Induced regulatory T cells suppress Tc1 cells through TGF-β signaling to ameliorate STZ-induced type

Type 1 diabetes mellitus (T1D) is a chronic autoimmune condition in which the immune system destroys insulin-producing pancreatic β cells. In addition to well-established pathogenic effector T cells, regulatory T cells (Tregs) have also been shown to be defective in T1D. Thus, an increasing number of therapeutic approaches are being developed to target Tregs. However, the role and mechanisms of TGF-β-induced Tregs (iTregs) in T1D remain poorly understood. Here, using a streptozotocin (STZ)-induced preclinical T1D mouse model, we found that iTregs could ameliorate the development of T1D and preserve β cell function. The preventive effect was associated with the inhibition of type 1 cytotoxic T (Tc1) cell function and rebalancing the Treg/Tc1 cell ratio in recipients. Furthermore, we showed that the underlying mechanisms were due to the TGF-β-mediated combinatorial actions of mTOR and TCF1. In addition to the preventive role, the therapeutic effects of iTregs on the established STZ-T1D and nonobese diabetic (NOD) mouse models were tested, which revealed improved β cell function. Our findings therefore provide key new insights into the basic mechanisms involved in the therapeutic role of iTregs in T1D.

T1D occurs when pancreatic β cells cannot produce enough insulin to maintain normal blood glucose levels, and children are the main group that develop T1D. It has been reported that the prevalence of T1D in children is doubling approximately every 20 years in Europe,1 and T1D currently causes an estimated loss of life expectancy of ~11–13 years.2 Unfortunately, no therapeutic approaches can prevent or cure T1D,3 and people with T1D need lifelong exogenous insulin therapy, which can only delay the progress of the disease. Thus, the development of new treatments that specifically target the pathogenesis of T1D and eventually prevent and cure T1D is urgently needed.

It is well recognized that β cell destruction is mainly mediated by autoimmunity. Signals generated by stressed, injured, or dying β cells recruit immune cells to the islets and activate these cells in the islets. Then, invading or resident macrophages and T cells release chemokines and cytokines in the islets and deliver proapoptotic signals to β cells, which ultimately destroy β cells.4 CD8+ T cells have been demonstrated to play a crucial role in this process. Papers have reported that in humans, CD8+ cytotoxic T cells are the most abundant population during insulitis5 and that islet-specific autoreactive CD8+ T cells infiltrate the islets of individuals with recent-onset or longstanding T1D.6 Tc1 cells (CD8-positive cells that secrete IFN-γ and TNF-α) mainly contribute to the cytotoxic activity of CD8+ T cells in T1D, as Tc1 cells are more frequent in T1D patients than in normoglycemic people.7 Furthermore, Tc1 cell adoptive transfer was shown to induce diabetes in nonirradiated adult rat insulin promoter-HA (RIP-HA) mice, while Tc2 cells (CD8-positive cells that secrete IL-4, IL-5, and IL-10) did not induce diabetes efficiently compared with Tc1 cells. Moreover, adoptive transfer of Tc1 cells from IFN-γ-deficient mice fails to induce diabetes in RIP-HA mice, suggesting that IFN-γ contributes to the pathogenic potential of cytotoxic Tc1 cells in T1D.8

Regulatory T cells (Tregs) are a class of suppressive T cells that can affect the function of CD8+T cells and are essential for inducing and maintaining immune tolerance. Tregs have deficiencies in frequency and/or function in T1D.9,10,11 Furthermore, boosting Treg numbers (either by adoptive transfer or in vivo induction) has been found to be protective in animal and human T1D,12,13,14 thus fueling interest in Tregs as a promising T1D immunotherapeutic target. Tregs are heterogeneous and can be divided into three populations: thymus-derived naturally occurring Tregs (nTregs), Tregs induced in vivo (pTreg), and Tregs induced ex vivo with IL-2 plus transforming growth factor-β (TGF-β) with or without retinoic acid or rapamycin (iTregs).15Among the three types of Tregs, nTregs are the most studied population in T1D treatment. Although work has been promising, there are several flaws that may significantly diminish the application of nTregs for T1D treatment, including (1) limited availability, (2) susceptibility to inflammation-induced apoptosis, (3) inability to suppress proinflammatory Th17 cells, and (4) self-conversion to effector T cells during inflammation.16,17,18 In contrast, we and others have shown that iTregs do not have the flaws of nTregs mentioned above19,20,21 and maintain their suppressive function similar to nTregs. Moreover, iTregs have a better therapeutic effect than nTregs in autoimmune diseases, such as airway allergic inflammation,18 rheumatoid arthritis,22,23 and experimental allergic encephalomyelitis.24 However, only a few studies have tested the role of iTregs in T1D,25,26,27 and the underlying mechanism is still unclear. Here, we investigated the preventive and therapeutic roles of iTregs in T1D and further explored the impact of iTregs on IFN-γ production by Tc1 cells and the underlying mechanisms using an STZ-induced T1D model.

STZ can specifically induce β cell damage by binding to the receptor GLUT2 on pancreatic β cells. In contrast to a single high-dose STZ injection, which induces necrosis of β cells without lymphocyte infiltration, injection of multiple low doses of STZ can mimic T cell-mediated pancreatic injury.28,29,30,31 Thus, we applied this well-established and broadly used STZ-T1D model32,33,34 to test the role of iTregs during the progression of diabetes. We found that iTregs significantly ameliorated the progression of T1D by inhibiting Tc1 cells and rebalancing Treg/Tc1 cells in recipients. Furthermore, we showed that the suppressive effect of iTregs on IFN-γ production by Tc1 cells was due to the TGF-β-mediated combinatorial actions of mTOR and TCF1. Finally, we applied the established STZ-T1D and nonobese diabetic (NOD) mouse models to further confirm the improved β cell function observed after iTreg treatment. Thus, our data indicate that iTregs have great potential as a cell-based therapy for T1D.

