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“Natural” regulatory T (nTreg) cells that express the transcription factor Foxp3 and produce IL-10 are required for systemic immunological tolerance. “Induced” Treg (iTreg) cells are non-redundant and essential for tolerance at mucosal surfaces, yet their mechanisms of suppression and stability are unknown. We investigated the role of iTreg cell-produced IL-10 and iTreg cell fate in a treatment model of inflammatory bowel disease. Colitis was induced in Rag1−/− mice by the adoptive transfer of naïve CD4+ T cells carrying a non-functional Foxp3 allele. At the onset of weight loss, mice were treated with both iTreg and nTreg cells where one marked subset was selectively IL-10-deficient. Body weight assessment, histological scoring, cytokine analysis, and flow cytometry were used to monitor disease activity. Transcriptional profiling and TCR repertoire analysis were used to track cell fate. When nTreg cells were present but IL-10 deficient, iTreg cell-produced IL-10 was necessary and sufficient for the treatment of disease, and vice versa. Invariably, ~85% of the transferred iTreg cells lost Foxp3 expression (ex-iTreg) but retained a portion of the iTreg transcriptome, which failed to limit their pathogenic potential upon retransfer. TCR repertoire analysis revealed no clonal relationships between iTreg and ex-iTreg cells, either within mice or between mice treated with the same cells. These data identify a dynamic IL-10-dependent functional reciprocity between Treg subsets that maintains mucosal tolerance. The niche supporting stable iTreg cells is limited and readily saturated, which promotes a large population of ex-iTreg cells with pathogenic potential during immunotherapy.
CD4+ CD25+ Foxp3+ regulatory T (Treg) cells are essential to the balance between pro- and anti-inflammatory responses at mucosal surfaces (1). There are two subsets of Treg cells, natural Treg (nTreg) cells that develop in the thymus and induced Treg (iTreg) cells that arise from conventional T (Tconv) cells in the periphery (2–6). Immunologic tolerance requires both Treg subsets, which act synergistically (6, 7). Additionally, iTreg cells can be generated in vitro by T cell activation in the presence of TGF-β1 and IL-2 which makes them an attractive alternative for the treatment of human autoimmune disorders unresponsive to current approaches (8–11). In vitro-derived iTreg cells also suppress inflammation in animal models of inflammatory bowel disease, diabetes, and autoimmune gastritis (12–14). Importantly, in vitro-derived iTreg cells contribute to tolerance in disease models where in vivo-derived iTreg cells are not present (6, 7). The pathways utilized by iTreg cells to support disease resolution in these models remain unknown and it is unclear to what extent the method of iTreg cell derivation influences the acquisition of the full complement of Treg cell suppressive mechanisms.
Although much work has been done to uncover the molecular mechanisms of Treg cell suppressive activity, the “division of labor” between nTreg cells and iTreg cells remains largely unresolved (2). Molecules such as CTLA-4, Granzyme B, IL-10 and TGF-β have been proposed as mechanisms of nTreg cell-mediated suppression (15–18). Of these mechanisms, the overall importance of IL-10 to immune homeostasis is exemplified by the chronic colitis that develops with complete or APC-specific IL-10 deficiency (19, 20). IL-10 is particularly important for Treg cells at environmental interfaces, as a Treg cell-specific inactivation of IL-10 results in spontaneous colitis (16). In the CD45RBhi transfer model of colitis, IL-10 present in the CD45RBlow population, which contains Treg cells, was found to be necessary for disease prevention and treatment (21, 22). Furthermore, Treg cell-derived IL-10 was shown to control Th17 and Th1+Th17 cells (23). To date, studies have been limited to the role of IL-10 in nTreg cell function, leaving unresolved the role of IL-10 as an iTreg cell suppressive mechanism.
Here, we used cells from mice that harbor a cassette encoding a mutant Foxp3-enhanced green fluorescent fusion protein within the Foxp3 locus (Foxp3ΔEGFP) to create colitis in an environment free of iTreg cells (6, 24). Following immunotherapy with nTreg and iTreg cells that were selectively deficient in IL-10, we investigated how the source of IL-10 impacted disease progression. We also examined the clonal and transcriptional relationships between those iTreg cells that maintained or lost Foxp3 expression (ex-iTreg cells). Our results identify in vivo selection of a potent iTreg cell pool that is clonally distinct from pathogenic ex-iTreg cells, and IL-10-dependent reciprocal compensation between Treg subsets as critical for the maintenance of mucosal tolerance.
Foxp3EGFP and Foxp3ΔEGFP mice on the BALB/c background were generated and screened as previously described (24). Thy1.2+ Foxp3ΔEGFP newborn mice were rescued by intraperiotneal (i.p.) transfer of 60×106 unfractionated Thy1.1+ BALB/c splenocytes to generate naive Thy1.2+ CD4+ CD45RBhi T cells with the nonfunctional Foxp3ΔEGFP allele. Rag1−/− and IL10−/− mice were obtained from The Jackson Laboratory. The Animal Resource Committee at the Medical College of Wisconsin approved all animal experiments.
