An evolved AAV variant enables efficient genetic engineering of murine T cells.

Precise targeting of large transgenes to T cells using homology-directed repair has been transformative for adoptive cell therapies and T cell biology. Delivery of DNA templates via adeno-associated virus (AAV) has greatly improved knockin efficiencies, but the tropism of current AAV serotypes restricts their use to human T cells employed in immunodeficient mouse models. To enable targeted knockins in murine T cells, we evolved Ark313, a synthetic AAV that exhibits high transduction efficiency in murine T cells. We performed a genome-wide knockout screen and identified QA2 as an essential factor for Ark313 infection. We demonstrate that Ark313 can be used for nucleofection-free DNA delivery, CRISPR-Cas9-mediated knockouts, and targeted integration of large transgenes. Ark313 enables preclinical modeling of Trac-targeted CAR-T and transgenic TCR-T cells in immunocompetent models. Efficient gene targeting in murine T cells holds great potential for improved cell therapies and opens avenues in experimental T cell immunology.

Modular pooled discovery of synthetic knockin sequences to program durable cell therapies.

Chronic stimulation can cause T cell dysfunction and limit the efficacy of cellular immunotherapies. Improved methods are required to compare large numbers of synthetic knockin (KI) sequences to reprogram cell functions. Here, we developed modular pooled KI screening (ModPoKI), an adaptable platform for modular construction of DNA KI libraries using barcoded multicistronic adaptors. We built two ModPoKI libraries of 100 transcription factors (TFs) and 129 natural and synthetic surface receptors (SRs). Over 30 ModPoKI screens across human TCR- and CAR-T cells in diverse conditions identified a transcription factor AP4 (TFAP4) construct that enhanced fitness of chronically stimulated CAR-T cells and anti-cancer function in vitro and in vivo. ModPoKI's modularity allowed us to generate an ∼10,000-member library of TF combinations. Non-viral KI of a combined BATF-TFAP4 polycistronic construct enhanced fitness. Overexpressed BATF and TFAP4 co-occupy and regulate key gene targets to reprogram T cell function. ModPoKI facilitates the discovery of complex gene constructs to program cellular functions.

Mitigation of chromosome loss in clinical CRISPR-Cas9-engineered T cells.

CRISPR-Cas9 genome editing has enabled advanced T cell therapies, but occasional loss of the targeted chromosome remains a safety concern. To investigate whether Cas9-induced chromosome loss is a universal phenomenon and evaluate its clinical significance, we conducted a systematic analysis in primary human T cells. Arrayed and pooled CRISPR screens revealed that chromosome loss was generalizable across the genome and resulted in partial and entire loss of the targeted chromosome, including in preclinical chimeric antigen receptor T cells. T cells with chromosome loss persisted for weeks in culture, implying the potential to interfere with clinical use. A modified cell manufacturing process, employed in our first-in-human clinical trial of Cas9-engineered T cells (NCT03399448), reduced chromosome loss while largely preserving genome editing efficacy. Expression of p53 correlated with protection from chromosome loss observed in this protocol, suggesting both a mechanism and strategy for T cell engineering that mitigates this genotoxicity in the clinic.

NUDT21 limits CD19 levels through alternative mRNA polyadenylation in B cell acute lymphoblastic leukemia.

B cell progenitor acute lymphoblastic leukemia (B-ALL) treatment has been revolutionized by T cell-based immunotherapies-including chimeric antigen receptor T cell therapy (CAR-T) and the bispecific T cell engager therapeutic, blinatumomab-targeting surface glycoprotein CD19. Unfortunately, many patients with B-ALL will fail immunotherapy due to 'antigen escape'-the loss or absence of leukemic CD19 targeted by anti-leukemic T cells. In the present study, we utilized a genome-wide CRISPR-Cas9 screening approach to identify modulators of CD19 abundance on human B-ALL blasts. These studies identified a critical role for the transcriptional activator ZNF143 in CD19 promoter activation. Conversely, the RNA-binding protein, NUDT21, limited expression of CD19 by regulating CD19 messenger RNA polyadenylation and stability. NUDT21 deletion in B-ALL cells increased the expression of CD19 and the sensitivity to CD19-specific CAR-T and blinatumomab. In human B-ALL patients treated with CAR-T and blinatumomab, upregulation of NUDT21 mRNA coincided with CD19 loss at disease relapse. Together, these studies identify new CD19 modulators in human B-ALL.

