The thioredoxin-1 system is essential for fueling DNA synthesis during T-cell metabolic reprogramming and proliferation

Muri, Jonathan; Heer, Sebastian; Matsushita, Mai; Pohlmeier, Lea; Tortola, Luigi; Fuhrer, Tobias; Conrad, Marcus; Zamboni, Nicola; Kisielow, Jan; Kopf, Manfred

doi:10.1038/s41467-018-04274-w

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Published: 10 May 2018

The thioredoxin-1 system is essential for fueling DNA synthesis during T-cell metabolic reprogramming and proliferation

Nature Communications volume 9, Article number: 1851 (2018) Cite this article

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Abstract

The thioredoxin-1 (Trx1) system is an important contributor to cellular redox balance and is a sensor of energy and glucose metabolism. Here we show critical c-Myc-dependent activation of the Trx1 system during thymocyte and peripheral T-cell proliferation, but repression during T-cell quiescence. Deletion of thioredoxin reductase-1 (Txnrd1) prevents expansion the CD4⁻CD8⁻ thymocyte population, whereas Txnrd1 deletion in CD4⁺CD8⁺ thymocytes does not affect further maturation and peripheral homeostasis of αβT cells. However, Txnrd1 is critical for expansion of the activated T-cell population during viral and parasite infection. Metabolomics show that TrxR1 is essential for the last step of nucleotide biosynthesis by donating reducing equivalents to ribonucleotide reductase. Impaired availability of 2′-deoxyribonucleotides induces the DNA damage response and cell cycle arrest of Txnrd1-deficient T cells. These results uncover a pivotal function of the Trx1 system in metabolic reprogramming of thymic and peripheral T cells and provide a rationale for targeting Txnrd1 in T-cell leukemia.

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Introduction

Aerobic organisms have developed a complex system of antioxidant proteins that maintains a reducing environment in the cell. The thioredoxin (Trx) system is one of the major cellular disulfide reducing systems comprising NADPH, Trx reductase (TrxR, encoded by Txnrd), and Trx (encoded by Txn). TrxR possesses the unique capacity to utilize electrons from NADPH to recycle oxidized Trx to its reduced form^1,2. The Trx system regulates multiple cellular processes, such as gene expression, antioxidant response, apoptosis, and proliferation. In particular, it acts as electron donor for the antioxidant enzymes peroxiredoxins and methionine sulfoxide reductases^3,4, modulates the activity of transcription factors such as NF-κB, Ref1, and HIF1α^5,6,7, and is involved in the regulation of apoptosis by suppressing ASK1 function⁸. In mammals, the glutathione (GSH)–glutaredoxin (Grx) system has many overlapping activities with the Trx system due to extensive crosstalk⁹. For instance, Trx and Grx redundantly deliver reducing equivalents for the enzymatic activity of ribonucleotide reductase (RNR)¹⁰. However, how antioxidant systems regulate immune cell function and to which extent the Trx and Grx systems can compensate for each other functions in vivo are poorly understood at present.

Mammals express three TrxR isozymes that differ in their intracellular localization and tissue-specific expression pattern: TrxR1 localized in the cytoplasm, TrxR2 localized in mitochondria, and TrxR3 specifically expressed in the testis. Moreover, Trx1 and Trx2 are isoforms localized in the cytoplasm and in mitochondria respectively⁹. This implicates distinct roles at different cellular compartments. Thioredoxin-interacting protein (Txnip) binds to reduced Trx and acts as a negative regulator of Trx functions¹¹. Txnip knockout in mice results in hypoglycemia, hyperinsulinemia, and liver steatosis¹². Moreover, disruption of Txnip in obese mice strikingly improves hyperglycemia and glucose intolerance¹³, demonstrating a crucial role of Txnip in metabolic disorders. Independently of its Trx-binding function, Txnip represses cellular glucose uptake by both reducing Glut1 expression and inducing Glut1 internalization^14,15,16,17. Txnip expression is mainly mediated by the glucose-sensing transcription complexes chREBP–Mlx and MondoA–Mlx, which bind to carbohydrate response element on the Txnip promoter^18,19.

Due to the critical functions of Txnip in regulating glucose metabolism, we hypothesized the Txnip–Trx system might play a role in the metabolic changes occurring upon T-cell stimulation. Notably, in contrast to naive T cells, activated T cells consume large amount of glucose and amino acids, thereby adjusting their metabolism toward increased glycolysis and glutaminolysis^20,21. Previously, the mitochondrial Trx system was described to be dispensable for development, maintenance, and proliferation of lymphocytes²². To establish a potential function the cytosolic Trx system in T-cell-mediated immunity and metabolism, we generated Txnrd1^fl/fl;Cd4-Cre αβ T-cell-specific and tamoxifen (TAM)-inducible (Txnrd1^fl/fl;Cre-ERT2) mice. We found that T-cell activation is linked to a drastic downregulation of Txnip and an increase in Txn1/Txnrd1 expression, which is absolutely required for synthesis of 2′-deoxyribonucleotides during T-cell metabolic reprogramming. These results consequently characterized a previously unknown function of the cytosolic Trx system in T-cell development and responses.

Results

Txnrd1 is essential for thymic iNKT cell development

To investigate the function of the Trx system in T cells, we generated Txnrd1^fl/fl;Cd4-Cre mice by crossing mice with loxP-flanked Txnrd1 alleles to mice expressing Cre recombinase from the Cd4 promoter. In these mice, deletion of Txnrd1 mainly occurs in CD4⁺CD8⁺ double positive (DP) thymocytes, and consequently both CD4⁺ and CD8⁺ αβ T cells and CD1d-resticted, invariant natural killer T (iNKT) cells lack Txnrd1. Cre-mediated deletion in DP and mature thymocytes, and in peripheral CD4⁺ and CD8⁺ T cells from Txnrd1^fl/fl;Cd4-Cre mice was complete at the genomic DNA and mRNA levels (Supplementary Fig. 1a,b). Wild-type (WT) and Txnrd1^fl/fl;Cd4-Cre mice showed comparable frequencies and numbers of thymic populations of CD4⁻CD8⁻ double-negative (DN), CD4⁺CD8⁺ DP and CD4⁺ and CD8⁺ single-positive (SP) T cells (Fig. 1a). Furthermore, Txnrd1 deficiency had no effects on peripheral αβ T cell numbers in spleen, lymph nodes (LNs), and liver (Fig. 1b and Supplementary Fig. 1c). Expectedly, a proportion of peripheral CD4⁺ and CD8⁺ T cell in naive WT mice displayed an activated/memory phenotype (i.e., CD62L^hiCD44^hi and CD62L^loCD44^hi). However, Txnrd1^fl/fl;Cd4-Cre mice had a considerably lower percentage of such cells in the spleen, LNs, and the liver (Fig. 1c and Supplementary Fig. 1d).