Pooled splenocytes and lymph node cells (axillary, brachial, inguinal, and mesenteric) were stained with either anti-CD4-allophycocyanin (RM4-5, BD Biosciences) or anti-CD4-Pacific Blue (RM4-5, Invitrogen), plus anti-CD90.1-PerCP (OX-7, BD Biosciences) and anti-CD45RB-allophycocyanin (C363.16A, eBioscience) as appropriate and sorted on the basis of Ab and EGFP fluorescence. All sorting was done on a FACSAria (BD Biosciences). The average purity and viability of the sorted CD4+ populations was 98.96 +/− 0.14 and 84.31 +/− 0.68 (n=120), respectively.
Colitis was induced in 6 to 8 week old Rag1−/− BALB/c mice by i.p. injection of 4×105 CD4+ CD90.1− EGFP− CD45RBhi cells. Mice were weighed twice weekly. In some experiments, when mice lost 2.5% (+/− 5.7%) of their initial body weight or began to exhibit symptoms of colitis (diarrhea, hunched posture), they were treated by i.p. injection of nTreg (Thy1.2+) plus iTreg (Thy1.1+) cells (5×105 each) purified by cell sorting. The nTreg cells were isolated on the basis of EGFP expression from the spleen and lymph nodes of Foxp3EGFP BALB/c mice or IL10−/− Foxp3EGFP BALB/c mice. The iTreg cells were generated in culture (as described below) and isolated on the basis of EGFP expression.
For serial adoptive transfer experiments 1×105 CD4+ Thy1.1+ EGFP− ex-iTreg cells were isolated by sorting cells from the MLN and spleen of mice that were successfully treated with Thy1.1+ WT iTreg cells plus Thy1.2+ WT nTreg cells and injected i.p. into Rag1−/− hosts. In some experiments the mice were treated with 0.5×106 Thy1.2+ nTreg cells plus 0.5×106 Thy1.2+ iTreg cells and were co-injected with 15,000 Thy1.1+ EGFP− T cells (based on a 3% sort impurity, 3 times the calculated impurity). The Thy1.1+ EGFP− T cells were isolated from iTreg cells that had lost Foxp3 expression in culture.
Sorted CD4+ EGFP− cells from Foxp3EGFP or IL10−/− FoxpEGFP mice (1×106/mL) were cultured with anti-CD3 mAb (clone 14-2C11 at 2.5 µg/mL) coated dishes in the presence of soluble anti-CD28 mAb (1 µg/mL; clone 37.51), TGF-β1 (5 ng/mL; R&D Systems), and 100 U/mL IL-2. After 72h, cells were resorted based upon EGFP fluorescence and used for adoptive transfer or maintained in culture with IL-2.
The entire colon and the distal 15 cm of the small intestine were used as the source of intra-epithelial lymphocytes (IELs) and lamina propria lymphocytes (LPLs) (25). IELs were removed by gentle shaking of 0.5-cm intestinal sections for 30 min in buffer containing 5% (vol/vol) FCS, 1 mM dithiothreitol (Sigma-Aldrich) and 5 mM EDTA. IELs were washed and isolated on a discontinuous Percoll gradient (67%, 44%). Washed intestinal sections were digested with collagenase D (1 mg/mL, Roche) in the presence of DNase I (Invitrogen). A discontinuous Percoll gradient (67%, 44%) was used to isolate washed LPLs.
Cells were collected from the spleen, MLN, colon and small intestine and stained as indicated. The anti-mouse antibodies used were Pacific Blue-conjugated anti-CD4 (RM4-5, Invitrogen); PerCP-conjugated anti-CD90.1 (OX-7, BD Biosciences); Alexa Fluor-conjugated anti-CD44 (IM7, Biolegend), PE-conjugated anti-CD62L (MEL-14, BD Biosciences); Alexa Fluor 647-conjugated anti-CD103 (2E7, Biolegend); PE-Texas Red-conjugated anti-CD25 (PC61 5.3, Invitrogen); PE-Cy7-conjugated Klrg1 (2F1, eBioscience); APC eFluor 780-conjugated anti-TCRβ (H57-597, eBioscience); and APC-conjugated anti-GITR (DTA-1, eBioscience). A four-laser custom LSRII was used to collect the data, and FlowJo software was used for analysis.
Intracellular cytokine staining was performed after a 5 hour restimulation with phorbol 12-myristate 13-acetate (PMA, 5 ng/mL, Sigma-Aldrich) and ionomycin (0.5 µM, Sigma-Aldrich) in the presence of brefeldin A (1 μL/mL; BD Biosciences). Surface staining of cells was performed using a modified FACS buffer containing 10 μg/mL brefeldin A. Cells were stained on ice for 30 min with the primary anti-mouse antibodies PE-Cy7-conjugated anti-CD4 (RM4-5, BD Biosciences), PerCP-conjugated anti-CD90.1 (OX-7, BD Biosciences), and APC eFluor 780-conjugated anti-TCRβ (H57-597, eBioscience) then washed with the modified FACS buffer and fixed in 1% paraformaldehyde overnight at 4°C. After this incubation, cells were washed with 1mL PBS and then permeabilized with 1mL 0.1% Triton-X. Intracellular staining was performed for 30 minutes at room temperature with APC conjugated anti-IFN-γ (XMG1.2, BD Biosciences), Alexa Fluor 700 conjugated anti-TNF-α (MP6-XT22, BD Biosciences), Pacific Blue-conjugated anti-IL-17A (TC11-18H10.1, Biolegend) or with Pacific Blue-conjugated anti-Helios (22F6, Biolegend), and APC-conjugated anti-CTLA-4 (UC10-4B9, Biolegend). A four-laser custom LSRII was used to collect the data, and FlowJo software was used for analysis. Serum cytokines were measured using the eBioscience FlowCytomix kit following the manufacturer’s recommendations.