Base-editing mutagenesis maps alleles to tune human T cell functions.

CRISPR-enabled screening is a powerful tool for the discovery of genes that control T cell function and has nominated candidate targets for immunotherapies1-6. However, new approaches are required to probe specific nucleotide sequences within key genes. Systematic mutagenesis in primary human T cells could reveal alleles that tune specific phenotypes. DNA base editors are powerful tools for introducing targeted mutations with high efficiency7,8. Here we develop a large-scale base-editing mutagenesis platform with the goal of pinpointing nucleotides that encode amino acid residues that tune primary human T cell activation responses. We generated a library of around 117,000 single guide RNA molecules targeting base editors to protein-coding sites across 385 genes implicated in T cell function and systematically identified protein domains and specific amino acid residues that regulate T cell activation and cytokine production. We found a broad spectrum of alleles with variants encoding critical residues in proteins including PIK3CD, VAV1, LCP2, PLCG1 and DGKZ, including both gain-of-function and loss-of-function mutations. We validated the functional effects of many alleles and further demonstrated that base-editing hits could positively and negatively tune T cell cytotoxic function. Finally, higher-resolution screening using a base editor with relaxed protospacer-adjacent motif requirements9 (NG versus NGG) revealed specific structural domains and protein-protein interaction sites that can be targeted to tune T cell functions. Base-editing screens in primary immune cells thus provide biochemical insights with the potential to accelerate immunotherapy design.

RASA2 ablation in T cells boosts antigen sensitivity and long-term function.

The efficacy of adoptive T cell therapies for cancer treatment can be limited by suppressive signals from both extrinsic factors and intrinsic inhibitory checkpoints1,2. Targeted gene editing has the potential to overcome these limitations and enhance T cell therapeutic function3-10. Here we performed multiple genome-wide CRISPR knock-out screens under different immunosuppressive conditions to identify genes that can be targeted to prevent T cell dysfunction. These screens converged on RASA2, a RAS GTPase-activating protein (RasGAP) that we identify as a signalling checkpoint in human T cells, which is downregulated upon acute T cell receptor stimulation and can increase gradually with chronic antigen exposure. RASA2 ablation enhanced MAPK signalling and chimeric antigen receptor (CAR) T cell cytolytic activity in response to target antigen. Repeated tumour antigen stimulations in vitro revealed that RASA2-deficient T cells show increased activation, cytokine production and metabolic activity compared with control cells, and show a marked advantage in persistent cancer cell killing. RASA2-knockout CAR T cells had a competitive fitness advantage over control cells in the bone marrow in a mouse model of leukaemia. Ablation of RASA2 in multiple preclinical models of T cell receptor and CAR T cell therapies prolonged survival in mice xenografted with either liquid or solid tumours. Together, our findings highlight RASA2 as a promising target to enhance both persistence and effector function in T cell therapies for cancer treatment.

Novel extragenic genomic safe harbors for precise therapeutic T-cell engineering.

Cell therapies that rely on engineered immune cells can be enhanced by achieving uniform and controlled transgene expression in order to maximize T-cell function and achieve predictable patient responses. Although they are effective, current genetic engineering strategies that use γ-retroviral, lentiviral, and transposon-based vectors to integrate transgenes, unavoidably produce variegated transgene expression in addition to posing a risk of insertional mutagenesis. In the setting of chimeric antigen receptor (CAR) therapy, inconsistent and random CAR expression may result in tonic signaling, T-cell exhaustion, and variable T-cell persistence. Here, we report and validate an algorithm for the identification of extragenic genomic safe harbors (GSH) that can be efficiently targeted for DNA integration and can support sustained and predictable CAR expression in human peripheral blood T cells. The algorithm is based on 7 criteria established to minimize genotoxicity by directing transgene integration away from functionally important genomic elements, maximize efficient CRISPR/Cas9-mediated targeting, and avert transgene silencing over time. T cells engineered to express a CD19 CAR at GSH6, which meets all 7 criteria, are curative at low cell dose in a mouse model of acute lymphoblastic leukemia, matching the potency of CAR T cells engineered at the TRAC locus and effectively resisting tumor rechallenge 100 days after their infusion. The identification of functional extragenic GSHs thus expands the human genome available for therapeutic precision engineering.

