The CRISPR Journal

Pooled genome-wide CRISPR-Cas9 knockout (CRISPR-KO) screening is a powerful approach for discovering new biology and identifying genetic vulnerabilities in cancers. This approach uses the Cas9 nuclease in combination with sgRNA libraries, typically consisting of 4-8 sgRNAs to induce mutations in each target gene. A critical assumption is that the effect of each sgRNA is solely due to Cas9 editing of the target gene. However, libraries can contain sgRNAs that direct Cas9 to multiple locations, thus potentially introducing bias into gene hit lists and leading to flawed biological hypotheses. Here we have developed GuideRefine, a pipeline to detect multi-targeting and off-targeting sgRNAs. GuideRefine outputs a virtual refined sub-library containing only on-target sgRNAs. Using GuideRefine with T2T-CHM13 as the reference genome, we surveyed the Brunello, TKOv3, Yusa, Avana, and Jacquere libraries, finding that ~7.5% to ~16% of sgRNAs are potentially problematic. We confirmed that multi-targeting sgRNAs disproportionately impair cell fitness and that sgRNAs aligning to more than one location with a single mismatch can also reduce fitness, although to a lesser extent. After flagging problematic sgRNAs and creating virtual ''on-target only'' sub-libraries, ~10% to ~16% of genes lose critical representation (< 3 sgRNAs per gene). Intriguingly, a set of 467 genes, characterised by short CDS length and lower PAM site density, have fewer than three sgRNAs in all sub-libraries, suggesting they cannot be well-targeted using current libraries. We anticipate that GuideRefine, together with caution in assessing the effects of problematic sgRNAs, will help prioritise biologically relevant hits.

The CRISPR/Cas12a system has emerged as a highly useful tool for genomic editing due to its ability to process multiple guide RNAs (gRNAs) from a single promoter. In this study, we explored the potential of combinatorial CRISPR/Cas12a to excise targeted loci in the mammalian genome by inducing double-strand breaks (DSBs) upstream and downstream of targeted exons. We designed a library of gRNA pairs targeting introns flanking 54 exons of 18 essential genes and 17 exons of 10 nonessential genes, aiming to assess the efficiency of Cas12 in excising targeted loci. While single gRNAs directly targeting essential exons produced expected fitness defects in cells, pairs of guides targeting flanking intronic regions resulted in only trivial viability effects, suggesting that excision rarely occurred. Further analysis showed no meaningful correlation between gRNA efficiency, orientation, GC content, or cut site geometry and the success of locus excision. These findings indicate that, under the conditions tested, the Cas12a system does not yield locus excision at the efficiency required for sensitive negative selection assays.

Genome-edited mice are widely used to elucidate molecular mechanisms in vivo and are an indispensable tool, particularly for studies aimed at clarifying gene function at the organismal level. Currently, there is an increasing demand for mice in which multiple genes are simultaneously modified in order to investigate interactions among multiple genes. Notably, generating conditional multiple-gene knockout mice with temporal and spatial specificity requires extensive crossing between multiple Cre-driver mice and floxed mice, resulting in a prolonged time frame for line establishment. To address this limitation, we developed Transposon-Enhanced Multi-Plex Orchestration Editing (TEMPO-editing), a single-step strategy for generating multiple-gene-edited mice. TEMPO-editing enables simultaneous modification of multiple genes through the integration of transposon, Cre-loxP, and CRISPR/Cas9 systems. Using the DNA transposon piggyBac, we constructed a single cassette harboring gRNAs targeting genes of interest together with a conditionally expressed Cas9 (lsl-Cas9). By injecting this cassette into fertilized eggs of Cre mice, we enabled the generation of temporally and spatially specific genome edited mice in the F0 generation. In this study, we generated double-gene-edited mice targeting Hoxa13 and Hoxd13, which are key regulators of embryonic body patterning and are essential for autopod development. The phenotype observed in these mice was consistent with the incomplete autopod phenotype previously reported in mice generated by crossing Hoxa13 knockout and Hoxd13 knockout mice. These results demonstrate the utility of TEMPO-editing for the generation of multiple-gene-edited mice in the F0 generation. Notably, this study establishes a simplified strategy for producing conditional multi-gene-edited mice, a process that has traditionally required substantial time and labor using conventional methods.

