The CRISPR Journal

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.

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.

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.

Efficient genome writing in mammalian cells requires robust methods for integrating large DNA payloads. The previously described method mammalian Switching Antibiotic resistance markers Progressively for Integration (mSwAP-In) enables iterative, biallelic genome rewriting in mammalian stem cells with DNA payloads exceeding 100 kb. However, the lack of standardized vectors and certain technical constraints have limited its broader adoption. Here we present an improved plasmid toolkit designed to streamline the implementation of mSwAP-In. The toolkit includes two core vectors. pLP-TK (pCTC174) is a landing-pad plasmid compatible with Golden Gate assembly of genomic homology arms and supports both mSwAP-In and the recombinase-mediated cassette exchange method Big-IN. mSwAP-In MC2v2 (pKBA135) is a versatile Big DNA assembly and delivery vector that supports Gibson-based assembly and incorporates positive, negative, and fluorescent selection markers, as well as a backbone counterselection cassette to minimize unwanted plasmid integration. The vector architecture also enables propagation in yeast and bacterial hosts, inducible plasmid copy-number amplification in standard E. coli strains, and CRISPR/Cas9-mediated payload release through preinstalled guide RNA target sites. We further characterize the FCU1/5-FC counterselection system in mouse embryonic stem cells and define conditions that minimize its bystander toxicity. Finally, we provide a set of Cas9-gRNA expression plasmids optimized for common mSwAP-In applications. Together, these reagents constitute a standardized and experimentally validated toolkit that simplifies large-scale genome writing using mSwAP-In.

Allele conversion describes a process where a heterozygous variant is made homozygous. Recently, it has been shown that allele conversion can be triggered by DNA damage at the heterozygous site. This process has the potential to repair pathogenic heterozygous mutations; however, the efficiency is low. Here, we endeavoured to understand the mechanism underlying allele conversion, ultimately to raise allele conversion efficiency to functionally relevant levels. To test this, we developed a Compound Heterozygous Allele Conversion Reporter (CHACR) cell line. This line comprises knocked-in fluorescent protein encoding genes, with heterozygous inactivating mutations resulting in different fluorescence profiles from each allele. These mutations create protospacer adjacent motifs (PAM) for Cas9 recognition, where allele-specific gRNAs (AS-gRNAs) target the heterozygous mutations. We showed that applying these AS-gRNAs with either Cas9 nuclease or Cas9(D10A) nickase can recover mCherry fluorescence. Sorting and sequencing these fluorescent cells revealed wild-type sequences, suggesting allele conversion repaired the mutation using the homologous allele as a template. Allele conversion can also be triggered using an adenine base editor with an AS-gRNA, and this allele conversion mechanism can be manipulated by inhibiting DNA-PKcs or overexpressing RAD51. This work introduces a model for measuring allele conversion, and modifiers of this mechanism.

In gene editing, CRISPR/Cas approaches are often limited by off-target effects. In in vivo approaches involving multiple cell types, off-targets may result from unintended targeting of the wrong cells. In this work, we propose a solution to this limitation by using a transcribed intron of the target gene as an endogenous trigger (intron triggers) for a novel conditional guide RNA (intcgRNA). In vitro, intcgRNAs were responsive to the presence of the trigger. As a proof-of-concept, the human IL2 receptor subunit gamma gene (IL2RG) was then targeted using both the intcgRNA and the corresponding conventional crRNA in two cell lines: the lymphocytic HPB-ALL cell line, where IL2RG is highly expressed, and the epithelial HeLa cell line, where it is not. Sanger sequencing revealed that the crRNA and intcgRNA Cas9 complexes edited IL2RG with similar efficiency in HPB-ALL, whereas only the crRNA edited IL2RG in HeLa. This shows that intcgRNA avoids targeting unwanted cells that do not express the target gene, which is particularly relevant for in vivo targeting. The triggers of choice for conditional guides have been microRNAs, but as short intronic RNAs are far more diverse, trigger introns could become biomarkers of cell identity that improve the precision of CRISPR-based manipulations in vivo. GRAPHICAL ABSTRACT O_FIG O_LINKSMALLFIG WIDTH=200 HEIGHT=83 SRC="FIGDIR/small/714022v1_ufig1.gif" ALT="Figure 1"> View larger version (17K): org.highwire.dtl.DTLVardef@1ae60cdorg.highwire.dtl.DTLVardef@1556c03org.highwire.dtl.DTLVardef@1264a0dorg.highwire.dtl.DTLVardef@c7d47d_HPS_FORMAT_FIGEXP M_FIG C_FIG

This protocol enables rapid CRISPR-Cas9 genome editing in Saccharomyces cerevisiae by replacing restriction/ligation guide cloning with PCR-based protospacer installation and seamless plasmid recircularization. It describes in silico HDR donor and SgRNA design, install guide sequences into cas9 plasmid by PCR and seamless assembly, plasmid cloning and sequence verification in E. coli, and LiAc/PEG co-transformation of yeast with Cas9-sgRNA plasmid plus HDR donor. The workflow selects yeast colonies on G418 and confirms edits by PCR and sequencing.

