The CRISPR Journal

Protospacer adjacent motifs (PAMs) and target-adjacent motifs (TAMs) are essential for target recognition by CRISPR-Cas and TnpB nucleases. Here we present TAMIPAMI, an efficient experimental and computational framework for rapid PAM/TAM identification. TAMIPAMI requires only a single control library and Cas or TnpB-treated library, simplifying experimental design, reducing cost, and providing greater accessibility for users. The platform interprets sequencing data with interactive visualizations and introduces a novel algorithm that determines the minimal exact set of degenerate IUPAC sequences describing the observed PAM/TAM patterns. Using this approach, we accurately recovered canonical motifs for several nucleases, including SpCas9, LbCas12a, AsCas12a, BrCas12b, Cas12i1, and AmaTnpB. TAMIPAMI is available as both a web application and command-line tool, ultimately providing an accessible and efficient platform for PAM/TAM discovery and characterization across CRISPR and OMEGA systems.

Abstract/SummaryFIG4 deficiency is the cause of Charcot Marie Tooth type 4J, a neurological disorder characterized by enlarged lysosomes. Our CRISPR activation genome wide screen found that upregulation of PIKFYVE rescued the enlarged lysosome phenotype in cultured cells. To assess PIKFYVE upregulation treatment in vivo, we generated Fig4 deficient mice with CRISPR activation of Pikfyve in neurons. Pikfyve was increased 2 fold in whole brain of CRISPR activated mice. Pikfyve upregulation did not extend the 3 week survival of Fig4 deficient mice. Vacuolization of brain was not rescued. The data demonstrates that a 2 fold increase of Pikfyve is not sufficient to treat Fig4 deficiency. Further testing will be required to determine if a higher increase of Pikfyve can ameliorate the effects of FIG4 deficiency in vivo.

Cas9-based tools enable the introduction of genetic lesions to investigate DNA repair outcomes and edit the genome at disease-relevant loci. DNA double-strand breaks (DSBs) induced by CRISPR/Cas9 result in frequent aneuploidy and large deletions, revealing a repair deficiency in early human embryos and limiting the clinical application of this technology. Here we evaluated the DNA repair outcomes of DNA nicks and mismatches introduced using base editors in human embryos at two targets, PCSK9 and HBG. Editing was efficient and, unlike Cas9-induced DSBs, did not result in either chromosomal abnormalities or large deletions. Small insertions or deletions after base editing were rare, and off-target activity was dependent on the guide RNA. Delivering the base editor as a protein at fertilization or at the pronuclear stage allowed normal development to the blastocyst stage and the derivation of edited stem cell lines. In stark contrast, introduction of the editor as RNA resulted in early embryo arrest. Our results demonstrated that, unlike DSBs, DNA nicks and mismatches are efficiently repaired in human embryos, allowing specific on-target changes without genotoxic consequences.

Plant genetic transformation technologies are essential for functional genomics and genome engineering in plants. While transient expression systems offer a rapid alternative to stable transformation, existing platforms are often constrained by low efficiency, technical complexity, and limited scalability. Here, we developed AgroGem, an efficient Agrobacterium-mediated transient transformation system utilizing a geminiviral replicon-based T-DNA vector for Arabidopsis and Brassicaceae species. AgroGem significantly outperformed existing transient approaches, including AGROBEST and protoplast-based assays, in CRISPR-mediated editing efficiency. Moreover, AgroGem recapitulated the mutation spectra and chromatin accessibility-dependent editing patterns observed in stable transformation across both Cas9 and Cas12a systems, indicating that it captures genome editing outcomes in native chromatin contexts. Leveraging this capability, we performed high-resolution profiling of CRISPR-induced mutation outcomes across a panel of DNA repair mutants and identified distinct repair signatures, including unexpected roles for KU80 and XRCC4 in regulating non-homologous end joining (NHEJ). AgroGem also supported bimolecular fluorescence complementation assays for protein-protein interaction studies in Arabidopsis and was readily adapted to plate-based formats for high-throughput applications. Together, these results establish AgroGem as a robust, scalable, and versatile platform for genome editing, DNA repair analysis, and functional genetics in plants.

