[Application of CRISPR/Cas9 gene editing technology in edible fungi: a review]

The CRISPR/Cas9 gene editing system is a versatile technology for modifying gene, playing a crucial role in the study of functional genes and genetic breeding of plants, animals, fungi, and microorganisms. This review provides a comprehensive analysis of the application of this technology in gene research and genetic breeding of edible fungi. The review covers various aspects, including the delivery and expression strategies of Cas9 and sgRNA, genetic transformation methods, mutant screening, and repair strategies for target sites following DNA double-strand breaks. Additionally, the review summarizes the main challenges and optimization strategies associated with the application of this technology in edible fungi. Lastly, the future application potential of this technology in edible fungi research is discussed, drawing from the authors' personal research background.

Expanding the flexibility of base editing for high-throughput genetic screens in bacteria

Genome-wide screens have become powerful tools for elucidating genotype-to-phenotype relationships in bacteria. Of the varying techniques to achieve knockout and knockdown, CRISPR base editors are emerging as promising options. However, the limited number of available, efficient target sites hampers their use for high-throughput screening. Here, we make multiple advances to enable flexible base editing as part of high-throughput genetic screening in bacteria. We first co-opt the Streptococcus canis Cas9 that exhibits more flexible protospacer-adjacent motif recognition than the traditional Streptococcus pyogenes Cas9. We then expand beyond introducing premature stop codons by mutating start codons. Next, we derive guide design rules by applying machine learning to an essentiality screen conducted in Escherichia coli. Finally, we rescue poorly edited sites by combining base editing with Cas9-induced cleavage of unedited cells, thereby enriching for intended edits. The efficiency of this dual system was validated through a conditional essentiality screen based on growth in minimal media. Overall, expanding the scope of genome-wide knockout screens with base editors could further facilitate the investigation of new gene functions and interactions in bacteria.

CoCas9 is a compact nuclease from the human microbiome for efficient and precise genome editing

The expansion of the CRISPR-Cas toolbox is highly needed to accelerate the development of therapies for genetic diseases. Here, through the interrogation of a massively expanded repository of metagenome-assembled genomes, mostly from human microbiomes, we uncover a large variety (n = 17,173) of type II CRISPR-Cas loci. Among these we identify CoCas9, a strongly active and high-fidelity nuclease with reduced molecular size (1004 amino acids) isolated from an uncultivated Collinsella species. CoCas9 is efficiently co-delivered with its sgRNA through adeno associated viral (AAV) vectors, obtaining efficient in vivo editing in the mouse retina. With this study we uncover a collection of previously uncharacterized Cas9 nucleases, including CoCas9, which enriches the genome editing toolbox.

Modular dual-color BiAD sensors for locus-specific readout of epigenome modifications in single cells

Dynamic changes in the epigenome at defined genomic loci play crucial roles during cellular differentiation and disease development. Here, we developed dual-color bimolecular anchor detector (BiAD) sensors for high-sensitivity readout of locus-specific epigenome modifications by fluorescence microscopy. Our BiAD sensors comprise an sgRNA/dCas9 complex as anchor and double chromatin reader domains as detector modules, both fused to complementary parts of a split IFP2.0 fluorophore, enabling its reconstitution upon binding of both parts in close proximity. In addition, a YPet fluorophore is recruited to the sgRNA to mark the genomic locus of interest. With these dual-color BiAD sensors, we detected H3K9me2/3 and DNA methylation and their dynamic changes upon RNAi or inhibitor treatment with high sensitivity at endogenous genomic regions. Furthermore, we showcased locus-specific H3K36me2/3 readout as well as H3K27me3 and H3K9me2/3 enrichment on the inactive X chromosome, highlighting the broad applicability of our dual-color BiAD sensors for single-cell epigenome studies.

Improving prime editing with an endogenous small RNA-binding protein

Prime editing enables the precise modification of genomes through reverse transcription of template sequences appended to the 3' ends of CRISPR-Cas guide RNAs1. To identify cellular determinants of prime editing, we developed scalable prime editing reporters and performed genome-scale CRISPR-interference screens. From these screens, a single factor emerged as the strongest mediator of prime editing: the small RNA-binding exonuclease protection factor La. Further investigation revealed that La promotes prime editing across approaches (PE2, PE3, PE4 and PE5), edit types (substitutions, insertions and deletions), endogenous loci and cell types but has no consistent effect on genome-editing approaches that rely on standard, unextended guide RNAs. Previous work has shown that La binds polyuridine tracts at the 3' ends of RNA polymerase III transcripts2. We found that La functionally interacts with the 3' ends of polyuridylated prime editing guide RNAs (pegRNAs). Guided by these results, we developed a prime editor protein (PE7) fused to the RNA-binding, N-terminal domain of La. This editor improved prime editing with expressed pegRNAs and engineered pegRNAs (epegRNAs), as well as with synthetic pegRNAs optimized for La binding. Together, our results provide key insights into how prime editing components interact with the cellular environment and suggest general strategies for stabilizing exogenous small RNAs therein.

Base-editing mutagenesis maps alleles to tune human T cell functions

CRISPR-enabled screening is a powerful tool for the discovery of genes that control T cell function and has nominated candidate targets for immunotherapies1-6. However, new approaches are required to probe specific nucleotide sequences within key genes. Systematic mutagenesis in primary human T cells could reveal alleles that tune specific phenotypes. DNA base editors are powerful tools for introducing targeted mutations with high efficiency7,8. Here we develop a large-scale base-editing mutagenesis platform with the goal of pinpointing nucleotides that encode amino acid residues that tune primary human T cell activation responses. We generated a library of around 117,000 single guide RNA molecules targeting base editors to protein-coding sites across 385 genes implicated in T cell function and systematically identified protein domains and specific amino acid residues that regulate T cell activation and cytokine production. We found a broad spectrum of alleles with variants encoding critical residues in proteins including PIK3CD, VAV1, LCP2, PLCG1 and DGKZ, including both gain-of-function and loss-of-function mutations. We validated the functional effects of many alleles and further demonstrated that base-editing hits could positively and negatively tune T cell cytotoxic function. Finally, higher-resolution screening using a base editor with relaxed protospacer-adjacent motif requirements9 (NG versus NGG) revealed specific structural domains and protein-protein interaction sites that can be targeted to tune T cell functions. Base-editing screens in primary immune cells thus provide biochemical insights with the potential to accelerate immunotherapy design.

In Silico Design of gRNA for CRISPR/Cas9-Mediated Gene Knockout

CRISPR/Cas9 stands as a revolutionary and versatile gene editing technology. At its core, the Cas9 DNA endonuclease is guided with precision by a specifically designed single-guide RNA (gRNA). This guidance system facilitates the introduction of double-stranded breaks (DSBs) within the DNA. Subsequent imprecise repairs, mainly through the non-homologous end-joining (NHEJ) pathway, yield insertions or deletions, resulting in frameshift mutations. These mutations are instrumental in achieving the successful knockout of the target gene. In this chapter, we describe all necessary steps to create and design a gRNA for a gene knockout to a target gene before to transfer it to a target plant.