CRISPR/Cas9 enables precision gene editing via HDR for mutation correction and disease modelling. This protocol describes an 8-day non-viral HDR workflow for editing primary patient and healthy donor T cells, including reagent design, editing, on-target detection, and flow cytometry. The protocol was developed under research-grade conditions but supports scaling up and the transition to preclinical and clinical GMP workflows. For complete details on the use and execution of this protocol, please refer to Mamia et al.[1]. Before you beginCRISPR/Cas9 gene editing is a promising tool to correct pathogenic variants for autologous cell therapies, targeting monogenic diseases such as inborn errors of immunity (IEI). Furthermore, it can be used as a tool for disease modelling to study normal and pathological variations of the immune system. Here we present a detailed protocol for an efficient and customizable T cell single nucleotide variant (SNV) correction platform based on homology-directed repair (HDR). The protocol details every step of the process, which starts with custom CRISPR/Cas9 reagent design of guide-RNAs (gRNAs) and repair templates for editing a novel target with no previously published reagents. Furthermore, we describe the strategy of reagent design to assess on-target HDR editing using droplet digital PCR (ddPCR). Next, we detail the T cell platform itself, and present effective strategies to stimulate PBMCs ex vivo to promote CD4+ and CD8+ T cell activation and proliferation, which we have validated in 32 unique IEI patients. Next, we present the workflow of gene editing T cells using nucleofection and CRISPR ribonucleoprotein (RNP) complexes for efficient editing that preserves high cell viabilities and up to 80% HDR. Finally, we present a flow cytometry panel that assesses the immune cells present at the end of the platform, including characterization of memory and effector T cell populations and status of T cell exhaustion. InnovationIn the study, we present a detailed protocol of performing highly efficient, non-viral and HDR-based precision editing in patient and healthy control T cells. The developed platform enables custom editing, such as correction of small pathogenic variants, with one workflow that we demonstrate to achieve up to 80% efficiency in multiple genomic loci and donors. Institutional permissionsThe study was conducted in accordance with the principles of the Helsinki Declaration and approved by the Helsinki University Central Hospital Ethics Committee, and the Regional Committee for Medical and Health Research Ethics South-East Norway. All participants have signed written informed consent.

CRISPR interference (CRISPRi) is a powerful technology for studying loss-of-function phenotypes, enabling transient and reversible control of gene expression without the introduction of double-stranded DNA breaks. The cost of conducting large-scale CRISPR screens necessitates the selection of effective and specific sgRNAs for the design of compact libraries. While several genome-wide Cas9 CRISPRi libraries have been created, updates to transcript annotations, the generation of higher-resolution chromatin accessibility datasets and the development of newer on-target prediction models motivate an updated CRISPRi library design approach. Here, we generate large CRISPRi datasets tiling essential and nonessential genes. We compare the performance of multiple KRAB domain systems, develop an updated CRISPRi-specific on-target scoring scheme, and quantitatively characterize off-target effects associated with seed sequence patterns. We leverage these findings to design an optimized Cas9 CRISPRi library, Katsano, and validate its performance with genome-wide viability screens.

Antisense vivo-morpholino oligonucleotides (vivo-MOs) allow transient gene knockdown in adult organisms with high specificity and low toxicity. Vivo-MOs are used in cell culture and in many established model organisms, but a method for their use has not been described in threepsine stickleback (Gasterosteus aculeatus (Linnaeus, 1758)). Stickleback are an emerging model system used in evolutionary and ecological genetic studies. While genomic techniques are commonly used in stickleback research, there are few studies and tools available to assess gene function in-vivo, especially for genes that may be difficult to knock out by CRISPR (e.g., lethal knock-outs). Here, we test the use of splice-blocking vivo-MOs for gene knockdown in stickleback using intraperitoneal injection of vivo-MOs targeting three candidate genes. Gene expression was assessed in the liver, spleen, and intestine. Successful knockdown of Spi1b was observed in the spleen, however, we observed no other significant knockdown at either timepoint tested. Injection of a fluorescently labeled control vivo-MO confirmed delivery to each target organ, validating this approach, but delivery was variable which may explain inconsistent effects. These results indicate that vivo-MOs have potential as a tool for in-vivo gene knockdown in stickleback. Optimizing delivery methods could improve reproducibility and knockdown efficiency in future studies.

