Autoimmunity can result when cells fail to properly dispose of DNA. Mutations in the three-prime repair exonuclease 1 (TREX1) cause a spectrum of human autoimmune diseases resembling systemic lupus erythematosus. The cytosolic dsDNA sensor, cyclic GMP–AMP synthase (cGAS), and the stimulator of IFN genes (STING) are required for pathogenesis, but specific cells in which DNA sensing and subsequent type I IFN (IFN-I) production occur remain elusive. In this study, we demonstrate that TREX1 D18N catalytic deficiency causes dysregulated IFN-I signaling and autoimmunity in mice. Moreover, we show that bone marrow–derived cells drive this process. We identify both innate immune and, surprisingly, activated T cells as sources of pathological IFN-α production. These findings demonstrate that TREX1 enzymatic activity is crucial to prevent inappropriate DNA sensing and IFN-I production in immune cells, including normally low-level IFN-α–producing cells. These results expand our understanding of DNA sensing and innate immunity in T cells and may have relevance to the pathogenesis of human disease caused by TREX1 mutation.

Compartmentalization and disposal of polynucleotides is critical to prevent autoimmunity in metazoans. Nucleic acids represent a conserved pathogen-associated molecular pattern in viruses and bacteria, and cells have evolved pattern-recognition receptors (PRRs) to detect the inappropriate presence of these macromolecules in the intra- and extracellular spaces (1). PRRs act as nucleic acid sensors to stimulate production of type I IFNs (IFN-I), a family of cytokines that exert proinflammatory and antiviral effects. These nucleic acid sensors detect both pathogen and self-derived polynucleotides, so cells have evolved mechanisms to precisely manage and dispose of self-genetic material to prevent the aberrant activation of nucleic acid sensors and the consequent development of autoinflammation. Mutations in TREX1, SAMHD1, RNaseH2, ADAR1, and MDA5, genes encoding nucleic acid–processing or sensing proteins, cause human autoimmune diseases involving inappropriate IFN-I signaling (24). These disorders exhibit clinical overlap with the autoimmune disease systemic lupus erythematosus (SLE), including the development of an IFN-stimulated gene (ISG) signature in peripheral blood, antinuclear autoantibodies, and inflammation of the skin. Inappropriate IFN-I activity is central to pathogenesis in these autoimmune disorders, leading to their classification as type I interferonopathies (5).

Mutations in the TREX1 DNA exonuclease cause a spectrum of human interferonopathies dependent upon the specific mutant allele (6). Frameshift mutations at positions within the TREX1 gene encoding aa 1–242, comprising the N-terminal catalytic core, cause the severe neuroinflammatory disorder Aicardi–Goutières syndrome (AGS), likely resulting from the complete absence of TREX1 protein (7). In contrast, the missense mutation TREX1 D18N results in a catalytically inactive TREX1 protein and causes familial chilblain lupus (8). Frameshift mutations positioned between the catalytic core and the C-terminal region that controls cellular localization to the endoplasmic reticulum cause retinal vasculopathy with cerebral leukodystrophy (9). Additionally, some TREX1 mutations have been associated with the more prevalent SLE, highlighting the importance of proper DNA degradation for the prevention of systemic autoimmunity (8). The various disease phenotypes caused by TREX1 mutation suggest that TREX1 might contribute to the suppression of autoimmunity through multiple mechanisms.

We have previously demonstrated that TREX1 is a potent dsDNA exonuclease (1012), but its precise in vivo substrate(s) have not been delineated. The biochemical activity of TREX1 suggests that dsDNA of genomic origin is a likely substrate, but other substrates, including ssDNA replication intermediates and retroelements, have been proposed (13, 14). Autoimmunity in TREX1 knockout mice requires the cytoplasmic dsDNA sensor cyclic GMP–AMP synthase (cGAS), providing additional evidence that dsDNA is a primary TREX1 substrate (15, 16). Binding of unprocessed dsDNA to cGAS activates the synthase to generate the cyclic dinucleotide cGAMP, which binds to and activates the stimulator of IFN genes (STING) protein. Active STING recruits TBK1, which phosphorylates the transcription factor IRF3, leading to its nuclear translocation and induction of IFN-I genes (17, 18). Identification of the specific cell populations that sense undegraded TREX1 substrates through cGAS–STING and respond by producing IFN-I could facilitate the development of more effective therapies for human type I interferonopathies.

Mouse models of TREX1-mediated interferonopathies recapitulate aspects of the genotype–phenotype relationship observed with TREX1 mutant alleles in human disease. The TREX1 knockout mice develop aggressive disease and rapid mortality, mirroring the severe AGS phenotype in humans (19). In contrast, mice expressing the catalytically inactive TREX1 D18N enzyme develop milder disease mimicking familial chilblain lupus (20). Mice expressing TREX1 C-terminal truncation mutants develop elevated autoantibodies but no additional indications of autoimmunity, mimicking retinal vasculopathy with cerebral leukodystrophy (21).

In this study, we demonstrate that mice expressing the catalytically inactive TREX1 D18N allele develop a lupus-like autoimmune phenotype associated with robust induction of the IFN-I system. This finding directly links TREX1 nucleic acid catabolism to DNA sensing, IFN-I production, and autoimmunity. We show that TREX1 inactivity within bone marrow–derived cells drives the immunological features of this phenotype, including lymphocyte activation, differentiation, and autoantibody production. Importantly, we demonstrate that TREX1 catalytic inactivity induces hematopoietic overexpression of IFN-α. Furthermore, we identify both innate immune cells and T cells as sources of this IFN-α. We show that TREX1 D18N T cells possess a functional STING signaling axis, which is chronically activated, and that they produce IFN-α protein following TCR stimulation. Our work suggests that chronic IFN-α production in TREX1-deficient T cells acts as an inflammatory signal to facilitate expansion of autoreactive clones, leading to autoimmunity.

TREX1 D18N mice were generated as described (20). Immunophenotyping and bone marrow transplant experiments used 6–8-wk-old female TREX1 wild-type (WT) and D18N mice on a pure 129S1/Sv background. IFN-α/β receptor (IFNAR)−/− mice on the C57BL/6J background were purchased from The Jackson Laboratory and crossed with TREX1 D18N animals to generate F1 hybrid TREX1 D18N IFNAR−/− mice. Pure C57BL/6J TREX1 D18N mice develop an autoimmune phenotype indistinguishable from 129S1/Sv TREX1 D18N mice (data not shown). Mixed-sex TREX1 D18N STING−/− mice on the C57BL/6J background were used at 8–12 wk of age. All other experiments used 8–12-wk-old mixed-sex animals on the pure 129S1/Sv background. All experiments were performed in accordance with the guidelines set forth by the Institutional Animal Care and Use Committee at Wake Forest Baptist Medical Center and the University of Virginia.

