Skip to main content
NCBI home page
Immunology logoLink to Immunology
. 2015 Feb 13;144(3):431–443. doi: 10.1111/imm.12388

Hypoxic culture conditions enhance the generation of regulatory T cells

Thi My Anh Neildez-Nguyen 1,2,3,✉, Jérémy Bigot 1,2,3, Sylvie Da Rocha 1,2,3, Guillaume Corre 1,2,3, Florence Boisgerault 1,2,3, Andràs Paldi 1,2,3, Anne Galy 1,2,3
PMCID: PMC4557680  PMID: 25243909

Abstract

The generation of large amounts of induced CD4+ CD25+ Foxp3+ regulatory T (iTreg) cells is of great interest for several immunotherapy applications, therefore a better understanding of signals controlling iTreg cell differentiation and expansion is required. There is evidence that oxidative metabolism may regulate several key signalling pathways in T cells. This prompted us to investigate the effects of oxygenation on iTreg cell generation by comparing the effects of atmospheric (21%) or of low (5%) O2 concentrations on the phenotype of bead-stimulated murine splenic CD4+ T cells from Foxp3-KI-GFP T-cell receptor transgenic mice. The production of intracellular reactive oxygen species was shown to play a major role in the generation of iTreg cells, a process characterized by increased levels of Sirt1, PTEN and Glut1 on the committed cells, independently of the level of oxygenation. The suppressive function of iTreg cells generated either in atmospheric or low oxygen levels was equivalent. However, greater yields of iTreg cells were obtained under low oxygenation, resulting from a higher proliferative rate of the committed Treg cells and higher levels of Foxp3, suggesting a better stability of the differentiation process. Higher expression of Glut1 detected on iTreg cells generated under hypoxic culture conditions provides a likely explanation for the enhanced proliferation of these cells as compared to those cultured under ambient oxygen. Such results have important implications for understanding Treg cell homeostasis and developing in vitro protocols for the generation of Treg cells from naive T lymphocytes.

Keywords: Glut1, oxygenation, PTEN, reactive oxygen species, regulatory T-cell induction, Sirt1

Introduction

Regulatory CD4+ T lymphocytes (Treg) are central elements of cell-mediated immune tolerance and include natural Treg (nTreg) cells, which develop early in the thymus, and induced Treg (iTreg) cells, which arise through the conversion of peripheral naive T cells. Both types of Treg cells express the transcription factor Forkhead box P3 (Foxp3), which is essential for their function. Treg cells have potential medical applications, notably in transplantation, auto-immunity or to facilitate tolerance to therapeutic proteins.1 As a consequence, detailed protocols have been developed for the in vitro expansion of nTreg or iTreg cells using cord blood or mobilized peripheral blood cells in the perspective of clinical applications.2,3 A variety of factors and signalling molecules are well known to influence the production of iTreg cells in humans and mice, in particular the cytokines transforming growth factor-β (TGF-β) and interleukin-2 (IL-2) (recently reviewed by Dons et al.4). The identification of additional factors involved in the cell differentiation and growth of Treg cells would therefore be highly pertinent.

It has been shown that T-cell activation requires increased glucose metabolism, which through support of bioenergetic and macromolecular synthesis, facilitates rapid cell growth and proliferation.5–7 In particular, T-cell receptor (TCR)/CD28 co-stimulation activates the phosphoinositide 3-kinase (PI3K)/Akt/ mammalian target of rapamycin (mTOR) pathway to promote the increase of glucose uptake by cell surface trafficking of the Glut1 glucose transporter, which is a limiting factor for T-cell proliferation upon activation.8,9 Signalling through the IL-2 receptor (IL-2R) has been shown to activate mTOR and is able to prevent or reverse anergy in activated T cells.10 While data identify Phosphatase and tensin homologue deleted on chromosome 10 (PTEN), a lipid phosphatase, as a negative regulator of IL-2R signalling in CD4+ T cells, which can inhibit the expansion of these activated T cells11 and their differentiation into Treg cells4,12 in response to IL-2. Activated CD4+ T cells are metabolically and developmentally flexible as they can differentiate into effector T cells (Teff) or Treg cells depending on the local environment. These two functionally distinct types of cells show distinct metabolic requirements for their specification and survival since Treg cells appear to be more prone to up-regulate lipid oxidation while Teff cells are more glycolytic.13 Downstream of PI3K/Akt, the mTOR pathway has been found to be essential for the determination of the CD4+ T-cell fate, being required for differentiation into Teff cells, while the lack of mTOR or its inhibition by rapamycin foster Treg cell differentiation.2,14–16 The signalling of mTOR can also be negatively regulated by metabolic and stress-response pathway mediators, notably by Sirt1,17 a class III NAD+-dependent histone deacetylase providing novel potential pathways linking metabolic regulation and iTreg cell differentiation.18 Besides, Sirt1 is known to mediate deacetylation of Foxp3, thereby enhancing its proteasomal degradation.19–21

Oxygen is therefore a key regulator of cellular metabolic states that may control iTreg generation. Standard in vitro cultures of mammalian cells are generally performed at atmospheric oxygen levels (21% O2). However, oxygen concentrations are normally much lower in mammalian organisms, ranging between 1% and 14% O2 depending on the tissue. It has been shown that culturing T cells under physiological oxygenation modulates their proliferation rate, function, activation status, surface receptor expression, intracellular reactive oxygen species (ROS) and the production of cytokines.22–26 A low-O2 environment enables the accumulation of extracellular adenosine, a factor recently implicated in the induction of Treg cells.27,28 However, it is not known precisely how oxygen levels affect the generation of Treg cells in culture. Only a few studies have examined the effect of hypoxia and of hypoxia-induced factors such as hypoxia-inducible factor-1α on the production of Treg cells29 but results are controversial – reporting either an essential role for promoting Treg cell differentiation30,31 or an inhibitory effect.32,33 The effects may be complex to interpret in cultures that include various types of precursor cells and of differentiated cells.13,30,31 Here we used a simplified in vitro culture system to assess how O2 level supply influences iTreg cell generation through the analysis of different proteins involved in the regulation of Treg cell differentiation. Experiments were conducted using bead-stimulated transgenic mouse T cells; these allow live detection of FoxP3 expression and are useful to study the autonomous signals involved in iTreg generation from naive T cells. In this study, we demonstrate that the commitment of CD4+ cells to the Treg cell lineage pathway is dependent on the production of superoxide anions and is accompanied by increased levels of Sirt1, PTEN and Glut1, which characterize the process of Treg differentiation. We also show that the generation of Treg cells is enhanced under low oxygenation due to a better cellular amplification of the committed cells as facilitated by a higher expression of Glut1 at the cell membrane. These novel results may help to find optimized cell culture parameters for expansion of suppressive T cells.

Materials and methods

Mice

Mice were housed under specific pathogen-free conditions and handled in accordance with French and European directives. C57BL/6 mice were purchased from Charles River (l’Arbresle, France). Simone transgenic mice with fluorescent Treg cells (Tg(TcraH-Y,TcrbH-Y)1Pas, Ptprc, Foxp3, Rag2) were generated and housed in our facility by crossing Foxp3-GFP-KI mice (B6.Cg-Foxp3tm1Mal/J)34 with Marilyn mice (B6.129-Ptprca Rag2tm1Fwa Tg(TcraH-Y,TcrbH-Y)1Pas/Pas).35 Simone mice are homozygous for mutations Foxp3eGFP, Rag-2–/– and for a TCR specific for a complex of the male antigen HY, Dby peptide with IA-b. For some experiments (only in Fig.4), we also used HY2 mice generated in our facility as F1 cross between Foxp3-GFP-KI mice and Simone mice. HY2 mice have homozygous Foxp3eGFP and TCR alleles, but heterozygous Rag-2 (Rag-2+/–). Animals were used in experiments at between 6 and 9 weeks of age.

Figure 4.