CAR T cell trogocytosis and cooperative killing regulate tumour antigen escape.

Chimeric antigen receptors (CARs) are synthetic antigen receptors that reprogram T cell specificity, function and persistence1. Patient-derived CAR T cells have demonstrated remarkable efficacy against a range of B-cell malignancies1-3, and the results of early clinical trials suggest activity in multiple myeloma4. Despite high complete response rates, relapses occur in a large fraction of patients; some of these are antigen-negative and others are antigen-low1,2,4-9. Unlike the mechanisms that result in complete and permanent antigen loss6,8,9, those that lead to escape of antigen-low tumours remain unclear. Here, using mouse models of leukaemia, we show that CARs provoke reversible antigen loss through trogocytosis, an active process in which the target antigen is transferred to T cells, thereby decreasing target density on tumour cells and abating T cell activity by promoting fratricide T cell killing and T cell exhaustion. These mechanisms affect both CD28- and 4-1BB-based CARs, albeit differentially, depending on antigen density. These dynamic features can be offset by cooperative killing and combinatorial targeting to augment tumour responses to immunotherapy.

Targeting a CAR to the TRAC locus with CRISPR/Cas9 enhances tumour rejection.

Chimeric antigen receptors (CARs) are synthetic receptors that redirect and reprogram T cells to mediate tumour rejection. The most successful CARs used to date are those targeting CD19 (ref. 2), which offer the prospect of complete remission in patients with chemorefractory or relapsed B-cell malignancies. CARs are typically transduced into the T cells of a patient using γ-retroviral vectors or other randomly integrating vectors, which may result in clonal expansion, oncogenic transformation, variegated transgene expression and transcriptional silencing. Recent advances in genome editing enable efficient sequence-specific interventions in human cells, including targeted gene delivery to the CCR5 and AAVS1 loci. Here we show that directing a CD19-specific CAR to the T-cell receptor α constant (TRAC) locus not only results in uniform CAR expression in human peripheral blood T cells, but also enhances T-cell potency, with edited cells vastly outperforming conventionally generated CAR T cells in a mouse model of acute lymphoblastic leukaemia. We further demonstrate that targeting the CAR to the TRAC locus averts tonic CAR signalling and establishes effective internalization and re-expression of the CAR following single or repeated exposure to antigen, delaying effector T-cell differentiation and exhaustion. These findings uncover facets of CAR immunobiology and underscore the potential of CRISPR/Cas9 genome editing to advance immunotherapies.

Low-Dose Radiation Conditioning Enables CAR T Cells to Mitigate Antigen Escape.

CD19 chimeric antigen receptors (CARs) have demonstrated great efficacy against a range of B cell malignancies. However, antigen escape and, more generally, heterogeneous antigen expression pose a challenge to applying CAR therapy to a wide range of cancers. We find that low-dose radiation sensitizes tumor cells to immune rejection by locally activated CAR T cells. In a model of pancreatic adenocarcinoma heterogeneously expressing sialyl Lewis-A (sLeA), we show that not only sLeA+ but also sLeA- tumor cells exposed to low-dose radiation become susceptible to CAR therapy, reducing antigen-negative tumor relapse. RNA sequencing analysis of low-dose radiation-exposed tumors reveals the transcriptional signature of cells highly sensitive to TRAIL-mediated death. We find that sLeA-targeted CAR T cells produce TRAIL upon engaging sLeA+ tumor cells, and eliminate sLeA- tumor cells previously exposed to systemic or local low-dose radiation in a TRAIL-dependent manner. These findings enhance the prospects for successfully applying CAR therapy to heterogeneous solid tumors. Local radiation is integral to many tumors' standard of care and can be easily implemented as a CAR conditioning regimen.

