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Due to their immunomodulatory properties, mesenchymal stem cells (MSC) are interesting candidates for cellular therapy for autoimmune disorders, graft-versus-host disease and allograft rejection. MSC inhibit the proliferation of effector T cells and induce T cells with a regulatory phenotype. So far it is unknown whether human MSC-induced CD4+CD25+CD127–forkhead box P3 (FoxP3)+ T cells are functional and whether they originate from effector T cells or represent expanded natural regulatory T cells (nTreg). Perirenal adipose-tissue derived MSC (ASC) obtained from kidney donors induced a 2·1-fold increase in the percentage of CD25+CD127–FoxP3+ cells within the CD4+ T cell population from allostimulated CD25–/dim cells. Interleukin (IL)-2 receptor blocking prevented this induction. The ASC-induced T cells (iTreg) inhibited effector cell proliferation as effectively as nTreg. The vast majority of cells within the iTreg fraction had a methylated FOXP3 gene Treg-specific demethylated region (TSDR) indicating that they were not of nTreg origin. In conclusion, ASC induce Treg from effector T cells. These iTreg have immunosuppressive capacities comparable to those of nTreg. Their induction is IL-2 pathway-dependent. The dual effect of MSC of inhibiting immune cell proliferation while generating de-novo immunosuppressive cells emphasizes their potential as cellular immunotherapeutic agent.
Mesenchymal stem cells (MSC) can be isolated from an abundance of human tissue sites, including adipose tissue and bone marrow, and their expansion is accomplished easily [1-4]. MSC possess immunosuppressive capacities and, as a consequence, over the past decennium MSC have been studied extensively as a prospective cellular therapeutic agent to prevent or treat autoimmune diseases, graft-versus-host disease (GVHD) and allograft rejection [5-12]. Upon activation, MSC prevent the proliferation of various immune cells, in particular T cell proliferation [13-19]. MSC mediate their suppressive effect through cell–cell contact and the secretion of various soluble factors such as transforming growth factor (TGF)-β, hepatocyte growth factor (HGF), interleukin (IL)-10, nitric oxide (NO), human leucocyte antigen G5 (HLA-G5), indoleamine 2,3-dioxygenase (IDO) and prostaglandins [20-28]. While MSC strongly inhibit T cell proliferation via these mechanisms, they preserve the function of CD4+CD25+CD127–forkhead box P3 (FoxP3)+ regulatory T cells (Treg) . Beyond this, in-vitro studies and studies in animal models have indicated that MSC have the capacity to generate Treg [9, 30-34]. Recent evidence was provided that this phenomenon might also occur in renal transplant patients undergoing MSC therapy . Intravenous administration of autologous MSC post-transplant led to a proportional increase of CD4+CD25+CD127–FoxP3+ T cells. Despite their regulatory phenotype, it remains essential to investigate the characteristics and function of these cells. Further, due to the heterogeneity of the Treg population these CD4+CD25+CD127–FoxP3+ T cells could represent expanded natural Treg (nTreg) or newly induced Treg (iTreg).
nTreg and iTreg are distinct from each other with regard to their place of origin, the stability of their transcription factor FOXP3 expression and in their methylation pattern of the Treg-specific demethylated region (TSDR) in the FOXP3 gene [36, 37]. nTreg develop intrathymically, express FOXP3 constitutively and have a fully demethylated FOXP3 TSDR. In contrast, iTreg development takes place in the periphery, their FOXP3 expression is inducible and their FOXP3 TSDR is fully methylated. The MSC-mediated generation of cells with an immunosuppressive function is of particular importance if one considers the fate of MSC after infusion; Eggenhofer et al.  showed recently that after intravenous administration into mice, MSC survive no longer than 24 h. This evident short lifespan of MSC in connection with their proven ability to prolong graft survival prompts further investigation to reveal how MSC accomplish long-term immunosuppression, and which mediators and mechanisms are involved in this phenomenon.
While we have studied previously the interaction between human adipose-tissue derived MSC (ASC) and natural Treg , the aim of this study was to determine whether human ASC can generate functional de-novo iTreg from CD25-/dim effector T cells and to find evidence for the mechanisms involved in MSC-mediated Treg induction.
