Abstract
Human T cells are central to physiological immune homeostasis, which protects us from pathogens without collateral autoimmune inflammation. They are also the main effectors in most current cancer immunotherapy strategies1. Several decades of work have aimed to genetically reprogram T cells for therapeutic purposes2–5, but as human T cells are resistant to most standard methods of large DNA insertion these approaches have relied on recombinant viral vectors, which do not target transgenes to specific genomic sites6, 7. 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), but to date in human T cells this still requires viral transduction8, 9. Here, we developed a non-viral, CRISPR-Cas9 genome targeting system that permits the rapid and efficient insertion of individual or multiplexed large (>1 kilobase) DNA sequences at specific sites in the genomes of primary human T cells while preserving cell viability and function. We successfully tested the potential therapeutic use of this approach in two settings. First, we corrected a pathogenic IL2RA mutation in primary T cells from multiple family members with monogenic autoimmune disease and demonstrated enhanced signalling function. Second, we replaced 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 recognized the tumour antigen, with concomitant cytokine release and tumour cell killing. Taken together, these studies provide preclinical evidence that non-viral genome targeting will 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 DNA10. While the introduction of short single stranded oligodeoxynucleotide (ssODN) HDR templates does not cause significant T cell death, it has been shown that larger linear double stranded (dsDNA) templates are toxic at high concentrations11, 12. Contrary to expectations, we found that co-electroporation of human primary T cells with CRISPR-Cas9 ribonucleoprotein (Cas9 RNP13, 14) complexes and long (>1kb) linear dsDNA templates reduced the toxicity associated with the dsDNA template (Extended Data Fig 1). 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). Systematic exploration of this approach while optimizing for both viability and efficiency (Fig. 1b and Extended Data Fig. 2) resulted in GFP expression in ~50% of cells in both primary human CD4+ and CD8+ T cells. The method was reproducibly efficient while maintaining high cell viability and expandability (Fig. 1c, d, e, and Extended Data Fig. 3). The system is also compatible with current manufacturing protocols for cell therapies as it could be applied to fresh or cryopreserved cells, bulk T cells or FACS-sorted sub-populations, and cells from whole blood or leukapheresis (Extended Data Fig. 4).
We next confirmed that the system could be applied broadly by targeting sequences in different locations throughout the genome. We efficiently engineered GFP+ primary T cells by generating fusions with different genes (Fig. 2a and Extended Data Fig. 5). Live-cell imaging with confocal microscopy confirmed the specificity of gene targeting, revealing the distinct sub-cellular locations of each resulting GFP-fusion protein15 (Fig. 2b). Appropriate chromatin binding of a transcription factor GFP-fusion protein was assessed by performing genome-wide CUT & RUN16 analysis with an anti-GFP antibody (Fig. 2c and Extended Data Fig. 6). Finally, we validated that gene targeting preserved endogenous gene regulation. Consistent with correct cell-type specific expression, a CD4-GFP fusion was selectively expressed in CD4+ T cells, but not in CD8+ 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. 7) or multiplex modification of two (Fig. 2f and Extended Data Fig. 8) or even three (Fig. 2e and Extended Data Fig. 8) different genes17, 18. Individual or multiple endogenous genes can therefore be directly engineered without virus in T cells, preserving gene and protein regulation.
For therapeutic use of genetically modified T cells, integrated sequences should be introduced specifically without unintended disruption of other critical genome sites19. We performed targeted locus amplification (TLA) sequencing20 and found no evidence of off-target integrations above the assay’s limit of detection (~1% of alleles) (Extended Data Fig. 9). We further assessed potential off-target integrations at the single cell level by quantifying GFP+ cells generated with a Cas9 RNP that cuts outside the homology site. Similar to what has been described with viral HDR templates8, 21, we found evidence to suggest that double-stranded templates could integrate independent of target homology22, 23, albeit at low rates (Extended Data Fig. 10). These rare events could be reduced almost completely by using single-stranded DNA templates24, 25 (Extended Data Fig. 11). As an additional safeguard that could be important for some applications, we demonstrated that 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 cuts26, 27 (Extended Data Fig. 12).
Having optimized this non-viral gene 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 for therapeutic effect in patients. Specifically, we tested the ability to rapidly and efficiently correct an inherited T cell genetic alteration and we also tested the targeted insertion of the two chains of a TCR to genetically redirect the specificity of T cells to recognize cancer.
