The CRISPR Journal

Clonal cell lines harbouring loss-of-function mutations in genes of interest are crucial for studying the cellular functions of the encoded proteins. Recent advances in genome engineering have converged on the CRISPR/Cas9 technology to quickly and reliably generate frameshift mutations in the target genes across various cell lines and species. Although high on-target cleavage efficiencies can be obtained reproducibly, screening and identifying clones with loss-of-function alleles remains a major bottleneck. Here, we describe a single sgRNA strategy to generate CRISPR/Cas9-mediated frameshift mutations in target genes of mammalian cell lines that can be easily and cost-effectively identified. Given the proliferation of workhorse cell lines such as HEK293 and N2a cells and the resulting clonal expansion of the cell type, our protocol can facilitate the isolation of knockout clonal cell lines and their genetic validation within a period of down to 3-4 weeks.

CRISPR/Cas9 mutagenesis is a revolutionary tool for genetics in organismal and cell culture systems. One notable caveat with this system is the potential for phenotype-inducing off-target/background mutations. There has been considerable success in modifying the methodology to minimize these potential confounds. Here we have developed a tool to functionally demonstrate that a targeted mutation of interest is responsible for the phenotype observed. This approach creates revertable mutations in cell culture systems using CRISPR/Cas9-induced homology-directed repair (HDR) to insert a LoxP-flanked transcriptional stop sequence into an early intron of a target gene. This method has the potential to be used in multiplexed and inducible scenarios to restore gene function within a given experiment.

The development of the central nervous system (CNS) depends on tightly regulated gene expression programs that guide neural progenitor differentiation and neuronal subtype specification. The tunicate Ciona robusta provides a powerful and simplified model for dissecting the genetic control of nervous system development, with a larval CNS composed of just over 200 neurons and sensory cells. Although CRISPR/Cas9-mediated mutagenesis is now routinely used in Ciona, validated single-guide RNAs (sgRNAs) have yet to be validated for key neural genes. Here, we report the design and experimental validation of 25 novel sgRNAs targeting eight conserved genes encoding conserved proteins involved in neurodevelopment and neural function, including six transcription factors (Cdx, Foxb, Sox1/2/3, Dmbx, Engrailed, and Mnx) and two neural effector genes (Tyrosinase and Slc18a3/VAChT). Candidate sgRNAs were selected using CRISPOR and tested for mutagenesis efficiency using Illumina-based target site amplicon sequencing. All sgRNAs induced insertions or deletions at their target loci, with most genes yielding at least one sgRNA with mutagenesis efficacy exceeding 30%, with the exception of Dmbx, for which maximal efficacy reached 25%. We further compared measured mutagenesis rates with predicted Doench 16 and Doench Ruleset 3 (RS3) scores, observing a modest but improved correlation with RS3 predictions. Based on these results, we recommend considering both scoring algorithms, with RS3 potentially offering improved predictive value for Ciona.

CRISPR/Cas9-mediated transcriptional activation (CRISPRa) is a powerful tool for investigating complex biological phenomena. Although CRISPRa approaches based on VP64 have been widely studied in both cultured cells and in animal models and exhibit great versatility for various cell types and developmental stages in vivo, different dCas9-VP64 versions have not been rigorously compared. Here, we compared different dCas9-VP64 constructs in identical contexts, including the cell lines used and the transfection conditions, for their ability to activate endogenous and exogenous genes. Moreover, we investigated the optimal approach for VP64 addition to VP64- and p300-based constructs. We found that MS2-MCP-scaffolded VP64 enhanced dCas9-VP64 and dCas9-p300 activity better than did direct VP64 fusion to the N-terminus of dCas9. dCas9-VP64+MCP-VP64 and dCas9-p300+MCP-VP64 were superior to VP64-dCas9-VP64 for all target genes tested. Furthermore, multiplexing gRNA expression with dCas9-VP64+MCP-VP64 or dCas9-p300+MCP-VP64 significantly enhanced endogenous gene activation to a level comparable to CRISPRa-SAM with a single gRNA. Our findings demonstrate improvement of the dCas9-VP64 CRISPRa system and contribute to development of a versatile, efficient CRISPRa platform.