CRISPR/Cas9-mediated homology-directed precise genome editing using long single-stranded DNA (ssDNA) donors has expanded the possibilities for generating defined genetic modifications. However, the required ssDNA donor preparation can be technically demanding and often requires extensive locus-specific sequence design. Here, we examined parameters influencing ssDNA donor-mediated genome editing and developed approaches to simplify donor preparation using {lambda}-exonuclease-mediated ssDNA generation. By evaluating donor designs across multiple genomic loci, we found that efficient genome editing can be achieved with relatively short homology regions for a range of insertion sizes. These findings provide a basis for simplifying ssDNA donor preparation and the overall gene-editing pipeline, potentially facilitating its application across species.

Precise temporal control of gene editing is essential for studying dynamic biological processes, interrogating essential gene function, and improving the interpretability of pooled perturbation screens. Cre-dependent single guide RNA (sgRNA) switches provide temporal regulation by coupling guide activation to site-specific recombination, but existing designs retain a loxP-derived 5' sequence (scar) on the mature sgRNA that can impair guide function. We developed a scarless conditional sgRNA platform that combines Cre-loxP recombination with endogenous RNA processing to restore the native sgRNA architecture following induction. A MALAT1-associated small cytoplasmic RNA (mascRNA) module was positioned upstream of the guide sequence such that, after Cre-mediated recombination, cellular RNase P and RNase Z remove the residual loxP-derived overhang, generating a mature sgRNA with an authentic 5' terminus. Using guides targeting endogenous cell-surface marker genes, the scarless design maintained stringent OFF-state control while improving ON-state editing performance compared with a conventional Cre-activated sgRNA switch, resulting in faster editing kinetics, greater perturbation penetrance, and more consistent editing efficiency. This modular strategy provides a simple approach for conditional CRISPR genome editing that preserves guide integrity and should be readily adaptable to time-resolved functional genomics and pooled screening applications.

Genome editing has revolutionized biomedical sciences and is now an essential tool to define molecular pathways through genetic interaction and loss-of-function studies. Through its diverse variations, it allows for the generation of specific knockout cell lines or organisms, as well as the creation of endogenously edited gene regions. While high-throughput methodologies exist to map CRISPR/Cas9 genetic modifications, the validation of guide efficiencies in cell populations is often performed through analysis of the targeted gene product by western blotting or by deconvolution of Sanger sequencing chromatograms using TIDE or ICE assays. Here, we highlight a rapid nanopore sequencing pipeline, which we have named NanoCutSight, to quantify the percentage of indels at a specific genomic locus and to identify the types of modifications generated. We also benchmarked the methodology on various guide RNAs and in both cultured cell and organoid models. We believe that NanoCutSight will simplify the analysis of complex sample editing and enable the rapid screening of edited samples.