Mice

Wild-type C57BL/6 mice (male, 6–8 weeks old) were purchased from Guangdong Medical Laboratory Animal Center (Guangzhou, China). C57BL/6-FoxP3GFP mice were generously provided by Dr. Talil Chatilla (University of Southern California, Los Angeles). C57BL/6 Thy1.1 transgenic mice were purchased from The Jackson Laboratory (Stock No: 000406). C57BL/6-FoxP3GFP mice were backcrossed to B6 Thy1.1 mice, yielding Thy1.1+ C57BL/6-FoxP3GFP mice. Six-week-old NOD/ShiLtj mice were purchased from GemPharmatech (JiangSu, China), and ten-week-old NOD/ShiLtj mice were purchased from HuaFuKang Bio Company (Beijing, China). All animals were housed in the SPF animal facility in a temperature (72 ± 3 °F)- and air (50 ± 20% relative humidity)-controlled room with a 12-h light/dark cycle and were given a standard diet and tap water. All experiments using mice were performed in accordance with protocols approved by the Institutional Animal Care and Use Committee of Sun Yat-sen University.

Induction of diabetes

T1D was induced in Thy1.2+ C57BL/6 mice (male, 6–8 weeks old) using multiple low-dose STZ (Sigma-Aldrich, St. Louis, MO, USA) injections. STZ was administered via the intraperitoneal route at a dose of 50 mg/kg in fresh cold 0.01 M citrate buffer (pH 4.5) within 20 min of dissolution for 5 consecutive days. Mice were fasted for 4–6 h prior to STZ injection, and 10% sucrose water was supplied overnight after the first injection to avoid sudden hypoglycemia.

Cell transfers and diabetes measurement

Naïve CD4+ CD62L+ T cells were MACS-sorted from the spleen or lymph nodes (the inguinal lymph nodes, mesenteric lymph nodes and axillary lymph nodes). In brief, the spleen or lymph nodes of Thy1.1+ C57BL/6-FoxP3GFP mice were collected, and single-cell suspensions were obtained by passing through a 40-µm cell strainer (352340, Corning, USA). Red blood cell lysis buffer (Sigma-Aldrich, St. Louis, MO, USA) was used to lyse splenic erythrocytes, and T cells were enriched using nylon wool (18369-50, Polysciences, USA). Then, the T cells were labeled with biotinylated anti-CD8 (100704, BioLegend), anti-B220 (103204, BioLegend), anti-CD11b (101204, BioLegend), anti-CD11c (117304, BioLegend), and anti-CD49b (103522, BioLegend) antibodies and anti-biotin microbeads (130-090-485, Miltenyi, Biotec, Germany). The cells were then subjected to depletion followed by positive selection after labeling with CD62L microbeads (130-049-701, Miltenyi Biotec, Germany) by an automatic magnetic cell sorter (MACS) (Miltenyi, Biotec, Germany). The purity of naïve CD4+ CD62L+ T cells was determined with flow cytometry, and naïve CD4+ T cells with a purity >90% (please see Supplementary Fig. S1) were used for iTregs induction. iTregs were induced with recombinant human IL-2 (50 U/mL, R&D Systems) in the presence of recombinant human TGF-β (2 ng/mL, R&D Systems) or Med cells (CD4+ Foxp3- cells) with only recombinant human IL-2 for 3 days. To determine intervention effects, 3.0 × 106 iTregs or Med cells suspended in 200 µl phosphate-buffered saline (PBS) were injected via the tail vein on the first day of STZ administration. In addition, STZ-induced model mice receiving 200 µl PBS were considered an additional control. Nonfasting blood glucose levels were monitored twice a week over the following 30 days using an Embrace One Touch blood glucometer (OMNIS Health, USA). Blood glucose concentrations exceeding 300 mg/dL or 16.7 mmol/L in two consecutive daily measurements were considered to indicate diabetes onset. Cages were changed every 3–4 days since mice developed symptoms of polyuria. For some experiments, an ALK5 inhibitor (Sigma-Aldrich) was given through i.p. injection at a dose of 0.5 mg per mouse on days 0, 7, 14, 21, and 28 to explore the role of TGF-β signaling in iTregs-mediated protection from T1D.

To determine the role of iTregs in established T1D, we made use of two models: STZ-T1D and NOD mice. For the STZ-T1D model, diabetes was induced in male C57BL/6 mice, and iTregs (3.0 × 106) induced from C57BL/6 mice under the conditions mentioned above were transferred into diabetic mice via the tail vein on day 7. The nonfasting blood glucose concentration was measured every other day within one week after cell transfer and 1–2 times per week thereafter. For some mice, we transferred the same dose of iTregs a second time on day 26, and blood glucose levels were monitored thereafter. For NOD mice, 3.0 × 106 iTregs induced in vitro from naïve CD4+ T cells isolated from C57BL/6 mice were injected into 6-week-old and 10-week-old female mice via the tail vein. The nonfasting blood glucose concentration was measured 1–2 times per week. IPGTT and immunohistochemistry were performed to evaluate the function of β cells.

