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Immunology. Apr 2003; 108(4): 481–492.
PMCID: PMC1782923

Mucosal CD8α+ DC, with a plasmacytoid phenotype, induce differentiation and support function of T cells with regulatory properties

Abstract

Repetitive stimulation of naïve T cells by immature splenic dendritic cells (DC) can result in the differentiation of T-cell lines with regulatory properties. In the present study we identified a population of DC in the mucosae that exhibit the plasmacytoid phenotype, secrete interferon-α (IFN-α) following stimulation with oligodeoxynucleotides containing certain cytosine-phosphate-guanosine (CpG) motifs and can differentiate naïve T cells into cells that exhibit regulatory properties. Although these DC appear to be present in both spleen and mesenteric lymph nodes (MLN), only CpG-matured DC from the MLN (but not the spleen) were able to differentiate naïve T cells into T regulatory 1-like cells with regulatory properties. The activity of these DC failed to sustain robust T-cell proliferation and thereby enhanced the suppressive efficacy of CD4+ CD25+ T regulatory cells. These DC are the major CD8α+ DC population in the Peyer's patches (PP). Given their significant presence in mucosal tissue, we propose that these DC may provide a mechanistic basis for the homeostatic regulation in the gut by eliciting regulatory cell suppressor function and poorly supporting T helper cell proliferation at a site of high antigenic stimulation like the intestine.

Introduction

Dendritic cells (DC) are central to the generation of adaptive immune responses and there is clear evidence to suggest that DC populations are not all equal in their capacity to activate T cells. CD8α+ and CD8α DC have been proposed by some to be derived from lymphoid and myeloid lineages, respectively,1,2 and have been demonstrated to prime distinct T helper responses.3 In situ experimentation has shown that lymphoid and myeloid DC lineages may also have particular localization patterns in mucosal lymphoid tissue, raising the possibility of defined immunological roles for different DC populations.4 Functionally, peripheral CD8α+ DC are less potent inducers of T-cell activation compared with CD8α DC, despite the expression of similar levels of costimulatory molecules.2 This inefficiency to stimulate T cells has been proposed to result from the inadequate induction of interleukin (IL)-2 and other cytokines from lymphocytes, thus limiting the subsequent expansion of antigen-specific cells.57 More recently, peripheral CD8α+ DC have been shown to actively suppress ongoing immune responses in vivo via the immunoregulatory enzyme, IDO, suggesting that this subset of DC may utilize multiple mechanisms to limit T-cell responsiveness.8

Most studies concerning CD8α+ DC have concentrated on peripherally derived DC, but there is clear evidence to suggest that DC populations within gut-associated lymphoid tissue (GALT) may be important in controlling T-cell responsiveness to mucosal antigens.9,10 In healthy individuals, the gut microenvironment exists in a continuous state of controlled inflammation, despite the presence of potent antigen-presenting cells (APC), like DC. How, then, can the maintenance of intestinal homeostasis in the presence of such overt stimulation and such competent stimulatory cells be reconciled? CD4+ CD25+ T regulatory (Treg) cells are believed to play a central role in inhibiting the development of intestinal inflammation and inflammatory bowel disease (IBD).11,12 While conventional Treg cells are generated in the thymus,13 recent studies on transplantation tolerance14,15 and mucosally induced tolerance16 suggest that naïve CD4+ T cells in peripheral tissues can be taught to exhibit regulatory function [i.e. Tr1 and T helper 3 (Th3) cells]. Cytokines such as IL-10, interferon-α (IFN-α) or transforming growth factor-β (TGF-β) have each been used to generate regulatory T cells in vitro,17,18 while recent intriguing data shows that stimulation with immature DC19 or DC that overexpress Serrate-1, a ligand for the Notch 1 receptor,20 can also generate regulatory T cells. Of particular relevance to these recent findings is the observation that a subset of CD8α+ DC can produce IFN-α following stimulation with CpG.21 Given that IFN-α exposure can generate T cells with regulatory properties, it has been postulated that IFN-α-producing DC may represent a unique DC subset dedicated to the priming of regulatory T cells in vivo,22 thereby providing a mechanism for the stringent control of immune responses in the periphery.