A Comprehensive Protocol for Assembly of Multiple gRNAs into a Direct Vector for Genome Editing in Tomato

The Clustered Regularly Interspaced Short Palindromic Repeats (CRISPR)-Cas 9 (CRISPR-associated protein 9) is a robust DNA-encoded, RNA-mediated sequence-specific nuclease system widely used for genome editing of various plants. Although there are many reports on the assembly of gRNAs and plant transformation, there is no single resource for the complete gene editing methodology in tomato. This chapter provides a comprehensive protocol for designing gRNAs, their assembly into the vector, plant transformation, and final mutant analysis in tomato.

CRISPR/Cas9 Vector Construction for Gene Knockout

This protocol outlines the construction of a plant transformation plasmid to express both the Cas9 nuclease and individual guide RNA (gRNA), facilitating the induction of double-stranded breaks (DSBs) in DNA and subsequent imprecise repair via the non-homologous end-joining (NHEJ) pathway. The gRNA expression cassettes are assembled from three components. First, the Medicago truncatula U6.6 (MtU6) promoter (352 bp) and scaffold (83 bp) sequences are amplified from a pUC-based plasmid. Additionally, a third fragment, corresponding to the target sequence, is synthesized as an oligonucleotide. The three gRNA expression fragments are then loosely assembled in a ligation-free cloning reaction and used as a template for an additional PCR step to amplify a single gRNA expression construct, ready for assembly into the transformation vector. The benefits of this design include cost efficiency, as subsequent cloning reactions only require 59 oligonucleotides and standard cloning reagents. Researchers engaged in CRISPR/Cas9-mediated genome editing in plants will find this protocol a clear and resource-efficient approach to create transformation plasmids for their experiments.

Precision Genome Editing with CRISPR-Cas9

The CRISPR/Cas9 system is a revolutionary technology for genome editing that allows for precise and efficient modifications of DNA sequences. The system is composed of two main components, the Cas9 enzyme and a guide RNA (gRNA). The gRNA is designed to specifically target a desired DNA sequence, while the Cas9 enzyme acts as molecular scissors to cut the DNA at that specific location. The cell then repairs the digested DNA, either through nonhomologous end joining (NHEJ) or homology-directed repair (HDR), resulting in either indels or precise modifications of DNA sequences with broad implications in biotechnology, agriculture, and medicine. This chapter provides a practical approach for utilizing CRISPR/Cas9 in precise genome editing, including identifying the target gene sequence, designing gRNA and protein (Cas9), and delivering the CRISPR components to target cells.

Dual-Luciferase Reporter Assay for Prescreening CRISPR (d)Cas9-Mediated Epigenetic Editing on a Plant Promoter Using Human Cells

Epigenetic editing, also known as EpiEdit, offers an exciting way to control gene expression without altering the DNA sequence. In this study, we evaluate the application of EpiEdit to plant promoters, specifically the MLO (mildew locus o) gene promoter. We use a modified CRISPR-(d)Cas9 system, in which the nuclease-deficient Cas9 (dCas9) is fused to an epigenetic modifier, to experimentally demonstrate the utility of this tool for optimizing epigenetic engineering of a plant promoter prior to in vivo plant epigenome editing. Guide RNAs are used to deliver the dCas9-epigenetic modifier fusion protein to the target gene sequence, where it induces modification of MLO gene expression. We perform preliminary experiments using a plant promoter cloned into the luciferase reporter system, which is transfected into a human system and analyzed using the dual-luciferase reporter assay. The results suggest that this approach may be useful in the early stages of plant epigenome editing, as it can aid in the selection of appropriate modifications to the plant promoter prior to conducting in vivo experiments under plant system conditions. Overall, the results demonstrate the potential of CRISPR (d)Cas9-based EpiEdit for precise and controlled regulation of gene expression.

Fanzor is a eukaryotic programmable RNA-guided endonuclease

RNA-guided systems, which use complementarity between a guide RNA and target nucleic acid sequences for recognition of genetic elements, have a central role in biological processes in both prokaryotes and eukaryotes. For example, the prokaryotic CRISPR-Cas systems provide adaptive immunity for bacteria and archaea against foreign genetic elements. Cas effectors such as Cas9 and Cas12 perform guide-RNA-dependent DNA cleavage1. Although a few eukaryotic RNA-guided systems have been studied, including RNA interference2 and ribosomal RNA modification3, it remains unclear whether eukaryotes have RNA-guided endonucleases. Recently, a new class of prokaryotic RNA-guided systems (termed OMEGA) was reported4,5. The OMEGA effector TnpB is the putative ancestor of Cas12 and has RNA-guided endonuclease activity4,6. TnpB may also be the ancestor of the eukaryotic transposon-encoded Fanzor (Fz) proteins4,7, raising the possibility that eukaryotes are also equipped with CRISPR-Cas or OMEGA-like programmable RNA-guided endonucleases. Here we report the biochemical characterization of Fz, showing that it is an RNA-guided DNA endonuclease. We also show that Fz can be reprogrammed for human genome engineering applications. Finally, we resolve the structure of Spizellomyces punctatus Fz at 2.7 Å using cryogenic electron microscopy, showing the conservation of core regions among Fz, TnpB and Cas12, despite diverse cognate RNA structures. Our results show that Fz is a eukaryotic OMEGA system, demonstrating that RNA-guided endonucleases are present in all three domains of life.

Binding to the conserved and stably folded guide RNA pseudoknot induces Cas12a conformational changes during ribonucleoprotein assembly

Ribonucleoproteins (RNPs) comprise one or more RNA and protein molecules that interact to form a stable complex, which commonly involves conformational changes in the more flexible RNA components. Here, we propose that Cas12a RNP assembly with its cognate CRISPR RNA (crRNA) guide instead proceeds primarily through Cas12a conformational changes during binding to more stable, prefolded crRNA 5' pseudoknot handles. Phylogenetic reconstructions and sequence and structure alignments revealed that the Cas12a proteins are divergent in sequence and structure while the crRNA 5' repeat region, which folds into a pseudoknot and anchors binding to Cas12a, is highly conserved. Molecular dynamics simulations of three Cas12a proteins and their cognate guides revealed substantial flexibility for unbound apo-Cas12a. In contrast, crRNA 5' pseudoknots were predicted to be stable and independently folded. Limited trypsin hydrolysis, differential scanning fluorimetry, thermal denaturation, and CD analyses supported conformational changes of Cas12a during RNP assembly and an independently folded crRNA 5' pseudoknot. This RNP assembly mechanism may be rationalized by evolutionary pressure to conserve CRISPR loci repeat sequence, and therefore guide RNA structure, to maintain function across all phases of the CRISPR defense mechanism.