Material and methods
Origin, isolation and culture of human ASC
Perirenal adipose tissue was removed surgically from living kidney donors and collected in minimum essential medium Eagle alpha modification (MEM-α) (Sigma-Aldrich, St Louis, MO, USA) supplemented with 2 mM L-glutamine (Lonza, Verviers, Belgium), 1% penicillin/streptomycin solution (P/S; 100 IU/ml penicillin, 100 IU/ml streptomycin; Lonza). Samples were obtained with written informed consent, as approved by the Medical Ethical Committee at Erasmus University Medical Center Rotterdam (protocol no. MEC-2006-190).
ASC were isolated, cultured and characterized as described previously . In brief, perirenal adipose tissue was disrupted mechanically and digested enzymatically with collagenase type IV (Life Technologies, Paisley, UK). ASC were expanded using ASC culture medium consisting of MEM-α with 2 mM L-glutamine, 1% P/S and 15% fetal bovine serum (FBS; Lonza) in a humidified atmosphere with 5% CO2 at 37°C. Culture medium was refreshed twice weekly. At subconfluency, ASC were removed from culture flasks using 0·05% trypsin-ethylenediamine tetraacetic acid (EDTA) (Life Technologies) and reseeded at 1000 cells/cm2. ASC were characterized by means of immunophenotyping and by their ability to differentiate into adipocytes and osteoblasts. ASC cultured between two and six passages were used. ASC from these passages did not differ in their ability to differentiate or to exert their immunosuppressive functions.
Isolation of peripheral blood mononuclear cells
Peripheral blood mononuclear cells (PBMC) were isolated from buffy coats of healthy blood donors (Sanquin, Rotterdam, the Netherlands) by density gradient centrifugation using Ficoll-Paque PLUS (density 1·077 g/ml; GE Healthcare, Uppsala, Sweden). Cells were frozen at −150°C until further use in Roswell Park Memorial Institute (RPMI)-1640 medium with GlutaMAXTM-I (Life Technologies) supplemented with 1% P/S, 10% human serum (Sanquin) and 10% dimethylsulphoxide (DMSO; Merck, Hohenbrunn, Germany).
Isolation of effector cells and nTreg from PBMC
CD25–/dim cells (effector cells) and CD25bright cells (nTreg) were separated by means of CD25 MicroBeads II (Miltenyi Biotec, Bergisch Gladbach, Germany) and magnetic cell sorting, as described previously . Cell fraction purity was determined by flow cytometry using monoclonal antibodies (mAbs) against CD3-AmCyan (clone SK7), CD4-Pacific Blue (RPA-TA), CD25-phycoerythrin (PE)-cyanin 7 (Cy7) (epitope B; M-A251), CD127-PE (HIL-7R-M21; all BD Biosciences, San Jose, CA, USA); and FoxP3-allophycocyanin (APC) (PCH101; eBioscience, San Diego, CA, USA). Intracellular FoxP3 staining was carried out following the manufacturer's instructions of the anti-human FoxP3 staining set APC (eBioscience). Flow cytometric analyses were performed using the BD FACSCanto II flow cytometer and BD FACSDiva software (both BD Biosciences).
Mixed lymphocyte reaction and suppression assay
Mixed lymphocyte reactions (MLR) consisted of 5 × 104 CD25–/dim effector cells stimulated with 5 × 104 γ-irradiated (40 Gy) allogeneic PBMC in round-bottomed 96-well plates (Nunc, Roskilde, Denmark) using PBMC culture medium (PCM) consisting of MEM-α supplemented with 2 mM L-glutamine, 1% P/S and 10% heat-inactivated human serum. Effector–stimulator cell combinations were chosen on the basis of a minimum of four HLA mismatches. The immunomodulatory capacities of ASC (various concentrations), nTreg (1:10), ASC-induced CD4+CD25+CD127– T cells (1:10) and control CD4+CD25– T cells (1:10) (all ratios: indicated cells/effector cells) on the MLR were determined in suppression assays. After an 8-h incubation period on day 7, [3H]-thymidine incorporation (0·25 μCi/well; PerkinElmer, Groningen, the Netherlands) was measured using the Wallac 1450 MicroBeta TriLux (PerkinElmer). When MLR were performed in microtitre plates with different well sizes, the number of cells was adjusted accordingly. When applicable, 50 μl cell-culture supernatant was harvested prior to the addition of [3H]-thymidine and frozen at −80°C until further use.
