Reprogramming human T cell function and specificity with non-viral genome targeting
Product# 1002: Recombinant tat HIV-1 IIIB
Decades of work have aimed to genetically reprogram T cells for therapeutic purposes1 using recombinant viral vectors, which do not target transgenes to specific genomic sites2,3. In addition, the need for viral vectors has slowed down research and clinical use as their manufacturing and testing is lengthy and expensive. Genome editing brought the promise of specific and efficient insertion of large transgenes into target cells through homology-directed repair (HDR)4,5. Here, we developed a CRISPR-Cas9 genome targeting system that does not require viral vectors, allowing rapid and efficient insertion of large DNA sequences (> 1kb) at specific sites in the genomes of primary human T cells, while preserving cell viability and function. This permits individual or multiplexed modification of endogenous genes. First, we apply this strategy to correct a pathogenic IL2RA mutation in cells from patients with monogenic autoimmune disease, demonstrating improved signaling function. Second, we replace the endogenous T cell receptor (TCR) locus with a new TCR redirecting T cells to a cancer antigen. The resulting TCR-engineered T cells specifically recognize tumour antigen and mount productive anti-tumour cell responses in vitro and in vivo. Taken together, these studies provide preclinical evidence that non-viral genome targeting can enable rapid and flexible experimental manipulation and therapeutic engineering of primary human immune cells.
The major barrier to effective non-viral T cell genome targeting of large DNA sequences has been the toxicity of the DNA6. While the introduction of short single-stranded oligodeoxynucleotide (ssODN) HDR templates does not cause significant T cell toxicity, it has been shown that larger linear double stranded (dsDNA) templates are toxic at high concentrations7,8. Contrary to expectations, we found that co-electroporation of human primary T cells with CRISPR-Cas9 ribonucleoprotein (Cas9 RNP9,10) complexes and long (>1kb) linear dsDNA templates reduced the toxicity associated with the dsDNA template (Extended Data Fig 1a-e). Cas9 RNPs were co-electroporated with a dsDNA HDR template designed to introduce an N-terminal GFP-fusion in the housekeeping gene RAB11A (Fig. 1a). Both cell viability and the efficiency of this approach were optimized by systematic exploration (Fig. 1b and Extended Data Fig. 1f-h) resulting in GFP expression in ~50% of both primary human CD4+ and CD8+ T cells. The method was reproducibly efficient with high cell viability (Fig. 1c, d, e). The system is also compatible with current manufacturing protocols for cell therapies. The method can be used with fresh or cryopreserved cells, bulk T cells or FACS-sorted sub-populations, and cells from whole blood or leukapheresis (Extended Data Fig. 2a-d).
We next confirmed that the system could be applied broadly by targeting sequences in different locations throughout the genome. We efficiently engineered primary T cells by generating GFP fusions with different genes (Fig. 2a and Extended Data Fig. 2e-g). Live-cell imaging with confocal microscopy confirmed the specificity of gene targeting, revealing the distinct sub-cellular locations of each of the resulting GFP-fusion proteins11 (Fig. 2b). Appropriate chromatin binding of a transcription factor GFP-fusion protein was confirmed by performing genome-wide CUT & RUN12 analysis with an anti-GFP antibody (Fig. 2c and Extended Data Fig. 2h). Finally, we showed that gene targeting preserved the regulation of the modified endogenous gene. Consistent with correct cell-type specific expression, a CD4-GFP fusion was selectively expressed in the CD4+ population of T cells (Fig. 2d). Using HDR templates encoding multiple fluorescent proteins, we demonstrated that we could generate cells with bi-allelic gene targeting (Fig. 2e and Extended Data Fig. 3a-d) or multiplex modification of two (Fig. 2f and Extended Data Fig. 3e-h) or even three (Fig. 2g and Extended Data Fig. 3i) different genes13,14. These results show that multiple endogenous genes can be directly engineered without virus in T cells, and that gene and protein regulation are preserved.