Complete colons were fixed in formalin, processed, and stained with H&E using a histology core facility. Blinded sections from the entire colon were examined by a pathologist (N.H.S.) and large intestine colitis scores were determined for the following inflammatory changes on a 4-point semiquantitative scale with 0 representing no change (26). The following features were considered: severity, depth and chronic nature of the inflammatory infiltrate, crypt abscess formation, granulomatous inflammation, epithelial cell hyperplasia, mucin depletion, ulceration, and crypt loss.
Spleen and MLN cells of mice that were successfully treated with 0.5×106 Thy1.1+ WT iTreg cells plus 0.5×106 Thy1.2+ WT nTreg cells were stained with Pacific Blue-conjugated anti-CD4 (RM4-5, Invitrogen) and PerCP-conjugated anti-CD90.1 (OX-7, BD Biosciences) and were sorted on the basis of Ab and EGFP fluorescence. ex-iTreg (CD4+ Thy1.1+ EGFP−), iTreg (CD4+ Thy1.1+ EGFP+), and nTreg cells (CD4+ Thy1.1– EGFP+) were isolated by flow cytometry sorting on a FACSAria. Total RNA was extracted with the RNeasy Micro Kit for <100,000 cells or the RNeasy Mini Kit for >100,000 cells (Qiagen) according to the manufacturer’s protocol. cDNA was synthesized with the Superscript III First Strand Synthesis System and oligo(dT) primers (Invitrogen) according to the manufacturer’s protocol. Isolated cDNA was used for quantitative PCR, spectratype analysis, or CDR3 sequencing.
qPCR was performed in an StepOnePlus PCR System (Applied Biosystems) using the TaqMan Fast Universal PCR Master Mix (Applied Biosystems) and pre-developed specific Taqman primers for Il10 (Mm00439616_m1). GAPDH was used (Mm99999915_g1) as a housekeeping gene (Applied Biosystems). The standard curve for Il10 was developed by isolating bone marrow cells from C57BL/6J and culturing the cells at 4×104 cell/mL in DMEM/F12-10 media (DMEM/F12 (GIBCO), 10% (v/v) FBS (Atlantic Biologics), 10 mM L-glutamine (GIBCO), 100 IU penicillin/mL (GIBCO), 100 µg/mL streptomycin (GIBCO)) at 4×106 cells/mL in the presence of 10 U/mL MCSF (eBioscience, 14-8983-80). The cells were incubated at 37°C for 7 days (media was changed on day 3). Macrophages were isolated using Cellstripper (Cellgro), washed, and resusupended in DMEM/F12-10 at 2×105 cells/mL. The cells were plated for 8 hours at 37°C in a 24 well plate containing 50 ng/mL LPS (Sigma), 1 µg/mL OVA (323–339 peptide), and 15 µL/mL anti-Ova (AbD Serotec, 0220-1682). cDNA was isolated as described and the Il10 gene from the exon 2/3 boundary (forward primer: 5’–AATGCAGGACTTTAAGGGTTACTTGGG–3’) to the exon 4/5 boundary (reverse primer: 5’– CTTGTAGACACCTTGGTCTTGGAG–3’) was cloned into the pCR4-TOPO vector (Invitrogen) and transformed into DH5α cells. The plasmid was isolated with a mini prep (Promega) and then purified by agarose gel extraction. The concentration was determined using log-dilutions and measured with a NanoDrop (Invitrogen). The standard curve was created with 10 fold dilutions based on copy number.
Total RNA for the iTreg, nTreg, and ex-iTreg sets was isolated with TRIzol (Invitrogen), according to the manufacturer’s protocol, from sorted cells pooled from 14 mice treated with 0.5×106 WT nTreg plus 0.5×106 WT iTreg cells. Labeled target was prepared and hybridized to Affymetrix 430 2.0 GeneChips in accordance with the manufacturer’s protocol. Two technical replicates were performed and the results were averaged. Probe sets that revealed a twofold difference (|log2 ratio| > 1.0) relative to Tconv cells were identified and used in subsequent analyses. The data was normalized with the Robust Mulit-array Analysis (RMA) algorithm derived by the Bioconductor group (http://www.bioconductor.org) (27). The mean fold change was calculated from 2 independent arrays for each cell type and was scored P<0.05 with a false discovery rate (FDR) <10% by the non-parametric rank product test (28). A false negative result is recovered for Foxp3+ Treg cells because the gene array probes for Foxp3 lie distal to the poly-A initiation site in the Foxp3EGFP allele. The microarray data are available in the Gene Expression Omnibus (GEO) database (http://www.ncbi.nlm.nih.gov/gds) under the accession number GSE35543. Tconv cell data was taken from GEO microarray data set GSE6875.