Development of novel CRISPR/Cas systems enhances opportunities for gene editing to treat infectious diseases, cancer, and genetic disorders. We evaluated CasX2 (PlmCas12e), a class II CRISPR system derived from Planctomycetes, a non-pathogenic bacterium present in aquatic and terrestrial soils. CasX2 offers several advantages over Streptococcus pyogenes Cas9 (SpCas9) and Staphylococcus aureus Cas9 (SaCas9), including its smaller size, distinct protospacer adjacent motif (PAM) requirements, staggered cleavage cuts that promote homology-directed repair, and no known pre-existing immunity in humans. A recent study reported that a three amino acid substitution in CasX2 significantly enhanced cleavage activity (1). Therefore, we compared cleavage efficiency and double-stranded break repair characteristics between the native CasX2 and the variant, CasX2Max, for cleavage of CCR5, a gene that encodes the CCR5 receptor important for HIV-1 infection. Two CasX2 single guide RNAs (sgRNAs) were designed that flanked the 32 bases deleted in the natural CCR5 {Delta}32 mutation. Nanopore sequencing demonstrated that CasX2 using sgRNAs with spacers of 17 nucleotides (nt), 20 nt or 23 nt in length were ineffective at cleaving genomic CCR5. In contrast, CasX2Max using sgRNAs with 20 nt and 23 nt spacer lengths, enabled robust genomic cleavage of CCR5. Structural modeling indicated that two of the CasX2Max substitutions enhanced sgRNA-DNA duplex stability, while the third improved DNA strand alignment within the catalytic site. These structural changes likely underlie the increased activity of CasX2Max in cellular gene excision. In sum, CasX2Max consistently outperformed native CasX2 across all assays and represents a superior gene-editing platform for therapeutic applications.

CRISPR/Cas is under development as a therapeutic tool for the cleavage, excision, and/or modification of genes in eukaryotic cells. While much effort has focused on CRISPR/Cas from Streptococcus pyogenes (SpCas9) and Staphylococcus aureus (SaCas9), alternative CRISPR systems have been identified using metagenomic datasets from non-pathogenic microbes, including previously unknown class 2 systems, adding to a diverse toolbox of gene editors. The Cas12e (CasX1, CasX2) endonucleases from non-pathogenic Deltaproteobacteria (DpeCas12e) and Planctomycetes (PlmCas12e) are more compact than SpCas9, have a more selective protospacer adjacent motif (PAM) requirement, and deliver a staggered cleavage cut with 5-7 base overhangs. We investigated varying guide RNA (spacer) lengths and alternative PAM sequences to determine optimal conditions for PlmCas12e cleavage of the cellular gene CCR5 (CC-Chemokine receptor-5). CCR5 encodes one of two chemokine coreceptors required by HIV-1 to infect target cells, and a mutation of CCR5 (delta-32) is responsible for HIV-1 resistance and reported cures following bone marrow transplantation. Consequently, CCR5 has been an important target for gene editing utilizing CRISPR, TALENs, and ZFNs. We determined that CCR5 cleavage activity varied with the target site, guide RNA length, and the terminal nucleotide in the PAM sequence. Our analyses demonstrated a PlmCas12e PAM preference for purines (A, G) over pyrimidines (T, C) in the fourth position of the CasX2 PAM (TTCN). These analyses have contributed to a better understanding of CasX2 cleavage requirements and will position us more favorably to develop a therapeutic that creates the delta-32 mutation in the CCR5 gene in hematopoietic stem cells.