Figure 4

Open in a new tab

Equivalent suppressive activities of induced regulatory T (iTreg) cells generated under different oxygen conditions. After 7 days of in vitro generation under either 5% or 21% O2, CD4+ GFP+ iTreg cells were FACS-sorted and then co-cultured with different ratios of CFSE-labelled T effector cells (responders) stimulated to proliferate by Dby peptide and antigen-presenting cells during 72 hr, in 21% O2 (a), or in 5% O2 (b). Cells were then harvested and CFSE signal was measured by flow cytometry. Bar diagrams represent the percentage of parental population corresponding to the percentage of responders that have not divided. Negative control of effector T-cell proliferation consisting of cultures of not-stimulated (– peptide) effector T cells, and positive control consisting of cultures of stimulated (+ peptide) effector T cells, both without iTreg cells, are included. Different Treg : Teff ratios (1 : 8, 1 : 4, 1 : 2 or 1 : 1) were studied.

Cell culture

For induction of CD4+ CD25+ Foxp3+ GFP+ Treg cells, CD4+ naive T cells were purified from Simone mouse spleen by Dynal negative isolation kit (Invitrogen, Saint Aubin, France), according to the manufacturer’s recommendations. Purity of isolated CD4+ cells was usually > 80%. These cells were cultured in RPMI-1640 containing 10% fetal calf serum, 100 U/ml penicillin/streptomycin, 2 mm glutamax, 50 μm β-mercaptoethanol, in the presence of recombinant human IL-2 (30 U/ml; Prometheus Laboratories, Inc., San Diego, CA), TGF-β (20 ng/ml; Miltenyi Biotec, Paris, France) and stimulated with Dynabeads anti-CD3/CD28 Mouse T Cell Expander (0·125 beads/cell; Dynal, Invitrogen). When indicated, Mn(III)tetrakis(4-benzoic acid) porphyrin chloride (MnTBAP; 200 μm; Calbiochem, Fontenay sous Bois, France) was added at the beginning of the culture described above, to neutralize ROS. Glycolytic inhibitor 2-deoxy-d-glucose (2-DG; Sigma, St-Quentin-Fallavier, France) or drugs to modulate Sirt1 or mTOR activities, all purchased from Enzo Life Sciences (Villeurbanne, France), were added, when indicated, at the beginning of the culture described above: 1 mm 2-DG, 10 μm Resveratrol (Sirt1 activator), 10 μm Sirtinol (Sirt1 inhibitor) and 50 nm rapamycin (mTOR inhibitor). At different time-points during cultures, cell counts and viability were determined by trypan blue exclusion, and Dynabeads were removed with magnetic columns before immunostaining. Cultures were maintained in a 37° incubator containing 5% CO2, and either atmospheric (21%) O2 or 5% O2, condition obtained by nitrogen injections (Thermo Electron Corporation incubator, Thermo Scientific, Villebon sur Yvette, France).

Antibodies and flow cytometric analysis

The following antibodies were used for flow cytometric analysis: anti-CD4 (RM4-5), CD25 (PC61) from BD Pharmingen (Le Pont de Claix, France), and Foxp3 (FJK-16s) from eBioscience (Paris, France). Fc receptors were blocked by the addition of unlabelled anti-CD16/32 (Fc block; BD Pharmingen). According to experiments, Foxp3 staining was performed using the intracellular Foxp3 staining kit as described in the protocol (eBioscience). Gates for expression of CD25 or Foxp3 in the CD4+ population were set according to isotype controls (BD Pharmingen and eBioscience). Cell viability was determined by staining with 7-aminoactinomycin-D (7-AAD; Sigma) or Fixable Viability Dye eFluor 780 (eBioscience) in intracellular labelling assays.

At the indicated times, intracellular ROS production was determined by dihydroethidium (DHE) (10 μm; Invitrogen) staining for 30 min at 37°. The cells were washed once, resuspended in PBS, 0·1% BSA with Sytox Red (5 μm; Invitrogen) for cell viability determination, and then subjected to flow cytometric analysis. Samples were acquired on an LSR II flow cytometer (BD Biosciences, Le Pont de Claix, France) and data were analysed with DIVA software (BD Biosciences).

Treg cell proliferation assay

CellTrace Violet (CTV)-fluorescent dye labelling of cells was performed according to the manufacturer’s recommendations (Invitrogen). Briefly, purified naive CD4+ T cells were resuspended to 1 × 106/ml in PBS, and a final concentration of 5 μm of CTV (Invitrogen) was added. Cells were incubated at 37° for 20 min, protected from light. The labelling reactions were stopped with complete RPMI-1640 medium, and cells were washed and resuspended in pre-warmed complete culture medium for Treg cell induction. CTV dye-labelled cells were cultured in 5% or 21% O2 before antibody and 7-AAD staining, and analysis by flow cytometry at the indicated time-points. Flow cytometric data files were analysed with the proliferation wizard module of ModFIT LT software (Verity Software House, Topsham, ME), by gating on 7-AAD– CD4+ CD25+ GFP+ living iTreg cells generated at the studied time-point of culture, to calculate the Proliferation Index (ratio of the total number of cells analysed to the calculated number of parental cells required for the observed cell number). Increasing numbers of replications are reflected by higher Proliferation Index values.

In vitro suppression assays

Induced Treg cells, generated after 7 days of culture in 5% or 21% O2, were purified as 7-AAD– CD4+ GFP+ with MoFlo cell cytometry sorting (Beckman-Coulter, Villepinte, France). The isolated Treg samples (CD45.1+) were then co-cultured for 72 hr in 5% or 21% O2, with different ratios of CFSE- (Invitrogen) labelled Teff cells (CD45.2+ responders) from HY2 mice (1 : 8, 1 : 4, 1 : 2 or 1 : 1 Treg/Teff ratios), in the presence of 2 × 105 irradiated antigen-presenting cells from C57BL/6 mice and 10 nm of Dby peptide (GENEPEP, St Jean de Védas, France). The suppressive capacity of iTreg cells was expressed as the percentage of the responders’ parental population, which increases with higher Treg :Teff ratios, reflecting Treg-mediated inhibition of responders T cell proliferation. The percentage of parental population defined as the percentage of responders that have not divided, and so have retained the initial CFSE fluorescence intensity level, was assessed by flow cytometry.

Fluorescence microscopy and image analysis

At the indicated time-points in culture, cells were collected and subjected to immunostaining as described earlier by Brock et al.36 Briefly, after adhesion on poly-l-lysine-coated slides (Thermo Scientific), the cells were fixed in 4% paraformaldehyde for 5 min at 4° and for 10 min at room temperature, and permeabilized with methanol for 6 min at −20°. Cells were stained overnight with the primary antibody, and then with the AlexaFluor 594 secondary antibody (Invitrogen) for 30 min at room temperature. The slides were mounted with Fluoromount/Dapi (SouthernBiotech, Montrouge, France). Antibodies used are: anti- Sirt1 (H-300; Santa Cruz Biotechnology, Santa Cruz, Nanterre, France), PTEN ((C-20)-R, Santa Cruz Biotechnology), Glut1 (Thermo Scientific) and Foxp3 (FJK-16s, eBioscience). Negative controls were performed on CD4+ naive T cells labelled with the corresponding isotype control (eBioscience and Southern Biotech). Confocal analysis was done on a Leica TCS SP2 DMRE confocal-laser-scanning microscope (Leica Microsystems, Nanterre, France), with 40× immersion objective. Images were analysed with the free open-source image analysis software CellProfiler (Broad Institute, Harvard, MA).37 Pipeline for the analysis includes modules used for cell identification, cell segmentation, cell size and fluorescence quantification. A minimum of 1000 cells were analysed for each studied group. Data were imported in R software (http://www.R-project.org/) and histograms were generated.

Statistics

Statistical analyses were performed using graphpad prism 5 (Logi Labo, Paris, France). Data were expressed as the mean ± SEM. Student’s paired t-test or Kolmogorov–Smirnov test were applied with significance levels set at *P < 0·05, **P < 0·005, ***P < 0·0005 and ****P < 0·0001.