Total RNA was collected from splenocytes of experimental animals using the RNeasy RNA Isolation Kit (QIAGEN), according to the manufacturer’s protocol. RNA quality was determined using an Agilent 2100 Bioanalyzer, and samples with a RNA integrity number >8 were used. RNA quality and concentration was determined using a Qubit fluorometer, and 4 μg of total DNase-treated RNA was used for cDNA library generation using the TruSeq Stranded Total RNA LT Sample Prep Kit (Illumina). Purified mRNA was fragmented, converted to cDNA, A-tailed and indexing adapters ligated, PCR amplified, purified with Ampure XP beads, and assessed again for quality using the Agilent 2100 Bioanalyzer. Samples were normalized, pooled, and run on the Illumina HiSeq 2500 using SBS v3 reagents. Collected reads were pseudo-aligned to the Ensembl mouse transcriptome (GRCm38.p6) (22) using Kallisto (23). Differential expression of aligned transcripts was measured using the DESeq2 R package (24). Gene ontology analysis was performed using the Database for Annotation, Visualization, and Integrated Discovering (25).

Spleens were harvested from experimental animals and processed into a single-cell suspension by mashing on a wire screen in RPMI 1640 and 10% FBS with a 5-ml syringe plunger. RBCs were lysed using ammonium–chloride–potassium lysing buffer (Lonza), and remaining splenocytes were washed by dilution with RPMI 1640 and 1% FBS, passed through a 70-μm nylon filter, and counted. For each stain, 1 × 106 cells were incubated with the fixable viability dye Zombie Red (BioLegend) diluted 1:200 in PBS for 15 min at room temperature.

For innate immune cell enumeration, cells were first stained with a mixture of biotinylated Abs against CD3 (17A2, 1:50), CD19 (6D5, 1:200), CD49b (DX5, 1:200), and TER-119 (1:200) for 30 min on ice, then washed. Cells were then stained with anti-B220 (RA3-6B2, APC/Fire750, 1:100), CD11b (M1/70, e450, 1:400; eBioscience), CD11c (N418, PE/Cy7, 1:400), Ly-6C (HK1.4, APC, 1:400), Ly-6G (1A8, BV510, 1:400), CD86 (GL1, FITC, 1:100), and streptavidin–PE in FACS buffer (PBS and 2% FBS) for 30 min on ice. For T cell enumeration, cells were stained with anti-CD3 (17A2, AF700, 1:50; eBioscience), CD90.2 (30-H12, PerCP/Cy5.5, 1:200), CD4 (RM4-4, APC/Fire 750, 1:200), CD8 (53-6.7, V500, 1:200; BD Biosciences), CD44 (IM7, BV421, 1:400), CD62L (MEL-14, FITC, 1:400), CD69 (H1.2F3, PE, 1:100), and SCA-1 (D7, PE/Cy7, 1:250). For B cell enumeration, cells were stained with anti-CD19 (6D5, PerCP/Cy5.5, 1:200), B220 (as above), CD138 (281-2, PE, 1:200), GL7 (AF647, 1:200), CD95 (Jo2, BV510, 1:100; BD Biosciences), and CD69 (H1.2F3, FITC, 1:100). Except where otherwise specified, all Abs were acquired from BioLegend. Cells were washed three times with FACS buffer after staining, fixed in a 1% paraformaldehyde (PFA) solution for 30 min on ice, then acquired on an LSRFortessa X-20 (BD Biosciences). Data were analyzed using FlowJo analytical software (FlowJo). Gating strategies can be found in Supplemental Fig. 1. Innate immune gating strategies were based on an established approach (26).

For measurement of protein phosphorylation, cells were processed as described above, then incubated for 1 h at 2 × 106 cells/ml and 37°C with or without 10 μg/ml 5,6-dimethylxanthenone-4-acetic acid (DMXAA) (Invivogen). After harvesting and pelleting, cells were stained with a viability dye. For p-TBK1 interrogation, cells were stained with surface Abs, allowing delineation of T cells and plasmacytoid dendritic cells (pDCs), then fixed and permeabilized using the eBioscience Foxp3 Transcription Factor Staining Buffer Set. Cells were then stained with anti–p-TBK1 (D52C2, PE, 1:200) or an equal concentration of rabbit IgG isotype control (DA1E, PE, 1:50). Cells were washed three times, then acquired. For measurement of p-IRF3, cells were first stained with anti-CD19 (6D5, FITC, 1:200), then fixed with 1% PFA. Cells were washed twice with PBS, resuspended in 100 μl of PBS, then slowly dripped into 900 μl of ice-cold 100% methanol while gently vortexing and stored at −20°C overnight. The next day, cells were pelleted, resuspended in FACS buffer, and stained with anti-CD3 (PE), CD4 (BUV737), CD8 (BUV395), B220 (APC/Fire750), CD11c (PE/Cy7), and CD11b (e450) at the concentrations described above. We verified that these Ags survive fixation with PFA and methanol and that the CD19-FITC signal does not degrade during the fixation process. The cells were then stained with anti–p-IRF3 (D6O1M, 1:100) or concentration-matched anti-rabbit IgG isotype control (DA1E, 1:500), washed three times, stained with goat anti-rabbit polyclonal Ab (Alexa Fluor 647, 1:1000; Abcam), then washed three times and acquired. All phospho-Abs and isotype controls were from Cell Signaling Technology.

Bone marrow transplants were conducted as three independent experiments with three to five mice per group in each experiment. Recipient animals were female 6–8-wk-old WT and D18N mice. In preparation for the transplant, animals were maintained on filtered antibiotic water containing 800 mg/l sulfamethoxazole and 160 mg/l trimethoprim (Sigma-Aldrich) for 3 d before irradiation. On the day of the transplant, bone marrow was harvested from female 6–8-wk-old WT and D18N donor mice by flushing the femur and tibia with RPMI 1640 and 1% FBS. RBCs were lysed using ammonium–chloride–potassium buffer, and remaining cells were resuspended in RPMI 1640 and 10% FBS and counted. T cells were depleted from isolated bone marrow using a CD3ε MicroBead magnetic depletion kit and LS Column (Miltenyi Biotec) to prevent transfer of autoreactive T cell clones. Successful depletion of T cells was confirmed by staining cells with anti-mouse CD3-PE (1:50; BioLegend) and CD90.2-APC (1:200; BioLegend) and verifying the absence of positive cells by flow cytometry. Depleted cells were resuspended in serum-free RPMI 1640 for injection. Recipient animals were administered a single sublethal dose of gamma radiation equaling 9 Gy using a cesium source. A total of 5 × 106 T cell–depleted bone marrow cells of either genotype were injected in a volume of 500 μl through the tail vein into irradiated animals. Animals were bled at 4-wk intervals following the transplant and euthanized 15 wk after the transplant procedure.

