is lethal, enabling robust counterselection, we successfully engineered a strain carrying the MAD7-gRNA machinery in a reversibly inactivated state, marking a significant advance in the field. We demonstrated that both MAD7 and its catalytically inactive variant (dMAD7) can be conditionally regulated by inactivation at elevated temperatures. In addition, the MAD7-gRNA complex is effective for multiplex genome editing, allowing for the simultaneous deletion, mutation, or insertion of up to four loci, and enabling the combination of gene deletion, gene insertion, and point mutations. Furthermore, we established a strategy that achieves the simultaneous removal of MAD7 and the gRNA along with the desired genome edits. Altogether, this comprehensive study underscores the versatility of MAD7 for complex, scarless genome engineering and lays a strong foundation for further advancing genetic manipulation in B. subtilis.
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Generation and validation of versatile inducible CRISPRi embryonic stem cell and mouse model.
Clustered regularly interspaced short palindromic repeat (CRISPR)-CRISPR-associated (Cas) 9 has been widely used far beyond genome editing. Fusions of deactivated Cas9 (dCas9) to transcription effectors enable interrogation of the epigenome and controlling of gene expression. However, the large transgene size of dCas9-fusion hinders its applications especially in somatic tissues. Here, we develop a robust CRISPR interference (CRISPRi) system by transgenic expression of doxycycline (Dox) inducible dCas9-KRAB in mouse embryonic stem cells (iKRAB ESC). After introduction of specific single-guide RNAs (sgRNAs), the induced dCas9-KRAB efficiently maintains gene inactivation, although it modestly down-regulates the expression of active genes. The proper timing of Dox addition during cell differentiation or reprogramming allows us to study or screen spatiotemporally activated promoters or enhancers and thereby the gene functions. Furthermore, taking the ESC for blastocyst injection, we generate an iKRAB knock-in (KI) mouse model that enables the shutdown of gene expression and loss-of-function (LOF) studies ex vivo and in vivo by a simple transduction of gRNAs. Thus, our inducible CRISPRi ESC line and KI mouse provide versatile and convenient platforms for functional interrogation and high-throughput screens of specific genes and potential regulatory elements in the setting of development or diseases.
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Molecular basis for the PAM expansion and fidelity enhancement of an evolved Cas9 nuclease.
Clustered regularly interspaced short palindromic repeats (CRISPR)-Cas systems have been harnessed as powerful genome editing tools in diverse organisms. However, the off-target effects and the protospacer adjacent motif (PAM) compatibility restrict the therapeutic applications of these systems. Recently, a Streptococcus pyogenes Cas9 (SpCas9) variant, xCas9, was evolved to possess both broad PAM compatibility and high DNA fidelity. Through determination of multiple xCas9 structures, which are all in complex with single-guide RNA (sgRNA) and double-stranded DNA containing different PAM sequences (TGG, CGG, TGA, and TGC), we decipher the molecular mechanisms of the PAM expansion and fidelity enhancement of xCas9. xCas9 follows a unique two-mode PAM recognition mechanism. For non-NGG PAM recognition, xCas9 triggers a notable structural rearrangement in the DNA recognition domains and a rotation in the key PAM-interacting residue R1335; such mechanism has not been observed in the wild-type (WT) SpCas9. For NGG PAM recognition, xCas9 applies a strategy similar to WT SpCas9. Moreover, biochemical and cell-based genome editing experiments pinpointed the critical roles of the E1219V mutation for PAM expansion and the R324L, S409I, and M694I mutations for fidelity enhancement. The molecular-level characterizations of the xCas9 nuclease provide critical insights into the mechanisms of the PAM expansion and fidelity enhancement of xCas9 and could further facilitate the engineering of SpCas9 and other Cas9 orthologs.
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Rapid and robust validation of pooled CRISPR knockout screens using CelFi.