Induction of CD4+CD25+CD127–FoxP3+ T cells by ASC
CD25–/dim effector cells were labelled using the PKH67 Green Fluorescent Cell Linker Kit (Sigma-Aldrich). For discrimination, allogeneic stimulator PBMC were labelled with PKH26 (PKH26 Red Fluorescent Cell Linker Kit; Sigma-Aldrich), according to the manufacturer's instructions. MLR were performed for 7 days in the absence or presence of ASC (1:40; ASC/effector cells). The PKH-label dilution caused by proliferation was measured by flow cytometry. After a 7-day incubation period in the absence or presence of 4 μg/ml basiliximab (Novartis Pharma, Nürnberg, Germany), cells were stained with mAbs against CD3-AmCyan (clone SK7), CD4-Pacific Blue (RPA-TA), CD8-peridinin chlorophyll (PerCP) (SK1), CD25-APC-Cy7 (epitope B; M-A251) and CD127-PE-Cy7 (HIL-7R-M21; all BD Biosciences) and FoxP3-APC (PCH101; eBioscience). Allogeneic stimulator PBMC were excluded from the analysis based on their PKH26-label. To confirm that basiliximab does not interfere with the binding of the monoclonal anti-CD25 antibody on epitope B, a competition staining was performed. In the presence of basiliximab no weakening of the CD25 staining was observed.
Isolation and function test of ASC-induced CD4+CD25+CD127– T cells
ASC-induced CD4+CD25+CD127– T cells were generated in primary MLR consisting of CD25–/dim effector cells and allogeneic stimulator PBMC in the presence or absence of ASC (1:40; ASC/effector cells). Allogeneic stimulator PBMC were labelled with PKH67 (Sigma-Aldrich). After 7 days, cells were stained with mAbs against CD3-AmCyan (clone SK7), CD4-Pacific Blue (RPA-TA), CD25-PE-Cy7 (epitope B; M-A251), CD127-PE (HIL-7R-M21) and BD Via-ProbeTM (7-AAD-PerCP) (all BD Biosciences). ASC-induced Treg were defined as PKH67–7-AAD–CD3+CD4+CD25+CD127– cells. Cell sorting was performed using the BD FACSAria II cell sorter (BD Biosciences).
Sorted PKH67–7-AAD–CD3+CD4+CD25+CD127– cells (1:10) were reseeded into secondary MLR. PKH67–7-AAD–CD3+CD4+CD25– cells (1:10) were used as negative control; nTreg (1:10) obtained from PBMC by magnetic cell separation served as positive control (all ratios: indicated cells/effector cells). After an 8-h incubation period on day 7, [3H]-thymidine incorporation (0·25 μCi/well; PerkinElmer) was measured. Alternatively, sorted cell samples were washed twice with phosphate-buffered saline-diethylpyrocarbonate (PBS-DEPC) and frozen at −80°C until further use.
Quantitative DNA methylation analysis of the FOXP3 gene TSDR
To quantify DNA methylation of the FOXP3 gene TSDR, the EZ DNA Methylation-Direct™ Kit (Zymo Research, Irvine, CA, USA) was used according to the manufacturer's instructions. Cell pellets were digested with proteinase K prior to bisulphite conversion. During DNA bisulphite treatment, unmethylated cytosines are converted into uracils while methylated cytosines remain unmodified. After bisulphite treatment, the TSDR of the FOXP3 gene was amplified by quantitative real-time PCR (qPCR) using the StepOnePlus™ real-time PCR system and the TaqMan® Genotyping Master Mix (all Applied Biosystems). Methylation-specific and demethylation-specific amplification primers and probes were chosen as suggested by Wieczorek et al. . The percentage of cells within a cell fraction with a methylated TSDR was calculated using the ratio of amplified methylated TSDR copies and the sum of amplified methylated and unmethylated TSDR copies. To correct for the X-linked nature of the FOXP3 gene, results obtained from female PBMC donors were multiplied by 2. Intra- and interassay variations were determined by negative controls and positive reference samples.