We next assessed the ability of Txnrd1-deficient T cells to refill the hematopoietic compartment of lethally irradiated hosts in a competitive situation with WT cells. Therefore, we reconstituted irradiated C57BL/6 mice with an equal ratio of congenically marked donor bone marrow (BM) cells from Txnrd1^fl/fl;Cd4-Cre (CD45.2⁺) and WT (CD45.1⁺) mice. In this setting, Txnrd1-deficient CD4⁺ and CD8⁺ T cells had a clear disadvantage compared to WT cells in peripheral organs but not in the thymus (Fig. 1d and Supplementary Fig. 1e). This suggests that Txnrd1 is required to refill the peripheral hematopoietic compartment but not for thymic selection and maturation. In line with the low number of activated/memory T cells in Txnrd1-deficient naive mice (Fig. 1c and Supplementary Fig. 1d), the defect in contribution of Txnrd1-deficient T cells to this compartment was even more pronounced in a competitive setting (Fig. 1e and Supplementary Fig. 1f). Taken together, these data suggest that Txnrd1 is dispensable for selection of conventional αβ DP T cells in the thymus and their homeostasis in the periphery. Moreover, Txnrd1 is required intrinsically for expansion of T cells in a lymphopenic environment and steady state generation of activated/memory T cells.

In contrast to conventional αβ T cells, we found that Txnrd1-deficient iNKT cells were massively reduced in the thymus, liver, and spleen (Fig. 1f), suggesting a crucial function of Txnrd1 in iNKT cell development. iNKT cells are known to arise from DP T cells and undergo massive thymic expansion thereafter²³. In the absence of Txnrd1, thymic CD1d-PBS57-tetramer⁺PLZF⁺ iNKT cell numbers were unaffected at stage 0 (CD24^hiCD44^loNK1.1⁻) but were reduced at stages 1 (CD24^loCD44^loNK1.1⁻), 2 (CD24^loCD44^hiNK1.1⁻), and 3 (CD24^loCD44^hiNK1.1⁺), indicating a defect in expansion rather than failed thymic selection (Fig. 1g). In contrast to thymic iNKT cells, expansion of T cell precursors occurs mainly at the DN stage, and therefore prior Txnrd1 deletion in Txnrd1^fl/fl;Cd4-Cre mice. Interestingly, by examining expression of the three main components of the Trx1 system including TrxR1, Trx1, and Txnip, the inhibitor of Trx1, we found that both Txn1 and Txnrd1 expression was increased in DN compared to DP and SP T cells. In contrast, Txnip was predominantly expressed in DP and SP compared to DN T cells (Supplementary Fig. 1g). DN precursors undergo two separate phases of proliferation. Pre-β proliferation occurs prior to TCR β-chain rearrangement at the DN2 stage followed by pre-α proliferation before TCRα chain rearrangement at the DN4 stage²⁴. Precursor proliferation ceases at a stage characterized as CD4⁺CD8⁺TCR⁻^/^loFCS^hi (DP blasts) and it comes to a rest at the CD4⁺CD8⁺TCR⁻^/^loFCS^lo stage (DP rest). To better resolve the expression profiles of the Trx system components during thymic development, we FACS-sorted distinct thymocyte populations. We consistently observed that phases of precursor proliferation (i.e., DN2 and DN4) were associated with low levels of Txnip but high levels of both Txn1 and Txnrd1. By contrast, cells in resting phases (i.e., DN1, DN3, DP, and SP) displayed high Txnip, but low Txn1 and Txnrd1 expression (Fig. 1h and Supplementary Fig. 1h). These data indicate that cell proliferation may be licensed by a high Txnrd1–Txn1:Txnip ratio and that TrxR1 may play a role for expansion of DN precursors, which would be missed in Txnrd1^fl/fl;Cd4-Cre mice.

Txnrd1 is essential for expansion of DN thymocytes

To directly assess the effects of Txnrd1 deletion in hematopoietic precursors, we generated Txnrd1^fl/fl;Cre-ERT2 mice that allow Txnrd1 gene deletion at will by TAM injection. We evaluated the function of Txnrd1 in early thymocyte development 2 weeks after TAM treatment. Txnrd1 was completely deleted in T cells from Cre-expressing mice at the genomic DNA and mRNA levels upon TAM administration (Supplementary Fig. 2a,b). These mice had visibly smaller thymi (Fig. 2a) due to a massively reduced cellularity (Fig. 2b), which affected all thymocyte subsets, including DN and DP precursors, mature SP αβ T cells, γδ T cells, as well as iNKTs (Fig. 2c, d). Importantly, we excluded the possibility that reduction of thymocytes occurred due to Cre-mediated toxicity rather than deletion of Txnrd1 in DN cells (Supplementary Fig. 2c). The massive reduction of DP precursors is consistent with the proliferation occurring during CD44⁻CD25⁻ (DN4) to DP phase (pre-α proliferation). In line with this, DN4 thymocytes lacking Txnrd1 showed defective proliferation, measured as incorporation of the thymidine analog 5-Ethynyl-2′-deoxyuridine (EdU) (Fig. 2e). Analysis of known DN precursor subsets (i.e., CD44⁺CD25⁻ [DN1], CD44⁺ CD25⁺ [DN2], CD44⁻CD25⁺ [DN3] and DN4) showed that each was present in reduced numbers in TAM-treated Txnrd1^fl/fl;Cre-ERT2 mice, although DN2–DN3 stage where pre-β proliferation occurs was affected the most, highlighting the significance of Txnrd1 during proliferation (Fig. 2f).

To assess whether Txnrd1 is cell-intrinsically required in T cells during development, we reconstituted irradiated WT mice (CD45.1⁺) with an equal ratio of congenically marked BM cells from WT (CD45.1⁺CD45.2⁺) and TAM-treated Txnrd1^fl/fl;Cre-ERT2 (CD45.2⁺) or control Txnrd1^fl/fl (CD45.2⁺) mice. In line with the previous observation, we found reduced contribution of Txnrd1-deficient cells during all stages of T cell development with the most prominent decrease from the DN3 stage (Supplementary Fig. 2d). As expected, abolished thymocyte development resulted in an almost complete absence of T cells in the periphery (Supplementary Fig. 2e). Taken all together, these results indicate that the Trx1 system has a critical function for proliferation during thymic development, in line with the expression levels of Txn1, Txnrd1, and Txnip in the different thymic populations (Fig. 1h and Supplementary Fig. 1h).

