View larger version (48K): [in this window] [in a new window] [Download PPT slide]   Figure 2. RA–T reg cells are suppressive in vitro and home to the small intestine in vivo. To examine the suppressive capacity of A-Tregs, RA–T reg cells, and nTregs, Ly5.2+Foxp3/GFP+ A-Tregs and RA–T reg cells were generated in vitro and sorted to >99% purity, with Ly5.2+ nTregs freshly isolated ex vivo from a Ly5.2+Foxp3/GFP reporter mouse. These T reg cell subsets were cultured with CFSE-labeled Ly5.1+CD4+ T effector cells at the indicated ratios under activating conditions. Representative CFSE plots of the T effector cells are shown (A), with the percentage of dividing T effector cells quantified (B). (C and D) Representative in vivo competitive homing assay. Naive CD4+Foxp3– cells were sorted from an Ly5.2+Foxp3/GFP reporter mouse and activated for 5 d in the presence of TGF-ß1, IL-2, and RA to generate RA–T reg cells expressing Foxp3/GFP. Naive CD4+CD25– cells were sorted from an Ly5.2+ mouse and cultured under the same conditions, minus the RA, to generate A-Tregs lacking the Foxp3/GFP allele. These two cell populations were mixed and injected into recipient mice. (C) Organs were harvested after 18 h and stained for donor Ly5.2+CD4+ T cells, and the ratio of GFP+/– cells within the donor population was analyzed, with a representative mouse shown. Numbers show the percentage of transfered cells that are FoxP3/GFP– and FoxP3/GFP+. (D) The HI (ration of [GFP+]tissue/[GFP–]tissue to [GFP+]input/[GFP–]input) was calculated, with bar graphs showing mean ± SEM of one representative experiment using three recipient mice. Peripheral lymph node (pLN): HI = 0.96 (SEM = 0.13); spleen: HI = 0.96 (SEM = 0.08); blood: HI = 0.56 (SEM = 0.05); lung: HI = 0.39 (SEM = 0.02); mesenteric lymph node (mLN): HI = 1.86 (SEM = 0.1); Peyer's patches (PP): HI = 3.46 (SEM = 0.33); and lamina propria (LP): HI = 12.27 (SEM = 0.77). All experiments are representative of at least two independent experiments. *, P < 0.05; **, P < 0.01; and ***, P < 0.001 compared with HI = 1 (dotted line).

Co-stimulation and A-T reg cell generation: RA allows DCs to drive TGF-ß1–mediated FoxP3 induction
To investigate the capacity of different APC subsets to induce A-Treg generation in vitro, peptide-pulsed splenic CD19⁺ B cells and CD11c⁺ DCs were analyzed for their ability to drive TGF-ß1–dependent FoxP3 induction in CD4⁺OTII⁺FoxP3^– cells harvested and purified by sorting from the OTII⁺FoxP3/GFP mouse. A difference was observed between these two APC subsets to mediate conversion: B cells repeatedly induced conversion at rates between 40–60%, with DCs inducing conversion at substantially lower rates of 0–14% (Fig. 3 A; and Fig. S1, available at http://www.jem.org/cgi/content/full/jem.20070719/DC1), consistent with a previous study (25). The ability of B cells and DCs to mediate conversion was further analyzed by titrating exogenous TGF-ß1, further sub stantiating the differential capacity of these APC subsets to medi ate T reg cell generation (Fig. S1).

View larger version (29K): [in this window] [in a new window] [Download PPT slide]   Figure 3. DC expression of CD80/86 co-stimulatory molecules prevents Foxp3 induction by T cells, with exogenous RA overcoming this inhibition and allowing T reg cell generation. Sorted CD4+OTII+Foxp3– T cells from the OTII+Foxp3/GFP reporter mouse (100,000 T cells/well) were cultured with either purified WT or CD80/86 knockout CD19+ B cells or CD11c+ DCs (100,000 APCs/well) pulsed with ISQ peptide, or under activating conditions. Media was supplemented with 100 U IL-2, ±20 ng/ml TGF-ß1, ±1 nM RA, ±10 µg/ml CD154 (unless otherwise indicated) for 5 d. (A) WT or CD80/86 knockout splenic B cells or DCs plus T cells were cultured under the depicted culture conditions, and the percentage of CD4+ T cells expressing Foxp3/GFP was measured. (B) Same experimental conditions as in A, except with absolute CD4+ T cells per well counted, with white bars representing CD4+Foxp3– cells and green bars representing CD4+Foxp3+ cells. (C) Freshly harvested B cells (quiescent B cells) or 48-h preactivated B cells with the indicated stimulus were co-cultured with T cells, and the percentage of CD4+ cells expressing Foxp3/GFP was measured after 5 d. (D) CD4+ T cells were activated using 10 µg/ml of plate-bound CD3 and titrating concentrations of CD28, with media supplemented with IL-2 and TGF-ß1 ±10 nM RA as indicated; Foxp3/GFP expression was plotted as a function of plate-bound CD28 concentration. Mean ± SEM is depicted in A, B, and D.