We identified a family with monogenic primary immune dysregulation with autoimmune disease caused by recessive loss-of-function mutations in the gene encoding the IL-2 alpha receptor (IL2RA)28(Extended Data Table 1), which is essential for healthy regulatory T cells (Tregs)29, 30 (Extended Data Fig. 13). Whole exome sequencing revealed that the IL2RA-deficient children harboured compound heterozygous mutations in IL2RA (Fig. 3a and Extended Data Fig. 14). One mutation, c.530A>G, creates a premature stop codon. With non-viral genome targeting, we were able to correct the mutation and observe 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 the efficiency of correction was lower than with the longer dsDNA template (Extended Data Fig. 15, 16). 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 (Extended Data Table 1 and Extended Data Fig. 17). 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 small indels restoring the reading frame (Extended Data Fig. 16). 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. 15, 16). In addition, following correction, the fraction of total T cells expressing both IL2RA and FOXP3, a critical transcriptional factor in Tregs, returned to levels similar to healthy controls (Extended Data Fig. 15, 16). We were also able to correct IL2RA in a sorted population of CD3+CD4+CD127loTIGIT+CD45RO+ Treg-like patient cells (Extended Data Fig. 15), a potential strategy for 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 c/s-regulatory elements that constitute a super-enhancer31, 32. Therefore, therapeutic correction of IL2RA is likely to depend on specifically 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 applied this advantage of the system 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 template8. Targeting of specific TCR sequences to this locus has not been achieved and is challenging because a cell must express paired TCR alpha (TCR-α) and beta chains (TCR-ß) 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-a sequence when integrated at the endogenous TRAC exon (Extended Data Fig. 18). To test this strategy, we introduced a TCR-β and TCR-α pair (1G4) recognizing the NY-ESO-1 tumour antigen into the TRAC locus of polyclonal T cells isolated from healthy human donors33. 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. 19). The majority of T cells that did not show NY-ESO-1 TCR expression 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 specifically 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-L and TNF-a and induced T cell degranulation (measured by CD107a surface expression) (Fig. 4c). Cytokine production and degranulation only occurred when the NY-ESO-1 TCR transgenic 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. 19).
Finally, we confirmed the anti-tumour cell killing ability of T cells following nonviral endogenous TCR replacement. NY-ESO-1 TCR knock-in T cells rapidly killed target M257-HLA-A*0201 cancer cells at similar rates to the positive control, retrovirally transduced T cells (Fig. 4d). Killing was NY-ESO-1 antigen-specific, consistent across donors, and dependent on the T cells being modified using both the correct gRNA and HDR template (Extended Data Fig. 20).
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 need for 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 hope to build 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.
METHODS
Data reporting
No statistical methods were used to predetermine sample size. The experiments were not randomized and the investigators were not blinded to allocation during experiments and outcome assessment.
Antibodies
All antibodies used in the study for fluorescence activated cell sorting, flow cytometry, and cellular stimulations are listed in Supplementary Table 1.
Guide RNAs
All guide RNAs used in the study are listed in Supplementary Table 2.
Isolation of human primary T cells for gene targeting
Primary human T cells were isolated from healthy human donors either from fresh whole blood samples, 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 with their informed consented 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 Internal Review Board (IRB). PBMCs for retroviral transduction experiments and corresponding TCR knockins were obtained from leukapheresis products of healthy donors. The leukapheresis product were collected either under UCLA Institutional Review Board (IRB) approval #10-001598 or purchased from AllCells, LLC.
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.
RNP production
RNPs were produced by complexing a two-component gRNA to Cas9, as previously described14. 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 highoutput 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 2.
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 described25. 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 cel 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 electroporatior 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-wel polypropylene V-bottom plate. RNPs were then added to the HDRTs and allowed tc 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 wel electroporation cuvette plate. Immediately following electroporation, 80 μLs of prewarmed 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 Accustic Focusing Cytometei (ThermoFisher) or an LSRII FACs machine (BD). Fluorescence activated cell sorting was performed on an Aria II Cell Sorter (BD). Surface staining for flow cytometry anc 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 1) for 20 minutes at 4C in the dark. Cells were washed once in FACS buffer before resuspension.
Confocal microscopy
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
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 described16, 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 described16. 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 described20. Similarly, data analysis of integration sites and transgene fusions was performed by Cergentis as previously described20. 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).
