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.

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.

Pooled CRISPR screening has become a widely used approach for functional genomics, yet the construction of screening libraries remains tightly coupled to specific vector architectures and experimental formats. Incorporation of barcodes, multiplexed guide configurations, alternative fluorophores, selectable markers, or distinct screening modalities often requires reconstruction of entire libraries, even when the underlying biological content remains unchanged. These limitations are particularly acute in engineered cellular systems that already contain reporters, knock-in alleles, or pre-existing selection markers. Here, we describe FORGE-CRISPR (Functional Oncology Research Genetic Engineering - CRISPR), a CRISPR library-construction system that separates biological guide content from screening-vector context. CRISPR knockout and CRISPR interference guide collections are first converted into reusable guide modules. Barcode modules and second-guide modules are generated separately and combined with guide modules during downstream assembly into screening acceptor vectors. This design allows guide-only, barcoded, and multiplexed libraries to be generated from shared physical components. To support this framework, we developed a manufacturing workflow in which synthetic oligonucleotide pools are converted into reusable FORGE module libraries through PCR amplification, Golden Gate cloning, and background suppression. Using this approach, we constructed nine reusable guide-module libraries ranging from 45 to 4,830 guides (11,303 guides in total), and evaluated library quality through PCR-NGS analysis of guide representation and abundance distributions. As content for these libraries, we defined a modular, non-overlapping set of focused target collections, termed the druggable oncology genome, by partitioning druggable targets according to clinical development status and dependency distribution in DepMap, while drawing guide sequences from established, experimentally validated genome-wide libraries. We report four resources: reusable guide, barcode, and second-guide modules; compatible screening acceptor vectors; a manufacturing and PCR-NGS quality-control workflow demonstrated across nine guide-module libraries; and a versioned set of focused druggable-oncology target collections.

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.

Hirschsprung disease (HSCR) is a complex developmental disorder of the enteric nervous system, primarily driven by regulatory variants within enhancer elements of the RET gene. To investigate how these variants lead to aganglionosis, we developed a humanized mouse model by inserting an intact 77kb human RET genomic locus into the Rosa26 safe-harbor locus. Utilizing "big DNA" synthetic biology and Bxb1-mediated recombination, we integrated the complete human locus including all exons, introns, and upstream regulatory elements which we validated via nanopore and short-read sequencing. Functional analysis confirmed in vivo human RET expression; however, our initial HSCR-associated "sensitive" haplotype expressed at only 21% of wild-type levels. This significant reduction proved insufficient to rescue the viability when endogenous mouse Ret was deleted. We identified that this deficiency is partially driven by five risk SNPs within established enhancers. Specifically, using CRISPR/Cas9 to restore a conserved Sox10 binding site (converting a sensitive SNP to a protective one) increased RET expression by 1.9-fold and restored transcription factor binding. This study provides a robust framework for modeling human-specific regulatory disorders and demonstrates the critical impact of non-coding variation on disease pathogenesis.

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.

Cas12a2 CRISPR nucleases, including SuCas12a2, have been shown to have extensive collateral activity towards RNA, ssDNA, and dsDNA. This collateral activity results in targeted cell elimination and has applications across biotechnology, agriculture, and human health. We explored the natural genetic diversity of Cas12a2 nucleases and characterized nine novel orthologs in a DNA damage kinetic assay in E. coli. Three new Cas12a2 orthologs (RsCas12a2, SdCas12a2, and HmCas12a2) were shown to have high collateral activity towards DNA. These nucleases are highly divergent from SuCas12a2, have conserved core RuvC catalytic residues, and have sequence diversity in the previously reported aromatic clamp residues required for nucleic acid positioning in the active site. We defined PFS preferences and mismatch tolerance for each high-activity Cas12a2 nuclease, expanding the available Cas12a2 toolbox, and discovered functional differences with obvious impacts on downstream applications.