Targeted protein degradation using conditional degron tag (CDT) technology is a powerful method for rapidly degrading a protein of interest (POI) upon the addition of a degrader drug. A prerequisite for the temporally controlled degradation of an endogenous POI is the generation of homozygous knock-in cells with the degron tag integrated at either the N- or C-terminus of their gene loci. However, obtaining those homozygous knock-in cells often requires selecting many single-cell clones, as human cells typically exhibit low homology-directed repair (HDR) activities. Additionally, tagging a degron to an endogenous protein may inadvertently reduce protein expression, potentially affecting protein function even before the drug is administered. Here, we develop a method for generating degron-tagged knock-in cells that allows us to skip the laborious single-cell cloning. This method arose from our observation that most knock-in cells carry the degron tag only in one allele (heterozygous), while the other allele typically harbors a frameshift insertion/deletion. This observation allowed us to bypass the need for single-cell cloning. We validated our method by knocking in degron tags at the N-terminus of cytoplasmic dynein1 subunits or Adaptor Protein 2 (AP2) subunit. Our experiments confirmed the rapid degradation of these proteins and their functional inhibition in bulk cell populations. Additionally, to mitigate the reduced expression often associated with the degron tagging, we established a method to control expression levels by inserting a mini-promoter immediately upstream of the knock-in cassette. Our method simplifies the workflow for degron tag knock-ins and enhances the versatility of these valuable technologies.

CRISPR-Cas9 technology is a powerful tool extensively used for genome editing in mouse and many other species. Streptococcus pyogenes Cas9 efficiently cuts both alleles in mouse zygotes leaving many edited embryos without a functional protein that might be needed to sustain development, to survive postnatally or to reproduce, thus complicating its overwhelmingly advantageous use in making gene modifications. About 25% of mouse genes are essential for embryonic development and another 7% are necessary for fertility, thus for these genes it is desirable to maintain a functional allele to establish viable lines from CRISPR-Cas9 edited mouse embryos. However, exclusive monoallelic editing is challenging to achieve with current CRISPR methods. Controlling the activity of Cas9 in genome editing is an ongoing research field focused on developing new methods to curtail its damage caused by excess of on-target and off-target editing. In this study we describe a novel and a simple method, we termed das-CRISPR, for dCas allele sequestration in combination with CRISPR system, that allows monoallelic editing of targeted allele in mouse and in cultured cell lines. This method incorporates the use of a nuclease deficient deadCas9 (dCas9) present at higher levels than an active Cas9, both complexed with the same single guide RNA (sgRNA) sequence. We showed the delivery of the two proteins as ribonucleoprotein complexes (RNP) into mouse zygotes leads to the generation of viable and fertile mice carrying lethal mutations in an essential gene. We found that greater amounts of dCas9 RNPs bind and protect a target site while the lower amount of functional Cas9 RNPs accessed the unoccupied target site resulting in higher frequency of monoallelic gene editing, compared to using just Cas9 alone. We also showed this method can mitigate and control the activity of Cas9 in mouse NIH3T3 cells in culture to achieve monoallelic editing. This method is a versatile approach to controlling excessive Cas9 activity on-target and off-target both in vitro and in vivo.

Plant virus-based gRNA delivery systems offer a rapid alternative to stable transformation for CRISPR-mediated genome editing, but potyvirus-based platforms in Cas9-expressing plants are still underexplored. Here, we developed a turnip mosaic virus (TuMV)-based system for gRNA delivery in Cas9-expressing Nicotiana benthamiana and tested whether Csy4-mediated gRNA processing could improve editing efficiency. A TuMV construct carrying a gRNA targeting PHYTOENE DESATURASE (NbPDS) induced detectable editing in both infiltrated and systemic tissues, although editing frequencies were low. Incorporation of the bacterial endoribonuclease Csy4 increased editing efficiencies in the two NbPDS genes, raising editing in infiltrated leaves to 7.1-13.8% for NbPDSa and 7.6-23.0% for NbPDSb, while lower but reproducible editing was detectable in systemic leaves. The TuMV-Csy4 platform also supported editing of a second endogenous target, MAGNESIUM CHELATASE SUBUNIT H (NbChlH), and enabled multiplex editing of NbPDS and NbChlH regardless of guide order. Editing efficiencies were consistently higher in infiltrated leaves than in systemic leaves, and no visible photobleaching or chlorosis was observed in systemic tissues despite confirmed molecular editing. To assess the potential for heritable editing, a tRNAIle mobility element was fused to the NbPDS gRNA. Although this construct increased somatic editing, no albino progeny were recovered after screening approximately 20,000 seedlings, indicating that heritable editing was not achieved under these conditions. Together, these results establish TuMV as a platform for Cas9-based gRNA delivery and show that Csy4-mediated processing improves editing efficiency, supports multiplex targeting, and demonstrates the feasibility of potyvirus-based genome editing systems in plants.