Histology and immunohistochemistry

Pancreas samples were fixed overnight in 10% formalin and embedded in paraffin. All embedding, sectioning, and H&E staining were performed by Servicebio Company (Wuhan, China). For assessment of insulin and glucagon production in the pancreas, two consecutive sections were stained with anti-insulin (1:1600, Abcam 63820) or anti-glucagon (1:8000, Abcam ab92517) antibodies, and the staining was visualized with anti-rabbit DAB-HRP (OMap) secondary antibodies; nuclei were counterstained with hematoxylin and a bluing reagent. Area measurement was performed with ImageJ software. For morphometric analysis of islet size and β cell area (positive insulin staining), each islet was traced and analyzed manually with ImageJ software. One section per pancreas in the preventive experiment (iTregs given at the same time of the STZ injection) and three sections per pancreas in the therapeutic experiment (iTregs given after diabetes onset) were used, and all islets seen on the sections were measured for analysis. Sections were evaluated by at least two different people to avoid subject bias. In brief, islets seen in pancreatic sections imaged at 200× magnification were captured with an Olympus microscope (Tokyo, Japan). For islet area measurement, the “Freehand selections” tool was used to draw the outline of islets in images with scale bars, and then the “measure” tool was used for area measurement. The size of islets is reported in the Results section and expressed as µm2. To quantify β cell area in relation to islet area, images were first converted to RGB stacks, and then the islet outline was drawn, followed by automatic thresholding. β cell area in relation to islet area could be read after using the “measure” tool.

Assessment of insulitis

Pancreatic tissue samples were stained with hematoxylin and eosin (H&E). The degree of insulitis was determined from three nonsequential slides from 3 to 5 individual mice. Every islet on each section was scored as previously described35: 0 = no lymphocytic infiltration; 1 = peri-insulitis (<30%); 2 = <50% islet infiltration; and 3 = >50% islet infiltration.

Enzyme-linked immunosorbent assay (ELISA)

Blood was collected from each mouse on day 15 after STZ injection and clotted at room temperature for 1 h, followed by incubation at 4 °C overnight. The serum was frozen at −80 °C until analysis. Serum insulin and C-peptide levels were measured by ELISA using a mouse C-peptide and insulin ELISA kit (Nanjing Herb Source Bio-Technology Co., Ltd, China) according to the manufacturer’s instructions.

Intraperitoneal glucose tolerance test (IPGTT)

Glucose tolerance was evaluated using a conventional IPGTT conducted 15 days after STZ administration. Mice were given 20% glucose via i.p. injection at a dose of 2.0 g/kg body weight. Blood samples were collected at 0, 15, 30, 60, and 120 min via tail vein bleeding. Blood glucose levels were measured using a glucometer (OMNIS Health, USA).

Murine naïve CD8+ T cell differentiation in vitro

Naïve CD8+ CD62L+ T cells were purified from the spleen or lymph nodes of Thy1.2+ C57BL/6 mice via magnetic beads isolation (Miltenyi, Biotec, Germany) in the same way that naïve CD4+CD62L+ T cells were isolated. iTregs induced in vitro were cocultured with naïve CD8+CD62L+ T cells (1:1) during in vitro differentiation. Naïve CD8+ T cells were stimulated with anti-CD3 (1 µg/ml) and anti-CD28 (1 µg/ml) antibodies (both from BioLegend) in the presence of irradiated (30 cGy) syngeneic non–T cells (1:1) and recombinant mouse IL-12 (20 ng/ml, R&D Systems). After 3 days in culture, differentiated cells were restimulated with PMA and ionomycin (Sigma-Aldrich) for 5 h, with brefeldin A (BFA, BioLegend) added for the last 4 h. The expression of IFN-γ was then measured by flow cytometry. To determine the possible mechanisms, an ALK5 inhibitor (10 µM) was added to the coculture systems, and the cell differentiation ratio, mTOR activation and TCF1 expression were determined.

Flow cytometry analysis

Antibodies against CD4 (GK1.5, PerCP/Cy5.5), CD8 (53–6.7, PerCP/Cy5.5), Thy1.1 (OX-7, PE), Foxp3 (MF-14, Alexa Fluor 647), IFN-γ (XMG1.2, APC), IL-17 (TC11-18H10.1, PE), and Neuropilin-1 (3E12, PE) were purchased from BioLegend. An antibody against Foxp3 (FJK-16s, PE-Cyanine7) was purchased from Invitrogen. Antibodies specific for phospho-S6 ribosomal protein (Ser240/244) (D68F8, Alexa Fluor 647) and TCF1 (C63D9, Alexa Fluor 647) were purchased from Cell Signaling Technology. An Annexin V-FITC apoptosis detection kit (BMS500FI-300, Invitrogen) was used to evaluate cell apoptosis. Results were obtained on a BD FACSCalibur or BD LSRFortessa flow cytometer and analyzed using FlowJo. For ex vivo IFN-γ detection, lymphocytes were isolated from the spleen, mesenteric and inguinal lymph nodes (LNs) and pancreatic lymph nodes (PLNs) of diabetic mice on day 15 and day 30 after STZ injection and then stimulated in vitro with PMA (50 ng/ml) and ionomycin (500 ng/ml) for 5 h, with BFA added for the last 4 h. The intracellular expression of IFN-γ in CD8+ T cells was analyzed by flow cytometry.

