CRISPR takes first steps in humans
CRISPR-Cas9 is a revolutionary gene-editing technology that offers the potential to treat diseases such as cancer, but the effects of CRISPR in patients are currently unknown. Stadtmauer et al. report a phase 1 clinical trial to assess the safety and feasibility of CRISPR-Cas9 gene editing in three patients with advanced cancer (see the Perspective by Hamilton and Doudna). They removed immune cells called T lymphocytes from patients and used CRISPR-Cas9 to disrupt three genes (TRAC, TRBC, and PDCD1) with the goal of improving antitumor immunity. A cancer-targeting transgene, NY-ESO-1, was also introduced to recognize tumors. The engineered cells were administered to patients and were well tolerated, with durable engraftment observed for the study duration. These encouraging observations pave the way for future trials to study CRISPR-engineered cancer immunotherapies.
Most cancers are recognized and attacked by the immune system but can progress owing to tumor-mediated immunosuppression and immune evasion mechanisms. The infusion of ex vivo engineered T cells, termed adoptive T cell therapy, can increase the natural antitumor immune response of the patient. Gene therapy to redirect immune specificity combined with genome editing has the potential to improve the efficacy and increase the safety of engineered T cells. CRISPR coupled with CRISPR-associated protein 9 (Cas9) endonuclease is a powerful gene-editing technology that potentially allows the ability to target multiple genes in T cells to improve cancer immunotherapy.
Our first-in-human, phase 1 clinical trial (clinicaltrials.gov; trial NCT03399448) was designed to test the safety and feasibility of multiplex CRISPR-Cas9 gene editing of T cells from patients with advanced, refractory cancer. A limitation of adoptively transferred T cell efficacy has been the induction of T cell dysfunction or exhaustion. We hypothesized that removing the endogenous T cell receptor (TCR) and the immune checkpoint molecule programmed cell death protein 1 (PD-1) would improve the function and persistence of engineered T cells. In addition, the removal of PD-1 has the potential to improve safety and reduce toxicity that can be caused by autoimmunity. A synthetic, cancer-specific TCR transgene (NY-ESO-1) was also introduced to recognize tumor cells. In vivo tracking and persistence of the engineered T cells were monitored to determine if the cells could persist after CRISPR-Cas9 modifications.
Four cell products were manufactured at clinical scale, and three patients (two with advanced refractory myeloma and one with metastatic sarcoma) were infused. The editing efficiency was consistent in all four products and varied as a function of the single guide RNA (sgRNA), with highest efficiency observed for the TCR α chain gene (TRAC) and lowest efficiency for the TCR β chain gene (TRBC). The mutations induced by CRISPR-Cas9 were highly specific for the targeted loci; however, rare off-target edits were observed. Single-cell RNA sequencing of the infused CRISPR-engineered T cells revealed that ~30% of cells had no detectable mutations, whereas ~40% had a single mutation and ~20 and ~10% of the engineered T cells were double mutated and triple mutated, respectively, at the target sequences. The edited T cells engrafted in all three patients at stable levels for at least 9 months. The persistence of the T cells expressing the engineered TCR was much more durable than in three previous clinical trials during which T cells were infused that retained expression of the endogenous TCR and endogenous PD-1. There were no clinical toxicities associated with the engineered T cells. Chromosomal translocations were observed in vitro during cell manufacturing, and these decreased over time after infusion into patients. Biopsies of bone marrow and tumor showed trafficking of T cells to the sites of tumor in all three patients. Although tumor biopsies revealed residual tumor, in both patients with myeloma, there was a reduction in the target antigens NY-ESO-1 and/or LAGE-1. This result is consistent with an on-target effect of the engineered T cells, resulting in tumor evasion.
Preliminary results from this pilot trial demonstrate that multiplex human genome engineering is safe and feasible using CRISPR-Cas9. The extended persistence of the engineered T cells indicates that preexisting immune responses to Cas9 do not appear to present a barrier to the implementation of this promising technology.
T cells (center) were isolated from the blood of a patient with cancer. CRISPR-Cas9 ribonuclear protein complexes loaded with three sgRNAs were electroporated into the normal T cells, resulting in gene editing of the TRAC, TRBC1, TRBC2, and PDCD1 (encoding PD-1) loci. The cells were then transduced with a lentiviral vector to express a TCR specific for the cancer-testis antigens NY-ESO-1 and LAGE-1 (right). The engineered T cells were then returned to the patient by intravenous infusion, and patients were monitored to determine safety and feasibility. PAM, protospacer adjacent motif.
