Article | Published:

CCR9 expression defines tolerogenic plasmacytoid dendritic cells able to suppress acute graft-versus-host disease

Nature Immunology volume 9, pages 12531260 (2008) | Download Citation

Subjects

Abstract

Dendritic cells (DCs) are 'professional' antigen-presenting cells that are key in the regulation of immune responses. Here we characterize a unique subset of tolerogenic DCs that expressed the chemokine receptor CCR9 and migrated to the CCR9 ligand CCL25, a chemokine linked to the homing of T cells and DCs to the gut. CCR9+ DCs were of the plasmacytoid DC (pDC) lineage, had an immature phenotype and rapidly downregulated CCR9 in response to maturation-inducing pDC-restricted Toll-like receptor ligands. CCR9+ pDCs were potent inducers of regulatory T cell function and suppressed antigen-specific immune responses both in vitro and in vivo, including inhibiting acute graft-versus-host disease induced by allogeneic CD4+ donor T cells in irradiated recipients. Our results identify a highly immunosuppressive population of pDCs present in lymphoid tissues.

Main

Dendritic cells (DCs) constitute a family of bone marrow–derived antigen-presenting cells that are pivotal in orchestrating immune responses either by maintaining tolerance to self antigens or by inducing potent immune responses to infectious agents1. After being exposed to microbial products, DCs mature by upregulating costimulatory molecules (CD40, CD80 and CD86) and migrate to T cell areas of organized lymphoid tissues, where they activate naive T cells and induce effector rather than tolerogenic immune responses. In the absence of such inflammatory or infectious signals, however, DCs present self antigens in secondary lymphoid tissues for the induction and maintenance of self-tolerance2. The ability of DCs to induce tolerance has led to many studies that have used these cells therapeutically in an effort to control unwanted immune responses in models of allograft rejection, graft-versus-host disease (GVHD) and autoimmune disorders3,4. Most studies have used myeloid DCs (mDCs) derived from mouse bone marrow or human monocytes cultured in vitro with the cytokine granulocyte-macrophage colony stimulating factor in the presence or absence of interleukin 4 (IL-4)4. Such studies have shown that in vitro–derived immature mDCs are able to dampen arthritis in an antigen-driven mouse model5 and prolong allograft survival in a mouse transplant model6,7. As the therapeutic effects have generally been incomplete, some studies have further manipulated mDCs through genetic modification8,9,10 or exposure to either immunosuppressive agents11 or cytokines such as IL-10 and transforming growth factor-β12 in an effort to generate more-potent tolerogenic mDC populations.

Subsequent studies, however, have suggested that naturally occurring DC subsets may also be effective in inducing peripheral tolerance. Lymphoid-related CD11c+CD8α+ DCs mobilized in vivo by the hematopoietic growth factor Flt3L have been shown to prolong the survival of vascularized heart allografts in rodents13. Plasmacytoid DCs (pDCs) have also been suggested to be critical regulators of immune responses14. These cells are best known for their copious production of type I interferons and subsequent induction of cell-mediated adaptive immune responses after viral activation15. However, in the absence of maturation signals, freshly isolated pDCs do not induce strong T cell responses; instead, they seem to prime naive CD4+ T cells to differentiate into IL-10-producing regulatory Tr1 cells in vitro in both humans16 and mice17,18. In vivo, unstimulated pDC precursors have been shown to facilitate allogeneic hematopoietic stem cell engraftment19,20 and delay allograft rejection in mice21. As pDCs have less capacity for foreign antigen uptake than their mDC counterparts do22, it has been postulated that pDCs present more self antigen–major histocompatibility complexes (MHC) than mDCs do. In addition, the observation that pDCs normally reside in the thymus and peripheral lymphoid tissues suggests that they may have greater involvement in tolerance than other DC subsets do23. Thus, resting or immature pDCs may represent a particularly important naturally occurring regulatory DC subset14,15.

Despite the promise of cellular therapy with DC populations, so far, no studies have taken advantage of specific tolerogenic phenotypes to sort immunosuppressive DCs from immune-activating DCs (on the basis of DC maturation markers, for example). Perhaps as a consequence, most DC populations studied have yielded only partial or transient amelioration of autoimmune symptoms or allograft survival3,4. Here we report that the chemokine receptor CCR9 (A000632), linked before to the homing of T cells to the gut24 and the homing of T cell precursors to the thymus25, was selectively expressed on pDCs of immature phenotype in vivo. CCR9 expression was rapidly downregulated in response to maturational signals and could be used to effectively distinguish endogenous pDCs of immature and mature phenotypes. CCR9+ pDCs constituted a sizeable fraction of the pDC compartment in resting secondary lymphoid tissues; in addition, they were much more efficient than CCR9 pDCs in inducing regulatory T cells and they inhibited antigen-specific immune responses both in vitro and in vivo. Finally, sorted and adoptively transferred CCR9+ (but not CCR9) pDCs effectively prevented acute GVHD, providing long-term suppression of graft-versus-host responses in a model of allogeneic T cell transfer.