Genome editing in plants typically relies on T-DNA plasmids that are mobilized by Agrobacterium-mediated transformation to deliver the CRISPR/Cas9 machinery. Here, we introduce a series of CRISPR/Cas9 T-DNA vectors for minimal lab settings, such as in the classroom or citizen science projects. Spacer sequences targeting genes of interest can be inserted as annealed short oligonucleotides in a single straightforward cloning step. Fluorescent markers expressed in mature seeds enable reliable selection of transgenic as well as transgene-free individuals using a combination of inexpensive LED lamps and colored-glass alternative filters. Testing these tools on the Arabidopsis GROWTH-REGULATING FACTOR (GRF) gene family, we found that Cas9 expression from an EGG CELL1 (EC1) promoter resulted in tenfold lower mutation rates than expression from a UBIQUITIN10 (UBQ10) promoter. A collection of bona fide null mutations in all nine GRF genes could be established with little effort. Finally, we explored the effects of simultaneously targeting two, four and eight GRF genes on the rate of induced mutations at each target locus. Multiplexing caused strong interference effects: while mutation rates at some loci remained consistently high, mutation rates at other loci dropped dramatically with increasing number of single guide RNA species. Our results suggest potential detrimental genetic interaction between induced mutations as well as competition of CRISPR RNAs for a limiting amount of Cas9 apoprotein.

The use of CRISPR-associated enzymes in iPSC-derived neurons for precise gene targeting and high-throughput gene perturbation screens offers great potential but presents unique challenges compared to dividing cell lines. CRISPRi screens in iPSC-derived neurons and glia have already been successful in relating gene function to neurological phenotypes; however, loss of dCas9-KRAB expression after differentiation has been observed by many labs and has been largely ascribed to transgene silencing after differentiation. Here, we investigated the expression levels of different CRISPR enzymes in iPSC and Ngn2-derived neurons using piggybac delivery. We found that the commonly used dCas9-KRAB (using the KOX1 domain) displayed dramatic reduction in protein expression levels following neuronal differentiation, yet surprisingly, nCas9 constructs retained comparable protein expression between iPSCs and neurons. We further found that CRISPR constructs, primarily relying on the SV40 Nuclear Localization Signal (NLS), fail to efficiently localize to the nuclei of neurons, despite having robust nuclear levels in iPSCs, leading to KRAB-specific cytoplasmic degradation. By adding a neuronal-specific NLS, we were able to correct neuronal nuclear localization and protein expression, confirming the contribution of mislocalization to the instability of dCas9-KRAB in neurons. As the lack of nuclear localization can have a profound impact on editing and gene perturbation efficiency, we suggest further investigation across both cultured and in-vivo postmitotic cell models.

CRISPR/Cas9 is currently the most powerful tool to generate mutations in plant genomes and more efficient tools are needed as the scale of experiments increases. In the model plant Arabidopsis, the choice of promoter driving Cas9 expression is critical to generate germline mutations. Several optimal promoters have been reported. However, it is unclear which promoter is ideal as they have not been thoroughly tested side-by-side. Furthermore, most plant vectors still use one of the two Cas9 nuclear localization sequence (NLS) configurations initially reported and can still be optimized. We genotyped more than 6,000 Arabidopsis T2 plants to test seven promoters and eleven NLS architectures across 14 targets to systematically improve the generation of single and multiplex inheritable mutations. We find that the RPS5A promoter and double-BP NLS architecture were individually the most efficient components. When combined, 99% of T2 plant contained at least one knockout mutation and 84% contained 4-7-plex knock-outs. These optimizations will be useful to generate higher-order knockouts in the germline of Arabidopsis and likely be applicable to other CRISPR systems as well.

Despite many advantages over Cas9, Cas12a has not been widely used in genome editing in mammalian cells largely due to its strict requirement of the TTTV protospacer adjacent motif (PAM) sequence. Here, we report that Mb3Cas12a (Moraxella bovoculi AAX11_00205) could edit the genome in murine zygotes independent of TTTV PAM sequences and with minimal on-target mutations and close to 100% editing efficiency when crRNAs of 23nt spacers were used.\n\nSummary statementCRISPR-Mb3Cas12a can target a broader range of sequences in murine zygotes compared to AsCas12a and LbCas12a, and has lower on-target effects than Cas9 and high overall knock-in efficiency.