Results

Yields of iTreg cells are increased in hypoxic cultures due to a better cellular amplification

Cells from Foxp3-KI-GFP TCR mice (so-called Simone mice) provide a useful model to study iTreg generation because they carry a transgenic CD4+ TCR specific for the Dby peptide of the male HY antigen, and also bear a reporter GFP transgene knocked into the Foxp3 locus, which serves to identify live iTreg cells by fluorescence microscopy or flow cytometry. In the steady-state, there are no Foxp3+ cells in peripheral lymphoid organs of Simone mice but splenic T cells can be induced to differentiate into iTreg cells in vitro following TCR stimulation in the presence of Dby peptides or under polyclonal stimulation using anti-CD3/CD28 beads, TGF-β and IL-2 as described in other systems.38 To test the hypothesis that oxygenation impacts on the efficiency of iTreg cell generation, we polyclonally-stimulated CD4+-enriched spleen cells from Simone mice in cultures using two different oxygen levels, i.e. 21% O2 (atmospheric O2 levels) generally used in standard in vitro culture systems and 5% O2, a level much closer to the physiological situation encountered by splenic T cells in vivo.24 The percentage of iTreg cells obtained in culture was defined as the percentage of CD25+ GFP+ cells gated on the viable CD4+ cell population. A higher efficiency of iTreg cell generation was observed under 5% O2 compared with 21% O2, starting to be discerned by day 3, although not yet significant (Fig.1a), but becoming clearer at later time points (Fig.1a) (at day 7: 68 ± 5% and 53 ± 4% of CD25+ GFP+, respectively; at day 11: 81 ± 3% and 65 ± 4% of CD25+ GFP+, respectively). This result was also validated on the basis of the intracellular marker Foxp3 detection. Indeed, at day 11 the highest percentages of CD25+ GFP+ Foxp3+ cells were found within the CD4+ cell population under 5% O2 (Fig.1b) and this was also confirmed by immunofluorescence microscopy (Fig.2a). Image immunofluorescence quantification showed that percentages of Foxp3+ cells were not only increased, but the majority of 5% O2-induced Treg cells also expressed higher levels of Foxp3 (Fig.2b) as confirmed by flow cytometry (Fig.2c), suggesting a better stability of their phenotype than cells cultured under 21% O2. Independently of O2 levels, CD25 expression was similar on iTreg cells (Fig.2d). In addition, CD4+ GFP− cell number was decreasing (data not shown), probably as a result of suppressive activity of iTreg cells generated in culture in both O2 conditions. Low oxygenation increased by about two-fold the absolute number of iTreg cells (CD4+ CD25+ GFP+) generated from day 7 of culture compared with 21% O2 (Fig.3a). This increase was not due to a better survival of iTreg cells under hypoxic culture conditions. Cell viability as assessed by trypan blue (data not shown) or 7-AAD labelling showed no significant difference between the two conditions of oxygenation (Fig.3b). However, cell mortality was observed at the beginning of culture (at day 3, average 40%, Fig.3b) but was independent of the level of oxygenation. Cell proliferation was assessed with the use of a stable cell membrane-permeable dye, CTV, which is a culture-compatible probe enabling the tracking of the in vitro cell division history of the live GFP+ cell population by FACS. This technique showed that after 7 days of culture when absolute numbers of iTreg cells differ clearly according to oxygen levels, the viable CD4+ CD25+ GFP+ cells had the highest proliferative activity under 5% O2, as shown by a shift to the left of the last generation of cells (Fig.3c). The calculated Proliferation Index of iTreg cells at 5% and 21% O2, respectively, was 26 and 14 at day 7 (data not shown) and 30 and 20 at day 11 (Fig.3c), thereby quantifying this effect. The suppressive capacity of purified cells generated in atmospheric or hypoxic culture conditions was tested in a stringent antigen-specific proliferation assay based on the suppression of Dby-induced immune responses. Flow cytometry cell-sorted iTreg cells derived from 7-day cultures either in 5% or 21% O2 were co-cultured for 72 hr with different ratios of CFSE-labelled Teff cells (responders) obtained from Dby-specific TCR transgenic mice and stimulated to proliferate by the Dby peptide (Fig.4). By measuring variations from the initial CFSE fluorescence intensity level, FACS analysis showed that iTreg cells derived from cultures either at 5% or 21% O2 were able to suppress T-cell proliferation in a dose-dependent manner and regardless of the oxygenation level at which co-cultures were performed (Fig.4a,b). These data show that equally functional iTreg cells are generated under 5% O2 than 21% O2, but greater amounts of cells are generated under 5% O2 due to higher cellular division of committed cells. These results prompted us to examine specific signalling pathways.

Figure 1.

Figure 1

Open in a new tab

Low O2 levels increase yield of induced regulatory T (iTreg) cell conversion from naive T cells. The iTreg cells were induced from Simone mice naive CD4+ T cells for 3, 7 or 11 days under either 5% or 21% O2. Culture samples were then analysed by flow cytometry to determine the level of iTreg cell conversion. (a) Percentage of CD25+ GFP+ gated on living (7-AAD−) CD4+ T cells (Treg cells) at days 3, 7 and 11 under either 5% or 21% oxygen. Results are from independent experiences. Lines between dots indicate paired samples. ns, not statistically significant, ****P < 0·0001. (b) Flow cytometry analysis of day 11 Treg cell induction cultures, done under either 5% or 21% O2. Cells gated on living CD4+ T-cell population were analysed for their expression of GFP (upper panels) or Foxp3 (bottom panels) versus CD25. Inset numbers indicate the percentage of cells in each quadrant. Data are representative of at least three independent experiments.

Figure 2.

Figure 2

Open in a new tab

Induced regulatory T (iTreg) cells produced under low O2 concentration express higher level of Foxp3. After 11 days of Treg cell induction in cultures under either 5% or 21% O2, cells were collected, stained with Foxp3-allophycocyanin antibody and analysed by fluorescence microscopy as described in Materials and methods. Negative controls were performed with an allophycocyanin-isotype control antibody. Images were acquired on LEICA SP2 confocal microscope, with a 40× oil immersion objective. Each image is a mosaic composed of nine images. (a) Panels show DAPI nuclear staining (blue) and Foxp3 staining (green) of day 11 Treg induction cultured cells produced at either 5% (bottom panels) or 21% (upper panels) O2. Scale bar, 50 μm. Data represent typical results from at least three independent experiments. (b) Density histograms of mean intensity of Foxp3 expression in day 11 Foxp3+ cells generated under either 5% (black line) or 21% (grey line) O2. The grey-shaded area represents staining with the isotype control antibody. Fluorescence was analysed using Cell Profiler software and density histograms were created using R. Data are representative of at least three independent experiments, n ≥ 1500. (c) Mean fluorescence intensity (MFI) determined by flow cytometry of intracellular Foxp3 antigen, (d) and surface CD25 antigen in living CD4+ T cells that co-express CD25 and Foxp3 (upper-right quadrant in Fig.1b) from 11 days Treg cell induction cultures under either 5% or 21% O2. In (c) and (d), lines between dots indicate paired samples. ns, not statistically significant, *P < 0·05.

Figure 3.

Figure 3

Open in a new tab

Enhanced induced regulatory T (iTreg) cell amplification under low O2 levels. After 3, 7 or 11 days of Treg cell induction in cultures under 5% or 21% O2, cells were collected, counted with trypan blue dye (not shown) and subjected to flow cytometry analysis. (a) Absolute number of living iTreg cells present after 3, 7 or 11 days in cultures under 5% (square) or 21% (circle) O2, and characterized as 7-AAD– CD4+ CD25+ GFP+ by flow cytometry (n = 6 or 7 independent experiments carried out in duplicate; ns, not statistically significant, *P < 0·05, **P < 0·005). (b) Percentage of cell viability determined by 7-AAD staining and flow cytometry analysis on day 3, 7 or 11 of cell cultures under 5% (square) or 21% (circle) O2. Bars represent mean cell viability of independent experiments carried out in duplicate. (c) Flow cytometric evaluation of day 11 iTreg cell proliferative capability in cultures performed under either 5% or 21% O2, using CTV-fluorescent dye and ModFit LT software. Cell proliferation analyses were performed on gated 7-AAD− CD4+ CD25+ GFP+ living iTreg cells. Histograms represent distinct iTreg cell generations, each represented by a unique peak colour, produced following 11-day culture under either 5% (right) or 21% (lefts) O2. The Proliferation Indices of iTreg cell cultures, calculated with the proliferation wizard of the ModFit software, are shown. Data are representative of two independent experiments with similar results.