Quantitative polymerase chain reaction

Total RNA was isolated from the mesenteric lymph nodes and inguinal lymph nodes using the RNeasy Mini Kit (Qiagen, Valencia, CA, USA) according to the manufacturer’s instructions; 1 μg RNA (OD260 nm/OD280 nm  = 1.8–2.2) was used to synthesize first-strand cDNA using the Prime-Script RT Reagent Kit (Takara Bio Inc., Shiga, Japan). Quantitative polymerase chain reaction (qPCR) was performed on a 7500 Fast (ABI) RT-PCR instrument using SYBR Premix Ex Taq (Takara Bio Inc., Shiga, Japan). For qPCR, the following program was performed: 95 °C for 30 s (predenaturation); 95 °C for 5 s and 60 °C for 34 s for 40 cycles (amplification); 95 °C for 15 s, 60 °C for 1 min and 95 °C for 15 s (melt curve). The 2^ (-delta CT) method was used for calculations, and glyceraldehyde-3-phosphate dehydrogenase (GAPDH) was used as the housekeeping gene for normalization.

Western blots

Protein isolation from the mesenteric lymph nodes and inguinal lymph nodes was performed as previously described.36 Western blotting was performed using the standard SDS-polyacrylamide gel electrophoresis method and enhanced chemiluminescence detection reagents (GE Healthcare Biosciences AB, Uppsala, Sweden). Antibodies against S6 ribosomal protein (S6), phospho-S6 ribosomal protein (Ser240/244) (pS6), TCF1 and GAPDH (all from Cell Signaling Technology, Beverly, MA) were used. Immunoreactivity was semiquantitatively measured by gel densitometric scanning and analyzed with an MCID image analysis system (Imaging Research, Inc.).

Statistical analysis

All results are presented as the mean ± standard deviation (s.d.). Differences among multiple groups were analyzed using one- or two-way analysis of variance (ANOVA). When ANOVA showed significant differences, pairwise comparisons between groups were performed with Dunnett’s test. A two-tailed unpaired t test or the Mann-Whitney test was used for two-group comparisons. 1The chi-square test was used to compare diabetes incidence between groups. In all analyses, P < 0.05 was considered statistically significant.

iTregs ameliorated the progression of T1D and improved β cell function

To generate a T1D preclinical animal model, C57BL/6 mice were injected with multiple low doses of streptozotocin (STZ), a widely accepted model (Fig. 1A). Consistent with the data reported in the literature, STZ-injected mice exhibited progressive hyperglycemia 9 days after the first administration (Fig. 1B). To test the therapeutic potential of iTregs in T1D, C57BL/6 mice were adoptively transferred with 3 × 106 iTregs during the initial STZ injection. Notably, the mice infused with iTregs exhibited significantly decreased blood glucose levels compared with those in the STZ-treated only (Model) and CD4+ Foxp3- cell infusion (Med) groups (Fig. 1B). Disease onset was markedly postponed following iTregs administration, and diabetes incidence was significantly decreased in the iTregs group compared to the control groups (Fig. 1C). To determine whether the improved glycemic control was associated with changes in β cells, we performed an intraperitoneal glucose tolerance test (IPGTT) on STZ-treated mice 15 days after iTreg administration. As shown in Fig. 1D, the blood glucose levels of mice from all three groups reached a peak 15 min after intraperitoneal glucose injection. However, the iTregs-treated mice exhibited dramatically faster clearance of the glucose load from the body than those in the Model and Med groups; the differences reached significance at 120 min (Fig. 1D). The area under the curve (AUC) results derived from the IPGTT also proved that iTregs treatment could preserve β cell function (Fig. 1E). To further assess β cell function, serum insulin and C-peptide levels were tested on day 15. Consistent with the previous results, iTregs-treated mice showed increased insulin secretion and C-peptide production (Fig. 1F, G). To directly reveal the protective role of iTregs, histochemical analyses were performed to show the conservation of insulin-producing β cells (Fig. 1H). Taking all the islets as individual samples, iTregs-treated mice maintained a larger islet size (Fig. 1I) and exhibited significantly higher conservation of insulin-producing β cells per islet than those in the other two groups (Fig. 1J). If the average islet size or β cell area in relation to the islet area for each sample was used for analysis, we could see that the mice in the iTregs group still showed a larger islet size (Supplementary Fig. S2A) and more insulin-producing β cells per islet (Supplementary Fig. S2B). Taken together, these results demonstrate that iTregs can ameliorate the progression of T1D and protect β cell function in the STZ-induced T1D mouse model.

Adoptively transferred iTregs migrate to the pancreas and immune organs and maintain their phenotypic features in vivo

