We evaluated a 33-year-old male of East-Asian origin. He originally presented to gastroenterology services at the age of 19 years with chronic diarrhea. Following initial assessment, which included normal colonoscopy and negative coeliac screen, he was found to have severe panhypogammaglobulinaemia and was referred to clinical immunology for further investigations. On further questioning the patient reported a 5-year long history of recurrent lower respiratory tract infections requiring treatment with antibiotics. He also reported having a left lower lobe pneumonia at 15 years of age. He had no other significant infection history and reported being generally well during his childhood. His only other past medical history of note was appendicectomy at the age of 10 years. He had no history of developmental difficulties, dental, skin or any musculoskeletal abnormalities. He was an only child from non-consanguineous parents with no family history of PID. Shortly after presenting to immunology, he developed a left hip and iliac fossa pain and was found to have multiloculated left psoas abscess which required a computerized tomography (CT) guided drainage and prolonged antibiotic treatment. This infective episode was complicated by pulmonary embolism (incidental finding on a repeat CT scan) for which he was anticoagulated for a period of 6 months. He had no subsequent history of thromboembolic events. A detailed CT scan of his chest showed cylindrical bronchiectasis of the left lower lobe, lingua and R middle lobe. This appearance remains unchanged as shown on a repeat scan 11 years later in 2016 (Figure 1A).

The initial immunological investigations showed profound panhypogammaglobulinaemia with drastically reduced IgG and undetectable IgA and IgM levels (Table 1). The basic lymphocyte profiling showed mild CD4⁺ T cell lymphopenia with adequate numbers of B and NK cells. Subsequent detailed phenotyping revealed reduced proportion of naïve T cells and seemingly adequate percentages of central and effector memory T cells. The proportion of TH1/TH2 and TH17 T cells was comparable to healthy controls (Figure 1—figure supplement 1) and TCR phenotyping showed a normal TCR repertoire (Figure 1B). The patient also had reduced proportion of class switched memory B cells (Table 1). Functional testing showed reduced T cell proliferation to both PHA and anti-CD3 stimulation (Figure 1C and D). Over the next 15 years, there was evidence of progressive CD4⁺ T cell and B cell lymphopenia. Interestingly, biopsy of his large bowel, which was performed in 2011 for investigation of intermittent diarrhoea, showed mild non-specific inflammatory changes, but also absence of the plasma cells.

Following the initial assessment, the patent was given a provisional diagnosis of common variable immunodeficiency (CVID) and treated with regular intravenous immunoglobulin (IVIG) replacement therapy. His clinical progress since the diagnosis has been uncomplicated. His chronic diarrhoea resolved following commencement of IVIG and the ongoing treatment with IVIG has resulted in excellent control of respiratory infections.

The patient was consented for genetic testing and whole-exome sequencing was performed. Full details regarding candidate variant selection are provided in methods. Three potentially damaging rare variants were identified in CRACR2A GRCh37 12:3715799–3873985 ENSG00000130038 ENST00000440314 (c.430A > G; p.R144G, c.898G > T; p.E300*, exons 6/20 and 10/20), which were maternal and c.834G > T; p.E278D in exon 9/20 on the other allele. Only c.834G > T is present in the gnomAD database, though it is considered rare (heterozygous only; 0.00095 in East Asian). All three amino acids are highly conserved in other species.

The variants were confirmed in the genomic DNA by Sanger sequencing the proband and the mother and maternal grandmother and the father, with the pedigree shown in Figure 2A. Compound heterozygous segregation of these variants was confirmed by subcloning of the proband cDNA to isolate the individual alleles. Sanger sequencing confirmed the c.834 G > T variant on a separate allele to the other variants (Figure 2B). The protein domains affected by the mutations are schematically depicted in Figure 2C.