Txnrd1 is critical for T-cell immune responses in vivo

Given the requirement for expansion of T-cell precursors but not the homeostatic maintenance of naive αβ T cells, we next investigated the requirement of the gene during an in vivo immune response. To address this, we infected mice with lymphocytic choriomeningitis virus (LCMV)-WE strain. We found a drastic reduction in activated CD62L⁻CD4⁺ and CD62L⁻CD8⁺ T cells in spleen and blood of Txnrd1^fl/fl;Cd4-Cre compared to control mice (Fig. 3a and Supplementary Fig. 3a). Moreover, virus-specific CD4⁺ and CD8⁺ T cells were undetectable by gp_61–80 and gp_33–41 tetramer staining, suggesting a crucial role of Txnrd1 in T-cell expansion (Fig. 3b and Supplementary Fig. 3b). To assess whether this impaired expansion in Txnrd1-deficient T cells arose from defective in vivo cycling or increased cell death, we next measured cell proliferation during the course of a viral infection by incorporation of EdU. A drastic reduction in EdU incorporation was observed in Txnrd1^fl/fl;Cd4-Cre T cells at all time points analyzed with the largest difference at the peak of expansion (7 days post infection; Fig. 3c and Supplementary Fig. 3c). Moreover, staining of 7-AAD and Annexin-V during infection showed comparable frequencies of dead/dying CD4⁺ and CD8⁺ T cells (Supplementary Fig. 3d,e). In line with the lack of virus-specific T cells, Txnrd1^fl/fl;Cd4-Cre mice failed to clear LCMV unlike control mice (Fig. 3d). Similarly, Txnrd1 was also essential for Th1-mediated protection against the protozoan parasite L. major, as judged by increased footpad swelling (Fig. 3e), elevated parasite load (Fig. 3f), and impaired numbers of activated CD4⁺ and CD8⁺ T cells (Fig. 3g) in foodtpad and draining LNs of infected Txnrd1^fl/fl;Cd4-Cre mice. Taken all together, these data suggest a pivotal role of the Trx1 system in expanding T cells during in vivo immune responses.

Induction of the Trx system upon T-cell activation

To further elucidate mechanisms of defective T-cell expansion, we studied responses of naive T cells to αCD3/CD28 stimulation in vitro. TCR and IL-2R triggering are known to induce c-Myc which is essential for proliferation of T cells (as well as cancer and embryonic stem cells)^25,26 and metabolic reprogramming²¹. Expectedly, c-Myc expression was upregulated upon T-cell activation (Fig. 4a and Supplementary Fig. 4a). Concomitantly, both Txn1 and Txnrd1 but not Grx1 expression were upregulated (Fig. 4b, c), while the prominent expression of Txnip in naive T cells was abrogated within 4 h of T-cell activation (Fig. 4d), suggesting that the Trx system is the critical pathway controlling T-cell activation and expansion. Txnrd1 deficiency did not alter Txnip, Txn1, and c-Myc expression indicating that TrxR1 acts downstream and is not involved in T-cell activation (Fig. 4a–d). To assess a potential regulation of Txnip by c-Myc²⁷, we used JQ1, a transcriptional inhibitor of c-Myc. Indeed, JQ1 treatment of T-cell stimulation cultures dose-dependently inhibited c-Myc protein production and cell surface expression of CD71, a well-established c-Myc target gene (Fig. 4e). Notably, c-Myc blockade prevented Txnip downregulation induced by T-cell activation (Fig. 4f), indicating that TCR-triggering induction of c-Myc is responsible for Txnip repression.

In line with the in vivo proliferation defect, we observed lower numbers of T cells lacking Txnrd1 compared to control T cells upon stimulation (Fig. 4g). This defect was confirmed by CFSE dilution analysis (Fig. 4h and Supplementary Fig. 4b). The application of the precursor cohort method²⁸ on this data allowed calculation of the mean division time (t_division) and the time to perform the first cell division (t_1°division) upon T-cell stimulation. We found that the mean division time in both CD4⁺ and CD8⁺ T cells lacking Txnrd1 was ~30% longer. Moreover, completion of the first cell division was also delayed in the absence of Txnrd1 (Fig. 4i and Supplementary Fig. 4c), suggesting that additional defects in cell cycle entry might play a role. Consistently, stimulated Txnrd1-deficient T cells had a delayed upregulation of Ki-67, an indicator of cell cycle entry (Fig. 4j and Supplementary Fig. 4d). Finally, we also evaluated whether increased cell death in T cells lacking Txnrd1 might contribute to impaired expansion. We found that viability of Txnrd1-deficient T cells was marginally affected (Supplementary Fig. 4e). Yet, the use of classical apoptosis, necroptosis, ferroptosis inhibitors, or anti-oxidants was insufficient to rescue this death (Supplementary Fig. 4f) and proliferation (not shown) indicating that absence of Txnrd1 did not activate a pathway of controlled cell death.

The Trx system is a major player in cellular antioxidant responses². To test whether the defective proliferation in Txnrd1-deficient T cells is a consequence of increased cellular reactive oxygen species (ROS) levels, we stimulated T cells from Txnrd1^fl/fl;Cd4-Cre and Txnrd1^fl/fl mice in the presence of classical antioxidants. However, supplementation failed to restore proliferation of Txnrd1-deficient T cells (Fig. 4k). Moreover, generation of cellular ROS detected by CM-H₂DCFDA staining in stimulated T cells was comparable (Supplementary Fig. 4g).

Since secreted Trx has been reported to induce IL-2R²⁹, we stimulated mixtures of T cells from congenically marked WT (CD45.1⁺) and Txnrd1^fl/fl;Cd4-Cre (CD45.2⁺) mice at different ratios. Presence of Txnrd1-competent T cells did not rescue the defective expansion of Txnrd1-deficient T cells (Supplementary Fig. 4h), indicating a cell-intrinsic requirement of TrxR1. Furthermore, CD25, CD44, and CD69 upregulation was comparable between Txnrd1-deficient and control T cells upon stimulation (Fig. 4l and Supplementary Fig. 4i), excluding impaired activation as a cause for defective expansion.

Overall, these results indicate that T-cell proliferation requires c-Myc-dependent unlocking of Trx1 from Txnip-mediated inhibition and that Txnrd1 deficiency delays cell cycle entry and slows down cell cycling independent of T-cell activation, cell death, and cellular ROS.

Txnrd1 is required for nucleotide biosynthesis in T cells

Growing evidence demonstrates that T-cell activation is coupled to drastic changes in cellular metabolism that regulate the differentiation and function of activated T cells^20,30. To explore whether the defective proliferation of Txnrd1-deficient T cells was the result of impaired metabolic reprogramming, we stimulated naive T cells in vitro and measured oxygen consumption rate (OCR), an indicator of oxidative phosphorylation (OXPHOS), and extracellular acidification rate (ECAR), a measure of aerobic glycolysis. We observed that Txnrd1-deficient T cells displayed substantially higher basal and maximal OCR compared to control cells. Moreover, they also had higher baseline ECAR as well as maximal glycolytic capacity (Fig. 5a), indicating that Txnrd1-deficient T cells were metabolically more active compared to control cells. Phosphorylation of mTOR was comparable between Txnrd1-deficient and control T cells upon stimulation (Supplementary Fig. 5a). However, we observed increased phosphorylation of the metabolic stress sensor 5′AMP-activated protein kinase (AMPK) in the absence of the Trx system (Fig. 5b). We additionally measured decreased ATP-to-AMP ratio in Txnrd1-deficient T cells upon stimulation, indicating that AMPK activation is due to energy crisis (Fig. 5c). AMPK is well known to respond to nutrient availability and to reprogram cellular metabolism toward increased glycolysis, β-oxidation and mitochondrial function³¹. These data are therefore consistent with and explain the higher ECAR and OCR observed in Txnrd1-deficient T cells (Fig. 5a).