The methylation status of the Treg cell-specific demethylation region (TSDR) of Foxp3 in donor male nTreg, iTreg cells, and ex-iTreg cells purified from mice treated with WT nTreg plus WT iTreg cells by cell sorting, was assessed by bisulphite sequence analysis (29). Briefly, genomic DNA was treated by bisulfite, to convert unmethylated cytosines into uracil leaving methylated cytosines unchanged. Bisulfite conversion of DNA was performed using EZ DNA methylation-Direct™ Kit (Zymo research) according to the manufacturer instructions. The TSDR of converted DNA was amplified by methylation-specific primer sequences 5'-TATTTTTTTGGGTTTTGGGATATTA-3' (forward) and 5'-AACCAACCAACTTCCTACACTATCTAT-3' (reverse). The PCR product was purified and inserted into a TOPO TA cloning vector (Invitrogen). The ligation product was used to transform competent bacteria (10–beta competent E. coli, New England Biolabs) and clones were selected on kanamycin. Plasmid DNA was extracted by Qiagen miniprep kit and clones that present a 300bp fragment after EcoRI digestion were selected. Sequencing was done with M13R primer. Blast analysis was done by comparing the M13R sequence and converted FoxP3 gene sequence.
cDNA was amplified by PCR with a Cβ primer (5’ CTCAAACAAGGAGACCTTGGGTGG– 3’) and a Vβ primer from one of 22 Vβs (30). An ABI 3100 Genetic Analyzer was used to analyze the length distribution of amplified cDNA products and Xplorer v2.4.2 (http://www.dnatools.com/download.html) was used to create histograms.
cDNA was amplified by PCR with the C primer and a V 8.2 primer (5’ – GCTACCCCCTCTCAGACATCAGTG– 3’). PCR products were purified using the QIAquick PCR purification kit (Qiagen) according to the manufacturer’s protocol and concentrated using ethanol precipitation. The DNA sample was purified using Agencourt AMpure beads. The purified samples were used to generate libraries for Next - Generation Sequencing using the Ion Torrent Personal Genome Machine (PGM) (Life Technologies, Carlsbad, CA) following the Ion Fragment Library Kit protocol. During preparation of the libraries, samples were size selected using the Sage Science Pippen Prep instrument and 2% agarose cassettes. Completed libraries were analyzed and quantified on the Agilent 2100 BioAnalyzer. The obtained libraries were processed for sequencing by dilution following the recommendations for the Ion Xpress Template Kit protocol that utilized an emulsion PCR, breaking and enrichment of each sample. Sequencing was performed using the Ion Torrent 314 or 316 chip and respective reagents. Sequence analysis and base calling was performed using the built in sequence software (v.1.9).
The purified PCR products were TA cloned into the pCR4-TOPO vector (Invitrogen) to create cDNA libraries. Individual colonies were subcloned and plasmids containing inserts were grown for 16 hours in LB media, frozen at -80°C in 50% glycerol and sent to Beckman Coulter Inc. for sequencing. The CDR3 regions were identified as the sequence between the second conserved cysteine encoded by the 5’ Vβ gene segment and the conserved phenylalanine encoded by the 3’ Jβ segment (IMGT).
The comparisons between groups for overall survival functions were done using the log-rank test. The random coefficient model was used to generate a quadratic fit of the weight change over time. For the colitis scores, cell frequencies and numbers, and serum cytokine levels a non-parametric Kruskal-Wallis test was used to compare the measurements between the groups. For the pairwise comparisons a Mann-Whitney U test was performed. The TCR repertoire data was analyzed using the Morisita-Horn Index, which was calculated using EstimateS software.
In order to establish the relevance of iTreg cell-produced IL-10 in a setting containing the nTreg cell subset, we induced colitis in BALB/c Rag1–/– mice by transferring 4×105 Thy 1.2+ EGFP– CD45RBhi cells isolated by cell sorting from BALB/c Thy1.2+ Foxp3ΔEGFP mice. The transferred CD45RBhi cells have a non-functional Foxp3 allele, and colitis develops with accelerated kinetics in the absence of in vivo derived iTreg cells (6). When mice lost ~2.5% of their initial body weight we treated them with different combinations of 5×105 iTreg cells plus 5×105 nTreg cells, where one or both of the Treg subsets lacked the capacity to produce IL-10 (Figure 1). For all experiments, the iTreg cells were derived in vitro from CD4+ EGFP– cells isolated from the spleens and lymph nodes of Foxp3EGFP or Il10–/– Foxp3EGFP mice and cultured with TGF-β1 and TCR crosslinking to upregulate Foxp3 expression. The nTreg cells were isolated by cell sorting from the spleen and lymph nodes of these same mice.