In the present study, we investigated whether CD8α+ DC in the gut have the capacity to secrete IFN-α and we further analysed whether this specialized DC population might have the capacity to support differentiation of naïve T cells into T cells with regulatory qualities, even in the presence of maturation stimuli. Finally, we determined whether CD8α+ DC from the gut can interact with thymically derived Treg cells to support their regulatory function.

Materials and methods

Mice

Female BALB/c mice (6–8 weeks old) were purchased from Taconic Laboratories (Garmantown, NY). BALB/c DO11.10 OVA TCR Tg mice, whose T cells express transgenic (Tg) T-cell receptor (TCR) specific for ovalbumin (OVA)-P323−339, were bred in-house (Immunex Corporation, Seattle, WA). All mice were maintained under specific pathogen-free conditions in accordance with Immunex IACUC guidelines.

Reagents

Fluorescein isothiocyanate (FITC)-conjugated KJ-126 and purified anti-FcγRII monoclonal antibodies (mAbs) were made in-house. Anti-CD4, anti-CD25, anti-CD11b, anti-CD11c, anti-B220, anti-CD40, anti-CD62L, anti-CD69, anti-CD80, anti-CD86 and anti-I/A antibodies were purchased from BD Pharmingen (San Diego, CA). Chinese hamster ovary (CHO)-derived human Flt3-ligand (Flt3L) was produced and purified as described previously.23 Purified and biotin-labelled anti-IL-4, anti-IL-10 and anti-IFN-γ antibodies were purchased from BD Pharmingen.

Generation and isolation of DC

Mice were injected with the DC expansion factor Flt3L (10 µg in 100 µl) intraperitoneally (i.p.) once daily for 10 consecutive days, a process known to expand all DC subsets without activation.23 All DC subsets investigated in this report are present in mice that have not been treated with Flt3-L (Table 1). A single-cell suspension was prepared from the spleen, mesenteric lymph nodes (MLN), Peyer's patches (PP) and peripheral lymph nodes (PLN) (inguinal, popliteal, axillary, brachial and cervical) of treated mice by mechanical disruption of tissue (n = 10 mice/group). Cells were spun over a Nycodenz (GibcoBRL, Gaithersburg, MD) gradient to enrich for DC. Cells were stained for the expression of CD11b, CD11c and CD8α, and the different DC populations were separated by fluorescence-activated cell sorter (FACS) analysis using the Vantage (BD, San Diego, CA) or MoFlo (Cytomation, Ft. Collins, CO) cell sorters.

Table 1
Percentage of CD8α+ dendritic cells in tissues of untreated and Flt3-ligand (Flt3L)-treated mice

Preparation of lipopolysaccharide blasts

A single-cell suspension was prepared from spleens of naïve BALB/c mice. Cells were washed twice and resuspended in Iscove's modified Dulbecco's medium (IMDM) (Invitrogen, Carlsbad, CA) supplemented with 5% fetal calf serum (FCS), 5·5 × 10−5 m 2-mercaptoethanol (2-ME), 1 m sodium pyruvate, 100 µm non-essential amino acids, 100 U/ml penicillin, 100 µg/ml streptomycin and 2 mm l-glutamine (complete IMDM). Lipopolysaccharide (LPS) was added to the cultures at 10 µg/ml (Sigma, St Louis, MO) for 48 hr. Following the incubation, cells were washed three times and irradiated at 3000 rads (137Cs source) before being used as APC.