To check whether the mutant proteins are expressed, we performed immunoblot analyses using T cells from patient and healthy control. Peripheral blood mononuclear cells (PBMCs) harvested from the patient and healthy control were stimulated with plate-coated anti-CD3 and anti-CD28 antibodies for 48 hr, following which they were further expanded for a further 4 days and then harvested for making cellular lysates. Western blot analysis using anti-CRACR2A antibodies detected reduced amounts of only the higher molecular weight band (corresponding to the long isoform) in patient samples (Figure 2D, left panel). The antibody was targeting the N-terminal 160 amino acids of CRACR2A, and hence it should detect the ~30 KDa truncated form of the p.E300* variant, if expressed. However, we could not detect presence of any band at ~30 KDa, suggesting possibly degradation of the truncated form. To check for CRACR2A transcript levels, we used primers that were common to both isoforms, however, we detected pronounced reduction in CRACR2A transcript levels from patient samples (Figure 2D, right panel). Further, in healthy control samples, expression of CRACR2A transcripts increased after stimulation with PMA and ionomycin by more than two-fold, but it was not upregulated in patient T cells. These data suggest possible nonsense-mediated mRNA decay for CRACR2A transcripts.

Since the patient exhibited profound panhypogammaglobulinaemia and a relative decline in memory B and plasma cell populations, we next checked whether patient B cells were functional. To test if the panhypogammaglobulinaemia is due to an intrinsic B cell defect, we set up an in vitro culture system with a T cell-dependent (TD) stimulus that enables the generation of mature plasma cells from primary B cells (Figure 3A; Cocco et al., 2012). The capacity of the patient’s B cells to differentiate into plasma cells was comparable to healthy controls, as judged by the acquisition of CD38 and CD138 surface expression, which is associated with the transition to an antibody-secreting state (Figure 3B). Furthermore, assessment of immunoglobin levels in cell culture showed adequate production of IgM and IgG (Figure 3C), suggesting that the failure of antibody production in this case might be secondary to inadequate in vivo T cell help.

Since the patient B cells produced antibodies in vitro, in response to TD antigen, we surmised that panhypogammaglobulinaemia is likely due to defect in T cell function. Previously we showed that Cracr2a KO mice had a defect in effector of T cell functions (Woo et al., 2018). Hence, we further analyzed phenotypes of patient T cells, including TCR-proximal signaling events and cytokine production. Patient CD4⁺ cells produced less IFN-γ than the ones isolated from a representative healthy donor (Figure 4A). We extended our findings using cells from multiple healthy donors and different batches of the patient cells by checking expression of IFN-γ, IL-2, and TNF using quantitative RT-PCR (Figure 4B) and ELISA (Figure 4C). Expression of all these cytokines was impaired in patient T cells compared to control T cells derived from multiple different healthy donors. Since CRACR2A is known to interact with ORAI1 and STIM1, to regulate SOCE and JNK phosphorylation (Srikanth et al., 2010a; Srikanth et al., 2016), we measured SOCE after anti-CD3 cross-linking as well as passive depletion of the ER Ca²⁺ stores using thapsigargin, a SERCA blocker. In both these analyses, patient cells showed significant reduction in SOCE, when compared to those from representative healthy controls (Figure 4D and E). Further, patient T cells also showed a profound defect in JNK phosphorylation upon TCR cross-linking (Figure 4F). To confirm that reduced SOCE in patient cells is not due to altered expression of CRAC channel subunits, we checked expression of ORAI1 and STIM1 in lysates of T cells from healthy controls and patient. Expression of ORAI1 and STIM1 were similar between control and patient T cells as judged by immunoblotting as well as real-time quantitative RT-PCR measurements (Figure 4—figure supplement 1). Together, these results show the conserved role of CRACR2A in effector T cell responses by regulating the Ca²⁺ and JNK pathways in human effector T cells, consistent with the previous finding in murine T cells (Woo et al., 2018).

To gain insight into the phenotypes of mutants derived from individual CRACR2A alleles in the patient, we first cloned the mutant cDNA in a lentiviral vector. When overexpressed in HEK293 cells, E278D and R144G/E300* mutants migrated at the expected molecular weights of ~90 and ~ 30 kDa, respectively (Figure 5—figure supplement 1A). To check their expression without interference from endogenous CRACR2A proteins, we generated CRACR2A KO Jurkat T cells using the lentivirus-based CRISPR/Cas9 system and found that sgRNA#1 and #2 had a great deletion efficiency (Figure 5—figure supplement 1B). When we stably expressed E278D and R144G/E300* mutants in CRACR2A KO Jurkat cells (sgRNA#2), these mutants migrated at their expected molecular weights, similarly with HEK293 cells. Notably, even after repeated transduction the expression of R144G/E300* mutant protein was much lower than WT or E278D CRACR2A, suggesting that the transcripts of this double mutant may be unstable, similar to our observation with patient cells (Figure 2D).

