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Peripheral regulatory T lymphocytes recirculating to the thymus suppress the development of their precursors Nicolas Thiault1–3,7, Julie Darrigues1–3,7, Véronique Adoue1–3,7, Marine Gros1–4, Bénédicte Binet1–3, Corine Perals1–3, Bertrand Leobon5, Nicolas Fazilleau1–3, Olivier P Joffre1–3, Ellen A Robey6, Joost P M van Meerwijk1–3 & Paola Romagnoli1–3 Most T lymphocytes, including regulatory T cells (Treg cells), differentiate in the thymus. The age-dependent involution of this organ leads to decreasing production of T cells. Here we found that the output of new Treg cells from the thymus decreased substantially more than that of conventional T cells. Peripheral mouse and human Treg cells recirculated back to the thymus, where they constituted a large proportion of the pool of Treg cells and displayed an activated and differentiated phenotype. In the thymus, the recirculating cells exerted their regulatory function by inhibiting interleukin 2 (IL-2)-dependent de novo differentiation of Treg cells. Thus, Treg cell development is controlled by a negative feedback loop in which mature progeny cells return to the thymus and restrain development of precursors of Treg cells. T lymphocytes have a major role in protecting the integrity of the organism. Most T cells differentiate from hematopoietic precursors in the thymus, where their precursors undergo lineage commitment, somatic rearrangements of genes encoding the α- and β-chains of the T cell receptor for antigen (TCR), and rigorous antigen-specificitydependent selection. These mechanisms lead to the generation of a population of T lymphocytes with a very diverse repertoire of TCRs capable of recognizing rapidly evolving infectious agents and largely devoid of autospecific cells. However, some autospecific T cells mature and migrate to the periphery, where they are controlled by various peripheral tolerance mechanisms. A crucial role in the suppression of deleterious effects of such autospecific T cells (and of other actors of adaptive and innate immunity) is served by Foxp3+CD4+ regulatory T cells (Treg cells)1,2. Similar to conventional T cells (Tconv cells), Treg cells are generated in the thymus, and these two populations share a common hemato­ poietic precursor. However, the population of Treg cells, but not that of Tconv cells, is enriched for autospecific cells3,4. The development of precursor cells into Tconv cells and Treg cells in the thymus therefore appears to be governed by distinct criteria. Indeed, whereas the recognition of agonist ligands consisting of peptide and major histocompatibility complex molecules leads to negative selection of precursors of Tconv cells, it favors the development of precursors of Treg cells2. Moreover, the cytokines interleukin 2 (IL-2) and transforming growth factor β (TGF-β) and co-stimulation via CD28 are required for the

development of Treg cells but not that of Tconv cells. These differences would theoretically allow differential modulation of the development of Tconv cells versus that of Treg cells in the thymus. Under the influence of sex hormones, the thymus involutes after puberty. This phenomenon, combined with the migration of fewer precursors of T cells from the bone marrow to the thymus, leads to a reduced output of new T cells in adults and is a major determinant to the reduced diversity of the naive T cell pool in terms of its specificity for different antigens5,6. We set out to investigate how Treg cell production by the thymus changes with age. RESULTS Thymic Treg cell production decreases substantially with age We used mice expressing a transgene encoding green fluorescent protein (GFP) under the control of the recombination-activating gene 2 (Rag2) promoter (Rag-GFP mice)7. Transcription from the Rag2 promoter in precursors of T cells terminates rapidly upon positive selection in the thymus8 and, in Rag-GFP mice, the GFP then decays with a half-life of 56 h (ref. 9). Recent thymic emigrants (RTEs) to the blood and peripheral lymphoid organs, such as the spleen, therefore, still express GFP, whereas cells that left the thymus earlier do not. We used flow cytometry to quantify GFP-expressing RTEs among the total CD4+ T cell population in the spleens of mice of various ages. As expected, in young mice, a very large fraction of splenic CD4+ T cells were GFP+ RTEs, and the proportion of these cells diminished

1Institut

National de la Santé et de la Recherche Médicale U1043, Toulouse, France. 2Centre National de la Recherche Scientifique U5282, Toulouse, France. de Toulouse, Université Paul Sabatier, Centre de Physiopathologie de Toulouse Purpan, Toulouse, France. 4Ecole Normale Supérieure de Lyon, Department of Biology, 69007 Lyon, France. 5Department of Pediatric Cardiology and Cardiovascular surgery, Children Hospital, University Hospital of Toulouse, Toulouse, France. 6Department of Molecular and Cell Biology, Division of Immunology and Pathogenesis, University of California, Berkeley, California, USA. 7These authors contributed equally to this work. Correspondence should be addressed to J.P.M.v.M. ([email protected]) or P.R. ([email protected]). 3Université

Received 18 December 2014; accepted 17 March 2015; published online 4 May 2015; doi:10.1038/ni.3150

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Peripheral Treg cells recirculate to the thymus The diminishing proportion of newly developing GFP+ Treg cells in the thymus of aging Rag-GFP mice was paralleled by an increasing proportion (from 10% to almost 90%) of mature Treg cells that no longer expressed GFP (Fig. 2a and Supplementary Fig. 1a). The clearly discernable but much smaller GFP− population among thymic Tconv cells increased slightly with age (Fig. 2a, Supplementary Fig. 1a). The population of GFP− Treg cells we noted increasing in the thymus with age may have been cells that never left the thymus and were retained for prolonged periods, as has been proposed13,14, and/or they may have emigrated to the periphery and later returned to the thymus9. To distinguish between these two possibilities, we first compared the gene expression of GFP+ and GFP− Treg cells in the thymus and in the spleen of 8-week-old mice. Principal-component analysis 