T-cell proliferation assays and generation of T-cell lines

PLN of DO11.10 TCR Tg animals were collected and CD4+ cells enriched by negative immunomagnetic selection. Non-CD4+ cells (Mac-1+, B220+, CD8+, Gr-1+ and Ter119+) were removed by incubating cell preparations with a CD4 T-cell isolation kit (Stem Cell Technologies, Vancouver, BC) and then passing these preparations over a magnetic column inside a VarioMACS (Miltenyi Biotec, Auburn, CA). For isolation of CD25 CD4+ cells, anti-CD25 mAb was added to the depleting antibody cocktail. Cells eluted in the flow-through fraction were enriched for CD4+ cells at >95% with less than 0·5% contaminating CD4+ CD25+ cells. For T-cell proliferation assays, 5 × 104 purified naïve CD4+ T cells were placed in wells of a 96-well plate with DC subsets, at a ratio of 5 : 1 (T cells : DC). OVA peptide (323–339) was added to cultures at a final concentration of 375 nm. Assays were pulsed with [3H]thymidine ([3H]TdR) after a 72-hr incubation. For the generation of T-cell lines, 1 × 106 total CD4+ T cells or CD4+ CD25+ Treg-depleted CD4+ CD25 T cells were placed in wells of a 48-well plate with DC subsets at a ratio of 10 : 1 (T cells : DC). In some experiments, DC were stimulated overnight in the presence or absence of 1 µg/ml CpG (Sigman-Genosys, The Woodlands, TX), counted and then pulsed with 375 nm of OVA peptide (323–339) before being added to T-cell cultures. Cell cultures were stimulated every 2 weeks, in the presence of 20 ng/ml of IL-2, by addition of appropriate DC subsets, pulsed with peptide, at a ratio of 10 : 1 (T cell : DC). At each fortnightly restimulation, a sample of cells was removed, phenotyped by FACS analysis and tested for regulatory activity in vitro.

Assays for regulatory activity by Tr-1 like cell lines

Tr1-like cell lines (as generated above) or freshly isolated ‘classical Treg’ CD4+ CD25+ cells, resuspended at 0·5 × 106 cells/ml, were distributed into 96-well round-bottom plates in a volume of 50 µl with an equal number of DO11.10-derived CD4+ CD25 T helper cells. LPS blasts or different DC populations were used as APC and were added to wells at 1·5 × 106 cells/ml in 50 µl. Each well contained a final volume of 200 µl of complete IMDM. Cellular proliferation was assayed over serial 10-fold dilutions of OVA peptide (323–339) from 500 nm to 0·5 nm. Cultures were incubated for a total of 96 hr at 37°, in an atmosphere of 5% CO2, and pulsed with 1 µCi [3H]TdR for the last 12 hr of culture. Cell cultures were harvested onto glass-fibre filtermats (Wallac, Turku, Finland) using a Brandel harvester (Brandel, Gaithersburg, MD) and [3H]TdR incorporation was assessed on either a TriLux 1450 MicroBeta counter (Wallac) (see Fig. 3a, Fig. 4, Fig. 5 and Fig. 6a) or a Packard beta counter (Packard, Meridan, CT) (Fig. 6b and Fig. 7b).

Figure 3
CD8+ plasmacytoid dendritic cells (pDC) are less efficient at inducing T helper cell proliferation than CD8+ non-pDC. Fresh CD8+ pDC and CD8+ non-pDC populations were isolated from the spleen (Sp), peripheral lymph nodes (PLN) and mesenteric lymph nodes ...
Figure 4
CD8+ plasmacytoid dendritic cells (pDC) favour T regulatory (Treg)-mediated suppression over T-cell proliferation. Fresh CD4+ CD25 T helper cells (black circles) and CD4+ CD25+ Treg cells (white circles) were isolated from DO11.10 T-cell receptor ...
Figure 5
T regulatory (Treg) cell-mediated suppression is more robust at high T helper : Treg cell ratios when CD8+ plasmacytoid dendritic cells (pDC) are the antigen-presenting cells (APC). Fresh CD4+ CD25 T helper and CD4+ CD25+ Treg cells were isolated ...
Figure 6
CD8+ plasmacytoid dendritic cells (pDC) induce de novo differentiation of T regulatory 1 (Tr1)-like cells from naïve T cells. T-cell lines were generated by culturing CD25-depleted CD4+ T cells for 3 weeks with either CD8+ pDC or CD8+ non-pDC ...
Figure 7
Mesenteric lymph node (MLN) CD8+ plasmacytoid dendritic cells (pDC) activated with CpG maintain the ability to induce T regulatory 1 (Tr1)-like T cells that exhibit regulatory T-cell function. T-cell lines were cultured with CD8+ pDC or CD8+ non-pDC for ...

Cytokine enzyme-linked immunosorbent assay

For assessment of IFN-α secretion by DC, freshly separated DC populations were placed in 96-well round-bottom plates at a concentration of 1 × 105 cells/well and stimulated with 1 µg/ml of CpG in a volume of 200 µl of complete IMDM. Supernatants were collected 18 hr later and tested for IFN-α levels using an IFN-α enzyme-linked immunosorbent assay (ELISA) kit (PBL Biomedical Laboratories, New Brunswick, NJ), according to the manufacturer's instructions.