Similar to patient T cells, CRACR2A KO Jurkat T cells showed reduced IL-2 production (Figure 5A). Stable expression of WT CRACR2A in these cells substantially rescued the IL-2 production defect, validating that the decrease was caused due to loss of CRACR2A. However, expression of E278D or R144G/E300* mutants showed very marginal rescue of IL-2 expression in CRACR2A KO Jurkat T cells under the same condition. We also validated these results with primary human CD4⁺ cells. We found that transduction with lentiviral vectors encoding sgRNA to delete CRACR2A resulted in reduced IFN-γ production in primary cells, which was completely rescued by expression of WT CRACR2A (Figure 5B). However, both E278D and R144G/E300* mutants did not rescue cytokine production in the KO primary T cells, similar to our observations with Jurkat T cells. Exogenous expression of WT CRACR2A in primary T cells from healthy donors (in the presence of WT CRACR2A) slightly enhanced IFN-γ production while the mutants did not (Figure 5C). On the contrary, overexpression of the R144G/E300* showed significant reduction in cytokine production, suggesting that this mutant may impair function of endogenous CRACR2A. Overall, our cytokine analysis suggested that expression of mutants derived from each of the CRACR2A alleles could not rescue cytokine production in CRACR2A KO T cells.

We next checked whether individual mutants could rescue TCR-proximal events in CRACR2A KO T cells. Earlier we showed that loss of CRACR2A impaired TCR stimulation-induced SOCE and JNK phosphorylation in T cells (Srikanth et al., 2016). Accordingly, CRACR2A KO Jurkat T cells showed reduced Ca²⁺ entry triggered by TCR stimulation (Figure 6A). Expression of WT CRACR2A fully rescued the decreased level of Ca²⁺ entry in KO cells while that of R144G/E300* and E278D mutants did not. We observed similar results when SOCE was triggered by passive depletion of intracellular Ca²⁺ stores using thapsigargin (Figure 6B). Interestingly, these two mutants behaved differently in activation of the JNK signaling pathway. While WT and the E278D mutant rescued phosphorylation of JNK, R144G/E300* mutant failed to do so (Figure 6C). These results indicate that while R144G/E300* mutant has a defect in both Ca²⁺ entry and JNK activation due to truncation of the coiled coil, proline-rich and Rab GTPase domains, E278D mutant has a selective defect in Ca²⁺ signaling.

The C-terminal Rab GTPase domain of CRACR2a contains the conserved residues required for GTP binding and hydrolysis as well as a prenylation site for membrane anchoring (Figure 2C). The N-terminal EF-hand motif and coiled-coil domains are involved in sensing intracellular Ca²⁺ levels and stabilizing ORAI1-STIM1 interaction (Srikanth et al., 2010a). Also, the longer isoform, CRACR2A, which localizes to intracellular vesicles, translocates to the immunological synapse, and interacts with a signaling adaptor molecule, Vav1, through its proline-rich domain (PRD) (Srikanth et al., 2016).

To gain insight into the mechanism underlying the defective function of the mutants, we checked the interaction of these mutants with three known interacting partners, ORAI1, STIM1, and Vav1 using co-immunoprecipitation. These results showed that the R144G/E300* mutant failed to interact with all the molecules, whereas the E278D mutant retained binding to all three partners, similar to WT CRACR2A (Figure 7A and B). Confocal analysis showed that WT CRACR2A and E278D mutant localized in the proximal Golgi area and translocated to the interface of anti-CD3 antibody-coated coverslips after stimulation (Figure 7C). However, the R144G/E300* mutant showed a diffuse cytoplasmic localization and was not enriched at the interface in stimulated cells. These results suggest that while E278 behaves similar to WT CRACR2A in interacting with binding partners (under our experimental conditions) as well as intracellular localization under resting and stimulated conditions, the R144G/E300* mutant is impaired in all these functions. Deletion of protein interaction domains including the PRD and the C-terminal prenylation site is likely the cause loss of binding to its partners and cytoplasmic localization of the DM. Even though E278D retained binding to ORAI1/STIM1, it could not reconstitute SOCE or cytokine production in CRACR2A KO Jurkat T cells, suggesting that subtle changes in the protein conformation due to this mutation may alter its affinity for interaction with ORAI1/STIM1 under physiological conditions, which is not recapitulated in our biochemical studies performed using overexpression.