Flow cytometric characterization of CD4+CD25+CD127–FoxP3+ T cells and CD4+CD25+FoxP3– T cells
The induction of CD4+CD25+CD127–FoxP3+ T cells by ASC (1:40; ASC/effector cells) was initiated as described above. Allogeneic stimulator PBMC were labelled with either PKH67 or BD Horizon Violet Cell Proliferation Dye 450 (VPD450; BD Biosciences). After 7 days, cells were stained with mAbs against CD3-AmCyan (clone SK7), CD4-PerCP (SK3), CD25-APC-Cy7 (epitope B; M-A251), CD127-PE-Cy7 (HIL-7R-M21), cytotoxic T lymphocyte antigen-4 (CTLA-4)-APC (BNI3; all BD Biosciences), glucocorticoid-induced TNFR-related protein (GITR)-fluorescein isothiocyanate (FITC) (110416; R&D Systems Europe Ltd, Abingdon, UK), Helios-Pacific Blue (22F6; BioLegend, San Diego, CA, USA) and FoxP3-PE or FoxP3-APC (both PCH101; eBioscience). Fluorescence minus one (FMO) controls were used to determine negative expression.
mRNA expression analysis
ASC were cultured alone or co-cultured with MLR (1:5; ASC/effector cells) for 7 days. In co-cultures, MLR were separated from ASC by cell culture inserts with permeable membrane supports (0·4 μm pore size; Greiner Bio-One, Alphen a/d Rijn, the Netherlands). ASC were harvested and washed twice with PBS-DEPC. Cells were either handled immediately or snap-frozen in liquid nitrogen and stored at −80°C. Total RNA was purified using the High Pure RNA Isolation Kit (Roche Diagnostics), according to the manufacturer's instructions. Complementary DNA (cDNA) was synthesized by reverse transcription using random primers. qPCR was performed using 500 ng cDNA, the StepOnePlus™ real-time PCR system, TaqMan Universal PCR Master Mix and the assay-on-demand primer/probes for IL-2 (Hs00174114.m1) (Applied Biosystems, Foster City, CA, USA). Glyceraldehyde-3-phosphate dehydrogenase (GAPDH) mRNA served as endogenous control for normalization (Hs99999905.m1; Applied Biosystems). Intra- and interassay variations were determined by negative controls and positive reference samples. Changes in target gene expression relative to the GAPDH gene were quantified using the comparative CT method . Fold changes fewer than three were considered insignificant.
Cytometric bead array (CBA)
Supernatants were obtained from ASC cultures, MLR and ASC-MLR co-cultures (1:40; ASC/effector cells) in the absence or presence of 4 μg/ml basiliximab (Novartis Pharma) after a 7-day incubation period. They were frozen until further use at −80°C. IL-2 concentrations were measured using the BD cytometric bead array human T helper type 1 (Th1)/Th2/Th17 Cytokine Kit (BD Biosciences), according to the manufacturer's instructions.
Flow cytometric analysis of IL-2 expression by CD4+CD25+CD127–FoxP3+ T cells and CD4+CD25+FoxP3– T cells
CD4+CD25+CD127–FoxP3+ T cells were induced by ASC (1:40; ASC/effector cells) as described above. CD25–/dim effector cells were labelled with VPD450 (BD Biosciences). For discrimination, allogeneic stimulator PBMC were labelled with PKH67 (Sigma-Aldrich). MLR were performed in the absence or presence of ASC and 4 μg/ml basiliximab (Novartis Pharma). After 7 days, cells were stimulated with 50 ng/ml phorbol 12-myristate 13-acetate (PMA) and 1 μg/ml calcium ionomycin (both Sigma-Aldrich) in the presence of BD GolgiStop (BD Biosciences) for 4 h. Subsequently cells were stained with mAbs against CD3-AmCyan (clone SK7), CD4-PerCP (SK3), CD25-APC-Cy7 (epitope B; SK1), CD127-PE-Cy7 (HIL-7R-M21), IL-2-APC (5344.111; all BD Biosciences) and FoxP3-PE (PCH101; eBioscience). FMO controls were used to determine negative expression.
Statistical analyses were performed by means of one-way analysis of variance (anova), Bonferroni's multiple comparison tests and (un)paired t-tests using GraphPad Prism 5 software (GraphPad Software, San Diego, CA, USA). A P-value lower than 0·05 was considered statistically significant. Two-tailed P-values are stated.