Levels of IL-2, IL-4, IL-10 and IFN-γ in 48-, 72- and 96-hr culture supernatants from T-cell proliferation assays were determined using the Beadlyte Mouse Multi-Cytokine Detection System (Upstate Biotechnology, Lake Placid, NY) and the Luminex100 plate reader (Luminex Corporation, Austin, TX) according to the manufacturer's instructions. Quantification of cytokines was performed by regression analysis from a standard curve generated using cytokine standards included in the kit. Lower limits of detection were: 2 pg/ml for IL-2, 0·2 pg/ml for IL-4, 35 pg/ml for IL-10 and 3 pg/ml for IFN-γ.

IL-4, IL-10 and IFN-γ cytokine secretion by Tr1-like cell lines cultured with CD8α+ DC populations was assessed by stimulating T cells overnight with plate-bound anti-CD3 (clone 500-A2 at 10 µg/ml). Culture supernatants were collected at 48 hr and tested in sandwich ELISAs with biotin-labelled anti-cytokine antibodies, as per the manufacturer's instructions. Briefly, Nunc (Rochester, NY) ELISA plates were coated with purified antibodies at 1 µg/ml (IL-4 and IFN-γ) or 2 µg/ml (IL-10) in 0·1 m NaHCO3 (pH 8·5) and blocked with phosphate-buffered saline (PBS) supplemented with 10% FCS (PBS–FCS) before the addition of samples and standards. Samples were tested in duplicate over a series of two- and threefold serial dilutions. The appropriate biotinylated anti-cytokine antibodies were added to wells at 1 µg/ml, and streptavidin–horseradish peroxidase (Zymed, San Francisco, CA) was diluted 1 : 2000 in PBS–FCS. 3,3′,5,5′ tetramethylebenzidine (TMB) microwell substrate was obtained from Kirkergard & Perry Laboratories (Gaithersberg, MD). Plates were read on an ELISA plate reader (Molecular Devices, Sunnyvale, CA) at 450 nm, and cytokine concentrations in the supernatant were calculated using DeltaSoft 3, version 2·14, software (Biometallics, Princeton, NJ).

Statistical analysis

Statistical analyses (Student's t-test and Welch t-test) were performed using the InStat software program (GraphPad Software Inc., San Diego, CA).

Results

Phenotypic diversity of CD8α+ DC populations in peripheral and gut lymphoid tissues

CD8α+ DC have been shown to be less potent inducers of T-cell activation than CD8α DC2 and may also actively suppress ongoing immune responses in vitro and in vivo.8 In light of these studies, we assessed the relative proportion of CD8α+ DC in different lymphoid tissues, with particular emphasis on the GALT, which has yet to be analysed for this particular DC population. FACS analysis on cells derived from the spleen, MLN and PLN revealed the presence of two major subsets of CD8α+ DC that could be distinguished based on intensity of CD11c expression (Fig. 1, R1 and R2). We found that the CD8α+ CD11clo population in each of these tissues was B220+ Gr1+ CD19 and appeared to have an immature phenotype based on the low levels of expression of Class II, CD40, CD80 and CD86 (Fig. 1). This phenotype is indicative of plasmacytoid DC (pDC).24,25 The CD11chi population of CD8α+ DC did not express B220 or CD19, and expressed little or no Gr-1. PP were also found to contain the CD8α+ CD11clo, plasmacytoid-like DC; however, the corresponding CD8α+ CD11chi population was greatly reduced in the PP (mean 1·48%, range 0·68–2·90%, n = 7) compared with the other tissues (MLN: mean 6·45%, range 3·73–7·95%, n = 6; PLN: mean 3·87%, range 3·0–4·74, n = 2; spleen: mean 6·44%, range 5·48–8·10%, n = 4)(Fig. 1). Thus, the average ratio of CD8α+ CD11clo DC to CD8α+ CD11chi DC was highest in the PP at 4·2 : 1 compared with the spleen (0·9 : 1), MLN (0·8 : 1) and PLN (2·3 : 1). There was no significant difference observed in the percentage of CD8α+ CD11clo DC recovered from the PP when we compared our mechanical disruption technique with collagenase treatment (2·0% of CD8α+ CD11clo DC for mechanical disruption and 2·2% of CD8α+ CD11clo DC for collagenase treatment); we therefore subsequently used mechanical disruption to isolate DC because of the variability in endotoxin contamination of collagenase batches.