Successful bone marrow engraftment was assessed by collecting PBMCs and employing a quantitative PCR (qPCR)–based genotyping strategy. In brief, blood was collected from experimental animals 12 wk after transplant by facial vein puncture. A total of five to six drops of blood were collected into tubes containing 1 ml 4% (w/v) sodium citrate in water, 1 ml of RPMI 1640 and 1% FBS was added, and the tubes mixed by inversion. This solution was underlaid with 1 ml of Histopaque 1077 (Sigma-Aldrich) and the mixture centrifuged at 400 × g for 20 min. PBMCs were aspirated from the resulting interface and washed twice by pelleting and resuspension in RPMI 1640 and 1% FBS. DNA was isolated from purified PBMCs using the DNeasy DNA isolation kit (QIAGEN) according to the manufacturer’s protocol and the quality and concentration of DNA assessed using a NanoDrop spectrophotometer.

The TREX1 genotype of engrafted hematopoietic cells was assessed using qPCR (Fig. 2A). The total concentration of DNA in each sample was quantitated using a TaqMan primer/spanning a single exon of ISG15 (Thermo Fisher Scientific) and a standard curve of genomic DNA of known concentrations, whereas the concentration of TREX1WT/WT DNA was determined using a primer/probe set specifically recognizing the TREX1 WT allele and a similar standard curve of genomic DNA (forward: 5′-CCC ATC TCC TCC CCA GGC-3′, reverse: 5′-GGC CAG TGG CTT CCA GGT C-3′, probe: 5′–FAM–CCC ATG GTC ACA TGC AGA CCC TCA TCT TC–[Black Hole Quencher 1 (BHQ1)]–3′). Each reaction consisted of 5 μl of 2× TaqMan Gene Expression Master Mix (Thermo Fisher Scientific), 0.5 μl of 20× Primer/Probe Mix (18 μM primer, 5 μM probe), and 5 μl of template DNA. Data were collected on a 7500 Real-Time qPCR system (Applied Biosystems) and analyzed using the 7500 analysis software. The ratio of TREX1 WT DNA concentration/total DNA concentration was used as a measure of successful engraftment.

Serum was collected from whole blood incubated at room temperature for 1 h and centrifuged at 1500 × g for 15 min at 4°C. For measurement of anti-dsDNA autoantibodies, serum was diluted 1:2000 with the supplied diluent and anti-dsDNA autoantibody titer measured using a commercial kit (Alpha Diagnostics), according to the manufacturer’s protocol. For measurement of IFN-α, serum or cell culture supernatants were diluted 1:125 (lymphocytic choriomeningitis virus [LCMV] samples) or 1:1 (all others) with supplied diluent and IFN-α concentration measured using a commercial kit (VeriKine High-Sensitivity IFN-α All-Subtype ELISA Kit; PBL Assay Science).

RNA was collected from splenocytes or purified cells as described above. On-column DNase treatment was performed during RNA isolation using a DNase kit (QIAGEN) according to the manufacturer’s supplemental protocols. Quality and concentration of RNA was assessed using a NanoDrop spectrophotometer. A maximum of 1 μg of RNA was converted to cDNA using the ProtoScript II cDNA conversion kit (Applied Biosystems) according to the manufacturer’s supplied protocol. The cDNA product was diluted five times with deionized water and reactions performed as described above. ISG, IFN-β, and RPLP0 TaqMan primer/probe mixes were from Thermo Fisher Scientific. The IFN-α assay consisted of six forward primers, listed 5′→3′ (5′-ATA CTT CCA CAG GAT CAC TGT GTA CCT G-3′, 5′-ATA TTT CCA CAG GAT CAC TGT GTA CCT G-3′, 5′-ATA CTT CCA CAG CAT CAC TGT GTT CCT G-3′, 5′-ATA CTT CCA CAG CAT CAC TGT GTA CCT G-3′, 5′-ATA CTT CCA CAG GAT CAC TGT TTA CCT G-3′, and 5′-ATA CTT CCA CAG GAT CAC TGT GTT CCT G-3′), four reverse primers (5′-GGC TCT CCA GAC TTC TGC TCT GA-3′, 5′-GGT TCT CTG GAC TTC TGC TCT GA-3′, 5′-GGC TCT CCA GAT TTC TGC TCT GA-3′, and 5′-GGT TCT CCA GAC TTC TGC TCT GA-3′), and a 5′-FAM, 3′-BHQ1–labeled hydrolysis probe (5′-[FAM]-AGA AGA AAC ACA GCC CCT GTG CCT GG-[BHQ1]-3′), all from Eurofin Genomics. Forward primers were used at 150 nM, reverse primers at 225 nM, and the probe at 250 nM. Data were collected on a 7500 Real-Time qPCR system (Applied Biosystems) and analyzed using the accompanying 7500 analysis software. Fold expression measurements were made using the ΔΔCT method, with normalization against the house-keeping gene RPLP0.

For pDC IFN-I expression measurements, low cell inputs necessitated concentration of cDNA products before downstream quantitative RT-PCR (qRT-PCR) analysis. RNA was collected from a total of 1 × 105 purified pDCs. Following reverse transcription, a 1:10 volume of 3 M sodium acetate was added, followed by glycogen to a final concentration of 0.25 μg/μl. This solution was diluted with 4 vol of ice-cold 100% ethanol and incubated at −80°C for 1 h. The solution was then centrifuged at 16,000 × g for 30 min and carefully decanted, avoiding the glycogen pellet. The pellet was washed twice by adding 500 μl ice-cold 70% ethanol, centrifuging for 10 min, and decanting. The pellet was then air dried and resuspended in 30 μl of nuclease-free water for downstream analysis. An equal number of splenocytes and T cells were subjected to this protocol in tandem with pDC samples to ensure that precipitation did not introduce any artifacts, which yielded similar results to larger cell inputs (data not shown).