Generation of retrovirally transduced PBMCs control cells
PBMCs from healthy donors were thawed, activated with CD3/CD28 beads (Invitrogen, Carlsbad, CA) at a 1:1 ratio and expanded in T cell media (RPMI supplemented with 10% fetal bovine serum, penicillin-streptomycin and 300IU/mL of hIL2). Forty-eight hours after activation, PBMCs were transduced with clinical grade MSGV-1-1G4 (NY-ESO-1 TCR transgene) retroviral vector (IUVPC, Indianapolis, IN) by spinoculation in retronectin (Clontech, Mountain View, CA) coated plates. Control mock-transduced PBMCs were generated. PBMCs were expanded for 8 days after transduction, aliquoted and stored in liquid nitrogen.
Antigen specific TCR expression analysis
Modified PBMCs were thawed and expanded in T cell media supplemented with 300IU/mL hIL2. To assess the expression of the NY-ESO-1 TCR cells were stained with NY-ESO-1 specific (SLLMWITQC) dextramer-PE (Immundex, Copenhagen, Denmark), 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 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 1) 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 1). Negative dextramer and Fluorescence minus one (FMOs) staining were used as controls.
T Cell killing assay
M257-nRFP, M257-A2-nRFP, A375-nRFP, and M407-nRFP melanoma cell lines stably transduced to express nuclear RFP (Zaretsky 2016 NEJM) were seeded approximately 16 hours before starting the coculture. 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. Biological quadruplicates per condition were assessed.
Data availability
CUT&RUN data will be publically deposited prior to publication. TLA sequencing data is available upon request.
AUTHOR INFORMATION
Contributions
T.L.R. and A.M. designed the study and wrote the manuscript. T.L.R. designed and performed all electroporation experiments. T.L.R., R.Y., E.S. J.H. J.L. V.T., D.N., and K.S. contributed to functional assays of edited T cells. R.Y. performed and analyzed CUT&RUN experiments. H.L., J.W., and M.L. developed the IVT-RT ssDNA production method. H.M., M.M, Y.M, B.S, and M.H. developed the exonuclease based ssDNA production method. R.Q. and C.G. advised on the use of ssDNA. A.M.F. and S.H.H. advised on methods of DNA introduction into T cells. A.P.M. advised on integration site analysis. J.C., J.N.S., L.P., D.C, G.A.A., D.D.G., G.K., S.G., R.B., E.M., M.G.R., N.R., and K.C.H. contributed to the clinical workup of the IL2RA deficient family and functional assays on unedited patient T cells. T.L.R., C.P.S., E.S., A.R., and A.M. designed the endogenous TCR knock-in strategy. T.L.R., C.P.S., J.C., J.S., A.A., and A.R. performed or supervised functional assays of T cells with endogenous TCR knock-ins.
Competing Financial Interests
The authors declare 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 from Juno Therapeutics and Epinomics. Patents have been filed based on the findings described here.
Corresponding Author
Correspondence and requests for materials should be addressed to alexander.marson{at}ucsf.edu.
ACKNOWLEDGEMENTS
We thank members of the Marson lab, Chris Jeans and the QB3 MacroLab, Kyle Marchuk and the UCSF Biological Imaging Development Center, Jeffrey Bluestone and Jacob Corn for suggestions and technical assistance. We thank Lonza for technical assistance and providing reagents to test electroporation conditions. This research was supported by NIH grants DP3DK111914-01 (A.M.), P50GM082250 (A.M.) and R35 CA197633 (A.R.), a grant from the Keck Foundation (A.M.), gifts from Jake Aronov and Galen Hoskin (A.M.), a gift from the Jeffrey Modell Foundation, and a National Multiple Sclerosis Society grant (A.M.; CA 1074-A-21). T.L.R. and J.H. were supported by the UCSF Medical Scientist Training Program (T32GM007618). T.L.R. was supported by the UCSF Endocrinology Training Grant (T32 DK007418). A.M. holds a Career Award for Medical Scientists from the Burroughs Wellcome Fund and is an investigator at the Chan Zuckerberg Biohub. C.P.S., J.S. and A.R. are funded by the Ressler Family Fund. A.R. is a member of the Parker Institute for Cancer Immunotherapy. The UCSF Flow Cytometry Core was supported by the Diabetes Research Center grant NIH P30 DK063720. SHH and AMF were supported by the NIH Intramural Program, Center for Cancer Research, National Cancer Institute.