Rhodnius prolixus is an insect vector of the protozoan Trypanosoma cruzi, the causative agent of debilitating Chagas disease, which is transmitted to humans during blood feeding. Identifying germline markers is a critical step in advancing vector control and transgenic technologies of these medically important insects. Transmission of genetic traits to the next generation requires proper differentiation of the germline that gives rise to gametes. Germline precursors are established during early stages of development as the primordial germ cell (PGC) population. Among the genes required for this process, vasa homologues exert a conserved role in germline specification. Here, we characterize and validate the genomic structure of the R. prolixus Rp-vasa locus and assess its expression during early embryogenesis. We observe widespread Rp-vasa expression in preblastoderm embryos. Later, during the cellular blastoderm and at the beginning of gastrulation, Rp-vasa and Rp-piwi2 expression is restricted to PGCs, morphologically identifiable as a cluster of cells at the posterior of the embryo. We also report, for the first time, the use of R. prolixus regulatory sequences to drive the expression of exogenous genes. We identify the Rp-vasa regulatory region and show that these cis-regulatory sequences are sufficient to drive Cas9 and dsRed expression in the early embryo. Together, these findings demonstrate that Rp-vasa has great potential for use as a PGC marker and as a driver for gene expression in transgenic and gene editing approaches for Triatomine vectors.

Promoters and vectors are critical components of gene therapy, enabling the delivery and expression of therapeutic genes to correct both loss- and gain-of-function mutations. Adeno-associated virus (AAV) vectors are the leading platform for in vivo gene delivery; however, the widely used Streptococcus pyogenes Cas9 (SpCas9, 4.1 kb) approaches the AAV packaging limit of 4.7 kb. This constraint often necessitates dual-vector systems, which reduce therapeutic efficiency, or the use of smaller nucleases such as SaCas9 (3.2 kb) and AacCas12b (3.4 kb), which have lower PAM site frequencies. To enhance promoter selection for gene therapy applications, we developed a strategy to identify compact, cell-preferred RNA polymerase II (Pol II) promoters. Analysis of approximately 300 compact Pol II promoters revealed that exogenous expression levels in one cell type correlate more strongly with those in other cell types than with endogenous expression, underscoring the importance of exogenous expression efficiency in promoter selection. Using this approach, we identified a compact Pol II promoter #2 (Pro2, 133 bp) that drives robust transgene expression in human retinal ganglion cells (RGCs). To enable single-AAV delivery of SpCas9, we analyzed three commonly used Pol III promoters (H1, 7SK and U6) and determined their minimal functional lengths using a CRISPR/Cas9 reporter assay. We further engineered three compact hybrid Pol II/III promoters which combined pro2 with minimal H1, 7SK and U6 (276, 294, and 323 bp) capable of co-expressing SpCas9 and gRNA, enabling efficient genome editing in both transfected HEK293 cells (approaching 100%) and human RGCs (up to 55.9%) from human stem cell-derived retinal ganglion cells (RGCs). Together, these findings establish a framework for developing single-AAV CRISPR-based gene therapy strategies. Authors contributionsPWZ and DJZ conceived the study, designed the experiments, performed data analysis and interpretation, and were the primary contributors to manuscript writing. STZ played a key role in data collection and correlation analysis. YYC, SL, LF, CJK, YD, CAB, JC, and DW contributed to the execution of essential experiments and subsequent data analysis. All authors have read and approved the final manuscript. Declaration of interestsThe authors declare no conflicts of interest.

CRISPR-Cas9 is a powerful tool for precise genome editing in plants, but the presence of foreign DNA, such as T-DNA, raises regulatory concerns and complicates mutant screening and field studies of edited material. Detecting plants with good transgene expression and later removing the T-DNA from edited plants is both time-consuming and costly. To address this, we developed a system that uses the non-destructive RUBY reporter, linked to the CRISPR-Cas9 cassette, and expressed under the ZmUbi1 promoter. To assess the applicability of the system, it was tested on two Triticum species, targeting three genes in either tetraploid or hexaploid wheat. Strong correlations were observed in both T0 and T1 plants between betalain content and Cas9 expression, allowing for the quick identification of plants likely to be edited. Furthermore, the RUBY reporter could be used to select against the transgenic CRISPR-Cas9 cassette in subsequent generations at both the seed and seedling stages, thereby reducing the number of plants that need to be screened to identify edited lines without a T-DNA. This approach, using a nondestructive reporter, enabled rapid distinction between transgene expression in primary transgenics and served as a negative selection in the T1 generation, streamlining selection towards edited and T-DNA-free progeny.