SummaryA catch-all intronic guide RNA pair excises the KIAA1549--BRAF oncofusion across its major variants, with productive junction excision confirmed by gain-of-function PCR in patient-derived glioma cells. An allele-specific guide selectively disrupts BRAF V600E, in patient-derived pediatric low-grade glioma cells. Pediatric low-grade glioma (pLGG) is the most common brain tumor of childhood, accounting for 30--50% of all pediatric central nervous system malignancies1. The disease is almost universally driven by activating mutations in the BRAF serine/threonine kinase: a chromosomal tandem duplication generating the KIAA1549--BRAF oncofusion in approximately 70% of cases, or the BRAF V600E gain-of-function point mutation in approximately 15%2. Current targeted pharmacotherapies, including the RAF inhibitor tovorafenib, require continuous dosing, are not allele-specific, and carry risks of long-term toxicity in children. A one-time genomic intervention that permanently disables the oncogenic BRAF alteration while preserving wild-type BRAF signaling represents a compelling therapeutic alternative. In this study, we describe the design and experimental validation of allele-specific CRISPR guide RNAs targeting both the KIAA1549--BRAF oncofusion and the BRAF V600E point mutation. For the oncofusion, we developed a double-cut intronic excision strategy in which a guide RNA targeting KIAA1549 intron 14 is paired with a guide RNA targeting BRAF intron 11. Because the genomic breakpoints of all four major fusion variants (KB 16:9, 15:9, 16:11, and 15:11) fall within these introns, a single guide pair can address the full landscape of fusion heterogeneity in a single intervention. For BRAF V600E, we exploited a unique PAM sequence created by the pathogenic TBA transversion at codon 600, enabling allele-specific SpCas9 and AsCas12a guide designs that distinguish the mutant from the wild-type allele at single-nucleotide resolution. We screened guide RNA candidates by ribonucleoprotein (RNP) nucleofection in A375 human melanoma cells (BRAF V600E homozygous) and in patient-derived 3635 PXA glioma cells (BRAF V600E heterozygous). The top KIAA1549 intron 14 guide, K9_i14_A_Cas9, achieved 66% indel frequency in A375 cells. The top BRAF intron 11 guides, B_i11_A_Cas9 and B_i11_D_Cas9, achieved 84% and 85% indel frequency, respectively. For BRAF V600E, the best allele-specific SpCas9 guide achieved l57% editing in A375 cells and l74% editing in 3635 PXA patient-derived glioma cells. Dual-cut excision of the KIAA1549--BRAF junction was confirmed by a gain-of-function PCR assay designed to detect the excision junction amplicon ([~]191 bp) produced by NHEJ-mediated rejoining of the KIAA1549 intron 14 and BRAF intron 11 cut ends.

In the last decade, there has been an explosion in the development of in vivo germline genetic editing technologies in arthropods, first pioneered by the Receptor-Mediated Ovary Transduction of Cargo (ReMOT Control) method. While ReMOT Control and related technologies (DIPA-CRISPR and SYNCAS) have shown great utility for gene knock-out in many different species, consistent gene insertion has been elusive, limiting widespread adoption. We developed a novel system based on the baculovirus Autographa californica multiple nucleopolyhedrovirus (AcMNPV) to deliver heritable CRISPR knock-out and knock-in constructs to the arthropod germline by oral feeding. We call this method "Germline Engineering by Viral Transduction" ("GEValT"). With GEValT, newly-eclosed female and male adult arthropods are fed on virus in sugar (and/or blood) and allowed to breed, resulting in easy generation of genetically modified offspring. GeValT is easy to use and has the potential to revolutionize gene editing methodology for researchers working across diverse arthropod species.

RNA-guided nucleases (RGNs), such as Cas9 from Streptococcus pyogenes (SpCas9), are widely used for plant genome editing. Previous surveys for off-targeting, the modification of unintended targets with similarity to the intended target, indicate high specificity of SpCas9 in plant cells. However, off-targeting has not been assessed for efficiency-optimized editing systems combined with extensive multiplexing, which might increase the likelihood of cleavage at unintended sites. We therefore analyzed Arabidopsis thaliana lines that had been extensively mutagenized using zCas9i and up to 29 gRNAs addressing >45 target sites over several rounds of editing. Genomes were sequenced by short- and long-read technologies, and genome-wide variants were catalogued. Our pipeline for variant calling reliably detected RGN-induced mutations at on-targets. When excluding these on-target modifications, variants were detected in edited lines at frequencies similar to those previously reported for spontaneous mutations. In further analyses, we did not find any evidence for an origin of these variants from RGN activity. Our data are thus consistent with high specificity of SpCas9. In contrast, we detected genomic reorganization events upon editing at two complex loci, RPP1 and RPP7, encompassing multiple homologous genes, and also identified an allele by WGS that had escaped detection by amplicon sequencing. We conclude that, while off-targets may efficiently be avoided by selection of specific gRNAs, on-target modifications may be more extensive than intended, especially at complex loci and/or during multiplexing.