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by approximately threefold from 4 to 30 weeks of age (Fig. 1a,b). This reduction in thymic output paralleled a similar decrease in the total number of cells in the thymus (Fig. 1c), which therefore probably accounted in large part for the decrease in RTEs in the spleen. Whereas most CD4+ T lymphocytes are conventional cells involved in protection of the organism against endogenous and exogenous aggression, only a small proportion (~10%) has a Foxp3+ Treg cell phenotype. To investigate whether the output of Treg cells from the thymus also decreases with age, we compared the proportion of RTEs among CD4+Foxp3+ Treg cells with that among CD4+ Foxp3− Tconv cells in the spleen. To facilitate the identification of Foxp3+ cells, we crossed Rag-GFP mice with mice expressing the cell-surface marker Thy-1.1 under the control of the promoter of the gene encoding the transcription factor Foxp3 (Foxp3-Thy-1.1) to generate Rag-GFPFoxp3-Thy-1.1 mice10. Foxp3+ cells could thus be identified by the expression of Thy-1.1 on their cell surface (Supplementary Fig. 1a,b). As expected from our findings reported above, in spleens from Rag-GFPFoxp3-Thy-1.1 mice, the proportion of RTEs among Tconv cells decreased approximately threefold with age (i.e., from 4 weeks to 47 weeks) (Fig. 1d). However, the proportion of RTEs in the Treg cell population decreased substantially more (approximately eightfold) (Fig. 1d). Our data suggested that the production of Treg cells by the thymus decreased considerably more than the production of Tconv cells in adult mice. We confirmed that finding by our observation that with increasing age, Treg cells comprised a diminishing proportion of developing GFP+ cells in the thymus, whereas the relative proportion of newly developing Tconv cells in the thymus decreased only slightly (Fig. 1e). Despite the decreasing production of Treg cells by the thymus, the proportion of Treg cells among CD4+ splenocytes increased with age (Supplementary Fig. 2), which confirmed the observation that peripheral mechanisms also control the number of T cells11,12.

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Figure 1  Thymic production of Treg cells decreases substantially with age. (a) Profiles of GFP expression by electronically gated CD4 +CD8− splenocytes from 5-, 8- and 47-week-old Rag-GFPFoxp3-Thy-1.1 mice (ages above plots), analyzed by flow cytometry (as in Supplementary Fig. 1). Numbers above bracketed lines (gate limits) indicate percent GFP+ cells. (b–e) Quantitative analysis of GFP+ RTEs among CD4+CD8− splenocytes (b), total thymocytes (c), GFP+ cells among Thy-1.1+ Treg cells and Thy-1.1− Tconv cells in spleen (d) and Tconv cells and Treg cells among developing GFP+ thymocytes (e) as a function of age of mice as in a; results in d,e are presented relative to maximum mean values. Each symbol (b–e) represents an individual mouse (data points indicate single measurements); lines (d,e) indicate ‘one-phase decay’ regression curves. *P < 0.001 (extra sum-of-square F test). Data are from 12 experiments with 56 mice from 20 litters.

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of data obtained by high-throughput sequencing technologies for cDNA (RNA-seq) showed that most of the variance in gene expression among the four populations (principal component 1) was for genes with similar low expression in the GFP+ Treg cell populations in the thymus and spleen, higher expression by GFP− Treg cells in the spleen and even higher expression by GFP− Treg cells in the thymus (Fig. 2b). These data indicated a progression in differentiation from GFP+ Treg cells in the thymus and spleen to GFP− Treg cells in the spleen and then to GFP− Treg cells in the thymus. This finding was most consistent with the postulate that the GFP− thymocytes were cells that had passed through the periphery and migrated back to the thymus. Principal component 2 correlated with the tissue from which the cells were derived: low gene expression in the thymus and high gene expression in the spleen (Fig. 2b). The TCR repertoire of Treg cells is shaped not only in the thymus but also by antigens they encounter in the periphery15. In the C57BL/6 (B6) strain of mice we used in this study, for example, T cells expressing the TCR variable-region segment Vβ5 are partially deleted in the periphery but not in the thymus16,17. If the thymic GFP− Treg cells had passed through the periphery, therefore, their TCR repertoire should have reflected such peripheral modulation. To investigate this possibility, we analyzed the expression of several TCR Vβ segments by Treg cells in the periphery and in the thymus from RagGFP mice by flow cytometry. Compared with the abundance of such cells among newly developing GFP+ Treg cells in the thymus, we discerned deletion of Vβ5-expressing cells from the peripheral Treg cell pool among GFP+ RTEs, and this reached its maximum in the GFP− population in the spleen (Fig. 2c and Supplementary Fig. 3a). The deletion of Vβ5-expressing Tconv cells was greater than that of Treg cells (Supplementary Fig. 3b), consistent with published data showing that Treg cells are relatively resistant to deletion in the periphery18. The peripheral deletion of Vβ5-expressing cells was entirely reflected in the thymic GFP− Treg cell population (Fig. 2c and Supplementary Fig. 3a). aDVANCE ONLINE PUBLICATION  nature immunology

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Figure 2  Peripheral Treg cells are able to recirculate to the thymus. (a) Frequency 0.8 0.8 of GFP− cells among CD4+CD8− Thy-1.1+ Treg cells and Thy-1.1− Tconv cells in the 0.6 0.6 thymus of Rag-GFPFoxp3-Thy-1.1 mice, as a function of age, assessed by flow 20 0.4 0.4 cytometry (as in Supplementary Fig. 1). Each symbol represents an individual 0.2 0.2 measurement. (b) Principal-component analysis of RNA-seq expression data 0 0 0 from GFP+ and GFP− Treg cell populations from the thymus (Thy) and spleen (Spl) of 8-week-old Rag-GFPFoxp3-Thy-1.1 mice. PC1 and PC2, principal components 1 and 2. Each symbol represents an individual biological sample (cells pooled from 12 mice). (c) Frequency of cells expressing the TCR segments V β5, Vβ6 or Vβ11 among GFP+ and GFP− CD4+CD8− Treg cell populations in the thymus and spleen. (d) Frequency of total Treg cells among CD4+CD8−TCRhi T cells (left) and of GFP− cells among Treg cells (right) in thymuses from GK-transgenic Rag-GFPFoxp3-Thy-1.1 mice (GK) and non–GK-transgenic Rag-GFPFoxp3-Thy-1.1 mice (WT). Each symbol represents an individual measurement; small horizontal lines indicate the mean (± s.d.). (e) Frequency of injected Treg cells present in the spleen and thymus 1 d after intravenous injection into wild-type mice, relative to total Treg cells in those organs. Each symbol represents an individual data point; small horizontal lines indicate the mean. (f) Ratio of pertussis toxin–treated Treg cells to mock-treated Treg cells (PTX/Mock) in the spleen, bone marrow (BM) and thymus of syngenic recipient mice given injection of Treg cells mock-pretreated or pretreated with pertussis toxin. (g) Ratio of Treg cells in AMD3100-treated mice to Treg cells in PBS-treated (control) mice (AMD3100/PBS) in the spleen, bone marrow and thymus of recipient mice given injection of Treg cells. NS, not significant (P > 0.05); *P < 0.05, **P < 0.01 and ***P < 0.001 (Wilcoxon matched-pairs signed-rank test (c,e), Mann-Whitney test (d) or t-test (f,g)). Data are from 12 experiments with 56 mice from 20 litters (a) or four populations from three biological replicates (cells pooled from 12 mice) (b) or are from three independent experiments (c,d,f,g; mean and s.d. of n = 9 mice in c,d or 3 mice in f,g) or eight independent experiments (e).