Accurate prediction of CRISPR-Cas9 guide RNA (gRNA) editing efficiency remains limited, particularly outside human systems, where models trained on exogenous human datasets show poor generalization. We analyzed Cas9 efficiency and repair outcomes using novel endogenous editing data from four human cell types, two tomato cell types, and cells from giant river prawn and black soldier fly. While integrating publicly available predictors via ensemble frameworks improved performance, our analysis revealed hundreds of novel features affecting activity. Crucially, dominant features related to sites competition for gRNA, and local geometric properties varied across systems, highlighting the strong context dependence of Cas9 efficiency and arguing against a universal model. Interestingly, codon usage bias-based features also emerged as informative predictors, as they are proxies for chromatin accessibility. In contrast, trends in repair outcomes remained conserved. This work provides essential resources for more generalizable CRISPR guide design.

The use of viral vectors offers a promising alternative to traditional transformation methods for creating gene-edited plants. In this study, we developed a novel plant genome editing system by delivering Cas9, Cas12f, and Cas12j nucleases along with their guide RNAs using a broad-host-range geminivirus, Wheat dwarf India virus (WDIV), in combination with Ageratum yellow leaf curl betasatellite (AYLCB). Cas9, Cas12f, and Cas12j nucleases were efficiently expressed along with corresponding guide RNAs under viral promoters. By leveraging tRNA spacers in place of external promoters and terminators, we significantly reduced the overall cargo size, streamlining vector design. Additionally, we compared the traditional AtU6-driven gRNA delivery with a novel spacer:gRNA:spacer format in Cas9-expressing lines and observed comparable editing efficiencies. The broad host range of WDIV and AYLCB, combined with this tissue culture-free genome editing platform, opens up possibilities for editing across a wide range of plant species.

Splice site mutations represent a major class of pathogenic mutations in many diseases, as these changes disrupt normal splicing leading to gene expression changes. Cystic fibrosis (CF) results from mutations to the cystic fibrosis transmembrane conductance regulator (CFTR) gene that encodes an essential ion channel. Approximately 13% of the over 2,100 known CFTR mutations disrupt 3 or 5 splice sites and are predicted to cause splicing defects. Because each splicing mutation is rare, developing individualized therapies to treat each one is financially challenging. Exon specific U1 snRNA (ExSpeU1) targets the non-conserved intronic region downstream the 5 splice site (ss) to rescue exon skipping. Because this approach is exon-rather than mutation-specific, a single agent can potentially rescue multiple mutations. In this study, we have developed a platform to systematically classify all patient variants associated with an exon that are predicted to affect splicing and then determine their rescue potential using ExSpeU1. Here we report the results of these studies. Our minigene reporter study shows that 7 of 10 exon 18 variants resulted in exon skipping. Four mutations at the 3 and 5 ss were rescued at least partially using a single ExSpeU1. Using a luciferase reporter, we observe that the splicing rescue is reflected at the protein level. Lastly, we demonstrate exon-targeting ExSpeU1s can also rescue 3 and 5 ss mutations. Overall, this study exemplifies the power of our platform to screen and rescue multiple patient-derived splicing mutations using a single agent.

Agrobacterium-mediated transformation relies on binary vectors in which T-DNA and virulence genes are maintained on separate replicons. While Golden Gate cloning has become standard for T-DNA assembly, no modular framework exists for systematic construction of Agrobacterium vector backbones. Here, we present BackBone Builder (B3), a Golden Gate-based platform for combinatorial backbone assembly. B3 uses the Type IIS enzyme PaqCI to minimize domestication and enables one-pot assembly of nine backbone modules plus a selectable cloning cassette. The system is compatible with GreenGate and remains independent of downstream cloning strategies. We generated a library of 42 backbone components, supporting a theoretical design space exceeding 370,000 constructs. A 4 x 4 origin-of-replication (ORI) matrix combining four Escherichia coli and four Agrobacterium ORIs assembled with 100% efficiency and functioned robustly in bacterial and plant contexts. Reporter expression reflected expected ORI-dependent patterns in E. coli, Agrobacterium, and Nicotiana benthamiana. A B3-derived maize transformation backbone achieved stable transformation efficiencies comparable to established vectors. B3 establishes a standardized and extensible framework for rational engineering of Agrobacterium binary vector architecture.