For therapeutic use of genetically modified T cells, integrated sequences should be introduced specifically without unintended disruption of other critical genome sites15. We performed targeted locus amplification (TLA) sequencing16 and found no evidence of off-target integrations above the assay’s limit of detection (~1% of alleles) (Extended Data Fig. 4a-b). We further assessed potential off-target integrations at the single cell level by quantifying GFP+ cells generated using a Cas9 RNP that cuts outside the homology site. Similar to what has been described with viral HDR templates4,17, we found evidence to suggest that double-stranded templates could integrate independent of target homology18,19, albeit at low rates (Extended Data Fig. 4c-i). These rare events could be reduced almost completely by using single-stranded DNA templates20,21 (Extended Data Fig. 5a-d). As an additional safeguard that could be important for some applications, we demonstrated that efficient non-viral T cell genome targeting also could be achieved using either a single-stranded or double-stranded template with a Cas9 “nickase” engineered to reduce potential off-target double-stranded cuts22,23 (Extended Data Fig. 5e-h).
Having optimized this non-viral genome engineering approach in primary human T cells, we demonstrated its utility it in two different clinically relevant settings where targeted replacement of a gene would provide proof-of-principle that the method can be used to create therapeutically relevant gene modifications. Specifically, we tested the ability to rapidly and efficiently correct an inherited genetic alteration in T cells and we also tested the targeted insertion of the two chains of a TCR to redirect the specificity of T cells to recognize cancer cells.
We identified a family with monogenic primary immune deficiency with autoimmune disease caused by recessive loss-of-function mutations in the gene encoding the IL-2 alpha receptor (IL2RA)24 (Supplementary Table 4), which is essential for healthy regulatory T cells (Tregs)25 (Extended Data Fig. 6a-h). Whole exome sequencing revealed that the IL2RA-deficient children harboured compound heterozygous mutations in IL2RA (Fig. 3a and Extended Data Fig. 6i). One mutation, c.530A>G, creates a premature stop codon. With non-viral genome targeting, we were able to correct the mutation and observed IL2RA expression on the surface of corrected T cells from the patient (Fig. 3b). Long dsDNA templates led to efficient correction of the mutations. Because only two base pair changes were necessary (one to correct the mutation and one to silently remove the gRNA’s PAM sequence), a short single-stranded DNA (~120 bps) could also be used to make the correction. These single-stranded DNAs were able to correct the mutation at high frequencies, although here the efficiency of correction was lower than with the longer dsDNA template (Extended Data Fig. 7a, ,8a).8a). Correction was successful in T cells from all three siblings, but lower rates of IL2RA expression were seen in compound het 3, which could be due to altered cell-state associated with the patient’s disease or the fact she was the only sibling treated with immunosuppressive therapy (Supplementary Table 4 and Extended Data Fig. 8f). The second mutation identified, c.800delA, causes a frameshift in the reading frame of the final IL2RA exon. This frameshift mutation could be corrected both by HDR as well as by RNP cutting alone, presumably due to some of the small indels restoring the reading frame (Extended Data Fig. 8). Taken together, these data show that distinct mutations can be corrected in patient T cells using HDR template-dependent and non-HDR template-dependent mechanisms.
Mutation correction improved cell signalling function. Following correction of the c.530A>G IL2RA mutation, IL-2 treatment led to increased STAT5 phosphorylation, a hallmark of productive signalling (Fig. 3c and Extended Data Fig. 7c, ,8c).8c). In addition, following correction, we found that the modified T cells expressed both IL2RA and FOXP3, a critical transcriptional factor in Tregs (Extended Data Fig. 7d, ,8d).8d). We were also able to correct the IL2RA mutation in a sorted population of CD3+CD4+CD127loTIGIT+CD45RO+ Treg-like cells from a patient (Extended Data Fig. 7e-f), a strategy that could potentially be used in a gene-modified cell therapy for the children in this family. Cell-type specific and stimulus responsive expression of IL2RA is under tight control by multiple endogenous cis-regulatory elements that constitute a super-enhancer26,27. Therefore, effective therapeutic correction of the IL2RA defect is likely to depend on repairing the gene in its endogenous genomic locus; off-target effects should be avoided. We therefore demonstrated that the c.800delA mutation could also be repaired using Cas9 nickase combined with a single-stranded HDR template (Fig. 3d).