In the absence of any therapeutic intervention, mice rapidly lost weight, became moribund and were sacrificed, with a mean survival of 45 days (Figure 1A,B). In treated mice, where the transferred iTreg cell compartment supplied the only source of Treg cell-derived IL-10, weight gain and survival were similar to control mice that were treated with WT iTreg cells plus WT nTreg cells (Figure 1A). Reversing the experiment, where the nTreg compartment supplied the IL-10, showed similar patterns of weight gain and survival. In contrast, complete absence of all IL-10 produced by Treg cells, while better than no therapy, still resulted in weight loss and only 25% of the mice survived to 125 days (Figure 1A,B). When Treg cell-derived IL-10 was limited to iTreg cells there was also less inflammatory infiltrate in the colons of treated mice (Figure 1C,D). Serum levels of IFN-γ were reduced when IL-10 was produced by at least one Treg cell compartment, while TNF-α was reduced in all treatment groups irrespective of the capacity of Treg cells to produce IL-10 (Figure 1E). Serum levels of IL-17A were not reduced with Treg cell treatment.
An increase in iTreg cell frequency and/ or number was seen in the mesenteric lymph nodes (MLN), colon, and spleen of mice treated with WT iTreg plus Il10–/– nTreg cells when compared to control mice treated with WT iTreg plus WT nTreg cells (Figure 2A,B, Figure S1A,B and data not shown). However, the number of nTreg cells recovered from these tissues was similar between all treatment groups, demonstrating that there was no effect of Treg cell-derived IL-10 on nTreg cell recovery (Figure 2B and S1B). These data indicate that a compensatory expansion of WT iTreg cells occurs when nTreg cells lack the capacity to produce IL-10 and that differences in clinical outcome between the treatment groups are not due to an overall reduction in Treg cell numbers.
Next, we examined the impact of iTreg cell-produced IL-10 on effector T cells in the target tissues. Overall, the number of CD4+ T cells was reduced in the MLN and spleen of treated mice compared to untreated controls, irrespective of the IL-10 status of the Treg cells used to treat the mice (Figure 2C and data not shown). CD4+ T cells were further reduced in the MLN of mice treated with WT iTreg plus Il10–/– nTreg cells, suggesting a modest effect of iTreg cell produced IL-10 in this location. Consistent with the reduction in CD4+ T cells, the number of IFN-γ+, IL-17A+ or IFN-γ+ IL-17A+ effector CD4+ T cells in the MLN of treated mice was also reduced when the iTreg compartment selectively produced IL-10 (Figure 2D,E), and closely matched that seen in mice treated with Il10–/– iTreg plus WT nTreg cells. In the colon, the frequency of IL-17A+ and IFN-γ+ IL-17A+ T cells was reduced when Treg IL-10 production was limited to iTreg cells (Figure S1C). These data demonstrate that iTreg cells are a potent source of IL-10 and that iTreg cell suppression of lymphoproliferation, and Th1 and Th17 cell differentiation is closely associated with IL-10 production. The data also confirm a similar outcome with a reverse in experimental design. Additional mechanisms of Treg cell-mediated suppression may be operative, as a modest Treg-associated effect on weight change and survival was observed in the complete absence of Treg cell-derived IL-10.
In all treatment groups, many Thy1.1+ Foxp3+ iTreg cells lost Foxp3 expression (ex-iTreg) and were maintained in the MLN, colon, spleen, and small intestine of treated mice (Figure 2A, Figure S1A, and data not shown). Contaminating EGFP– cells from the sorted iTreg cultures contributed little (12%) to the ex-iTreg population (Figure S1D,E). When comparing groups, the frequency of ex-iTreg cells was highest in the MLN and spleen of mice treated with IL-10 sufficient iTreg cells (Figure 3A,B). This pattern was reversed in the colon and small intestine (Figure S1F). The ex-iTreg cells did not continue to produce IL-10 as demonstrated by RT-PCR of sorted cells (Figure S1G). However, in vitro stimulation of ex-iTreg cells isolated from multiple tissues resulted in the production of IFN-γ and IL-17A (Figure 3C and Figure S1H). Importantly, the frequency of ex-iTreg cells in the MLN that were IL-17A+ or IFN-γ+ IL-17A+ was reduced when iTreg cells produced IL-10 (Figure 3C). The frequency of ex-iTreg cells producing IL-17A was increased (18–27%) in the small intestines of mice from all treatment groups relative to the other tissues (data not shown). These data demonstrate that ex-iTreg cells can produce proinflammatory cytokines, and that the capacity of ex-iTreg cells to produce IL-17A and the localization of ex-iTreg cells may be influenced by iTreg cell-produced IL-10.
We investigated the phenotypic and molecular relationship between nTreg, iTreg and ex-iTreg cells by examining their cell surface phenotype, gene expression profile, and their methylation status at the Treg cell specific demethylation region (TSDR) of conserved noncoding sequence 2 (CNS2) within the Foxp3 promoter. Flow cytometric analysis of cells from the MLN of mice treated with WT iTreg plus WT nTreg cells revealed that iTreg and nTreg cells had similar expression of several cell surface markers including CD25, CD62L, CD44, CD103, KLRG1, GITR, and CTLA-4 (Figure 4A). Helios was expressed in 20.6% (+/− 5.8%) of the iTreg cells in the spleen, but was generally not found in iTreg cells in the MLN or in ex-iTreg cells (data not shown). Several differences were noted between ex-iTreg and iTreg cells found in the MLN. Compared to iTreg cells, the ex-iTreg cells had lower levels of CD25, CD103, KLRG1, and CTLA-4, although they had similar expression of CD62L, CD44, and GITR (Figure 4B).
