Human Liver Dendritic Cells Promote T Cell Hyporesponsiveness

doi:10.4049/jimmunol.0803404

J Immunol. Author manuscript; available in PMC 2012 January 10.

Published in final edited form as:

J Immunol. 2009 February 15; 182(4): 1901–1911.

doi: 10.4049/jimmunol.0803404

PMCID: PMC3254024

NIHMSID: NIHMS347029

Human Liver Dendritic Cells Promote T Cell Hyporesponsiveness^¹

Zubin M. Bamboat,^*² Jennifer A. Stableford,^*² George Plitas,^* Bryan M. Burt,^* Hoang M. Nguyen,^* Alexander P. Welles,^* Mithat Gonen,^† James W. Young,^‡ and Ronald P. DeMatteo^*³

^*Hepatopancreatobiliary Service, Memorial Sloan-Kettering Cancer Center, New York, NY 10065

^†Department of Epidemiology and Biostatistics, Memorial Sloan-Kettering Cancer Center, New York, NY 10065

^‡Laboratory of Cellular Immunobiology and Adult Allogeneic Bone Marrow Transplantation Service, Division of Hematologic Oncology, Department of Medicine, Memorial Sloan-Kettering Cancer Center, New York, NY 10065

³ Address correspondence and reprint requests to Dr. Ronald P. DeMatteo, Memorial Sloan-Kettering Cancer Center, Box 203, 1275 York Avenue, New York, NY 10065. Email: dematter/at/mskcc.org

²Z.M.B. and J.A.S. contributed equally to this work.

The publisher's final edited version of this article is available free at J Immunol

See other articles in PMC that cite the published article.

Abstract

The liver is believed to promote tolerance, which may be beneficial due to its constant exposure to foreign Ags from the portal circulation. Although dendritic cells (DCs) are critical mediators of immune responses, little is known about human liver DCs. We compared freshly purified liver DCs from surgical specimens with autologous blood DCs. Liver and blood DCs were equally immature, but had distinct subset compositions. BDCA-1⁺ DCs represented the most prevalent liver DC subset, whereas the majority of peripheral blood DCs were CD16⁺. Upon TLR4 ligation, blood DCs secreted multiple proinflammatory cytokines, whereas liver DCs produced substantial amounts of IL-10. Liver DCs induced less proliferation of allogeneic T cells both in a primary MLR and after restimulation. Similarly, Ag-specific CD4⁺ T cells were less responsive to restimulation when initially stimulated by autologous liver DCs rather than blood DCs. In addition, liver DCs generated more suppressive CD4⁺ CD25⁺FoxP3⁺ T regulatory cells and IL-4-producing Th2 cells via an IL-10-dependent mechanism. Our findings are critical to understanding hepatic immunity and demonstrate that human liver DCs promote immunologic hyporesponsiveness that may contribute to hepatic tolerance.

The liver is charged with regulating differential immune responses to both harmless foreign Ags and invasive pathogens (1). Autoimmune hepatitis and clearance of certain hepatotropic pathogens like hepatitis A represent the liver’s capability to manifest a robust immune response. The overall balance in hepatic immunity, however, appears to favor the induction of immunologic hyporesponsiveness and sometimes tolerance. For example, the lack of systemic inflammation despite continuous exposure to ingested food Ags highlights the liver’s role in oral tolerance (2). The liver is a frequent site of chronic viral and parasitic infection and is the most common site of distant metastatic disease in cancer. Furthermore, whereas HLA-disparate heart, lung, and kidney transplants are readily rejected, the acceptance of allogeneic liver transplants in HLA-mismatched patients is typical. In addition, the occurrence of T cell-mediated rejection in liver transplant recipients is low, and in some cases patients can be weaned off immunosuppression altogether (3). Although these phenomena have led to wide-ranging hypotheses to explain the liver’s tolerogenic predisposition, none have been definitively established in humans.

The liver contains a distinct repertoire of immune cells. It is enriched in NKT cells that make up 30% of all hepatic CD3⁺ lymphocytes in human nonparenchymal cells (NPC)^⁴ compared with less than 1% of PBMCs. Furthermore, the 1:3.5 ratio of CD4⁺ to CD8⁺ T cells in the liver is distinct from the 2:1 ratio in the blood (4). Hepatic NK cells also contain a larger fraction of CD16⁻ cells than does the blood (5). A variety of APCs, including Kupffer cells and dendritic cells (DCs), also reside in the liver.

DCs play a vital role in the initiation of innate and adaptive immune responses, and their role as immune mediators in cancer and infection has been studied extensively (6). More recently, several lines of evidence have emerged supporting the importance of DCs in the induction and maintenance of tolerance (7–9). The use of such tolerogenic DCs in animal models has revealed that their adoptive transfer can limit the incidence of transplant rejection and curb the progression of certain autoimmune diseases (10, 11). In humans, evidence of the tolerogenic capacity of DCs is limited to work using immature, cytokine-generated monocyte-derived DCs (MoDCs) that are prepared ex vivo. Injection of Ag-loaded immature MoDCs into patients decreased immunity and induced the generation of T regulatory cells (T_reg) (12, 13). The precise mechanisms governing development of tolerance in situ in humans, however, remain unknown.

Human DCs have been defined traditionally as HLA-DR⁺ cells that lack the hematopoietic cell lineage (lin) markers CD3, CD14, CD16, CD19, and CD56. DCs have been subdivided into conventional myeloid (CD11c⁺CD123^low) and plasmacytoid (CD11c⁻ CD123^high) populations (14). Four additional surface Ags specific to human DCs have further distinguished subsets of DCs. BDCA-2 (CD303) and BDCA-4 (CD304) are expressed on all plasmacytoid DCs, whereas BDCA-1 (CD1c) and BDCA-3 (CD141) are each expressed by a subgroup of conventional myeloid DCs (15, 16). Meanwhile, a population of CD16⁺ DCs has been recognized to comprise the majority of freshly isolated blood DCs (17–21). CD16⁺ DCs are M-DC8⁺(6-sulfo LacNAc⁺)CD11c⁺BDCA-1⁻CD123⁻, which in addition to inducing T cell proliferation, mediate Ab-dependent cellular cytotoxicity, stimulate NK cells, and produce large amounts of TNF-α and IL-12p70 in response to LPS (18, 19, 21). Overall, at least four distinct DC subgroups are now recognized after fresh isolation from human peripheral blood: BDCA-1⁺, BDCA-3⁺, CD16⁺, and CD123^high.