Non-viral genome targeting not only allows the correction of point mutations, but also enables integration of much larger DNA sequences. We were able to use a large DNA construct to rapidly reprogram the antigen specificity of human T cells, which is critical for many cellular immunotherapy applications. Recent work demonstrates that chimeric antigen receptors (CARs) have enhanced efficacy when they are genetically encoded in the endogenous TCR locus using CRISPR-Cas9 gene cutting and an adeno-associated virus vector as a repair template4. Targeting of specific TCR sequences to this locus is a more challenging problem because T cells must express paired TCR alpha (TCR-α) and beta chains (TCR-β) to make a functional receptor.
We developed a strategy to replace the endogenous TCR using non-viral genome targeting to integrate an approximately 1.5 kb DNA cassette into the first exon of the TCR-α constant region (TRAC) (Fig. 4a). This cassette encoded the full-length sequence of a TCR-β separated by a self-excising 2A peptide from the variable region of a new TCR-α, which encodes the full TCR-α sequence when appropriately integrated at the endogenous TRAC exon (Extended Data Fig. 9a-d). To test this strategy, we introduced a TCR-β and TCR-α pair (1G4) that recognizes the NY-ESO-1 tumour antigen28 into the TRAC locus of polyclonal T cells isolated from healthy human donors. Antibody staining for total TCR-α/β expression and NY-ESO-1-MHC dextramer staining for the NY-ESO-1 TCR expression revealed that non-viral genome targeting enabled reproducible replacement of the endogenous TCR in both CD8+ and CD4+ primary human T cells (Fig. 4b and Extended Data Fig. 9k). NY-ESO-1 TCR cells could also be generated with a similar targeting strategy at the TCR-β constant region (TRBC1/2) or with multiplexed simultaneous replacement of both endogenous TCR-α and TCR-β (Extended Data Fig. 9e-i). The majority of the T cells that did not express NY-ESO-1 TCR were TCR knockouts (Fig. 4b), presumably due to NHEJ events induced by the Cas9-mediated double-stranded breaks in TRAC exon 1. Up to ~70% of resulting TCR-positive cells recognized the NY-ESO-1 dextramer.
Next, we assessed the tumour antigen-specific function of targeted human T cells. When the targeted T cells were co-cultured with two different NY-ESO-1+ melanoma cell lines, M257 and M407, the modified T cells robustly and specifically produced IFN-ɣ and TNF-α and induced T cell degranulation (measured by CD107a surface expression) (Fig. 4c). Cytokine production and degranulation only occurred when the NY-ESO-1 TCR T cells were exposed to cell lines expressing the appropriate HLA-A*0201 class I MHC allele required to present the cognate NY-ESO-1 peptide. Both the CD8+ and CD4+ T cell response was consistent across healthy donors, and was comparable to the response of T cells from the same healthy donor in which the NY-ESO-1 TCR was transduced by gamma retrovirus and heterologously expressed using a viral promoter (Fig. 4c and Extended Data Fig. 9j). NY-ESO-1 TCR knock-in T cells rapidly killed target M257-HLA-A*0201 cancer cells in vitro at rates similar to the positive control, retrovirally transduced T cells (Fig. 4d). Killing was selective for target cells expressing NY-ESO-1 antigen and the HLA-A*0201 allele, consistent across donors, and depended on the T cells being modified using both the correct gRNA and HDR template (Extended Data Fig. 9n-q).
Finally, we confirmed that non-viral genome targeting could be used to generate NY-ESO-1 TCR cells at scale and that these cells have in vivo anti-tumour function (Fig. 4e and Extended Data Fig. 10a). Given that knock-in efficiency was lower with non-viral targeting than with comparable sized AAV templates4, we first wanted to ensure that we could generate sufficient numbers of NY-ESO-1 positive cells for adoptive cell therapies. We electroporated 100 million T cells from six healthy donors, which after ten days of expansion yielded an average of 385 million NY-ESO-1 TCR T cells per donor (Fig. 4f and Extended Data Fig. 9i-m). NY-ESO-1 TCR knock-in T cells preferentially localized to, persisted at, and proliferated in the tumour rather than the spleen, similar to positive control lentivirally-transduced T cells (Fig. 4g and Extended Data Fig. 10b-f). Adoptive transfer of sorted NY-ESO-1 TCR T cells also reduced the tumour burden in treated animals (Fig. 4h).