Our analysis of the TCR Vβ repertoire also revealed peripheral polyclonal expansion of the Vβ6+ Treg cell population and the Vβ11+ Treg cell population (Fig. 2c and Supplementary Fig. 3a,c). Again, this modulation of the TCR repertoire of peripheral Treg cells was reflected in the thymic GFP− Treg cell population (Fig. 2c). We did not observe any modulation of the expression of the Vβ3, Vβ4, and Vβ14 segments of the TCR (Supplementary Fig. 3d). Together these data supported our hypothesis that the population of GFP− Treg cells in the thymus of Rag-GFP mice consisted largely of cells that had passed through the periphery and returned to the thymus. To further investigate our hypothesis, we analyzed B6 mice with transgenic expression of an antibody to the coreceptor CD4 that results in depletion of the CD4+ T cell pool in the periphery but not in the thymus19 (GK-transgenic mice) (Fig. 2d and Supplementary Fig. 4a–c). We found approximately threefold fewer GFP− cells among Treg cells in thymuses from Rag-GFPFoxp3-Thy-1.1 mice expressing than in those of mice not expressing the GK transgene (Fig. 2d and Supplementary Fig. 4d). Cell-cycle analysis showed that the proliferative activity of GFP+ and GFP− thymic Treg cells was similar in GK-transgenic and non–GK-transgenic Rag-GFPFoxp3-Thy-1.1 mice (approximately 2%; Supplementary Fig. 4e). These results substantiated our conclusion that the GFP− Treg cell population contained a very large proportion of cells that had recirculated from the periphery back to the thymus. To assess directly whether peripheral Treg cells are able to migrate to the thymus, we isolated Treg cells from Foxp3-Thy-1.1 mice, activated them in vitro with antibody to the invariant signaling protein CD3 (anti-CD3) and anti-CD28 and injected them intravenously into wild-type host mice. One day later, a small number of these donor cells was readily detectable in the thymuses of the host mice (Fig. 2e and Supplementary Fig. 5a). Adoptive-transfer experiments with in vitro–activated Tconv cells showed that these cells were also able nature immunology  aDVANCE ONLINE PUBLICATION

to reenter the thymus (Supplementary Fig. 5b), consistent with published reports20. Treg cells generated in vitro by the activation of peripheral Tconv cells in presence of TGF-β also recirculated upon injection into hosts, to the thymus (Supplementary Fig. 5c). To investigate the molecular mechanisms involved in the migration of peripheral Treg cells to the thymus, we treated Treg cells with pertussis toxin, a potent inhibitor of chemokine-receptor signaling, before injecting them into host mice. The next day, we found these Treg cells in the spleen, consistent with the fact that T cells enter this organ via sinusoids and not by chemokine-mediated extravasation through endothelium; however, we found very few of the injected cells in the thymus (Fig. 2f). One of the chemokine receptors expressed by GFP− Treg cells in the thymus was CXCR4 (Supplementary Fig. 5d), which is involved in homing of Treg cells to the bone marrow and of precursors of T cells to the thymus21,22. This finding suggested that CXCR4 might be involved in homing of peripheral Treg cells to the thymus. Injection of Treg cells into hosts treated with the CXCR4 inhibitor AMD3100 did not prevent the accumulation of Treg cells in the spleen, but homing to the bone marrow was somewhat reduced, and homing to the thymus was reduced by approximately half (Fig. 2g). These data unambiguously demonstrated that peripheral Treg cells were able to migrate back to the thymus and indicated that chemokine receptors, in particular CXCR4, were involved in this. Recirculating Treg cells have an activated phenotype To gain insight into the potential functional consequences of Treg cell recirculation to the thymus, we first assessed if recirculating Treg cells in the thymus were activated. Flow cytometry demonstrated that GFP− Treg cells had higher expression of the Treg cell–activation markers CD44, CD103, TIGIT and Nrp1 than did GFP + thymic Treg cells (Fig. 3a). GFP− Treg cells also had lower expression of the 

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Figure 3  Recirculating Treg cells in the thymus have an activated and differentiated phenotype. (a) Flow cytometry analyzing markers (horizontal axes) on GFP + and GFP− CD4+CD8− Treg cells in the thymus of Rag-GFPFoxp3-Thy-1.1 mice. Numbers above bracketed lines (gate limits) indicate percent CD103 + cells or TIGIT+ cells; numbers in other plots indicate mean fluorescence intensity of the marker. (b) Expression scores (as determined by RNA-seq) of the 394 genes upregulated (P < 0.05 (t-test)) in TIGIT+ Treg cells relative to their expression in TIGIT− Treg cells23, in various Treg populations (horizontal axis). Boxes indicate first and third quartiles; small horizontal lines indicate the median (with 10th and 90th percentiles); each symbol represents an individual data point. (c) Heat maps of the normalized expression-scores (row mean value of 0 and variance of 1) of genes encoding markers of Treg cell effector functions (top), helper T cell polarization (middle) and Treg cells in fat and muscle (bottom) in GFP+ and GFP− Treg populations from the thymus and spleen, relative to mean row values (bibliographic references, Supplementary Table 1). *P < 0.001 (t-test). Data are from one experiment representative of at least three independent experiments (a) or one experiment with three biological replicates (b,c).