Recent advances in CRISPR technology have enabled us to perform gene knock-in in various species and cell lines. CRISPR-mediated knock-in requires donor DNA which serves as a template for homology-directed repair (HDR). For knock-in of short sequences or base substitutions, ssDNA donors are frequently used among various other forms of HDR donors, such as linear dsDNA. However, for insertion of long transgenes such as fluorescent reporters in human cells, the optimal type of HDR donors remains unclear. In this study, we established a simple and efficient CRISPR-mediated knock-in method for long transgenes using linear dsDNA and ssDNA donors, and systematically compared the performance of these two donors for endogenous gene tagging in human non-transformed diploid cells. Quantification using flow cytometry revealed higher efficiency of fluorescent tagging with dsDNA donors than with ssDNA. By analyzing knock-in outcomes using long-read amplicon sequencing and a classification framework, a variety of mis-integration events were detected regardless of the donor type. Importantly, the ratio of precise insertion was higher with dsDNA donors than with ssDNA. Moreover, in off-target integration analyses, dsDNA and ssDNA were comparably prone to non-homologous integration. These results indicate that ssDNA is not superior to dsDNA as long HDR donors for gene knock-in in human cells.

Cas-assisted lambda Red recombineering techniques have rapidly become a mainstay of bacterial genome editing. Such techniques have been used to construct both individual mutants and massive libraries to assess the effects of genomic changes. We have found that a commonly used Cas9-assisted editing method results in unintended mutations elsewhere in the genome in 26% of edited clones. The unintended mutations are frequently found over 200 kb from the intended edit site and even over 10 kb from potential off-target sites. We attribute the high frequency of unintended mutations to error-prone polymerases expressed in response to dsDNA breaks introduced at the edit site. Most unintended mutations occur in regulatory or coding regions and thus may have phenotypic effects. Our findings highlight the risks associated with genome editing techniques involving dsDNA breaks in E. coli and likely other bacteria and emphasize the importance of sequencing the genomes of edited cells to ensure the absence of unintended mutations. GRAPHICAL ABSTRACT O_FIG O_LINKSMALLFIG WIDTH=200 HEIGHT=61 SRC="FIGDIR/small/584922v1_ufig1.gif" ALT="Figure 1"> View larger version (15K): org.highwire.dtl.DTLVardef@476263org.highwire.dtl.DTLVardef@8c891eorg.highwire.dtl.DTLVardef@7e32b4org.highwire.dtl.DTLVardef@132d3fc_HPS_FORMAT_FIGEXP M_FIG C_FIG

RNA-targeting CRISPR-Cas systems enable modulation of gene expression without permanent genome modification, making them useful for sensitive cell types such as neurons. While CRISPR-Cas technologies have been most extensively applied and validated in primary hippocampal and cortical neurons, their use in sensory neurons remains largely unexplored. Sensory neurons are an established cellular model for studying axon growth and regeneration, pain mechanisms, sensory transduction, and neuron-environment interactions. Here, we evaluated the performance of compact RNA-targeting CRISPR-Cas effectors Cas7-11S, hfCas13X, and hfCas13d in primary rat sensory neurons in culture. Using an endogenous mRNA as the target, we compared knockdown efficiency and assessed the effects of CRISPR-Cas expression on neuronal health. The systems showed distinct differences in performance, with Cas7-11S inducing toxicity, hfCas13X showing minimal knockdown, and hfCas13d providing robust gene silencing with minimal adverse effects on neuronal health. These findings identify hfCas13d as an effective and well-tolerated RNA-targeting CRISPR-Cas tool for sensory neurons and provide important insight into its suitability for neuroscience research and potential therapeutic applications.