ASC inhibit the proliferation of CD25–/dim effector cells dose-dependently and induce CD4+CD25+CD127–FoxP3+ T cells
The immunosuppressive effect of ASC on the proliferation of CD25–/dim effector cells was examined by means of [3H]-thymidine incorporation. CD25–/dim effector cells were stimulated with γ-irradiated allogeneic PBMC for 7 days resulting in a strong proliferative activity of these cells (Fig. 1a). Co-culture with third-party ASC suppressed the effector cell proliferation in a dose-dependent manner (one-way anova, P < 0·0001), confirming our previously published data . ASC reduced significantly the proliferation from a ratio of 1:80 (ASC/effector cells). ASC suppressed the proliferation of CD4+ T cells and CD8+ T cells (Fig. 1b).
The effect of ASC on the generation of CD4+CD25+CD127–FoxP3+ T cells from CD25–/dim cells was determined by flow cytometry. At a 1:40 ratio, ASC reduced the proliferation of CD25–/dim effector cells by 59% (Fig. 1a) and mediated a 2·1-fold increase in the percentage of CD25+CD127–FoxP3+ cells within the CD4+ T cell population (Fig. 2).
ASC-induced CD4+CD25+CD127– T cells are immunosuppressive
The suppressive capacity of ASC-induced CD4+CD25+CD127– T cells was determined by means of [3H]-thymidine incorporation and compared to the effect of nTreg. Sorted ASC-induced CD4+CD25+CD127– T cells (median purity: 98·7%; range 95·3%–99·7%) were added at a 1:10 ratio to secondary MLR. They inhibited the proliferation of CD25–/dim effector cells as effectively as nTreg (1:10; 61% suppression versus 48%, respectively; P = 0·402; Fig. 3). Hence, ASC induce functional regulatory T cells. MLR-induced CD4+CD25+CD127– T cells (median purity: 99·5%; range 98·5–99·8%) also suppressed proliferation (1:10; 87%). Sorted CD4+CD25– T cells from an MLR or MLR–ASC co-culture (1:10; median purities 99·8 and 99·7%; ranges 99·5–99·8% and 99·3–99·9%), as negative controls, did not inhibit the proliferative activity of the effector cells.
Functional ASC-induced CD4+CD25+CD127– Treg are de-novo cells
To determine the origin of the ASC-induced Treg, i.e. iTreg or nTreg, the methylation status of the TSDR was investigated. The percentage of cells with a methylated FOXP3 gene TSDR present in different cell fractions was determined by means of quantitative DNA methylation analysis. The CD25–/dim fraction, obtained from PBMC by magnetic cell separation, consisted almost entirely (98·1%) of cells with a methylated FOXP3 TSDR (Fig. 4a). In contrast, the nTreg fraction consisted to 67·3% of cells with a demethylated FOXP3 TSDR, confirming their thymic origin. After 7-day MLR in the presence of ASC (1:40; ASC/effector cells) using allostimulated CD25–/dim cells as effector cells, sorted fractions of CD4+CD25+CD127– iTreg and CD4+CD25– T cells contained 83·7 and 99·9% cells with a methylated FOXP3 TSDR, respectively. The small percentage of cells (16·3%) with a demethylated FOXP3 TSDR in the CD4+CD25+CD127– iTreg fraction probably represents proliferated nTreg which were present in the initial CD25-/dim fraction (1·9%). This demonstrates that ASC expand nTreg, but that the majority of the CD4+CD25+CD127– Treg found after 7-day MLR–ASC co-culture are induced from CD25–/dim cells.
To characterize further the ASC-induced CD4+CD25+CD127–FoxP3+ Treg (Fig. 4b), their expression of GITR, CTLA-4 and Helios was investigated (Fig. 4c,d). While the expression levels of GITR were similar for iTreg and CD4+CD25+FoxP3– T cells, iTreg showed the tendency to have higher Helios levels and expressed 3·2-fold more CTLA-4 than FoxP3– T cells (P = 0·003).