Results

DCs express gut-specific T cell–homing receptors

Using a gene-expression profiling approach, we evaluated the expression of tissue-specific homing receptor transcripts in DCs from various tissues in an effort to explore the homing patterns of DCs and the effect they might have on tissue-specific immune responses. We characterized the expression of selected trafficking receptor transcripts in CD11c+ DCs from mesenteric lymph nodes (MLNs), Peyer's patches and peripheral lymph nodes (PLNs), as well as in memory CD4+ T cells from MLNs and the lamina propria of the small intestine, for comparison. We focused on the expression of key gut- and skin-specific homing receptors because many studies have used this known dichotomy between tissue-homing lymphocytes in various inflammatory disorders26,27. Gut-associated T cells had high expression of the gut-selective homing receptors CCR9 and the β7 integrin (part of the α4β7 heterodimer), whereas their expression of the skin-homing receptors CCR4 and CCR10 was low (data not shown), which confirmed the utility of the assay. However, we unexpectedly found high expression of CCR9 transcript and protein on DCs from lymphoid tissues, including not only lymph nodes that drain the gut (MLNs) but also those that do not (PLNs; Fig. 1 and data not shown). In contrast, transcripts for the skin-homing-associated chemokine receptors CCR4 and CCR10 were low in all DCs tested here (data not shown). Flow cytometry confirmed high expression of CCR9 protein on immature CD11c+ MHC class II–intermediate cells in lymphoid tissues (Fig. 1), whereas only a small percentage of DCs in the blood were CCR9+.

Figure 1: Tissue-specific CCR9 expression profiles of DCs.
Figure 1

Surface expression of CCR9 and MHC class II (MHC II) on CD11c+ DCs from the MLNs, PLNs, spleen and blood of normal BALB/c mice, gated on CD11c+ DCs. Numbers adjacent to outlined areas indicate percent CCR9+, MHC class II–intermediate cells. Data are from one of two representative experiments.

CCR9 defines an immature population of pDCs

Using flow cytometry, we subdivided the DC populations from different lymphoid tissues into pDCs (CD11cintB220+) and non-pDCs (CD11chiB220). The non-pDC group included CD11chiB220CD8αCD11bhi mDCs and the so-called CD11chiB220CD8αhiCD11blo 'lymphoid' DCs. Almost all CCR9+ DCs in the lymphoid tissues examined were in the pDC subset (Fig. 2a) and almost all had an 'immature' phenotype, as shown by their low expression of the costimulatory molecules CD80, CD86 and CD40 and intermediate expression of MHC class II molecules (Fig. 2a). The mainly CCR9-deficient mDC (non-pDC) compartment contained DCs with slightly higher expression of costimulatory molecules (Fig. 2a, bottom). The pDC compartment in lymphoid tissues contained a sizeable population of CCR9+ pDCs (Fig. 2b), with the highest percentage (about 70–80%) of pDCs expressing CCR9 in the PLNs and MLNs. The lowest proportion of CCR9+ DCs among pDCs was in the blood and thymus. As a whole, pDCs are less abundant than mDCs, so CCR9+ pDCs represent about 12–18% of total CD11c+ DCs in the PLNs, about 5–10% of DCs in the MLNs and spleen, 2–3% of DCs in the blood and less than 1% of DCs in the thymus.

Figure 2: CCR9+ DCs reside in the pDC compartment and have a mainly immature phenotype.
Figure 2

(a) Flow cytometry of the expression of MHC class II and the costimulatory markers CD80, CD86 and CD40, correlated with CCR9 expression, to assess the maturation state of DCs from pooled PLNs isolated from normal C57BL/6 mice. Cell suspensions were stained and gated on pDC (CD11cintB220+) and non-pDC (CD11chi B220) lineage-negative DCs; gates were set on the basis of isotype controls for MHC class II and costimulatory markers. Numbers adjacent to outlined areas and in quadrants indicate percent positive cells in each. Data are representative of results from two experiments with three mice in each. (b) CCR9+ cells in gated pDC and non-pDC subsets from spleen, PLNs, MLNs, blood and thymus. Data are from one of two experiments with similar results (mean and s.e.m. of three mice). (c) Flow cytometry of the expression of MHC class II and the costimulatory markers CD80 and CD40, correlated with CCR9 expression, to assess the maturation state of DCs from pooled PLNs expanded by Flt3L-secreting B16 melanoma cells in C57BL/6 mice, gated on pDC (CD11cint B220+) and non-pDC (CD11chi B220) lineage-negative DCs; gates were set on the basis of isotype controls for MHC class II, CD80 and CD40. Numbers adjacent to outlined areas and in quadrants indicate percent positive cells in each. Data are representative of three separate experiments with a total of approximately ten mice.

To determine whether additional population expansion of pDCs in vivo would alter CCR9 expression on these cells, we transplanted C57BL/6 mice with a B16 melanoma cell line secreting Flt3L (called 'Flt3L-treated B6 mice' here). This system allowed us to expand the pDC population without activation, as Flt3L is an important growth and differentiation factor for the development of pDCs from hematopoietic stem cells in mice28. Flt3L treatment in vivo increased the frequency and number of pDCs in lymphoid tissues by almost tenfold after 10–14 d (data not shown). In addition to the CCR9+ pDC population, there was a distinct population of CCR9 pDCs with higher expression of costimulatory molecules (Fig. 2c). However, most CCR9+ pDCs remained phenotypically immature even after in vivo population expansion with Flt3L. These results collectively show that CCR9 defines an immature population of pDCs in peripheral lymphoid tissues that is distinct from most mature pDCs and from mDCs.

CCR9+ pDCs migrate to CCL25

We next sought to determine whether CCR9 on pDCs was functional by assessing the chemotactic responses of various DC subsets to the CCR9 ligand CCL25 (a chemokine also called 'thymus-expressed chemokine'; A002265). Because the number of pDCs that can be recovered from normal lymphoid tissues is limited, we expanded the DC population in Flt3L-treated B6 mice as described above and examined the migration of pooled PLN cells in response to various chemokines across a Transwell membrane. As expected, pDCs migrated more efficiently than other DC populations toward CXCL12 (the ligand for CXCR4; also called SDF-1), identified before as a potent chemoattractant for pDCs29,30 (Fig. 3a). Notably, pDCs were the only DC subset to migrate efficiently in response to CCL25, and they did so with a higher chemotactic response than their response to CXCL12 (Fig. 3a). As for the CCR9+ and CCR9 pDC subsets, only the CCR9+ pDC subset migrated efficiently to CCL25 (Fig. 3b). In contrast, consistent with their immature status, CCR9+ pDCs did not migrate to CCL21 (SLC; Fig. 3b), a ligand for CCR7 that is upregulated on mature DCs after activation31.