CRISPR-Cas9 gene editing provides a powerful tool to enhance the natural ability of human T cells to fight cancer. We report a first-in-human phase 1 clinical trial to test the safety and feasibility of multiplex CRISPR-Cas9 editing to engineer T cells in three patients with refractory cancer. Two genes encoding the endogenous T cell receptor (TCR) chains, TCRα (TRAC) and TCRβ (TRBC), were deleted in T cells to reduce TCR mispairing and to enhance the expression of a synthetic, cancer-specific TCR transgene (NY-ESO-1). Removal of a third gene encoding programmed cell death protein 1 (PD-1; PDCD1), was performed to improve antitumor immunity. Adoptive transfer of engineered T cells into patients resulted in durable engraftment with edits at all three genomic loci. Although chromosomal translocations were detected, the frequency decreased over time. Modified T cells persisted for up to 9 months, suggesting that immunogenicity is minimal under these conditions and demonstrating the feasibility of CRISPR gene editing for cancer immunotherapy.
Gene editing offers the potential to correct DNA mutations and may offer promise to treat or eliminate countless human genetic diseases. The goal of gene editing is to change the DNA of cells with single–base pair precision. The principle was first demonstrated in mammalian cells when it was shown that expression of a rare cutting endonuclease to create double-strand DNA breaks resulted in repair by homologous and nonhomologous recombination (1). A variety of engineered nucleases were then developed to increase efficiency and enable potential therapeutic applications, including zinc finger nucleases, homing endonucleases, transcription activator–like effector nucleases, and CRISPR-Cas9 (clustered regularly interspaced short palindromic repeats associated with Cas9 endonuclease) (2). The first pilot human trials using genome editing were conducted in patients with HIV/AIDS and targeted the white blood cell protein CCR5, with the goal of mutating the CCR5 gene by nonhomologous recombination and thereby inducing resistance to HIV infection (3, 4). The incorporation of multiple guide sequences in CRISPR-Cas9 permits, in principle, multiplex genome engineering at several sites within a mammalian genome (5–9). The ability of CRISPR to facilitate efficient multiplex genome editing has greatly expanded the scope of possible targeted genetic manipulations, enabling new possibilities such as simultaneous deletion or insertion of multiple DNA sequences in a single round of mutagenesis. The prospect of using CRISPR engineering to treat a host of diseases, such as inherited blood disorders and blindness, is moving closer to reality.
Recent advances in CRISPR-Cas9 technology have also permitted efficient DNA modifications in human T cells, which holds great promise for enhancing the efficacy of cancer therapy. T lymphocytes are specialized immune cells that are largely at the core of the modern-day cancer immunotherapy revolution. The T cell receptor (TCR) complex is located on the surface of T cells and is central for initiating successful antitumor responses by recognizing foreign antigens and peptides bound to major histocompatibility complex molecules. One of the most promising areas of cancer immunotherapy involves adoptive cell therapy, whereby the patient’s own T cells are genetically engineered to express a synthetic (transgenic) TCR that can specifically detect and kill tumor cells. Recent studies have shown safety and promising efficacy of such adoptive T cell transfer approaches using transgenic TCRs specific for the immunogenic NY-ESO-1 tumor antigen in patients with myeloma, melanoma, and sarcoma (10–12). One limitation of this approach is that the transgenic TCR has been shown to mispair and/or compete for expression with the α and β chains of the endogenous TCR (13–15). Mispairing of the therapeutic TCR α and β chains with endogenous α and β chains reduces therapeutic TCR cell surface expression and potentially generates self-reactive TCRs.
A further shortcoming of adoptively transferred T cells has been the induction of T cell dysfunction or exhaustion leading to reduced efficacy (16). Programmed cell death protein 1 (PD-1)–deficient allogeneic mouse T cells with transgenic TCRs showed enhanced responses to alloantigens, indicating that the PD-1 protein on T cells plays a negative regulatory role in antigen responses that are likely to be cell intrinsic (17). The adoptive transfer of PD-1–deficient T cells in mice with chronic lymphocytic choriomeningitis virus infection initially leads to enhanced cytotoxicity and later to enhanced accumulation of terminally differentiated T cells (18). Antibody blockade of PD-1, or disruption or knockdown of the gene encoding PD-1 (i.e., PDCD1), improved chimeric antigen receptor (CAR) or TCR T cell–mediated killing of tumor cells in vitro and enhanced clearance of PD-1 ligand–positive (PD-L1+) tumor xenografts in vivo (19–23). In preclinical studies, we and others found that CRISPR-Cas9–mediated disruption of PDCD1 in human T cells transduced with a CAR increased antitumor efficacy in tumor xenografts (24–26). Adoptive transfer of transgenic TCR T cells specific for the cancer antigen NY-ESO-1, in combination with a monoclonal antibody targeting PD-1, enhanced antitumor efficacy in mice (27). We therefore designed a first-in-human, phase 1 human clinical trial to test the safety and feasibility of multiplex CRISPR-Cas9 genome editing for a synthetic biology cancer immunotherapy application. We chose to target endogenous TRAC, TRBC, and PDCD1 on T cells to increase the safety and efficacy profile of NY-ESO-1 TCR–expressing engineered cells. In principle, this strategy allowed us to increase exogenous TCR expression and reduce the potential for mixed heterodimer formation (i.e., by deleting the α and β TCR domain genes TRAC and TRBC, respectively) and to limit the development of T cell exhaustion, which can be triggered by the checkpoint ligands PD-L1 and PD-L2 (i.e., by deleting PDCD1).