ROS are essential for iTreg cell generation in vitro but are not modulated by O2 levels

T lymphocytes, expressing a non-phagocytic NADPH oxidase (NOX)39,40 can produce ROS that serve to regulate signal transduction. Constitutively formed at a low level in T cells, increased production of ROS can be observed, consecutive to a change in oxygen concentration. To determine the potential contribution of ROS in the production of Treg cells, a cell-permeable superoxide dismutase mimetic MnTBAP,41 which neutralizes intracellular ROS,42 was included in the Treg cell induction cultures performed at atmospheric O2 levels. Superoxide anion (Inline graphic ), a precursor of most other ROS, is involved in the propagation of oxidative chain reactions. Intracellular Inline graphic can be detected and quantified by staining with DHE, which becomes fluorescent upon oxidation by Inline graphic.43,44 Used at 200 μm, MnTBAP significantly reduced intracellular Inline graphic production in day 7-Treg induction-cultured cells, as shown by the decrease by roughly a half of the DHE Geometric Fluorescence Intensity (Geomean) in comparison with not-treated cultured cells (Geomean = 4209 and 8634 in MnTBAP-treated and not-treated cultured cells, respectively, Fig.5b upper panel). Neutralization of intracellular Inline graphic during the culture hampered the induction of Treg cells. The presence of 200 μm of MnTBAP in culture reduced (almost three-fold) the percentage of CD4+ CD25+ GFP+ iTreg cells at day 7 (Fig.5a, and 5b, bottom panel), irrespective of O2 levels, while the overall cell number was unchanged (data not shown). Following MnTBAP treatment, the few remaining Foxp3+ cells retained high levels of CD25, indicating that this treatment did not attenuate T-cell activation (Fig.5a). This strongly suggests that ROS signalling is important for the induction of Treg cells. However, the quantity of intracellular ROS produced under 5% or 21% O2 was similar in cultures at all time-points examined (days 3, 7 and 11) (Fig.5c). Hence, the higher Treg induction efficiency observed at 5% in comparison to 21% O2 was not related to a difference in the intracellular levels of Inline graphic.

Figure 5.

Figure 5

Open in a new tab

Intracellular reactive oxygen species (ROS) impact on the conversion of induced regulatory T (iTreg) cells but are not modulated by oxygenation. Induction of Treg cells was performed for 7 days under 21% O2, in the presence or not of the ROS scavenger Mn(III)tetrakis(4-benzoic acid) porphyrin chloride (MnTBAP; 200 μm). Cells were then collected and subjected to flow cytometric analysis to determine the level of iTreg cell conversion or of intracellular superoxide anions by dihydroethidium (DHE) staining. (a) iTreg cell conversion from CD4+ naive T cells in day 7 of culture without (upper panel) or with MnTBAP (bottom panel). Cells gated on a living CD4+ T cell population were analysed for their expression of GFP versus CD25. Inset numbers indicate the percentage of cells in each quadrant. Representative data of three independent experiments with similar results are shown. (b) Quantification of intracellular superoxide anions and of iTreg conversion in day-7 cultures done in the presence or not of MnTBAP (200 μm). Bar diagrams represent the level of intracellular superoxide anions expressed as DHE geometric mean fluorescence intensity in living 21% O2 Treg induction cultured cells (upper panel), and the percentage of CD4+ CD25+ GFP+ living iTregs under either 5% or 21% O2 (bottom panel). Data were obtained from three independent experiments. *P < 0·05, **P < 0·005. (c) Bar diagrams represent the level of intracellular superoxide anions, expressed as DHE geometric mean fluorescence intensity, in living Treg induction cultured cells at different time points performed at either 5% or 21% O2. Data were obtained from four independent experiments.

Sirt1, PTEN and Glut1 levels are modulated during Treg commitment and by oxygenation

Different proteins are involved in the process of Treg cell differentiation including PTEN, Sirt1 and Glut1. PTEN acts upstream of the Treg cell induction process as a negative regulator of IL-2 receptor signalling,4,11,12 Sirt1 acts downstream of the mechanism by having a deleterious effect on iTreg phenotype stability19–21 and Glut1 controls T-cell proliferation.8,9 Immunofluorescence microscopy analysis enables not only the determination of the expression levels of these biomarkers but also their subcellular localizations, which may affect differently their functions.9,17,20,45 At day 11, Sirt1 and PTEN expressions were found to be cytoplasmic and predominantly nuclear, while Glut1 was found only at the cell membrane (Fig.6a). No difference could be observed in subcellular distribution of these proteins depending on the level of oxygenation (data not shown). Nevertheless, the analysis of Foxp3 expressing cells revealed higher levels of Sirt1, PTEN and Glut1 compared with non-Foxp3-expressing cells, regardless of the level of oxygenation (Fig.6b). These results suggest that these markers accompany the engagement of CD4+ cells in the Treg cell differentiation pathway. However, by focusing the analysis only on Foxp3+ cells, it becomes evident that these molecules are differently regulated by oxygenation on the committed cells. While PTEN remains unchanged by oxygen levels, Sirt1 expression is reduced and Glut1 is higher on Foxp3+ cells cultured at 5% O2 compared with 21% O2 (Fig.6c). Hence, Sirt1 and Glut1, which are regulated in opposite manners by oxygenation, could play important roles in the proliferation of committed Treg cells. The higher expression of Glut1 under 5% O2 is probably linked to enhanced cell proliferation, as Glut1 is already known to promote glucose uptake and cell proliferation in various types of cells. To emphasize the role of glycolysis in the promotion of iTreg cell generation, we added the glucose analogue 2-DG to Treg cell induction cultures (Fig.7a). Inhibition of glycolysis with 1 mm of 2-DG during 7 days of naive T cells induced under 21% or 5% O2 significantly diminished CD4+ CD25+ GFP+ cell production, demonstrating that glycolysis promotes iTreg cell generation through enhancement of cell proliferation (Fig.3c and 7a). We also tested the effects of rapamycin, a well-known inhibitor of mTOR that has been shown to promote the in vitro induction of Treg cells, with suppressive properties, from human or mice effector T cells.2,15 However, inhibitory effects of rapamycin have also been reported on the growth of both non-Treg and Treg CD4+ cell populations,46 prompting us to test this agent in our system. Indeed, we observed that rapamycin treatment of cells, while having no effect on the percentage of Treg cell induction regardless of the level of oxygenation (Fig.7b), decreased significantly the total cellularity of iTreg cells at 5% and 21% O2 (Fig.7c).

Figure 6.