The characterization of human liver DCs and the precise role they play in modulating the distinct immune environment within the liver have yet to be fully elucidated. To investigate whether DCs contribute to the tolerogenic environment of the liver, we analyzed freshly isolated DCs from human liver in 56 patients and compared them with freshly isolated autologous blood DCs.

Materials and Methods

Cell isolation

Fresh blood and liver samples were collected from 56 individual patients undergoing elective hepatic resection at Memorial Sloan-Kettering Cancer Center. Operations were performed for benign disease (n = 7), primary cancer (n = 9), and metastatic disease (n = 40). Informed consent was obtained according to an Institutional Review Board-approved protocol. Blood was drawn intraoperatively, and PBMC were isolated by density centrifugation over Ficoll-Paque^Plus (LPS free; GE Healthcare). Liver tissue was procured at least 5 cm away from any tumor and processed immediately following removal from the patient. To remove contaminating circulating blood cells, liver vessels were flushed with HBSS containing type IV collagenase (1 mg/ml; Sigma-Aldrich), DNase (50 ng/ml; Roche Diagnostics), and 2% endotoxin-free human AB serum (Invitrogen). Liver tissue was then morselized and digested in the collagenase solution at 37°C for 30 min. The digestion was quenched with cold HBSS containing DNase (25 ng/ml), and then passed through a 100-μm filter. The cell suspension was centrifuged twice at 250 × g for 10 min to remove fat and debris. The cell pellet was then reconstituted with HBSS and DNase, and the parenchymal hepatocytes were removed with a low speed spin (30 × g for 1 min). Mononuclear cells from the remaining NPC were isolated by Ficoll-Paque^Plus density centrifugation.

Spleen samples were processed directly following removal from patients undergoing elective pancreatic resections in which the spleen was also removed. There was no pathologic evidence of disease in any of the spleen specimens removed. Five gram samples were digested and processed by an identical process, as described above. After quenching and filtering, the cell suspension was centrifuged at 300 × g for 7 min. The supernatant was discarded, and the cell pellet was resuspended in PBS and layered over a Ficoll-Paque^Plus density gradient, after which the interface layer was harvested.

Flow cytometry

Liver NPC and PBMCs were stained with FITC, PE, PerCP-Cy5.5, allophycocyanin, allophycocyanin-Cy7, and AF700 conjugated to Abs against CD3 (UCHT1), CD4 (RPA-T4), CD14 (M5E2), CD19 (HIB19), CD56 (NCAM 16.2), HLA-DR (L243), CD11c (B-ly6), CD16 (3G8), CD34 (581), CD123 (9F5), CD40 (5C3), CD80 (L307.4), CD25 (M-A251), CD83 (HB15e), CD86 (2331) (all BD Pharmingen), BDCA-1 (AD5-8E7), and BDCA-3 (AD5-14H12) (Miltenyi Biotec). We defined a set of lin-specific markers using a combination of anti-CD3, anti-CD14, anti-CD19, and anti-CD56 Abs. Appropriate isotype controls were used. Flow cytometry was performed on a FACS Aria flow cytometer (BD Biosciences), and data were analyzed with FlowJo software (Tree Star). Intracellular FoxP3 expression was assessed using the FITC anti-human FoxP3 staining kit (eBiosciences), and rat IgG2a FITC (BD Biosciences) was used as an isotype control. Sorting was performed on a MoFlo (DakoCytomation) or BD FACSAria (BD Biosciences) cell sorter. Bulk DCs were defined as lin⁻HLA-DR⁺. Postsort purities were routinely >98%.

Ag uptake

A total of 1 × 10⁶ blood PBMCs and liver NPC was incubated at 37°C for 0, 5, 15, or 30 min in a solution of PBS with 1 mg/ml 10,000 m.w. FITC dextran or 50 μg/ml DQ OVA (Molecular Probes). The cells were then quenched with ice-cold PBS, washed twice, stained with Abs to lin and DC surface markers, and analyzed on a FACSCalibur flow cytometer (BD Biosciences).

Allogeneic MLR and coculture cytokine measurement

FACS-purified bulk liver and blood DCs were cultured in triplicate at varying concentrations with 1 × 10⁵ allogeneic naive CD4⁺CD45RA⁺ T cells purified by immunomagnetic selection using the naive CD4⁺ T cell isolation kit (Miltenyi Biotec). The T cells were isolated from donor buffy coat leukocytes from healthy adult volunteers (Greater NY Blood Center, American Red Cross). Cells were routinely greater than 93% pure (for CD4⁺ and CD45RA⁺) and displayed no FoxP3 staining. On day 5 of coculture, supernatants were harvested and cytokine levels were measured using the human Th1/Th2 cytometric bead array kit (BD Biosciences). Cells were then pulsed with [³H]thymidine (1 μCi/well; PerkinElmer), and radioactive uptake was measured 18 h later with a TopCount NXT micro-plate scintillation and luminescence counter (PerkinElmer). For secondary MLRs, liver and blood DC-primed T cells were harvested on day 6, recounted, and restimulated with irradiated (2500 rad) PBMCs that were enriched for APCs by depletion of CD3⁺ cells using immunomagnetic beads (Miltenyi Biotec). These CD3⁻ PBMC APCs for restimulations were from the same donor as the DCs used in the primary MLR. [³H]Thymidine (1 μCi/well) was added to wells at the indicated time points after restimulation, and proliferation was measured 18 h later.

DC cytokine measurement

A total of 5 × 10⁴ freshly sorted bulk DCs or DC subsets from liver or blood was cultured in triplicate with medium alone or with LPS (1 μg/ml from Escherichia coli 055:B5; Sigma-Aldrich). Medium consisted of RPMI 1640, 2 mM L-glutamine, 1% nonessential amino acids, 100 U/ml penicillin, 100 μg/ml streptomycin, 0.1% 2-ME, and 10% heat-inactivated, endotoxin-free human AB⁺ serum (Media Preparation Core Facility, Memorial Sloan-Kettering Cancer Center). After 24 h, supernatant was harvested and cytokine levels were measured using the human inflammation cytometric bead array kit (BD Biosciences).