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 discovery of sensory ion channels, such as thermosensitive transient receptor potential (TRP) channels and mechanosensitive PIEZOs, have transformed our understanding of mammalian sensory biology. However, the sensory receptor landscape remains incomplete, as many physiologically relevant sensory stimuli still lack identified molecular targets. Here, we describe a novel screening strategy utilizing FM 1-43, a fluorescent marker for activity of various cation channels, with a CRISPRa library (MPCL) targeting multi-transmembrane domain proteins. We validate this method by focusing on allyl isothiocyanate (AITC) and its putative receptor TRPA1. Specifically, we show that CRISPRa-mediated overexpression of TRPA1 is sufficient for FM 1-43 labeling when co-treated with AITC. Furthermore, we show that using FM 1-43 and AITC, we can efficiently FACS enrich TRPA1-expressing cells from a pool of MPCL-expressing cells. Collectively, this presents a novel method for rapidly screening select cation-dependent sensory stimuli.

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.

Gene therapy has been the subject of extensive research following the advent of gene-editing technologies. Genetic disorders with difficult-to-target tissues, such as cystic fibrosis (CF), still face many challenges in developing efficacious gene therapy. The potential universal approach of gene replacement involves inserting a functional CFTR gene after generating DNA double strand breaks using gene editors such as CRISPR/Cas9. However, this strategy has not achieved clinical significance, as CRISPR/Cas9-mediated integration of CFTR is limited primarily by the infrequent activity of the homology-directed repair (HDR) pathway. To circumvent this limitation and improve CFTR transgene integration and expression, we explored a method of adding a second integration site, which we termed the dual-locus-targeting method. Using a helper-dependent adenoviral vector (HDAd)-delivered CRISPR/Cas9 system in porcine epithelial cells, we found that sequential delivery of two vectors, one targeting the CFTR locus and the other the genomic safe harbour site GGTA1, enhanced the integration efficiency of lacZ and CFTR donor genes to 16.5% and 3.4%, respectively. These results demonstrated a potential strategy to improve the efficacy of CFTR replacement for the development of a universal and permanent gene therapy treatment for CF lung disease. GRAPHICAL ABSTRACT O_FIG O_LINKSMALLFIG WIDTH=200 HEIGHT=76 SRC="FIGDIR/small/731381v1_ufig1.gif" ALT="Figure 1"> View larger version (17K): org.highwire.dtl.DTLVardef@1774590org.highwire.dtl.DTLVardef@1782915org.highwire.dtl.DTLVardef@1d13b12org.highwire.dtl.DTLVardef@17d3f93_HPS_FORMAT_FIGEXP M_FIG C_FIG

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.

Spacer efficacy generally declines with distance from the leader sequence, but the scarcity of fine-scale studies hampers comparisons across taxa. Here, we investigated positional effects across an exceptionally long 121-spacer CRISPR array associated with the type I-C cas operon of a Myxococcus xanthus natural isolate. In plasmid-interference assays, we found that interference rapidly declined with distance from the leader sequence, with only the proximal ~4% of spacers conferring measurable interference. This contrasts strikingly with a study in Vibrio cholerae, in which it was shown that ~95% of spacers in a shorter (39-spacer) array were effective. Our results suggest that there is great variation in the effective proportion of spacers across species, highlighting the need for fine-scale studies of CRISPR-array activity across diverse bacterial lineages.

Precision proteome engineering requires tools that leverage CRISPR technologies, yet most current design platforms remain gene-centric. To bridge this gap, we introduce CORe Designer. Featuring an intuitive graphical user interface, CORe Designer enables researchers to design guide RNAs and homology-directed repair templates in a protein-centric manner. By automating these workflows, CORe Designer enables the design of targeted mutagenesis experiments for precision proteome engineering to be completed in minutes.

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.

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.