The phase 1 human trial (clinicaltrials.gov; trial NCT03399448) was designed to assess the safety and feasibility of infusing autologous NY-ESO-1 TCR–engineered T cells in patients after CRISPR-Cas9 editing of the TRAC, TRBC, and PDCD1 loci. During the manufacturing process, cells were taken out of the cancer patient, engineered, and then infused back into the individual. The genetically engineered T cell product was termed “NYCE” (NY-ESO-1–transduced CRISPR 3X edited cells) and is referred to as NYCE hereafter. During clinical development of the protocol, we elected to use a TCR rather than a CAR because the incidence of cytokine release syndrome is generally less prevalent using TCRs (11). In principle, this allowed a more discriminating assessment of whether gene editing with Cas9 was potentially immunogenic or toxic when compared with the baseline low level of adverse events observed in our previous clinical trial targeting NY-ESO-1 with transgenic TCRs (11). The autologous T cells were engineered by lentiviral transduction to express an HLA-A2*0201–restricted TCR specific for the SLLMWITQC peptide in NY-ESO-1 and LAGE-1. The manufacturing process, vector design, and clinical protocol for NYCE T cells are described in the materials and methods and are depicted schematically (figs. S1 and S2). Of the six patients who were initially enrolled, four patients had successfully engineered T cells that were subjected to detailed release criteria testing as specified in the U.S. Food and Drug Administration (FDA)–accepted Investigational New Drug application (table S1) (see fig. S3 for the consort diagram). Of the four patients with cell products available, one patient assigned unique patient number (UPN) 27 experienced rapid clinical progression and was no longer eligible for infusion owing to the inability to meet protocol-mandated safety criteria (see supplementary materials). Of the three patients who were infused with CRISPR-Cas9–engineered T cells, two patients had refractory advanced myeloma and one patient had a refractory metastatic sarcoma not responding to multiple prior therapies (Table 1). The patients were given lymphodepleting chemotherapy with cyclophosphamide and fludarabine on days −5 to −3 (i.e., before administration with CRISPR-Cas9–engineered T cells) and a single infusion of 1 × 108 manufactured CRISPR-Cas9–engineered T cells per kilogram on day 0 of the protocol (fig. S2). No cytokines were administered to the patients.
Characteristics of infused CRISPR-Cas9–engineered T cell products
The T cell product was manufactured by electroporation of ribonucleoprotein complexes (RNPs) comprising recombinant Cas9 loaded with equimolar mixtures of single guide RNA (sgRNA) for TRAC, TRBC, and PDCD1 followed by lentiviral transduction of the transgenic TCR (Fig. 1A). All products were expanded to >1 × 1010 T cells by the time of harvest (Fig. 1B). The transgenic TCR could be detected by flow cytometric staining for Vβ8.1 or dextramer staining, ranging from 2 to 7% of T cells in the final product (Fig. 1C). The frequency of editing, as determined by digital polymerase chain reaction (PCR), varied according to the sgRNA and was about 45% for TRAC, 15% for TRBC, and 20% for PDCD1 (Fig. 1D). Final product transduction efficiency, CD4:CD8 ratio, and dosing are shown in table S2.
(A) Schematic representation of CRISPR-Cas9 NYCE T cells. (B) Large-scale expansion of NYCE T cells. Autologous T cells were transfected with Cas9 protein complexed with sgRNAs (RNP complex) against TRAC, TRBC (i.e., endogenous TCR deletion), and PDCD1 (i.e., PD-1 deletion) and subsequently transduced with a lentiviral vector to express a transgenic NY-ESO-1 cancer-specific TCR. Cells were expanded in dynamic culture for 8 to 12 days. On the final day of culture, NYCE T cells were harvested and cryopreserved in infusible medium. The total number of enriched T cells during culture is plotted for all four subjects (UPN07, UPN27, UPN35, and UPN39). (C) NY-ESO-1 TCR transduction efficiency was determined in harvested infusion products by flow cytometry. Data are gated on live CD3-expressing and Vβ8.1- or dextramer-positive lymphocytes and further gated on CD4-positive and/or CD8-positive cells. (D) The frequencies of TRAC, TRBC, and PDCD1 gene-disrupted total cells in NYCE infusion products were measured using chip-based digital PCR. All data are representative of at least two independent experiments. Error bars represent mean ± SEM.