TCR and of CD62L (Fig. 3a), another indication of their activated state. We compared our RNA-seq data with the expression of 394 genes reported to be upregulated in activated and/or differentiated Treg cells that express the marker TIGIT23. Whereas the GFP+ Treg cells in thymus and spleen expressed small amounts of mRNA from these genes, the GFP− Treg cells in the spleen expressed larger amounts, and the GFP− Treg cells in the thymus expressed even larger amounts (Fig. 3b). Notably, recirculating GFP− cells expressed abundant amounts of mRNAs from many genes encoding Treg cell effector molecules, markers of specialized Treg subpopulations (i.e., Treg cells that control polarized cells of the TH1, TH2, TH17 or TFH subset of helper T cells), and markers found in Treg cells from adipose tissue or muscle (Fig. 3c and Supplementary Table 1). The Treg cell pool that passed through the periphery and recirculated to the thymus thus had the characteristics of activated and differentiated cells, which suggested that it might exert a suppressive function in this organ of T lymphopoiesis. Recirculating Treg cells inhibit thymic Treg cell development We next analyzed the localization of recirculating and newly developing Treg cells within the thymus. Confocal microscopy showed that most Foxp3+ Treg cells were located in the medulla, the main site of Treg cell differentiation10,24 (Fig. 4a,b, left). Approximately half of the Foxp3+ Treg cells were GFP−, in reasonable agreement with flow cytometry of cells from 8-week-old mice (Fig. 4b, right). Moreover, the proportions of recirculating GFP− and newly developing GFP+ cells among Treg cells were similar in the cortex and in the medulla (Fig. 4b, right). The activated and differentiated phenotype of recirculating Treg cells in the thymus and the similar intrathymic distribution of recirculating and newly developing Treg cells suggested that recirculating cells might interfere with the de novo development of their precursors. To test this hypothesis, we added mature Treg cells to fetal thymus organ cultures and assessed the effect of this on de novo T cell 

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development. Exogenous mature Treg cells inhibited the development of Treg cells by 34–60% (Fig. 4c,d, left), but Tconv cells did not (Supplementary Fig. 5e). In contrast, exogenous Treg cells did not inhibit the development of Tconv cells (Fig. 4d, middle). Moreover, in GK-transgenic mice, which lack peripheral CD4+ T cells, the proportion of Treg cells among newly developing GFP+ mature thymic CD4+ cells was significantly higher than that in non–GK-transgenic control mice (Fig. 4e, left), which demonstrated that recirculating cells also inhibited the differentiation of Treg cells in vivo. In Rag-GFPFoxp3Thy-1.1 mice of various ages, we observed an inverse correlation between the proportion of recirculating Treg cells among thymocytes and the de novo development of Treg cells (Fig. 4f). Together these data demonstrated that the recirculation of Treg cells to the thymus inhibited the de novo development of Treg cells. Recirculating GFP− thymic Treg cells had high expression of several genes encoding products involved in the regulatory functions of peripheral Treg cells (Fig. 3c), which suggested that one or more of these regulatory function(s) might be involved in inhibiting the de novo development of Treg cells. For example, mature Treg cells can induce T cell apoptosis by adsorbing IL-2 onto their high-affinity IL-2 receptors25, and our data demonstrated that recirculating Treg cells in the thymus expressed the IL-2 receptor α-chain (CD25; encoded by Il2ra) (Fig. 3c), which confers high affinity on the IL-2 receptor complex. Published studies have indicated that in the absence of signaling through the IL-2 receptor, the development of Treg cells in the thymus is reduced by approximately twofold (ref. 26). IL-2 seems to be involved in the final phases of Treg cell development, when CD4+CD25+Foxp3− precursor cells develop into CD4+CD25+Foxp3+ mature Treg cells27. In our fetal thymus organ cultures, exogenous Treg cells inhibited the development of CD4+Foxp3+ Treg cells but not that of CD4+CD25+Foxp3− precursor cells (Fig. 4d, right). We found more aDVANCE ONLINE PUBLICATION  nature immunology

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Figure 4  Mature peripheral Treg cells that GFP+ GFP– reenter the thymus inhibit the differentiation Superposition Foxp3 80 60 of Treg cells. (a) Confocal microscopy 60 40 of medullary sections of the thymus from 40 8-week-old Rag-GFP mice (n = 3), stained 20 20 for CD4-expressing T cells (magenta), 0 0 differentiating T cells expressing GFP (green) Cortex Medulla Cortex Medulla and Foxp3-expressing Treg cells (blue), as well as a merged image (superposition), with Ctrl Ab: + + Treg cells: – + cortical and medullary regions distinguished 5 10 5.5 0 3.3 1.2 on the basis of the GFP-staining intensity of 4 10 the thymocytes (cortex ≥ 83 > medulla). Total 3 area analyzed: medulla, 3.12 mm 2; cortex, 10 3.44 mm2. White arrows indicate a GFP + 2 10 0 Foxp3+ Treg cell; yellow arrows indicate a 2 3 4 5 2 3 4 5 GFP− Foxp3+ Treg cell. (b) Density of GFP + 0 10 10 10 10 0 10 10 10 10 or GFP− Foxp3+CD4+ Treg cells (left) and b CD4 GFP H-2K frequency of GFP + or GFP− cells among WT NS NS 4 4 6 6 20 GK Foxp3+CD4+ Treg cells (right) in the cortex NS * * and medulla of the thymus sections in a. 3 3 15 4 4 (c) Flow cytometry of electronically gated 2 2 10 CD4+CD8−TCRhi cells from NMRI fetal 2 2 thymuses cultured alone (Treg cells −) or with 5 1 1 exogenous, mature H-2K b-positive B6 Treg 0 0 0 0 0 cells (Treg cells +), with the addition of isotypeTreg cells: – + – + – + matched control antibody (Ctrl Ab) (control for Treg cells the antibody in g) throughout the duration mAb to Treg cells – + + CD25: Treg cell precursors of culture. Numbers above outlined areas Treg cells: Treg cells + – + 3 4 indicate percent Foxp3 + H-2Kb-negative cells 5 NS NS + 1.5 1.1 1.5 0 10 20 3 3 (endogenous developing Foxp3 Treg cells) 2 104 15 (left) or percent Foxp3 + H-2Kb-positive cells 2 2 3 10 10 1 (right). (d) Frequency of endogenous Treg 1 2 1 10 + − hi cells among CD4 CD8 TCR cells (left), of 5 0 0 0 endogenous Tconv cells among total thymocytes 0 0 (middle), and of CD25 +Foxp3− precursors of – b GFP Treg cells (%) H-2K Treg cells among CD4 +CD8−TCRhi cells (right) in fetal thymus organ cultures as in c. Each pair of symbols connected by a line is from the same experiment. (e) Frequency of Foxp3 + Treg cells and of CD25 +Foxp3− precursors of Treg cells among differentiating GFP + CD4+CD8−TCRhi thymocytes in GK-transgenic and non–GK-transgenic Rag-GFPFoxp3-Thy-1.1 mice. (f) Frequency of differentiating GFP + Treg cells (left vertical axis) and of CD25 +Foxp3− precursors of Treg cells (right vertical axis) among CD4 +CD8− cells, as a function of the frequency of recirculating GFP − Treg cells among total thymocytes in the thymus of Rag-GFPFoxp3-Thy-1.1 mice 4 –47 weeks of age. Each symbol represents an individual mouse. (g) Flow cytometry of CD4 +CD8−TCRhi cells from fetal thymus organ cultures as in c but with the addition of monoclonal antibody to CD25 (mAb to CD25) throughout the duration of culture (numbers in plots as in c). (h) Frequency of endogenous Foxp3 + Treg cells among CD4 +CD8−TCRhi cells (left) and of CD4 +CD8−TCRhi Foxp3− Tconv cells among total thymocytes (right) in fetal thymus organ culture as in g. Each symbol (e,h) represents an individual data point; small horizontal lines indicate the mean (± s.d.). NS, not significant; *P < 0.01 (Wilcoxon matched-pairs signed-rank test (d,h) or Mann-Whitney test (e)). Data are representative of three experiments (a,c,g) or are from three experiments (b,e,h; mean and s.d.), eight experiments (d) or 12 experiments with 56 mice from 20 litters (Treg cells) or 7 experiments with 31 mice from 11 litters (Treg cell precursors) (f). Foxp3