Figure 1
Phenotypic diversity of plasmacytoid dendritic cells (DC) in tissues of Flt3-ligand (Flt3L)-treated mice. A single-cell suspension was prepared from the spleen, mesenteric lymph nodes (MLN), peripheral lymph nodes (PLN) and Peyer's patches (PP) of Flt3L-treated ...

Gut-derived CD8α+ CD11clo DC exhibit increased capacity for IFN-α production compared with peripherally derived DC

Peripheral plasmacytoid DC are known to produce IFN-α following stimulation with CpG,21,25 and we confirmed that the CD8α+ CD11clo DC identified in the spleen secrete IFN-α following an 18-hr stimulation with CpG (Fig. 2). We further extended these studies to PLN and the GALT and found that CD8α+ CD11clo DC isolated from the PLN, MLN and PP also produce IFN-α following CpG stimulation (Fig. 2). Furthermore, we consistently found that CD8α+ CD11clo DC isolated from the MLN have an increased capacity for the production of IFN-α, on a per-cell basis, compared with the same population of cells isolated from the spleen, PLN or PP (Fig. 2).

Figure 2
Interferon-α (IFN-α) secretion by CD8α+ CD11chi and CD8α+ CD11clo dendritic cell (DC) populations. CD8α+ CD11chi and CD8α+ CD11clo DC populations were isolated from the spleen, mesenteric lymph nodes (MLN) ...

Based on the expression of B220, Gr-1, CD11c and the production of IFN-α, we conclude that there are two major populations of CD8α+ DC in the tissues we examined. We therefore refer to the CD8α+ CD11clo B220+ DC as CD8+ plasmacytoid DC (CD8+ pDC) and the CD8α+ CD11chi B220 DC population as CD8+ non-plasmacytoid DC (CD8+ non-pDC).

CD8+ pDC are less efficient at supporting CD4+ T helper cell proliferation

Compared with conventional DC, CD8α+ DC and plasmacytoid DC populations from the spleen and PLN have a decreased capacity to support T helper cell function.1,25 As described above, in our studies CD8α+ DC were divided into CD8+ pDC and CD8+ non-pDC and these populations were compared, within different tissues, for their capacity to support the proliferation of OVA TCR Tg T cells following stimulation with OVA peptide. [For PP, we compared the CD8+ pDC population with myeloid (CD11c+ CD11b+ CD8α) PP DC, as the CD8+ non-pDC population is rare.] Compared with CD8+ non-pDC, CD8α+ pDC from all tissues tested were observed to be less efficient at inducing T helper cell proliferation, although only CD8+ pDC derived from the MLN reached statistical significance (Fig. 3a). CD8α+ pDC from PP were significantly less efficient at inducing T helper proliferation compared with PP myeloid DC (Fig. 3a).

Cell culture supernatants were collected at 48 and 72 hr and assessed for the presence of IFN-γ, IL-10, IL-4 and IL-2. In cultures where CD8+ pDC were used as T-cell stimulators, higher levels of IL-10 (Fig. 3b) and lower levels of IFN-γ (Fig. 3c) were found compared with cultures where CD8+ non-pDC were used as T-cell stimulators. No significant difference was found in the IL-4 or IL-2 levels in supernatant, with either DC population as the APC (data not shown). These data suggest that stimulation of antigen-specific T cells by CD8+ pDC may result in a cytokine milieu that is more immunosuppressive rather than immunostimulatory.