Figure 3: CCR9 expression allows pDCs to migrate to CCL25.
Figure 3

Chemotactic responses of DC subsets from pooled lymph node cells of Flt3L-treated B6 mice to medium or to the CXCR4 ligand CXCL12, the CCR7 ligand CCL21 or the CCR9 ligand CCL25. (a) DC subsets subdivided into pDCs and non-pDCs as described in Figure 2c. *, P = 0.001, or **, P < 0.001, migration of pDCs versus non-pDCs to CXCL12 or CCL25, respectively (t-test). (b) The pDC subset subdivided on the basis of CCR9 expression. *, P = 0.008, or **, P < 0.001, migration of CCR9+ versus CCR9 pDCs to CCL21 or CCL25, respectively. Data are from one of two experiments (error bars, s.e.m. of triplicate wells).

Activated pDCs produce type I interferon and downregulate CCR9

To determine if CCR9 expression is confined to immature pDCs or instead is maintained on the CCR9+ subset during maturation, we stimulated in vitro–sorted CCR9+ pDCs from Flt3L-treated B6 mice with an array of Toll-like receptor (TLR) ligands. Unlike mDCs, pDCs do not express TLR2, TLR4, TLR5 or TLR3, which makes them unresponsive to bacterial products such as peptidoglycan, lipopolysaccharide (LPS) and flagellin or viral double-stranded RNA mimics, respectively15. However, pDCs are equipped with microbial sensors such as TLR7 or TLR9 that detect the presence of single-stranded RNA or microbial DNA, respectively15. As expected, activation of sorted CCR9+ pDCs with LPS induced no DC activation (Fig. 4a) or cytokine production (Fig. 4b), and the production of interferon-α (IFN-α) and tumor necrosis factor (TNF) as well as the expression of MHC class II and the costimulatory molecules CD80 and CD40 remained low and similar to that of untreated cells (Fig. 4). However, treatment with the pDC-specific TLR ligands R-837 (synthetic TLR7 ligand) or bacterial CpG oligonucleotides (TLR9 ligand) downregulated CCR9 on half or more of the cells, with a concomitant increase in the expression of MHC class II, CD80 and CD40 on the CCR9-downregulated population (Fig. 4a). In addition, overnight treatment with CpG resulted in a burst of production of IFN-α and TNF by both the CCR9+ and CCR9 pDC subsets (Fig. 4b). These results further support the idea of the plasmacytoid identity of CCR9-expressing DCs and define CCR9 as a marker for immature pDCs, as CCR9 expression was lost after TLR-dependent activation of these cells.

Figure 4: CCR9+ DCs downregulate CCR9 after being activated with pDC-specific TLR ligands.
Figure 4

(a) Flow cytometry of CCR9, MHC class II (I-Ab) and the costimulatory ligands CD80 and CD40 in CCR9+ pDCs sorted from pooled PLNs isolated from Flt3L-treated B6 mice, then left untreated or activated for 8–12 h in the presence of LPS (1 ng/ml), CpG (1 μM) or R-837 (10 μg/ml). Gates were based on isotype controls with control antibody staining, which yielded less than 1% of cells in the positive gates. Numbers in quadrants indicate percent CCR9+I-Ab cells (top left), CCR9+I-Ab+ cells (top right) or CCR9I-Ab+ cells (bottom right). Data are from one of three experiments with similar results. (b) IFN-α and TNF in supernatants of sorted CCR9+ and CCR9 pDCs activated for 16 h in the presence of LPS (10 ng/ml) or increasing doses of CpG (2 or 20 μg/ml; wedges). Data are from one of two (IFN-α) or three (TNF) experiments with similar results (mean of duplicate cultures).

CCR9+ DCs suppress immune responses

We next sought to determine whether CCR9+ pDCs, with a characteristic immature phenotype, were potent in suppressing immune responses in vitro and in vivo. Using an antigen-specific approach, we sorted CCR9+ and CCR9 pDCs from Flt3L-treated B6 mice and cultured them for 2–4 h with an ovalbumin peptide of amino acids 323–339 (pOVA) before injecting them intravenously into naive B6 mice. We boosted recipient mice 1 week later with similar antigen-loaded pDCs and immunized them 1 week after the final boost with pOVA in complete Freund's adjuvant. After 10 d, we examined draining lymph nodes for in vitro recall responses to pOVA. Lymphoid populations from mice that initially received CCR9+ pDCs were impaired in their ability to proliferate in response to pOVA in vitro relative to those from mice that had received CCR9 pDCs or no pDCs at all (Fig. 5a).

Figure 5: CCR9+ DCs suppress immune responses in vivo and in vitro.
Figure 5

(a) CCR9+ and CCR9 pDCs were sorted from pooled PLNs isolated from Flt3L-treated B6 mice, pulsed for 2–4 h with pOVA, then administered intravenously into naive C57BL/6 mice (control mice did not receive pDCs); recipient mice were boosted 1 week later with the same antigen-loaded pDCs and were immunized subcutaneously 1 week after the final boost with pOVA emulsified in complete Freund's adjuvant. After 10 d, cell suspensions from draining lymph nodes were stimulated for 72 h with a 'dose range' of pOVA before the addition of [3H]thymidine. Data are from one of two experiments with similar results (mean and s.e.m. (error bars) of quadruplicate cultures). (b) CCR9+ and CCR9 pDCs were sorted from pooled lymph nodes of Flt3L-treated B6 mice, then cultured a ratio of 1:5 (pDCs:T cells) with CD4+ T cells purified by magnetic-activated cell sorting from spleens of BALB/c mice. Cells were then stimulated for 72 h with a 'dose range' of pOVA and cultured with [3H]thymidine for an additional 18 h. Data are from one of two experiments (mean and s.e.m. (error bars) of triplicate cultures).