Figure 6

Open in a new tab

Sirt1, PTEN and Glut1 expressions in day-11 regulatory T lymphocyte (Treg) induction cultures. After 11 days of Treg-induction cultures under either 5% or 21% O2, cells were collected, stained with Sirt1, PTEN or Glut1 and Foxp3-Allophycocyanin antibodies, and subjected to fluorescence microscopy analysis as described in Materials and Methods. (a) Sirt1, PTEN and Glut1 subcellular distributions in day-11 Treg induction cultured-cells produced at 21% O2. Panels show DAPI nuclear staining (blue, left panels), Alexa fluor 594 staining for the corresponding antibody detection (red, middle panels) and merged images (right panels). Images were acquired on a LEICA SP2 confocal microscope, with a 100× oil immersion objective and zoom 2×. Scale bar, 10 μm. (b) Higher expression levels of Sirt1, PTEN and Glut1 in Foxp3+ as compared to Foxp3− cells. Immunostaining images of Sirt1, PTEN and Glut1 acquired on LEICA SP2 confocal microscope, with a 40× oil immersion objective, were analysed using CellProfiler software. Density histograms, created in R software, represent mean fluorescence intensity of Sirt1, PTEN and Glut1 detected in Foxp3+ (grey line) and Foxp3– (black line) cell populations from day 11 cultures perofrmed at either 5% (bottom panels) or 21% (upper panels) O2. Grey-shaded areas represent staining with isotype control antibodies. (c) Expression levels of Sirt1, PTEN and Glut1 in converted Foxp3+ Treg cells from day 11 cultures performed at either 5% or 21% O2. Density histograms, created in R software, represent mean fluorescence intensity of Sirt1, PTEN and Glut1 detected in Foxp3+ cells generated from day 11 cultures performed at 5% (black line) or 21% (grey line) O2. Data represent typical results from at least three independent experiments. Statistical analyses of mean intensity fluorescence curves were performed using the Kolmogorov–Smirnov test. ns, not statistically significant, ***P < 0·0005.

Figure 7.

Figure 7

Open in a new tab

Effect of modulation of glycolytic or mammalian target of rapamycin (mTOR) activities on regulatory T (Treg) cell induction. Treg cell induction cultures were performed under either 5% or 21% O2, and in the presence or not of 1 mm 2-deoxy-d-glucose (2-DG) or 50 nm rapamycin (mTOR inhibitor). Vehicle controls were performed with H2O (for 2-DG) or ethanol (for rapamycin). After 7 days of culture, cells were collected, counted with trypan blue dye, subjected to flow cytometry analysis to determine the percentage of CD25+ GFP+ gated on living (7-AAD−) CD4+ T cells (Tregs) (a, b), or the absolute number of living iTreg cells obtained at 5% (black diamonds) or 21% (grey circles) O2 in rapamycin treatment (c). Data were obtained from three to five independent experiments carried out in duplicate. *P < 0·05.

Discussion

Metabolic factors and oxygen levels are shown here to play a key role in the process of generating murine iTreg cells in culture by regulating the differentiation and amplification of naive cells. We emphasize that most published works dealing with oxygenation and iTreg cell generation document only relative and not absolute effects on Treg cell output.30,31,47,48 Our work shows for the first time that low oxygenation enhances the number of generated iTreg cells and not only their relative abundance, therefore emphasizing the importance of regulating cell growth in addition to cell differentiation in this complex process. Our data also provide a novel description of this process by combining the analysis of three biomarkers involved in T-cell differentiation under different oxygen concentration levels. The first of which, histone deacetylase Sirt1, showed up-regulated expression by the committed cells in our model, which may facilitate conformational changes in chromatin structure49 required at the onset of Treg cell specification. Yet, a critical role of Sirt-1 could not be established in the generation of iTreg cells in our experimental system. Treatment of cultures with inhibitor (sirtinol) or activator (resveratrol) of Sirt-1 had no significant effect on percentages or absolute numbers of iTRegs generated (data not shown). The effects of Sirt1 on T-cell differentiation may be complex, as published data assign to Sirt1 both a positive14,17 and a negative19–21,50 effect on Treg cell differentiation. PTEN up-regulation observed upon differentiation in our model may represent a requisite to the conversion of naïve CD4+ T cells into Treg cells through inhibition of the PI3K/Akt/mTOR signalling axis.51–53 Committed Treg cells were shown to up-regulate Sirt1 and PTEN independently of the level of oxygenation and at the initial step of culture (day 3) no clear difference in the induction of Treg cells could be observed (percentage or absolute number of induced Treg cells). Therefore oxygenation appears to play a role not in the induction but later in the higher amplification of induced Treg cells. Augmented Glut1 expression, mediated by hypoxia-inducible factor 1α, is observed in response to low O2 in several cell types54,55 and recently on CD4+ T-cell surfaces.56 Our results are in line with these studies and those indicating a beneficial effect of low O2 levels on iTreg cell generation.30,31 Indeed following differentiation, the amplification of the committed iTreg cells is unequivocally favoured by low oxygenation and by glycolysis, probably through induction of Glut1 on the cell membrane. We speculate that the induction of Glut1 on iTreg cells increases glucose uptake and causes the observed cellular proliferation, thereby enhancing the efficiency of iTreg cell generation. This effect was probably not controlled by IL-2 signalling as oxygenation levels had no impact on the intensity of CD25 expression or on the levels of PTEN. In contrast, the higher expression of Glut1 on the committed Treg cells suggests that increased glucose uptake is responsible for the beneficial effects of low oxygenation in the process of iTreg cell generation. Up-regulation of Glut1 was also observed in CD4+ Foxp3− cells, indicating that it is not specific to iTreg cells (Fig.6b). However these effector T cells do not seem to proliferate much more under low oxygenation as shown by experiments with the CTV-fluorescent dye. We obtained quite similar Proliferative Index values with analyses performed on day 11 gated CD4+ GFP– living effector T cells under 5% or 21% O2; respectively, 5·9 ± 0·3 and 4·9 ± 1·0. Enhanced Glut1 expression was recently found to increase mTOR activity through the GAPDH/Rheb axis independently of Tuberous Sclerosis Complex 2 (TSC2).57 Altogether, our results suggest that a glycolytic metabolism, concomitant with activated mTOR signalling, may contribute to a selective advantage for induced-Foxp3+ T-cell proliferation. This contrasts with other studies in which an oxidative metabolism has been proposed to be more suited to the survival and specification of Treg cells.9,13,58 Consistent with our findings, the addition of rapamycin in our system decreased the absolute number of generated iTreg cells. This is in agreement with the proposed dynamic model in which oscillating mTOR activity enables physiological adaptations to environmental changes.58–61 Hence, transient inhibition of mTOR signalling is required to overcome Treg cell anergy, but its sustained activation is necessary for these cells to proliferate.

Another important finding from our study is to identify a direct role of ROS in the Treg cell induction process. Usually considered as harmful for cells, ROS are not solely mediators of oxidative stress, but when produced in lower amounts, ROS may sustain some immune regulatory functions. Indeed, upon activation by TCR stimulation, T cells produce low levels of ROS via its NOX system.39,62 Interestingly, our results show that the scavenging of Inline graphic is profoundly detrimental to the process of Treg cell induction. The contribution of ROS to the production of iTreg cells has already been highlighted in a model of human Treg cell induction42 and recently in NOX2 knockout mice.63,64 Induction of Foxp3 in human T cells was shown by Amarnath et al. to depend on ROS, for the activation of latent endogenous TGF-β produced upon CD3/CD28 triggering. Results in NOX2 knockout mice emphasize the role of macrophages on the induction Treg cells via a ROS-dependent pathway63 as well as via a ROS-dependent decrease of Treg cell development.64 In this latter study, Lee et al. used diphenyleneiodonium (DPI), a broad spectrum flavoenzyme inhibitor, to lower the ROS level and demonstrate its critical role in controlling the development of Treg cells. However, DPI has been shown to present conflicting properties in its ability to inhibit or to stimulate the production of Inline graphic or downstream products.65–67 Therefore, data with DPI should be interpreted with caution. However, we reproduced the findings with the addition of MnTBAP in cultures, which depletes Inline graphic, and showed consequently a dramatic decrease of induction of Treg cells. Our simplified experimental model based on bead-stimulated purified T cells allows us to propose a direct role of T-cell-originating ROS in the Treg cell induction process, and not only via the production by macrophages as described previously.63 As our cultures were also supplemented with exogenous TGF-β in contrast to the human model system,42 the observed effect of MnTBAP cannot be attributed to the indirect abrogation of latent TGF-β. However, no overall change in the quantity of intracellular ROS produced at 5% or 21% O2 was detected, indicating that increased iTreg cell generation under low but physiological oxygenation is mediated independently of ROS and hence of the extent of induction process, but is done through preferential expansion of differentiated Treg cells, contributing to the increased numbers and frequencies.