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newly developed GFP+ mature Treg cells, but not CD4+CD25+Foxp3− precursor cells, in GK-transgenic mice than in non–GK-transgenic mice (Fig. 4e, right). The increasing frequency of recirculating Treg cells with age correlated with the decreasing frequency of newly developing Treg cells but not of CD4+CD25+Foxp3− precursor cells (Fig. 4f). Recirculating Treg cells therefore seemed to inhibit the development of mature Treg cells but not of CD4+CD25+Foxp3− precursor cells. As expected, the addition of anti-CD25 substantially reduced the development of Treg cells in fetal thymus organ cultures (Fig. 4g). Exogenous Treg cells did not further inhibit the remaining IL-2-independent development of Treg cells (Fig. 4g,h). These data indicated that mature recirculating Treg cells inhibited IL-2-dependent but not IL-2-independent development of Treg cells in the thymus.

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Treg cell precursors (%)

e

Treg cell precursors (%)

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d

development of Treg cells. To determine whether peripheral Treg cells also recycle to the thymus in humans, we analyzed the marker CD31 (PECAM-1), which is expressed by de novo developing thymocytes and RTEs and whose expression is downregulated in the periphery upon activation28. In human thymuses, we found a population of CD31− cells among the mature CD4+ thymocytes (Fig. 5a), which probably corresponded to peripheral T cells that had migrated back to the thymus. The proportion of recirculating CD31− cells was substantially greater among CD4+Foxp3+ Treg cells than among CD4+Foxp3− Tconv cells (Fig. 5a). We found Treg cells expressing the co-stimulatory molecule ICOS and with high expression of Foxp3 (which had therefore been activated29) in the recirculating CD31− thymic Treg cell population but not in the newly developing CD31+ thymic Treg cell population (Fig. 5b,c). A subpopulation of CD31− thymic Treg cells, but not CD31+ thymic Treg cells, expressed the transcription factor T-bet (Fig. 5d), indicative of their differentiated phenotype30,31. These findings indicated that in humans, peripheral Treg cells recirculated to the thymus and that these cells had an activated and differentiated phenotype. 

Articles

11.4 Cells

104 3

CD25

10

20.4

8.1

0 85.3 0102 Foxp3

103 104 105

0 CD31

103

**

35 30 25 20 15 10 5 0

b

Tconv cells

CD31

–

70

CD31+

60

26.8

–

**

CD31

*

CD31–

CD31+

50 40 30 20 10 0

104 105

0

c

ICOS+CD31–

d

103 104 105 ICOS CD31–

60

DISCUSSION The data presented here have demonstrated that in the mouse, peripheral Treg cells were able to migrate to the thymus, where they inhibited the de novo development of this key immunoregulatory T cell subset. The proportion of recirculating cells among thymic Treg cells augmented with age, which led to an increasingly reduced output of newly developed Treg cells. In the human thymus, we found recirculating cells with an activated and differentiated phenotype, which suggested that a similar negative feedback loop of Treg cell development may also operate in humans. Parabiosis experiments have shown that thymuses from mice lacking the pre-TCR component pTα, in which T cell development is arrested at a very early stage and the thymus therefore contains very few cells, contain a substantial fraction of T cells derived from the wild-type partner32. Mature Treg cells adoptively transferred into irradiated mice or mice deficient in the RAG recombinase have been found in the thymus of the hosts14. GFP− Foxp3+ Treg cells are present in the Rag-GFP thymus9,14,33,34. Such results suggest that peripheral Treg cells might recirculate to the thymus. We have demonstrated that peripheral Treg cells were able to migrate back to the thymus: GFP− Treg cells in the thymus expressed a TCR repertoire reflective of peripheral deletion and expansion; these cells expressed genes that are also expressed in fat and muscle Treg cells; in mice with depletion of peripheral CD4+ T cells, including Treg cells, the number of GFP− (but not GFP+) Treg cells in the thymus was substantially lower than that in mice without depletion of peripheral CD4+ T cells; and Treg cells adoptively transferred into otherwise unmanipulated wild-type mice were able to home to the thymus. Although we cannot exclude the possibility that some of the GFP − Treg cells were long-time resident cells that had never left the thymus, as has been suggested13,14, our results also indicated that the majority of these cells were peripheral cells that had recirculated back to the thymus. Our finding that recirculating Treg cells expressed many genes encoding products involved in the regulatory functions of peripheral Treg cells suggested that these cells exert a suppressive function 

T-bet+ cells (%)