The in vitro and in vivo activation of both B cells and DCs through CD40 engagement by its ligand, CD154, increases the expression of CD80/86 by these APCs. The inclusion of blocking CD154 antibody to our co-cultures enabled DCs to have an increased ability to mediate conversion (7.94% [SD = 1.14] compared with 15.26% [SD = 0.55] with CD154; Fig. 3 A). Similar results were observed using CD40 knockout DCs (unpublished data). Interestingly, CD154 antibody appeared to decrease B cell–mediated conversion, possibly implying a requirement for very low levels of co-stimulation in this co-culture system (63% [SD = 7.81] compared with 39.33% [SD = 4.5] with CD154).

We next addressed whether B cell activation and the acquisition of heightened co-stimulatory molecule expression impaired the ability to induce conversion. When B cells were preactivated for 48 h by either LPS or agonistic-CD40 supplemented with IL-4 and then used as APCs, B cell–mediated conversion was impaired (Fig. 3 C).

When CD4⁺ T cells were activated using WT or CD80/86 knockout DCs in the presence of exogenous IL-2, considerably more T cells per well were generated by WT DCs in comparison with CD80/86 knockout DCs (19.64 x 10⁴ cells [SD = 2.42 x 10⁴] compared with 9.13 x 10⁴ cells [SD = 2.48 x 10⁴]; Fig. 3 B). The addition of RA or TGF-ß1 or both did not affect this observed difference (Fig. 3 B). Upon the addition of TGF-ß1 to these cultures, the decrease of T cell numbers between WT and CD80/86 knockout DCs inversely correlated with a substantial increase in the number of FoxP3-expressing T cells in the CD80/86 knockout group versus the WT group (6.35 x 10⁴ cells [SD = 0.15 x 10⁴] compared with 1.33 x 10⁴ cells [SD = 0.25 x 10⁴]; Fig. 3 B).

The addition of CD154 did not lead to a considerable change in T cell numbers generated by DCs with the addition of exogenous IL-2 (19.64 x 10⁴ [SD = 2.42 x 10⁴] compared with 11.13 x 10⁴ [SD = 7.75 x 10⁴] with CD154; Fig. 3 B). No difference in T cell number was again observed with the further addition of RA or TGF-ß1, or both. Additionally, there was no considerable increase in the numbers of FoxP3-expressing cells because of the inclusion of CD154 to our DC co-cultures supplemented with TGF-ß1 (1.33 x 10⁴ [SD = 0.25 x 10⁴] compared with 1.56 x 10⁴ [SD = 0.36 x 10⁴]; Fig. 3 B). Despite the enhanced conversion imparted by CD154 in our DC/T cell co-cultures in Fig. 3 A, this result was not entirely unexpected. First, even with a CD154/CD40 blockade, DCs will still express levels of CD80/86 that are sufficient in expanding T cells. Second, the increase in DC-mediated conversion caused by CD154 may not be enough to net an absolute increase in T reg cell numbers in this experimental system.

B cells, CD80/86 knockout B cells, and B cells cultured with CD154 all generated similar total numbers of T cells in all culture conditions. This was expected and is likely caused by the low levels of co-stimulatory molecule expression on the surface of resting B cells (26). Collectively, these data support our hypothesis that under conditions favoring conversion, the greater the absolute yield of T cells generated by an APC and the fewer the number of T cells expressing FoxP3. Further substantiating this is our finding that increasing levels of plate-bound co-stimulation inversely correlates with the absolute number of A-Tregs generated (Fig. S2, available at http://www.jem.org/cgi/content/full/jem.20070719/DC1).