Suppression assay

For isolation of T cells primed by liver DCs for 7 days, staining was performed using anti-CD4 (RPA-T4), and anti-CD25 (M-A251) conjugated to PE or allophycocyanin, respectively. The CD4⁺CD25^high (2–3% of highest CD25⁺ cells) and CD4⁺CD25⁻ fractions were purified via FACS and used as suppressors or negative controls, respectively. Naive CD4⁺CD45RA⁺ T cells were freshly isolated by immunomagnetic selection (Miltenyi Biotec) and labeled with CFSE (5 μM; Invitrogen). These naive T cell responders were cultured alone or in the presence of anti-CD3/CD28 mAb-coated Dynabeads (Invitrogen; 2:1 bead-total cell ratio) with varying concentrations of CD4⁺ CD25^high or CD4⁺ CD25⁻ cells. Proliferation of the CFSE-labeled cells was determined after 2 or 3 days of coculture.

CD4⁺ T cell stimulation and analysis of T cell cytokine production

A total of 5 × 10⁴ allogeneic peripheral blood naive CD4⁺CD45RA⁺ T cells isolated from buffy coats was cultured in triplicate with FACS-purified bulk blood or liver DCs (DC:T cell ratio of 1:10) for 7 days. In some cultures, IL-10-blocking Ab (1 μg/ml; BD Biosciences) was added. Rat IgG1 (OX86, 1 μg/ml; mAb Core Facility, Memorial Sloan-Kettering Cancer Center) was used as an isotype control. Cells were collected after 7 days of primary stimulation with DCs, recounted, and then restimulated with 5 μg/ml plate-bound anti-CD3 and 1 μg/ml soluble anti-CD28 (BD Biosciences) for 24 h. Culture supernatants were harvested 18 h after restimulation, and the levels of IL-4 and IFN-γ were measured using a human Th1/Th2 cytometric bead array (BD Biosciences). For intracellular cytokine analysis, the primed CD4⁺ T cells were restimulated, as described above, with the addition of brefeldin A (10 μg/ml; BD Biosciences) during the final 6 h of culture. The cells were then harvested and stained with a combination of fluorochrome-labeled Abs to CD3 and CD4, fixed, permeabilized, and stained with 0.5 μg of FITC-conjugated anti-IFN-γ, 0.03 μg of PE-conjugated anti-IL-4, or the appropriate isotype controls (BD Biosciences).

Ag-specific CD4⁺ T cell assay

A total of 2.0 × 10⁵ to 4 × 10⁵ FACS-purified bulk liver and blood DCs was pulsed for 3 h with a pool of 23 peptides corresponding to HLA class II-restricted T cell epitopes from CMV, EBV, influenza virus, and tetanus toxin (selected MHC class II Peptide Pool^Plus, 40 μg/ml; Axxora). Autologous peripheral blood CD4⁺ T cells were obtained using the CD4⁺ T isolation kit (Mitenyi Biotec). Peptide-pulsed DCs were washed twice in PBS and cultured with CD4⁺ T cells at a ratio of 1:10 for 7 days. IL-2 (12.5 IU/ml; R&D Systems) was added on day 2 and every other day for 7 days. Blood and liver DC-primed T cells were restimulated on day 7 using irradiated (2500 rad), peptide-pulsed (10 μg/ml) autologous CD3⁻ PBMCs at a ratio of 1:1. ELISPOT assays were performed 7 days later after 24 h of restimulation with irradiated, peptide-pulsed CD3⁻ PBMCs at a ratio of 2 APC:1 CD4⁺ T cell in X-VIVO 15 serum-free medium (BioWhittaker). Induction of IFN-γ and TNF-α production was measured using the human IFN-γ and TNF-α ELISPOT kits (BD Biosciences), and cells were counted with an automated immunospot device (Cellular Technology). T cells cultured in PHA (5 μg/ml; Sigma-Aldrich) were used as positive controls.

Statistics

Data were analyzed for statistical significance with the nonparametric, unpaired, two-tailed Mann-Whitney U test or the two-tailed Wilcoxon matched pairs test, as indicated (Prism statistical software; GraphPad). Values of p <0.05 were considered statistically significant in all cases except Fig. 5C, in which the α level was adjusted via the Bonferroni method to avoid an inflated type I error rate. In Fig. 5C, in which two simultaneous comparisons were made, p values <0.025 were considered statistically significant.

FIGURE 5

Liver, blood, and spleen DC cytokine production. FACS-purified, freshly isolated bulk liver and autologous blood DCs (A), bulk spleen and autologous blood DCs (B), or liver DC subsets (C) were cultured in medium or LPS (A) or LPS alone (B and C). Supernatant (more ...)

Results

Bulk liver DCs have different subset composition than blood DCs

The subset composition of freshly isolated human liver DCs is unknown. We therefore analyzed liver lin⁻HLA-DR⁺ cells for the DC subsets known to exist in the blood. All four major subsets of freshly isolated blood DCs were also present in the liver (Fig. 1A). CD16⁺ DCs were more prevalent in the blood (median 64 vs 30%), whereas BDCA-1⁺ DCs were more common in the liver (median 36 vs 15%). The percentage of CD123^high DCs was similar between blood and liver (median 13 vs 15%), whereas BDCA-3⁺ DCs (median 3% in blood and 6% in liver) were infrequent (Fig. 1B). Overall, the trend of liver DC composition and the differences between liver and blood DCs were similar among patients, except for one patient with hemochromatosis and hepatocellular carcinoma who had a high proportion of CD123^high DCs in both sites (Fig. 1B). The average yields of hepatic NPC and PBMCs were 1.8 × 10⁶ cells/g liver and 1.2 × 10⁶ cells/ml blood processed, respectively. The average yield of bulk DCs after sorting was 4.2 × 10³ cells/g liver and 4.3 × 10³ cells/ml peripheral blood (Fig. 1C).