CD8+ pDC support CD4+ CD25+ Treg cell suppression

Given the decreased efficiency of CD8+ pDC to induce T helper cell proliferation (Fig. 3), it was of interest to determine whether the CD8+ pDC identified in the GALT could support CD4+ CD25+ Treg activity, as gut homeostasis is believed to be maintained, at least in part, by the activity of Treg cells.26 In vitro, Treg cells suppress the proliferation of T helper cells responding to antigen and APC.13,27 We therefore compared the ability of CD8+ pDC and CD8+ non-pDC from different lymphoid tissues to support Treg cell-mediated suppression of OVA TCR Tg T cells responding to OVA peptide. Our findings were twofold: first, and in support of our previous data, CD8+ pDC did not induce comparable levels of T-cell proliferation compared with CD8+ non-pDC over a range of peptide concentrations. Second, CD4+ CD25+ Treg cell function was observed, regardless of which population of DC was used as APC; however, CD8+ pDC appeared to favour T-cell suppression over T-cell proliferation (Fig. 4). In order to ensure that the suppressive effect we observed in Fig. 4 was the result of suppressive activity and not a dilution effect of responding T cells, we titrated the number of CD4+ CD25+ Treg cells and tested T-cell proliferation at two different OVA concentrations using our different DC populations as APC. Data in Fig. 5 demonstrate that when CD8+ pDC are used as APC in the presence of Treg cells, Treg cell suppression is more robust at higher T helper : Treg cell ratios compared with the suppression observed when using CD8+ non-pDC as APC. The enhanced suppression at higher T helper : Treg ratios is probably a result of the poor stimulation of T helper responses by CD8+ pDC, as indicated by the difference in the maximum proliferation of T helper cells when using either CD8+ pDC or CD8+ non-pDC as APC (≈ 40 000 and 200 000 counts per minute, respectively). These data suggest that poor stimulation by CD8+ pDC, combined with suppression by CD4+ CD25+ Treg cells, results in low overall T helper cell proliferation.

CD8+ pDC induce the differentiation of naïve T cells into Tr1-like cells

IFN-α and immature DC have both been demonstrated, independently, to support the differentiation of cells that exhibit regulatory properties in vitro and in vivo.17,19,28 Given the high levels of IFN-α secreted by the MLN CD8+ pDC subset in our studies (Fig. 2), and the decreased expression of costimulatory molecules (Fig. 1), it was of interest to determine whether CD8+ pDC, particularly from the MLN, could induce the differentiation of T cells with regulatory properties from naïve CD4+ cells. CD4+ T cells depleted of CD25+ cells from OVA TCR Tg mice were cultured and stimulated weekly with OVA peptide-pulsed CD8+ pDC for three consecutive weeks. The suppressive activity of the resulting T-cell lines was assessed by the ability of these lines to inhibit proliferation of naïve OVA TCR Tg CD4+ CD25 T cells over a range of OVA peptide concentrations using irradiated splenocytes as APC. We found that T-cell lines derived from the CD8+ pDC could completely inhibit OVA TCR Tg T-cell proliferation, while T-cell lines derived from CD8+ non-pDC cultures did not (Fig. 6a). T-cell lines derived from both the CD8+ pDC and CD8+ non-pDC cultures proliferated poorly in response to antigen stimulation (Fig. 6a). If LPS-activated splenocytes were used as a potent APC in these restimulation assays, it was found that T-cell lines derived from the CD8+ pDC were still able to suppress helper T-cell proliferation (Fig. 6b), indicating that the strength of the APC stimulus influences the ability of these differentiated T cells to exhibit their regulatory properties. Similar results were obtained from T-cell lines derived from repetitive stimulation with splenic CD8+ pDC (data not shown). CD8+ pDC therefore bear the ability to support differentiation of T-cell lines with suppressive activity, which we have termed Tr1-like cells.

T cells with regulatory properties (referred to as Tr1 cells) are reported to produce high levels of IL-10, moderate amounts of IFN-γ and no IL-4,28 while immature human pDC have been shown to polarize T cells to produce IL-4, IL-10 and IFN-γ.29 We therefore tested the CD4+ Tr1-like cell lines, derived via culture with the different DC populations in this study, for their capacity to produce IL-4, IL-10 and IFN-γ following stimulation with plate-bound anti-CD3 antibodies. The Tr1-like cell lines generated by repetitive stimulation with CD8+ pDC were found to produce IL-10 at similar levels to Tr1 cells described in other studies (1·587 ± 0·28 ng/ml).30 However, the Tr1-like cell lines also produced IL-4 (7·814 ± 0·32 ng/ml) and IFN-γ (172·3 ± 10·8 ng/ml) upon activation, distinguishing them from bona fide Tr1 cells. Hence, we refer to our T-cell lines as Tr1-like. The T-cell lines that were derived via culture with CD8+ non-pDC did not produce detectable IL-4 or IL-10 (data not shown), but did secrete IFN-γ (368·04 ± 24·5 ng/ml). Our data suggest therefore that both CD8+ non-pDC and CD8+ pDC can induce the differentiation of poorly proliferative T-cell lines; however, only those T-cell lines differentiated by stimulation with CD8+ pDC can produce significant levels of IL-4 and IL-10 and suppress T helper proliferation.