Precision phage therapeutics provide a promising strategy to combat multidrug-resistant pathogens, including Staphylococcus aureus. Efficient, specific packaging of genetic cargoes remains challenging. Using modular design principles, we report a minimal phagemid packaging signal consisting of the phage terminase small subunit under its native promoter that significantly outperforms conventional packaging signals. The utility of this synthetic terS operon was demonstrated through production of highly concentrated and genetically pure CRISPR-Cas antimicrobials. To circumvent CRISPR-mediated self-targeting during antimicrobial generation, a terS-deficient strain was engineered to express the anti-CRISPR protein AcrIIA4, enabling titers above 1010 transducing units per milliliter (TRU/mL) with over 94% purity. With a high-copy origin of replication module, CRISPR-Cas phage-like particle titers could approach 1012 TRU/mL. We discovered that pure CRISPR-Cas antimicrobials are potent and can be amplified in hosts possessing prophages. Taken altogether, this study defines the minimal and optimal genetic requirements for efficient, specific creation of phage-based technologies. Technological ReadinessThe described system for engineering phage-like particles has reached a technological readiness level (TRL) of 4-5 based on the provided laboratory validation and strong literature support from other engineered phage therapies applied to in vivo models. Previous systems have demonstrated high-purity phage-like particle preparation, but always at the cost of severely reduced productivity. Therefore, our demarcation of a specific, efficient, and minimal system for packaging nucleic acid cargoes into phage vectors is a critical step toward real-world use. Despite this, low levels of contaminating host/phage DNA remain a key barrier to phage-based therapies. Protein and strain engineering efforts can help mitigate terminase nonspecificity, but care must be taken to not compromise productivity. More generally, widespread adoption will require deeper understanding of host-pathogen-phage interactions, development of scalable GMP manufacturing processes, and harmonized regulatory guidance that recognizes the dynamic nature of phage-derived technologies.

Targeted knock-in technologies have enabled precise insertion of reporters, affinity tags, degrons, and other functional payloads into endogenous genomic loci. Over the past decade, a diverse collection of genome engineering strategies has emerged, including approaches based on homology-directed repair (HDR), microhomology-mediated end joining (MMEJ), homology-mediated end joining (HMEJ), and related methodologies. While these advances have greatly expanded the capabilities of endogenous genome engineering, they have also increased the complexity of donor design, assembly, and validation. Here, we describe FORGE-KI (Functional Oncology Research Genetic Engineering - Knock in), a pathway-matched design workflow for endogenous knock-in engineering that aligns the assembly strategy with the underlying repair mechanism. For large-cargo insertions, we use a modular five-component framework that separates gene-specific targeting arms from reusable functional modules, allowing rapid assembly of HDR donor constructs targeting AHR, IRF1, and FOSL1 from a shared reagent collection. For MMEJ/PITCh applications, where short targeting elements permit rapid fabrication, we developed a streamlined one-step pipeline in which the entire donor and selection payload is synthesized as a single continuous fragment for direct cloning, compressing the design-to-reagent cycle time. This MMEJ workflow is paired with a dual-promoter nuclease vector (pForge-KI-MMEJ-Cas9-DualGuide) that drives the PITCh-release and locus-specific guides from distinct promoters, a design intended to reduce the repeated-promoter instability associated with some dual-guide vectors. We also established a standardized workflow for donor assembly, generation of knock-in cell populations, molecular validation, and selectable-cassette removal, and we demonstrate it by generating a functional, selection-marker-free, cytokine-inducible IRF1 HDR reporter line and an inducible IRF1 PITCh/MMEJ reporter pool with confirmed junction enrichment. In parallel, we developed forgeKI, an R package that automates C-terminal reporter knock-in design across both HDR and PITCh/MMEJ repair pathways, including guide selection, target-biology validation, targeting-arm design, domestication, donor-assembly planning, and generation of synthesis-ready constructs. Together, the reagents and software provide a practical system for endogenous knock-in engineering that supports multiple payloads, selection strategies, and repair pathways within a shared donor organization. Rather than replacing existing knock-in technologies, this framework provides a modular foundation for incorporating, extending, and automating the published knock-in methods.

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.

Background: Rapid detection of viral pathogens can be challenging, especially when routine PCR fails. Conventional assays typically detect known viruses which are specifically targeted by the assay, which may result in the failure to identify novel or non-targeted viruses. Broad-range hybrid-capture sequencing enables unbiased detection of viruses, including those that are uncommon or divergent. Methods: We combined the TWIST Comprehensive Viral Research Panel (>3,000 virus species) with Oxford Nanopore Flongle sequencing for easy and quick viral genome detection. The workflow includes random-primed cDNA synthesis, dsDNA conversion, TWIST probe enrichment, and Nanopore sequencing. Performance was evaluated using the QCMD 2024 Viral Metagenomics EQA panel and one clinical sample. Results: All expected targets of the QCMD 2024 Viral Metagenomics EQA panel were detected; eight of thirteen viruses achieved [&ge;]90% genome coverage. The negative control showed no targeted viral reads. Mixed infections of DNA and RNA viruses were resolved accurately. The workflow from nucleic acid extraction to obtaining sequence data was completed within 3 days.