Genomic DNA was purified from whole blood using QIAamp DNA Blood kit (Qiagen) with quality and concentration assayed using a TapeStation (Agilent). Exome sequence data was produced by batch sequencing with other non-related samples. Samples were sonicated using a Covaris E220 Focused-ultrasonicator to fragment the DNA to a median size of approximately 250–300 bp. Libraries were generated using Agilent SureSelect XT Human All Exon V5, using a standard protocol, receiving a unique index sequence and were then pooled to form equimolar pools. These were sequenced on a 150 bp paired end Illumina HiSeq 3,000 (Illumina, San Diego, CA, United States) with an average depth of coverage of x30.

Exome sequences were trimmed and checked for quality control using CutAdapt (Andrews, 2010) and FastQC (Andrews, 2010) via Trim-galore (Krueger, 2015). Reads were aligned to GrCh37 using BWA-MEM (Li and Durbin, 2009). GATKv3 best practices were applied for genomic analysis. Reads were sorted and marked with Picard SortSam and MarkDuplicates (Broad Institute), respectively. The read alignments were readjusted for optimal local alignment using IndelRealigner (GATK), Base Quality Score Recalibration (GATK) was used to estimate base call accuracy by detecting systematic sequencing errors, followed by (GATK) PrintReads. HaplotypeCaller was used to produce gVCFs via local re-assembly of haplotypes. GenotypeGVCFs (GATK) was used for joint genotyping with other non-related samples and in-house controls for an analysis group size of 34 (Auwera et al., 2018). Variant Quality Score Recalibration (GATK) was used to filter out probable artifacts from the callset. A total of 719,868 variants were present for the combined cohort, with 49,891 of these variants occurring in the proband sample.

Filtering and prediction of functional consequences was performed by annotating data using Variant Effect Predictor (VEP) (McLaren et al., 2016), Exome Variant Server (EVS), The Single Nucleotide Polymorphism database (dbSNP), ClinVar, and the Exome Aggregation Consortium and the Genome Aggregation Database (gnomAD) (Lek et al., 2016). Filtering of common variations and annotation terms was performed using the tool VCFhacks (Parry, 2015). Candidate variants were required to pass the following filtering conditions: read frequency (count/coverage) between 20 and 100%, according to VEP-annotation at least one canonical transcript is affected with one of the following consequence: variants of the coding sequence, frameshift, missense, protein altering, splice acceptor, splice donor, or splice region; an in-frame insertion or deletion; a start lost, stop gained, or stop retained, or according to VEP a gnomAD frequency of unknown, less of equal to 0.01, or with clinical significance ‘path’.Cohort-specific filtering retained functional variants that were present in at least one case but absent for in-house filtering controls or having indication of misalignment or index swapping. The resulting candidate list of variants consisted of rare variants that were of unknown or potentially damaging significance.

Candidate variants were also annotated with terms from Gene Ontology, GeneRIF, Biogrid interaction, OMIM, NCBI summary, MGI phenotype, MorbidMap, VOC MammalianPhenotype, UniProtKB, PDB, and HMD human phenotype (Parry, 2015). Variants were flagged if they had any annotation related to the disease phenotype, were predicted pathogenic, or were in a gene known to cause immunological responses. A filter for all known PID genes was also used to flag any candidates that may have been otherwise excluded. A total of 48 recessive candidate variants (compound heterozygous of homozygous) were identified in 34 genes. Filtering based on gene annotation removed genes with no known immunological significance.