Because pDCs are important in inducing distinct CD4+ T helper cell phenotypes15, we next examined the function of CCR9+ DCs in priming T cell responses. We used an in vitro allogeneic stimulation system in which we primed splenic CD4+ T cells from BALB/c mice with sorted CCR9+ and CCR9 pDC subsets from pooled PLNs of Flt3L-treated B6 mice. CCR9+ pDCs failed to support the proliferation of allogeneic T cells, in contrast to their CCR9 counterparts (Fig. 5b). Phenotypic analysis of the cultured T cells showed that CCR9+ pDCs induced fewer activated Foxp3CD4+CD25+ T cells than did CCR9 pDCs (Fig. 6a). Instead, a higher percentage and a predominant population of Foxp3+CD4+CD25+ T cells, which phenotypically resembled regulatory T cells, appeared after 5 d of culture with CCR9+ pDCs (Fig. 6a). In addition, the CCR9+ pDC–induced T cells suppressed the proliferation of freshly isolated CD4+CD25 effector T cells in coculture experiments, whereas T cells primed by the CCR9 DC subset were inefficient in suppressing effector T cell responses (Fig. 6b). Given those results, we propose that the CCR9+ pDC population is the main pDC subset that contributes to T cell tolerance, as these cells induced regulatory T cells, had an immature phenotype and represented almost the entire immature pDC pool in lymphoid tissues.

Figure 6: CCR9+ pDCs are potent inducers of regulatory T cells in vitro.
Figure 6

(a) Flow cytometry of the expression of intracellular Foxp3 and cell surface CD25 induced by CCR9+ and CCR9 pDCs (0.2 × 106 cells) sorted from pooled lymph nodes isolated from Flt3L-treated B6 mice and cultured for 5 d with splenic CD4+ BALB/c T cells (1 × 106). Cells were gated on CD4+ lymphocytes; gates were set on the basis of isotype controls for mAb to Foxp3 and mAb to CD25. Numbers in quadrants indicate percent CD25+Foxp3 cells (top left) or CD25+Foxp3+ cells (top right). (b) Proliferation of effector cells after CD4+ T cells (primed T cells) cultured in vitro for 5 d with CCR9+ pDCs or CCR9 pDCs were added in increasing numbers to 1 × 105 freshly isolated BALB/c CD4+CD25 effector T cells in wells coated with anti-CD3 and anti-CD28, followed by culture for 48 h and a pulse for an additional 18 h with 1 μCi [3H]thymidine. Data are from one of two experiments with similar results (a,b; error bars (b), s.e.m. of triplicate cultures).

CCR9+ DCs suppress acute GVHD

Because CCR9+ pDCs suppressed alloresponses in vitro, we next examined the effect of CCR9+ DCs in vivo in an animal model of GVHD induced by the transplantation of allogeneic bone marrow. To determine whether CCR9+ pDCs from Flt3L-treated B6 mice could suppress GVHD induced by CD4+CD25 BALB/c T cells, we injected the two populations together at a ratio of 1:2 (DCs:T cells), along with T cell–depleted BALB/c bone marrow, into C57BL/6 host mice within 24 h of lethal total-body irradiation (900 rads). All mice that received CD4+CD25 effector T cells and bone marrow developed clinical signs of GVHD, including diarrhea, skin ulcerations and weight loss; approximately 50% died after 5 weeks (Fig. 7a). We obtained similar results with mice that received CCR9 pDCs together with CD4+CD25 effector T cells. The addition of CCR9+ pDCs with effector T cells to bone marrow–transplanted hosts 'rescued' all the mice from death (100% in two separate experiments; Fig. 7a) and ameliorated their clinical signs, including diarrhea, weight loss and hunched posture.

Figure 7: Lethal GVHD of C57BL/6 recipients induced by BALB/c CD4+CD25 effector T cells can be suppressed by coinjected C57BL/6 CCR9+ DCs.
Figure 7

(a) Survival of C57BL/6 mice given two doses of 450 rads of total-body irradiation, BALB/c T cell–depleted bone marrow (2 × 106 cells) and BALB/c splenic CD4+CD25 T cells (0.5 × 106 (first experiment) or 1 × 106 (second experiment)) coinjected with sorted CCR9+ pDCs, CCR9 pDCs or no pDCs (No DC control) from pooled PLNs of Flt3L-treated B6 mice (0.2 × 106 or 0.5 × 106 DCs per mouse, respectively). *, P = 0.027, CCR9+ pDCs versus no pDCs; **, P = 0.010, CCR9+ pDCs versus CCR9 pDCs (log-rank test). Data are from two independent experiments with similar results, with a total of nine to ten mice per treatment group. (b) Intracellular cytokine staining of splenocytes and PLNs from irradiated C57BL/6 (Thy-1.2+) mice 3 weeks after intravenous transfer of BALB/c (Thy-1.2+) T cell–depleted bone marrow (2 × 106 cells), splenic Thy-1.1+ effector CD4+CD25 T cells from congenic BALB/c.Thy-1.1 mice (0.5 × 106 to 1 × 106 cells) and either no pDCs or sorted CCR9+ or CCR9 pDCs from Flt3L-treated B6 (Thy-1.2+) mice. Cells were gated on CD4+Thy-1.1+ effector T cells. Data are from one of two experiments with similar results, with pooled tissues from two to three mice. (c) Flow cytometry of the expression of intracellular Foxp3 and surface CD25 by unstimulated splenocytes and MLN cells from the mice in b. Cells were gated on CD4+ Thy-1.1+ effector T cells; gates were set on the basis of isotype controls for the mAb to Foxp3 and mAb to CD25 to include less than 1% in the positive gates. Numbers in quadrants indicate percent CD25+Foxp cells (top left), CD25+Foxp+ cells (top right) or CD25Foxp+ cells (bottom right). Data are representative of two experiments.