Our therapeutic gene editing in human T cells is a process that takes only a short time from target selection to production of the genetically modified T cell product. In approximately one week, novel guide RNAs and DNA repair templates can be designed, synthesized, and the DNA integrated into primary human T cells that remain viable, expandable, and functional. The whole process and all required materials can be easily adapted to good manufacturing practices (GMP) for clinical use. Avoiding the use of viral vectors will accelerate research and clinical applications, reduce the cost of genome targeting, and potentially improve safety.
Looking forward, the technology could be used to “rewire” complex molecular circuits in human T cells. Multiplexed integration of large functional sequences at endogenous loci should allow combinations of coding and non-coding elements to be corrected, inserted, modified, and rearranged. Much work remains to be done to improve our understanding endogenous T cell circuitry if we are going to create synthetic circuits. Rapid and efficient non-viral tagging of endogenous genes in primary human cells will facilitate live-cell imaging and proteomic studies to decode T cell programs. Non-viral genome targeting provides an approach to re-write these programs in cells for the next generation of immunotherapies.
No statistical methods were used to predetermine sample size. For all in vivo experiments experimental conditions were allocated randomly at the time of adoptive transfer, and experimental conditions were mixed among littermates. For in vivo tumour sizing experiments the investigator was blinded to experimental condition. No power analysis was used to determine sample sizes.
All antibodies used in the study for fluorescence activated cell sorting, flow cytometry, and cellular stimulations are listed in Supplementary Table 2.
All guide RNAs used in the study are listed in Supplementary Table 3.
Isolation of human primary T cells for gene targeting
Primary human T cells were isolated from healthy human donors either from fresh whole blood, residuals from leukoreduction chambers after Trima Apheresis (Blood Centers of the Pacific), or leukapheresis products (StemCell). Peripheral blood mononuclear cells (PBMCs) were isolated from whole blood samples by Ficoll centrifugation using SepMate tubes (STEMCELL, per manufacturer’s instructions). T cells were isolated from PBMCs from all cell sources by magnetic negative selection using an EasySep Human T Cell Isolation Kit (STEMCELL, per manufacturer’s instructions). Unless otherwise noted, isolated T cells were stimulated as described below and used directly (fresh). When frozen cells were used, previously isolated T cells that had been frozen in Bambanker freezing medium (Bulldog Bio) per manufacturer’s instructions were thawed, cultured in media without stimulation for 1 day, and then stimulated and handled as described for freshly isolated samples. Fresh blood was taken from healthy human donors under a protocol approved by the UCSF Committee on Human Research (CHR #13-11950). Patient samples used for gene editing were obtained under a protocol approved by the Yale Human Investigation Committee (HIC). Additional leukapheresis products from healthy donors were collected either under UCLA Institutional Review Board (IRB) approval #10-001598 or purchased from AllCells, LLC. All patients and healthy donors provided informed consent.
Primary human T cell culture
Unless otherwise noted, bulk T cells were cultured in XVivo15 medium (STEMCELL) with 5% Fetal Bovine Serum, 50 mM 2-mercaptoethanol, and 10 mM N-Acetyl L-Cystine. Immediately following isolation, T cells were stimulated for 2 days with anti-human CD3/CD28 magnetic dynabeads (ThermoFisher) at a beads to cells concentration of 1:1, along with a cytokine cocktail of IL-2 at 200 U/mL (UCSF Pharmacy), IL-7 at 5 ng/mL (ThermoFisher), and IL-15 at 5 ng/mL (Life Tech). Following electroporation, T cells were cultured in media with IL-2 at 500 U/mL. Throughout the culture period T cells were maintained at an approximate density of 1 million cells per mL of media. Every 2-3 days post-electroporation additional media was added, along with additional fresh IL-2 to bring the final concentration to 500 U/mL, and cells were transferred to larger culture vessels as necessary to maintain a density of 1 million cells/mL.
RNPs were produced by complexing a two-component gRNA to Cas9, as previously described10. Briefly, crRNAs and tracrRNAs were chemically synthesized (Dharmacon, IDT), and recombinant Cas9-NLS, D10A-NLS, or dCas9-NLS were recombinantly produced and purified (QB3 Macrolab). Lyophilized RNA was resuspended in 10 mM Tris-HCL (7.4 pH) with 150 mM KCl at a concentration of 160 μM, and stored in aliquots at −80C. crRNA and tracrRNA aliquots were thawed, mixed 1:1 by volume, and annealed by incubation at 37C for 30 min to form an 80 μNM gRNA solution. Recombinant Cas9 or the D10A Cas9 variant were stored at 40 μM in 20 mM HEPES-KOH pH 7.5, 150 mM KCl, 10% glycerol, 1 mM DTT, were then mixed 1:1 by volume with the 80 μM gRNA (2:1 gRNA to Cas9 molar ratio) at 37C for 15 min to form an RNP at 20 μM. RNPs were electroporated immediately after complexing.