The top gene candidate, calcium release activated channel regulator 2 A (CRACR2A) ENSG00000130038 (also annotated by previous gene symbol EFCAB4B), contained three variants; ENST00000440314 (1) missense c.430A > G, p.R144G, exon 6/20, novel, 12:3788175, Ref T, Alt C, (2) missense c.834G > T, p.E278D, exon 9/20, gnomAD 0.0001647 rs541326469, 12:3765501 Ref C, Alt A, and (3) stop-gained c.898G > T p.E300*, exon 10/20, novel, 12:3763526 Ref C, Alt A. LDpop Tool via LDlink (with appropriate population EAS) was used to guide interpretation prior to confirmation by genomic DNA sequencing of relatives. Chr12:3763526 (p.E300*) and chr12:3788175 (p.R144G) were not in dbSNP build 151, indicating potential LD, inherited together from one parent and in trans with 12:3765501 (p.E278D). The inheritance pattern (p.[R144G;E300*];[E278D]) was confirmed by Sanger sequencing of genomic DNA in the proband, mother, maternal grandmother, and the father. Amplification of genomic DNA for Sanger sequencing was performed by the standard PCR method. PCR clean-up was performed with ExoSAP-IT (Affymetrix). Sanger sequencing was then done using the same primers; Exon 6, (F- ATGATTCCTGGCAGGTGAGA, R- ATTCCAGTGCAGGGACCAG), Exon 9 (F- GGCCCTGATGTTGAGTAGGT, R- GTGAATGGCAGGGAAAGTGG), and Exon 10 (F- AAACAAGGTGAGGCCAGGG, R- AGCCCAAATCCTCTTTTCACAG). Sanger sequencing using BigDye Terminator Cycle Sequencing Kit, version 3.1 (Applied Biosystems) and analysis on an ABI 3130XL DNA analyzer (Applied Biosystems).

Two candidate variants were also identified in UNC13D, a gene reported as causal for the recessive PID Familial hemophagocytic lymphohistiocytosis syndromes (as reported in Genomics England panelapp). However, one variant was present at ~1% in the patient genetic ancestry population and not considered as potentially damaging. Variant 1: ENST00000207549.4 p.Arg411Gln, Polyphen - benign, SIFT – tolerated. East Asian allele frequency 0.008. gnomAD link 1; variant 2: East Asian allele frequency 0. gnomAD link 2 ENST00000207549.4 HGVSpc.753 + 1 G> T, pLoF - high-confidence, East Asian allele frequency 0, gnomAD link 2. No other candidate biallelic variants were considered causal bases on annotation or variant effect. No dominant gain-of-function variants were considered as causal candidates.

Fura 2-AM was purchased from Invitrogen (Carlsbad, CA). Thapsigargin, PMA (Phorbol 12-myristate 13-acetate), and ionomycin were purchased from EMD Millipore. Brefeldin A was purchased from eBioscience. Antibody for detection of CRACR2A (15206–1-AP) was purchased from Proteintech. Antibodies for detection of Vav1 (2502), STIM1 (5668) and His tag (12,698 S) were purchased from Cell Signaling Technologies. Antibodies for detection of FLAG tag (F3040) and ORAI1 (AB9868) were purchased from Millipore Sigma. Antibody for detection of β-actin (sc-47778) was obtained from Santa Cruz Biotechnology.

Full-length cDNA of human CRACR2A-a (NCBI Reference Sequence: NM_001144958.1) was cloned into a lentiviral vector, FGllF (kind gift from Dr. Dong Sun An, UCLA) with a N-terminal FLAG tag as previously described (Srikanth et al., 2010a; Srikanth et al., 2016). Various mutants of CRACR2A-a were generated by PCR amplification and site-directed mutagenesis using primers described in Supplementary file 1. All the clones were verified by sequencing. Myc-ORAI1 and Myc-STIM1 plasmids had been described previously (Gwack et al., 2007; Srikanth et al., 2019). Vav1-GFP clone was purchased from Addgene. HEK293 and Jurkat E6-1 T cell lines were obtained from American Type Culture Collection Center (ATCC, Manassas, VA).

PBMCs were isolated from peripheral blood by density dependent centrifugation with Lymphoprep (Axis Shield, Norway). Total B cells were isolated by negative selection with a memory B cell isolation kit according manufacturer’s instructions (Miltenyi Biotech, USA). B cells were cultured as previously described (Cocco et al., 2012) using gamma-irradiated CD40L-L cells and 2 µg/ml F(ab′)two goat anti-human IgM/IgG/IgA (Jackson ImmunoResearch) supplemented with 20 U/ml IL-2 (Roche) and 50 ng/ml IL-21 (Peprotech) as the initial T-depedent stimulus. Cells were monitored for changes in cell surface antigen expression using anti-CD38-APC-Cy7 (HB-7; BD Biosciences), anti-CD138-APC (44F9; Miltenyi Biotec), as well as 7-AAD-PerCP-Cy5 (BD Biosciences) to identify live cells. Flow cytometry was performed using a or a CytoFLEX S or LX (Beckman Coulter). Analysis was performed using FlowJo v10 (TreeStar). Supernatants were collected at day 6 and day 13 from the differentiation assays and assessed for Ig levels using human IgM ELISA Quantitation Set (E80-100) or Human IgG ELISA Quantitation Set (E80-104) (Bethyl Laboratories Inc, USA) according to manufacturer’s instructions on a Berthold 96-well plate reader. ELISA absorbance values were analyzed at 450 nm and Ig concentrations calculated from standard curves.