To perform a more comprehensive analysis of the metabolism in proliferating Txnrd1-deficient T cells and to understand the cause of the observed metabolic stress, we next utilized high-throughput, non-targeted, flow injection time-of-light metabolomics³². This method allows rapid and deep profiling of metabolites involved in central carbon metabolism and in peripheral pathways, such as urea cycle, amino acid metabolism, and nucleotide biosynthesis. We stimulated WT and Txnrd1-deficient T cells in vitro and extracted polar metabolites for metabolomics analysis. We identified a total of 989 distinct ion species and putatively annotated them to metabolites listed in the human metabolome database (HMDB) version 3.0³³. Out of 989 metabolites, 101 were significantly changed between Txnrd1-deficient and control T cells upon stimulation (|Log2(fold-change)| > 0.5; adjusted p[Benjamini–Hochberg] < 0.01), but only eight in unstimulated conditions, indicating that the absence of the Trx system influences the metabolism of T cells mainly after stimulation (Supplementary Fig. 5b and Supplementary Data 1). To obtain a general overview of the metabolic differences, we performed a pathway enrichment analysis of the metabolites changing in Txnrd1-deficient versus control T cells both before and after stimulation. This revealed significantly increased levels of metabolites involved in purine and pyrimidine metabolisms in Txnrd1-deficient T cells (Fig. 5d). In addition, Txnrd1-deficient T cells also showed significant accumulation of metabolites involved in the aspartate and glutamate metabolism (Fig. 5d and Supplementary Fig. 5c). However, no significant enrichment for metabolic pathways that have been described to regulate immune cell function was observed, such as glycolysis, tricarboxylic acid (TCA) cycle, and fatty acid synthesis and oxidation (Fig. 5d and Supplementary Fig. 5d,e). Among the components of purine and pyrimidine biosynthetic pathways, we observed accumulation of nucleotides containing ribose sugar, suggesting that the final reduction of RNA into DNA building blocks during DNA biosynthesis was defective (Fig. 5e). To confirm this, we performed a targeted metabolomics study measuring the cellular levels of the abundant purine ribonucleotides and 2′-deoxyribonucleotides. As expected, T-cell stimulation in the absence of the Trx system led to accumulation of ribonucleotides but reduced abundance of their 2′-deoxyribonucleotide forms (Fig. 5f). Taken together, these data imply that the Trx system in T cells regulates the reduction of RNA into DNA building blocks at the last step of DNA biosynthesis.

Increased Cdkn1a expression in Txnrd1-deficient T cells

To further investigate the cellular function of TrxR1 in T cells, we performed RNA sequencing (RNA-seq) of in vivo-activated T cells. Since virus-specific T cells failed to expand in the absence of Txnrd1 during LCMV infection (Fig. 3b and Supplementary Fig. 3b), this model would not have allowed us to isolate enough cells for the experiment. To circumvent this issue, we injected SEB superantigen to obtain enough activated T cells for transcriptomics. After 2 days post SEB injection, Vβ8.1/8.2⁺CD8⁺ T cells from Txnrd1^fl/fl;Cd4-Cre and Txnrd1^fl/fl controls were FACS-sorted for analysis. Total CD8⁺ cells from PBS-treated animals served as a control. Similar to αCD3/CD28 T cell stimulation in vitro, superantigen-mediated T cell activation in vivo resulted in downregulation of Txnip and upregulation of Txnrd1 and Txn1 expression (Fig. 6a). Moreover, the frequency of Vβ8.1/8.2⁺ among CD8⁺ T cells was significantly reduced in Txnrd1^fl/fl;Cd4-Cre upon SEB administration but not in the PBS-treated group, indicating defective expansion in the absence of Txnrd1 (Fig. 6b and Supplementary Fig. 6).

Among the top differentially expressed gene sets (|GFOLD| > 1.5; false discovery rate [FDR] < 0.2) between SEB-stimulated Txnrd1-deficient and control T cell groups, the cell cycle inhibitor Cdkn1a (the gene encoding p21; FDR = 0.05) was significantly enriched in Txnrd1-deficient T cells (Fig. 6c). Analysis of the differentially expressed gene sets involved in cell-cycle progression and regulation pathways revealed that Cdkn1a was the only gene significantly enriched in Txnrd1-deficient T cells upon SEB stimulation (Fig. 6d). Conversely, levels of Cdkn1a expression in unstimulated T cells were comparable between Txnrd1-deficient and -sufficient T cells (Fig. 6d). Furthermore, Cdkn1a also appeared as the top single best predictor, when we used a linear regression model on all detected genes in SEB-treated groups as an additional statistical tool (Supplementary Table 1). Consistent with in vivo RNA-seq results, in vitro T cell stimulation with αCD3/CD28 and real-time quantitative PCR (RT-PCR) confirmed higher expression levels of Cdkn1a in the absence of Txnrd1 (Fig. 6e). Since p21 is known to play a critical role in the DNA damage response by inducing cell cycle arrest³⁴, these data are consistent with the slower proliferative capacity and indicate increased DNA damage response in Txnrd1-deficient T cells.

In addition to Cdkn1a, the expression levels of the Nrf2 target genes Nqo1 and Srxn1 were also increased in Txnrd1-deficient T cells, as suggested by RNAseq (Fig. 6c) and confirmed by RT-PCR (Supplementary Fig. 7a,b), indicating oxidative stress-induced Nrf2 activation as the response to maintain redox balance.

Increased replication stress in Txnrd1-deficient T cells

It is well established that low levels of DNA building blocks contribute to replication fork stalling and consequently activation of the DNA damage response. This process involves phosphorylation of the histone family member H₂AX, leading to accumulation of its phosphorylated form, γH₂AX^35,36. Consistently, stimulation of normal T cells in the presence of hydroxyurea (HU) dose-dependently increased H₂AX phosphorylation (Fig. 7a). HU specifically blocks the catalytic activity of RNR, the enzyme that reduces RNA into DNA building blocks in the last step of nucleotide biosynthesis^37,38. Indeed, T cells with higher replication stress accumulated in the S-phase of the cell cycle (i.e., intermediate levels of DNA in the FACS plot) during which DNA building blocks are most required (Fig. 7a).