iTregs migration to diseased organs would be of great significance for therapeutic efficacy. To investigate the fate of adoptively transferred iTregs in vivo, iTregs derived from Thy1.1+Foxp3GFP reporter mice were transferred into Thy1.2+ STZ-induced T1D mice, and the distribution of iTregs in different organs was examined by flow cytometry on days 15 and 30 postinjection. The results showed that injected iTregs (CD4+ Foxp3-GFP+) could be found in the spleen, lymph nodes (mesenteric lymph nodes and inguinal lymph nodes) and, importantly, PLNs on both day 15 and day 30 (Fig. 2A). Although the frequencies in the spleen and lymph nodes were decreased on day 30 regardless of whether gating on total live cells or CD4+ T cells was used (Fig. 2B and C), the numbers of iTregs in these immune tissues remained stable (Fig. 2D). The frequency of iTregs among live PBMCs was almost the same on day 30 compared with day 15 but decreased when gated on total CD4+ T cells (Fig. 2E–G). Most importantly and strikingly, iTregs migrated to the pancreas and persisted for at least 30 days (Fig. 2H–J), indicating a strong ability to home to the target organ. To further investigate whether transferred iTregs could maintain their phenotypic features in vivo, we first examined the expression of Foxp3, the master transcription factor of regulatory T cells, which showed no changes between before (Supplementary Fig. S3) and after adoptive transfer (Fig. 2K, L). We next determined whether iTregs could convert into Th1 or Th17 cells under inflammatory conditions. Lymphocytes from the spleen, mesenteric lymph nodes, inguinal lymph nodes and pancreatic lymph nodes were harvested on day 30 postinjection and stimulated with PMA and ionomycin in vitro. IFN-γ and IL-17A production in iTregs was detected by intracellular staining. As shown in Fig. 2M, very few iTregs produced IFN-γ or IL-17A up to 30 days postinjection, whereas Thy1.1+ CD4+ Foxp3− cells from the Med group produced the above cytokines, indicating that iTregs did not convert into Th1 or Th17 cells in vivo.

Adoptively transferred iTregs migrate to the pancreas and immune organs and maintain their phenotypic features in vivo. iTregs (CD4+ Foxp3+) from Thy1.1+ FoxP3GFP mice were injected into Thy1.2+ T1D mice, and the iTregs organ distribution, engraftment and stability were then examined on day 15 and day 30. A Representative flow cytometry data showing the distributions of iTregs (CD4+GFP+ cells) on day 15 (upper) and day 30 (lower) in the spleen (SP), mesenteric and inguinal lymph nodes (LNs) and pancreatic lymph nodes (PLNs). Gated on total live cells. Analysis of the iTregs cell percentage in the total live cells (n = 5) (B) and in relation to the CD4+ T cells (n = 5) (C) in the SP, LNs and PLNs. Two-tailed unpaired t-test; data are shown as the mean ± s.d. D Analysis of iTregs cell numbers (n = 7) in the SP, LNs and PLNs. Two-tailed unpaired t-test; data are shown as the mean ± s.d. Representative flow cytometry data (E) and statistical data (F) showing the iTregs distribution in PBMCs gated on total live cells (n = 5). Two-tailed unpaired t-test; data are shown as the mean ± s.d. GAnalysis of the iTregs cell percentage in relation to CD4+ T cells (n = 5) in PBMCs. Two-tailed unpaired t-test; data are shown as the mean ± s.d. Representative flow cytometry data (H) and statistical data (I) showing the iTregs distribution in the pancreas gated on total live cells (n = 5). Two-tailed unpaired t-test; data are shown as the mean ± s.d. J Analysis of the iTregs cell percentage in the CD4+ T cell population (n = 5) in the pancreas. Two-tailed unpaired t-test; data are shown as the mean ± s.d. Representative flow data (K) and statistical data (L) showing Foxp3 expression on day 15 and day 30 in cells in different tissues gated on CD4+ Thy1.1+ cells (n = 3). Two-way ANOVA; data are shown as the mean ± s.d. M Representative flow cytometry data showing the conversion of iTregs or Med cells into Th1 and Th17 cells on day 30; gated on CD4+ Thy1.1+ cells. Experiments were repeated three times. *P < 0.05, **P < 0.01, and ***P < 0.001; ns indicates no significance

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iTregs rebalance endogenous IFN-γ-producing CD8 T cells and regulatory T cells in recipient STZ-induced T1D mice

To investigate the mechanism through which iTregs ameliorate the progression of T1D, we detected endogenous IFN-γ- and IL-17A-producing T cells and Tregs in recipient mice, as multiple lines of evidence show that the balance between effector T cells and Tregs is critical in T1D. The production of IFN-γ and IL-17A by endogenous CD4+ or CD8+ T cells (Thy1.2+) from the spleen and lymph nodes (mesenteric, inguinal and pancreatic) of STZ-induced T1D mice was determined by intracellular staining after iTregs or control Med cells injection. Among the three groups (Model, Med, and iTregs), no differences were seen in the frequencies of IFN-γ- or IL-17A-producing CD4+ T cells or IL-17A-producing CD8+ T cells in the LNs (Supplementary Fig. S4), spleen, or PLNs (data not shown). However, the frequency of IFN-γ-producing CD8+ T cells was significantly decreased in the iTregs group on both day 15 and day 30, indicating that iTregs might suppress the progression of T1D by inhibiting the IFN-γ-producing ability of CD8+ T cells (Fig. 3A–C).

Next, we examined the effect of iTregs on endogenous Tregs, the frequency of which is decreased during T1D. As shown in Fig. 3D, E, the frequencies of endogenous Tregs (Thy1.2+) in the spleen and lymph nodes (mesenteric, inguinal, and pancreatic) were significantly increased in iTregs-treated mice on day 15 compared to those in the Model and Med control groups, indicating that iTregs can improve the homeostasis of regulatory T cells. Neuropilin-1 (Nrp-1) can be used as a specific cell marker to distinguish nTregs from iTregs,37 so we costained for Nrp-1 and Foxp3 to determine whether the increased population of endogenous Tregs was primarily generated in the thymus (tTreg) or induced at peripheral sites (pTreg). As shown in Fig. S5, Nrp-1 expression on recipient Tregs was almost the same among the three groups, suggesting that iTregs mainly expanded endogenous nTregs, thus increasing the Tregs percentage in T1D mice. To confirm that iTregs can regulate the balance between endogenous regulatory T cells and IFN-γ-producing CD8+ T cells (Tc1 cells), the Treg/Tc1 cell ratio was calculated. Although the frequencies of endogenous Tregs on day 30 were not elevated in iTreg-treated mice (Fig. 3F), the Treg/Tc1 cell ratio was significantly increased on both day 15 and day 30 in the iTregs group compared to the control groups (Fig. 3G, H). Thus, iTregs can rebalance Tregs and Tc1 cells in the T1D mouse model.