Next, we sorted nTreg, iTreg, and ex-iTreg cells from the spleens and MLN of mice treated with WT iTreg plus WT nTreg cells, and examined their gene expression profiles. From the three groups, we identified a total of 3250 probe sets that were differentially regulated relative to naive CD4+ T cells (Table S1). This analysis captured 319 of the 603 probe sets found in the canonical Treg cell transcriptional signature (31). Interestingly, 1437 probe sets (44%) were common to all three groups and included about 40% (132/319) of the Treg probe sets that we recovered (Figure 4C). These included directionally concordant expression of the Treg signature genes Itgae, Lef1, Dusp4, Ctla4, Ahr, Gzmb, Ccr6, and Rora. In most cases, however, the ex-iTreg cells had the smallest fold changes in level of expression (Figure 4D). Most of the remaining Treg signature genes were suppressed or not differentially regulated in the ex-iTreg expression profile, including Ikzf4, Gpr83, Nrp1, Ikzf2, Pde3b, Il2ra, Ebi3, and Klrg1. The genetic signatures of in vitro-derived iTreg that were maintained in vivo and nTreg cells were remarkably similar (64% overlap, Figure 4C), consistent with our previous data examining the relationship between in vivo-derived iTreg cells and nTreg cells (7).
Ex-iTreg cells also differentially regulated a number of genes not contained within the canonical Treg signature (Figure 4E). Ex-iTreg cells showed induction of Il17a (30 fold versus Tconv cells), and Ifng (11 fold versus Tconv cells), consistent with the intracellular cytokine staining. In contrast, Il10 expression was largely confined to nTreg and iTreg cells (52 fold and 27 fold increases versus Tconv cells, respectively). The ex-iTreg cells expressed Il2, Il22, Ccl5, and Gzmk. Several other genes not associated with the Treg signature were expressed in ex-iTreg cells as well as both Treg subsets, such as Lyz1, Gp49a, S100a8, S100a9, and Klf4. In all three cell types, Ikzf1 (Ikaros) and Sox4 were repressed.
To further investigate the molecular relationship between the nTreg, iTreg and ex-iTreg cell subsets we analyzed the methylation status of the TSDR within CNS2 of the Foxp3 promoter of mice rescued with WT nTreg cells plus WT iTreg cells. As expected, the CpG islands in the TSDR of WT nTreg cells were demethylated (Figure 4F, top panel) (7, 29). Consistent with their lack of Foxp3 expression, the CpG islands in ex-iTreg cells showed extensive methylation at the examined CpG motifs. In aggregate, CpG islands in the TSDR of stable in vitro-derived iTreg cells isolated from successfully treated mice showed partial demethylation, although there was considerable variability in the methylation patterns found in individual mice. Interestingly, iTreg cells isolated from at least 2 mice were largely demethylated (Figure 4F, bottom panel). The extensive iTreg cell demethylation seen in these 2 mice did not correlate with Treg or ex-iTreg cell recovery, rate of weight loss following colitis induction, or the weight gained after treatment (data not shown).
Taken together, these gene and protein expression data indicate that ex-iTreg cells retain a component of the Treg canonical signature. However, loss of Foxp3 expression is associated with changes in the expression of several genes important for regulatory T cell suppressive function and in the acquisition of genes related to the inflammatory response and T cell effector capability (32–36). The general failure of most in vitro-derived iTreg cells to demethylate CpG islands in the TSDR likely contributed to the large number of ex-iTreg cells present in the treated mice, as demethylation of these residues has been shown to be important to iTreg cell stability (37). Fully demethylated iTreg cell TSDRs were occasionally observed, suggesting that iTreg cells per se are not excluded from stable Foxp3 expression.
The ex-iTreg cells expressed genes associated with regulation but also expressed genes associated with inflammation. Because of the dual nature exhibited by ex-iTreg cells, we tested their pathogenic potential by the adoptive transfer of 1×105 sorted ex-iTreg cells into Rag1–/– hosts. Recipients rapidly lost weight, became moribund and were sacrificed after ~40 days (Figure 5A,B). Histological analysis of the colon revealed lymphocytic infiltration and severe colitis (Figure 5C). Interestingly, an average of 2% of the transferred cells found in the MLN regained Foxp3 expression in vivo. This frequency of iTreg cells matched that seen after the transfer of 1×105 CD4+ EGFP– CD45RBhi cells into Rag1–/– mice (Figure 5D). Thus, in the absence of regulatory T cells, ex-iTreg cells cause severe disease. The capacity of ex-iTreg cells to upregulate Foxp3 is retained and is similar to that of naïve CD4+ T cells.
