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Cas13d Is a Compact RNA-Targeting Type VI CRISPR Effector Positively Modulated by a WYL-Domain-Containing Accessory Protein.
Bacterial class 2 CRISPR-Cas systems utilize a single RNA-guided protein effector to mitigate viral infection. We aggregated genomic data from multiple sources and constructed an expanded database of predicted class 2 CRISPR-Cas systems. A search for novel RNA-targeting systems identified subtype VI-D, encoding dual HEPN domain-containing Cas13d effectors and putative WYL-domain-containing accessory proteins (WYL1 and WYL-b1 through WYL-b5). The median size of Cas13d proteins is 190 to 300 aa smaller than that of Cas13a-Cas13c. Despite their small size, Cas13d orthologs from Eubacterium siraeum (Es) and Ruminococcus sp. (Rsp) are active in both CRISPR RNA processing and targeting, as well as collateral RNA cleavage, with no target-flanking sequence requirements. The RspWYL1 protein stimulates RNA cleavage by both EsCas13d and RspCas13d, demonstrating a common regulatory mechanism for divergent Cas13d orthologs. The small size, minimal targeting constraints, and modular regulation of Cas13d effectors further expands the CRISPR toolkit for RNA manipulation and detection.
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Rapid modelling of cooperating genetic events in cancer through somatic genome editing.
Cancer is a multistep process that involves mutations and other alterations in oncogenes and tumour suppressor genes. Genome sequencing studies have identified a large collection of genetic alterations that occur in human cancers. However, the determination of which mutations are causally related to tumorigenesis remains a major challenge. Here we describe a novel CRISPR/Cas9-based approach for rapid functional investigation of candidate genes in well-established autochthonous mouse models of cancer. Using a Kras(G12D)-driven lung cancer model, we performed functional characterization of a panel of tumour suppressor genes with known loss-of-function alterations in human lung cancer. Cre-dependent somatic activation of oncogenic Kras(G12D) combined with CRISPR/Cas9-mediated genome editing of tumour suppressor genes resulted in lung adenocarcinomas with distinct histopathological and molecular features. This rapid somatic genome engineering approach enables functional characterization of putative cancer genes in the lung and other tissues using autochthonous mouse models. We anticipate that this approach can be used to systematically dissect the complex catalogue of mutations identified in cancer genome sequencing studies.
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Fast but not furious: a streamlined selection method for genome-edited cells.
In the last decade, transcription activator-like effector nucleases and CRISPR-based genome engineering have revolutionized our approach to biology. Because of their high efficiency and ease of use, the development of custom knock-out and knock-in animal or cell models is now within reach for almost every laboratory. Nonetheless, the generation of genetically modified cells often requires a selection step, usually achieved by antibiotics or fluorescent markers. The choice of the selection marker is based on the available laboratory resources, such as cell types, and parameters such as time and cost should also be taken into consideration. Here, we present a new and fast strategy called magnetic-activated genome-edited cell sorting, to select genetically modified cells based on the ability to magnetically sort surface antigens (i.e., tCD19) present in Cas9-positive cells. By using magnetic-activated genome-edited cell sorting, we successfully generated and isolated genetically modified human-induced pluripotent stem cells, primary human fibroblasts, SH-SY5Y neuroblast-like cells, HaCaT and HEK 293T cells. Our strategy expands the genome editing toolbox by offering a fast, cheap, and an easy to use alternative to the available selection methods.
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The democratization of gene editing: Insights from site-specific cleavage and double-strand break repair.
DNA double-strand breaks (DSBs) are dangerous lesions that if not properly repaired can lead to genomic change or cell death. Organisms have developed several pathways and have many factors devoted to repairing DSBs, which broadly occurs by homologous recombination, which relies on an identical or homologous sequence to template repair, or nonhomologous end-joining. Much of our understanding of these repair mechanisms has come from the study of induced DNA cleavage by site-specific endonucleases. In addition to their biological role, these cellular pathways can be co-opted for gene editing to study gene function or for gene therapy or other applications. While the first gene editing experiments were done more than 20 years ago, the recent discovery of RNA-guided endonucleases has simplified approaches developed over the years to make gene editing an approach that is available to the entire biomedical research community. Here, we review DSB repair mechanisms and site-specific cleavage systems that have provided insight into these mechanisms and led to the current gene editing revolution.
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Comparison of CRISPR and Marker-Based Methods for the Engineering of Phage T7.
With the recent rise in interest in using lytic bacteriophages as therapeutic agents, there is an urgent requirement to understand their fundamental biology to enable the engineering of their genomes. Current methods of phage engineering rely on homologous recombination, followed by a system of selection to identify recombinant phages. For bacteriophage T7, the host genes cmk or trxA have been used as a selection mechanism along with both type I and II CRISPR systems to select against wild-type phage and enrich for the desired mutant. Here, we systematically compare all three systems; we show that the use of marker-based selection is the most efficient method and we use this to generate multiple T7 tail fibre mutants. Furthermore, we found the type II CRISPR-Cas system is easier to use and generally more efficient than a type I system in the engineering of phage T7. These results provide a foundation for the future, more efficient engineering of bacteriophage T7.
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Interrogation of Functional miRNA-Target Interactions by CRISPR/Cas9 Genome Engineering.