iTregs suppress Tc1 cells through TGF-β-dependent inhibition of mTOR signaling and promotion of TCF1 expression in vitro

To investigate the basis of the inhibition of IFN-γ-producing CD8+ T cells by iTregs, we first examined the proliferative ability of CD8+ T cells following iTregs administration by staining for Ki-67 and found that iTregs did not alter the proliferation of CD8+ T cells (Fig. 4A, B). In addition, we did not observe different amounts of annexin V staining among the three groups, indicating that iTregs did not affect the apoptosis of CD8+ T cells either (Fig. 4C, D). Then, we analyzed whether iTregs can inhibit Tc1 cell differentiation. As shown in Fig. 4E, iTregs significantly suppressed IFN-γ production during the in vitro polarization of Tc1 cells. Since the TGF-β signaling pathway has been suggested to modulate iTregs-mediated suppressive function,38,39 we then examined whether TGF-β itself can also suppress Tc1 cell polarization and whether interfering with the TGF-β signaling pathway fails to inhibit IFN-γ production by CD8+ T cells. Strikingly, TGF-β alone inhibited Tc1 cell differentiation, and moreover, the suppression of IFN-γ production by iTregs was completely abrogated when an inhibitor of activin-like kinase 5 (ALK5, also known as type 1 TGF-β receptor) was added to the in vitro coculture system (Fig. 4E, F), suggesting that the suppressive function of iTregs is dependent on TGF-β signaling.

Previous studies have shown that mTOR activity is required for CD8+ T cells to maintain effector cell differentiation and function.40,41,42 Thus, to determine whether iTregs suppress IFN-γ production by inhibiting mTOR signaling, we examined the phosphorylation of the S6 protein, an indicator of mTOR activation, in the Tc1 cell differentiation system with or without iTregs. We found that iTregs treatment led to a significant inhibition of S6 phosphorylation and this inhibition was partially dependent on TGF-β signaling, as the ALK5 inhibitor partially rescued the suppression of S6 phosphorylation (Fig. 4G, H).

Since T cell factor 1 (TCF1) is also involved in Tc1 cell differentiation, as inflammatory signals can inhibit TCF1 expression during CD8+ T cell polarization and primed CD8+ T cells lacking Tcf1 undergo default effector differentiation,43,44,45 we evaluated TCF1 protein levels by flow cytometry to determine whether TCF1 is involved in the iTreg-mediated suppression of IFN-γ production in CD8+ T cells. We found that TCF1 expression was indeed decreased during effector CD8+ T cell differentiation, while iTregs treatment maintained high expression of the TCF1 protein, and this effect also partially relied on TGF-β (Fig. 4I, J). These results collectively suggest that the TGF-β-dependent inhibition of mTOR signaling and promotion of TCF1 are critical for the suppression of Tc1 cells by iTregs.

The therapeutic role of iTregs in STZ-induced T1D mice is dependent on TGF-β signaling

To extend our finding that TGF-β signaling is critical to the therapeutic role of iTregs in T1D, we investigated the effect of an ALK5 inhibitor on iTregs treatment. As described in Fig. 5A, the ALK5 inhibitor was injected five times at 1-week intervals in the context of the iTregs treatment protocol, as shown in Fig. 1A. Consistent with the in vitro results, disrupting TGF-β signaling with the ALK5 inhibitor abrogated the protective role of iTregs in STZ-induced T1D development, as suggested by elevated blood glucose levels (Fig. 5B) and an increased diabetes incidence (Fig. 5C). Moreover, the iTregs group showed a tendency toward amelioration of insulitis compared with the Model group, and the mice in the TGF-β signaling blockade group had more severe insulitis than those in the iTregs group (Fig. 5D-F). Decreases in islet size and β cell area in relation to islet area (Fig. 5D, G, H and Supplementary Fig. S6) further proved that the protective role of iTregs in STZ-T1D was dependent on TGF-β signaling. We further investigated whether mTOR signaling and TCF1 were involved in the effects of iTregs treatment, as suggested in vitro, and found that consistent with our in vitro data, mTOR mRNA and S6 phosphorylation levels were significantly suppressed, while TCF1 mRNA and protein expression was promoted and that inhibition of TGF-β signaling reversed these phenotypes (Fig. 5I–N). Taken together, these results suggested that the TGF-β-dependent inhibition of mTOR signaling and promotion of TCF1 were critical for the therapeutic effect of iTregs on the STZ-induced T1D mouse model.