The capacity of ex-iTreg cells to reacquire Foxp3 expression after transfer into Rag1–/– hosts (Figure 5D) suggested that the ex-iTreg and iTreg cells could be interconverting and thus clonally related. To test this hypothesis we isolated iTreg and ex-iTreg cells from six mice treated with WT iTreg cells plus WT nTreg cells and characterized the TCR repertoires. Notably, mouse 1 and mouse 2 received equivalent aliquots drawn from the same population of pooled Treg cells. In a separate experiment, we also treated mouse 4 and mouse 6 by splitting a pooled Treg cell sample. Mouse 3 and mouse 5 each received an unrelated population of cells. This experimental design allowed us to compare the iTreg and ex-iTreg populations that were maintained within a single individual and between two individuals that received equal fractions of the same inoculum. Fragment analysis of 12 Vβ CDR3 regions from six treated mice showed both skewed and Gaussian distributions with limited similarity between the iTreg and ex-iTreg cell populations, both within individuals and between those individuals that received equivalent Treg cell populations (Figure S2A). For example, only the Vb1 and Vb2 analyses showed a dominant peak at the same CDR3 length distribution in more than half of the samples. Spectratypes dominated by a single CDR3 length were occasionally observed and in general did not correlate between cell populations and mice.
We next examined the clonal relationship between iTreg and ex-iTreg cells by comparing TCR sequences from mouse 4 and mouse 6. Vb8.2 cDNA fragments were selected for TCR CDR3 sequencing based on the broad distribution of CDR3 lengths with both Gaussian and skewed distributions seen in all analyzed mice (Figure 6A). CDR3 regions from each population were amplified by PCR, linked to beads and examined using non-optical integrated semiconductor sequencing. We obtained 416,207 in-frame reads encoding 5,162 iTreg and 3,738 ex-iTreg CDR3 polypeptides (Table S2). Repertoire comparisons based on amino acid sequences revealed minimal overlap (~2%) between unique iTreg and ex-iTreg CDR3 regions (Figure 6B). Consistent with this observation, the Morisita Horn Index (MHI) was 0.11 and 0.12 for the two comparisons, indicating little similarity. Ln-rank frequency versus ln-rank plots revealed a two-component distribution consisting of clones that follow a power law relationship and multiple high rank clonotypes. To determine if the same high rank clonotypes expanded after transfer, we compared the two iTreg and the two ex-iTreg repertoires that developed in treated mice that received aliquots from a common Treg cell pool (Figure 6C). There was little overlap between the iTreg populations (~1%, MHI=0.092). The ex-iTreg populations also showed a small 1.4% overlap, although the MHI for this comparison was higher (MHI=0.488). The higher MHI here indicates that a few of the high rank clonotypes were recovered from both mice. For control experiments, we compared the iTreg clonotypes recovered from the same samples run on two different semiconductor chips or by direct cloning and sequencing of all four populations (Figure 6D, Figure S2B, and Table S2). As expected, the TCR repertoires recovered by the different methods were highly similar (average MHI~0.8) and in some instances nearly identical (MHI=0.971). These data validate non-optical integrated semiconductor sequencing in this application and demonstrate that we can find similarity between repertoires where it exists. Collectively, these data indicate that each mouse expanded unique populations of iTreg and ex-iTreg cells, while the ex-iTreg populations also shared a few high rank clonotypes between the two mice. Within an individual mouse the iTreg and ex-iTreg populations were clonally unrelated.
Here, we modified a T cell transfer model of colitis to create chimeric mice where nTreg and iTreg cells were selectively deficient in IL-10. Although iTreg cells comprised a fraction (~20%) of all Treg cells recovered from treated mice, the IL-10 supplied by iTreg cells could replace nTreg cell-derived IL-10 in the cure of disease. In the reverse experiment, nTreg cell-derived IL-10 was equally effective. These results demonstrate the principle of reciprocal compensation between Treg subsets, which is an essential tolerogenic mechanism. Importantly, we identified iTreg cells as a particularly potent source of IL-10 in the gastrointestinal tract that can be utilized to augment regulatory function in situations where nTreg cells are defective or depleted. Reductions in Treg cell numbers and/or function have been implicated in many human autoimmune diseases (38). This therapeutic approach may therefore have broad application, as long-term stable tolerance can be achieved through iTreg cell transfers when nTreg cells are also present.
Prior work has shown that the iTreg cell transcriptional signature 72 hours after in vitro induction of Foxp3 with TGF-β was largely independent of Foxp3 expression and distinct from that of nTreg cells (6, 39). Here we extended these studies by determining the transcriptional profile of in vitro-derived iTreg cells that were maintained in treated mice for approximately three months. We now demonstrate that nTreg cells and in vitro-derived iTreg cells that are stable in vivo share similar transcriptional profiles, including the expression of many genes associated with Treg cell suppressive function in addition to Il10. These data provide further support for the hypothesis that the two Treg subsets utilize similar suppressive mechanisms. This result also illustrates that the in vitro-derived iTreg and nTreg transcriptional signatures converge as the iTreg cells are selected and maintained in vivo. One caveat to these studies is that we have compared in vivo-derived nTreg cells to in vitro-derived iTreg cells and not to in vivo-derived iTreg cells. In vitro-derived iTreg cells that persist in treated mice may not be equivalent to in vivo-derived iTreg cells under the same conditions. Further studies are needed to compare the function and stability of iTreg and nTreg cells derived in vivo. This caveat notwithstanding, we have identified important characteristics of stable in vitro-derived iTreg relevant to their use in immunotherapy.