The CRISPR-Cas technology has sparked a new technological revolution, significantly enhancing our ability to understand and engineer organisms. The nuclease that underpins this technology is evolving from the "One Cas9 for all" model to a diverse CRISPR toolbox. Identifying PAM sequences is a critical bottleneck in developing novel Cas proteins. Given the limitations of experimental methods, bioinformatics approaches are essential for predicting PAM sequences of Cas proteins in advance. To date, there are only a few PAM sequence prediction programs, and their accuracy is relatively low due to the limited number of spacers in CRISPR-Cas systems. To overcome this challenge, we have developed a pipeline named PAMPHLET, which innovatively utilizes homology searches of Cas proteins to identify additional spacers. PAMPHLET was tested on 20 CRISPR-Cas systems with known PAMs, increasing the number of spacers by up to 18-fold compared to the original datasets and successfully predicting 18 PAM sequences for protospacers. For rigorous and high-quality wet-lab validation of the predictions made by PAMPHLET, we employed the published DocMF platform. This platform leverages next-generation sequencing chips to profile protein-DNA interactions and can simultaneously screen both 5 and 3 PAMs with high throughput. The PAMPHLET predictions showed high consistency with the DocMF results for four novel Cas proteins. We expect that PAMPHLET will overcome the current limitations in PAM sequence prediction, expedite the discovery of PAM sequences, and help to shorten the development cycle for CRISPR tools. Remarkably, PAMPHLET has revealed an intriguing genomic phenomenon: the C2c9 and C2c10 systems, which lack the canonical adaptation module, possess identical PAM sequences to those found in co-occurring type I systems, suggesting potential shared spacer acquisition mechanisms. This finding highlights the complex evolutionary relationships of CRISPR-Cas systems and propels us toward a deeper understanding of their mechanistic diversity and adaptability.

Type I-E CRISPR-Cas3 derived from Escherichia coli (Eco CRISPR-Cas3) can introduce large deletions in target sites and is available for mammalian genome editing. The use of Eco CRISPR-Cas3 to plants is challenging because 7 CRISPR-Cas3 components (6 Cas proteins and CRISPR RNA) must be expressed simultaneously in plant cells. To date, application has been limited to maize protoplasts, and no mutant plants have been produced. In this study, we developed a genome editing system in rice using Eco CRISPR-Cas3 via Agrobacterium-mediated transformation. Deletions in the target gene were detected in 39-48% and 55-71% of calli transformed with 2 binary vectors carrying 7 expression cassettes of Eco CRISPR-Cas3 components and a compact all-in-one vector carrying 3 expression cassettes of Cas proteins fused to 2A self-cleavage peptide, respectively. The frequency of alleles lacking a region 7.0 kb upstream of the PAM sequence was estimated as 21-61% by quantifying copy number by droplet digital PCR, suggesting that mutant plants could be obtained with reasonably high frequency. Deletions were determined in plants regenerated from transformed calli and stably inherited to the progenies. Sequencing analysis showed that deletions of 0.1-7.2 kb were obtained, as reported previously in mammals. Interestingly, deletions separated by intervening fragments or with short insertion and inversion were also determined, suggesting the creation of novel alleles. Overall, Eco CRISPR-Cas3 could be a promising genome editing tool for gene knockout, gene deletion, and genome rearrangement in plants.

Single-guide RNA lentiviral infection with Cas9 protein electroporation (SLICE) enables CRISPR screening in primary cell types that require transient Cas9 expression yet is limited by scalability and robustness. Here, we introduce dual-guide RNA infection with Cas9 electroporation (DICE), which expresses two guides from the same lentiviral construct that target the same gene. In genome-wide screens, DICE outperformed SLICE in defining essential genes and modulators of PD-L1 expression in IFN gamma activated THP1 cells. Collectively, these data demonstrate that DICE can be utilized for reduced-scale CRISPR screens in cell types with transient Cas9 protein expression without sacrificing screening quality.

While genome editing has been revolutionized by the advent of CRISPR-based nucleases, difficulties in achieving efficient, nuclease-mediated, homology-directed repair (HDR) still limit many applications. Commonly used DNA donors such as plasmids suffer from low HDR efficiencies in many cell types, as well as integration at unintended sites. In contrast, single-stranded DNA (ssDNA) donors can produce efficient HDR with minimal off-target integration. Here, we describe the use of ssDNA phage to efficiently and inexpensively produce long circular ssDNA (cssDNA) donors. These cssDNA donors serve as efficient HDR templates when used with Cas9 or Cas12a, with integration frequencies superior to linear ssDNA (lssDNA) donors. To evaluate the relative efficiencies of imprecise and precise repair for a suite of different Cas9 or Cas12a nucleases, we have developed a modified Traffic Light Reporter (TLR) system [TLR-Multi-Cas Variant 1 (MCV1)] that permits side-by-side comparisons of different nuclease systems. We used this system to assess editing and HDR efficiencies of different nuclease platforms with distinct DNA donor types. We then extended the analysis of DNA donor types to evaluate efficiencies of fluorescent tag knock-ins at endogenous sites in HEK293T and K562 cells. Our results show that cssDNA templates produce efficient and robust insertion of reporter tags. Targeting efficiency is high, allowing production of biallelic integrants using cssDNA donors. cssDNA donors also outcompete lssDNA donors in template-driven repair at the target site. These data demonstrate that circular donors provide an efficient, cost-effective method to achieve knock-ins in mammalian cell lines.