For each separation, splenocyte single-cell suspensions were stained on ice for 10 min in FACS buffer and 2 mM EDTA containing a mixture of biotinylated Abs. For innate immune negative selection, cells were stained with the same biotinylated Ab mixture used for innate immune cell surface staining, described above. For T cell–negative selection, splenocytes were stained with anti-CLEC9a (7H11, 1:50; Miltenyi Biotec), CD19 (6D5, 1:200), CD138 (281-2, 1:200), Ly-6G (1A8, 1:400), TER-119 (1:220), CD11c (N418, 1:200), and CD11b (M1/70, 1:400). For B cell negative selection, cells were stained with anti-CD3 (17A2, 1:50), CD49b (DX5, 1:200), CD115 (AFS98, 1:200), and CLEC9a, Ly-6G, and TER-119, as above. Stained and unstained cells were separated using Anti-Biotin MicroBeads and an LS column, according to the manufacturer’s protocol (Miltenyi Biotec). B cells underwent a secondary round of purification, using a positive selection against CD19 and CD138, as above. Except where otherwise specified, biotinylated Abs were from BioLegend. Purity of cell preparations was assessed after every separation using flow cytometry and similar panels to those described above (Supplemental Fig. 3C–F).

To enrich for T cells before sorting, splenocyte single-cell suspensions were stained on ice for 10 min in FACS buffer and 2 mM EDTA containing anti-CD19 (1:200), TER-119 (1:200), and Ly-6G (1:400) biotinylated Abs, then washed. Stained and unstained cells were separated using Anti-Biotin MicroBeads and an LS column, according to the manufacturer’s protocol (Miltenyi Biotec). For each sorting sample, enriched T cells from two mice were pooled to increase sorting yield. Cells were then stained with a T cell–specific panel of fluorochrome-conjugated Abs similar to that described above. CD3+ CD4+ or CD8+ cells were sorted based on CD44 expression at 4°C on a Beckman Coulter MoFlo Astrios EQ using a 70-μm nozzle, and immediately pelleted and lysed for subsequent RNA isolation and qRT-PCR analysis, as described above. Purity of all sorted populations was >95%.

For sorting of pDCs, pooled splenocyte suspensions were generated from three mice for each sample. Magnetic depletion of lymphocytes and neutrophils was accomplished as described above, with the addition of biotinylated anti-CD3 (1:50). Cells were stained with viability dye and anti-CD90.2, CD4, CD8, B220, Ly-6C, and CD11c. Contaminating T cells were excluded as CD90.2+ CD4+ or CD8+ events, whereas contaminating B cells were excluded as B220+ Ly-6C events. pDCs were sorted as B220+ Ly-6C+ CD11cint cells using a Becton Dickinson FACSAria with a 100-μm nozzle. Purity was routinely >95%.

The IFN-α expression assay was validated using virally challenged mice. Mixed-sex 8–12-wk-old WT mice on the 129 background were challenged with 2 × 104 PFU LCMV, Armstrong strain (LCMV-Arm) by i.p. injection. After 48 h, animals were euthanized and splenocyte single-cell suspensions generated. A total of 5 × 106 cells were set aside for RNA isolation, and the remainder of the cells subjected to innate immune cell purification as described above. RNA was isolated with on-column DNase treatment from whole splenocytes, innate cells, and noninnate cells, and subjected to IFN-α and ISG expression measurements, as described above.

TREX1 WT and D18N T cells were purified by column purification or sorted, as described above. For transwell experiments, source cells were incubated at a concentration of 1 × 106 cells/ml and 37°C for 1 h with or without 10 μg/ml DMXAA. In addition, TREX1 WT reporter cells were incubated under similar conditions in either media or 20 μg/ml anti-mouse IFNAR (MAR1-5A3; Leinco Technologies). Cells were then washed four times by dilution with 1 ml media, pelleting, and decanting. Cells were added to a 96-well transwell apparatus with a 0.4-μm pore size (Corning). A total of 235 μl of 1 × 106 cells/ml source cell solution was added to the bottom chamber of each well, and then the upper chamber gently overlaid. A total of 80 μl of 2 × 106 cells/ml reporter cell solution was added to the upper chamber of each well, and the plate covered. Cells were incubated for 24 h at 37°C, then reporter cells from the upper chamber collected for RNA isolation and qRT-PCR analysis, as described above.

For long-term in vitro measurement of the ISG signature, 2 × 105 purified WT or D18N T cells in 100 μl media were added to a 96-well plate. This was then supplemented with 100 μl media, 40 μg/ml anti-IFNAR, 40 μg/ml mouse IgG1 isotype control (Leinco Technologies), 2 × 106 beads/ml anti-CD3/CD28–coated beads (Miltenyi Biotec), or T cell activation beads and anti-IFNAR/isotype control. For experiments using sorted naive T cells, wells were precoated with 5 μg/ml anti-CD3/CD28 overnight and then washed once with PBS in lieu of activation beads. Cells were incubated for 72–96 h at 37°C, then harvested and RNA collected for qRT-PCR analysis. For in vitro measurement of IFN-α production, 2 × 105 purified WT or D18N T cells in 100 μl media, or 2.5 × 104 pDCs in 50 μl media were added to a 96-well plate. T cells were supplemented with 100 μl of media, 20 μg/ml DMXAA, 2 μM CpG DNA, or 2 × 106 beads/ml anti-CD3/CD28–coated beads, whereas pDCs were supplemented with 50 μl of media, 20 μg/ml DMXAA, or 2 μM CpG DNA. Supernatants were collected after 24 h and IFN-α measured as described above.

TREX1 catalysis–dependent and –independent mechanisms contribute to inflammation and autoimmunity in the TREX1 null mouse (13, 14, 27, 28). We used TREX1 D18N catalytically deficient mice to specifically address the effects of failed DNA degradation in TREX1-mediated disease and to demonstrate a definitive link between DNA sensing, IFN-I signaling, and inappropriate immune activation. We began by assessing differences in splenic gene expression between TREX1 WT and D18N mice using RNA sequencing. Seventeen of the nineteen most significantly and robustly upregulated genes in TREX1 D18N animals (false discovery rate < 0.001 and fold change > 4) were identified as ISGs (Fig. 1A). Consistent with this finding, gene ontology analysis demonstrated significant enrichment in signaling pathways associated with immune responses and response to viral infection. To identify the extent of IFN signaling caused by the TREX1 D18N mutation, we measured the expression of a subset of ISGs in tissues of TREX1 WT and D18N mice using qRT-PCR. ISG induction was readily detected in multiple tissues, indicating systemic IFN-I sensing (Fig. 1B). Thus, TREX1 catalytic inactivity is sufficient to induce DNA sensing and inappropriate ISG induction.