Double stranded DNA HDRT production
Novel HDR sequences were constructed using Gibson Assemblies to insert the HDR template sequence, consisting of the homology arms (commonly synthesized as gBlocks from IDT) and the desired insert (such as GFP) into a cloning vector for sequence confirmation and future propagation. These plasmids were used as templates for high-output PCR amplification (Kapa Hotstart polymerase). PCR amplicons (the dsDNA HDRT) were SPRI purified (1.0X) and eluted into a final volume of 3 μL H2O per 100 μL of PCR reaction input. Concentrations of HDRTs were determined by nanodrop using a 1:20 dilution. The size of the amplified HDRT was confirmed by gel electrophoresis in a 1.0% agarose gel. All homology directed repair template sequences used in the study, both dsDNA and ssDNA, are listed in Supplementary Table 3.
Single stranded DNA HDRT production by exonuclease digestion
To produce long ssDNA as HDR templates, the DNA of interest was amplified via PCR using one regular, non-modified PCR primer and a second phosphorylated PCR primer. The DNA strand that will be amplified using the phosphorylated primer, will be the strand that will be degraded using this method. This makes it possible to prepare either a single-stranded sense or single-stranded antisense DNA using the respective phosphorylated PCR primer. To produce the ssDNA strand of interest, the phosphorylated strand of the PCR product was degraded by treatment with two enzymes, Strandase Mix A and Strandase Mix B, for 5 minutes (per 1kb) at 37C, respectively. Enzymes were deactivated by a 5 minute incubation at 80C. The resulting ssDNA HDR templates were SPRI purified (1.0X) and eluted in H2O. A more detailed protocol for the Guide-it™ Long ssDNA Production System (Takara Bio USA, Inc. #632644) can be found at the manufacturer’s website.
Single stranded DNA HDRT production by reverse synthesis
ssDNA HDR templates were synthesized by reverse transcription of an RNA intermediate followed by hydrolysis of the RNA strand in the resulting RNA:DNA hybrid product, as described21. Briefly, the desired HDR donor was first cloned downstream of a T7 promoter and the T7-HDR donor sequence amplified by PCR. RNA was synthesized by in vitro transcription using HiScribe T7 RNA polymerase (New England Biolabs) and reverse-transcribed using TGIRT-III (InGex). Following reverse transcription, NaOH and EDTA were added to 0.2 M and 0.1 M respectively and RNA hydrolysis carried out at 95C for 10 min. The reaction was quenched with HCl, the final ssDNA product purified using Ampure XP magnetic beads (Beckman Coulter) and eluted in sterile RNAse-free H2O. ssDNA quality was analysed by capillary electrophoresis (Bioanalyzer, Agilent).
Primary T cell electroporation
RNPs and HDR templates were electroporated 2 days following initial T cell stimulation. T cells were harvested from their culture vessels and magnetic anti-CD3/anti-CD28 dynabeads were removed by placing cells on an EasySep cell separation magnet for 2 minutes. Immediately prior to electroporation, de-beaded cells were centrifuged for 10 minutes at 90g, aspirated, and resuspended in the Lonza electroporation buffer P3 using 20 μL buffer per one million cells. For optimal editing, one million T cells were electroporated per well using a Lonza 4D 96-well electroporation system with pulse code EH115. Alternate cell concentrations from 200,000 up to 2 million cells per well resulted in lower transformation efficiencies. Alternate electroporation buffers were used as indicated, but had different optimal pulse settings (EO155 for OMEM buffer). Unless otherwise indicated, 2.5 μL of RNPs (50 pmols total) were electroporated, along with 2 μL of HDR Template at 2 μg/μL (4 μg HDR Template total).