CpG matured MLN CD8+ pDC retain the ability to induce differentiation of Tr1-like cells

Reduced expression of costimulatory molecules by pDC is thought to contribute to the ability of these DC to support differentiation of T cells with regulatory function.22 We therefore stimulated MLN-derived CD8+ pDC with CpG to upregulate surface costimulatory molecules (Fig. 7a) and tested whether CpG-matured CD8+ pDC could still induce differentiation of Tr1-like cells. Our data indicate that CD8+ pDC from MLN can indeed support differentiation of Tr1-like cells with suppressive activity, whether or not the DC exhibit a mature or immature phenotype at the onset of culture as a consequence of CpG exposure (Fig. 7b). As expected, when we tested normal irradiated splenocytes as the APC (Fig. 7b, right panel), rather than the more potent LPS-blasted splenocytes (Fig. 7b, left panel), we found an enhanced inhibition of T-cell proliferation by the Tr1-like cell lines that were generated using repetitive stimulation of CpG-matured CD8+ pDC (Fig. 7b). T-cell lines generated by CpG-matured CD8+ pDC from the spleen were not able to suppress T-helper cell proliferation (data not shown).

Consistent with the T-cell lines generated by immature CD8+ pDC, T-cell lines generated by CpG-matured CD8+ pDC produced significant amounts of IL-4 and IL-10 upon stimulation with plate-bound anti-CD3 compared to those lines generated with CpG-matured CD8+ non-pDC (Fig. 7c). IFN-γ production by the CD8+ pDC-generated T-cell lines was only slightly lower than that of CD8+ non-pDC-generated T-cell lines (data not shown). These data suggest that the CD8+ pDC present in the MLN have the capacity to induce Tr1-like cells, even in the presence of maturation stimuli.

Discussion

In this study, we identified a particular subset of DC within the GALT that is inefficient at inducing antigen-mediated proliferation and secretes high levels of IFN-α following stimulation with CpG. These characteristics are similar to the recently described murine equivalent of human plasmacytoid DC, which produce IFN-α following stimulation with CpG and generally express low levels of costimulatory molecules.24,25 More importantly, we showed that the mucosally derived population of plasmacytoid DC supports CD4+ CD25+ Treg-mediated suppression of T-cell responses and induces de novo differentiation of Tr1-like cells that secrete IL-4 and IL-10 and suppress the proliferation of T helper cells in vitro. Most significant is our finding that when these DC are matured with CpG, they can still induce the differentiation of Tr1-like cells that exhibit suppressive properties. These findings therefore suggest that a particular population of DC within the GALT may be less likely to induce robust T-cell expansion, even in the presence of overt stimulation, and may be more likely to support regulatory T-cell activity that is reported to predominate at mucosal sites.31

T cells with regulatory properties can be elicited after oral administration of antigen16,3234 and are implicated in the maintenance of oral tolerance. Furthermore, IL-10-driven Tr1 cells and CD4+ CD25+ Treg cells have both been clearly shown to play a role in vivo in the suppression of colitis.26,28 Therefore, suppression mediated by regulatory T cells is probably a major mechanism by which the immune system of the gut is able to maintain a state of ‘controlled inflammation’. The role that gut DC might play in maintaining intestinal homeostastis is, as yet, unclear but has been alluded to through the finding that DC expansion in vivo enhances oral tolerance.35 Recent data monitoring T-cell expansion at local tissue sites following oral antigen administration shows that T cells at mucosal sites undergo fewer divisions than T cells in the periphery.9 It is intriguing to postulate that mucosal DC might be involved in this limitation of T-cell expansion. Indeed, DC from different mucosal sites are known to produce immunosuppressive cytokines such as IL-10 or TGF-β,10,31,32 and can differentiate naïve CD4+ T cells into cells that secrete IFN-γ, IL-4 or IL-10 following stimulation with CD40 ligand (CD40L) or Staphylococcus aureus Cowan 1 (SAC) combined with IFN-γ.10 In our studies, we differentiated naïve CD4+ T cells into cells that secrete IFN-γ, IL-4 and IL-10 using DC that express high levels of toll-like receptor (TLR) 9 and are responsive to CpG. This differential responsiveness to pathogen-associated molecular patterns (PAMPs) conferred by different TLR, expressed on different DC populations, may be central in maintaining selective non-responsiveness to certain microbes and might play a role in discriminating between pathogens and commensal microflora in the gut.36