To directly implicate CD28 as a major mediator affecting FoxP3 induction, we varied the magnitude of CD28 co-stimulation in an in vitro assay in which plate-bound agonistic CD28 was titrated against saturating concentrations of plate-bound CD3. It was observed that CD28 concentration had an inverse relationship with A-Treg generation, with optimal conversion occurring at CD28 concentrations of 1 µg/ml and below. At saturating concentrations (10 µg/ml CD28), no conversion was observed (Fig. 3 D). Collectively, these data show that high levels of CD80/86 co-stimulation, such as those observed on splenic CD11c⁺ DCs, impairs TGF-ß1–driven FoxP3 induction and A-Treg generation while inducing maximal expansion of T cells. In the context of minimal to no CD80/86 co-stimulation, such as levels present on resting B cells and CD80/86 knockout DCs, little to no expansion occurs, with TGF-ß1–dependent FoxP3 expression and T reg cell generation the result.

The addition of RA to our assays greatly enhanced both the percentage and total numbers of T cells expressing FoxP3 in a TGF-ß1–dependent manner (Fig. 3, A and B; Fig. S1; and Fig. S2). Notably, the addition of RA to our DC/T cell co-cultures increased the percentage of T cells expressing FoxP3 from 7.94% (SD = 1.14) to 72% (SD = 2; Fig. 3 A). Additionally, the combination of RA with TGF-ß1 resulted in a net increase in the numbers of FoxP3-expressing cells in all DC cultures in comparison with TGF-ß1 alone (WT DCs: from 1.33 x 10⁴ [SD = 0.25 x 10⁴] to 9.96 x 10⁴ [SD = 0.62 x 10⁴]; CD80/86 knockout DCs: from 6.35 x 10⁴ [SD = 0.15 x 10⁴] to 8.7 x 10⁴ [SD = 0.19 x 10⁴]; DCs with CD154: from 1.56 x 10⁴ [SD = 0.36 x 10⁴] to 9.9 x 10⁴ [SD = 0.19 x 10⁴]; Fig. 3 B and Fig. S1). Furthermore, RA reversed co-stimulation from suppressing, in dose-dependent manner, the net yield of FoxP3⁺ T cells, as concentrations of plate-bound CD28 now positively correlate with FoxP3⁺ T cell numbers (Fig. S2). Across all concentrations of plate-bound CD28, as well as in the presence of only a TCR signal with no CD28, RA greatly enhanced the percentage of T cells expressing FoxP3 (Fig. 3 D). In this experiment, co-stimulation still affected FoxP3 induction in a dose-dependent manner, albeit to a substantially attenuated degree, as a result of the inclusion of RA. This suggests that RA lowers an as yet undefined threshold within the T cell, with this threshold determining whether the T cell can convert into a T reg cell. Thus, the presence of RA during antigen presentation allows TGF-ß1–dependent FoxP3 induction to occur by T cells, even in the face of overt CD80/86 co-stimulation afforded by the DC.

RA enhances FoxP3 expression without impeding T cell activation and proliferation
The observation that RA–T reg cells can be induced in the presence of high levels of co-stimulation led us the question whether this occurs by RA functioning to interrupt T cell activation and proliferation. To visualize this, CFSE-labeled CD4⁺CD25^– cells were stimulated in the presence and absence of 1 nM RA, with T cell activation determined by CD62L expression and proliferation by CFSE dye dilution. The data shows that 1 nM RA does not suppress T cell activation, as indicated by CD62L expression. Instead, RA appears to enhance T cell activation, as indicated by the decreased expression of CD62L across the three levels of activating conditions tested (Fig. 4 A). Additionally, when CFSE histograms between different culture conditions are overlaid, equivalent profiles indicate that the addition of RA does not impede T cell proliferation (Fig. 4 B). The addition of TGF-ß1 to these cultures generated FoxP3⁺ cells at expected frequencies throughout all peaks of cell division, with no impediment in cell division observed because of the presence of RA (Fig. 4 C). Importantly, RA enhances TGF-ß1–mediated FoxP3 expression in the absence of any CD28 signal and just the presence of CD3 (Fig. 4 C and Fig. 3 D). Collectively, these data show that RA enhances FoxP3 induction and attenuates co-stimulation from interfering with T reg cell generation through a mechanism independent of dampening T cell activation and proliferation. Fur thermore, co-stimulation is not needed for RA to enhance FoxP3 induction.

View larger version (30K): [in this window] [in a new window] [Download PPT slide]   Figure 4. RA enhances Foxp3 expression without impeding T cell activation and proliferation. Purified CFSE-labeled CD4+CD25– T cells from a WT B6 mouse were cultured with IL-2, TGF-ß1, and 1 nM RA, as indicated, under activating conditions for 4 d, after which CD62L and Foxp3 expression was measured. (A) CFSE dye dilution versus CD62L expression is shown, with T cells cultured with IL-2 or IL-2 plus 1 nM RA under different activating conditions. The percentage of T cells within each quadrant is depicted. (B) Histograms depicting CFSE dye dilution profiles of the data shown in A (IL-2, blue; IL-2 with 1 nM RA, red). (C) CFSE dye dilution versus Foxp3 expression is shown with the indicated activating and culture conditions, with the percentage of cells in each quadrant shown. Representative experiments of at least n = 3 mice are shown.