The order of cell, RNP, and HDRT addition appeared to matter (Extended Data Fig. 1). For 96-well experiments, HDRTs were first aliquoted into wells of a 96-well polypropylene V-bottom plate. RNPs were then added to the HDRTs and allowed to incubate together at RT for at least 30 seconds. Finally, cells resuspended in electroporation buffer were added, briefly mixed by pipetting with the HDRT and RNP, and 24 μLs of total volume (cells + RNP + HDRT) was transferred into a 96 well electroporation cuvette plate. Immediately following electroporation, 80 μLs of pre-warmed media (without cytokines) was added to each well, and cells were allowed to rest for 15 minutes at 37C in a cell culture incubator while remaining in the electroporation cuvettes. After 15 minutes, cells were moved to final culture vessels.
Flow cytometry and cell sorting
Flow cytometric analysis was performed on an Attune NxT Acoustic Focusing Cytometer (ThermoFisher) or an LSRII flow cytometer (BD). Fluorescence activated cell sorting was performed on the FACSAria platform (BD). Surface staining for flow cytometry and cell sorting was performed by pelleting cells and resuspending in 25 μL of FACS Buffer (2% FBS in PBS) with antibodies at the indicated concentrations (Supplementary Table 2) for 20 minutes at 4C in the dark. Cells were washed once in FACS buffer before resuspension.
Samples were prepared by drop casting 10 μl of a solution of suspended live T cells onto a 3×1” microscope slide onto which a 25 mm2 coverslip was placed. Imaging was performed on an upright configuration Nikon A1r laser scanning confocal microscope. Excitation was achieved through a 488 nm OBIS laser (Coherent). A long working distance (LWD) 60x Plan Apo 1.20 NA water immersion objective was used with additional digital zoom achieved through the NIS-Elements software. Images were acquired under “Galvano” mirror settings with 2x line averaging enabled and exported as TIFF to be analyzed in FIJI (ImageJ, NIH).
CUT&RUN was performed using epitope-tagged primary human T cells 11 days after electroporation and 4 days after re-stimulation with anti-CD3/anti-CD28 dynabeads (untagged cells were not electroporated). Approximately 20% and 10% of electroporated cells showed GFP-BATF expression as determined by flow cytometry in donor 1 and donor 2 samples, respectively. CUT&RUN was performed as described12, using anti-GFP (ab290), anti-BATF (sc-100974), and rabbit anti-mouse (ab46540) antibodies. Briefly, 6 million cells (30 million cells for anti-GFP CUT&RUN in GFP-BATF-containing cells) were collected and washed. Nuclei were isolated and incubated rotating with primary antibody (GFP or BATF) for 2 hours at 4C. BATF CUT&RUN samples were incubated an additional hour with rabbit anti-mouse antibody. Next, nuclei were incubated with proteinA-micrococcal nuclease (kindly provided by the Henikoff lab) for one hour at 4C. Nuclei were equilibrated to 0C and MNase digestion was allowed to proceed for 30 minutes. Solubilized chromatin CUT&RUN fragments were isolated and purified. Paired-end sequencing libraries were prepared and analysed on Illumina Nextseq machines and sequencing data was processed as described12. For peak calling and heatmap generation, reads mapping to centromeres were filtered out.
TLA sequencing and analysis
TLA sequencing was performed by Cergentis as previously described16. Similarly, data analysis of integration sites and transgene fusions was performed by Cergentis as previously described16. TLA sequencing was performed in two healthy donors, each edited at the RAB11A locus with either a dsDNA or ssDNA HDR template to integrate a GFP fusion (Fig. 1b). Sequencing reads showing evidence of primer dimers or primer bias (i.e. greater than 99% of observed reads came from single primer set) were removed.
In vitro Treg suppression assay
CD4+ T cells were enriched using the EasySep Human CD4+ T cell enrichment kit (STEMCELL Technologies). CD3+CD4+CD127loCD45RO+TIGIT+ enriched Treg-like cells from IL2RA-deficient subjects and HD as well as CD3+CD4+IL2RAhiCD127lo Tregs from IL2RA+/− individuals were sorted by flow cytometry. CD3+CD4+IL2RA-CD127+ responder T cells (Tresps) were labeled with CellTrace CFSE (Invitrogen) at 5 μM. Tregs and HD Tresps were co-cultured at a 1:1 ratio in the presence of beads loaded with anti-CD2, anti-CD3 and anti-CD28 (Treg Suppression Inspector; Miltenyi Biotec) at a 1 bead: 1 cell ratio. On days 3.5 to 4.5, co-cultures were analyzed by FACS for CFSE dilution. % inhibition is calculated using the following formula: 1 - (% proliferation with Tregs / % proliferation of stimulated Tresps without Tregs).