We hypothesized that defective nucleotide biosynthesis in Txnrd1-deficient T cells (Fig. 5) would cause increased replication stress and phosphorylation of H₂AX, leading to enhanced expression of p21 (as shown in Fig. 6), similar to HU-treatment of WT T cells. To explore this possibility, we first assessed the role of Txnrd1 in the progression of in vitro activated T cells through the different cell cycle stages. Notably, T cells lacking Txnrd1 accumulated in the S-phase upon stimulation and were conversely reduced in the G₀/G₁- and G₂/M-phases (Fig. 7b). Furthermore, Txnrd1-deficient T cells accumulating in the S-phase had higher levels of γH₂AX (Fig. 7c) and increased expression of the DNA damage sensors Ddb2 and Xpc compared to control cells (Supplementary Fig. 8a). Comparable γH₂AX levels in T cells from Txnrd1^fl/fl and Cd4-Cre mice excluded the possibility that Cre-mediated toxicity/DNA damage rather than Txnrd1 deletion was responsible for increased H₂AX phosphorylation (Supplementary Fig. 8b). In line with these in vitro results, in vivo activation using α-CD3 administration similarly led to increased phosphorylation of H₂AX among proliferating cells (Ki-67⁺). As a control, unstimulated T cells did not accumulate γH₂AX irrespectively of the presence or absence of Txnrd1 (Fig. 7d). To address whether the increased γH₂AX level in the absence of the Trx system was due to stalled replication forks, we measured phosphorylation of ATR, the sensor that responds to this type of DNA damage³⁹. We observed higher ATR phosphorylation in Txnrd1-deficient T cells (Supplementary Fig. 8c), in line with the lower amount of 2′-deoxyribonucleotides and the increased phosphorylation of H₂AX. Taken together, these data show increased replication stress in the absence of the Trx system, thereby mimicking the HU-mediated phenotype caused by reduction of DNA building block pools.

Our results are consistent with previous reports indicating the Trx and the GSH systems as electron donors for RNR in bacteria^40,41. Therefore, we next investigated to which extent the two cellular antioxidant systems are involved in nucleotide biosynthesis in T cells. We stimulated Txnrd1-deficient and control T cells in the presence of L-buthionine-sulfoximine (BSO), which blocks GSH synthesis by inhibiting glutamate-cysteine ligase, thereby reducing the cellular GSH levels⁴². We observed that BSO treatment further reduced the expansion capacity of Txnrd1-deficient T cells in a concentration-dependent manner. Conversely, control T cells were resistant to BSO treatment and were able to expand at BSO concentrations that completely blocked proliferation of cells lacking Txnrd1 (Fig. 7e). BSO-treatment further increased the already higher γH₂AX levels in Txnrd1-deficient T cells in a concentration-dependent manner, leading to cell-cycle arrest in the S-phase (Supplementary Fig. 8d). This demonstrated that blocking GSH in addition to the Trx system led to even higher replication stress. Notably, BSO treatment on control T cells did not affect levels of γH₂AX (Supplementary Fig. 8d). Taken all together, these data demonstrate that the Trx system is the major electron donor for RNR during nucleotide biosynthesis in stimulated T cells. In contrast, the GSH system is dispensable and only inefficiently compensates for the lack of the Trx system in T cells.

Discussion

T-cell activation and proliferation are coupled to drastic changes in cellular metabolism to support their increased bioenergetic and biosynthetic demands. Although less efficient than OXPHOS at producing ATP, activated T cells switch to aerobic glycolysis to generate metabolic intermediates that are crucial for cell growth and proliferation^20,30,43,44. In particular, glucose-6-phosphate generated in the first step of glycolysis can be metabolized in the pentose phosphate pathway to generate ribose, thereby promoting nucleotide biosynthesis^20,21. RNR catalyzes the rate-limiting step of nucleotide biosynthesis by reducing ribonucleotides into the corresponding 2′-deoxyribonucleotides¹⁰. Both the Trx and Grx systems are known electron donors for this reaction^40,41. Previous studies showed that proliferation and development of embryonic fibroblasts, cardiomyocytes, and hepatocytes was unaffected in the absence of Txnrd1^45,46,47 and redundantly covered by the Grx system⁴⁸. Through a mass-spectrum-based metabolomics approach, we found impaired pyrimidine and purine biosynthesis after T-cell stimulation in the absence of Txnrd1 (Fig. 5), consistent with the Trx1 system at providing reducing equivalents to RNR. Glutamine and aspartate are crucial nitrogen donors for synthesis of purines and pyrimidines and can potentially accumulate in case RNR-catalyzed reaction is impaired^49,50. Interestingly, metabolomics revealed also the accumulation of metabolites in the glutamate and aspartate metabolism, further indicating defective nucleotide biosynthesis in the absence of the Trx1 system. Seahorse-based measurements showed that Txnrd1-deficient T cells were metabolically more active, in line with a recent publication indicating that mouse embryonic fibroblasts lacking Txnrd1 displayed increased metabolic flux, glycogen storage, lipogenesis, and adipogenesis⁵¹. Moreover, the higher metabolic activity in Txnrd1-deficient T cells is consistent with the observed increased phosphorylation of the metabolic stress sensor AMPK, thereby boosting glycolytic flux, β-oxidation, and mitochondrial function³¹. Increased glutamine levels in the absence of Txnrd1 may also fuel the TCA cycle and potentially contribute to the increased OCR. Consistent with impaired nucleotide biosynthesis, we detected increased phosphorylation of H₂AX in the S-phase of the cell cycle in the absence of Txnrd1 after stimulation (Fig. 7). During normal cell-cycle progression, γH₂AX decorates single-stranded DNA at the replication forks. The higher levels of γH₂AX in the absence of Txnrd1, which was mimicked by direct inhibition of RNR by HU, is consistent with replication fork stalling events due to reduced availability of DNA building blocks^35,36. Moreover, expression of the cyclin-dependent kinase inhibitor p21, known to participate in DNA damage response downstream of γH₂AX³⁴, was upregulated (Fig. 6). Thus, in the absence of Txnrd1, cellular pools of DNA building blocks are depleted, thereby increasing replication stress, which in turn promotes p21-dependent inhibition of cell cycle progression. Taken together, our results demonstrate that the Trx system is the major electron donor for RNR during nucleotide biosynthesis in rapidly proliferating T cells during development and immune responses. Indeed, blockade of Grx system by targeting gamma-glutamylcysteine synthetase with BSO did not affect proliferation of WT T cells, while it abrogated the remaining proliferative capacity observed in activated Txnrd1-deficient T cells in vitro. Notably, this Grx-dependent partial capacity of Txnrd1-deficient T cells to proliferate in vitro was not observed upon LCMV infection in vivo, where expansion of anti-viral T cells was completely abrogated in the absence of Txnrd1. By contrast, Grx1 was poorly expressed in peripheral T cells and not upregulated upon T-cell activation, thereby providing an explanation why the Grx system cannot fuel DNA synthesis required for T-cell expansion.