The induction of CD4+CD25+CD127–FoxP3+ Treg by ASC coincides with increased IL-2 levels and is IL-2 pathway-dependent
IL-2 is known to be required for the expansion and function of nTreg. Therefore, its involvement in the ASC-mediated induction of CD4+CD25+CD127–FoxP3+ Treg was investigated (Fig. 5). Seven-day MLR were cultured in the absence or presence of ASC (1:40; ASC/effector cells) and concentrations of secreted IL-2 in the supernatant were analysed. IL-2, 20·1 pg/ml, was detected in the supernatant of an MLR of CD25–/dim effector cells stimulated with allogeneic PBMC. Non-activated ASC and MLR-activated ASC do not express IL-2 (data not shown). However, activated ASC mediated a 15·1-fold increase in IL-2 levels by effector cells during MLR suppression and induction of CD4+CD25+CD127–FoxP3+ Treg. The monoclonal anti-IL-2 receptor antibody basiliximab (4 μg/ml) effectively inhibited the IL-2 consumption in MLR and MLR–ASC co-cultures (1:40; ASC/effector cells; Fig. 5a); in the presence of basiliximab IL-2 concentrations in MLR and MLR–ASC co-cultures accumulated to 217 and 525 pg/ml, respectively.
To gain knowledge regarding which cell subset contributed most to the IL-2 concentrations detected in the supernatant, the percentage of IL-2-expressing cells within the CD4+CD25+CD127–FoxP3+ iTreg and CD4+CD25+FoxP3– T cells was investigated. The majority of the IL-2-expressing cells were CD4+CD25+FoxP3– T cells (Fig. 5b,c). The IL-2 levels produced were similar between the IL-2 expressing CD4+CD25+CD127–FoxP3+ iTreg and CD4+CD25+FoxP3– T cells (Fig. 5d).
To examine the effect of the IL-2 pathway on the ASC-mediated generation of functional CD4+CD25+CD127–FoxP3+ iTreg from CD25–/dim cells, IL-2 binding to its receptor was blocked with basiliximab. MLR were performed for 7 days in the absence and presence of ASC (1:40; ASC/effector cells). Basiliximab reduced the percentage of CD25+CD127–FoxP3+ cells within the CD4+ T cell population (Fig. 6). In the presence of basiliximab, ASC did not induce a proportional increase of CD25+CD127–FoxP3+ cells within the CD4+ T cell population.
Knowledge about the underlying mechanisms of how ASC contribute to a reduced responsiveness of immune effector cells is scarce. The present study provides the first evidence that human ASC mediate their immunosuppressive effect via the formation of functional de-novo iTreg. Our results are in line with earlier in-vivo studies with bone-marrow MSC in animal models and a recent case report of MSC administration to renal transplant patients; these studies also observed proportional increases of Treg [9, 35, 41, 42]. However, as these groups did not investigate the Treg origin and Treg functionality, it is not clear whether the observed rises in Treg percentages are a consequence of the de-novo formation of Treg or a result of the expansion of existing nTreg. We found that ASC mediate the generation of iTreg from effector T cells and that these newly formed cells have immunosuppressive capacities. iTreg also formed in MLR without ASC, but at lower numbers. While ASC-induced Treg showed similar suppressive capacities to nTreg, MLR-induced Treg had a stronger inhibitory effect. The induction of a more potent immunosuppressive phenotype in these cells might be due to their generation under highly proinflammatory conditions. In the presence of ASC effector cell proliferation is reduced. Because of this milder environment higher numbers of less effective iTreg are generated. Therefore, the suppressive strength of iTreg might be proportional to the stimulus under which they are induced. Further, due to the presence of a few nTreg in our initial effector T cell population we were able to infer that ASC also mediate an increase in nTreg.
Although the first evidence of extrathymic conversion of conventional T cells into iTreg was found almost a decade ago, no iTreg-specific marker has yet been identified [43-45]. The only tool currently available to distinguish iTreg from nTreg is the determination of the methylation status of the TSDR. While nTreg have a demethylated TSDR, this specific region of the FOXP3 gene is methylated in iTreg . The high percentage of cells with a methylated TSDR in the ASC-induced Treg fraction indicates that the vast majority of these cells originated from CD25–/dim effector cells. To characterize further the ASC-induced Treg we investigated their expression of GITR, CTLA-4 and Helios. GITR expression was similar in both iTreg and CD4+CD25+FoxP3– T cells. In contrast, iTreg showed a tendency towards a higher Helios expression and expressed significantly higher CTLA-4 levels than FoxP3– T cells. It has been described that CTLA-4 expression is up-regulated upon binding of FoxP3 to the promoter of the CTLA-4 gene [46-50]. Therefore, the observed difference in CTLA-4 expression between iTreg and FoxP3– T cells can be attributed to their differing FoxP3 expression. Whether Helios is a potential marker for human iTreg is an ongoing debate in the field, as conflicting results have been reported [51-53].