Homing endonuclease genes spread by cleaving homologous chromosomes that lack the endonuclease cassette, after which repair from the endonuclease-containing chromosome converts the cut allele into a copy of the drive allele. This mechanism has provided a conceptual foundation for synthetic gene drive systems, including CRISPR-based drives, that represent promising strategies for the genetic control of insect pests. However gene drive performance depends critically on the repair pathways available in the germline of the target organism. Here, we report a set of transgenic assays originally developed as part of an attempt to establish gene targeting in the malaria mosquito Anopheles gambiae using an in vivo-generated linear targeting molecule. Although the intended FLP-mediated excision step was not achieved in the mosquito germline, analysis of the component strains revealed efficient germline activity of the rare-cutting homing endonuclease I-SceI and a striking bias towards homology-based repair of I-SceI-induced double-strand breaks. Across reporter and donor configurations, cleavage outcomes were dominated by single-strand annealing, microhomology-mediated repair, synthesis-dependent strand annealing and gene conversion-like events, with comparatively limited evidence for classical non-homologous end joining. In reciprocal crosses designed to distinguish gene conversion from gamete loss, I-SceI cleavage also produced inheritance distortion consistent with both conversion of the cleaved allele and reduced recovery of gametes carrying extensively damaged donor alleles. These findings indicate that the An. gambiae germline can strongly favour homology-dependent repair following homing endonuclease cleavage and that cleavage can also generate meiotic drive-like distortion through selective loss of damaged gametes. The results have direct relevance for the design and interpretation of homing endonuclease and CRISPR-based gene drives in malaria mosquitoes, where the balance between homology-directed repair, end joining and gamete viability will determine drive efficiency, resistance formation and transmission bias. Author summaryGene drives depend on a simple but demanding principle: a nuclease cuts one chromosome, and the cell repairs the break using the homologous chromosome as a template, copying the drive element in the process. Before CRISPR, this type of system was explored using naturally occurring homing endonucleases such as I-SceI. We attempted to develop a gene targeting system in Anopheles gambiae based on the Rong and Golic strategy, in which FLP recombinase would excise a donor molecule and I-SceI would linearise it to stimulate recombination. The full knockout technology did not work because FLP-mediated excision was not detected in the mosquito germline. However, the component tests revealed something more broadly important: I-SceI-induced breaks were repaired predominantly through homology-based pathways rather than simple end joining. We also observed inheritance distortion consistent with both gene conversion and loss of damaged gametes. These results help explain why homing-based systems can work in mosquitoes, while also highlighting why repair pathway choice and gamete viability need to be measured directly in any new drive configuration.

Cas9 precision editing is increasingly predictable because guide, donor and target-context effects have been systematically characterized. Extending this framework to other nucleases is essential for installing variants outside convenient Cas9 target space. Cas12a provides a T-rich protospacer-adjacent motif (PAM) alternative, but determinants of efficient donor-templated Cas12a editing remain poorly defined. Here, we systematically dissected Cas12a precision editing in Saccharomyces cerevisiae across nuclease, direct repeat, expression, crRNA, donor, genomic context and time-course variables. Reporter and amplicon-sequencing assays showed that cleavage activity alone did not predict precise editing. Highly active configurations often reduced viability or lost edited alleles over time, whereas attenuated configurations better preserved programmed edits. Enhanced AsCas12a edited rapidly and tolerated shorter crRNAs, resulting in a narrower editing window, while an attenuated FnCas12a configuration edited more slowly but maintained higher viability and better distal-edit recovery. Alternative repair outcomes were rare, target-dependent, and further suppressed by LexA-FHA donor recruitment. To define design parameters at scale, we established a pooled Cas12a platform with 530 barcoded edit cassettes and recovered programmed edits for 70.2% of designs. Successful editing was reduced with TTTG PAMs, a C upstream of the PAM and at distal edit positions. Excluding these features increased the edited fraction to 85.4% and adding high predicted cleavage scores further elevated it to 91.4%. Applied retrospectively, these criteria also identified poorly edited loci in the targeted panels. Together, these data define design principles for Cas12a-mediated precision editing and establish a scalable platform for genome-scale pooled variant engineering and phenotyping in yeast.