In summary, our work based on cell cultures conducted under low oxygenation shows that conditions close to physiological conditions can be used to enhance the abundance of iTreg cells converted from naive T cells, not by influencing the extent of differentiation but their subsequent amplification. Hence, interactions between the metabolic and immune systems can be exploited to develop improved protocols for optimization of the production of iTreg cells. Indeed such protocols should consider the use of low O2 levels and focus on parameters that could enhance the expression of Glut1 on iTreg cells. In addition, the beneficial effects of low oxygenation on iTreg cells could have important implications in the pathophysiology of conditions involving immune response regulations.

Acknowledgments

We are very grateful to Dr David Gross for generating the Simone transgenic mice. We thank Fanny Onodi and Justine Trimouillas for technical assistance and Thibaut Viard for helpful advice on the statistical analysis.

Glossary

7-AAD

7-aminoactinomycin-D

CTV

CellTrace violet

2-DG

2-deoxy-d-glucose

DHE

dihydroethidium

DPI

diphenyleneiodonium

Foxp3

Forkhead box P3

IL-2

interleukin-2

iTreg

induced regulatory T lymphocyte

MnTBAP

Mn(III)tetrakis(4-benzoic acid) porphyrin chloride

mTOR

mammalian target of rapamycin

nTreg

natural regulatory T lymphocytes

NOX

NADPH oxidase

PI3K

phosphoinositide 3-kinase

ROS

reactive oxygen species

TCR

T-cell receptor

Teff

effector T

TGF

transforming growth factor

Treg

regulatory T lymphocyte

Author contributions

TMANN, JB and SDR performed the experiments; TMANN, GC, FB, AP and AG contributed to study design, data analysis and interpretation. TMANN and AG wrote the manuscript.

Disclosures

The authors declare no conflict of interest.