Recent data, showing that CD8+ pDC demonstrate selective usage of chemokine receptors, also supports a role for CD8+ pDC, particularly those present in the PP, in maintaining control of local immune responses.37 Murine CD8α+ DC primarily and constitutively express the chemokine receptors, CXCR4 and CCR7.37 The ligand for CXCR4 is stromal-cell-derived factor 1 (SDF-1), a chemokine expressed within the lymph node,38 suggesting that these DC may reach lymph nodes independently of inflammatory stimuli. Moreover, expression of CCR7 by these DC has been documented in the PP and perhaps indicates that they are uniquely attracted to T-cell areas of secondary lymphoid tissues in response to macrophage inflammatory protein-3β (MIP-3β)/EBI 1-ligand chemokine (ELC).4,37 Interestingly, in humans, plasmacytoid DC are refractory to inflammatory chemotactic signals and demonstrate a preferential bias towards lymph node-homing chemokines.37 Given their inefficiency in generating T-cell proliferation, their imperviousness to inflammatory stimuli37 and their significant presence in the interfollicular T-cell rich region of PP4 (at least in mice), we hypothesize that under static, non-inflammatory conditions, these CD8+ pDC may play a significant role in dampening local T-cell responses to normal luminal antigens.39 Under conditions of pathogenic assault, we hypothesize that these DC encourage limited T-cell responsiveness to pathogens in the face of selected PAMPs. This concept is supported by our finding that CpG-matured CD8+ pDC can still induce the differentiation of Tr1-like cells. Limiting inflammatory responses in the intestine, even in situations mimicked by CpG, would be highly desirable in order to maintain homeostasis and barrier integrity. Furthermore, as pDC have the innate ability to produce anti-pathogenic factors, such as IFN-α, such characteristics might ensure that ‘a first-line defence’ scenario could be employed without the inflammation associated with active adaptive immunity. Under conditions where the pathogenic assault is more robust, we hypothesize that other, more immunostimulatory, DC populations may be employed to generate a more active immune response.

Our findings support studies suggesting that murine plasmacytoid DC induce poor stimulation of T-cell responses, and we provide new evidence suggesting that murine CD8+ pDC have the ability to both differentiate IL-4- and IL-10-secreting Tr1-like cells with suppressive activity and support CD4+ CD25+ Treg cell mediated suppression by stimulating poor T helper responses. We hypothesize that these DC not only play a pivotal role in the downregulation of immune responses in the gut, but are also involved in the differentiation and maintenance of the resident regulatory T-cell population. The significant presence of these cells in the GALT suggests that they are likely to be dedicated to maintaining the gut environment in a state of controlled inflammation whilst simultaneously dealing with microbial invasion.

Acknowledgments

We thank Steve Brady, Julie Hill and Daniel Hirschstein for assistance with flow cytometry. We also thank Laurent Galibert, Thibaut de Smedt, David Cosman, John Sims and Anne Aumell for informative discussions and for critically reviewing this manuscript.

Abbreviations

CpG
cytosine-phosphate-guanosine
DC
dendritic cell
Flt3L
Flt3-ligand
GALT
gut-associated lymphoid tissue
IBD
inflammatory bowel disease
MLN
mesenteric lymph nodes
PLN
peripheral lymph nodes
PP
Peyer's patches
pDC
plasmacytoid dendritic cell
Th3
T helper 3
Tr1
T regulatory 1
transgenic
Tg
Treg
T regulatory

References

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