A platform for rapid prototyping of synthetic gene networks in mammalian cells.

Mammalian synthetic biology may provide novel therapeutic strategies, help decipher new paths for drug discovery and facilitate synthesis of valuable molecules. Yet, our capacity to genetically program cells is currently hampered by the lack of efficient approaches to streamline the design, construction and screening of synthetic gene networks. To address this problem, here we present a framework for modular and combinatorial assembly of functional (multi)gene expression vectors and their efficient and specific targeted integration into a well-defined chromosomal context in mammalian cells. We demonstrate the potential of this framework by assembling and integrating different functional mammalian regulatory networks including the largest gene circuit built and chromosomally integrated to date (6 transcription units, 27kb) encoding an inducible memory device. Using a library of 18 different circuits as a proof of concept, we also demonstrate that our method enables one-pot/single-flask chromosomal integration and screening of circuit libraries. This rapid and powerful prototyping platform is well suited for comparative studies of genetic regulatory elements, genes and multi-gene circuits as well as facile development of libraries of isogenic engineered cell lines.

Localized in vivo gene editing of murine cancer-associated fibroblasts.

Discovering the role of fibroblasts residing in the tumor microenvironment (TME) requires controlled, localized perturbations because fibroblasts play critical roles in regulating immunity and tumor biology at multiple sites. Systemic perturbations can lead to unintended, confounding secondary effects, and methods to locally genetically engineer fibroblasts are lacking. To specifically investigate murine stromal cell perturbations restricted to the TME, we developed an adeno-associated virus (AAV)-based method to target any gene-of-interest in fibroblasts at high efficiency (>80%). As proof of concept, we generated single (sKO) and double gene KOs (dKO) of Osmr, Tgfbr2, and Il1r1 in cancer-associated fibroblasts (CAFs) and investigated how their cell states and those of other cells of the TME subsequently change in mouse models of melanoma and pancreatic ductal adenocarcinoma (PDAC). Furthermore, we developed an in vivo knockin-knockout (KIKO) strategy to achieve long-term tracking of CAFs with target gene KO via knocked-in reporter gene expression. This validated in vivo gene editing toolbox is fast, affordable, and modular, and thus holds great potential for further exploration of gene function in stromal cells residing in tumors and beyond.

Peptide-enabled ribonucleoprotein delivery for CRISPR engineering (PERC) in primary human immune cells and hematopoietic stem cells.

Peptide-enabled ribonucleoprotein delivery for CRISPR engineering (PERC) is a new approach for ex vivo genome editing of primary human cells. PERC uses a single amphiphilic peptide reagent to mediate intracellular delivery of the same pre-formed CRISPR ribonucleoprotein enzymes that are broadly used in research and therapeutics, resulting in high-efficiency editing of stimulated immune cells and cultured hematopoietic stem and progenitor cells (HSPCs). PERC facilitates nuclease-mediated gene knockout, precise transgene knock-in, and base editing. PERC involves mixing the CRISPR ribonucleoprotein enzyme with peptide and then incubating the formulation with cultured cells. For efficient transgene knock-in, adeno-associated virus (AAV) bearing homology-directed repair template DNA may be included. In contrast to electroporation, PERC is appealing as it requires no dedicated hardware and has less impact on cell phenotype and viability. Due to the gentle nature of PERC, delivery can be performed multiple times without substantial impact to cell health or phenotype. Here we report methods for improved PERC-mediated editing of T cells as well as novel methods for PERC-mediated editing of HSPCs, including knockout and precise knock-in. Editing efficiencies can surpass 90% using either Cas9 or Cas12a in primary T cells or HSPCs. Because PERC calls for only three readily available reagents - protein, RNA, and peptide - and does not require dedicated hardware for any step, PERC demands no special expertise and is exceptionally straightforward to adopt. The inherent compatibility of PERC with established cell engineering pipelines makes this approach appealing for rapid deployment in research and clinical settings.