To monitor the effects of CCR9+ DCs on coinjected effector T cells, we used CD4+CD25 effector T cells from congenic BALB/c Thy-1.1 donor mice. All other mice (irradiated recipients and donor mice for the sorted DC subsets and bone marrow) were Thy-1.2+. We found that 3 weeks after transfer, Thy-1.1+ CD4+ effector T cells from PLNs produced more IL-17 and IFN-γ in GVHD mice that received no pDCs (Fig. 7b) than in unmanipulated healthy controls (percentages of IL-17- and IFN-γ-producing T cells less than 1% of that in PLNs of untreated controls). Coinjection of CCR9+ pDCs suppressed the frequency of IL-17-producing effector T cells to about 25% of that in the GVHD group that received no pDCs, without substantially decreasing the frequency of IFN-γ-producing effector cells (Fig. 7b, left). Studies suggest that the development of IL-17-producing T helper cells and the development of T helper type 1 (IFN-γ-producing) cells are antagonistic to each other32,33; we found that coinjected CCR9 pDCs suppressed the appearance of IFN-γ- but not IL-17-producing effector T cells (Fig. 7b, left). IL-17 production in the spleen was less prominent, but we still found few IL-17- and many IFN-γ-producing splenic effector T cells after cotransfer of CCR9+ pDCs (Fig. 7b, right). The frequency of cytokine-producing effector T cells was higher in the PLNs and spleen (Fig. 7b) than in the MLNs (data not shown). Examination of Thy-1.1+ CD4+ effector T cells for the regulatory T cell marker Foxp3 showed an expansion of the Foxp3+CD25 T cell population in the MLNs and spleens of recipient mice that received CCR9+ pDCs (Fig. 7c). In contrast, CCR9 pDCs failed to induce Foxp3+ effector T cells, similar to results obtained with the GVHD control mice that did not receive DCs. These results collectively show that CCR9+ DCs are potent suppressors of in vivo alloresponses; they decrease the clinical severity of allogeneic GVHD, suppress effector T cell responses (in particular IL-17 production) and induce new development of Foxp3+ regulatory T cells from effector cells.

Discussion

We have shown here that the chemokine receptor CCR9 selectively marked immature pDCs and that these CCR9+ pDCs were normally present as a resident pDC population in resting secondary lymphoid tissues. CCR9+ DCs underwent maturation by upregulating costimulatory and MHC class II molecules in response to TLR7 and TLR9 ligands but not TLR4 ligands; moreover, they produced IFN-α after TLR activation, which confirmed their plasmacytoid lineage. Notably, CCR9 expression was lost after activation, which indicated that CCR9 can be used as a reliable marker of immature pDCs. Moreover, we found that the CCR9+, but not CCR9 pDCs, potently inhibited immune responses in vivo in an antigen-driven immunization model and an animal model of acute GVHD induced by the transplantation of allogeneic bone marrow. Immune suppression by CCR9+ DCs involved the inhibition of T cell proliferation and inflammatory cytokine production, which presumably reflected the 'preferential' ability of CCR9+ pDC to induce Foxp3+ regulatory T cells. Our findings suggest that CCR9 expression defines a physiologically important tolerogenic DC subset well positioned in lymphoid tissues to participate in homeostatic immune regulation.

What functional relevance might CCR9 have for the tolerogenic capacity of pDCs? CCR9 expression on pDCs permits their chemotaxis to the CCR9 ligand CCL25, as shown here and in another study34. The two main cell types with abundant CCL25 expression in vivo are the small intestinal epithelium and thymic epithelium35. CCR9 mediates the migration of intestinal memory T cells and immunoglobulin A–positive plasma cells to the small intestines and the migration of T cell precursors to the thymus25,36,37. Thus, it is reasonable to postulate that CCR9 might allow tolerogenic DCs to migrate either to the thymus or the gut, where they can present peripheral antigens and induce T cell tolerance. As CCR9 was rapidly downregulated by pDC-specific TLR ligands, activation of CCR9+ pDCs by infectious agents would eliminate their thymic- or gut-specific homing capabilities, ensuring that they would not induce T cell tolerance to foreign antigens in those sites. Consistent with that hypothesis, CCR9 has been linked to the localization of pDCs to the gut wall34. Our findings, however, indicate a more widespread distribution and function of CCR9+ pDCs. Published work has suggested that resting DCs have the ability to sample tissue-specific antigens and carry them into the thymus, where they induce clonal deletion of antigen-specific T cells38. Although it is possible to propose a critical function for CCR9 on tolerogenic pDCs in this context, in our studies, GVHD was mediated by adoptively transferred mature effector T cells, which rules out central tolerance as an important mechanism of GVHD suppression. The thymic generation of regulatory T cells might be involved but seems unlikely given the time frame required for suppression of the acute graft-versus-host response. Moreover, other studies have found no involvement of CCL25 in the localization of peripheral DCs to the thymus38. These findings collectively rule out the importance of thymic mechanisms in the tolerogenicity of CCR9+ DCs in our model. Instead, our data are more consistent with a mechanism involving CCR9+ pDC–induced development of Foxp3+ regulatory T cells from the mature peripheral T cell pool.