Pooled CRISPR screens are vital in the unbiased interrogation of gene function and are instrumental in uncovering therapeutic targets and biological processes. However, follow-up hit validation is critical to confirm observed results. Researchers need a simple and robust approach to rapidly verify putative hits and test resulting observations. Thus, we developed a CRISPR-based method for hit validation that tests the effect of a genetic perturbation on cell fitness. By editing target loci and monitoring the indel profiles over time, we have created a Cellular Fitness (CelFi) assay that can elucidate cellular vulnerabilities and verify hits from pooled CRISPR knockout screens. Unlike traditional cellular fitness assays that evaluate viability over time, the CelFi assay correlates changes in the indel profile at the target gene with a selective growth advantage or disadvantage in individual cells over time. Moreover, the CelFi assay can be utilized to evaluate gene dependencies and test new hypotheses, regardless of variations in single guide RNA optimization, ribonucleoprotein concentration, and gene copy number.
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Functional characterization of SlitPBP3 in Spodoptera litura by CRISPR/Cas9 mediated genome editing.
Functional gene analysis by using genome editing techniques is limited only in few model insects. Here, we reported an efficient and heritable gene mutagenesis analysis in an important lepidopteran pest, Spodoptera litura, using the CRISPR/Cas9 system. By using this system, we successfully obtained the homozygous S. litura strain by targeting the pheromone binding protein 3 gene (SlitPBP3), which allowed us to elucidate the role of this gene in the olfaction of the female sex pheromones. By co-injection of Cas9 mRNA and sgRNA into S. litura eggs, highly efficient chimera mutation in SlitPBP3 loci was detected both in injected eggs (39.1%) and in the resulting individual moths (87.5%). We used the mutant moths as parents to obtain the G1 offspring and the homozygous mutant strain in G2. The function of SlitPBP3 was explored by Electroantennogram (EAG) recordings with a homozygous mutant strain. The result showed that the EAG responses were significantly decreased in mutant males than in control males when treated with the major sex pheromone component (Z9,E11-14:Ac) and a minor component (Z9-14:Ac) at higher dosages. The results demonstrate that s SlitPBP3 gene plays a minor role in the perception of the female sex pheromones. Furthermore, our study provides a useful methodology with the CRISPR/Cas9 system for gene in vivo functional study, particular for lepidopteran species in which the RNAi approach is not efficient.
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CrisprBuildr: an open-source application for CRISPR-mediated genome engineering in Drosophila melanogaster.
CRISPR/Cas9 is a powerful tool for targeted genome editing experiments. Using CRISPR/Cas9, genes can be deleted or modified by inserting specific DNA sequences, encoding for fluorescent proteins, small peptide tags, or other modifications. Such experiments are essential for detailed gene and protein characterization. However, designing and cloning the corresponding constructs can be repetitive, time-consuming, and laborious. To assist users in CRISPR/Cas9-based genome engineering, we developed CrisprBuildr, an open-source, web-based application for designing modifications to their target genes. CrisprBuildr guides users through creating guide RNAs and repair template vectors to generate cloning maps. The application is designed for the Drosophila melanogaster genome but can serve as a template for other available genomes. We also created new tagging vectors using EGFP and mCherry combined with the small peptide SspB-Q73R for use in iLID-based optogenetic experiments.
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Large-scale GMP-compliant CRISPR-Cas9-mediated deletion of the glucocorticoid receptor in multivirus-specific T cells.
Virus-specific T cells have proven highly effective for the treatment of severe and drug-refractory infections after hematopoietic stem cell transplant (HSCT). However, the efficacy of these cells is hindered by the use of glucocorticoids, often given to patients for the management of complications such as graft-versus-host disease. To address this limitation, we have developed a novel strategy for the rapid generation of good manufacturing practice (GMP)-grade glucocorticoid-resistant multivirus-specific T cells (VSTs) using clustered regularly interspaced short palindromic repeats (CRISPR)-CRISPR-associated protein 9 (Cas9) gene-editing technology. We have shown that deleting the nuclear receptor subfamily 3 group C member 1 (NR3C1; the gene encoding for the glucocorticoid receptor) renders VSTs resistant to the lymphocytotoxic effect of glucocorticoids. NR3C1-knockout (KO) VSTs kill their targets and proliferate successfully in the presence of high doses of dexamethasone both in vitro and in vivo. Moreover, we developed a protocol for the rapid generation of GMP-grade NR3C1 KO VSTs with high on-target activity and minimal off-target editing. These genetically engineered VSTs promise to be a novel approach for the treatment of patients with life-threatening viral infections post-HSCT on glucocorticoid therapy.
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