CRISPR interference (CRISPRi) has emerged as a versatile approach for targeted gene repression in many organisms, including microbes and bacteria, due to the simple design of sequence-specific transcriptional silencing of gene expression. However, the strain-specific effects on repression efficiency and the host when translating a CRISPRi system from a laboratory strain to non-model strains are not well understood, yet they can present important limitations to its use. Here, we investigated the repression efficiency and toxicity of three CRISPRi systems (one dCas9 and two dCas12a variants) across four different Escherichia coli strains, including a laboratory K-12 strain (MG1655) and three non-model strains that are clinical isolates (probiotic Nissle 1917, uropathogenic CFT073, and uropathogenic UMN026). We evaluated the repression in each strain using sets of guide RNAs (gRNAs) targeting along the gene sequence and assayed cytotoxicity of expressing each dCas protein. Growth toxicity from expression of the different dCas proteins notably differed and showed high variation between some host strains. We also observed variable repression among the strains and notably poorer repression in multiple clinical strains. Therefore, we developed a dual gRNA CRISPRi system for enhanced gene silencing among the strains, which achieved up to 824-fold repression in CFT073. The results demonstrate that strain-specific design considerations can arise when a CRISPRi genetic system is transferred to a closely related bacterial strain. These findings provide insight into the relationships between criteria used for CRISPRi genetic design and in vivo activity across non-model E. coli strains, providing guidelines for diverse applications of these tools.

Autism spectrum disorder (ASD) is a highly heritable neurodevelopmental disorder associated with deficits in social communication and stereotypical behaviors. Numerous ASD-related genetic mutations have been identified and genome editing methods have been developed but successful genome editing in the whole-brain scale to alleviate autistic-like behaviors in animal models has not been achieved. Here we report the development of a new CRISPR-mediated cytidine base editor (CBE) system, which converts C{middle dot}G base pairs to T{middle dot}A. We demonstrate the effectiveness of this system by targeting an ASD-associated de novo mutation in the MEF2C gene (c.104T>C, p.L35P). We constructed a Mef2c L35P knock-in mouse and observed that Mef2c L35P heterozygous mice displayed autistic-like behaviors, including deficits in social behaviors and repetitive behaviors. We programmed the CBE to edit the C{middle dot}G base pairs of the mutated Mef2c gene (c.104T>C, p.L35P) to T{middle dot}A base pairs and delivered it via a single dose intravenous injection of blood brain barrier (BBB)-crossing AAV-PHP.eB vector into the mouse brain. This treatment restored MEF2C protein levels and reversed impairments in social interactions and repetitive behaviors in Mef2c L35P heterozygous mice. Together, this work presents an in vivo gene editing strategy in which correcting a single nucleotide mutation in the whole-brain scale could be successfully achieved, further providing a new therapeutic framework for neurodevelopmental disorders.

Cas-mediated genome editing has enabled researchers to perform mutagenesis experiments with relative ease. Effective genome editing requires tools for guide RNA selection, off-target prediction, and genotyping assay design. While independent tools exist for these functions, there is still a need for a comprehensive platform to design, view, evaluate, store, and catalogue guides and their associated primers. The Finding Optimizing and Reporting Cas Targets (FORCAST) application integrates existing open source tools such as JBrowse, Primer3, BLAST, bwa, and Silica to create a complete allele design and quality assurance pipeline. FORCAST is a fully integrated software that allows researchers performing Cas-mediated genome editing to generate, visualize, store, and share information related to guides and their associated experimental parameters. It is available from a public GitHub repository and as a Docker image, for ease of installation and portability.