Cryo-EM structure of the transposon-associated TnpB enzyme

The class 2 type V CRISPR effector Cas12 is thought to have evolved from the IS200/IS605 superfamily of transposon-associated TnpB proteins1. Recent studies have identified TnpB proteins as miniature RNA-guided DNA endonucleases2,3. TnpB associates with a single, long RNA (ωRNA) and cleaves double-stranded DNA targets complementary to the ωRNA guide. However, the RNA-guided DNA cleavage mechanism of TnpB and its evolutionary relationship with Cas12 enzymes remain unknown. Here we report the cryo-electron microscopy (cryo-EM) structure of Deinococcus radiodurans ISDra2 TnpB in complex with its cognate ωRNA and target DNA. In the structure, the ωRNA adopts an unexpected architecture and forms a pseudoknot, which is conserved among all guide RNAs of Cas12 enzymes. Furthermore, the structure, along with our functional analysis, reveals how the compact TnpB recognizes the ωRNA and cleaves target DNA complementary to the guide. A structural comparison of TnpB with Cas12 enzymes suggests that CRISPR-Cas12 effectors acquired an ability to recognize the protospacer-adjacent motif-distal end of the guide RNA-target DNA heteroduplex, by either asymmetric dimer formation or diverse REC2 insertions, enabling engagement in CRISPR-Cas adaptive immunity. Collectively, our findings provide mechanistic insights into TnpB function and advance our understanding of the evolution from transposon-encoded TnpB proteins to CRISPR-Cas12 effectors.

Computational design of CRISPR guide RNAs to enable strain-specific control of microbial consortia

Microbes naturally coexist in complex, multistrain communities. However, extracting individual microbes from and specifically manipulating the composition of these consortia remain challenging. The sequence-specific nature of CRISPR guide RNAs can be leveraged to accurately differentiate microorganisms and facilitate the creation of tools that can achieve these tasks. We developed a computational program, ssCRISPR, which designs strain-specific CRISPR guide RNA sequences with user-specified target strains, protected strains, and guide RNA properties. We experimentally verify the accuracy of the strain specificity predictions in both Escherichia coli and Pseudomonas spp. and show that up to three nucleotide mismatches are often required to ensure perfect specificity. To demonstrate the functionality of ssCRISPR, we apply computationally designed CRISPR-Cas9 guide RNAs to two applications: the purification of specific microbes through one- and two-plasmid transformation workflows and the targeted removal of specific microbes using DNA-loaded liposomes. For strain purification, we utilize gRNAs designed to target and kill all microbes in a consortium except the specific microbe to be isolated. For strain elimination, we utilize gRNAs designed to target only the unwanted microbe while protecting all other strains in the community. ssCRISPR will be of use in diverse microbiota engineering applications.

A Practical Guide for CRISPR-Cas9-Induced Mutations in Axolotls

Clustered regularly interspaced short palindromic repeats (CRISPR) is a powerful tool that enables editing of the axolotl genome. In this chapter, we will cover how to retrieve gene sequences, confirm annotation, design CRISPR targets, analyze indels, and screen for Crispant axolotls. This is a comprehensive guide on how to use CRISPR on your favorite gene and gain insights into its function.

Simplified CRISPR-Mediated DNA Editing in Multicellular Eukaryotes

The CRISPR-Cas9 system is becoming an imperative tool to edit the genome of various organisms. The gene-editing study by the CRISPR-Cas9 system has revolutionized the diverse field of biomedical research, genome engineering, and gene therapy. CRISPR-Cas9 system has been modified to induce genome editing by small-guide RNAs, which function together with Cas9 nuclease for sequence-specific cleavage of target sequences. Here, we describe the simplified protocol of CRISPR-Cas9-mediated DNA editing in multicellular eukaryotes, including the construction of gRNA plasmids into vectors, screening of positive clones, transfections into 293FT cell line, and transduction into Jurkat cells. We also describe different bioinformatic tools to design suitable gRNAs with increased efficiency and decreased off-target events. Further, we describe the assessments of DNA editing by indel mutations and sequencing in transduced cells.

Efficient Targeted DNA Methylation with dCas9-Coupled DNMT3A-DNMT3L Methyltransferase

Epigenome editing is a powerful approach for the establishment of a chromatin environment with desired properties at a selected genomic locus, which is used to influence the transcription of target genes and to study properties and functions of gene regulatory elements. Targeted DNA methylation is one of the most often used types of epigenome editing, which typically aims for gene silencing by methylation of gene promoters. Here, we describe the design principles of EpiEditors for targeted DNA methylation and provide step-by-step guidelines for the realization of this approach. We focus on the dCas9 protein as the state-of-the-art DNA targeting module fused to 10×SunTag as the most frequently used system for editing enhancement. Further, we discuss different flavors of DNA methyltransferase modules used for this purpose including the most specific variants developed recently. Finally, we explain the principles of gRNA selection, outline the setup of the cell culture experiments, and briefly introduce the available options for the downstream DNA methylation data analysis.

Design, Construction, and Validation of Targeted Gene Activation with TREE System in Human Cells

Genome editing technologies can be diverted into artificial transcription activators. In particular, researchers have improved dCas9-based technologies by tandem-fusing or trans-accumulating effector domains. Previously, we developed a hierarchical effector accumulation system named "TREE," enabling robust activation of target genes even when strongly silenced. In this chapter, we describe our protocol to design, construct, and validate the TREE-mediated target gene activation in cultured human cells.

Multiplexed measurement of cell type-specific calcium kinetics using high-content image analysis combined with targeted gene disruption

Kinetic analysis of intracellular calcium (Ca2+) in cardiomyocytes is commonly used to determine the pathogenicity of genetic mutations identified in patients with dilated cardiomyopathy (DCM). Conventional methods for measuring Ca2+ kinetics target whole-well cultured cardiomyocytes and therefore lack information concerning individual cells. Results are also affected by heterogeneity in cell populations. Here, we developed an analytical method using CRISPR/Cas9 genome editing combined with high-content image analysis (HCIA) that links cell-by-cell Ca2+ kinetics and immunofluorescence images in thousands of cardiomyocytes at a time. After transfecting cultured mouse cardiomyocytes that constitutively express Cas9 with gRNAs, we detected a prolonged action potential duration specifically in Serca2a-depleted ventricular cardiomyocytes in mixed culture. To determine the phenotypic effect of a frameshift mutation in PKD1 in a patient with DCM, we introduced the mutation into Cas9-expressing cardiomyocytes by gRNA transfection and found that it decreases the expression of PKD1-encoded PC1 protein that co-localizes specifically with Serca2a and L-type voltage-gated calcium channels. We also detected the suppression of Ca2+ amplitude in ventricular cardiomyocytes with decreased PC1 expression in mixed culture. Our HCIA method provides comprehensive kinetic and static information on individual cardiomyocytes and allows the pathogenicity of mutations to be determined rapidly.