We next used high-dimensional flow cytometry to examine cellular indications of inflammation and IFN-I signaling in TREX1 D18N mice (Supplemental Fig. 1A–C). TREX1 D18N animals exhibited significantly increased numbers of innate immune cells, including macrophages, monocytes, lymphoid dendritic cells, and pDCs (Fig. 1C). Innate immune cells from these mice also expressed higher levels of the costimulatory molecule CD86, indicative of inflammation-associated maturation (Fig. 1D). In addition to changes within innate immune cells, lymphocytes were also dramatically altered by the TREX1 D18N mutation. All T cells from TREX1 D18N mice uniformly expressed high levels of SCA-1 (Fig. 1E). T cells strongly upregulate SCA-1 following IFN-I exposure (29), but this protein can also be found on T memory stem cells (30, 31) and on classical memory cells (32). We confirmed that SCA-1+ T cells did not express other markers of T memory stem cells, and observed that SCA-1 was universally expressed by naive D18N T cells (data not shown). Thus, this observation likely reflects exposure to IFN-I. IFN-I sensing by T cells acts as a potent inflammatory and proactivation signal (33), and IFN-I signaling is required for T cell–mediated autoimmunity in TREX1-null animals (27). TREX1 D18N animals exhibited robust CD4 T cell differentiation, similar to that reported in TREX1 knockout mice. The CD4 compartment skewed toward a more Ag-experienced phenotype in both the spleen (Fig. 1F) and peripheral blood as early as 10 wk and became increasingly pronounced with age (Fig. 1G). We also observed increased numbers of total and activated B cells, increased germinal centers, and increased numbers of Ab-producing plasma cells in the spleens of TREX1 D18N animals, consistent with the autoreactive B cell response of lupus-like disease (Fig. 1H). Together, these results establish a definitive link between loss of TREX1 exonuclease activity, aberrant activation of IFN-I signaling, and development of cellular correlates of lupus-like autoimmunity.

To identify the cellular origins of IFN-I signaling and immune activation in TREX1 D18N mice, we generated bone marrow chimeric animals expressing TREX1 D18N in only hematopoietic cells, only somatic cells, or in both compartments. TREX1 WT and D18N mice were irradiated and injected with T cell depleted bone marrow from TREX1 WT and D18N donors, and a qPCR strategy was used to confirm successful engraftment of the desired hematopoietic genotypes (Fig. 2A). We first assessed immune phenotypes in experimental animals after 15 wk by measuring ISG expression and comparing with unmanipulated TREX1 WT and D18N mice. Expression of TREX1 D18N in either compartment alone was sufficient to induce a modest ISG signature, whereas the presence of TREX1 D18N within both compartments induced a more robust response similar to unmanipulated TREX1 D18N animals (Fig. 2B). This initial finding suggested that catalytic inactivity of TREX1 in both hematopoietic and nonhematopoietic cells contributes to IFN-I signaling. However, further immune cell analyses revealed increased numbers of innate immune cells only within TREX1 D18N bone marrow recipients (Fig. 2C), accompanied by significantly increased expression of CD86 within these populations (Fig. 2D), consistent with autoinflammation observed in unmanipulated TREX1 D18N animals. We also observed significant induction of T cell SCA-1 expression in all mice that received TREX1 D18N bone marrow (Fig. 2E), accompanied by increased CD4 and CD8 T cell activation (Fig. 2F), increased differentiation of circulating CD4 T cells (Fig. 2G), and splenic CD4 T cells (data not shown). As expected, this CD4 T cell response coincided with increased B cell activation and proliferation. TREX1 D18N bone marrow recipients exhibited increased numbers of total and activated B cells, as well as a trend toward increased germinal center and plasma cells (Fig. 2H). Importantly, increased serum concentrations of anti-dsDNA autoantibodies were only observed in animals expressing TREX1 D18N in hematopoietic cells (Fig. 2I). This major diagnostic criterion of SLE in humans has been previously observed by us in unmanipulated TREX1 D18N animals (20). Thus, ISG signaling results from TREX1 D18N expression in either bone marrow or somatic cells, but the cellular correlates of autoimmunity require expression only in hematopoietic cells.

Having identified a molecular signature of IFN-I signaling and a hematopoietic origin of immune activation in the TREX1 D18N mice, we next sought to identify specific cell populations producing IFN-I. IFN-Is are induced by DNA sensing through the cGAS–STING-signaling axis and other PRRs (34, 35) and act through the common IFNAR (36). IFNAR expression was required for development of lupus-like disease in TREX1 D18N mice, supporting the IFN-I and autoimmunity connection. TREX1 D18N IFNAR−/− mice were completely rescued from autoimmune mortality, exhibited no detectible ISG overexpression, and showed no indications of increased lymphocyte activation, differentiation, or antinuclear autoantibody production (Supplemental Fig. 2A–E). Similarly, TREX1 D18N STING−/− mice exhibited no evidence of ISG induction, antinuclear autoantibody production (Supplemental Fig. 2B, 2E), or accelerated mortality (data not shown), and TREX1 D18N cGAS−/− mice similarly show no evidence of autoimmunity (37). Thus, exonuclease deficiency in TREX1 D18N mice causes cGAS–STING-dependent IFN-I production, which is sensed through IFNAR, leading to autoimmunity.

IFN-α has been implicated as a key molecule in the pathophysiology of SLE (38, 39) and has been found in the cerebrospinal fluid and serum of AGS patients (40, 41). As such, we sought to measure IFN-α production in TREX1 D18N animals. We were able to detect IFN-α protein in cells stimulated with TLR or STING agonists using an established flow cytometry protocol (42), but levels in unstimulated TREX1 D18N cells were below the limit of detection, consistent with a low-level IFN-I response (data not shown). As such, we designed a qRT-PCR assay to simultaneously measure expression of all 14 IFN-α genes. To validate this assay, IFN-α expression was measured in the spleens of WT mice infected with LCMV-Arm, which potently induces IFN-α production predominantly within innate immune dendritic cells (4345). As expected, we observed significantly increased levels of IFN-α expression in LCMV-Arm–infected splenocytes relative to uninfected animals. Further, we measured an 11-fold enrichment of IFN-α expression in purified innate immune cells from LCMV-infected splenocytes (Fig. 3A, Supplemental Fig. 3A, 3B), consistent with a predominantly innate immune origin. Thus, this assay represents an appropriate method for measuring IFN-α expression.

We next sought to identify active isoforms and cellular sources of IFN-I in the TREX1 D18N mouse. IFN-α and IFN-β were upregulated 5-fold in the spleens of TREX1 D18N animals, with IFN-α transcripts ∼25-fold more abundant than IFN-β transcripts (Fig. 3B, Supplemental Fig. 4). Where measured, the relative expression levels of IFN-α and IFN-β were well correlated, so we focused on the more abundant IFN-α because of its hypothesized role in the pathogenesis of lupus and lupus-like disease. Elevated levels of IFN-α protein were observed in the serum of TREX1 D18N mice (Fig. 3C), confirming that it is both expressed and secreted. The absolute concentration of IFN-α was several orders of magnitude below that observed in acute LCMV-Arm infection, consistent with a low-level, “smoldering” inflammatory response caused by TREX1 inactivation.