Genome engineering via CRISPR/Cas9 has been widely applied in biological research and developing gene therapies. However, genome editing flexibility is strongly constrained by the position and quality of guide RNAs relative to the intended edit: desired edit sites are frequently positioned far from efficient protospacer adjacent motifs (PAM). SMART (Silently Mutate And Repair Template) technique relaxes this constraint by silently mutating the gap between the Cas9 cut and the edit in the repair donor, preserving the encoded protein while expanding the usable guide-RNA design space. Designing such donors manually requires coordinated reasoning over transcript structure, reading frame, candidate protospacers, codon usage, homology arms and the expected edited allele. Auto-SMART is a web platform that automates this process for CRISPR/Cas9-mediated insertion, deletion, point mutation and sequence replacement in coding exons and in introns. Users choose a genome assembly, search a gene symbol or transcript, select a modification site and select an editing operation. Auto-SMART finds nearby Streptococcus pyogenes Cas9 (SpCas9) guides, ranks them by predicted on-target activity and cut-to-edit geometry, builds repair templates with synonymous substitutions, automatically recodes residual on-target sites to prevent re-cutting and renders interpretable panels for the target locus, donor template and locus after HDR, together with a downloadable report and a Primer3 genotyping handoff. The web application for Auto-SMART is freely available at https://auto-smart.chuanping.org/.

The dCas9 system has rapidly been developed into many tools to explore different aspects of the human genome; the high binding specificity, coupled with the inactive nuclease enzyme, allows for precise recruitment of molecules to specific sequences of DNA. We sought to exploit these capabilities to create a tool for assessing the real-time proximity of two DNA sequences within a biological system. By incorporating aptamers into gRNAs, dCas9 molecules can be used to recruit the {beta}9 or {beta}10 strands of split-NanoLuc(R) to specific DNA sequences and quantify the proximity of those sequences based on their ability to complex with the luciferase fragment ({Delta}11S) and produce luminescence. While many tools exist to detect a single DNA sequence, this system is uniquely capable of assessing how two DNA sequences interact with each other. As expected, we found that the interaction of two dCas9 molecules was affected by their linear distance from each other on dsDNA. Surprisingly, we found that their interaction was also strongly influenced by rotational orientation, even for sequences that are close together in linear space. This finding indicates that dCas9 rotational alignment is an important consideration for designing dCas9 systems that target multiple DNA sequences simultaneously. Beyond the findings presented herein, we believe this DNA proximity detection tool has the potential to be adapted for applications involving the proximity and orientation of two DNA sequences.

Compact Cas nucleases offer advantages over the widely used SpCas9 due to their smaller size, which enables more efficient delivery for in vivo applications. Among these, the phage-encoded Cas{Phi}2 (Cas12j2) is highly promising due to its relaxed PAM requirement (5-TTN-3) and compact size (757 aa); however, its translational potential is limited by low editing activity. To enhance the efficacy of Cas{Phi}2, we optimized the previously reported EPICA system, developing EPICA.2, a eukaryotic directed evolution platform to improve nucleases with nearly undetectable activity. EPICA.2 integrates additional yeast evolution rounds to enrich for active variants along with a low background mammalian reporter system that improves detection and selection of enhanced variants. Finally, we set up a long-read sequencing protocol which uses unique molecular identifiers (UMIs) to reduce sequencing errors, enabling accurate identification of the mutation combinations in each evolved variant. Among the most frequent variants, we obtained evoCas{Phi}2, which contains six activity-boosting mutations with a synergistic effect not predictable by rational engineering. Overall, evoCas{Phi}2 showed up to 70-fold increased activity in human cells compared to wild-type and outperformed variants generated through rational approaches, highlighting the potential of EPICA.2 as a powerful strategy to evolve genome editing tools with low native activity.

While directed differentiation of human pluripotent stem cells commonly relies on the timed delivery of extracellular factors, these uniform treatments often yield heterogeneous responses across cell populations. Linking intracellular gene regulation directly to an emerging developmental state offers a complementary strategy to coordinate these differentiation signals from within the cell. Here, we explored this approach by coupling a T/Brachyury promoter to CRISPR interference targeting CTNNB1, which encodes the canonical Wnt signaling mediator {beta}-catenin. A T-promoter EGFP reporter line exhibited transiently increased activity during early differentiation, supporting the use of this promoter as a developmentally responsive input. We then combined the promoter with dCas9-KRAB and a CTNNB1-targeting guide RNA. Cas9-mediated integration was accompanied by indels at the CTNNB1 target site, whereas a Cas12a-mediated integration strategy yielded clones with no indels detected by ICE analysis. During differentiation, the selected T-dCas9 CTNNB1 clone showed reduced CTNNB1 expression and attenuated induction of Wnt associated genes. Together, these findings provide a proof of concept for combining a developmental promoter with a programmable intracellular regulator and identify a strategy for separating circuit integration from unintended target-site editing. This modular approach provides a foundation for developing genetic interventions whose expression is linked to developmental state.