Polygenic traits require the coordinated effects of multiple genes. Such complex traits have been a long-term target of study for geneticists, but multiplex CRISPR--the editing of multiple loci in the genome via multiple guide RNAs--is in its infancy. Reviewing 106 plant studies using multiplex CRISPR, we find that the multiplexing capacity has doubled every 5.4 years; furthermore, a systematic experiment with 8, 16, and 24 simultaneous targets in Arabidopsis thaliana reveals that efficiency of 24-plex editing can reach up to 73% across over one hundred third-generation transformed plants sequenced. Our experiment revealed that the level of multiplexing, or the number of the targets, causes minor efficiency reduction compared to the other uncontrolled factors such as gRNA design or variation across plants. When we model the decay in editing efficiency as a function of the gRNA number, actual efficiency is higher than the expectation from both Cas9 competition interference and simple joint editing stochasticity models. Rather, efficiency decayed with diminishing interference with more gRNAs with substantial overdispersion attributed to other efficiency factors, such as PAM identity. We predict that editing close to 100 genes in a plant can be feasible with reasonably large plant screens; however, feasible and reliable polygenic genome engineering will necessitate developments outside of [insert novelty of this study in how multiplex CRISPR was implemented, here]. Author ContributionsM.E.-A. conceived the project and secured funding. M.E.-A. and M.Es. designed the experimental strategy. M.Es. established the multiplex CRISPR and transformation pipelines in the laboratory, propagation through the T1 and T2 generations, and oversaw the first amplicon sequencing. Y.P. established the in-house iSeq amplicon sequencing protocol and contributed to cloning and genotyping pilots. M.Es supervised K.P. to construct cloning, bacterial transformations, plant growth, floral-dip transformations, and selection of T1 plants. E.K. contributed to early amplicon genotyping. J.K. propagated and sampled the T3 and J.K. and J.M. conducted the final amplicon sequencing panel. J.K. and M.E.-A. performed gene editing variant mapping, dataset quality control, and summarized results from published multiplex CRISPR studies. M.E.-A. modeled editing efficiency. J.K and M.E.-A. generated figures and wrote the first draft. All authors revised and improved the manuscript. J.K. and M.Es. contributed equally to this work and are designated as co-first authors.

Current cytosine base editors (CBEs) are limited to unidirectional C to T conversions, restricting their applications. Retrons, bacterial genetic elements, encode a reverse transcriptase that generates multicopy single-stranded DNA (msDNA) by reverse transcribing specific non-coding RNA (ncRNA). This msDNA mimics Okazaki fragments during DNA replication, making retrons promising for gene editing. Here, we developed a retron-based cytosine base editor (RCBE) by fusing cytosine deaminase with reverse transcriptase (RT-CDA) within the retron system. RCBE first transcribes ncRNA, allowing RT-CDA to deaminate cytosine on the ncRNA. The modified ncRNA is then reverse transcribed into msDNA, where RT-CDA induces further cytosine deamination. This mutant msDNA introduces specific mutations into target gene sequences, enabling both C to T and G to A conversions. Using RCBE, we demonstrated accelerated molecular evolution of the rpoB gene in Escherichia coli. High-throughput sequencing confirmed that RCBE achieves a mutation rate of up to 0.2% in regions with high GC content. Our findings establish RCBE as a versatile tool, particularly suitable for directed evolution in GC-rich regions, with broad potential applications across various bacterial and eukaryotic hosts.

The SCN1A gene encodes NaV1.1, a voltage-gated sodium channel protein that is necessary for neuronal excitability and whose loss-of-function mutations cause Dravet syndrome, a treatment-resistant childhood onset epilepsy. Gene replacement strategies for this syndrome are challenged by the large size of SCN1A and difficulty achieving stable cellular expression. Lentiviral vectors (LVVs) offer sufficient packaging capacity and genomic integration for defective SCN1A gene replacement. Here, we evaluated LVV-mediated delivery of different engineered SCN1A transgene sequences in human cells. LVV-transduced cells expressed full-length NaV1.1 protein that trafficked to the membrane and produced functional sodium currents. However, SCN1A transgene expression declined over time despite stable vector copy number, indicating post-integration regulatory limitations. Expression efficiency varied by SCN1A transgene sequence, with a codon-optimized variant showing higher expression despite lower LVV copy number. Treatment with sodium butyrate, a histone deacetylase inhibitor, significantly enhanced SCN1A transgene expression and partially rescued expression decay in a sequence-dependent manner. Incorporation of a ubiquitous chromatin opening element (UCOE) upstream of the promoter to maintain expression resulted in a trend of increased expression and increased responsiveness to butyrate. These findings demonstrate that sequence-specific and epigenetic factors may influence expression of large transgenes following lentiviral delivery, highlighting key challenges and design considerations for therapeutic SCN1A transgene expression.