To generate lentiviruses for transduction, HEK293T cells were transfected with plasmid(s) encoding sgRNA and packaging vectors (pMD2.G and psPAX2 – from Addgene) using calcium phosphate transfection method. Cas9 encoding lentivirus was generated using the same technique. Culture supernatants were harvested at 48 and 72 hr post transfection and used for infection (50% of cas9-encoding virus +50% of sgRNA-encoding virus) of Jurkat T cells together with polybrene (8 µg/ml) using the spin-infection method. Cells were selected with puromycin (1 µg/ml) and blasticidin (5 µg/ml) 48 hrs post infection. The sgRNA sequences are described in Supplementary file 1.

Jurkat or human PBMCs were loaded at 1 × 10⁶ cells/ml with 1 μM Fura 2-AM for 30 min at 25 °C and attached to poly-D-lysine-coated coverslips. Intracellular [Ca²⁺] measurements were performed using essentially the same methods as previously described (Srikanth et al., 2010b). Confocal laser scanning microscopy was performed using Fluoview FV10i Confocal Microscope (Olympus), images were captured with a 60 x oil objective. Images were processed for enhancement of brightness or contrast using Fluoview software.

For immunoprecipitation, 6 x His-tagged ORAI1 and 6xHis-tagged STIM1 or Vav1-GFP together with empty vector or FLAG-tagged WT CRACR2A-a, FLAG-tagged CRACR2A^E278D or CRACR2A^{R144G, E300*} was transfected into HEK293T cells. Transfected cells (2 × 10⁷) were lysed in lysis buffer (20 mM Tris-Cl, 2 mM EDTA, 135 mM NaCl, 10% (vol/vol) glycerol, 0.5% Igepal CA-630, protease inhibitor mixture, pH 7.5) and centrifuged at 100,000 x g for 1 hr before preclearing with protein G-Sepharose. Lysates were immunoprecipitated with anti-FLAG antibody-conjugated resin for 6 hr. Immunoprecipitates were washed five times in lysis buffer and analyzed by immunoblotting. For detection of CRACR2A-a, 5 × 10⁶ HEK293 or CRACR2A-deficient Jurkat T cells stably expressing empty vector, FLAG-tagged WT CRACR2A-a, CRACR2A^E278D, or CRACR2A^{R144G, E300*} were lysed in in RIPA buffer (10 mM Tris-Cl pH 8.0, 1% Triton X-100, 0.1% SDS, 140 mM NaCl, 1 mM EDTA, 0.1% sodium deoxycholate and protease inhibitor cocktail [Roche]) and centrifuged to remove debris. For immunoblot analyses, lysates were separated on 10% SDS-PAGE and proteins were transferred to nitrocellulose membranes and subsequently analyzed by immunoblotting with relevant antibodies. Chemiluminescence images were acquired using an Image reader LAS-3000 LCD camera (FujiFilm).