FIGURE 1

Liver and blood DCs have distinct composition. A, DCs were gated as lin⁻HLA-DR⁺ cells from freshly isolated PBMCs and hepatic NPC. DC subsets were then identified based on the expression of CD11c, CD16, BDCA-1, CD123, and BDCA-3, compared with (more ...)

Liver DCs are immature and inefficient at Ag capture and processing

Maturation is critical in determining the functional ability of DCs to orchestrate protective immunity to infection and tumors (22, 23). Despite the differences in subset composition between bulk liver and blood DCs, we found that freshly isolated populations expressed similarly high surface levels of HLA-DR, low levels of CD86, and essentially no CD40, CD80, or CD83 (Fig. 2, A and B). Specific liver and blood DC subsets also had similar maturation. Nevertheless, using FITC-conjugated dextran to investigate Ag uptake, we found that bulk, CD16⁺, and BDCA-1⁺ DCs from the liver were less efficient at Ag uptake (Fig. 2, C and D). Additional experiments using DQ OVA, which requires uptake and proteolytic cleavage to become fluorescent, revealed that bulk liver DCs as well as the CD16⁺ and BDCA-1⁺ DC subsets, were less efficient at Ag processing than bulk blood DCs and their respective subsets (Fig. 2, C and D). Thus, liver DCs were as phenotypically immature as blood DCs, but functionally less able to capture and process Ag.

FIGURE 2

Liver and blood DCs are equally immature, but liver DCs acquire and process Ag less efficiently than blood DCs. A, The expression of maturation markers of freshly isolated bulk liver and blood DCs (open histograms) relative to their respective isotype (more ...)

Liver DCs induce less alloproliferation and promote T cell hyporesponsiveness

One of the hallmarks of DC function is the ability to stimulate naive T cells. Freshly isolated bulk liver DCs were weaker than autologous bulk blood DCs at inducing the proliferation of allogeneic naive CD4⁺ T cells (Fig. 3A). Furthermore, supernatants harvested from MLR cultures stimulated by bulk liver DCs contained significantly lower levels of IFN-γ and TNF-α and a greater amount of IL-10 (Fig. 3B). To determine whether disparity in the isolation procedure for blood and liver DCs may account for the observed differences between the two DC populations, we subjected whole blood to the collagenase-based procedure we used to isolate liver DCs. Both subset composition and activation state of blood DCs were unchanged (Fig. 3C). In addition, there was no difference in T cell allostimulation by blood DCs isolated by the collagenase method compared with autologous blood DCs isolated via the standard protocol described in Materials and Methods (Fig. 3D).

FIGURE 3

Liver DCs are weaker T cell stimulators. A, FACS-purified, freshly isolated bulk liver and blood DCs were cultured in varying concentrations with 1 × 10⁵ allogeneic naive CD4⁺CD45RA⁺ T cells. DCs alone (DC) and T cells alone (T) served as negative (more ...)

To determine whether priming by liver or blood DCs had a differential effect on T cell proliferative capacity following re-stimulation, we performed secondary MLRs. Irradiated PBMCs enriched for APCs by depletion of CD3⁺ cells were used to re-stimulate liver or blood DC-primed T cells isolated from primary MLRs. Liver DC-primed naive CD4⁺ T cells were significantly less responsive to restimulation than blood DC-primed T cells at serial time points and at different APC concentrations (Fig. 4).

FIGURE 4

Liver DCs render T cells less responsive to restimulation. A, Liver and blood DC-primed T cells from a 6-day MLR as in Fig. 3A were isolated and restimulated with irradiated CD3⁻ PBMCs (APCs) from the same DC donor of the primary MLR. Thymidine (more ...)

Liver DCs make substantial quantities of IL-10

To investigate why liver DCs accomplished less alloproliferation, we examined their ability to produce cytokines that may modulate their interaction with T cells. Consistent with the previous findings, freshly sorted blood DCs activated by LPS produced high levels of inflammatory cytokines, but nearly undetectable amounts of IL-10 (24). In contrast, liver DCs made 10 times more IL-10 than blood DCs upon culture alone and over 1 ng/ml IL-10 after culture with LPS (Fig. 5A). In addition, liver DCs made much less IL-1β, IL-6, and TNF-α than blood DCs after activation. To determine whether the differences in cytokine production were unique to liver DCs, we performed similar experiments with human spleen and autologous blood DCs. Although bulk spleen DCs made more IL-10 than their blood counterparts, they made comparable amounts of inflammatory cytokines upon LPS stimulation (Fig. 5B) and did not induce T cell hyporesponsiveness (data not shown). Further analysis of liver DCs revealed that the liver BDCA-1⁺/BDCA-3⁺ DC subsets were largely responsible for IL-10 production, whereas liver CD16⁺ DCs accounted for IL-1β, IL-6, and TNF-α secretion (Fig. 5C). As expected, CD123^high DCs did not produce cytokines in response to LPS because they do not express TLR4.

Liver DC IL-10 enhances Th2 polarization

IL-10 is a potent immunosuppressive cytokine and is known to shape CD4⁺ Th cell responses (25). We therefore tested whether IL-10 from liver DCs conditions naive CD4⁺ T cells. After polyclonal restimulation of naive CD4⁺ T cells, which had been cultured for 7 days with allogeneic DCs freshly isolated from either liver or blood, we found that liver DCs generated a substantially lower percentage of IFN-γ⁺ T cells (Fig. 6A). This effect depended on IL-10, because the presence of an IL-10-blocking Ab restored the percentage of IFN-γ⁺ T cells to that generated by blood DCs (Fig. 5A). Pooled data from multiple patients revealed that the average percentage of IFN-γ⁺ T cells in liver DC cocultures was one-third of that in blood DC-T cell cultures (Fig. 6B). We substantiated our intracellular cytokine data by measuring the amount of supernatant IFN-γ protein from T cells that were primed for 7 days with either freshly isolated blood or liver DCs and then polyclonally restimulated for 24 h. Liver DC-T cell cultures had significantly lower levels of IFN-γ unless IL-10 was blocked (Fig. 6C). These findings were consistent with the lower supernatant IFN-γ levels in the liver DC MLR cultures (Fig. 3B). In addition to the decreased Th1 cytokine profile, liver DC-T cell cultures yielded significantly higher percentages of IL-4⁺ T cells (Fig. 6, D and E). Similarly, supernatant amounts of IL-4 following poly-clonal restimulation were higher in the T cell cultures primed with liver DCs (Fig. 6F). Consistent with the increase in the Th1 response by liver DCs in the presence of an anti-IL-10 Ab, IL-10 blockade also abrogated the Th2 response induced by liver DCs. Simultaneous analysis of IFN-γ - and IL-4-producing effectors following polyclonal restimulation revealed that the cytokines were produced by distinct CD4⁺ T cells (Fig. 6G).