A notable finding of our studies was the long-term suppression of disseminated GVHD by CCR9+ DCs. Studies have shown that lethal GVHD is initiated mainly by alloreactive CD4+ donor T cells39 but that disease can be inhibited by the cotransfer of CD4+CD25+ regulatory T cells of donor origin40,41. These regulatory T cells must recognize alloantigens of the recipient to mediate their protective effects. In our studies, transferred donor CCR9+ DCs were potent inducers of allogeneic Foxp3+ regulatory T cells both in vitro and in vivo. We also found suppressed T cell proliferation in vitro and showed that the ratio of IL-17-producing effector T cells to IFN-γ-producing effector T cells was altered in vivo. In a lymphopenic T cell–mediated mouse model of systemic autoimmunity that resembles GVHD, IL-17 has been shown to be a key mediator of autoimmune pathology, whereas IFN-γ has a protective effect42. Furthermore, regulatory T cells in this setting inhibit the accumulation of effector T cells and attenuate disease42. Other studies have also suggested an inhibitory function for IFN-γ in polyclonal models of GVHD43,44, and cross-regulation between T helper type 1 responses and IL-17 production has also been demonstrated in vitro45,46. Those results42,43,44,45,46 might explain the protective effects of high production of IFN-γ and low production of IL-17 on GVHD noted after cotransfer of tolerogenic CCR9+ pDCs. In summary, our data suggest that donor T cell recognition of host alloantigens on CCR9+ DCs induces regulatory T cells that inhibit the accumulation of IL-17-producing effector T cells and thereby contribute to potent and prolonged disease suppression.

Studies have shown that a variety of DC populations can inhibit immune responses2,15 and can have therapeutic effects in animal models of transplantation4 and autoimmunity47,48,49,50. Although direct comparisons are complicated by the diversity of animal models and DC sources used, two potentially related features distinguish our studies. First, the tolerogenic effects reported before have been impermanent and generally partial, in contrast to the prolonged GVHD suppression we found. Second, no study before has used CCR9, or indeed any rigorous sorting method based on DC activation markers, to purify a tolerogenic subset from immune-stimulatory DCs before therapeutic transfer. Studies of the tolerogenic properties of adoptively transferred mDCs5 or pDCs20,21 have instead used total DCs generated in vitro from bone marrow cells cultured with granulocyte-macrophage colony-stimulating factor or Flt3L, respectively, or unsorted DC populations expanded in vivo with granulocyte-macrophage colony-stimulating factor, granulocyte colony-stimulating factor or Flt3L13,51, as used in our studies here. Full tolerance across MHC barriers in these studies has been achieved only with additional interventions such as pharmacological immunosuppression or antibody treatments to block costimulatory molecules4, and although experimentally quiescent DCs have been used, these cells were not sorted on the basis of immature phenotype. In contrast, by segregating immature pDCs on the basis of their robust CCR9 expression, we achieved 100% survival of irradiated hosts after transfer of these cells with allogeneic bone marrow and effector T cells. Transfer of CCR9 pDCs instead, or no pDC transfer at all, resulted in a vigorous alloimmune response and subsequent wasting due to the graft-versus-host response in most mice. The survival of about 50% of control mice may have reflected incomplete myeloablative conditions, as seen in other studies40. Our results thus suggest that the suppressive effects of tolerogenic DCs may be counteracted by the immune-stimulatory effects of mature DCs present in most experimental DC populations. Our findings may also be related to those of a published report describing the therapeutic use of in vivo–derived CD8α+CD11c+ 'lymphoid-related' DC populations expanded by Flt3L, in which these cells prolong allograft survival in adoptive recipients13. Notably, CD8α expression has been demonstrated on subsets of mouse pDCs, varying according to tissue source and state of activation18,52,53. We consider that these CD8α+ DC populations might include CCR9+ as well as CCR9 pDCs and therefore represent a heterogenous population of pDCs ranging from immature to activated cells13. It will be useful to compare the efficiency and duration of immune suppression by sorted immature and mature pDC populations in this and other models in the future.

In conclusion, here we have used phenotypic criteria, in particular CCR9 expression, to segregate in vivo–derived tolerogenic pDCs from other DCs. We have also shown that this purified subset was very effective in suppressing GVHD. The phenotypic characterization and isolation of tolerance-inducing DC subsets may be of therapeutic benefit in adoptive immunotherapy for a wide range of inflammatory disorders, including autoimmunity, allergic disorders and transplantation.

Methods

Mice.

C57BL/6 (CD45.2), congenic CD45.1 (B6.SJL-Ptprca Pep3b/BoyJ) and BALB/cJ mice were from the Jackson Laboratory. BALB/c.Thy-1.1 congenic mice were bred in the Veterinary Medical Unit facility of the Veterans Affairs Palo Alto Health Care Systems. Mice were housed in specific pathogen–free conditions and were used according to the guidelines set forth by the animal committee of the Veterans Affairs Palo Alto Health Care Systems.

Flow cytometry.