References

  1. Bilate AM, Lafaille JJ. Induced CD4+Foxp3+ regulatory T cells in immune tolerance. Annu Rev Immunol. 2012;30:733–58. doi: 10.1146/annurev-immunol-020711-075043. [DOI] [PubMed] [Google Scholar]
  2. Hippen KL, Merkel SC, Schirm DK, et al. Generation and large-scale expansion of human inducible regulatory T cells that suppress graft-versus-host disease. Am J Transplant. 2011;11:1148–57. doi: 10.1111/j.1600-6143.2011.03558.x. [DOI] [PMC free article] [PubMed] [Google Scholar]
  3. Hippen KL, Merkel SC, Schirm DK, et al. Massive ex vivo expansion of human natural regulatory T cells (Tregs) with minimal loss of in vivo functional activity. Sci Transl Med. 2011;3:83ra41. doi: 10.1126/scitranslmed.3001809. [DOI] [PMC free article] [PubMed] [Google Scholar]
  4. Dons EM, Raimondi G, Cooper DK, Thomson AW. Induced regulatory T cells: mechanisms of conversion and suppressive potential. Hum Immunol. 2012;73:328–34. doi: 10.1016/j.humimm.2011.12.011. [DOI] [PMC free article] [PubMed] [Google Scholar]
  5. Macintyre AN, Gerriets VA, Nichols AG, et al. The glucose transporter Glut1 is selectively essential for CD4 T cell activation and effector function. Cell Metab. 2014;20:61–72. doi: 10.1016/j.cmet.2014.05.004. [DOI] [PMC free article] [PubMed] [Google Scholar]
  6. Pearce EL. Metabolism in T cell activation and differentiation. Curr Opin Immunol. 2010;22:314–20. doi: 10.1016/j.coi.2010.01.018. [DOI] [PMC free article] [PubMed] [Google Scholar]
  7. Sitkovsky M, Lukashev D. Regulation of immune cells by local-tissue oxygen tension: HIF1α and adenosine receptors. Nat Rev Immunol. 2005;5:712–21. doi: 10.1038/nri1685. [DOI] [PubMed] [Google Scholar]
  8. Jacobs SR, Herman CE, Maciver NJ, Wofford JA, Wieman HL, Hammen JJ, Rathmell JC. Glucose uptake is limiting in T cell activation and requires CD28-mediated Akt-dependent and independent pathways. J Immunol. 2008;180:4476–86. doi: 10.4049/jimmunol.180.7.4476. [DOI] [PMC free article] [PubMed] [Google Scholar]
  9. MacIver NJ, Michalek RD, Rathmell JC. Metabolic regulation of T lymphocytes. Annu Rev Immunol. 2013;31:259–83. doi: 10.1146/annurev-immunol-032712-095956. [DOI] [PMC free article] [PubMed] [Google Scholar]
  10. Dure M, Macian F. IL-2 signaling prevents T cell anergy by inhibiting the expression of anergy-inducing genes. Mol Immunol. 2009;46:999–1006. doi: 10.1016/j.molimm.2008.09.029. [DOI] [PMC free article] [PubMed] [Google Scholar]
  11. Walsh PT, Buckler JL, Zhang J, et al. PTEN inhibits IL-2 receptor-mediated expansion of CD4+ CD25+ Tregs. J Clin Invest. 2006;116:2521–31. doi: 10.1172/JCI28057. [DOI] [PMC free article] [PubMed] [Google Scholar]
  12. Horwitz DA, Zheng SG, Wang J, Gray JD. Critical role of IL-2 and TGF-β in generation, function and stabilization of Foxp3+CD4+ Treg. Eur J Immunol. 2008;38:912–5. doi: 10.1002/eji.200738109. [DOI] [PubMed] [Google Scholar]
  13. Michalek RD, Gerriets VA, Jacobs SR, et al. Cutting edge: distinct glycolytic and lipid oxidative metabolic programs are essential for effector and regulatory CD4+ T cell subsets. J Immunol. 2011;186:3299–303. doi: 10.4049/jimmunol.1003613. [DOI] [PMC free article] [PubMed] [Google Scholar]
  14. Delgoffe GM, Kole TP, Zheng Y, et al. The mTOR kinase differentially regulates effector and regulatory T cell lineage commitment. Immunity. 2009;30:832–44. doi: 10.1016/j.immuni.2009.04.014. [DOI] [PMC free article] [PubMed] [Google Scholar]
  15. Gao W, Lu Y, El Essawy B, Oukka M, Kuchroo VK, Strom TB. Contrasting effects of cyclosporine and rapamycin in de novo generation of alloantigen-specific regulatory T cells. Am J Transplant. 2007;7:1722–32. doi: 10.1111/j.1600-6143.2007.01842.x. [DOI] [PMC free article] [PubMed] [Google Scholar]
  16. Peter C, Waldmann H, Cobbold SP. mTOR signalling and metabolic regulation of T cell differentiation. Curr Opin Immunol. 2010;22:655–61. doi: 10.1016/j.coi.2010.08.010. [DOI] [PubMed] [Google Scholar]
  17. Ghosh HS, McBurney M, Robbins PD. SIRT1 negatively regulates the mammalian target of rapamycin. PLoS ONE. 2010;5:e9199. doi: 10.1371/journal.pone.0009199. [DOI] [PMC free article] [PubMed] [Google Scholar]
  18. Salminen A, Kaarniranta K, Kauppinen A. Crosstalk between oxidative stress and SIRT1: impact on the aging process. Int J Mol Sci. 2013;14:3834–59. doi: 10.3390/ijms14023834. [DOI] [PMC free article] [PubMed] [Google Scholar]
  19. Kwon HS, Lim HW, Wu J, Schnolzer M, Verdin E, Ott M. Three novel acetylation sites in the Foxp3 transcription factor regulate the suppressive activity of regulatory T cells. J Immunol. 2012;188:2712–21. doi: 10.4049/jimmunol.1100903. [DOI] [PMC free article] [PubMed] [Google Scholar]
  20. van Loosdregt J, Brunen D, Fleskens V, Pals CE, Lam EW, Coffer PJ. Rapid temporal control of Foxp3 protein degradation by sirtuin-1. PLoS ONE. 2011;6:e19047. doi: 10.1371/journal.pone.0019047. [DOI] [PMC free article] [PubMed] [Google Scholar]
  21. van Loosdregt J, Vercoulen Y, Guichelaar T, et al. Regulation of Treg functionality by acetylation-mediated Foxp3 protein stabilization. Blood. 2010;115:965–74. doi: 10.1182/blood-2009-02-207118. [DOI] [PubMed] [Google Scholar]
  22. Atkuri KR, Herzenberg LA, Herzenberg LA. Culturing at atmospheric oxygen levels impacts lymphocyte function. Proc Natl Acad Sci U S A. 2005;102:3756–9. doi: 10.1073/pnas.0409910102. [DOI] [PMC free article] [PubMed] [Google Scholar]
  23. Atkuri KR, Herzenberg LA, Niemi AK, Cowan T, Herzenberg LA. Importance of culturing primary lymphocytes at physiological oxygen levels. Proc Natl Acad Sci U S A. 2007;104:4547–52. doi: 10.1073/pnas.0611732104. [DOI] [PMC free article] [PubMed] [Google Scholar]
  24. Caldwell CC, Kojima H, Lukashev D, Armstrong J, Farber M, Apasov SG, Sitkovsky MV. Differential effects of physiologically relevant hypoxic conditions on T lymphocyte development and effector functions. J Immunol. 2001;167:6140–9. doi: 10.4049/jimmunol.167.11.6140. [DOI] [PubMed] [Google Scholar]
  25. Carswell KS, Weiss JW, Papoutsakis ET. Low oxygen tension enhances the stimulation and proliferation of human T lymphocytes in the presence of IL-2. Cytotherapy. 2000;2:25–37. doi: 10.1080/146532400539026. [DOI] [PubMed] [Google Scholar]
  26. Ohta A, Madasu M, Subramanian M, Kini R, Jones G, Chouker A, Sitkovsky M. Hypoxia-induced and A2A adenosine receptor-independent T-cell suppression is short lived and easily reversible. Int Immunol. 2014;26:83–91. doi: 10.1093/intimm/dxt045. [DOI] [PMC free article] [PubMed] [Google Scholar]
  27. Ehrentraut H, Westrich JA, Eltzschig HK, Clambey ET. Adora2b adenosine receptor engagement enhances regulatory T cell abundance during endotoxin-induced pulmonary inflammation. PLoS ONE. 2012;7:e32416. doi: 10.1371/journal.pone.0032416. [DOI] [PMC free article] [PubMed] [Google Scholar]
  28. Ohta A, Kini R, Subramanian M, Madasu M, Sitkovsky M. The development and immunosuppressive functions of CD4+ CD25+ FoxP3+ regulatory T cells are under influence of the adenosine-A2A adenosine receptor pathway. Front Immunol. 2012;3:190. doi: 10.3389/fimmu.2012.00190. [DOI] [PMC free article] [PubMed] [Google Scholar]
  29. Ohta A, Sitkovsky M. Extracellular adenosine-mediated modulation of regulatory T cells. Front Immunol. 2014;5:304. doi: 10.3389/fimmu.2014.00304. [DOI] [PMC free article] [PubMed] [Google Scholar]
  30. Ben-Shoshan J, Maysel-Auslender S, Mor A, Keren G, George J. Hypoxia controls CD4+CD25+ regulatory T-cell homeostasis via hypoxia-inducible factor-1α. Eur J Immunol. 2008;38:2412–8. doi: 10.1002/eji.200838318. [DOI] [PubMed] [Google Scholar]
  31. Clambey ET, McNamee EN, Westrich JA, et al. Hypoxia-inducible factor-1α-dependent induction of FoxP3 drives regulatory T-cell abundance and function during inflammatory hypoxia of the mucosa. Proc Natl Acad Sci U S A. 2012;109:E2784–93. doi: 10.1073/pnas.1202366109. [DOI] [PMC free article] [PubMed] [Google Scholar]
  32. Dang EV, Barbi J, Yang HY, et al. Control of TH17/Treg balance by hypoxia-inducible factor 1. Cell. 2011;146:772–84. doi: 10.1016/j.cell.2011.07.033. [DOI] [PMC free article] [PubMed] [Google Scholar]
  33. Shi LZ, Wang R, Huang G, Vogel P, Neale G, Green DR, Chi H. HIF1α-dependent glycolytic pathway orchestrates a metabolic checkpoint for the differentiation of TH17 and Treg cells. J Exp Med. 2011;208:1367–76. doi: 10.1084/jem.20110278. [DOI] [PMC free article] [PubMed] [Google Scholar]