FIGURE 6

Liver DCs induce fewer IFN-γ⁺ effectors and promote a Th2 response. A, Freshly isolated bulk liver or blood DCs were cultured for 7 days with allogeneic naive CD4⁺CD45RA⁺ T cells in the presence or absence of anti-IL-10 or an isotype control Ab. (more ...)

Liver DCs promote the induction of T_reg

To investigate additional immunomodulatory effects of liver DCs, we analyzed MLR cultures on day 7 for the presence of CD4⁺FoxP3⁺ T cells. We found that blood DC-T cell cultures contained relatively low percentages of CD4⁺FoxP3⁺ T cells, whereas liver DC cultures generated up to 6% CD4⁺FoxP3⁺ T cells (Fig. 7A). Further phenotypic characterization of the CD4⁺FoxP3⁺ T cells that had been primed by fresh liver DCs revealed that 98 ± 2% stained positively for CD25 (data not shown). The generation of CD4⁺FoxP3⁺ T cells in liver DC-T cell cultures depended on IL-10, as demonstrated with a blocking IL-10 Ab (Fig. 7A). Data pooled from multiple patients revealed that the percentage of CD4⁺FoxP3⁺ T cells generated by liver DCs was 3 times more than blood DCs (Fig. 7B). Similarly, the absolute number of CD4⁺FoxP3⁺ T cells in the liver DC-T cell cultures also increased on day 7 in an IL-10-dependent manner (Fig. 7C). To confirm the regulatory function of the CD4⁺FoxP3⁺ T cells, we sorted CD4⁺CD25^high and CD4⁺CD25⁻ T cells from the liver DC-T cell cultures and incubated them with third-party allogeneic naive CD4⁺ T cells and anti-CD3/CD28 beads. CD4⁺ CD25^high T cells were able to inhibit T cell proliferation in a dose-dependent manner as measured by CFSE dilution (Fig. 7, D and E).

FIGURE 7

Liver DCs promote the generation of T_reg. A, Freshly isolated bulk liver or blood DCs were cultured for 7 days with allogeneic naive CD4⁺CD45RA⁺ T cells, at which time the expression of intracellular FoxP3 protein was determined. Contour plots are shown (more ...)

Liver DCs induce fewer Ag-specific CD4⁺ T cell effectors

To extend our findings that allogeneic T cells conditioned by liver DCs were hyporesponsive after restimulation (Fig. 4), we tested how Ag-specific stimulation of CD4⁺ T cells by liver DCs affected subsequent Ag rechallenge. Sort-purified liver and blood DCs were pulsed with a pool of HLA class II-restricted peptides and then cultured with bulk CD4⁺ T cells for 7 days. T cells were then incubated with autologous, peptide-loaded, CD3⁻ PBMCs for another 7 days. An ELISPOT assay was then performed after 24 h of restimulation with Ag-loaded CD3⁻ PBMCs. Presentation of such class II-restricted peptides by APCs can induce the secretion of IFN-γ and TNF-α by Ag-specific CD4⁺ T cells. We consistently found that liver DCs induced significantly fewer IFN-γ - and TNF-α -producing cells when compared with blood DCs (Fig. 8). The differential effect on cytokine production was abrogated when CD4⁺ T cells stimulated by liver and blood DCs were restimulated nonspecifically with PHA.

FIGURE 8

Liver DCs generate fewer Ag-specific CD4⁺ T cells. Au-tologous Ag-specific CD4⁺ T cells recognizing a pool of HLA class II-restricted peptides were generated by two rounds of stimulation. The frequencies of IFN-γ (A)- and TNF-α (B)-producing (more ...)

Discussion

Our understanding of human DCs is based almost exclusively on cultured peripheral blood DCs and various DC types generated with cytokines in vitro. There are scant data on the function of DCs from human organs because of limited access to tissue specimens and difficulty in isolating sufficient numbers of cells for analysis. It is becoming increasingly apparent, however, that organ-specific immune responses differ and adapt to their local environment. For example, the lung and gut, which are laden with foreign Ags and symbionts, display unique regulatory mechanisms in response to TLR activation (26, 27). Similarly, to protect against potentially harmful immune responses, the liver must adapt to the portal circulation, which is rich in endotoxin as well as harmless foreign Ags. Our results highlight the importance of studying organ-specific immunity and demonstrate that freshly isolated liver DCs most likely contribute to hepatic tolerance based on their distinct subset composition and function.

Although we showed that the majority of freshly isolated blood DCs were CD16⁺, as previously reported (20), BDCA-1⁺ DCs were the most prevalent subset within the liver. Similarly, a prior report of two patients also found that lin⁻CD11c⁺CD16⁻ CD123^low hepatic DCs (which include BDCA-1⁺ and BDCA-3⁺ DCs) constituted the majority of liver DCs (28). Not only did liver and blood DC composition differ, but liver DC composition was also distinct from spleen (data not shown). It is unclear whether differences in subset composition within the liver are a consequence of selective recruitment of DC subsets or the result of differential intrahepatic development from circulating DC precursors. Importantly, the differential results between liver and blood DCs did not depend on the method of isolation.

The distinct cytokine profile of liver DCs compared with both blood and spleen DCs supports the hypothesis that DCs undergo local conditioning within the liver. Previous reports of cytokine production by human liver DCs have been confounded by over 50% contamination with CD14⁺ cells, which contain Kupffer cells (29, 30). We found that activated liver DCs produced IL-10, but comparatively low levels of proinflammatory cytokines. The predominance of liver BDCA-1⁺ DCs and their production of IL-10 may be relevant to chronic viral infection within the liver. Natterman et al. (31) demonstrated an elevation in the number of intrahepatic BDCA-1⁺ cells in patients with chronic hepatitis B or C when compared with normal subjects. In addition, in mouse models of chronic lymphocytic choriomeningitis, reduction of DC IL-10 production by a blocking Ab was sufficient to restore T cell function and eliminate viral infection (32). In contrast, the potential hepatoprotective role of IL-10 has been revealed using murine models of Con A-induced hepatitis, because IL-10 blockade resulted in increased hepatic inflammation and higher systemic levels of proinflammatory cytokines (33).