Peripheral blood mononuclear cells (PBMCs) were obtained under federal and state regulations from the CFAR Virology core Laboratory at UCLA that were prepared from buffy coats from healthy, unidentified adult donors using Ficoll-PAQUE gradients. PBMCs from healthy control and patient were activated for 48 hr on a plate coated with 10 µg/ml of anti-CD3 antibody (OKT3, Bio X Cell) and cultured in T cell media (DMEM containing 20% fetal bovine serum and 1% Pen-Strep) supplemented with 5 µg/ml of anti-CD28 antibody (Bio X cell), and 20 U/ml IL-2 (Peprotech) for ThN differentiation. The cells were expanded for a further 4 days with IL-2 and on day 6, cells were extensively washed and activated with 20 nM of PMA, 1 µM of ionomycin, and Brefeldin A (3 µg/ml) for 5 hr, surface stained with anti-CD4-FITC, and intracellularly stained with anti-IFN-γ-PE. CD4-FITC-positive cells were gated for analysis. For reconstitution of CRACR2A-a in CRACR2A-a-deficient PBMCs, naive CD4+ T cells were enriched by magnetic sorting from single-cell suspensions using MagniSort naive CD4+ T cell enrichment kit (catalog # 8804-6814-74) according to manufacturer’s instructions (ThermoFisher Scientific). Cells were activated and cultured as described above. Cells were infected with lentiviruses encoding cas9 together with those encoding sgRNAs targeting CRACR2A-a on day 1 and with lentiviruses encoding WT CRACR2A-a, CRACR2A^E278Dor CRACR2A^{R144G, E300*} together with empty vector on day 2. 48 hr after infection, cells were selected with 1 µg/ml of puromycin and 5 µg/ml blasticidin for 16 hr and then expanded for further 2 days with fresh media. On day 7, cells were stimulated with 10 µg/ml of anti-CD3 antibody, 5 µg/ml of anti-CD28 antibody and Brefeldin A (3 µg/ml) for 5 hr, intracellularly stained with anti-IFN-γ-PE (4 S.B3). For cytokine staining, 1 × 10⁶ Jurkat T cells were fixed with 4% p-formaldehyde for 15 mins, permeabilized with 0.5% saponin, blocked and stained with anti-IL-2-PE antibody from Thermofisher Scientific. For flow cytometry, the following human specific antibodies were used: CD4-FITC (OKT4, eBioscience), IFN-γ-PE-Cy7 (45.B3, eBioscience) and IL-2-PE (MQ1-17H12, eBioscience). Data were acquired using FACSCalibur (Becton Dickinson) or BD LSRFortessa cell analyzers and analyzed using FlowJo software (Tree Star). ELISA was performed on human PBMCs harvested from healthy control and patient for detection of IL-2 (ThermoFisher, #88-7025-88) and TNF (ThermoFisher, # 88-7324-88).

Whole EDTA blood was stained with a combination of CD3-V500, CD4-BV421, CCR6-Pe and CXCR3-Alexa-Fluoro 647 (all antibodies from Becton Dickinson, UK) at room temperature protected from light. Erythrocytes were subsequently lysed using BD red cell lysis solution and following a further 10 min incubation the samples were washed by centrifugation (1500 rmp for 6 min) using PBS/1% FBS. Following the final wash, cells were resuspended in 400 µl of PBS + 0.5% formaldehye and cells analysed using a FACSCanto II flow cytometer (BD) using FACSDIVA software. The gating strategy is show in Figure 1—figure supplement 1B.

Total RNA from cells harvested in TRIzol Reagent (Thermofisher Scientific) was isolated using the Direct-zol RNA isolation kit (Zymo Research). RNA quantity and quality were confirmed with a NanoDrop ND-1000 spectrophotometer. cDNA was synthesized using 1–2 µg of total RNA using oligo(dT) primers and Maxima Reverse Transcriptase (Thermofisher Scientific). Real-time PCR was performed using iTaq Universal SYBR Green Supermix (Bio-Rad) and an iCycler IQ5 system (Bio-Rad) using gene-specific primers described in Supplementary file 1. Threshold cycles (CT) for all the candidate genes were normalized to those of 36B4 to obtain ΔCT. The specificity of primers was examined by melt-curve analysis and agarose gel electrophoresis of PCR products.

CRACR2A-deficient Jurkat T cells stably expressing empty vector, FLAG-tagged CRACR2A-a, CRACR2A^E278D, or CRACR2A^{R144G, E300*} were left untreated or stimulated with 10 µg/ml anti-CD3 antibody (OKT3) for indicated times. Human PBMCs isolated from healthy control and patient were stimulated as described above. Cells were fixed with 4% PFA, permeabilized with ice-cold methanol and stained with phospho-SAPK/Jnk mAb (Cell Signaling, #9257). Cells were washed twice with PBS and analyzed with a FACSCalibur flow cytometer (Becton Dickinson) and FlowJo software.

Statistical analysis was performed using the Origin2018b software (OriginLab, Northampton, MA, USA). Data are presented as mean  ±  s.e.m. Statistical significance to compare two quantitative groups was evaluated using two-tailed/unpaired t-test. A critical value for significance of p  <  0.05 was used throughout the study, and statistical thresholds of 0.05, 0.005 as well as 0.0005 are indicated on the figures by stars (see legends for detail).