With the expansion of therapeutic gene editing technology, small animal models provide essential platforms to evaluate the function of these new approaches in vivo. As part of the Somatic Cell Genome Editing (SCGE) Consortium, we developed next-generation murine reporters that overcome current model limitations and broaden detectable in vivo editing outcomes. These include two mouse models built on the "traffic light" reporter concept. This system enables fluorescent detection of both gene repair (green) and CRISPR-generated indels (red) events following editing by a single guide and either dsDNA or single-stranded oligonucleotide donor. We also generated a third reporter model that efficiently detects A-base editor activity. Reporters were validated in cultured embryos, via germline editing, and through activation in vivo by AAV transduction or direct ribonucleoprotein delivery. Together, these new models provide a valuable resource for improved detection of genome editing events in vivo.

Precision engineering of CRISPR/Cas9 components has advanced genome editing toward therapeutic applications. Completely chemically stabilized guide RNAs (gRNAs) have the potential to improve in vivo editing efficacy while enabling greater flexibility in delivery strategies. However, previous generations of fully modified guides have been associated with reduced Cas9 activity. Here, we employed an iterative, structure-guided optimization strategy to systematically introduce chemical modifications at each position of SpyCas9 gRNAs. Extending beyond commonly used nucleotide modifications, we incorporated 2-amino-RNA, 4-thio-RNA, and extended nucleic acid (exNA) to generate gRNA designs in which 90-100% of the nucleotides are sugar- or backbone-modified. Although certain modification patterns exhibit sequence-dependent variability, we have established a growing repertoire of guides that consistently maintain or enhance editing efficacy when applied both in vitro and in vivo. Collectively, our heavily and fully modified gRNAs hold potential for applications in nuclease editing, base editing, and other genome editing tools. Graphical abstract O_FIG O_LINKSMALLFIG WIDTH=200 HEIGHT=132 SRC="FIGDIR/small/725424v1_ufig1.gif" ALT="Figure 1"> View larger version (23K): org.highwire.dtl.DTLVardef@1118f08org.highwire.dtl.DTLVardef@1c57069org.highwire.dtl.DTLVardef@1572148org.highwire.dtl.DTLVardef@14a111e_HPS_FORMAT_FIGEXP M_FIG C_FIG

Bacillus thuringiensis (Bt) is a Gram-positive bacterium that during sporulation produces insecticidal Crystal (Cry) proteins, which play a major role in insect control today. Some Bt Cry proteins, e.g., Cry5Ba, target nematodes and, when given orally, can cure animals of gastrointestinal nematode (GIN) parasites. To eliminate concerns about treating humans and animals with spores and live bacteria, we developed an asporogenous system for scalable and safe Cry protein delivery called IBaCC (Inactivated Bacteria with Cytosolic Crystal(s)), which results in production of a bioactive crystal and a dead bacterium. However, to date, IBaCC involves expression of Cry proteins from antibiotic-selectable plasmids to ensure maintenance. Here, we develop and validate tools for markerless and stable integration and expression of Cry proteins in Bt. We markerlessly integrate an expression construct for Cry5Ba into either the spo0A or the sigK locus and demonstrate robust Cry5Ba expression. We also integrate our Cry5Ba expression construct into both loci simultaneously, increasing expression further. We demonstrate that an expression construct for a second anthelmintic Cry protein, Cry21Aa, can be integrated either alone or in combination with Cry5Ba in a single Bt strain. We furthermore show that these markerless integrants are stable in the absence of a selectable marker. These integrated strains, processed to IBaCC, demonstrate excellent ex vivo nematicidal bioactivity toward the larval stages of the sheep GIN parasite Haemonchus contortus and adult stages of the human hookworm GIN parasite Ancylostoma ceylanicum. This study demonstrates the successful markerless integration of 1-2 identical or dissimilar Cry proteins into Bt. These Cry integrants, in which genes essential to sporulation are deleted or replaced, provide robust Cry expression, stability, and bioactivity. These studies represent an important advance in Bt genetics and toward a safe, deployable, and cost-effective anthelmintic therapy to treat GIN parasitic infections in humans and animals.