Samples were first incubated with the 2.4.G2 antibody to the Fc receptor (BD Biosciences) in DC studies to prevent nonspecific binding of monoclonal antibody (mAb). The following mAbs were used for staining: peridinine chlorophyll protein complex–conjugated antibody to B220 (anti-B220; RA3-6B2), phycoerythrin–anti-CD11c (HL3), phycoerythrin-indotricarbocyanine–anti-CD3 (145-2C11), phycoerythrin-indotricarbocyanine–anti-CD19 (1D3), biotin-conjugated anti-IA-IE (2G9), fluorescein isothiocyanate–anti-I-Ab (AF6-120.1), allophycocyanin–anti-CD25 (PC61), phycoerythrin–anti-CD4 (RM4-5), peridinine chlorophyll protein complex–cyanine 5.5–anti-CD4 (RM4-5), biotin-conjugated anti-Thy-1.1 (OX-7) and peridinine chlorophyll protein complex–cyanine 5.5–anti-CD3 (145-2C11; all from BD Biosciences), and fluorescein isothiocyanate–conjugated anti-CD40 (HM40-3), anti-CD80 (16-10A1) and anti-CD86 (GL1; all from eBioscience). Allophycocyanin–anti-CCR9 was used according to the manufacturer's recommendations (242503; R&D Systems). Secondary reagents for the visualization of biotinylated mAbs included streptavidin–Pacific blue (Invitrogen).

DC isolation and sorting.

DCs were isolated from the lymphoid tissues of normal C57BL/6 and BALB/c mice by incubation for 1–2 h at 37 °C in collagenase IV (Worthington Biochemical) and DNase I (Sigma) in protein-free media at final concentrations of 500 U/ml and 1 U/ml, respectively. Tissues were resuspended, passed over a wire mesh and washed, then cells were counted and stained with conjugated mAbs. For isolation of Flt3L-expanded DC populations, C57BL/6 mice were injected subcutaneously with 5 × 106 Flt3L-secreting B16 melanoma cells that promote the expansion of DC populations in vivo54. After 14 d, lymph nodes were isolated and passed through a 70-μm nylon mesh. For sorting of pure populations of CCR9+ and CCR9 pDCs, cells were first enriched with CD11c microbeads (Miltenyi), followed by sorting of lineage-negative (CD3CD19) CD11cintB220+ cells on the basis of their CCR9 expression.

Chemotaxis assay.

Pooled lymph node cell suspensions from C57BL/6 mice transplanted with Flt3L-secreting B16 melanoma cells were resuspended in 100 μl complete RPMI-1640 medium and were loaded into collagen-coated Transwell devices (pore size, 5 μm; 3421; Corning) placed in 24-well plates containing 600 μl medium alone or medium supplemented with 250–500 nM CCL25, 100 nM CCL21 or 50 nM CXCL12 (R&D Systems). After plates had incubated for 2 h at 37 °C, a constant number of polystyrene beads (Polysciences) was added to each sample to control for recovery of cells from different wells. Cells that had migrated were collected and counted and were stained with mAb to determine the number of migrated pDCs and mDCs by flow cytometry. The ratio of the number of pDC that migrated in the presence of chemokine to the number of cells that migrated to control media was calculated and is presented as the percentage of migrated cells relative to the input.

DC stimulation with TLR ligands.

After pDCs from pooled PLNs of Flt3L-treated B6 mice were purified by magnetic-activated cell sorting and sorted by CCR9 expression, 0.2 × 106 to 0.5 × 106 pDCs were cultured for 8–12 h in 200 μl complete RPMI-1640 medium supplemented with 10% (vol/vol) FCS in the presence or absence of LPS (1 ng/ml), R848 (10 μg/ml) and CpG (1 μM; ODN1826; all three from Invivogen). After being stimulated, DCs were stained for expression of MHC class II (IA-IE) and CD80, CD86 or CD40.

Intracellular Foxp3 and cytokine assays.

Single-cell suspensions of lymph node cells and erythrocyte-free splenocytes were stimulated in vitro for 4 h at 37 °C with phorbol 12-myristate 13-acetate (5 ng/ml; Sigma) and ionomycin (1 μg/ml; Sigma). Brefeldin A (eBioscience) was added at a final concentration of 1 μg/ml at 2 h after the addition of phorbol 12-myristate 13-acetate and ionomycin. Cells were collected and their surfaces were stained with peridinine chlorophyll protein complex–cyanine 5.5–anti-CD4 and biotin-conjugated anti-Thy-1.1, followed by the secondary reagent, streptavidin–Pacific blue (Invitrogen). For visualization of Foxp3, cells were not stimulated. After surface staining, cells were washed, fixed and made permeable according to the manufacturer's recommendation (eBioscience). Cells were then stained in permeabilization buffer (eBioscience) with the following fluorochrome-labeled mAbs: fluorescein isothiocyanate–anti-IFN-γ (XMG1.2; eBioscience), phycoerythrin–anti-IL-17 (TC11-18H10; BD Biosciences) and allophycocayanin–anti-IL-10 (JES5-16E3; BD Biosciences), or fluorescein isothiocyanate–anti-Foxp3 (for the visualization of regulatory T cells; FJK-16s; eBioscience). Cells were washed in permeabilization buffer and were resuspended in staining buffer for analysis on a flow cytometer. Supernatants of overnight (16-hour) cultures of pDCs stimulated with TLR ligands were examined for the presence of IFN-α with a standardized kit (PBL Biomedical Laboratories) or for TNF by Luminex bead technology with a standardized kit (Millipore).

In vitro T cell stimulation and suppressor T cell assays.