Txnip plays a central role in the regulation of glucose metabolism and glycolysis by repressing cellular glucose uptake^14,15,16,17. Naive T cells utilizing low glucose and OXPHOS for energy generation²⁰ showed high Txnip expression, while T-cell stimulation resulted in rapid Txnip downregulation, in line with activated T cells requiring glucose uptake during metabolic reprogramming³⁰. We found that the transcription factor c-Myc, which controls the metabolic reprogramming occurring in activated T cells²¹, links T-cell activation with downregulation of Txnip, in line with other reports demonstrating the potential of c-Myc to directly repress Txnip in tumor cells²⁷. Increased glycolysis due to Txnip downregulation may dynamically deplete glucose metabolites that are sensed by chREBP–Mlx or MondoA–Mlx complexes^16,17 preventing Txnip gene expression and thereby generating a positive feedback loop further promoting glucose uptake and augmenting the flux of glycolysis⁵². In addition to facilitate glucose import, diminishment of Txnip releases Trx1 from inhibition. Concomitant with Txnip downregulation, T-cell activation resulted in Txn1 upregulation. Trx1 is essential for the activation of the low energy sensor AMPK during metabolic stress by suppressing its oxidation⁵³. Activated AMPK phosphorylates Txnip resulting in its degradation, which, in turn, increases Glut1 function and glucose import¹⁵. This mechanism may sustain proliferation in activated T cells. Consistently, T cells lacking AMPKα1 show strikingly reduced expansion upon viral and bacterial infection in vivo⁵⁴, similar to Txnrd1-deficient T cells.

The Trx system has also been proposed to inhibit apoptosis by binding and inactivating ASK1^9,55, and to maintain survival of DP thymocytes⁵⁶. However, our data revealed no relevant increase of cell death in Txnrd1-deficient T cells upon activation in vitro and in vivo, and the numbers and viability of DP cells were unaffected in thymi of Txnrd1^fl/fl;Cd4-Cre mice. Furthermore, as the Trx system is one of the main antioxidant pathways in mammalian cells, we originally speculated that increased ROS levels and consequent aberrant cellular functions were the major cause of the impaired proliferation of Txnrd1-deficient T cells. Yet, supplementation of antioxidants was found ineffective and ROS levels were comparable in WT and Txnrd1-deficient T cells, indicating that defective proliferation is independent of cellular ROS. It is feasible that the GSH system acts as the main cellular antioxidant⁹ and protects T cells from ROS-mediated cell death. Indeed, crucial roles of the GSH system in the regulation of the antioxidant response and in promoting survival of activated T cells have been shown recently. GSH buffers ROS in order to promote c-myc-dependent metabolic reprogramming⁵⁷. Moreover, deficiency of Glutathione peroxidase 4 results in fatal accumulation of membrane lipid peroxides and death by ferroptosis upon T-cell activation⁵⁸. Thus, our results point to the essential requirement of the Trx system for donating electrons to RNR during rapid T-cell proliferation, while the GSH system is indispensable as antioxidant and for survival of activated T cells.

Trx and TrxR upregulation has been reported in several types of cancers, including lymphomas and aggressive T-cell acute lymphoblastic leukemia (T-ALL), and it was associated with a poor clinical outcome^59,60. The Trx system has been suggested to promote development and propagation of cancer by various mechanisms including inhibition of apoptosis, promotion of cell growth, and sustainment of angiogenesis^59,60,61. Several chemotherapeutic drugs that target TrxR are in the clinics and are considered to mediate antitumor activity primarily through induction of oxidative stress and apoptosis^62,63. However, abrogation of T-cell expansion in the absence of Txnrd1 was not associated with a significant increase in ROS production and cell death possibly indicating off-target effects of the inhibitors. Indeed, currently, there is no clinical drug that targets TrxR1 specifically. Our data suggest that such inhibitors may be efficacious and sufficient in treatment of lymphoma and leukemia by direct inhibition of DNA synthesis.

In conclusion, we propose a model where c-Myc-dependent Txnip downregulation by T-cell activation is critical for release of active Trx1 that in turn provides electrons for the synthesis of 2′-deoxyribonucleotides independently of the GSH–Grx system.

Methods

Mice

Txnrd1^fl/fl mice⁴⁶ were provided by M. Conrad (Helmholtz Zentrum, Munich, Germany) and were backcrossed for more than eight generations to C57BL/6. To obtain Txnrd1^fl/fl;Cd4-Cre and Txnrd1^fl/fl;Cre-ERT2 mice, Txnrd1^fl/fl mice were crossed with Cd4-Cre⁶⁴ and Cre-ERT2 mice⁶⁵, respectively. B6 Ptprc^a (CD45.1) animals were purchased from The Jackson Laboratory (Bar Harbor, Maine, USA). About 6–12-week-old age- and sex-matched mice (either male or female) were used for the experiments. For deletion of Txnrd1 in Txnrd1^fl/fl;Cre-ERT2 mice, mice were treated with 2 mg TAM (Sigma-Aldrich) intraperitoneally (i.p.) on two consecutive days. For EdU labeling, mice were injected i.p. with 0.5 mg EdU (Life Technologies) and analyzed 4 or 16 h later for thymic expansion or T-cell proliferation during LCMV infection, respectively. Mice were housed in individually ventilated cages under specific pathogen free conditions at ETH Phenomics Center (EPIC) (Zurich, Switzerland). All animal experiments were approved by the local animal ethics committee (Kantonales Veterinärsamt Zürich, licenses 167/2011, ZH270/2014, 113/2012, ZH135/15, and ZH161/15), and performed according to local guidelines (TschV, Zurich) and the Swiss animal protection law (TschG).

Bone marrow chimeras

CD45.1⁺ or CD45.1⁺ CD45.2⁺ recipients were irradiated twice with 4.75 Gy with a 4-h break in a RS 2000 (Rad Source Technologies Inc., Alpharetta, USA). The following day, mice were reconstituted by intravenous (i.v.) injection of bone marrow cells from hind legs of donor mice, treated with antibiotics (0.024% Borgal, MSD Animal Health, in the drinking water) for 6 weeks, and used for experiments 8 or more weeks after reconstitution. Txnrd1^fl/fl;Cre-ERT2 (and Txnrd1^fl/fl as control) donor mice were pre-treated twice with 2 mg TAM i.p. to delete Txnrd1 gene before BM transfer.

LCMV and Leishmania major infection

LCMV WE was originally provided by Rolf Zinkernagel (University of Zurich, Zurich, Switzerland) and propagated in L929 cells with virus aliquots stored at −80 °C. Mice were i.v. infected with 200 or 500 FFU LCMV WE. Blood was diluted 1:5 in MEM medium (Life Technologies) + 2% FCS (Gibco) was stored at −80 °C until LCMV titers were determined in MC57 cells as previously described⁶⁶.