Sorting and TSDR analysis of corrected Tregs
Ex-vivo expanded Tregs and T effector cells from a healthy control and a patient with IL2RA compound heterozygous mutations (D6) were thawed and stained. Live cells were sorted based on expression of CD25 and CD62L markers directly into ZymoResearch M-digestion Buffer (2x) (cat# D5021-9) supplemented with proteinase K. The lysate was incubated at 65°C for greater than 2 hours and then frozen. Bisulfite conversion and pyrosequencing of the samples was performed by EpigenDx (assay ID ADS783-FS2) to interrogate the methylation status of 9 CpG sites intron 1 of the FOXP3 gene, spanning −2330 to −2263 from ATG.
Generation of retrovirally and lentivirally transduced control T cells
For retroviral infections, clinical grade MSGV-1-1G4 (NY-ESO-1 TCR transgene) retroviral vector (IUVPC, Indianapolis, IN) was used. For lentiviral production, HEK 293 cells were plated at 18 million cells in 15 cm dishes the night before transfection. Cells were transfected using the lipofectamine 3000 reagent following the manufacturer’s protocol (L3000001). Transfection media was changed the following day to fresh HEK 293 media (DMEM + 5% FBS + 1% pen/strep) with viral boost reagent per the manufacturer’s protocol at 500x (Alstem viral boost reagent #VB100). 48 hours after transfection the viral supernatant was collected, filtered, and the Alstem precipitation solution was added, mixed, and refrigerated at four degrees for four hours, concentrated by centrifugation, and the viral pellet was then resuspended at 100x in cold PBS following the manufacturer’s protocol (lentivirus precipitation solution #VC100).
T cells for viral infection were activated similarly to non-virally edited cells. Both retroviral and lentiviral transductions occurred 48 hours after TCR/cytokine stimulus, followed by expansion in IL-2 similarly to non-virally edited cells. For retroviral transduction, T cells were infected by spinoculation in retronectin (Clontech, Mountain View, CA) coated plates. Control mock-transduced T cells were also generated. For lentiviral transduction, viral concentrate was added to 1X final concentration.
Antigen specific TCR expression analysis
The expression of the NY-ESO-1 TCR was assessed in virally and non-virally modified cells with an NY-ESO-1 specific (SLLMWITQC) dextramer-PE (Immundex, Copenhagen, Denmark) according to the manufacturer’s protocol. Negative dextramer (Immudex, Copenhagen, Denmark) was used as a negative control.
T cell activation and cytokine production analysis
Melanoma cell lines were established from the biopsies of melanoma patients under the UCLA IRB approval #11-003254. Cell lines were periodically screened for mycoplasma contamination as well as authenticated using GenePrint® 10 System (Promega, Madison, WI), and were matched with the earliest passage cell lines. M257 (NY-ESO-1+ HLA-A*0201-), M257-A2 (NY-ESO-1+ HLA-A*0201+) and M407 (NY-ESO-1+ HLA-A*0201+) were cocultured 1:1 with the modified PBMCs in cytokine free media. The recommended amount per test of CD107a-APC-H7 (Supplementary Table 2) antibody was added to the coculture. After 1 hour, half the recommended amount of BD Golgi Plug and BD Golgi Stop (BD bioscience, San Jose, CA) was added to the coculture. After 6 hours, surface staining was performed followed by cell permeabilization using BD cytofix/cytoperm (BD bioscience, San Jose, CA) and intracellular staining according to manufacturer instructions (Supplementary Table 2). Negative dextramer and Fluorescence minus one (FMOs) staining were used as controls.
T Cell in vitro killing assay
M202-nRFP (NY-ESO-1-, HLA-A*0201+), M257-nRFP (NY-ESO-1+ HLA-A*0201-), M257-A2-nRFP (NY-ESO-1+ HLA-A*0201+), M407-nRFP (NY-ESO-1+ HLA-A*0201+), and A375-nRFP (NY-ESO-1+ HLA-A*0201+) melanoma cell lines stably transduced to express nuclear RFP (Zaretsky 2016 NEJM) were seeded approximately 16 hours before starting the coculture (~1500 cells seeded per well). Modified T cells were added at the indicated E:T ratios. All experiments were performed in cytokine free media. Cell proliferation and cell death was measured by nRFP real time imaging using an IncuCyte ZOOM (Essen, Ann Arbor, MI) for 5 days.