In search of possible underlying mechanisms involved in MSC-mediated iTreg generation, it has been reported recently that programmed death ligand-1 (PD-L1), a protein expressed by MSC, promotes the differentiation of Th1 cells into Treg [22, 54]. Other molecules that were suggested to play a role in the induction of Treg by MSC are IDO, prostaglandin E2 (PGE2), transforming growth factor (TGF)-β and haem oxygenase-1 (HO-1) [41, 55-58]. In this study we focused on the role of IL-2 in ASC-mediated Treg induction, as IL-2 drives T cell proliferation and, paradoxically, is also essential for tolerance induction by regulating nTreg function . We observed that in the presence of ASC, the IL-2 concentration in the cell culture supernatant increased. Because IL-2 is not expressed by ASC, the IL-2 surplus originated from the alloactivated effector cells. Upon T cell-receptor engagement and co-stimulation, activated effector T cells consume IL-2. During the inhibition of effector T cell proliferation by ASC, IL-2 concentrations in the cell culture supernatant rose. This indicates that although their proliferation is suppressed, effector T cells remain activated and continue to secrete IL-2. The diminished proliferation rate of effector T cells causes a reduced IL-2 consumption and hence a surplus of IL-2. High levels of IL-2 are instrumental for Treg induction. In the presence of basiliximab, we found further accumulation of IL-2 in the supernatant. In its function as anti-rejection therapy in kidney transplant patients, basiliximab prevents the binding of IL-2 to its receptor and the subsequent uptake of IL-2. As a result, basiliximab inhibits T cell proliferation but also iTreg generation, indicating that Treg induction by ASC is IL-2 pathway-dependent. Surprisingly, IL-2 was expressed not only by CD4+CD25+FoxP3– T cells but also by a smaller percentage of immunosuppressive ASC-induced Treg. Phenotypic adaptation of Treg in response to the cytokines present in their environment is a known phenomenon . The inhibition of T cell proliferation is not an exclusive effect of MSC, as other stromal cells share this characteristic [61, 62] Hence, it is possible that these cells can also create an environment that favours iTreg generation.
The graft-supporting effects of MSC in animal models and the positive results of MSC treatment for graft-versus-host disease (GVHD) in patients who underwent haematopoietic stem cell transplantation (HSCT) are strongly convincing [8, 63]. The reported short lifespan of MSC after infusion, however, indicates that MSC are only initiators of these effects. Long-term, MSC do not actively promote immunosuppression themselves. Nevertheless, they are able to pass on their immunosuppressive capabilities through the induction of functional de-novo Treg, the expansion of nTreg or possibly through other immune cells which remain to be identified.
Despite continuous efforts, the translation of the experimental success of Treg-mediated immune regulation to its application in clinical settings proves difficult. In animal models, ex-vivo- or in-vivo-generated Treg prevent type 1 diabetes, reduce the severity of experimental autoimmune encephalomyelitis (EAE), control the acute and chronic rejection of allografts and attenuate or prevent GVHD [64-70]. In humans, thus far, ex-vivo-expanded Treg have been used only for the treatment of patients undergoing HSCT. In these few clinical studies, Treg were able to reduce the incidence of GVHD or to prevent GVHD [71-73]. Due to the heterogeneity of the Treg population and the lack of a specific marker for human Treg, ex-vivo Treg expansion bears the risk of contamination with effector T cells. Therefore, in-vivo induction of functional Treg mediated by MSC represents a beneficial effect of MSC therapy in addition to its immunosuppressive effect on effector T cells. Once the functionality of MSC-induced Treg has been confirmed in autoimmune disease patients and allograft recipients, strategies to enhance the MSC-mediated in-vivo generation of iTreg may be considered. Possible approaches could be MSC treatment combined with low-dose IL-2 therapy or the use of rapamycin and rabbit anti-thymocyte globulin (rATG), anti-rejection drugs which were found to advance nTreg expansion and to allow Treg induction, respectively [70, 74-76].
In conclusion, our study demonstrates that human adipose tissue-derived MSC induce Treg from effector T cells and that these de-novo Treg are immunosuppressive. In conjunction with the well-known MSC function of preventing immune cell proliferation, our findings encourage the advancement of MSC therapy into clinical development for autoimmunity, GVHD and allograft rejection.
The authors of this manuscript have no conflicts of interest to disclose.