L. major parasite (MHOM/IL/81/FEBNI) were grown in Schneider’s Drosophila medium (Invitrogen) with 20% FBS and 100 U/ml penicillin 6-potassium and 100 μg/ml streptomycin sulfate. Mice were injected with 2 millions promastigote parasites into the left and right hind footpad, and they were analyzed 25 days after infection. Footpad swelling was monitored every 3–4 days. To quantify parasite load in footpads and draining LNs, single-cell suspensions were plated on 96-well plates in duplicates of threefold serial dilutions and were cultured at 30 °C for 7 days.

In vivo stimulation

In total, 100 μg Staphylococcal enterotoxin B (Sigma-Aldrich) was injected i.p. per mouse and blood/spleen were analyzed at indicated days. For DNA damage analysis in vivo, 10 μg of anti-CD3 (145-2C11; home-made) was injected i.v., and splenocytes were isolated 24 h later for staining.

Flow cytometry

The LCMV glycoprotein peptides gp_33–41 (KAVYNFATM), gp_61–80 (GLNGPDIYKGVYQFKSVEFD), and CD1d-PBS57 tetramer were kindly provided by the National Institutes of Health (Bethesda, MD). Fixable viability dye eFluor® 780 (eBioscience) and LIVE/DEAD® fixable yellow dead cell stain (Life Technologies) were stained in pure PBS before antibody staining. For EdU proliferation experiments, cells from EdU-injected mice were labeled before staining with antibodies with the Click-iT Plus EdU Alexa Fluor® 488 Flow Cytometry Assay Kit (Life Technologies) according to manufacture’s instructions. Antibodies for extracellular stains were incubated with cells for 15 min in FACS buffer (PBS + 2% FCS). For c-Myc, γH₂AX, PLZF, pATR, and Ki-67 intracellular stainings, cells were fixed and permeabilized with Foxp3 Staining Buffer Set (eBioscience) according to manufacture’s instructions and subsequently stained for 30 min. The anti-pATR and the anti-c-Myc antibodies were detected with an anti-rabbit secondary antibody labeled with the FITC fluorophore. To analyze cell death, cells were stained with Annexin-V-APC (BD Bioscience) and 7-AAD (eBioscience) in Annexin-V binding buffer. For cell cycle analysis, DNA was stained with Fxcycle violet stain (Thermo Scientific) according to manufacture’s instructions. Total cellular ROS was quantified by adding 1 μM CM-H₂DCFDA (Life Technologies) to cells and measuring fluorescein signal after 45 min recovery in supplemented medium. Cells were acquired on FACSCanto II or LSRFortessa (BD Bioscience), or sorted on FACSAria III (BD Bioscience). Data were analyzed in FlowJo software (Tree Star). A complete list of all antibodies and staining reagents used in this study can be found in Supplementary Table 2. All the gating strategies used for flow cytometry plots are shown in Supplementary Figs 9–13.

Magnetic cell sorting

For analysis of the different iNKT developmental stages, single thymi were MACS-enriched with APC-labeled CD1d-PBS57-tetramer and anti-APC microbeads (MACS, Miltenyi Biotec). For T-cell stimulation experiments in vitro, T cells were enriched from spleen and lymph node (inguinal, axillary, brachial) by positive selection using a MACS system with microbeads conjugated to monoclonal anti-mouse CD4 (L3T4), CD8a (Ly-2), or CD90.2 (Miltenyi Biotec) following the manufacturer’s instructions. Briefly, 100 millions cells/ml were stained with microbeads (1:10 dilution) for 15 min at 4 °C. After washing, 100 millions cells were resuspended in 500 μl of buffer, and the cell suspension was applied to LS columns and positively sorted in the magnetic field of a MACS separator.

In vitro T-cell stimulation

MACS-sorted T cells (10⁵/well) were seeded in IMDM + GlutaMAX, 10 % FCS, 100 U/mL penicillin, 100 μg/mL streptomycin, 50 μM β-mercaptoethanol (all Gibco) into 96 well plates pre-coated with anti-CD3 (4 μg/ml; 145-2C11; home-made) and anti-CD28 (2 μg/ml; 37.51; home-made) for stimulation. For time-course experiments, cells were fixed with 4% Formaldehyde in PBS (Sigma-Aldrich). For CFSE-labeling, 10⁷/ml PBS-washed cells were incubated for 10 min at 37 °C with 5 μM CFSE (5-(and-6)-Carboxyfluorescein Diacetate, Succinimidyl Ester; life technologies) in PBS + 0.1% BSA (bovine serum albumin; Sigma-Aldrich). Analysis of proliferation kinetics of CFSE-labeled cells was performed as described elsewhere (precursor cohort method²⁸). Inhibitors and compounds were added at the beginning of culture. The following substances were used (purchased from Sigma-Aldrich unless indicated otherwise): l-ascorbic acid, l-Buthionine-sulfoximine, Catalase-polyethylene glycol, Ciclopirox olamine, Deferoxamine, diphenyleneiodonium chloride, dl-Dithiothreitol, Ferrostatin-1 (ChemBridge Corporation), Hydroxyurea, N-Acetyl-l-Cysteine, Necrostatin-1 (inactive) (Merck), (±)-α-Tocopherol, Z-Val-Ala-Asp(OMe)-CH2F (zVAD-FMK; Peptitde Institute), (+)-JQ1.

RNA analysis by real-time quantitative PCR

Total RNA was extracted using TRIzol (Life Technologies), followed by reverse transcription using GoScript Reverse Transcriptase (Promega) according to the manufacturer’s instructions. Real-time quantitative PCR (RT-PCR) was performed using Brilliant SYBR Green (Stratagene) on an i-Cycler (Bio-Rad Laboratories) according to manufacturer’s protocol. Expression was normalized to the housekeeping gene Tbp for mRNA expression, or to genomic Txnrd1 for addressing DNA recombination efficiency in Txnrd1-deficient cells. The sequences of all used primers are listed in Supplementary Table 3.

Immunoblotting

Purified splenic T cells were lysed on ice with RIPA buffer (20 mM Tris-HCl, pH 7.5, 150 mM NaCl, 5 mM EDTA, 1 mM Na₃VO₄, 1% Triton X-100, supplemented with protease inhibitor [Sigma-Aldrich] and phosphatase inhibitor [Sigma-Aldrich]). Samples were then spun for 10 min at 4 °C to remove all cell debris. Protein concentrations were determined using the Pierce^TM BCA Protein Assay Kit (Thermo Scientific). Whole cell extracts (30 μg of proteins) were fractionated by SDS-PAGE and transferred to a Polyvinylidene difluoride (PVDF) membrane using a transfer apparatus according to manufacturer’s instructions (Bio-Rad). After blocking with 4% nonfat milk in TBST (50 mM Tris, pH 8.0, 150 mM NaCl, 0.1% Tween 20) for 45 min, the membrane was washed once with TBST and incubated with primary antibodies (1:1000 in TBST with 4% BSA) at 4 °C for 12 h. Membranes were washed three times for 10 min and incubated with a 1:2000 dilution (in TBST with 4% nonfat milk) of horseradish peroxidase-conjugated anti-rabbit antibody for 1 h. Blots were washed with TBST three times and developed with the ECL system (Thermo Scientific) according to manufacturer’s instructions. Antibodies for phospho-AMPKα (Thr172; 40H9; Cell Signalling Technology) and β-Actin (AC-15; Sigma-Aldrich) were used in this study. The uncropped immunoblots are shown in Supplementary Fig. 14.