Plant viruses are recognized as rapid and effective vectors to deliver CRISPR-Cas reaction components into plants, a strategy termed virus-induced gene editing (VIGE). However, VIGE is limited by the host range of the viral vectors. Development of new viral vectors to target a broad range of plant species will potentially enable the delivery of the editing components to new cultivars. Potyviruses (genus Potyvirus) comprises the largest group of plant RNA viruses. The main limitation of potyviral vectors to express a non-coding RNA consists of potential insertion of stop codons that interrupt the large open reading frame that encompass most potyviral genome. This is the case with the Streptococcus pyogenes Cas9 sgRNA scaffold, which contains stop codons in all three possible frames. In this work, we first built on a visual reporter system targeting the two homeologs of Nicotiana benthamiana Magnesium chelatase subunit I (CHLI). Second, we developed a tobacco etch virus (genus Potyvirus)-derived vector for VIGE by engineering a modified Cas9 scaffold, free of stop codons, to maintain the potyviral polyprotein reading frame while ensuring effective editing. This vector self-replicates and moves systemically, delivering sgRNAs efficiently throughout the plant. This allowed to obtain plants exhibiting a white phenotype with their four alleles edited through in vitro regeneration from infected leaves, and also to produce edited progeny. We further demonstrated the vector utility in tomato. Given the conserved biological properties within the genus Potyvirus, these findings must be broadly applicable to other potyviruses, expanding the reach of the VIGE technology.

Gene therapy offers the potential for long-term treatment or cures for a range of chronic diseases. However, permanent gene therapy expression may not be desirable. Efforts have been made to create systems which can be switched on/off by stimuli including light, designer drugs, or cellular contexts such as increased electrical activity. Here, we designed a novel plasmid system in which ion channel expression, and therefore function, is regulated by microRNA (miR) - an endogenous class of short noncoding RNAs which negatively regulate gene expression via binding the 3 untranslated region of target transcripts. We modified an existing voltage-gated potassium channel gene therapy with a binding cassette for miR-193a-3p, and transfected this miR-193-OFF system in neuro2A cells. Co-transfection with an inhibitor or mimic of miR-193a-3p respectively enhanced or repressed expression of our transgene, assessed using a GFP marker. Using whole-cell voltage clamp, we observed enhanced voltage-gated potassium currents in cells co-transfected with a miR-193a-3p inhibitor, compared with a non-targeting control oligonucleotide. Together, this demonstrates the concept of a novel miR-mediated molecular switch which can bias therapeutic ion channel expression based on a specific miR signal. As miRs are a ubiquitous molecular mechanism, our approach could be applied to a wide range of cellular and disease contexts, potentially expanding gene therapy to new patient populations.

Extrachromosomal arrays are unique chromosome-like structures created from DNA injected into the C. elegans germline. Arrays are easy to create and allow for high expression of multiple transgenes. They are, however, unstable unless integrated into a chromosome. Current methods for integration, such as X-rays and CRISPR, damage DNA and are low-efficiency. Here, we demonstrate that the viral integrase PhiC31, which mediates a non-mutagenic recombination between short attB and attP sequences, can be used for extremely efficient and targeted integration of arrays. In this method, a transgene, a selectable marker, and attP sites are injected into the gonad of a strain that (1) has an attB site in its genome, and (2) expresses PhiC31 in its germline. F1 extrachromosomal arrays are cloned, grown for multiple generations with selection, and then screened for homozygous array integrations. The procedure is simple, requires less time than screening for extrachromosomal arrays, and arrays can be screened for transgene function after stable integration. Arrays that transmit are integrated by PhiC31 with 50-95% efficiency, allowing for the isolation of many unique integrants from a single injection. Arrays can also be integrated at fluorescent landing pads and arbitrary sites in the genome. Using nanopore sequencing, we show that three new integrated arrays are between 1.6 and 18 megabases in length, assemble with large repeats, and can contain hundreds of copies of injected transgenes. We have built a collection of strains and plasmids to enable array integration at multiple sites in the genome using various selections. PhiC1-mediated Integration of Arrays of Transgenes (PhiAT) will allow C. elegans researchers to shift from using unstable extrachromosomal arrays to directly integrating arrays.