iTregs improved β cell function in mice with established T1D

In addition to assessing the preventive role, we further evaluated the therapeutic effect of iTregs on established T1D using both STZ-T1D and NOD mice. A majority of mice developed diabetes on day 7 after the first STZ injection. These mice were randomly assigned to the iTregs-treated group or Model group. The working model is shown in Fig. 6A. As shown in Fig. 6B and C, six and two out of the ten mice in the iTregs-treated group had a nonfasting blood glucose level lower than 16.7 mmol/L 2 days and 4 days after cell intervention, respectively, while the mice in the Model group showed no relief of diabetes. Blood glucose levels rebounded to a level similar to that of the Model group 6 days after iTregs injection. The results of an IPGTT performed 1 week after cell transfer are shown in Fig. 6D. Although the blood glucose concentrations of all the mice treated with iTregs rebounded after 6 days, three mice still showed more powerful β cell function than the mice in the Model group, which contributed to the lower blood glucose level observed at 120 min during the IPGTT. The pancreas was isolated from these three mice for analysis one day after the IPGTT experiment to evaluate islet size and β cell function. All islets seen in three sections from each pancreas were evaluated for insulitis, islet area and β cell area in relation to islet area. Insulitis was scored from 0 to 3 based on the severity of infiltration. As shown in Fig. 6E–G, the severity of insulitis was markedly reduced 1 week after iTregs transfer. Taking all the islets as individual samples, iTregs-treated mice retained a significantly larger islet size (Fig. 6E and H) and had more insulin-producing β cells per islet than mice in the Model group (Fig. 6E and I). If the average islet size or β cell area in relation to islet area for each sample was used for analysis, we found that the mice in the iTregs group still had a larger islet size (Supplementary Fig. S7A), while the results for the β cell area in relation to the islet area did not reach statistical significance (Supplementary Fig. S7B). As the protective role for iTregs did not seem to be sustained for a long time, we reinjected 3 × 106 iTregs into the remaining mice 26 days after STZ injection. The blood glucose level did not decrease until 29 days after the second iTregs transfer, when one mouse out of seven showed a blood glucose level lower than 16.7 mmol/L (Fig. 6B).

We also evaluated the protective role of iTregs in NOD mice, which spontaneously develop T1D as a result of autoimmune-mediated destruction of pancreatic β cells. Although the difference in the blood glucose level between the two groups did not reach statistical significance (Supplementary Fig. S8A), iTregs remarkably slowed disease onset and decreased diabetes incidence (Fig. 6J). An IPGTT was also conducted to compare β cell function between the two groups 40 days after cell transfer. The differences in blood glucose levels at different time points during the IPGTT experiment and the AUC of the IPGTT did not reach statistical significance (Supplementary Fig. S8B, C). Insulin staining revealed that mice treated with iTregs retained a remarkably larger islet area and more insulin-producing pancreatic β cells (Fig. 6K–M and Supplementary Fig. S8D, E). In summary, these results indicated that iTregs could improve β cell function to a certain degree when transferred in the context of established T1D.

T1D patients have been shown to have defects in the frequency and/or function of Treg cells,9,10,11 which are protective in animal and human T1D.12,13,14 To date, the role of a more stable Treg population, iTregs, in T1D has seldom been studied. In this paper, we found that iTregs were stable, which is consistent with the findings in other papers,15,19 as they maintained high levels of Foxp3 expression while seldom expressing proinflammatory cytokines, such as IL-17A and IFN-γ. These iTregs induced in vitro were protective in STZ-induced T1D. They could delay T1D onset and ameliorate disease severity, as shown by reduced blood glucose levels, decreased islet damage, and improved β cell morphology and function.

The T1D model established with multiple low-dose STZ injections is an extensively used T1D model. Regardless of a direct toxic effect, a subsequent autoimmune reaction has been demonstrated to lead to β cell apoptosis, which then induces infiltration of T cells into the islets, suggesting a role for immune imbalance in this model.46 Therefore, it is not surprising that iTregs are functional in STZ-T1D since iTregs have a strong immunosuppressive function.

Although some previous studies have reported the therapeutic potential of iTregs in T1D,25,26,27 the mechanism has not been thoroughly investigated, except for one paper that mentioned that iTregs might work by inhibiting Th1 cells via FasL-dependent cytotoxicity.25 As T1D is widely recognized as an autoimmune disease and iTregs mainly function through immunosuppression, we further elucidated the mechanisms by which iTregs protect against T1D. Among proinflammatory T cell subsets, Tc1 cells are the population primarily suppressed by iTregs. As cytotoxic Tc1 cells accumulate with disease progression in prediabetic mice and patients newly diagnosed with type 1 diabetes, we concluded that prompt subversion of CD8+T cell effector function underlies the efficacy of iTregs therapy in preserving the function of islet β cells. The possible mechanisms could be attributed to decreased Tc1 cell proliferation, increased Tc1 cell apoptosis, reduced Tc1 cell infiltration and/or inhibited Tc1 cell differentiation. Our results showed that iTregs did not affect the proliferation or apoptosis of CD8+ T cells in the spleen or lymph nodes in vivo but significantly suppressed Tc1 cell polarization in vitro. The elevated expression of CCR5, whose ligands are CCL3, CCL4, and CCL5, on Tc1 cells contributes to the preferential recruitment of Tc1 cells in target tissues.8,47Previous reports by Morlacchi et al. and others have demonstrated that Tregs can target dendritic cell-mediated proinflammatory chemokine (CCL3 and CCL4) production, thus mediating a decrease in the recruitment of target T cells.48,49 Whether iTregs have the ability to decrease the migration of Tc1 cells to target tissues in T1D mice remains elusive and is worthy of further study in the near future.