Post-transcriptional silencing by microRNAs (miRNAs) is a critical constituent of eukaryotic gene regulation. miRNAs are short (~22nt) noncoding RNAs capable of specifically targeting the miRNA-induced-silencing-complex (miRISC) to transcripts bearing a complementary miRNA response element (MRE). Although recent methodological advances have greatly improved our understanding of miRNA biogenesis and the mechanisms by which miRNAs repress their cognate targets, exploring the physiological relevance of direct miRNA-target interactions in vivo has remained an outstanding challenge. Here we describe the experimental protocol underlying a novel approach, which allows direct interrogation of specific miRNA-MRE interactions by CRISPR/Cas9-mediated genome engineering. In this instance, the CRISPR/Cas9 system is first used to catalyze homology-directed replacement of candidate MREs with molecular barcodes at endogenous loci. Subsequently, the effect of MRE mutation on transcript abundance (i.e., MRE activity) can be rapidly evaluated by routine quantitative PCR. This strategy enables functional investigation of a putative miRNA-target pair in a pool of transiently transfected cells, obviating the need for generation of clonal cell lines or transgenic animals. This protocol can be implemented in any cell line in less than 2 weeks, and can readily be scaled up for multiplex studies. To facilitate the conceptual workflow underlying this strategy, we also describe a genome-wide resource for automated design and computational evaluation of CRISPR/Cas9 guide RNAs targeting all predicted MREs in various species (miR-CRISPR).
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High-resolution characterization of gene function using single-cell CRISPR tiling screen.
Identification of novel functional domains and characterization of detailed regulatory mechanisms in cancer-driving genes is critical for advanced cancer therapy. To date, CRISPR gene editing has primarily been applied to defining the role of individual genes. Recently, high-density mutagenesis via CRISPR tiling of gene-coding exons has been demonstrated to identify functional regions in genes. Furthermore, breakthroughs in combining CRISPR library screens with single-cell droplet RNA sequencing (sc-RNAseq) platforms have revealed the capacity to monitor gene expression changes upon genetic perturbations at single-cell resolution. Here, we present "sc-Tiling," which integrates a CRISPR gene-tiling screen with single-cell transcriptomic and protein structural analyses. Distinct from other reported single-cell CRISPR screens focused on observing gene function and gene-to-gene/enhancer-to-gene regulation, sc-Tiling enables the capacity to identify regulatory mechanisms within a gene-coding region that dictate gene activity and therapeutic response.
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CRISPR-Edited Cell Lines: A New Era in Functional Oncology Research.
The use of CRISPR-Cas9 to engineer cancer cell lines has made it possible to precisely examine how cancer cells react to different drugs and therapies. Some of the key improvements are in the use of Mediator Complex Subunit 12 (MED12)-knockout cells to study cell resistance to BRAF inhibitors, CRISPR models of epithelial-mesenchymal transition for breast cancer, and pharmacogenomic analysis in various cancer cell lines. CRISPR is used in immunotherapy to help Chimeric Antigen Receptor T (CAR-T) cells function better by disrupting the immune checkpoints like Programmed Cell Death Protein 1 (PD-1) and Cytotoxic T-lymphocyte- associated protein 4 (CTLA-4) and to adapt T cells to react with various antigens. As a result of these innovations, it is now possible to track how cancers like non-small cell lung cancer (NSCLC) and ovarian cancer evolve, change their epigenetic features, and find strategies to reverse their resistance. Moving forward, integrating AI analytics, single-cell multi-omics, patient-derived organoids, and CRISPR mechanisms will help improve precision oncology and speed up effective treatment planning.
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Optimized CRISPR/Cas9-mediated in vivo genome engineering applicable to monitoring dynamics of endogenous proteins in the mouse neural tissues.
To analyze the expression, localization, and functional dynamics of target proteins in situ, especially in living cells, it is important to develop a convenient, versatile, and efficient method to precisely introduce exogenous genes into the genome, which is applicable for labeling and engineering of the endogenous proteins of interest. By combining the CRISPR/Cas9 genome editing technology with an electroporation technique, we succeeded in creating knock-in alleles, from which GFP (RFP)-tagged endogenous proteins are produced, in neurons and glial cells in vivo in the developing mouse retina and brain. Correct gene targeting was confirmed by single-cell genotyping and Western blot analysis. Several gene loci were successfully targeted with high efficiency. Moreover, we succeeded in engineering the mouse genome to express foreign genes from the endogenous gene loci using a self-cleaving 2A peptide. Our method could be used to monitor the physiological changes in localization of endogenous proteins and expression levels of both mRNA and protein at a single cell resolution. This work discloses a powerful and widely applicable approach for visualization and manipulation of endogenous proteins in neural tissues.
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A CRISPR-SpCas9M-reporting system for efficient and rapid genome editing in Caulobacter crescentus.
As members of the α-proteobacteria group, Caulobacter crescentus and its relatives are wildly studied for their unique asymmetric life cycle and versatile applications in industry, agriculture, and biomedicine. However, genetic manipulation in these bacteria remains challenging, typically requiring time-consuming and labor-intensive procedures. Here, we report a practical CRISPR-SpCas9M-reporting system that overcomes the limitations of SpCas9 expression and CRISPR escape, enabling efficient, markerless, and rapid genome editing in C. crescentus. Two genes encoding for a pair of scaffold proteins were knocked out individually or iteratively, demonstrating their direct involvements in cellular signaling asymmetry. Key components, including the Cas protein, Cas inducer, sgRNA, homologous arms, and reporter, were systematically analyzed and optimized in the system, finally achieving the apparent editing efficiency up to 80% in C. crescentus. Furthermore, we applied the CRISPR-SpCas9M-reporting system to two C. crescentus relatives, Agrobacterium fabrum and Sinorhizobium meliloti, establishing it as an efficient and general editing strategy. We anticipate that this system could be applied to other CRISPR-Cas-recalcitrant organisms, accelerating both basic and applied research in α-proteobacteria.
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