TP53 is mutated in roughly half of all human cancers. Eight recurrent missense substitutions in the DNA-binding domain (R175H, Y220C, G245S, R248Q, R248W, R249S, R273H, R282W) account for most of the mutational burden. Homology-directed repair (HDR) with a wild-type donor template is one of the few feasible routes to revert these alleles, but existing CRISPR sgRNA design tools rank candidates without reference to the cancer cohort being treated. We built a reproducible pipeline that prioritizes SpCas9 sgRNAs for HDR-mediated correction of TP53 hotspot codons. The pipeline uses NM 000546.6 from NCBI, GRCh38 off-target search via Cas-OFFinder with the published Doench-2016 CFD matrices, on-target Doench-2016 (Rule Set 2) scores from CRISPOR, and per-cohort hotspot prevalence from three TCGA Pan-Cancer Atlas studies (HGSOC, n = 523; PDAC, n = 179; CRC, n = 534) accessed through cBioPortal. We enumerate guides whose cut sites fall within {+/-}10 nt of each hotspot codon, exclude any candidate that fails to map to GRCh38, and score the remainder. The final set contains 21 SpCas9 NGG sgRNAs across the seven hotspots, with no PAM-desert residues. A single candidate at R248 (TP53-248-P-ad878223; spacer GCATGGGCGGCATGAACCGG, AGG PAM; off-target specificity 0.913 over 806 reference-genome hits) ranks first in all three cohorts and holds rank 1 in 97% of 147 weight settings tested. Four additional residues (R175, Y220, R273, R282) yield within-residue tier-1 picks robust in 100% of weight settings. Cohort-specific differences appear only in cross-residue ordering: R175 and R282 climb in CRC, consistent with the higher prevalence of R175H and R282W in colorectal tumors.

CRISPR interference screens use catalytically inactive dCas9 fused to a repressor domain to enable genetic perturbations at the transcriptomic level. Interpretation of results involves identification of guide RNAs associated with the screen phenotype, followed by secondary analysis. During validation of a genetic screen, we observed different phenotypes from non-overlapping guide RNAs targeting one gene. Here, we developed POCKET-seq to map the binding of dCas9 genome-wide. We show that off-target binding occurs frequently and can generate false-positive interactions when it occurs near the promoter of genes associated with the screen phenotype. POCKET-seq classifies these false-positive and true-positive interactions using gene ontology.

Reported CRISPR/SpCas9 editing efficiencies in crops span 0-100%, but no quantitative synthesis has separated taxonomic family from delivery method, ploidy, clustering or publication bias. This meta-analysis estimated pooled per-T0-line editing efficiency across Cucurbitaceae, Brassicaceae, Solanaceae and Poaceae, and tested whether family is an independent moderator after adjustment for delivery and ploidy. A PRISMA 2020 systematic review identified peer-reviewed studies using SpCas9 with extractable per-line T0 edit counts; data were extracted independently by two reviewers, with inter-rater agreement reported. Logit proportions were synthesised with a binomial-normal generalised linear mixed model, and the family-as-moderator hypothesis was tested by a small-sample CR2 cluster-robust F-test on a three-level model with study-level clustering. Publication bias was assessed by Eggers regression and trim-and-fill. Twenty-two studies contributed 172 per-line effect sizes (Cucurbitaceae k=14, Brassicaceae k=20, Solanaceae k=68, Poaceae k=70). Pooled editing efficiency was 61.8% (95% CI 54.5-68.6%) with I{superscript 2}=93.4% ({tau}{superscript 2}=3.21) and a 95% prediction interval of approximately 5-98%. Per-family estimates ranged from 47.8% (Poaceae) to 73.8% (Brassicaceae); the univariate Q test was significant (p=0.0016), but family did not survive cluster-robust adjustment (F=0.73, p=0.63). Intraclass correlation placed 64.4% of variance at the study level, and Solanaceae remained dominated by a single study (58/68 rows). Funnel asymmetry was severe (Egger p<0.0001), and trim-and-fill reduced the bias-adjusted estimate to 45.2% (95% CI 39.0-51.5%). Apparent crop-family differences dissolve once within-study clustering and methodological covariates are accounted for; the bias-adjusted pooled estimate is closer to 45% than to 62%, and reported editing efficiencies reflect study-level factors more than taxonomic family. Key MessageApparent between-family differences in CRISPR/SpCas9 editing efficiency across four crop families reflect within-study clustering and publication bias, not intrinsic biology; family is not an independent moderator after cluster-robust adjustment.