  34. Wang Y, Kissenpfennig A, Mingueneau M, et al. Th2 lymphoproliferative disorder of LatY136F mutant mice unfolds independently of TCR-MHC engagement and is insensitive to the action of Foxp3+ regulatory T cells. J Immunol. 2008;180:1565–75. doi: 10.4049/jimmunol.180.3.1565. [DOI] [PubMed] [Google Scholar]
  35. Lantz O, Grandjean I, Matzinger P, Di Santo JP. Gamma chain required for naive CD4+ T cell survival but not for antigen proliferation. Nat Immunol. 2000;1:54–8. doi: 10.1038/76917. [DOI] [PubMed] [Google Scholar]
  36. Brock R, Hamelers IH, Jovin TM. Comparison of fixation protocols for adherent cultured cells applied to a GFP fusion protein of the epidermal growth factor receptor. Cytometry. 1999;35:353–62. doi: 10.1002/(sici)1097-0320(19990401)35:4<353::aid-cyto8>3.0.co;2-m. [DOI] [PubMed] [Google Scholar]
  37. Carpenter AE, Jones TR, Lamprecht MR, et al. Cell Profiler: image analysis software for identifying and quantifying cell phenotypes. Genome Biol. 2006;7:R100. doi: 10.1186/gb-2006-7-10-r100. [DOI] [PMC free article] [PubMed] [Google Scholar]
  38. Fantini MC, Dominitzki S, Rizzo A, Neurath MF, Becker C. In vitro generation of CD4+ CD25+ regulatory cells from murine naive T cells. Nat Protoc. 2007;2:1789–94. doi: 10.1038/nprot.2007.258. [DOI] [PubMed] [Google Scholar]
  39. Jackson SH, Devadas S, Kwon J, Pinto LA, Williams MS. T cells express a phagocyte-type NADPH oxidase that is activated after T cell receptor stimulation. Nat Immunol. 2004;5:818–27. doi: 10.1038/ni1096. [DOI] [PubMed] [Google Scholar]
  40. Kwon J, Shatynski KE, Chen H, Morand S, de Deken X, Miot F, Leto TL, Williams MS. The nonphagocytic NADPH oxidase Duox1 mediates a positive feedback loop during T cell receptor signaling. Sci Signal. 2010;3:ra59. doi: 10.1126/scisignal.2000976. [DOI] [PMC free article] [PubMed] [Google Scholar]
  41. Patel M, Day BJ. Metalloporphyrin class of therapeutic catalytic antioxidants. Trends Pharmacol Sci. 1999;20:359–64. doi: 10.1016/s0165-6147(99)01336-x. [DOI] [PubMed] [Google Scholar]
  42. Amarnath S, Dong L, Li J, Wu Y, Chen W. Endogenous TGF-β activation by reactive oxygen species is key to Foxp3 induction in TCR-stimulated and HIV-1-infected human CD4+CD25– T cells. Retrovirology. 2007;4:57. doi: 10.1186/1742-4690-4-57. [DOI] [PMC free article] [PubMed] [Google Scholar]
  43. Dikalov S, Griendling KK, Harrison DG. Measurement of reactive oxygen species in cardiovascular studies. Hypertension. 2007;49:717–27. doi: 10.1161/01.HYP.0000258594.87211.6b. [DOI] [PMC free article] [PubMed] [Google Scholar]
  44. Zielonka J, Vasquez-Vivar J, Kalyanaraman B. Detection of 2-hydroxyethidium in cellular systems: a unique marker product of superoxide and hydroethidine. Nat Protoc. 2008;3:8–21. doi: 10.1038/nprot.2007.473. [DOI] [PubMed] [Google Scholar]
  45. Chang CJ, Mulholland DJ, Valamehr B, Mosessian S, Sellers WR, Wu H. PTEN nuclear localization is regulated by oxidative stress and mediates p53-dependent tumor suppression. Mol Cell Biol. 2008;28:3281–9. doi: 10.1128/MCB.00310-08. [DOI] [PMC free article] [PubMed] [Google Scholar]
  46. Valmori D, Tosello V, Souleimanian NE, Godefroy E, Scotto L, Wang Y, Ayyoub M. Rapamycin-mediated enrichment of T cells with regulatory activity in stimulated CD4+ T cell cultures is not due to the selective expansion of naturally occurring regulatory T cells but to the induction of regulatory functions in conventional CD4+ T cells. J Immunol. 2006;177:944–9. doi: 10.4049/jimmunol.177.2.944. [DOI] [PubMed] [Google Scholar]
  47. Deng B, Zhu JM, Wang Y, et al. Intratumor hypoxia promotes immune tolerance by inducing regulatory T cells via TGF-β1 in gastric cancer. PLoS ONE. 2013;8:e63777. doi: 10.1371/journal.pone.0063777. [DOI] [PMC free article] [PubMed] [Google Scholar]
  48. Wei J, Wu A, Kong LY, et al. Hypoxia potentiates glioma-mediated immunosuppression. PLoS ONE. 2011;6:e16195. doi: 10.1371/journal.pone.0016195. [DOI] [PMC free article] [PubMed] [Google Scholar]
  49. Vaquero A, Scher M, Lee D, Erdjument-Bromage H, Tempst P, Reinberg D. Human SirT1 interacts with histone H1 and promotes formation of facultative heterochromatin. Mol Cell. 2004;16:93–105. doi: 10.1016/j.molcel.2004.08.031. [DOI] [PubMed] [Google Scholar]
  50. Akimova T, Xiao H, Liu Y, et al. Targeting sirtuin-1 alleviates experimental autoimmune colitis by induction of Foxp3 T-regulatory cells. Mucosal Immunol. 2014;7:1209–20. doi: 10.1038/mi.2014.10. [DOI] [PMC free article] [PubMed] [Google Scholar]
  51. Haxhinasto S, Mathis D, Benoist C. The AKT-mTOR axis regulates de novo differentiation of CD4+Foxp3+ cells. J Exp Med. 2008;205:565–74. doi: 10.1084/jem.20071477. [DOI] [PMC free article] [PubMed] [Google Scholar]
  52. Sauer S, Bruno L, Hertweck A, et al. T cell receptor signaling controls Foxp3 expression via PI3K, Akt, and mTOR. Proc Natl Acad Sci U S A. 2008;105:7797–802. doi: 10.1073/pnas.0800928105. [DOI] [PMC free article] [PubMed] [Google Scholar]
  53. Weichhart T, Saemann MD. The multiple facets of mTOR in immunity. Trends Immunol. 2009;30:218–26. doi: 10.1016/j.it.2009.02.002. [DOI] [PubMed] [Google Scholar]
  54. Bashan N, Burdett E, Hundal HS, Klip A. Regulation of glucose transport and GLUT1 glucose transporter expression by O2 in muscle cells in culture. Am J Physiol. 1992;262:C682–90. doi: 10.1152/ajpcell.1992.262.3.C682. [DOI] [PubMed] [Google Scholar]
  55. Chen C, Pore N, Behrooz A, Ismail-Beigi F, Maity A. Regulation of glut1 mRNA by hypoxia-inducible factor-1. Interaction between H-ras and hypoxia. J Biol Chem. 2001;276:9519–25. doi: 10.1074/jbc.M010144200. [DOI] [PubMed] [Google Scholar]
  56. Loisel-Meyer S, Swainson L, Craveiro M, et al. Glut1-mediated glucose transport regulates HIV infection. Proc Natl Acad Sci U S A. 2012;109:2549–54. doi: 10.1073/pnas.1121427109. [DOI] [PMC free article] [PubMed] [Google Scholar]
  57. Buller CL, Heilig CW, Brosius FC., 3rd GLUT1 enhances mTOR activity independently of TSC2 and AMPK. Am J Physiol Renal Physiol. 2011;301:F588–96. doi: 10.1152/ajprenal.00472.2010. [DOI] [PMC free article] [PubMed] [Google Scholar]
  58. Zeng H, Chi H. The interplay between regulatory T cells and metabolism in immune regulation. Oncoimmunology. 2013;2:e26586. doi: 10.4161/onci.26586. [DOI] [PMC free article] [PubMed] [Google Scholar]
  59. Cobbold SP. The mTOR pathway and integrating immune regulation. Immunology. 2013;140:391–8. doi: 10.1111/imm.12162. [DOI] [PMC free article] [PubMed] [Google Scholar]
  60. Procaccini C, De Rosa V, Galgani M, et al. An oscillatory switch in mTOR kinase activity sets regulatory T cell responsiveness. Immunity. 2010;33:929–41. doi: 10.1016/j.immuni.2010.11.024. [DOI] [PMC free article] [PubMed] [Google Scholar]
  61. Procaccini C, Matarese G. Regulatory T cells, mTOR kinase, and metabolic activity. Cell Mol Life Sci. 2012;69:3975–87. doi: 10.1007/s00018-012-1058-6. [DOI] [PMC free article] [PubMed] [Google Scholar]
  62. Lambeth JD. NOX enzymes and the biology of reactive oxygen. Nat Rev Immunol. 2004;4:181–9. doi: 10.1038/nri1312. [DOI] [PubMed] [Google Scholar]
  63. Kraaij MD, Savage ND, van der Kooij SW, et al. Induction of regulatory T cells by macrophages is dependent on production of reactive oxygen species. Proc Natl Acad Sci U S A. 2010;107:17686–91. doi: 10.1073/pnas.1012016107. [DOI] [PMC free article] [PubMed] [Google Scholar]
  64. Lee K, Won HY, Bae MA, Hong JH, Hwang ES. Spontaneous and aging-dependent development of arthritis in NADPH oxidase 2 deficiency through altered differentiation of CD11b+ and Th/Treg cells. Proc Natl Acad Sci U S A. 2011;108:9548–53. doi: 10.1073/pnas.1012645108. [DOI] [PMC free article] [PubMed] [Google Scholar]
  65. Balcerczyk A, Soszynski M, Rybaczek D, Przygodzki T, Karowicz-Bilinska A, Maszewski J, Bartosz G. Induction of apoptosis and modulation of production of reactive oxygen species in human endothelial cells by diphenyleneiodonium. Biochem Pharmacol. 2005;69:1263–73. doi: 10.1016/j.bcp.2005.01.010. [DOI] [PubMed] [Google Scholar]
  66. Li N, Ragheb K, Lawler G, Sturgis J, Rajwa B, Melendez JA, Robinson JP. DPI induces mitochondrial superoxide-mediated apoptosis. Free Radic Biol Med. 2003;34:465–77. doi: 10.1016/s0891-5849(02)01325-4. [DOI] [PubMed] [Google Scholar]
  67. Riganti C, Gazzano E, Polimeni M, Costamagna C, Bosia A, Ghigo D. Diphenyleneiodonium inhibits the cell redox metabolism and induces oxidative stress. J Biol Chem. 2004;279:47726–31. doi: 10.1074/jbc.M406314200. [DOI] [PubMed] [Google Scholar]

Articles from Immunology are provided here courtesy of British Society for Immunology

RESOURCES