Cells

Figure 5  Activated and differentiated peripheral Treg cells also CD31+ CD31+ ICOS–CD31– 50 recirculate to the thymus in humans. (a) Gating strategy for CD31+ Tconv cells and Treg cells (left) among human thymocytes (electronically 40 gated CD4+CD8−TCRhi cells), expression of CD31 on Treg cells and 23.0 30 Tconv cells (middle) gated as at left, and frequency of recirculating − 20 (CD31 ) Treg cells and Tconv cells in the thymus (right), all assessed by flow cytometry. Numbers adjacent to outlined areas (far left) 10 indicate percent CD25neg–loFoxp3− (Tconv) cells (bottom left) or 0 hi + CD25 Foxp3 (Treg) cells (top right); numbers above bracketed lines 0 103 104 105 0 102 103 104 105 (middle) indicate percent CD31− (recirculating) Treg cells or T-bet Foxp3 Tconv cells. Each symbol (far right) represents an individual data point; small horizontal lines indicate the mean. (b) Expression of ICOS on electronically gated CD31− and CD31+ thymic Treg cells (left), and frequency of ICOS+ cells among CD31− and CD31+ Treg cells (right). Number above bracketed line (electronic gate) indicates percent ICOS+ cells (left). (c) Expression of Foxp3 on electronically gated ICOS+ and ICOS− CD31− cells and on CD31+ thymic Treg cells. (d) T-bet expression by electronically gated CD31− and CD31+ thymic Treg cells (left), and frequency of T-bet+ cells among CD31− and CD31+ Treg cells (right). Number above bracketed line (electronic gate) indicates percent T-bet+ cells (left). Each symbol (b,d, right) represents an individual data point; small horizontal lines indicate the mean. *P < 0.05 and **P < 0.001 (Wilcoxon matched-pairs signed-rank test). Data are representative of sixteen experiments (a,b,c) or seven experiments (d). Cells

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Treg cells

ICOS+ cells (%)

Tconv cells

Cells

Treg cells 105

CD31– cells (%)

a

in the thymus. Adding Treg cells to fetal thymus organ cultures indeed inhibited the de novo development of Treg cells (but not that of Tconv cells). We also observed increased development of Treg cells in mice in which recirculation to the thymus was experimentally reduced. The naturally occurring increased proportion of recirculating Treg cells in the thymus with age also correlated with a decrease in the de novo development of Treg cells. Together these data showed that recirculating Treg cells inhibited the development of Treg cells in the thymus. However, recirculating Treg cells did not inhibit the development of Foxp3−CD25+ precursors of Treg cells. Further differentiation of these precursors into Treg cells requires IL-2 (ref. 27), which suggests that recirculating Treg cells might act through adsorption of IL-2, which is thought to be available in limiting amounts in the thymus35. This hypothesis is further supported by our observation that in fetal thymus organ cultures, mature Treg cells inhibited IL-2-dependent but not IL-2-independent development of Treg cells. This is, to our knowledge, the first direct demonstration of the homing of a population of mature leukocytes back to its organ of origin to inhibit the differentiation of its precursors. Similar inhibitory recirculation mechanisms may control the de novo development of other cell types of the immune system. A decreased frequency of RTEs (measured by quantification of TCR excision circles, which are gene-recombination products generated during T cell differentiation) has been found in the peripheral T cell population from patients with adult T cell leukemia relative to that in healthy subjects 36, which may be explained by a similar negative feedback loop for Tconv cells. Moreover, patients with plasma cell myeloma, in which transformed plasma cells home back to the bone marrow, suffer from B cell lymphopenia37, which suggests that plasma cells might inhibit the differentiation of their own precursors in the bone marrow. Upon emigrating from the thymus, Treg cells that encounter their cognate major histocompatibility complex–peptide ligand are stimulated to proliferate38. This mechanism is thought to enrich the peripheral Treg cell repertoire for ‘useful’ cells. Thus, the function of the recirculation of Treg cells to the thymus may be to reduce, with aDVANCE ONLINE PUBLICATION  nature immunology

Articles age, the production of new Treg cells and thus prevent dilution of the peripheral Treg cells repertoire with cells of unproven specificity. The recirculation of Treg cells to the thymus might also convey information about immunoregulation in the periphery to this organ of T lymphopoiesis and thus potentially adapt T cell differentiation to peripheral immune responses. Methods Methods and any associated references are available in the online version of the paper. Accession codes. NCBI Sequence Read Archive: RNA-seq data, SRP056191.

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Note: Any Supplementary Information and Source Data files are available in the online version of the paper. Acknowledgments We thank A. Lew (Walter & Eliza Hall Institute of Medical Research) and A. Egle (Salzburg Cancer Research Institute) for GK-transgenic mice; P. Fink (University of Washington, Seattle) for B6 Rag-GFP mice; A. Rudensky (Memorial Sloan-Kettering Cancer Center) and A. Liston (University of Leuven) for Foxp3-Thy-1.1 mice; S. Guerder and J.-C. Guéry for critical reading of the manuscript; F. Auriol and A. Garnier for help in obtaining human thymuses; F.-E. L’Faqihi-Olive and V. Duplan-Eche for technical assistance at the flow-cytometry facility; S. Allart and A. Canivet for technical assistance at the cellular imaging facility of Inserm U1043, Toulouse; the personnel of the Inserm US006 ANEXPLO/CREFRE animal facility for expert animal care; and the GeT-PlaGe and GenoToul bioinformatics platforms Toulouse Midi-Pyrénées for sequencing, computing and storage resources. Supported by the Fondation pour la Recherche Médicale (O.P.J. and J.P.M.v.M.), the IdEx Toulouse (E.A.R. and P.R.), the Région Midi Pyrénées (O.P.J. and J.P.M.v.M.) and the Agence Nationale pour la Recherche (O.P.J.). AUTHOR CONTRIBUTIONS N.T., J.D. and V.A. performed experiments, analyzed data and contributed to writing the manuscript; M.G. and B.B. performed experiments and analyzed data; C.P. performed experiments; B.L. provided clinical samples; N.F., O.P.J. and E.A.R. designed experiments and analyzed data; J.P.M.v.M. and P.R. designed experiments, analyzed the data and wrote the manuscript; and all authors critically reviewed the manuscript. COMPETING FINANCIAL INTERESTS The authors declare no competing financial interests. Reprints and permissions information is available online at http://www.nature.com/ reprints/index.html. 1. Sakaguchi, S., Miyara, M., Costantino, C.M. & Hafler, D.A. FOXP3+ regulatory T cells in the human immune system. Nat. Rev. Immunol. 10, 490–500 (2010). 2. Josefowicz, S.Z., Lu, L.F. & Rudensky, A.Y. Regulatory T cells: mechanisms of differentiation and function. Annu. Rev. Immunol. 30, 531–564 (2012). 3. Romagnoli, P., Hudrisier, D. & van Meerwijk, J.P.M. Preferential recognition of self-antigens despite normal thymic deletion of CD4+CD25+ regulatory T cells. J. Immunol. 168, 1644–1648 (2002). 4. Hsieh, C.S. et al. Recognition of the peripheral self by naturally arising CD25+ CD4+ T cell receptors. Immunity 21, 267–277 (2004). 5. Chidgey, A., Dudakov, J., Seach, N. & Boyd, R. Impact of niche aging on thymic regeneration and immune reconstitution. Semin. Immunol. 19, 331–340 (2007). 6. Nikolich-Žugich, J. Ageing and life-long maintenance of T-cell subsets in the face of latent persistent infections. Nat. Rev. Immunol. 8, 512–522 (2008). 7. Yu, W. et al. Continued RAG expression in late stages of B cell development and no apparent re-induction after immunization. Nature 400, 682–687 (1999). 8. Borgulya, P., Kishi, H., Uematsu, Y. & von Boehmer, H. Exclusion and inclusion of alpha and beta T cell receptor alleles. Cell 69, 529–537 (1992). 9. McCaughtry, T.M., Wilken, M.S. & Hogquist, K.A. Thymic emigration revisited. J. Exp. Med. 204, 2513–2520 (2007).