Drosophila suzukii (Matsumura, 1931, Diptera: Drosophilidae) is a globally invasive pest of soft-skinned fruits that is currently controlled largely through the use of broad-spectrum insecticides. Increasing resistance to pesticides and regulatory pressures have motivated the development of genetic control strategies. We previously developed a CRISPR/Cas9-based homing gene drive targeting the coding sequence of the female-specific exon of the sex-determination gene doublesex, achieving highly efficient inheritance (94-99%) in both male and female germlines. A major limitation of homing gene drives is the formation of resistant alleles that evade cleavage yet retain gene function. Multiplexing guide RNAs (gRNAs) could reduce the formation of such functional resistance alleles. Here, we generated and tested homing constructs expressing one, two, or three gRNAs targeting different regions of the female-specific exon of doublesex, including a splice-junction target site. A single gRNA targeting the splice junction supported high inheritance in males but showed reduced efficiency in females. Combining this gRNA with a coding sequence-targeting guide further reduced drive efficiency, particularly in the female germline. Constructs expressing two gRNAs performed similarly whether guides were linked by transfer RNA (tRNA) sequences or expressed from independent promoters. Constructs expressing three gRNAs using tRNA processing showed consistently low drive inheritance in both sexes, likely a consequence of reduced cleavage efficiency due to inefficient gRNA production. Inheritance was significantly higher in male than female germlines for several constructs, indicating that germline context strongly influences drive performance. Our findings highlight that the approach used for multi-gRNA expression, target site choice and sex-specific germline environments can impact gene drive efficiency, and emphasize the need to optimize construct design within the biological context of the target species.

Multidrug-resistant pathogens pose a major threat to One Health. Within the past decade, CRISPR-Cas systems have been explored as sequence-specific antimicrobials. While chromosomal injury has been considered the primary mechanism underlying pathogen killing by CRISPR-Cas antimicrobials, the synergistic role of gene disruption together with chromosomal injuries remains poorly understood. In this study, we characterized a new class of CRISPR-Cas antimicrobials that simultaneously cotarget essential and defensive genes to enhance potency against the clinically relevant pathogen Staphylococcus aureus. High-throughput CRISPR screening identified top-performing guide RNAs for twenty functionally diverse essential and defensive genes across the S. aureus genome. CRISPR-Cas antimicrobials were modularly formulated to target single or multiple gene loci and packaged in phage-like particles for specific delivery. By engineering an S. aureus production host with a chromosomally integrated anti-CRISPR protein, we demonstrated efficient production of CRISPR-Cas antimicrobials targeting any S. aureus chromosomal locus without self-targeting. Characterization of CRISPR-Cas antimicrobials with single guide RNA designs revealed that potency varied according to targeted gene function, achieving up to a 4-log10 reduction in viability and outperforming traditional antibiotics. Multiplexed configurations were consistently more effective than single-targeting designs, with the top-performing design demonstrating a 4.7-log10 reduction in viability. Cotargeting essential and defensive genes revealed synergies that led to improved lethality and attenuated resistance, with enhanced activity in biofilms compared to traditional antibiotics. Genes involved in signaling and stress responses were important defensive targets for developing cotargeting CRISPR-Cas antimicrobials. Overall, this study establishes design principles for synergistic CRISPR-Cas antimicrobials applicable to next-generation precision antimicrobial development. SIGNIFICANCEThe ability to effectively combat multidrug-resistant pathogens is of primary importance to One Health. This study develops a generalizable design principle for formulating potent CRISPR-Cas antimicrobials that exploit synergistic cotargeting strategies for enhanced pathogen killing. In addition to chromosomal injuries, we found that disruption of gene function plays a crucial role in determining the lethality of CRISPR-Cas antimicrobials, providing a generalizable framework for effective CRISPR-Cas antimicrobial design. The development of a CRISPR-Cas antimicrobial production host with stable, chromosomally integrated anti-CRISPR genes greatly expands the modularity, adaptability, and efficiency of formulating CRISPR-Cas antimicrobials and enables deeper insights into the molecular mechanisms involved in eliminating multidrug-resistant pathogens.