Characterization of Impurities in Therapeutic RNAs at the Single Nucleotide Level

The chemistry of guide RNA (gRNA) affects the performance of the clustered regularly interspaced short palindromic repeats (CRISPR)-Cas9 genome editing technique. However, the literature is very scarce about the study of gRNA degradation and in particular at the single nucleotide level. In this work, we developed a workflow to characterize the impurities of large RNAs at the single nucleotide level and identified the residues prone to degradation. Our strategy involves (i) the reduction of RNA length, (ii) a chromatographic mode able to capture subtle changes in impurity polarity, and (iii) a streamlined data treatment. To illustrate the approach, stressed gRNA samples were analyzed by coupling an immobilized ribonuclease T1 cartridge to a hydrophilic interaction liquid chromatography (HILIC) column hyphenated with tandem mass spectrometry (MS/MS). Critical findings were made possible by the presented technology. In particular, the desulfurization of phosphorothioate (PS) linkages was the major degradation observed at the single nucleotide level while no change in purity profile could be observed when using conventional ion-pairing reversed-phase (IPRP) liquid chromatography. To our knowledge, this is the first time that several impurity types are screened for a large RNA molecule using an automated online digestion analysis approach.

De novo assembly and annotation of the CHOZN® GS-/- genome supports high-throughput genome-scale screening

Chinese hamster ovary (CHO) cells have been used as the industry standard for the production of therapeutic monoclonal antibodies for several decades. Despite significant improvements in commercial-scale production processes and media, the CHO cell has remained largely unchanged. Due to the cost and complexity of whole-genome sequencing and gene-editing it has been difficult to obtain the tools necessary to improve the CHO cell line. With the advent of next-generation sequencing and the discovery of the CRISPR/Cas9 system it has become more cost effective to sequence and manipulate the CHO genome. Here, we provide a comprehensive de novo assembly and annotation of the CHO-K1 based CHOZN® GS-/- genome. Using this platform, we designed, built, and confirmed the functionality of a whole genome CRISPR guide RNA library that will allow the bioprocessing community to design a more robust CHO cell line leading to the production of life saving medications in a more cost-effective manner.

Capturing nucleic acid variants with precision using CRISPR diagnostics

CRISPR/Cas systems have the ability to precisely target nucleotide sequences and enable their rapid identification and modification. While nucleotide modification has enabled the therapeutic correction of diseases, the process of identifying the target DNA or RNA has greatly expanded the field of molecular diagnostics in recent times. CRISPR-based DNA/RNA detection through programmable nucleic acid binding or cleavage has been demonstrated for a large number of pathogenic and non-pathogenic targets. Combining CRISPR detection with nucleic acid amplification and a terminal signal readout step allowed the development of numerous rapid and robust nucleic acid platforms. Wherever the Cas effector can faithfully distinguish nucleobase variants in the target, the platform can also be extended for sequencing-free rapid variant detection. Some initial PAM disruption-based SNV detection reports were limited to finding or integrating mutated/mismatched nucleotides within the PAM sequences. In this review, we try to summarize the developments made in CRISPR diagnostics (CRISPRDx) to date emphasizing CRISPR-based SNV detection. We also discuss the applications where such diagnostic modalities can be put to use, covering various fields of clinical research, SNV screens, disease genotyping, primary surveillance during microbial infections, agriculture, food safety, and industrial biotechnology. The ease of rapid design and implementation of such multiplexable assays can potentially expand the applications of CRISPRDx in the domain of affinity-based target sequencing, with immense possibilities for low-cost, quick, and widespread usage. In the end, in combination with proximity assays and a suicidal gene approach, CRISPR-based in vivo SNV detection and cancer cell targeting can be formulated as personalized gene therapy.

Enzyme-free targeted DNA demethylation using CRISPR-dCas9-based steric hindrance to identify DNA methylation marks causal to altered gene expression

DNA methylation involves the enzymatic addition of a methyl group primarily to cytosine residues in DNA. This protocol describes how to produce complete and minimally confounded DNA demethylation of specific sites in the genome of cultured cells by clustered regularly interspaced short palindromic repeats (CRISPR)-dCas9 and without the involvement of an epigenetic-modifying enzyme, the purpose of which is the evaluation of the functional (i.e., gene expression or phenotypic) consequences of DNA demethylation of specific sites that have been previously implicated in particular pathological or physiological contexts. This protocol maximizes the ability of the easily reprogrammable CRISPR-dCas9 system to assess the impact of DNA methylation from a causal rather than correlational perspective: alternative protocols for CRISPR-dCas9-based site-specific DNA methylation or demethylation rely on the recruitment of epigenetic enzymes that exhibit additional nonspecific activities at both the targeted site and throughout the genome, confounding conclusions of causality of DNA methylation. Inhibition or loss of DNA methylation is accomplished by three consecutive lentiviral transductions. The first two lentiviruses establish stable expression of dCas9 and a guide RNA, which will physically obstruct either maintenance or de novo DNA methyltransferase activity at the guide RNA target site. A third lentivirus introduces Cre recombinase to delete the dCas9 transgene, which leads to loss of dCas9 from the target site, allowing transcription factors and/or the transcription machinery to interact with the demethylated target site. This protocol requires 3-8 months to complete owing to prolonged cell passaging times, but there is little hands-on time, and no specific skills beyond basic molecular biology techniques are necessary.

Self-cleaving guide RNAs enable pharmacological selection of precise gene editing events in vivo

Expression of guide RNAs in the CRISPR/Cas9 system typically requires the use of RNA polymerase III promoters, which are not cell-type specific. Flanking the gRNA with self-cleaving ribozyme motifs to create a self-cleaving gRNA overcomes this limitation. Here, we use self-cleaving gRNAs to create drug-selectable gene editing events in specific hepatocyte loci. A recombinant Adeno Associated Virus vector targeting the Albumin locus with a promoterless self-cleaving gRNA to create drug resistance is linked in cis with the therapeutic transgene. Gene expression of both are dependent on homologous recombination into the target locus. In vivo drug selection for the precisely edited hepatocytes allows >30-fold expansion of gene-edited cells and results in therapeutic levels of a human Factor 9 transgene. Importantly, self-cleaving gRNA expression is also achieved after targeting weak hepatocyte genes. We conclude that self-cleaving gRNAs are a powerful system to enable cell-type specific in vivo drug resistance for therapeutic gene editing applications.