Degron tags enable rapid, tuneable and reversible control of protein expression, and have become a critical tool for target validation and elucidating biological mechanisms. Tag engineering uses genome editing to generate biallelic gene fusions, though this process remains laborious and requires new donor vectors to be generated for each locus. Here, we have generated a series of universal donor vectors to enable biallelic integration of different degron tags (AID, dTAG, BromoTag, SMASh) using different selection strategies (FACS, drug selection). Using this system, tags can be fused in-frame to endogenous protein C-termini via homology-independent repair following Cas9-induced double strand breaks, as shown here using human lymphoma cells, mouse embryonic stem cells and colonic tumour organoids. This workflow simplifies the genetic engineering of degron tags and is applicable in contexts where homology-dependent repair is challenging.

Antimicrobial resistance (AMR) is a growing public health threat, where drug-resistant Candida infections are associated with increased hospital costs, lengths of stay, and high mortality rates. Current diagnostic methods are slow and labor-intensive, causing delays in diagnosis and patient treatment. In this research, we propose a rapid CRISPR-based diagnostic for identifying Candida pathogens and antifungal resistance biomarkers in clinical applications. One-pot CRISPR assay designs are optimized for three clinically relevant Candida species, achieving attomolar sensitivity with synthetic DNA and validating performance on cultured isolates. Strategies are developed to engineer CRISPR gRNA to control one-pot reaction kinetics, enhancing discrimination of single nucleotide polymorphism (SNP) resistance mutations associated with azole and echinocandin antifungals. This research expands our understanding of the design of one-pot CRISPR reactions for sensitive detection of DNA, while establishing a foundation for faster, more precise AMR diagnostics to improve surveillance and outbreak management of fungal infections.

Virus-like particles (VLPs) enable transient, non-integrating delivery of CRISPR-Cas9 ribonucleoprotein cargo. Although VLPs have been reported for efficient DNA editing via base editors RNP delivery, the diversity of base editors tested as VLPs remains limited. We generated and benchmarked a panel of 12 base editors on the v5 eVLP backbone, targeting three genomic loci (HEK3, B2M, PDCD1) across five VLP dosages in LentiX-293T cells. Editing efficiency was generally dosage-dependent across all editors and varied by editor class and identity; PAM-flexible variants had lower editing efficiency than NGG-restricted counterparts, and the dual-function SPACE base editors showed reduced efficiency. We further characterized position-specific editing efficiencies and outcomes of the base editor VLP collection, revealing that a wide variety of mutation types are possible with the base editors in this collection.

CD8 T cells play a central role in immune responses to infection and cancer. However, the diversity of T cell receptor (TCR) specificities makes it challenging to study the mechanisms that regulate T cell activation, differentiation, and effector function. Beyond TCR transgenic mouse models, various complex genome-editing approaches have been employed to overcome this challenge. However, these strategies are often technically demanding, time-intensive, and difficult to adapt. Investigators who are interested in testing de novo TCRs under their chosen experimental conditions would benefit from a standardized and accessible method. Here, we describe a protocol that combines ribonucleoprotein (RNP)-based CRISPR-Cas9 editing with retroviral transduction to enable efficient genetic manipulation of murine CD8 T cells. We show that T cells engineered via this protocol can be generated at sufficient scale for downstream in vitro assays and in vivo adoptive transfer experiments. We expect this method will be useful for investigators who require a standardized and accessible way to study how TCR specificity impacts CD8 T cell responses.