In vivo mouse solid tumour model
All mouse experiments were completed under a UCSF Institutional Animal Care and Use Committee protocol. We used 8 to 12 week old NOD/SCID/IL-2Rɣ-null (NSG) male mice (Jackson Laboratory) for all experiments. Mice were seeded with tumours by subcutaneous injection into a shaved right flank of 1×106 A375 human melanoma cells (ATCC CRL-1619). At seven days post tumour seeding, tumour size was assessed and mice with tumour volumes between 15-30 mm3 were randomly assigned to experimental and control treatment groups. Indicated numbers of T cells were resuspended in 100 μl of serum-free RPMI and injected retro-orbitally. For tumour sizing experiments, the length and width of the tumour was measured using electronic calipers and volume was calculated as v = 1/6 * π * length * width * (length + width) / 2. The investigator was blinded to experimental treatment group during sizing measurements. A bulk edited T cell population (5×106) or a sorted NY-ESO-1 TCR+ population (3×106) was transferred as indicated in figures and legends. For bulk edited T cell transfers, lentivirally edited cells generally had a higher percentage of NY-ESO-1 positive cells, so mock-infected cells were added to normalize the percentage of total T cells NY-ESO-1+ to equal that of the bulk population of non-virally edited T cells (~10% NY-ESO-1+). For sorted T cell transfers, NY-ESO-1+ T cells were FACS sorted eight days following electroporation, expanded for two additional days, and frozen (Bambanker freezing medium, Bulldog Bio). Non-virally or lentivirally modified human T cells were then thawed and rested in media overnight prior to adoptive transfer. For flow cytometric analysis of adoptively transferred T cells, single-cell suspensions from tumours and spleens were produced by mechanical dissociation of the tissue through a 70 μm filter. All animal experiments were performed in compliance with relevant ethical regulations per an approved IACUC protocol (UCSF), including a tumor size limit of 2.0 cm in any dimension.
Data and reagent availability
CUT&RUN data has been deposited in GEO as record GSE108600. TLA and amplicon sequencing data is available upon request. Source data for animal experiments (Fig. 4g, h and Extended Data Fig. 10) is provided. Plasmids containing the HDR template sequences used in the study are available through AddGene.
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We thank members of the Marson Lab, Chris Jeans (QB3 MacroLab), Kyle Marchuk (UCSF BIDC), Jeffrey Bluestone and Qizhi Tang (UCSF Regulatory T Cell Therapy Program), Ryan Wagner (Parnassus CAT), the UCSF Flow Cytometry Core (NIH P30 DK063720 and 1S10OD021822-01), Lonza, Jacob Corn and Sarah Pyle for suggestions and assistance. This research was supported by NIH grants DP3DK111914-01 (A.M.), P50GM082250 (A.M.), R35 CA197633 (A.R.), K23 DK094866 (S.W.G.), T32GM007618 (T.L.R, J.H.), T32 DK007418 (T.L.R.), and P30 DK020595 (S.W.G.), the NIH NCI Intramural Program (A.L.F., S.H.H.), grants from the Keck Foundation (A.M.), National Multiple Sclerosis Society (A.M.; CA 1074-A-21), gifts from Jake Aronov, Galen Hoskin, the Jeffrey Modell Foundation (A.M), and awards from the Burroughs Wellcome Fund (A.M.) and the Ressler Family Fund (C.P.S., J.S., A.R.). A.M. is a Chan Zuckerberg Biohub investigator. A.R. is a Parker Institute for Cancer Immunotherapy member.
Competing Financial Interests
A.M. is a co-founder of Spotlight Therapeutics. A.M. serves as an advisor to Juno Therapeutics and PACT Pharma and the Marson laboratory has received sponsored research support (Juno Therapeutics, Epinomics, Sanofi) and a gift from Gilead. T.L.R., C.P.S., E.S., A.R., and A.M. are inventors on new patent applications related to this manuscript.