Spleen Tyrosine Kinase (SYK) negatively regulates ITAM-mediated human NK cell signaling and CD19-CAR NK cell efficacy.

NK cells express activating receptors that signal through ITAM-bearing adapter proteins. The phosphorylation of each ITAM creates binding sites for SYK and ZAP70 protein tyrosine kinases to propagate downstream signaling including the induction of Ca 2 + influx. While all immature and mature human NK cells co-express SYK and ZAP70, clonally driven memory or adaptive NK cells can methylate SYK genes and signaling is mediated exclusively using ZAP70. Here, we examined the role of SYK and ZAP70 in a clonal human NK cell line KHYG1 by CRISPR-based deletion using a combination of experiments and mechanistic computational modeling. Elimination of SYK resulted in more robust Ca + + influx after cross-linking of the CD16 and NKp30 receptors and enhanced phosphorylation of downstream proteins, whereas ZAP70 deletion diminished these responses. By contrast, ZAP70 depletion increased proliferation of the NK cells. As immature T cells express both SYK and ZAP70 but mature T cells often express only ZAP70, we transduced the human Jurkat cell line with SYK and found that expression of SYK increased proliferation but diminished TCR-induced Ca 2 + flux and activation. We performed transcriptional analysis of the matched sets of variant Jurkat and KHYG1 cells and observed profound alterations caused by SYK expression. As depletion of SYK in NK cells increased their activation, primary human NK cells were transduced with a CD19-targeting CAR and were CRISPR edited to ablate SYK or ZAP70. Deletion of SYK resulted in more robust cytotoxic activity and cytokine production, providing a new therapeutic strategy of NK cell engineering for cancer immunotherapy.

Ultra-high efficiency T cell reprogramming at multiple loci with SEED-Selection.

Multiplexed reprogramming of T cell specificity and function can generate powerful next-generation cellular therapies. However, current manufacturing methods produce heterogenous mixtures of partially engineered cells. Here, we develop a one-step process to enrich for unlabeled cells with knock-ins at multiple target loci using a family of repair templates named Synthetic Exon/Expression Disruptors (SEEDs). SEED engineering associates transgene integration with the disruption of a paired endogenous surface protein, allowing non-modified and partially edited cells to be immunomagnetically depleted (SEED-Selection). We design SEEDs to fully reprogram three critical loci encoding T cell specificity, co-receptor expression, and MHC expression, with up to 98% purity after selection for individual modifications and up to 90% purity for six simultaneous edits (three knock-ins and three knockouts). These methods are simple, compatible with existing clinical manufacturing workflows, and can be readily adapted to other loci to facilitate production of complex gene-edited cell therapies.

Scalable intracellular delivery via microfluidic vortex shedding enhances the function of chimeric antigen receptor T-cells.

Adoptive chimeric antigen receptor T-cell (CAR-T) therapy is transformative and approved for hematologic malignancies. It is also being developed for the treatment of solid tumors, autoimmune disorders, heart disease, and aging. Despite unprecedented clinical outcomes, CAR-T and other engineered cell therapies face a variety of manufacturing and safety challenges. Traditional methods, such as lentivirus transduction and electroporation, result in random integration or cause significant cellular damage, which can limit the safety and efficacy of engineered cell therapies. We present hydroporation as a gentle and effective alternative for intracellular delivery. Hydroporation resulted in 1.7- to 2-fold higher CAR-T yields compared to electroporation with superior cell viability and recovery. Hydroporated cells exhibited rapid proliferation, robust target cell lysis, and increased pro-inflammatory and regulatory cytokine secretion in addition to improved CAR-T yield by day 5 post-transfection. We demonstrate that scaled-up hydroporation can process 5 x 108 cells in less than 10 s, showcasing the platform as a viable solution for high-yield CAR-T manufacturing with the potential for improved therapeutic outcomes.

Characterization of three loci for homologous gene targeting and transgene expression.