RNA preparation and sequencing

Mice were injected with 100 μg Staphylococcal enterotoxin B i.p. (Sigma-Aldrich; or PBS as control). Vβ8.1/8.2⁺ CD8⁺ and total CD8⁺ T cells were FACS-sorted from SEB- and PBS-injected mice, respectively. Approximately 300,000 sorted cells were resuspended in TRIzol (life technologies), phase separation was achieved by addition of chloroform (Sigma-Aldrich) and RNA was precipitated from the aqueous layer with isopropanol (Sigma-Aldrich) using glycogen (Roche) as a carrier. The TruSeq RNA Stranded Sample Prep kit (Illumina) was used to construct the sequencing libraries. In brief, total RNA samples (100 ng) were poly A enriched and reverse transcribed into double-stranded cDNA. TruSeq adapters were then ligated to double-stranded cDNA. Fragments containing TruSeq adapters on both ends were selectively enriched with PCR and subsequently sequenced on the Illumina NextSeq 500 in single-end mode, 150 cycles at the Functional Genomics Center Zurich (FGCZ). The fragments were mapped to the ensemble mouse reference genome GRCm38 (Version25.06.2015) using the STAR aligner⁶⁷. For normalization, the read counts were scaled with the use of the trimmed mean of M-values (TMM) method proposed by Robinson and Oshlack⁶⁸. For differential expression analysis, the Bioconductor package edgeR was used. For the generation of the heat maps, genes with a |generalized fold change (GFOLD)| larger than 1.5 and a false-discovery rate of <0.2 were chosen for visualization. For the analysis of genes involved in the cell cycle, the KEGG cell cycle pathway (mmu04110) was used.

Extracellular flux analysis

Extracellular flux assay was performed with a Seahorse XF-96 Extracellular Flux Analyzer (Seahorse Bioscience) according to protocols described elsewhere⁶⁹. Oxygen consumption rates (OCR) and extracellular acidification rates (ECAR) were measured in unbuffered RPMI 1640 with 2 mM l-glutamine containing 25 mM or without glucose. Baseline levels and responses to the indicated compounds were determined. To address mitochondrial stress, 1 μM oligomycin, 1.5 μM FCCP, and 500 nM rotenone + 1 μM antimycin A were provided (all from Seahorse Bioscience). For glycolysis stress analysis, 25 mM glucose (Sigma-Aldrich), 1 μM oligomycin, and 45 mM 2′-deoxy-d-glucose (Sigma-Aldrich) were added.

Metabolomics

Freshly-isolated, MACS-sorted (with CD90.2 microbeads) T cells were either analyzed directly after isolation or at different time points after αCD3/CD28 stimulation. Approximately 500,000 cells were washed twice with PBS and snap frozen in liquid nitrogen. Intracellular metabolites were extracted twice with cold (−20 °C) 40:40:20 acetonitrile:methanol:water. Non-targeted metabolite profiling was performed by flow injection analysis on an Agilent 6550 QTOF instrument in negative mode as previously reported³². Ions were putatively annotated to metabolites based on the measured mass using the HMDM v3.0 database^32,33. For targeted metabolomics, intracellular nucleotides were extracted with the same procedure, dried in a speed-vac and resuspended in 60 μl of ddH₂O. Relative levels of XMP and dXTP were quantified by ion-pairing LC–MS/MS in negative mode with multiple reaction monitoring using the MRM settings listed in Supplementary Table 4. The data were analyzed as previously described⁷⁰.

Statistical analysis

Two-group comparisons were assessed with a Student’s t test (two-tailed, unpaired). One-sample t test was used to compare the mean of a group with specified value. Multi-group comparisons were assessed by one-way ANOVA followed by either Tukey’s or Dunnett’s corrections. For metabolomics data, the p-values were adjusted using the Benjamini–Hochberg multiple testing correction. The data are represented as mean + standard deviation, and the method of statistical evaluation and the significance levels are indicated in each figure legend.

Data availability

Sequence data that support the findings of this study have been deposited in GEO with the primary accession code GSE107090. The authors declare that all the other data supporting the findings of this study are available within the article and its supplementary information files and from the corresponding authors upon reasonable request.

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Acknowledgements

We thank Christian Wolfrum and Miroslav Baláz for help in using the Seahorse instrument; Wilhelm Krek and Andrea Aloia for sharing pAMPK and pATR antibodies; Gianluca Figlia and Georg Bornkamm for discussions about AMPK activation and the thioredoxin system, respectively; Anette Schütz and Malgorzata Kisielow at the ETH Flow Cytometry Core Facility for cell sorting. We acknowledge the use of the Immgen database as an informative tool for our study. We are grateful for research grants from SNF (310030_163443/1) and ETH Zurich (ETH-23-16-2).

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These authors contributed equally: Jonathan Muri, Sebastian Heer.

Authors and Affiliations

Institute of Molecular Health Sciences, ETH Zurich, 8093, Zürich, Switzerland
Jonathan Muri, Sebastian Heer, Mai Matsushita, Lea Pohlmeier, Luigi Tortola, Jan Kisielow & Manfred Kopf
Institute of Molecular Systems Biology, ETH Zurich, 8093, Zurich, Switzerland
Tobias Fuhrer & Nicola Zamboni
Institute of Developmental Genetics, Helmholtz Center, Neuherberg, 85764, Germany
Marcus Conrad

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Jonathan Muri
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Sebastian Heer
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Mai Matsushita
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Lea Pohlmeier
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Luigi Tortola
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Tobias Fuhrer
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Marcus Conrad
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Nicola Zamboni
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Jan Kisielow
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Manfred Kopf
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Contributions

J.M. and S.H. designed, performed, and analyzed the majority of experiments. M.M., L.P. and T.F. contributed to important methods and technologies. M.C. provided the Txnrd1^fl/fl mice. L.T., J.K., N.Z. and M.K. analyzed data and conceptualization. J.M. and M.K. wrote the manuscript.

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Correspondence to Manfred Kopf.

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Muri, J., Heer, S., Matsushita, M. et al. The thioredoxin-1 system is essential for fueling DNA synthesis during T-cell metabolic reprogramming and proliferation. Nat Commun 9, 1851 (2018). https://doi.org/10.1038/s41467-018-04274-w

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Received: 19 December 2017
Accepted: 17 April 2018
Published: 10 May 2018
DOI: https://doi.org/10.1038/s41467-018-04274-w

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