The potency of the final engineered T cells was assessed by coculture with HLA-A2+ tumor cells engineered to express NY-ESO-1 (Fig. 2A). The engineered T cells had potent antigen-specific cytotoxicity over a wide range of effector-to–target cell ratios. Interestingly, the cells treated with CRISPR-Cas9 were more cytotoxic than control cells transduced with the TCR but electroporated without CRISPR-Cas9 (i.e., cells that retained endogenous TCR). This is consistent with previous findings in mouse T cells, when a transgenic TCR was inserted into the endogenous locus, ablating expression of the endogenous TCR (15). Further studies will be required to determine if PD-1 knockout contributes to the increased potency afforded by knockout of the endogenous TCR.
(A) Cytotoxicity of NYCE T cells cocultured with HLA-A*0201–positive Nalm-6 tumor cells engineered to express NY-ESO-1 and luciferase. Patient T cells transduced with the NY-ESO-1 TCR without CRISPR-Cas9 editing (NY-ESO-1 TCR) and untransduced T cells with CRISPR-Cas9 editing of TRAC, TRBC, and PDCD1 (labeled CRISPR) were included as controls (n = 4 patient T cell infusion products). Asterisks indicate statistical significance determined by paired Student’s t tests between groups (*P < 0.05). Error bars represent SEM. (B) Levels of soluble interferon-γ produced by patient NYCE T cell infusion products (labeled NYCE) after a 24-hour coculture with anti-CD3 and anti-CD28 antibody-coated beads or NY-ESO-1–expressing Nalm-6 target cells. Patient NY-ESO-1 TCR–transduced T cells (NY-ESO-1 TCR) and untransduced, CRISPR-Cas9–edited T cells (labeled CRISPR) served as controls. Error bars represent SEM. (C) Quantification of residual Cas9 protein in NYCE T cell infusion products in clinical-scale manufacturing is shown over time. Asterisks indicate statistical significance determined by paired Student’s t tests between time points (*P < 0.05). (D) Results from the fluorescence-based indirect ELISA screen performed to detect antibodies against Cas9 protein in the sera of three patients treated with NYCE T cells. Each dot represents the amount of anti-Cas9 signals detected in patient serum before T cell infusion (indicated by a vertical black arrow) and at various time points after NYCE T cell transfer. RFU, relative fluorescent units. (E) Immunoreactive Cas9-specific T cells in baseline patient leukapheresis samples were detected. Representative flow cytometry plots (left) from two patients whose T cells were positive for interferon-γ in response to Cas9 peptide stimulation. Unstimulated T cells treated with vehicle alone (dimethyl sulfoxide, DMSO) served as a negative control, whereas matched T cells stimulated with phorbol myristate acetate (PMA) and ionomycin served as a positive control. Bar graphs (right) show the frequency of ex vivo CD4+ and CD8+ T cells from patients or healthy donor controls (n = 6) that secrete interferon-γ in response to stimulation with three different Cas9 peptide pools. The background frequency of interferon-γ–expressing T cells (unstimulated control group, DMSO alone) is subtracted from the values shown in the bar graph. Error bars represent SD.
We developed a sensitive immunoassay for detection of Streptococcus pyogenes Cas9 protein and quantified Cas9 early in the manufacturing process, showing declining levels that were <0.75 fg per cell in the harvested final product (Fig. 2C). Using a competitive fluorescence enzyme-linked immunosorbent assay (ELISA) screen, we found that healthy donors have humoral reactivity to Cas9 in serum (data not shown) and T cells (Fig. 2E), confirming previous reports (28–30). Interestingly, we found that the three patients tested at a variety of time points after infusion of the engineered T cells did not develop humoral responses to Cas9. The lack of immunization to Cas9 is consistent with the extended persistence of the infused cells (Fig. 3) and could be a consequence of the low content of Cas9 in the infused product and/or to the immunodeficiency in the patients as a result of their extensive previous treatment histories (Table 1).
(A) The total number of vector copies per microgram of genomic DNA of the NY-ESO-1 TCR transgene in the peripheral blood (UPN07, UPN35, and UPN39), bone marrow (UPN07 and UPN35; multiple myeloma), and tumor (UPN39; sarcoma) is shown pre– and post–NYCE T cell infusion. (B) Calculated absolute numbers of NY-ESO-1 TCR–expressing T cells per microliter of whole blood from the time of infusion to various postinfusion time points in the study are shown. The limit of detection is about 2.5 cells per microliter of whole blood. (C) Frequencies of CRISPR-Cas9–edited T cells (TRAC, TRBC, and PDCD1 knockout) before and after adoptive cell transfer are depicted. Error bars represent SD.