View larger version (13K): [in this window] [in a new window] [Download PPT slide]   Figure 5. RA–T reg cells are refractory to reversion in vivo. The ability of RA–T reg cells and A-Tregs to revert in vivo was analyzed by transferring 1.5–2 x 106 FACS-sorted congenically marked OTII+ RA–T reg cells or A-Tregs generated from a Ly5.2+OTII+Foxp3/GFP mice into Ly5.1+ hosts, who were either left untouched or intraperitoneally injected on the same day with either CFA or 1 mg OVA/CFA, as indicated. Donor RA–T reg cells or A-Tregs within the spleen were analyzed for Foxp3 expression on day 5 by staining for CD4+Ly5.2+ cells. RA–T reg cells are red, whereas A-Tregs are black (A). Horizontal lines represent the means. To examine relative donor T cell expansion 5 d after transfer, the percentage of donor OTII+ cells within the recipient splenic CD4+ compartment is shown in a box and whisker plot depicting the median, 25th, and 75th percentiles and range of values (B). Data are pooled from n = 3 independent experiments. ***, P < 0.001.

The implications of these data are important. First, Powrie et al. and Agace et al. jointly described CD103⁺CD11c⁺ DCs present within the mesenteric lymph nodes, Peyer's Patches, and small and large bowel lamina propria as composing the DC subset responsible for mediating gut imprinting of T cells through CCR9 and 4ß7 induction. Second, these CD103⁺ DCs likely imprint through an RA-dependent mechanism, although this has not yet been directly shown (9, 10). Therefore, our data suggest a new role for RA in mediating not only gut imprinting but also in driving gut tolerance through RA–T reg cell induction. Finally, with the use of A-Tregs in cellular adoptive therapies on the horizon, the use of RA in vitro to facilitate the production of fully committed, gut-homing A-Tregs is an attractive technology to meet the needs for high numbers of these cells in the therapeutic treatment of autoimmune disorders such as inflammatory bowel disease.

	MATERIALS AND METHODS

Top ABSTRACT RESULTS AND DISCUSSION MATERIALS AND METHODS REFERENCES

 
Mice and immunizations.
This study was approved by the Institutional Animal Care and Use Committee of Dartmouth College. C57BL/6 and C57BL/6 CD45.1 (Ly5.2⁺) mice were purchased from the National Cancer Institute. C56BL/6 CD40^–/– mice were purchased from the Jackson Laboratory. FoxP3/GFP reporter mice were previously described and were provided by A. Rudensky (University of Washington School of Medicine, Seattle, WA) (20). FoxP3/GFP mice were bred onto OTII TCR-Tg mice purchased from the Jackson Laboratory. CD80/86 double knockout mice on the C57BL/6 background were a gift from E. Usherwood (Dartmouth Medical School, Lebanon, NH). All animals were maintained in a pathogen-free facility at Dartmouth Medical School. For immunizations, OVA (Sigma-Aldrich) resuspended in PBS or PBS alone was emulsified in CFA (Sigma-Aldrich) at a 1:1 ratio and intraperitoneally injected in a volume of 200 µl.

Cell preparation.
B cells and DCs were harvested from the spleens, and CD4⁺FoxP3^– cells were isolated from spleens and peripheral and mesenteric lymph nodes. For B cell and T cells, single-cell suspensions were generated by crushing organs by sterile slides and were purified by positive selection using either CD4- or CD19-labeled magnetic beads (Miltenyi Biotec). T cells were further purified by FACS sorting (FACSAria; BD Biosciences) of the CD4FoxP3/GFP^– fraction, with purity always >99.9%. For CD11c⁺ selection, spleens were harvested and incubated at 37°C in RPMI 1640 with 50 µg/ml DNase I (Sigma-Aldrich) and 250 µg/ml Liberase (Roche) for 30 min, pushed through a 100-µM filter to create a single-cell suspension, and purified using CD11c magnetic beads (Miltenyi Biotec). Cell preparations were always >98% in purity. For small intestine lamina propria lymphocyte preparations, intestines were removed, and Peyer's Patches were excised and used for further analysis. The intestines were washed with cold PBS, split open, and cut into 1-cm pieces. After 30 min of incubation in unsupplemented RPMI to release the intestinal epithelial lymphocytes, intestines were vortexed, filtered using a 100-µM filter, and extensively washed. Intestines were digested for 2 h using 50 µg/ml DNase I and 250 µg/ml Liberase, after which they were pushed through a 100-µM filter. The cellular suspension was centrifuged and suspended in 40% Percoll, and overlaid on 60% Percoll. The Percoll gradient was spun at 400 g at room temperature for 25 min with no brake, with the buffy (lymphocyte) coat removed for further use. As indicated in the figures, APCs were pulsed with ISQ peptide at 10 µg/ml for 1 h in cRPMI in sixwell plates at a concentration of 10 x 10⁶ cells/ml, and were washed, counted, and used.