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10. Liston, A. et al. Differentiation of regulatory Foxp3+ T cells in the thymic cortex. Proc. Natl. Acad. Sci. USA 105, 11903–11908 (2008). 11. Almeida, A.R. et al. Quorum-sensing in CD4+ T cell homeostasis: a hypothesis and a model. Front Immunol. 3, 125 (2012). 12. Liston, A. & Gray, D.H. Homeostatic control of regulatory T cell diversity. Nat. Rev. Immunol. 14, 154–165 (2014). 13. Cuss, S.M. & Green, E.A. Abrogation of CD40–CD154 signaling impedes the homeostasis of thymic resident regulatory T cells by altering the levels of IL-2, but does not affect regulatory T cell development. J. Immunol. 189, 1717–1725 (2012). 14. Yang, E., Zou, T., Leichner, T.M., Zhang, S.L. & Kambayashi, T. Both retention and recirculation contribute to long-lived regulatory T-cell accumulation in the thymus. Eur. J. Immunol. 44, 2712–2720 (2014). 15. Attridge, K. & Walker, L.S. Homeostasis and function of regulatory T cells (Tregs) in vivo: lessons from TCR-transgenic Tregs. Immunol. Rev. 259, 23–39 (2014). 16. Fink, P.J., Swan, K., Turk, G., Moore, M.W. & Carbone, F.R. Both intrathymic and peripheral selection modulate the differential expression of Vβ5 among CD4+ and CD8+ T cells. J. Exp. Med. 176, 1733–1738 (1992). 17. McMahan, C.J. & Fink, P.J. RAG reexpression and DNA recombination at T cell receptor loci in peripheral CD4+ T cells. Immunity 9, 637–647 (1998). 18. Papiernik, M., de Moraes, M.L., Pontoux, C., Vasseur, F. & Penit, C. Regulatory CD4 T cells: expression of IL-2Rα chain, resistance to clonal deletion and IL-2 dependency. Int. Immunol. 10, 371–378 (1998). 19. Zhan, Y., Corbett, A.J., Brady, J.L., Sutherland, R.M. & Lew, A.M. Delayed rejection of fetal pig pancreas in CD4 cell deficient mice was correlated with residual helper activity. Xenotransplantation 7, 267–274 (2000). 20. Agus, D.B., Surh, C.D. & Sprent, J. Re-entry of T cells to the adult thymus is restricted to activated T cells. J. Exp. Med. 173, 1039–1046 (1991). 21. Leng, Q., Nie, Y., Zou, Y. & Chen, J. Elevated CXCL12 expression in the bone marrow of NOD mice is associated with altered T cell and stem cell trafficking and diabetes development. BMC Immunol. 9, 51 (2008). 22. Bajoghli, B. et al. Evolution of genetic networks underlying the emergence of thymopoiesis in vertebrates. Cell 138, 186–197 (2009). 23. Joller, N. et al. Treg cells expressing the coinhibitory molecule TIGIT selectively inhibit proinflammatory Th1 and Th17 cell responses. Immunity 40, 569–581 (2014). 24. Lee, H.M. & Hsieh, C.S. Rare development of Foxp3+ thymocytes in the CD4+CD8+ subset. J. Immunol. 183, 2261–2266 (2009). 25. Pandiyan, P., Zheng, L., Ishihara, S., Reed, J. & Lenardo, M.J. CD4+CD25+Foxp3+ regulatory T cells induce cytokine deprivation-mediated apoptosis of effector CD4+ T cells. Nat. Immunol. 8, 1353–1362 (2007). 26. Cheng, G., Yu, A. & Malek, T.R. T-cell tolerance and the multi-functional role of IL-2R signaling in T-regulatory cells. Immunol. Rev. 241, 63–76 (2011). 27. Lio, C.W. & Hsieh, C.S. A two-step process for thymic regulatory T cell development. Immunity 28, 100–111 (2008). 28. Kimmig, S. et al. Two subsets of naive T helper cells with distinct T cell receptor excision circle content in human adult peripheral blood. J. Exp. Med. 195, 789–794 (2002). 29. Miyara, M. et al. Functional delineation and differentiation dynamics of human CD4+ T cells expressing the FoxP3 transcription factor. Immunity 30, 899–911 (2009). 30. Dominguez-Villar, M., Baecher-Allan, C.M. & Hafler, D.A. Identification of T helper type 1-like, Foxp3+ regulatory T cells in human autoimmune disease. Nat. Med. 17, 673–675 (2011). 31. Duhen, T., Duhen, R., Lanzavecchia, A., Sallusto, F. & Campbell, D.J. Functionally distinct subsets of human FOXP3+ Treg cells that phenotypically mirror effector Th cells. Blood 119, 4430–4440 (2012). 32. Bosco, N., Agenes, F., Rolink, A.G. & Ceredig, R. Peripheral T cell lymphopenia and concomitant enrichment in naturally arising regulatory T cells: the case of the pre-Tα gene-deleted mouse. J. Immunol. 177, 5014–5023 (2006). 33. Romagnoli, P. et al. The thymic niche does not limit development of the naturally diverse population of mouse regulatory T lymphocytes. J. Immunol. 189, 3831–3837 (2012). 34. Smigiel, K.S. et al. CCR7 provides localized access to IL-2 and defines homeostatically distinct regulatory T cell subsets. J. Exp. Med. 211, 121–136 (2014). 35. Tai, X. et al. Foxp3 transcription factor is proapoptotic and lethal to developing regulatory T cells unless counterbalanced by cytokine survival signals. Immunity 38, 1116–1128 (2013). 36. Yasunaga, J. et al. Impaired production of naive T lymphocytes in human T-cell leukemia virus type I-infected individuals: its implications in the immunodeficient state. Blood 97, 3177–3183 (2001). 37. Pilarski, L.M., Mant, M.J., Ruether, B.A. & Belch, A. Severe deficiency of B lymphocytes in peripheral blood from multiple myeloma patients. J. Clin. Invest. 74, 1301–1306 (1984). 38. Fisson, S. et al. Continuous activation of autoreactive CD4+ CD25+ regulatory T cells in the steady state. J. Exp. Med. 198, 737–746 (2003).