Base editors act on the DNA strand displaced within an R-loop, and the set of positions they convert, the activity window, has been engineered for a decade on the assumption that linker length and attachment geometry determine it. We built a geometric model that predicts where a tethered deaminase acts from linker statistics, steric exclusion, and R-loop geometry alone; fitted three parameters to two previously reported profiles, held them fixed, and scored predictions against 50 architectures from seven studies, with each substrate coordinate withheld. Varying the contour length by a factor of 16 does not shift the predicted window at all, whereas changing the attachment site does: across 185 buildable single-linker designs at thirteen attachment sites, the predicted peak never leaves protospacer positions 3 to 12, positions 1, 2 and 13 to 20 are reached by no design, and transfer to Cas12a fails by five to six nucleotides in a way that localizes to the fusion junction rather than to reach, sterics or substrate. Tether geometry, therefore, bounds where a fused deaminase can act without predicting where it does so, making attachment site rather than linker length the effective design variable.

Background: DNA damage response (DDR) pathways are central regulators of genome maintenance and major determinants of cancer cell survival. The extensive genomic instability and high replicative stress that characterize glioblastoma render tumor cells highly dependent on DDR pathways to preserve genome integrity and sustain proliferation. This reliance creates potential therapeutic vulnerabilities, making the systematic identification of essential DDR genes a promising strategy for uncovering novel therapeutic targets. Methods: We developed DNA Damage Response KnockOut Library (DDRKOL), a custom CRISPR/Cas9 sgRNA library targeting 819 DDR genes with approximately 10 sgRNAs per gene, together with positive (essential), negative (non-essential) and non-targeting controls. Parallel depletion screens were performed in Cas9-expressing U87-MG and A172 cells cultured for 15 population doublings. Hits were prioritized utilizing TCGA and DepMap databases and validated by viability, clonogenic, apoptosis and GFP competition assays. Clinically relevant patient-derived glioblastoma spheroids and an orthotopic xenograft model was employed to characterize the effects of hit genes. Results: Sequencing confirmed near-complete recovery of the designed sgRNAs from the plasmid pool, with uniform representation across the library and complexity preserved through transduction and selection. Essential-gene controls depleted strongly while non-targeting controls remained neutral, confirming screen performance in both cell lines. The screens identified DDR dependencies in each line and defined a shared core composed of 20 genes belonging to homologous recombination, nucleotide excision repair and ATM/DSB signaling pathways. This shared dependency landscape highlighted four high-confidence candidate genes (TOP2A, CDK1, XRCC6, and RAD21), which were successfully validated across multiple orthogonal assays. These genes displayed grade-associated expression and their expressions were positively correlated with proliferation markers in TCGA. Individual knockouts reduced viability, colony formation and competitive fitness, induced apoptosis, and impaired growth of patient-derived glioblastoma spheroids. Both genetic depletion and pharmacological inhibition of TOP2A induced S/G2-M cell cycle arrest. In orthotopic xenografts, TOP2A depletion prevented tumor progression, and led to significantly prolonged survival. Conclusion: DDRKOL represents a robust and versatile focused CRISPR platform for systematic functional interrogation of the DDR associated genes. Using glioblastoma, we demonstrate that the library reliably identifies biologically significant and clinically relevant genetic dependencies through multiple orthogonal validation approaches. As a reusable platform rather than a disease-specific tool, DDRKOL can be broadly applied across diverse biological contexts to discover context-dependent DDR vulnerabilities, therapeutic targets, and mechanisms of treatment resistance.

The CRISPR-Cas12a system, primarily used for nuclear genome editing, operates through RNA-guided cleavage of target DNA. Given the high copy number of mitochondrial DNA (mtDNA) in cells and the pathogenic potential of mtDNA mutations, there is substantial interest in developing methods to selectively eliminate mutant mtDNA without affecting wild-type molecules. We suggest using AsCas12a, with its shorter CRISPR RNA (crRNA) and proven ability to be targeted into human mitochondria, to perform selective cleavage of mutant mtDNA. Here, we assess the potential of AsCas12a and various engineered crRNAs to discriminate between wild-type and mutant mtDNA in vitro and in living human cells. By exploring chemically synthesized full and split crRNAs with various modifications, we identified crRNAs capable of targeting specific mutant mtDNA. We demonstrated the strong influence of magnesium concentration on the ability of crRNAs to discriminate point mutations. Finally, we show that AsCas12a-fused cytosine base editor can be addressed to mitochondria and can direct low but significant specific base editing of mtDNA in human cells. Our study suggests AsCas12a as a promising tool for targeted mutant mtDNA elimination or editing.