EasyGuide Plasmids Support in Vivo Assembly of gRNAs for CRISPR/Cas9 Applications in Saccharomyces cerevisiae

Most CRISPR/Cas9 applications in yeast rely on a plasmid-based expression of Cas9 and its guide RNA (gRNA) containing a 20-nucleotides (nts) spacer tailored to each genomic target. The lengthy assembly of this customized gRNA requires at least 3-5 days for its precloning in Escherichia coli, purification, validation, and cotransformation with Cas9 into a yeast strain. Here, we constructed a series of 12 EasyGuide plasmids to simplify CRISPR/Cas9 applications in Saccharomyces cerevisiae. The new vectors provide templates for generating PCR fragments that can assemble up to six functional gRNAs directly into yeasts via homologous recombination between the 20-nts spacers. By dispensing precloning in E. coli, yeast in vivo gRNA assembly significantly reduces the CRISPR/Cas9 experimental workload. A highly efficient yeast genome editing procedure, involving PCR amplification of gRNAs and donors, followed by their transformation into a Cas9-expressing strain, can be easily accomplished through a quick protocol.

Host Cell Transcriptional Tuning with CRISPR/dCas9 to Mitigate the Effects of Toxin Exposure

Anthrax infection is caused byBacillus anthracis, a bacterium that once established within the host releases lethal toxin (LeTx). Anthrax LeTx is internalized by the capillary morphogenesis protein 2/anthrax toxin receptor 2 (CMG2/ANTXR2) cell surface receptor on mammalian cells. Once inside the cell, LeTx cleaves mitogen-activated protein kinases (MAPKs), ultimately leading to cell death. Previous reports have shown that decreased expression of ANTXR2 reduces cell susceptibility to LeTx. By ablating the ANTXR2 gene in cells in vitro, we observed complete resistance to LeTx-induced cell death. Here, we directed CRISPR/dCas9-based tools to the ANTXR2 promoter to modulate ANTXR2 expression without altering the underlying gene sequence in human cell lines that express the receptor at high levels. We hypothesized that downregulating the expression of the ANTXR2 gene at the genomic level would mitigate the impact of toxin exposure. In one epigenetic editing approach, we employed the fusion of DNMT3A DNA methyltransferase and dCas9 (dCas9-DNMT3A) to methylate CpGs within the CpG island of the ANTXR2 promoter and found this repressed ANTXR2 gene expression resulting in significant resistance to LeTx-induced cell death. Furthermore, by multiplexing gRNAs to direct dCas9-DNMT3A to multiple sites in the ANTXR2 promoter, we applied a broader distribution of CpG methylation along the gene promoter resulting in enhanced repression and resistance to LeTx. In parallel, we directed the dCas9-KRAB-MeCP2 transcriptional repressor to the ANTXR2 promoter to quickly and robustly repress ANTXR2 expression. With this approach, in as little as two weeks, we created resistance to LeTx at a similar level to ANTXR2 gene-ablated cells. Overall, we present a transcriptional tuning approach to inhibit the effects of LeTx and provide a framework to repress toxin-binding cell surface receptors.

New Effective Method of Lactococcus Genome Editing Using Guide RNA-Directed Transposition

Lactococcus lactis is an important industrial microorganism and a widely used model object for research in the field of lactic acid bacteria (LAB) biology. The development of new L. lactis and related LAB strains with improved properties, including phage-resistant strains for dairy fermentation, LAB-based vaccines or strains with altered genotypes for research purposes, are hindered by the lack of genome-editing tools that allow for the easy and straightforward incorporation of a significant amount of the novel genetic material, such as large genes or operons, into the chromosomes of these bacteria. We recently employed a suggested system based on the CRISPR-Cas-associated transposon for the editing of the L. lactis genome. After the in-depth redesign of the system, we were able to achieve the stable incorporation of the fragments that were sized up to 10 kbp into the L. lactis beta-galactosidase gene. The efficiency of editing under the optimized conditions were 2 × 10-4 and 4 × 10-5 for 1 kbp and 10 kbp, respectively, which are sufficient for fast and easy modifications if a positive selection marker can be used.

Identification and Analysis of Small Molecule Inhibitors of CRISPR-Cas9 in Human Cells

Genome editing tools based on CRISPR-Cas systems can repair genetic mutations in situ; however, off-target effects and DNA damage lesions that result from genome editing remain major roadblocks to its full clinical implementation. Protein and chemical inhibitors of CRISPR-Cas systems may reduce off-target effects and DNA damage. Here we describe the identification of several lead chemical inhibitors that could specifically inhibit the activity of Streptococcus pyogenes Cas9 (SpCas9). In addition, we obtained derivatives of lead inhibitors that could penetrate the cell membrane and inhibit SpCas9 in cellulo. Two of these compounds, SP2 and SP24, were able to improve the specificity of SpCas9 in cellulo at low-micromolar concentration. Furthermore, microscale thermophoresis (MST) assays showed that SP24 might inhibit SpCas9 activity by interacting with both the SpCas9 protein and the SpCas9-gRNA ribonucleoprotein complex. Taken together, SP24 is a novel chemical inhibitor of SpCas9 which has the potential to enhance therapies that utilize SpCas9.

Site-specific CRISPR-based mitochondrial DNA manipulation is limited by gRNA import

Achieving CRISPR Cas9-based manipulation of mitochondrial DNA (mtDNA) has been a long-standing goal and would be of great relevance for disease modeling and for clinical applications. In this project, we aimed to deliver Cas9 into the mitochondria of human cells and analyzed Cas9-induced mtDNA cleavage and measured the resulting mtDNA depletion with multiplexed qPCR. In initial experiments, we found that measuring subtle effects on mtDNA copy numbers is challenging because of high biological variability, and detected no significant Cas9-caused mtDNA degradation. To overcome the challenge of being able to detect Cas9 activity on mtDNA, we delivered cytosine base editor Cas9-BE3 to mitochondria and measured its effect (C →  T mutations) on mtDNA. Unlike regular Cas9-cutting, this leaves a permanent mark on mtDNA that can be detected with amplicon sequencing, even if the efficiency is low. We detected low levels of C → T mutations in cells that were exposed to mitochondrially targeted Cas9-BE3, but, surprisingly, these occurred regardless of whether a guide RNA (gRNA) specific to the targeted site, or non-targeting gRNA was used. This unspecific off-target activity shows that Cas9-BE3 can technically edit mtDNA, but also strongly indicates that gRNA import to mitochondria was not successful. Going forward mitochondria-targeted Cas9 base editors will be a useful tool for validating successful gRNA delivery to mitochondria without the ambiguity of approaches that rely on quantifying mtDNA copy numbers.

Genome-wide base editor screen identifies regulators of protein abundance in yeast

Proteins are key molecular players in a cell, and their abundance is extensively regulated not just at the level of gene expression but also post-transcriptionally. Here, we describe a genetic screen in yeast that enables systematic characterization of how protein abundance regulation is encoded in the genome. The screen combines a CRISPR/Cas9 base editor to introduce point mutations with fluorescent tagging of endogenous proteins to facilitate a flow-cytometric readout. We first benchmarked base editor performance in yeast with individual gRNAs as well as in positive and negative selection screens. We then examined the effects of 16,452 genetic perturbations on the abundance of eleven proteins representing a variety of cellular functions. We uncovered hundreds of regulatory relationships, including a novel link between the GAPDH isoenzymes Tdh1/2/3 and the Ras/PKA pathway. Many of the identified regulators are specific to one of the eleven proteins, but we also found genes that, upon perturbation, affected the abundance of most of the tested proteins. While the more specific regulators usually act transcriptionally, broad regulators often have roles in protein translation. Overall, our novel screening approach provides unprecedented insights into the components, scale and connectedness of the protein regulatory network.