Liver DCs promoted the induction of functional T_reg from naive CD4⁺ precursors. Their immature state alone does not explain the capacity of fresh liver DCs to generate T_reg, because freshly isolated blood DCs were equally immature (Fig. 2, A and B). The generation of T_reg de novo by liver DCs is consistent with previous reports of CD4⁺ CD25⁺ T_reg generated from naive (CD4⁺ CD25⁻ CD45RA⁺) T cell precursors using MoDCs (34). Liver DC-derived T_reg do not appear to fall into any of the currently known T_reg subsets. Their CD4⁺CD25⁺FoxP3⁺ phenotype is consistent with the naturally occurring subset of T_reg induced in the periphery, but their IL-10 dependency suggests that they may also share some similarities to the type 1 T regulatory subset of T_reg (35). Immature MoDCs possess the capacity to induce the generation of type 1 T_reg from allogeneic naive CD4⁺ precursors, but only following repetitive rounds of in vitro restimulation (7, 36). Furthermore, MoDCs rendered tolerogenic via treatment with exogenous IL-10 have also been shown to induce the differentiation of T_reg in vitro (37). The fact that freshly isolated liver DCs reproducibly generated higher numbers of T_reg in the absence of any exogenous treatment suggests that they may possess an inherent tolerogenic capacity. Additional studies will determine whether T_reg induced by liver DCs comprise a known or entirely novel subset of CD4⁺ T_reg.

In combination with an increase in T_reg, liver DCs simultaneously promoted a Th2 response. Although there is an extensive understanding of the specific pathways governing the differentiation of Th1 cells, the events controlling Th2-driven responses remain unclear. Although there are several models for the selective induction of a Th2 response by DCs, none have been definitively established (38). The current paradigm for DC-driven Th cell development is based on a cascade of events that occur after maturation of resting DCs (39). Notably, we found that bulk liver and blood DCs had similar increases in maturation during culture (data not shown). The three seminal events thought to mediate naive CD4⁺ T cell polarization include the following: stable Ag-MHC-II presentation, adequate costimulation, and a third signal, which in the case of Th1 differentiation is secretion of IL-12p70. Although alternate models exist that propose different sets of signaling cascades for both Th1 and Th2 development (40, 41), none have conclusively determined the third signal controlling Th2 development. Our data demonstrate that the Th2-polarizing capacity of liver DCs depends on IL-10. Evidence supporting such a role for IL-10 in murine models has been well established, because previous reports have revealed that IL-10 production by DCs in hepatic NPC and Peyer’s patches promoted Th2 responses (42, 43). Similarly, work on human MoDCs showed that IL-10 released after TLR2 stimulation was responsible for blocking the production of a number of Th1 cytokines that would otherwise be produced by stimulating MoDCs with TLR3 or 4 (44).

Although we provide direct functional assessment of human liver DCs supporting their tolerogenic role, this study does not rule out the possibility that other organ-resident DCs may exhibit similar properties. Our data are consistent with reports by Lu et al. (45), who provided early insight into the inherent tolerogenicity of murine liver DCs. Furthermore, in congruence with important observations on the regulatory role of liver DCs and the significance of donor-derived DCs in animal models of liver transplantation, our results lend support to the notion that liver DCs are critical regulators of alloimmunity and may contribute to liver allograft acceptance in humans (46–48). We demonstrate that freshly isolated human liver DCs promote T cell hyporesponsiveness after allogeneic and Ag-specific restimulation. The relative proportions of CD16⁺ and BDCA-1⁺ DC subsets are reversed in the liver compared with freshly isolated blood DCs. Paradoxically, liver DCs secrete more IL-10 and less proinflammatory cytokines such as IL-1β and TNF-α, which is unexpected from a population containing a large proportion of BDCA-1⁺ conventional DCs. Finally, freshly isolated liver DCs simultaneously promote a Th2 response and generate suppressive allogeneic CD4⁺FoxP3⁺CD25⁺ T_reg by an IL-10-dependent mechanism.

Although these properties of liver DCs may limit unnecessary immune responses to Ags in the portal circulation or to hepatic allografts, they may hinder immune responses to pathogens or tumors. Our results, therefore, have important implications to understanding hepatic immunity and should facilitate the development of therapeutic strategies for the treatment of liver diseases.

Acknowledgments

We are grateful to J. Hendrikx, M. Kweens, and P. Anderson (all from Flow Cytometry Core Facility, Memorial Sloan-Kettering Cancer Center) for their assistance with cell sorting.

Footnotes

¹This work was supported by National Institutes of Health Grants DK068346 and AI70658, (to R.P.D.), CA083070 (to J.W.Y.), Louis Berkowitz Family Foundation (to R.P.D.), and the New York State Empire Clinical Research Investigator Program (to J.A.S.).

⁴Abbreviations used in this paper: NPC, nonparenchymal cell; DC, dendritic cell; lin, lineage; MoDC, monocyte-derived DC; T_reg, T regulatory cell.

Disclosures

The authors have no financial conflict of interest.