Integrative gene transfer is widely used for bioproduction, drug screening, and therapeutic applications but usual viral methods lead to random and multicopy insertions, contribute to unstable transgene expression and can disturb endogenous gene expression. Homologous targeting of an expression cassette using rare-cutting endonucleases is a potential solution; however the number of studied loci remains limited. Furthermore, the behavior and performance of various types of gene cassettes following gene targeting is poorly defined. Here we have evaluated three loci for gene targeting, including one locus compatible with the proposed Safe Harbor criteria for human translational applications. Using optimized conditions for homologous gene targeting, reporter genes under the control of different promoters were efficiently inserted at each locus in both sense and antisense orientations. Sustainable expression was achieved at all three loci without detectable disturbance of flanking gene expression. However, the promoter, the integration locus and the cassette orientation have a strong impact on transgene expression. Finally, single targeted integrations exhibited greatly improved transgene expression stability versus multicopy or random integration. Taken together, our data suggest a potential set of loci for site-specific transgene integration, suitable for a variety of biotechnological applications.

Peptide-mediated delivery of CRISPR enzymes for the efficient editing of primary human lymphocytes.

CRISPR-mediated genome editing of primary human lymphocytes is typically carried out via electroporation, which can be cytotoxic, cumbersome and costly. Here we show that the yields of edited primary human lymphocytes can be increased substantially by delivering a CRISPR ribonucleoprotein mixed with an amphiphilic peptide identified through screening. We evaluated the performance of this simple delivery method by knocking out genes in T cells, B cells and natural killer cells via the delivery of Cas9 or Cas12a ribonucleoproteins or an adenine base editor. We also show that peptide-mediated ribonucleoprotein delivery paired with an adeno-associated-virus-mediated homology-directed repair template can introduce a chimaeric antigen receptor gene at the T-cell receptor α constant locus, and that the engineered cells display antitumour potency in mice. The method is minimally perturbative, does not require dedicated hardware, and is compatible with multiplexed editing via sequential delivery, which minimizes the risk of genotoxicity. The peptide-mediated intracellular delivery of ribonucleoproteins may facilitate the manufacturing of engineered T cells.

High-yield genome engineering in primary cells using a hybrid ssDNA repair template and small-molecule cocktails.

Enhancing CRISPR-mediated site-specific transgene insertion efficiency by homology-directed repair (HDR) using high concentrations of double-stranded DNA (dsDNA) with Cas9 target sequences (CTSs) can be toxic to primary cells. Here, we develop single-stranded DNA (ssDNA) HDR templates (HDRTs) incorporating CTSs with reduced toxicity that boost knock-in efficiency and yield by an average of around two- to threefold relative to dsDNA CTSs. Using small-molecule combinations that enhance HDR, we could further increase knock-in efficiencies by an additional roughly two- to threefold on average. Our method works across a variety of target loci, knock-in constructs and primary human cell types, reaching HDR efficiencies of >80-90%. We demonstrate application of this approach for both pathogenic gene variant modeling and gene-replacement strategies for IL2RA and CTLA4 mutations associated with Mendelian disorders. Finally, we develop a good manufacturing practice (GMP)-compatible process for nonviral chimeric antigen receptor-T cell manufacturing, with knock-in efficiencies (46-62%) and yields (>1.5 × 109 modified cells) exceeding those of conventional approaches.

Mitigation of chromosome loss in clinical CRISPR-Cas9-engineered T cells.

CRISPR-Cas9 genome editing has enabled advanced T cell therapies, but occasional loss of the targeted chromosome remains a safety concern. To investigate whether Cas9-induced chromosome loss is a universal phenomenon and evaluate its clinical significance, we conducted a systematic analysis in primary human T cells. Arrayed and pooled CRISPR screens revealed that chromosome loss was generalizable across the genome and resulted in partial and entire loss of the chromosome, including in pre-clinical chimeric antigen receptor T cells. T cells with chromosome loss persisted for weeks in culture, implying the potential to interfere with clinical use. A modified cell manufacturing process, employed in our first-in-human clinical trial of Cas9-engineered T cells, 1 dramatically reduced chromosome loss while largely preserving genome editing efficacy. Expression of p53 correlated with protection from chromosome loss observed in this protocol, suggesting both a mechanism and strategy for T cell engineering that mitigates this genotoxicity in the clinic.