Innate immune cells are major IFN-α–producing cells, with pDCs specialized for its production (46). As such, we hypothesized that exonuclease deficient TREX1 D18N pDCs sensing undegraded cytosolic DNA might be a major source of IFN-α production. To test this, we purified a mixed population of innate immune cells from TREX1 WT and D18N spleens, resulting in an ∼11-fold enrichment of pDCs (Supplemental Fig. 3A, 3B). Surprisingly, IFN-α expression in TREX1 D18N innate immune cells was similar to levels seen in TREX1 D18N whole splenocytes (Fig. 3D), suggesting that noninnate populations also contribute to IFN-α expression. To more specifically assess the contribution of pDCs, we sorted pDCs to a purity of >95% and measured IFN-α expression. Interestingly, we could not detect elevated expression of IFN-α in TREX1 D18N pDCs (Fig. 3D). This population, therefore, likely does not contribute to spontaneous IFN-α expression in the TREX1 D18N spleen.

We next asked if other TREX1 D18N splenic cell populations might exhibit elevated IFN-α expression. Although not traditionally regarded as major IFN-α–producing cells, T and B cells represent 70–80% of splenic leukocytes, and therefore strongly influence whole-spleen gene expression measurements. We purified TREX1 WT and D18N T and B cells to >99 and >97% purity, respectively (Supplemental Fig. 3C–E), and measured IFN-α induction. TREX1 D18N T cells exhibited robust enrichment of IFN-α expression relative to whole splenocytes, whereas expression was significantly reduced in B cells (Fig. 3D). We also noted significantly higher ISG expression in T cells relative to both splenocytes and B cells, consistent with the observed pattern of IFN-I induction (Fig. 3E). To determine if T cell IFN-α expression tracked to a specific cell population, we sorted naive or CD44high effector/memory (E/M) CD4 and CD8 T cells from the spleens of TREX1 WT and D18N animals and measured induction of IFN-α. Relative to TREX1 D18N splenocytes, significant enrichment of IFN-α expression was observed only in naive CD4 and CD8 T cells (Fig. 3F). Interestingly, although IFN-α induction in TREX1 D18N E/M CD8 T cell did not enrich relative to splenocytes, expression was still elevated over WT E/M CD8s. In contrast, we observed no significant induction in TREX1 D18N E/M CD4 T cells relative to their WT counterparts. Thus, the TREX1 D18N T cell IFN-α response appears to be delocalized across various populations and differentiation states but is most strongly enriched within naive cells. These findings demonstrate that catalytic inactivity of TREX1 D18N within both innate immune and T cells activates DNA sensing and IFN-α/β expression.

TREX1-mediated autoimmunity involves inappropriate T cell activation, and IFN-I can act as an inflammatory signal supporting T cell activation. As such, we elected to further explore TREX1 D18N T cell IFN-I production, as a potentially self-supplied, pro-autoimmune signal. We first sought to confirm that TREX1 D18N and WT T cells possess the capacity to respond to the presence of undegraded cytosolic DNA through a cGAS–STING-dependent mechanism. We found that all T cell subsets expressed cGAS, STING, and IRF3/IRF7, which control IFN-β and IFN-α/β transcription, respectively (47) (Fig. 4A, 4B). Relative to whole TREX1 WT splenocytes, WT E/M and D18N naive and E/M T cells were significantly enriched for cGAS expression. Interestingly, we observed slight enrichment of STING expression in both TREX1 WT and D18N E/M CD4, but not CD8 T cells. cGAS–STING expression levels were therefore not well correlated with the observed pattern of TREX1 D18N T cell IFN-α expression (Fig. 3F), suggesting alternative regulatory mechanisms. Similar to whole splenocytes (Fig. 1A), IRF7 expression was strongly expressed within all TREX1 D18N T cell subsets. Thus, TREX1 D18N T cells express all molecular components of the DNA sensing and signaling pathways required for cytosolic DNA detection and IFN-α/β production.

We next examined the functionality and activation state of the STING pathway in TREX1 WT and D18N T cells and pDCs. We attempted to measure basal IRF7 phosphorylation in TREX1 D18N cells by Western blot, but it was below the limit of detection in all animals tested (data not shown). Using flow cytometry, we were able to observe increased phosphorylation of TBK1 (Fig. 4C, 4D) and IRF3 (Fig. 4E, 4F) in both T cells and pDCs following stimulation with the small-molecule STING agonist DMXAA (48), confirming the functionality of the STING pathway within T cells (4951). Furthermore, TREX1 D18N T cells exhibited a heightened response to stimulation, suggesting that STING activation in TREX1 D18N T cells may lead to enhanced cytokine production. We noted that pDCs exhibited a smaller relative change in p-TBK1/IRF3 signal following DMXAA stimulation compared with T cells, but this may have been a consequence of higher background staining. We also noted that TREX1 D18N pDCs exhibited a dampened response to DMXAA stimulation relative to WT, with potential downstream consequences for cytokine production. Importantly, we observed increased basal TBK1/IRF3 phosphorylation in TREX1 D18N T cells (Fig. 4C–F), indicating spontaneous activation of the primary STING signaling transducers. Thus, TREX1 inactivation in T cells results in constitutive activation of the cGAS–STING pathway.

We next asked if this evidence of STING activation coincided with the spontaneous production of IFN-I protein. TREX1 WT or D18N T cells (source cells) were placed in the bottom chamber of a transwell apparatus, separated from an upper chamber by a pore-containing membrane. IFN-I produced by source cells could diffuse across this membrane to induce ISG expression in TREX1 WT T cells (reporter cells) placed in the upper chamber (Fig. 5A). A ∼2-fold upregulation of reporter ISG expression was observed following incubation of reporter cells with TREX1 D18N T cells for 24 h, indicating that a soluble factor produced by these cells can induce modest ISG expression (Fig. 5B). Induction of reporter ISGs was augmented by preincubation of TREX1 WT or D18N T cells with DMXAA, again confirming the functionality of the STING pathway in T cells. Interestingly, DMXAA-stimulated TREX1 D18N T cells produced a 5-fold more potent reporter ISG response than WT T cells (Fig. 5B), indicating more robust IFN-I production, which is consistent with our p-TBK1/IRF3 data (Fig. 4C–F). To confirm that these effects were dependent on IFN-I sensing, we preincubated reporter cells with anti-IFNAR Ab before measuring reporter ISG expression. IFNAR blockade returned reporter ISG expression to near baseline using both unstimulated and DMXAA-stimulated source cells, indicating that T cell–produced IFN-I was the source of ISG induction in both conditions (Fig. 5B). Together, these results demonstrate that small-molecule activation of STING or sensing of unprocessed TREX1 nucleic acid substrates can lead to IFN-I protein synthesis and secretion in T cells.