References

1. Crispe IN, Giannandrea M, Klein I, John B, Sampson B, Wuensch S. Cellular and molecular mechanisms of liver tolerance. Immunol Rev. 2006;213:101–118. [PubMed]

2. Mowat AM, Parker LA, Beacock-Sharp H, Millington OR, Chirdo F. Oral tolerance: overview and historical perspectives. Ann NY Acad Sci. 2004;1029:1–8. [PubMed]

3. Mazariegos GV, Reyes J, Marino IR, Demetris AJ, Flynn B, Irish W, McMichael J, Fung JJ, Starzl TE. Weaning of immunosuppression in liver transplant recipients. Transplantation. 1997;63:243–249. [PMC free article] [PubMed]

4. Norris S, Collins C, Doherty DG, Smith F, McEntee G, Traynor O, Nolan N, Hegarty J, O’Farrelly C. Resident human hepatic lymphocytes are phenotypically different from circulating lymphocytes. J Hepatol. 1998;28:84–90. [PubMed]

5. Hata K, Zhang XR, Iwatsuki S, Van Thiel DH, Herberman RB, Whiteside TL. Isolation, phenotyping, and functional analysis of lymphocytes from human liver. Clin Immunol Immunopathol. 1990;56:401–419. [PubMed]

6. Steinman RM, Banchereau J. Taking dendritic cells into medicine. Nature. 2007;449:419–426. [PubMed]

7. Jonuleit H, Schmitt E, Schuler G, Knop J, Enk AH. Induction of interleukin 10-producing, nonproliferating CD4⁺ T cells with regulatory properties by repetitive stimulation with allogeneic immature human dendritic cells. J Exp Med. 2000;192:1213–1222. [PMC free article] [PubMed]

8. Bonifaz L, Bonnyay D, Mahnke K, Rivera M, Nussenzweig MC, Steinman RM. Efficient targeting of protein antigen to the dendritic cell receptor DEC-205 in the steady state leads to antigen presentation on major histocompatibility complex class I products and peripheral CD8⁺ T cell tolerance. J Exp Med. 2002;196:1627–1638. [PMC free article] [PubMed]

9. Liu K, Iyoda T, Saternus M, Kimura Y, Inaba K, Steinman RM. Immune tolerance after delivery of dying cells to dendritic cells in situ. J Exp Med. 2002;196:1091–1097. [PMC free article] [PubMed]

10. Tarbell KV, Yamazaki S, Olson K, Toy P, Steinman RM. CD25⁺ CD4⁺ T cells, expanded with dendritic cells presenting a single autoantigenic peptide, suppress autoimmune diabetes. J Exp Med. 2004;199:1467–1477. [PMC free article] [PubMed]

11. Morelli AE, Thomson AW. Tolerogenic dendritic cells and the quest for transplant tolerance. Nat Rev Immunol. 2007;7:610–621. [PubMed]

12. Dhodapkar MV, Steinman RM, Krasovsky J, Munz C, Bhardwaj N. Antigen-specific inhibition of effector T cell function in humans after injection of immature dendritic cells. J Exp Med. 2001;193:233–238. [PMC free article] [PubMed]

13. Dhodapkar MV, Steinman RM. Antigen-bearing immature dendritic cells induce peptide-specific CD8⁺ regulatory T cells in vivo in humans. Blood. 2002;100:174–177. [PubMed]

14. Shortman K, Liu YJ. Mouse and human dendritic cell subtypes. Nat Rev Immunol. 2002;2:151–161. [PubMed]

15. Dzionek A, Fuchs A, Schmidt P, Cremer S, Zysk M, Miltenyi S, Buck DW, Schmitz J. BDCA-2, BDCA-3, and BDCA-4: three markers for distinct subsets of dendritic cells in human peripheral blood. J Immunol. 2000;165:6037–6046. [PubMed]

16. Dzionek A, Sohma Y, Nagafune J, Cella M, Colonna M, Facchetti F, Gunther G, Johnston I, Lanzavecchia A, Nagasaka T, et al. BDCA-2, a novel plasmacytoid dendritic cell-specific type II C-type lectin, mediates antigen capture and is a potent inhibitor of interferon α/β induction. J Exp Med. 2001;194:1823–1834. [PMC free article] [PubMed]

17. Schakel K, Mayer E, Federle C, Schmitz M, Riethmuller G, Rieber EP. A novel dendritic cell population in human blood: one-step immunomagnetic isolation by a specific mAb (M-DC8) and in vitro priming of cytotoxic T lymphocytes. Eur J Immunol. 1998;28:4084–4093. [PubMed]

18. Schakel K, Kannagi R, Kniep B, Goto Y, Mitsuoka C, Zwirner J, Soruri A, von Kietzell M, Rieber E. 6-Sulfo LacNAc, a novel carbohydrate modification of PSGL-1, defines an inflammatory type of human dendritic cells. Immunity. 2002;17:289–301. [PubMed]

19. Schmitz M, Zhao S, Schakel K, Bornhauser M, Ockert D, Rieber EP. Native human blood dendritic cells as potent effectors in antibody-dependent cellular cytotoxicity. Blood. 2002;100:1502–1504. [PubMed]

20. MacDonald KP, Munster DJ, Clark GJ, Dzionek A, Schmitz J, Hart DN. Characterization of human blood dendritic cell subsets. Blood. 2002;100:4512–4520. [PubMed]

21. Schakel K, von Kietzell M, Hansel A, Ebling A, Schulze L, Haase M, Semmler C, Sarfati M, Barclay AN, Randolph GJ, et al. Human 6-sulfo LacNAc-expressing dendritic cells are principal producers of early interleukin-12 and are controlled by erythrocytes. Immunity. 2006;24:767–777. [PubMed]

22. Mayordomo JI, Zorina T, Storkus WJ, Zitvogel L, Celluzzi C, Falo LD, Melief CJ, Ildstad ST, Kast WM, Deleo AB, et al. Bone marrow-derived dendritic cells pulsed with synthetic tumor peptides elicit protective and therapeutic antitumor immunity. Nat Med. 1995;1:1297–1302. [PubMed]

23. Dhodapkar MV, Krasovsky J, Steinman RM, Bhardwaj N. Mature dendritic cells boost functionally superior CD8⁺ T-cell in humans without foreign helper epitopes. J Clin Invest. 2000;105:R9–R14. [PMC free article] [PubMed]

24. Rescigno M, Martino M, Sutherland CL, Gold MR, Ricciardi-Castagnoli P. Dendritic cell survival and maturation are regulated by different signaling pathways. J Exp Med. 1998;188:2175–2180. [PMC free article] [PubMed]

25. Moore KW, de Waal Malefyt R, Coffman RL, O’Garra A. Interleukin-10 and the interleukin-10 receptor. Annu Rev Immunol. 2001;19:683–765. [PubMed]