Improving Precise Genome Editing Using Donor DNA/gRNA Hybrid Duplex Generated by Complementary Bases

In precise genome editing, site-specific DNA double-strand breaks (DSBs) induced by the CRISPR/Cas9 system are repaired via homology-directed repair (HDR) using exogenous donor DNA templates. However, the low efficiency of HDR-mediated genome editing is a barrier to widespread use. In this study, we created a donor DNA/guide RNA (gRNA) hybrid duplex (DGybrid) that was composed of sequence-extended gRNA and single-stranded oligodeoxynucleotide (ssODN) combined with complementary bases without chemical modifications to increase the concentration of donor DNA at the cleavage site. The efficiency of genome editing using DGybrid was evaluated in Saccharomyces cerevisiae. The results show a 1.8-fold (from 35% to 62%) improvement in HDR-mediated editing efficiency compared to genome editing in which gRNA and donor DNA were introduced separately. In addition, analysis of the nucleic acid introduction efficiency using flow cytometry indicated that both RNA and ssODNs are efficiently incorporated into cells together by using the DNA/RNA hybrid. Our technique would be preferred as a universal and concise tool for improving the efficiency of HDR-mediated genome editing.

A comprehensive Bioconductor ecosystem for the design of CRISPR guide RNAs across nucleases and technologies

The success of CRISPR-mediated gene perturbation studies is highly dependent on the quality of gRNAs, and several tools have been developed to enable optimal gRNA design. However, these tools are not all adaptable to the latest CRISPR modalities or nucleases, nor do they offer comprehensive annotation methods for advanced CRISPR applications. Here, we present a new ecosystem of R packages, called crisprVerse, that enables efficient gRNA design and annotation for a multitude of CRISPR technologies. This includes CRISPR knockout (CRISPRko), CRISPR activation (CRISPRa), CRISPR interference (CRISPRi), CRISPR base editing (CRISPRbe) and CRISPR knockdown (CRISPRkd). The core package, crisprDesign, offers a user-friendly and unified interface to add off-target annotations, rich gene and SNP annotations, and on- and off-target activity scores. These functionalities are enabled for any RNA- or DNA-targeting nucleases, including Cas9, Cas12, and Cas13. The crisprVerse ecosystem is open-source and deployed through the Bioconductor project ( https://github.com/crisprVerse ).

Marker counter-selection via CRISPR/Cas9 co-targeting for efficient generation of genome edited avian cell lines and germ cells

Efficient isolation of genetically modified cells that are phenotypically indistinguishable from the unmodified cells remains a major technical barrier for the broader utilization of CRISPR/Cas9. Here, we report a novel enrichment approach to select the genome engineered cells by co-targeting a genomically integrated GFP gene along with the endogenous gene of interest (GOI). Using this co-targeting approach, multiple genomic loci were successfully targeted in chicken (DF1) and quail (CEC-32) fibroblast cell lines by transient transfection of Cas9 and guide RNAs (gRNAs). Clonal isolation of co-targeted DF1 cells showed 75% of cell clones had deletion of GFP and biallelic deletion of the GOI. To assess the utility of this approach to generate genome modified animals, we tested it on chicken primordial germ cells (PGCs) expressing GFP by co-targeting with gRNAs against GFP and endogenous ovomucoid (OVM) gene. PGCs enriched for loss of GFP and confirmed for OVM deletion, derived by co-targeting, were injected into Hamburger and Hamilton stage 14-15 chicken embryos, and their ability to migrate to the genital ridge was confirmed. This simple, efficient enrichment approach could easily be applied to the creation of knock-out or edited cell lines or animals.

Frankenstein Cas9: engineering improved gene editing systems

The discovery of CRISPR-Cas9 and its widespread use has revolutionised and propelled research in biological sciences. Although the ability to target Cas9's nuclease activity to specific sites via an easily designed guide RNA (gRNA) has made it an adaptable gene editing system, it has many characteristics that could be improved for use in biotechnology. Cas9 exhibits significant off-target activity and low on-target nuclease activity in certain contexts. Scientists have undertaken ambitious protein engineering campaigns to bypass these limitations, producing several promising variants of Cas9. Cas9 variants with improved and alternative activities provide exciting new tools to expand the scope and fidelity of future CRISPR applications.

G-quadruplex-guided RNA engineering to modulate CRISPR-based genomic regulation

It is important to develop small moelcule-based methods to modulate gene editing and expression in human cells. The roles of the G-quadruplex (G4) in biological systems have been widely studied. Here, G4-guided RNA engineering is performed to generate guide RNA with G4-forming units (G4-gRNA). We further demonstrate that chemical targeting of G4-gRNAs holds promise as a general approach for modulating gene editing and expression in human cells. The rich structural diversity of RNAs offers a reservoir of targets for small molecules to bind, thus creating the potential to modulate RNA biology.

CRISPR Spherical Nucleic Acids

The use of CRISPR/Cas9 systems in genome editing has been limited by the inability to efficiently deliver the key editing components to and across tissues and cell membranes, respectively. Spherical nucleic acids (SNAs) are nanostructures that provide privileged access to both but have yet to be explored as a means of facilitating gene editing. Herein, a new class of CRISPR SNAs are designed and evaluated in the context of genome editing. Specifically, Cas9 ProSNAs comprised of Cas9 cores densely modified with DNA on their exteriors and preloaded with single-guide RNA were synthesized and evaluated for their genome editing capabilities in the context of multiple cell lines. The radial orientation of the DNA on the Cas9 protein surface enhances cellular uptake, without the need for electroporation or transfection agents. In addition, the Cas9 proteins defining the cores of the ProSNAs were fused with GALA peptides on their N-termini and nuclear localization signals on their C-termini to facilitate endosomal escape and maximize nuclear localization and editing efficiency, respectively. These constructs were stable against protease digestion under conditions that fully degrade the Cas9 protein, when not transformed into an SNA, and used to achieve genome editing efficiency between 32 and 47%. Taken together, these novel constructs and advances point toward a way of significantly broadening the scope of use and impact of CRISPR-Cas9 genome editing systems.