To further confirm that TREX1 D18N T cells produce IFN-I protein, T cells were cultured for 72 h, and ISG expression was assessed at regular intervals. We reasoned that if TREX1 D18N T cells could not degrade self-derived DNA and were producing IFN-I as a result, then the ISG signature would continue in vitro and would be sensitive to IFNAR blockade. We noted that ISG expression in unstimulated TREX1 D18N T cells fell relative to WT cells over the initial 24 h but then stabilized at a 5-fold relative induction over subsequent days. Surprisingly, this diminished ISG signature was not sensitive to IFNAR blockade (Fig. 5C), suggesting that it is not dependent on IFN-I. This may indicate that resting TREX1 D18N T cells require additional signals to stimulate IFN-I production or ISG expression in vitro and that unstimulated ISG induction measured in the transwell system (Fig. 5B) represented the residual effect of deteriorating IFN-I production. A recent report demonstrated that TCR ligation can amplify STING signaling in T cells (51). As such, we examined ISG expression in T cell cultures activated with anti-CD3/CD28 over the same time frame. Activated TREX1 D18N T cells maintained higher ISG expression relative to unstimulated cells, and this signature was significantly diminished by IFNAR blockade (Fig. 5D). To determine if this evidence of IFN-I production was dependent on a specific T cell subset or the presence of differentiated cells, we sorted naive CD4 and CD8 T cells to >95% purity and repeated the experiment. ISG expression in both populations remained elevated and sensitive to IFNAR blockade (Fig. 5E, 5F), indicating that activation-dependent IFN-I production does not require already differentiated cells and occurs in both CD4 and CD8 T cells. Interestingly, IFNAR blockade in activated TREX1 D18N cultures did not return ISGs to WT levels but lowered expression to levels similar to unstimulated cultures, suggesting that a component of the ISG signature is IFNAR independent in both conditions.

To confirm that IFNAR-dependent ISG expression in activated cultures truly reflected IFN-I protein production, we assayed T cell culture supernatants for IFN-α. Unsurprisingly, IFN-α was below the limit of detection in unstimulated and CpG DNA-stimulated wells. Consistent with our p-TBK1/IRF3 and transwell data, DMXAA-stimulated T cells produced measurable quantities of IFN-α, and TREX1 D18N T cells secreted more protein than WT. Importantly, activated TREX1 D18N T cells, but not WT, produced detectable quantities of IFN-α protein (Fig. 5G). We compared the magnitude of these responses to TREX1 WT and D18N pDCs, as the canonical primary IFN-α–producing cells. pDCs stimulated with DMXAA or CpG DNA produced substantially more IFN-α on a per-cell basis than T cells, as expected (Fig. 5H). Interestingly, WT pDCs produced significantly more IFN-α in response to CpG than to DMXAA, but this response was inverted in D18N pDCs, potentially indicating alteration of STING and TLR signaling pathways due to chronic inflammatory signaling. Importantly, however, we observed no measurable spontaneous IFN-α production in TREX1 D18N pDCs. Thus, activated TREX1 D18N T cells likely represent a major source of IFN-α in the TREX1 D18N mouse.

Failed processing of self-DNA triggers cytosolic DNA sensors, inflammation, and autoimmunity. Mice expressing catalytically inactive TREX1 D18N fail to process cytosolic DNA, resulting in inflammation and spontaneous lupus-like disease (20). In this study, we show that TREX1 inactivity in T cells causes IFN-I signaling and production. TREX1 D18N mice exhibit IFNAR-dependent ISG signaling and expansion of differentiated lymphocytes. The cellular correlates of autoimmunity require TREX1 inactivity only in hematopoietic cells, indicating that TREX1-mediated DNA disposal must occur within immune cells to prevent autoinflammation. Unprocessed DNA in TREX1 D18N T cells is sensed through cGAS–STING, and activated T cells aberrantly produce IFN-I protein. Thus, TREX1 DNA degradation in T cells is critical to prevent cytosolic self-DNA sensing and spontaneous IFN-I production, providing important new insights into DNA processing and innate immunity in T cells.

The TREX1 D18N mice reveal that catalytic inactivity within bone marrow–derived cells is the primary determinant of autoinflammation. TREX1 is expressed in all murine tissues tested, suggesting that TREX1-mediated DNA disposal occurs in many cell populations (20, 52). Indeed, our data indicate that hematopoietic or somatic TREX1 inactivation induces modest ISG expression (Fig. 2B), potentially reflecting IFN-I production by many cell populations following sensing of damaged DNA (13), retroelements (14), or unprocessed erythroblast DNA (53). It is therefore curious that only hematopoietic TREX1 inactivation results in indications of autoimmunity. This may reflect hematopoietic IFN-α production as the key pathogenic isoform driving autoimmunity in TREX1 D18N mice. Supporting this concept, fibroblast-specific TREX1 deletion and global SAMHD1 deletion in mice cause IFN-β–dependent ISG expression, but no evidence of autoimmunity (5456), suggesting that IFN-β is not sufficient to cause interferonopathy. In contrast, IFN-α appears to be the primary pathogenic isoform in the NZB mouse model of lupus-like disease (57), demonstrating its potential as a pro-autoimmune signal. We propose that somatic TREX1 inactivation induces IFN-β, whereas hematopoietic TREX1 inactivation induces IFN-α and -β synthesis to promote autoimmunity. Alternatively, it may be that IFN-I is necessary, but not sufficient, to promote loss of self-tolerance. This is supported by the fact that an adenoviral vector encoding IFN-α was sufficient to induce autoantibodies in NZBxW/F1 mice, but not WT mice (58). TREX1 inactivity and chronic DNA sensing in immune cells may stimulate production of other STING-inducible cytokines, such as TNF-α (59) or IL-6 (60), which could work in tandem with IFN-I to potentiate loss of self-tolerance.

TREX1 D18N immune cells express IFN-α and -β. Surprisingly, T cells exhibit more robust IFN-I expression than innate immune cells, and we observe no evidence of a spontaneous pDC IFN-I response (Figs. 35). This finding challenges conventional thought that pDCs are the predominant source of IFN-I signaling in lupus-like disease (61, 62) and might more accurately reflect the large capacity for IFN-α synthesis by pDCs during viral infection and through TLR-dependent sensing mechanisms (46). Indeed, CpG-stimulated WT pDCs produced substantial quantities of IFN-α (Fig. 5H). In contrast, TREX1 D18N pDCs produced significantly more IFN-α in response to DMXAA than to CpG (Fig. 5H), despite exhibiting weaker phosphorylation of STING signaling intermediates following DMXAA treatment (Fig. 4C–F). Chronic inflammation in the TREX1 D18N mouse may alter the regulation of pDC DNA–sensing pathways, augmenting some but dampening others, which may suggest that pDC-dependent responses against DNA viruses are altered in the context of interferonopathy. Although this finding merits further study, we observed no evidence of spontaneous STING activation or IFN-α production in TREX1 D18N pDCs, suggesting that they likely do not measurably contribute to systemic IFN-I signaling in this model system. Other TREX1 D18N innate immune populations, such as macrophages or conventional dendritic cells, likely do sense self-DNA and produce IFN-α/β (Fig. 3D), but T cells consistently exhibited the most robust signature of IFN-I expression and are highly abundant relative to these cell types, suggesting that they may be the predominant source in vivo.