26. Rimoldi M, Chieppa M, Salucci V, Avogadri F, Sonzogni A, Sampietro GM, Nespoli A, Viale G, Allavena P, Rescigno M. Intestinal immune homeostasis is regulated by the crosstalk between epithelial cells and dendritic cells. Nat Immunol. 2005;6:507–514. [PubMed]

27. Takabayshi K, Corr M, Hayashi T, Redecke V, Beck L, Guiney D, Sheppard D, Raz E. Induction of a homeostatic circuit in lung tissue by microbial compounds. Immunity. 2006;24:475–487. [PubMed]

28. Wertheimer AM, Bakke A, Rosen HR. Direct enumeration and functional assessment of circulating dendritic cells in patients with liver disease. Hepatology. 2004;40:335–345. [PubMed]

29. Goddard S, Youster J, Morgan E, Adams DH. Interleukin-10 secretion differentiates dendritic cells from human liver and skin. Am J Pathol. 2004;164:511–519. [PubMed]

30. Bosma BM, Metselaar HJ, Mancham S, Boor PP, Kusters JG, Kazemier G, Tilanus HW, Kuipers EJ, Kwekkeboom J. Characterization of human liver dendritic cells in liver grafts and perfusates. Liver Transpl. 2006;12:384–393. [PubMed]

31. Nattermann J, Zimmermann H, Iwan A, von Lilienfeld-Toal M, Leifeld L, Nischalke HD, Langhans B, Sauerbruch T, Spengler U. Hepatitis C virus E2 and CD81 interaction may be associated with altered trafficking of dendritic cells in chronic hepatitis C. Hepatology. 2006;44:945–954. [PubMed]

32. Ejrnaes M, Filippi CM, Martinic MM, Ling EM, Togher LM, Crotty S, von Herrath MG. Resolution of a chronic viral infection after interleukin-10 receptor blockade. J Exp Med. 2006;203:2461–2472. [PMC free article] [PubMed]

33. Louis H, Le Moine O, Peny MO, Quertinmont E, Fokan D, Goldman M, Deviere J. Production and role of interleukin-10 in concanavalin A-induced hepatitis in mice. Hepatology. 1997;25:1382–1389. [PubMed]

34. Walker MR, Carson BD, Nepom GT, Ziegler SF, Buckner JH. De novo generation of antigen-specific CD4⁺CD25⁺ regulatory T cells from human CD4⁺CD25⁻ cells. Proc Natl Acad Sci USA. 2005;102:4103–4108. [PubMed]

35. Groux H, Bigler M, de Vries JE, Roncarolo MG. Interleukin-10 induces a long-term antigen-specific anergic state in human CD4⁺ T cells. J Exp Med. 1996;184:19–29. [PMC free article] [PubMed]

36. Levings MK, Gregori S, Tresoldi E, Cazzaniga S, Bonini C, Roncarolo MG. Differentiation of Tr1 cells by immature dendritic cells requires IL-10 but not CD25⁺CD4⁺ Tr cells. Blood. 2005;105:1162–1169. [PubMed]

37. Steinbrink K, Graulich E, Kubsch S, Knop J, Enk AH. CD4⁺ and CD8⁺ anergic T cells induced by interleukin-10-treated human dendritic cells display antigen-specific suppressor activity. Blood. 2002;99:2468–2476. [PubMed]

38. MacDonald AS, Maizels RM. Alarming dendritic cells for Th2 induction. J Exp Med. 2008;205:13–17. [PMC free article] [PubMed]

39. Reis e Sousa C. Dendritic cells in a mature age. Nat Rev Immunol. 2006;6:476–483. [PubMed]

40. Van der Kleij D, Latz E, Brouwers JF, Kruize YC, Schmitz M, Kurt-Jones EA, Espevik T, de Jong EC, Kapsenberg ML, Golenbock DT, et al. A novel host-parasite lipid cross-talk: schistosomal lyso-phosphatidylserine activates Toll-like receptor 2 and affects immune polarization. J Biol Chem. 2002;277:48122–48129. [PubMed]

41. Rogers NC, Slack EC, Edwards AD, Nolte MA, Schulz O, Schweighoffer E, Williams DL, Gordon S, Tybulewicz VL, Brown GD, Reis e Sousa C. Syk-dependent cytokine induction by Dectin-1 reveals a novel pattern recognition pathway for C type lectins. Immunity. 2005;22:507–517. [PubMed]

42. Iwasaki A, Kelsall BL. Freshly isolated Peyer’s patch, but not spleen, dendritic cells produce interleukin 10 and induce the differentiation of T helper type 2 cells. J Exp Med. 1999;190:229–239. [PMC free article] [PubMed]

43. Khanna A, Morelli AE, Zhong C, Takayama T, Lu L, Thomson AW. Effects of liver-derived dendritic cell progenitors on Th1- and Th2-like cytokine responses in vitro and in vivo. J Immunol. 2000;164:1346–1354. [PubMed]

44. Re F, Strominger JL. IL-10 released by concomitant TLR2 stimulation blocks the induction of a subset of Th1 cytokines that are specifically induced by TLR4 or TLR3 in human dendritic cells. J Immunol. 2004;173:7548–7555. [PubMed]

45. Lu L, Woo J, Rao AS, Li Y, Watkins SC, Qian S, Starzl TE, Demetris AJ, Thomson AW. Propagation of dendritic cell progenitors from normal mouse liver using granulocyte/macrophage colony-stimulating factor and their maturational development in the presence of type-1 collagen. J Exp Med. 1994;179:1823–1834. [PMC free article] [PubMed]

46. Lu L, Rudert WA, Qian S, McCaslin D, Fu F, Rao AS, Trucco M, Fung JJ, Starzl TE, Thomson AW. Growth of donor-derived dendritic cells from the bone marrow of murine liver allograft recipients in response to granulocyte/macrophage colony-stimulating factor. J Exp Med. 1995;182:379–387. [PMC free article] [PubMed]

47. Morelli AE, Thomson AW. Dendritic cells: regulators of alloimmunity and opportunities for tolerance induction. Immunol Rev. 2003;196:125–146. [PubMed]

48. Solari MG, Thomson AW. Human dendritic cells and transplant outcome. Transplantation. 2008;85:1513–1522. [PubMed]