CRISPRactivation-SMS, a message for PAM sequence independent gene up-regulation in Escherichia coli

Governance of the endogenous gene regulatory network enables the navigation of cells towards beneficial traits for recombinant protein production. CRISPRactivation and interference provides the basis for gene expression modulation but is primarily applied in eukaryotes. Particularly the lack of wide-ranging prokaryotic CRISPRa studies might be attributed to intrinsic limitations of bacterial activators and Cas9 proteins. While bacterial activators need accurate spatial orientation and distancing towards the target promoter to be functional, Cas9-based CRISPR tools only bind sites adjacent to NGG PAM sequences. These circumstances hampered Cas9-guided activators from mediating the up-regulation of endogenous genes at precise positions in bacteria. We could overcome this limitation by combining the PAM independent Cas9 variant SpRY and a CRISPRa construct using phage protein MCP fused to transcriptional activator SoxS. This CRISPRa construct, referred to as SMS, was compared with previously reported CRISPRa constructs and showed up-regulation of a reporter gene library independent of its PAM sequence in Escherichia coli. We also demonstrated down-regulation and multi-gene expression control with SMS at non-NGG PAM sites. Furthermore, we successfully applied SMS to up-regulate endogenous genes, and transgenes at non-NGG PAM sites, which was impossible with the previous CRISPRa construct.

Specificity of oligonucleotide gene therapy (OGT) agents

Oligonucleotide gene therapy (OGT) agents (e. g. antisense, deoxyribozymes, siRNA and CRISPR/Cas) are promising therapeutic tools. Despite extensive efforts, only few OGT drugs have been approved for clinical use. Besides the problem of efficient delivery to targeted cells, hybridization specificity is a potential limitation of OGT agents. To ensure tight binding, a typical OGT agent hybridizes to the stretch of 15-25 nucleotides of a unique targeted sequence. However, hybrids of such lengths tolerate one or more mismatches under physiological conditions, the problem known as the affinity/specificity dilemma. Here, we assess the scale of this problem by analyzing OGT hybridization-dependent off-target effects (HD OTE) in vitro, in animal models and clinical studies. All OGT agents except deoxyribozymes exhibit HD OTE in vitro, with most thorough evidence of poor specificity reported for siRNA and CRISPR/Cas9. Notably, siRNA suppress non-targeted genes due to (1) the partial complementarity to mRNA 3'-untranslated regions (3'-UTR), and (2) the antisense activity of the sense strand. CRISPR/Cas9 system can cause hundreds of non-intended dsDNA breaks due to low specificity of the guide RNA, which can limit therapeutic applications of CRISPR/Cas9 by ex-vivo formats. Contribution of this effects to the observed in vivo toxicity of OGT agents is unclear and requires further investigation. Locked or peptide nucleic acids improve OGT nuclease resistance but not specificity. Approaches that use RNA marker dependent (conditional) activation of OGT agents may improve specificity but require additional validation in cell culture and in vivo.

Detection of Cancer Marker Flap Endonuclease 1 Using One-Pot Transcription-Powered Clustered Regularly Interspaced Short Palindromic Repeat/Cas12a Signal Expansion

As a critical functional protein in DNA replication and genome stability, flap endonuclease 1 (FEN1) has been considered a promising biomarker and druggable target for multiple cancers. We report here a transcription-powered clustered regularly interspaced short palindromic repeat (CRISPR)/Cas12a signal expansion platform for rapid and sensitive detection of FEN1. In this method, the probe cleavage by FEN1 generated a free 5' flap single-stranded DNA which could hybridize with the single-stranded T7 promoter-bearing template and trigger the extension. Then, the CRISPR guide RNA (crRNA) transcribed from the extended template activated the collateral DNase activity of Cas12a, releasing the fluorophore from the quenched DNA signal probe to report the FEN1 detection result. The high specificity for FEN1 was validated by comparing with other repair-relevant proteins. The limit of detection (LOD) could be as low as 0.03 mU, which is sensitive enough to detect the FEN1 activity in biological samples. In addition, the inhibition assay of FEN1 was also successfully achieved with this platform, proving its potential in inhibitor screening. In summary, this study provides a novel biosensor for FEN1 activity analysis and provides new insights into the development of CRISPR-based biosensors for non-nucleic acid targets.

Molecular mechanism of active Cas7-11 in processing CRISPR RNA and interfering target RNA

Cas7-11 is a Type III-E CRISPR Cas effector that confers programmable RNA cleavage and has potential applications in RNA interference. Cas7-11 encodes a single polypeptide containing four Cas7- and one Cas11-like segments that obscures the distinction between the multi-subunit Class 1 and the single-subunit Class-2 CRISPR Cas systems. We report a cryo-EM (cryo-electron microscopy) structure of the active Cas7-11 from Desulfonema ishimotonii (DiCas7-11) that reveals the molecular basis for RNA processing and interference activities. DiCas7-11 arranges its Cas7- and Cas11-like domains in an extended form that resembles the backbone made up by four Cas7 and one Cas11 subunits in the multi-subunit enzymes. Unlike the multi-subunit enzymes, however, the backbone of DiCas7-11 contains evolutionarily different Cas7 and Cas11 domains, giving rise to their unique functionality. The first Cas7-like domain nearly engulfs the last 15 direct repeat nucleotides in processing and recognition of the CRISPR RNA, and its free-standing fragment retains most of the activity. Both the second and the third Cas7-like domains mediate target RNA cleavage in a metal-dependent manner. The structure and mutational data indicate that the long variable insertion to the fourth Cas7 domain has little impact on RNA processing or targeting, suggesting the possibility for engineering a compact and programmable RNA interference tool.

Optimization of specific RNA knockdown in mammalian cells with CRISPR-Cas13

Prokaryotic adaptive immune systems use Clustered Regularly Interspaced Short Palindromic Repeats (CRISPRs) and CRISPR Associated (Cas) proteins to target and cleave foreign genetic elements in an RNA-guided manner [1-3]. Type VI CRISPR-Cas systems contain a single effector ribonuclease, Cas13, that binds and processes a CRISPR-RNA (crRNA; also known as a guide-RNA), forming an RNA-guided RNA-targeting effector complex [4,5]. Previous studies have shown that Cas13 can be engineered to target and modulate RNA processes in human cells, illustrating the versatility and specificity of Cas13 as an RNA knockdown (KD), splicing, editing, or imaging tool [6-8]. While Cas13 has been successfully used by several groups, our lab has observed significant variability in Cas13 KD ability depending which protocol is being followed [9-12]. To further understand this variability and generate a robust Cas13 KD protocol we thoroughly tested which Cas13 ortholog to use, the duration of KD experiments, the amount of plasmid DNA transfected, methods for analyzing KD efficiency, and report an optimized method for carrying out and analyzing Cas13 mediated RNA KD experiments. The method outlined in this paper illustrates a faster and more reliable protocol to iteratively test gRNA performance and target gene KD.