ONLINE METHODS

Mice. Rag2-GFP mice7 on a B6 genetic background39 were provided by P. Fink; B6 Foxp3-Thy-1.1 mice10 were provided by A. Rudensky and A. Liston; and B6 GK mice19 were provided by A. Lew and A. Egle. All comparisons were performed with groups with identical sex ratios. NMRI mice at day 15 of pregnancy, used for fetal thymus organ culture, were from Janvier. All experiments involving animals were performed in compliance with the relevant laws and institutional guidelines (Regional approval 310955545; ethical review MP/01/40/11/06).

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Human samples. Human thymus tissue from children undergoing cardiac surgery was obtained from the Cardiology Department of the Purpan Children’s Hospital in Toulouse. In line with French guidelines, French authorities authorized the study (ministry approval DC-2014-2088), and the participants’ legal representatives did not oppose the use of the tissue for research purposes. Antibodies. The following mAbs and secondary reagents were used for phenotypic analysis of mouse T cells: Pacific Blue–labeled anti-CD4 (GK1.5), phycoerythrin-CF594– or Alexa Fluor 700–labeled anti-CD8 (53.6.7), phycoerythrin-indotricarbocyanine–labeled anti-CD25 (PC61), allophycocyanin-labeled anti-Thy-1.1 (HIS51), biotin- or phycoerythrinlabeled anti-Vβ3 (KJ25), biotin-labeled anti-Vβ4 (K74), biotin- or phycoerythrin-labeled anti-Vβ5 (MR9-4), biotin- or phycoerythrin-labeled anti-Vβ6 (RR4-7), biotin-labeled anti-Vβ11 (RR3-15) and biotin-labeled anti-Vβ14 (14-2) (all from BD Bioscience); biotin-labeled anti-TCRβ (H57-597), phycoerythrin-indotricarbocyanine–labeled anti-CD44 (IM7), allophycocyanin-labeled anti-CD62L (MEL-14), peridinin chlorophyll protein–eFluor 710–labeled anti-TIGIT (GIGD7), phycoerythrinindotricarbocyanine– or eFluor 710–labeled streptavidin and biotin-labeled anti-CD103 (2E7) (all from eBioscience); and phycoerythrin-labeled anti-neuropilin (761705; R&D Systems). The following mAbs were used for phenotypic analysis of human T cells: fluorescein isothiocyanate–labeled anti-CD4 (RPA-T4), Brilliant Violet 405–labeled anti-CD8 (RPA-T8), phycoerythrin-indotricarbocyanine–labeled anti-CD3 (SK7), phycoerythrinlabeled anti-CD31 (WM59), peridinin chlorophyll protein–cyanine 5.5–labeled anti-ICOS (DX29), Brilliant Violet 605–labeled anti-CD31 (WM59), phycoerythrin-CF594–labeled anti-CD25 (M-A251) and phycoerythrin-CF594–labeled anti-CD45RA (Hl100) (all from BD Bioscience); and Alexa Fluor 647–labeled anti-Foxp3 (259D/C7), and phycoerythrinlabeled anti-TBX21 (eBio4B10) (both from eBioscience). Flow cytometry. Control experiments with mice given injection of India Ink 30 min before being killed were performed initially to ensure the surgical removal of the thymus without parathymic lymph nodes. Thymi, spleens and lymph nodes were homogenized, then were washed once in complete culture medium and resuspended in medium containing rat and mouse IgG (10 µg/ml) for blockade of nonspecific mAb binding. After 20 min of incubation on ice, saturating concentrations (empirically determined) of mAb to the relevant marker were added. Twenty min later, cells were washed three times in PBS containing 2.5% FCS and 0.02% NaN3 and were incubated with the appropriate secondary reagent. Intracellular staining was performed with Foxp3 staining kits (eBioscience). Labeled cells were analyzed on an LSRII or a Fortessa flow cytometer (BD Bioscience), and the acquired list-mode data were analyzed with FlowJo software (TreeStar). Doublets and dead cells were excluded by use of the appropriate forwardand side-scatter gates. Cell-cycle analysis. Thymocyte samples from GK-transgenic and non–GKtransgenic Rag-GFPFoxp3-Thy-1.1 mice were depleted of CD8+ cells as described33 and were stained with Pacific Blue–conjugated anti-CD4 and phycoerythrin-conjugated mAb to Thy-1.1 (each identified above). After two washes in PBS, cells were incubated for 30 min on ice with the viability dye eFluor 506 (eBioscience). Labeled cells were washed and resuspended in complete cell culture medium and were incubated at 37 °C with 5 µM Vibrant Dyecycle Ruby Stain according to the manufacturer’s instructions (Life Technologies). Cells were then analyzed with a Fortessa flow cytometer (BD Bioscience) and FlowJo software (TreeStar). Doublets were excluded

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through use of the appropriate forward and side scatter, as well as Ruby-H and Ruby-A gates (pulse height (H) and area (A) assessed with Ruby Stain), and dead cells were excluded through use of the viability dye eFluor 506. RNA-seq analysis. GFP+ and GFP− CD4+CD8− Thy-1.1+ Treg cells from thymus and spleen of Rag-GFPFoxp3-Thy-1.1 mice were sorted by flow cytometry, total RNA was immediately extracted with an RNeasy Micro Kit (Qiagen) (including a step of treatment with DNase I) and the quality of the RNA was assessed with an Agilent 2100 BioAnalyzer (Agilent Technologies). Biological replicates included cells pooled from 12 8-week-old male mice. Libraries for RNA-seq were prepared according to the TotalScript RNA-seq protocol (Epicentre), starting with 5 ng high-quality total RNA (i.e., RNA integrity number, >7), by the oligo(dT) primer–synthesis strategy. The quality of each library was assessed with an Agilent 2100 BioAnalyzer. Samples were indexed and sequenced on an Illumina HiSeq 2000 (two paired-end sequences of 100 base pairs each). ‘Reads’ were trimmed through use of the Cutadapt tool (version 1.3), with removal of low-quality bases (−q value,