TREX1-mediated autoimmunity requires cGAS–STING activation (27). Our finding of IFN-α/β induction in TREX1 D18N T cells therefore requires that these cells possess a functioning cGAS–STING axis. We have shown that T cells express all molecular components of the cGAS–STING pathway and that chemical activation of STING induces TBK1/IRF3 phosphorylation (Fig. 4) and IFN-α production (Fig. 5G), supporting the functionality of this innate immune pathway in T cells (4951). Further, we demonstrate that TREX1 D18N T cells are actively sensing DNA through cGAS–STING, as indicated by increased basal TBK1/IRF3 phosphorylation (Fig. 4C–F). Thus, failure to process DNA in TREX1 D18N T cells triggers cGAS to activate fully functional STING, leading to IFN-I production.

Chronic IFN-I signaling in TREX1 D18N mice likely sustains high-level IRF7 expression in T cells, licensing them to produce IFN-α following DNA sensing. TREX1 D18N T cells produce considerably more IFN-I than WT cells upon STING activation (Fig. 5G). This likely results from chronic overexpression of the IFN-α regulator IRF7 (Fig. 4A, 4B). Innate immune cells require cGAS–STING to produce IFN-α in response to cytosolic DNA (35, 60), supporting the idea that STING activation results in IRF7 phosphorylation and activation. During homeostasis, IRF7 is expressed at high levels only in innate immune cells, priming them to produce IFN-α in response to nucleic acid sensing. IRF7 is also an ISG, however, creating a positive feedback loop of IFN-α production (47). Self-DNA sensing in TREX1 D18N T cells may, therefore, perpetually maintain this feedback loop, allowing chronic IFN-α production.

IFN-α production by TREX1-deficient T cells likely contributes to autoimmunity. IFN-I is a cytokine signal 3 in the three-signal hypothesis of T cell activation, promoting survival, expansion, and function of activated cells (33, 6365). Inappropriate IFN-α production by TREX1 D18N T cells following TCR stimulation (Fig. 5G) may therefore have important consequences for autoreactive T cell responses. IFN-I production in response to the STING agonist cGAMP is strongly augmented by TCR ligation in WT T cells (51). Our data indicate a similar relationship between cGAS–STING-dependent sensing of undegraded TREX1 substrates and TCR signaling. We would propose that TCR engagement in vivo stimulates TREX1 D18N T cells to aberrantly produce IFN-α/β, which may act as a self-provided inflammatory signal to facilitate expansion and survival of autoreactive clones. Potentially consistent with this model, TREX1−/− mice lacking T cells never develop autoimmune disease, whereas those lacking B cells do (27), and deletion of RAG2 reduces IFN-β expression in TREX1−/− hearts (14). Curiously, we noted that TREX1 D18N naive T cells exhibited the highest transcriptional signature of IFN-α ex vivo (Fig. 3F), but the ISG signature decayed in unstimulated cells in vitro (Fig. 5C). TCR ligation was sufficient to induce in vitro IFN-I production in naive T cells, however (Fig. 5E, 5F), suggesting that TCR contacts may indeed be critical for driving naive T cell IFN-α expression in vivo. Interestingly, we also noted that in vitro ISG expression in unstimulated TREX1 D18N cultures decayed, but never reached WT levels, and was resistant to IFNAR blockade. Similarly, ISG expression in TCR-stimulated cultures was only partially IFNAR dependent (Fig. 5C–F). This could suggest that, in addition to IFN-I, TREX1 D18N T cells produce and sense IFN-III, which signals through an alternate receptor and can also drive ISG expression (66, 67).

In summary, our findings demonstrate that T cells act as an important source of IFN-α in a model of IFN-I–dependent lupus-like autoimmune disease. T cells exhibited a signature of IFN-α/β overexpression, evidence of spontaneous TBK1/IRF3 phosphorylation, and IFN-α protein synthesis following TCR stimulation in vitro. Although the mass of IFN-α protein produced on a per-cell basis is small, T cells are highly abundant relative to rare innate immune cells such as pDCs, suggesting that they possess the cumulative capacity to meaningfully contribute to systemic IFN-I levels. We propose a model in which chronic cGAS–STING-dependent DNA sensing in TREX1-deficient T cells, in conjunction with contact with self- or foreign Ag, induces inappropriate IFN-α production to potentiate autoimmunity. These findings add to our understanding of DNA sensing and innate immunity in T cells and may have relevance to the pathogenesis of human interferonopathy.

We thank Dr. Jessica Grieves (Takeda Pharmaceuticals) for preparing samples for RNA sequencing and tissue ISG expression and Dr. John Whitesides (Wake Forest University School of Medicine Flow Cytometry Core) for assistance with cell sorting.

This work was supported by the National Institutes of Health (NIH) (R01AI116725, T32AI007401, and T32GM095440), the Alliance for Lupus Research, the Cowgill and Artom Memorial Fellowships, and the Comprehensive Cancer Center of Wake Forest University National Cancer Institute Cancer (Center Support Grant P30CA012197). R.S. and R.V. were also supported by NIH awards (R01DK104963, R01DK105833, and R21DK112105).

The online version of this article contains supplemental material.

Abbreviations used in this article:

AGS

Aicardi–Goutières syndrome

BHQ1

Black Hole Quencher 1

cGAS

cyclic GMP–AMP synthase

DMXAA

5,6-dimethylxanthenone-4-acetic acid

E/M

effector/memory

IFN-I

type I IFN

IFNAR

IFN-α/β receptor

ISG

IFN-stimulated gene

LCMV

lymphocytic choriomeningitis virus

LCMV-Arm

LCMV, Armstrong strain

pDC

plasmacytoid dendritic cell

PFA

paraformaldehyde

PRR

pattern-recognition receptor

qPCR

quantitative PCR

qRT-PCR

quantitative RT-PCR

SLE

systemic lupus erythematosus

STING

stimulator of IFN genes

WT

wild-type.

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The authors have no financial conflicts of interest.

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