Optimizing CRISPR/Cas9-based gene manipulation in echinoderms

The impact of new technology can be appreciated by how broadly it is used. Investigators that previously relied only on pharmacological approaches or the use of morpholino antisense oligonucleotide (MASO) technologies are now able to apply CRISPR-Cas9 to study biological problems in their model organism of choice much more effectively. The transitions to new CRISPR-based approaches could be enhanced, first, by standardized protocols and education in their applications. Here we summarize our results for optimizing the CRISPR-Cas9 technology in a sea urchin and a sea star, and provide advice on how to set up CRISPR-Cas9 experiments and interpret the results in echinoderms. Our goal through these protocols and sharing examples of success by other labs is to lower the activation barrier so that more laboratories can apply CRISPR-Cas9 technologies in these important animals.

Application of CRISPR/Cas9 system and the preferred no-indel end-joining repair in tardigrades

Tardigrades are small aquatic animals known for the tolerant ability against various extreme stresses. Recent studies identified several tardigrade-unique proteins as protective factors of biomolecules from extreme stresses. Due to the limitation of the technique available in tardigrades, the function of these protective molecules has largely been studied utilizing the systems of in vitro and the heterologous expression in other organisms. Although RNAi is feasible in tardigrades, their effects are variable and not always sufficient. To analyze the functions of the tardigrade protective proteins, in vivo genetic manipulations have been desired. In this study, we used a tardigrade Hypsibius exemplaris as a model whose genome is available, and developed the delivery method of Cas9 ribonucleoproteins (RNPs) to adult tardigrade cells. Cas9 RNPs containing two kinds of crRNAs were injected to the body cavity of adult tardigrades and subjected to the subsequent electroporation to facilitate the incorporation of RNPs to the cells. Using this delivery method, we detected the deletion of the intervening region between two crRNAs from the genome. Intriguingly, all examined joining sites exhibited no incorporation of insertions/deletions (indels), suggesting that no-indel end-joining is dominant repair system in this tardigrade. We also detected similar removal of the intervening region even in the tardigrades injected with Cas9 RNPs without electroporation and in this case the no-indel end-joining is detected in still dominant but not all examined joining sites. This study provides the development of the delivery method of Cas9 RNPs to tardigrade cells and our data also suggested that simultaneous application of more than two crRNAs/gRNAs are recommended to disrupt the target gene by CRISPR/Cas9 system to avoid scarless repair in the tardigrade.

Target-enriched nanopore sequencing and de novo assembly reveals co-occurrences of complex on-target genomic rearrangements induced by CRISPR-Cas9 in human cells

The CRISPR-Cas9 system is widely used to permanently delete genomic regions via dual guide RNAs. Genomic rearrangements induced by CRISPR-Cas9 can occur, but continuous technical developments make it possible to characterize complex on-target effects. We combined an innovative droplet-based target enrichment approach with long-read sequencing and coupled it to a customized de novo sequence assembly. This approach enabled us to dissect the sequence content at kilobase scale within an on-target genomic locus. We here describe extensive genomic disruptions by Cas9, involving the allelic co-occurrence of a genomic duplication and inversion of the target region, as well as integrations of exogenous DNA and clustered interchromosomal DNA fragment rearrangements. Furthermore, we found that these genomic alterations led to functional aberrant DNA fragments and can alter cell proliferation. Our findings broaden the consequential spectrum of the Cas9 deletion system, reinforce the necessity of meticulous genomic validations, and introduce a data-driven workflow enabling detailed dissection of the on-target sequence content with superior resolution.

Genome editing using a versatile vector-based CRISPR/Cas9 system in Fusarium species

Fusarium species include important filamentous fungal pathogens that can infect plants, animals, and humans. Meanwhile, some nonpathogenic Fusarium species are promising biocontrol agents against plant pathogens. Here, we developed a genome editing technology using a vector-based CRISPR/Cas9 system for Fusarium oxysporum f. sp. lycopersici (Fol). This optimized CRISPR/Cas9 system, harboring an endogenous U6 small nuclear RNA promoter for the expression of single-guide RNA and an endogenous H2B nuclear localization signal for the localization of Cas9, enabled efficient targeted gene knock-out, including in the accessory chromosomal regions in Fol. We further demonstrated single crossover-mediated targeted base editing and endogenous gene tagging. This system was also applicable for genome editing in F. oxysporum f. sp. spinaciae and F. commune without any modifications, suggesting that this CRISPR/Cas9 vector has a potential application for a broad range of researches on other Fusarium species.

Insights Gained from RNA Editing Targeted by the CRISPR-Cas13 Family

Clustered regularly interspaced short palindromic repeat (CRISPR)/CRISPR-associated protein (Cas) systems, especially type II (Cas9) systems, have been widely developed for DNA targeting and formed a set of mature precision gene-editing systems. However, the basic research and application of the CRISPR-Cas system in RNA is still in its early stages. Recently, the discovery of the CRISPR-Cas13 type VI system has provided the possibility for the expansion of RNA targeting technology, which has broad application prospects. Most type VI Cas13 effectors have dinuclease activity that catalyzes pre-crRNA into mature crRNA and produces strong RNA cleavage activity. Cas13 can specifically recognize targeted RNA fragments to activate the Cas13/crRNA complex for collateral cleavage activity. To date, the Cas13X protein is the smallest effector of the Cas13 family, with 775 amino acids, which is a promising platform for RNA targeting due to its lack of protospacer flanking sequence (PFS) restrictions, ease of packaging, and absence of permanent damage. This study highlighted the latest progress in RNA editing targeted by the CRISPR-Cas13 family, and discussed the application of Cas13 in basic research, nucleic acid diagnosis, nucleic acid tracking, and genetic disease treatment. Furthermore, we clarified the structure of the Cas13 protein family and their molecular mechanism, and proposed a future vision of RNA editing targeted by the CRISPR-Cas13 family.

Allosteric activation of CRISPR-Cas12a requires the concerted movement of the bridge helix and helix 1 of the RuvC II domain

Nucleases derived from the prokaryotic defense system CRISPR-Cas are frequently re-purposed for gene editing and molecular diagnostics. Hence, an in-depth understanding of the molecular mechanisms of these enzymes is of crucial importance. We focused on Cas12a from Francisella novicida (FnCas12a) and investigated the functional role of helix 1, a structural element that together with the bridge helix (BH) connects the recognition and the nuclease lobes of FnCas12a. Helix 1 is structurally connected to the lid domain that opens upon DNA target loading thereby activating the active site of FnCas12a. We probed the structural states of FnCas12a variants altered in helix 1 and/or the bridge helix using single-molecule FRET measurements and assayed the pre-crRNA processing, cis- and trans-DNA cleavage activity. We show that helix 1 and not the bridge helix is the predominant structural element that confers conformational stability of FnCas12a. Even small perturbations in helix 1 lead to a decrease in DNA cleavage activity while the structural integrity is not affected. Our data, therefore, implicate that the concerted remodeling of helix 1 and the bridge helix upon DNA binding is structurally linked to the opening of the lid and therefore involved in the allosteric activation of the active site.