CD4+ T cells were enriched from spleens of BALB/c mice with a CD4+ T cell isolation kit (Miltenyi) and were cultured with CCR9+ and CCR9 pDCs, sorted from pooled lymph nodes of Flt3L-treated B6 mice, at a ratio of 5:1. For analysis of T cell proliferation, cultures were set up in 96-well flat-bottomed microtiter plates with 2 × 105 sorted T cells and 0.4 × 105 DCs and were stimulated for 3 d with a 'dose range' of pOVA before the addition of [3H]thymidine (1 μCi/well). After a further 18 h, cultures were collected and [3H]thymidine incorporation was measured with a liquid scintillation β-counter (Wallac). Results are presented as mean counts per minute of triplicate cultures. For T cell–suppression assays, cultures were set up for 5–7 d with larger numbers of sorted T cells (5 × 106) and DCs (1 × 106). Aliquots of T cells were analyzed for expression of CD25 and Foxp3 as described above. The remaining cells were cultured together with CD4+CD25 effector T cells isolated from spleens of BALB/c mice by negative selection over LD columns (Miltenyi) with the CD4+CD25+ regulatory T cell isolation kit (Miltenyi). Cultures were set up for 48 h in 96-well plates coated with anti-CD3 (3 μg/ml; 2C11; eBioscience) and anti-CD28 (3 μg/ml; 37.51 clone; eBioscience) before the addition of [3H]thymidine and subsequent analysis of T cell proliferation. Results are presented as mean counts per minute of triplicate cultures.

Adoptive transfer and immunization.

CCR9+ and CCR9 pDCs were sorted from pooled lymph node cells from Flt3L-treated B6 mice and were cultured for 2–4 h with 50 μM pOVA before being administered intravenously to naive C57BL/6 mice (0.5 × 106 DCs per mouse). Recipient mice were boosted 1 week later with the same antigen-loaded pDCs (0.5 × 106 DCs per mouse) and were immunized subcutaneously 1 week after the final boost with 20 μg pOVA emulsified in complete Freund's adjuvant (Sigma). After 10 d, cell suspensions from draining lymph nodes, in 96-well plates at a density of 0.5 × 106 lymph node cells per well, were stimulated for 72 h with a 'dose range' of pOVA before the addition of [3H]thymidine and subsequent analysis of cellular proliferation. Results are presented as mean counts per minute of quadruplicate cultures.

GVHD model.

C57BL/6 hosts were given total-body irradiation twice, 4 h apart, with a 131Cs source at 450 rads per dose for a cumulative dose of 900 rads. Irradiated mice were injected intravenously with donor cells within 24 h. All mice received 2 × 106 cells of T cell–depleted bone marrow with 0.5 × 106 to 1 × 106 splenic CD4+ CD25 donor T cells, both from BALB/c mice. Bone marrow was depleted of T cells with anti-Thy-1.2 microbeads followed by negative selection through LD columns (Miltenyi). Samples were enriched for CD4+CD25 effector T cells with the CD4+CD25+ regulatory T cell isolation kit (Miltenyi) by purification of total CD4+ T cells followed by negative selection of CD4+CD25+ T cells through LD columns (Miltenyi). MACS bead enrichment of CD4+ CD25 T cells and T cell–depleted bone marrow resulted in over 99% elimination of potential CD4+CD25+ regulatory and CD3+ effector T cells, respectively. Some groups received, in addition to T cells and bone marrow, 0.2 × 106 to 0.5 × 106 sorted CCR9+ or CCR9 pDCs from pooled lymph nodes of Flt3L-treated B6 mice. For the analysis of effector T cells after transfer, CD4+CD25 effector T cells were isolated from BALB/c.Thy-1.1 congenic mice. Mice were given water containing antibiotics for the first month. The survival and appearance of mice were monitored daily and body weight was measured weekly. For the analysis of effector T cell responses in vivo, mice were evaluated on days 10, 20 and 30 for cytokine production and induction of regulatory T cells in lymphoid tissues.

Statistical analysis.

The statistical significance of differences between sets of data was assessed with the two-tailed unpaired Student's t-test for comparison of two groups. The significance of differences in survival curves in GVHD studies was assessed with the log-rank test. P values of less than 0.05 were considered statistically significant.

Accession codes.

UCSD-Nature Signaling Gateway (http://www.signaling-gateway.org): A000632 and A002265.

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Acknowledgements

We thank L. Rott for assistance with flow cytometry and cell sorting; M. BenBarak for assistance with cytokine analysis with Luminex technology; and B. Zabel for discussions. Supported by the Arthritis Foundation (H.H.), the National Institutes of Health (AI07290 to H.H.; R03DK069395 to T.S.; K08DK069385 to A.H.; E.C.B.), the Wenner-Gren Foundation, Sweden (C.O.) and the Veterans Administration (E.C.B.).

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Affiliations

  1. Laboratory of Immunology and Vascular Biology, Department of Pathology, Stanford University School of Medicine, Stanford, California 94305, USA.

    • Husein Hadeiba
    • , Tohru Sato
    • , Aida Habtezion
    • , Cecilia Oderup
    • , Junliang Pan
    •  & Eugene C Butcher
  2. The Center for Molecular Biology and Medicine, Veterans Affairs Palo Alto Health Care System, Palo Alto, California 94304, USA.

    • Husein Hadeiba
    • , Tohru Sato
    • , Aida Habtezion
    • , Cecilia Oderup
    • , Junliang Pan
    •  & Eugene C Butcher

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Contributions

H.H. designed and did most of the experiments and wrote the manuscript; E.C.B. designed experiments and wrote the manuscript; T.S. prepared the irradiated mice, bone marrow and effector T cells in the GVHD studies and monitored the mice; A.H. was involved in the initial microarray studies and characterization of pDCs by flow cytometry; C.O. assisted with the TLR-activated pDC cytokine assays; J.P. helped with RNA preparation and microarray data analysis; and all authors discussed the results and read and provided comments on the manuscript.

Corresponding authors

Correspondence to Husein Hadeiba or Eugene C Butcher.

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https://doi.org/10.1038/ni.1658

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