Interestingly, ~85% of the surviving Thy1.1+ population no longer expressed Foxp3 three months after transfer. Although these ex-iTreg cells retained some elements of the canonical Treg transcriptional signature, the TCRβ CDR3 repertoires of iTreg and ex-iTreg cells were essentially non-overlapping. Thus the generation of a large population of ex-iTreg cells in vivo may be the unintended consequence of in vitro conversion based on non-specific TCR crosslinking. In the absence of Treg cell control, the prior history of Foxp3 expression was not sufficient to control the pathogenic potential of ex-iTreg cells upon re-transfer into Rag1–/– hosts. However, iTreg cell-derived IL-10 limited the frequency of ex-iTreg cells adopting a Th1, Th17 or Th1+Th17 cell phenotype. This is consistent with previous studies showing that IL-10 provided exogenously or by CD4+ Foxp3+ Treg cells, constrains Th17 and Th1+Th17 cell frequencies (23). Additionally, IL-10 signaling in Treg and Th17 cells limits Th17 cell-mediated inflammation (23, 40). Collectively, these data support a specific role for iTreg cell-produced IL-10 in the control of Th17 responses.
After re-transfer into Rag1–/– hosts, ~2% of the ex-iTreg cells reacquired Foxp3 expression. Thus ex-iTreg cells retain the potential to re-express Foxp3 and irrespective of their pathogenic potential, may serve as an iTreg cell reservoir in instances when iTreg responses are exhausted or diminished. Unstable Foxp3 expression in both nTreg and iTreg cells has been described, although others have concluded that Foxp3 expression in nTreg cells is heritable and stable (41–48). Our results clearly demonstrate that many in vitro-derived iTreg cells have unstable Foxp3 expression in vivo. It has been shown that iTreg cells derived in vitro are unstable due to a failure to fully demethylate the TSDR (29, 37). This incomplete demethylation could contribute to the high number of ex-iTreg cells observed in treated mice. It is interesting, however, that even mice with iTreg cells that had an extensively demethylated TSDR, still had a sizeable fraction of in vitro-derived iTreg cells that lost Foxp3 expression and became ex-iTreg cells. Furthermore, stable iTreg cells that never fully acquired the nTreg cell TSDR signature were still recovered from successfully treated mice, both in this experimental colits model and in a model of Foxp3 deficiency, suggesting that pathways other than promoter demethylation may impact the maintenance of iTreg cells in vivo (7). The amount of inter-mouse variability in the extent of iTreg cell TSDR demethylation further supports the notion that factors such as subclinical inflammation and disease status may contribute to iTreg cell stability. Since iTreg cells can be specific for gut bacterial antigens, strategies to derive polyclonal populations of antigen-specific iTreg cells prior to transfer may enhance the stability of Foxp3 expression (49). Treatment with inhibitors of DNA methyltransferases and histone deacetylases have been shown to enhance the stability of Foxp3 expression and could be incorporated into in vitro induction protocols (37, 50, 51).
When a pooled population of nTreg and iTreg cells was used to treat two mice, iTreg populations emerged that comprised a similar fraction of the Treg compartment but contained distinct TCR repertoires. This result supports two conclusions. First, a limited and fixed pool of in vitro-derived iTreg cells contains a surprisingly large number of clones with TCRs that can be maintained within the iTreg cell niche. In fact, it has been shown that high TCR diversity is important for the optimal function of Treg cells in a model of experimental acute graft versus host disease (52). Second, the iTreg cell niche is restricted, since selection of iTreg cells is likely to be stochastic and there was minimal overlap in the iTreg cell repertoires. This constraint is probably not due to the number of available antigens, given the complexity of the microbiome. Thus, other factors such as the number of tolerogenic APCs, local concentrations of TGF-β1 and IL-2, and signaling via the programmed death (PD) 1-PD- ligand (PD-L) pathway may determine the size of the iTreg cell population (53–57). In the absence of nTreg cells, the iTreg cell compartment expands ~5-fold indicating that nTreg cells, either directly or indirectly, also influence the size of the iTreg cell niche (6). Manipulation of these factors may provide mechanisms to expand the iTreg cell compartment thereby decreasing the propensity for the formation of pathogenic ex-iTreg cells while improving therapeutic outcome.
In conclusion, recent work has established that iTreg and nTreg cells are functionally non-redundant, based in part on differences in TCR repertoire and presumably TCR specificity (7, 49, 58). We now identify the capacity for reciprocal compensation between the two Treg subsets that relies on a mutual suppressive mechanism. Thus the answer to the recently posited question “Natural and adaptive Foxp3+ regulatory T cells: more of the same or a division of labor?” (2) is that both shared methods and unique characteristics are needed to enforce tolerance.
James Verbsky kindly provided the Foxp3EGFP IL10–/– mice. We thank Brandon Edwards and Kyle Upchurch for assistance with cell sorting. We also thank Mary Williams for administrative assistance and William Drobyski and Jack Gorski for critical review of the manuscript.
1This work was supported by the National Institutes of Health grants RO1 AI073731 and RO1 AI085090 (C.B.W. and T.A.C.), RO1 AI078713 (M.J.H.), a senior research award from the Crohn’s and Colitis Foundation of America (C.B.W.), the D.B. and Marjorie Reinhart Family Foundation (C.B.W.), and the Children’s Hospital of Wisconsin (C.B.W.).