Cell culture/reagents.
Cells were cultured in RPMI 1640 media supplemented with 10% FBS (Atlanta Biologicals), Hepes, 50 µM ß-ME, and penicillin/streptomycin/L-glutamine. LPS was purchased from Sigma-Aldrich. CD40 (clone FGK-45), CD154 (clone MR1), CD28 (clone PV-1), and CD3 (clone 2C11) were purchased from ISC BioExpress. LE540 was purchased from Wako Chemicals. For 96-well plate cultures, 200,000 cells in APC/T cell co-cultures (1:1 ratio) in round-bottom plates or 100,000 T cells in flat-bottom plates were cultured in 200 µl of media. In each experiment, triplicate wells of each experimental condition were set up. For the bulk RA–T reg cell and A-Treg cultures in Fig. 2 and Fig. 5, 24-well flat-well plates were used with 100,000 CD4⁺FoxP3^– cells/well in 1 ml of media. For all cultures, unless indicated otherwise in the figures, T cells were activated with 1 µg/ml CD28 and 10 µg/ml CD3 plate-bound antibody in the presence of 20 ng/ml hTGF-ß1 (PeproTech), 100 U hIL-2 (PeproTech), and RA (Sigma-Aldrich). For the in vitro suppressor assay, 50,000 CFSE-labeled CD4⁺ T cells were co-cultured with 100,000 irradiated T cell–depleted splenocytes, 5 µg/ml CD3, and the numbers of T reg cells indicated in the figures for 4 d, as previously described (27, 28). For Fig. 3 and Fig. S2, cells were counted by Guava according to instructions provided by the manufacturer (Guava Technologies)

Flow cytometry.
The following antibodies were used: CD11c (clone N418), CD25 (clone PC61), and CD62L (clone MEL-14; all obtained from BioLegend); B220 (clone 6B2), CD4 (clone L3T4), and ₄ß₇ (clone DATK32; all obtained from BD Biosciences); CCR9 (clone 242503; obtained from R&D Systems); and CD103 (clone 2E7) and the FoxP3 intracellular staining kit (obtained from eBioscience). For CFSE dye dilution, cells were labeled with 5 µM CFSE (Invitrogen). Flow cytometric analysis was performed on a refurbished machine (FACScan; Becton Dickinson) running CellQuest software (BD Biosciences), and data analysis was performed using FlowJo software (TreeStar, Inc.).

Homing assay.
Competitive homing experiments of RA–T reg cells and A-Tregs were performed as previously described, with modifications (3). In brief, 10 x 10⁶ RA–T reg cells generated from sorted CD4⁺FoxP3^– cells purified from a Ly5.2⁺FoxP3/GFP reporter mouse were mixed with an approximately equal number of A-Tregs generated from FACS-sorted CD4⁺CD25^– cells harvested from a Ly5.2⁺ mouse (determined to be 20% FoxP3⁺, as gauged by intracellular FoxP3 staining) and intravenously injected into Ly5.1⁺ C57BL/6 mice. An aliquot was saved to determine the input ratio: IR = (GFP⁺)_input/(GFP^–)_input). The homing index (HI) was calculated as the ratio of (GFP⁺)_tissue/(GFP^–)_tissue to IR. Homing indices were tested versus HI = 1 using a one-sample Wilcoxon signed-rank test. Significance was set at P < 0.05.

Online supplemental material.
Fig. S1 shows that DCs, with the addition of exogenous RA, can induce FoxP3 expression in T cells in a TGF-ß1–dependent manner and over a dose titration of TGF-ß1. Fig. S2 shows that the addition of exogenous RA to TGF-ß1 and IL-2 induces a net increase in the total number of T reg cells generated under various levels of plate-bound co-stimulation. Online supplemental material is available at http://www.jem.org/cgi/content/full/jem.20070719/DC1.

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