Transforming growth factor-β is an important pleiotropic cytokine with potent immunoregulatory properties that has been reported to be crucial for iTregs-mediated suppressive function.38,39 TGF-β can inhibit IFN-γ expression during Th1 cell development.50Moreover, a paper reported that TGF-β signals were critical for CD4+ CD25+ Treg cell-mediated regulation of autoreactive islet-specific cytotoxic T lymphocytes in a CD8+ T cell-mediated model of T1D.51 Unsurprisingly, we proved that the regulatory effect of iTregs on Tc1 cells was also dependent on TGF-β since blocking TGF-β signaling using an ALK5 inhibitor completely eliminated the suppression of IFN-γ production in CD8+ T cells in vitro.

Follicular regulatory T cells can significantly suppress the activation of CD8+ T cells in a manner dependent on IL-10 and probably TGF-β.52 IL-10 has also been reported to be important for iTregs’ immunomodulatory function.38 However, in our study, we found that IL-10 signaling was dispensable for the suppressive function of iTregs during Tc1 cell polarization in vitro, since IFN-γ expression was not significantly changed when we added serial doses of an anti-IL-10 antibody to the culture system (data not shown). This difference could be explained by the different cell culture systems and different index parameters used.

Since studies have shown a requirement for mTOR activity in the maintenance of effector CD8+ T cell differentiation and mTOR was reported to be a critical target of TGF-β in mouse and human NK cells leading to arrest of NK cell development and cytotoxic activity,53,54 it is reasonable to conclude that iTregs suppress Tc1 cell differentiation through TGF-β-mediated mTOR inactivation. TCF1 deficiency resulted in increased IFN-γ production in CD4+ T cells, while overexpression of TCF1 led to suppression of IFN-γ production.55 Recent studies have shown that TCF1 suppresses effector CD8+ T cell differentiation and function directly or indirectly via epigenetic modification of the IFN-γ enhancer and that the Tcf1 dose and protein stability are critical in suppressing IFN-γ production.56 All of these papers support our findings that iTregs might inhibit Tc1 cells by maintaining high expression of the TCF1 protein. Our results showed that TGF-β was also required for this effect. One possible mechanism underlying the sustained expression of TCF1 mediated by TGF-β might be inhibiting the expression of the transcription factor signal transducer and activator of transcription (STAT) 4, since the inflammatory cytokine IL-12 used in the Tc1 cell polarization system can induce TCF1 downregulation via STAT4,45 and TGF-β was reported to inhibit Th1 cell differentiation specifically through downregulation of STAT-4 expression.57 Nevertheless, further experiments, especially with transgenic mice, are needed to provide direct evidence for how TGF-β inhibits mTOR activation and maintains TCF1 expression and define the relationship between mTOR and TCF1 signaling in Tc1 cells during diabetes.

Since iTregs showed such powerful protection against diabetes onset, we evaluated their role in established T1D. As the data revealed, iTregs decreased the blood glucose level and improved β cell function soon after cell transfer, but these effects lasted for only several days. iTregs mainly function through immune suppression, and after cell transfer, iTregs could significantly inhibit the autoimmune response in diabetic mice, which led to the decreased blood glucose levels and improved β cell function. However, the autoimmune response in early-onset diabetic mice might be so strong that the iTregs failed to persistently suppress it, which resulted in the blood glucose level rebounding. Increasing the iTregs cell number and frequency in products for cell transfer might help to solve this problem, which could be further tested. Surprisingly, one mouse achieved a normal blood glucose level approximately two months after iTregs injection. One explanation might be that iTregs have the ability to repair damaged tissue in addition to performing their immunosuppressive function. Previous papers have reported that Tregs can promote tissue repair through direct interactions with tissue progenitor cells or through interactions with supportive cells to create an appropriate stem cell niche for tissue repair,58 and amphiregulin,59 keratinocyte growth factor,60 CCN3,61 VEGFA,62Jag1,63 and IL-1064 play important roles in the process of tissue repair. In the near future, it would be of interest to determine whether iTregs have a regenerative role in T1D mice or other disease models.

In addition to the STZ-T1D model, NOD mice, which spontaneously develop diabetes, were also used to explore the therapeutic role of iTregs. Due to the limitation of the small sample size, we could see only a potential decrease in blood glucose levels and delay of diabetes onset. Strikingly, iTregs indeed maintained larger islet sizes and more functional β cells, which motivated us to optimize the dose and frequency of the cell intervention and use a larger number of NOD mice in the future to further clarify the therapeutic role of iTregs.

In conclusion, the current work demonstrated the therapeutic role of iTregs in autoimmune responses and β cell protection in the STZ-T1D model. The underlying mechanisms might be associated with a TGF-β-dependent reduction in IFN-γ expression in CD8+ T cells. At the molecular level, the repression of mTOR activation and maintenance of high expression of the TCF1 protein by TGF-β could both be involved in the iTregs-mediated suppression of Tc1 cells (Fig. 7). Our work indicates a potential novel therapeutic strategy for delaying the occurrence and development of diabetes.

Model of iTregs migration, stability and function in STZ-T1D mice. iTregs migrated to PBMCs, the pancreas, the spleen and the lymph nodes. iTregs survived for at least 30 days in STZ-T1D mice and were stable in vivo, as indicated by the maintained expression of Foxp3 and resistance to secretion of IL-17 and IFN-γ. iTregs could protect β cell function, possibly by upregulating Treg levels in recipient mice while inhibiting type 1 cytotoxic T cell (Tc1) function, thus rebalancing the Treg/Tc1 cell ratio in recipients. The effect of iTregs on Tc1 cells was dependent on TGF-β-mediated combinational action on mTOR and TCF1 signaling

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