Decoy receptor 3 (DcR3), a soluble receptor belonging to the TNFR superfamily, is a receptor for both Fas ligand (FasL) and LIGHT. It has been demonstrated that DcR3 is up-regulated in lung and colon cancers, thus promoting tumor growth by neutralizing the cytotoxic effects of FasL and LIGHT. In this study, we found that DcR3.Fc profoundly modulated dendritic cell differentiation and maturation from CD14+ monocytes, including the up-regulation of CD86/B7.2, and the down-regulation of CD40, CD54/ICAM-1, CD80/B7.1, CD1a, and HLA-DR. Moreover, DcR3-treated dendritic cells suppressed CD4+ T cell proliferation in an allogeneic MLR and up-regulated IL-4 secretion of CD4+CD45RA+ T cells. This suggests that DcR3.Fc may act not only as a decoy receptor to FasL and LIGHT, but also as an effector molecule to skew T cell response to the Th2 phenotype.

Decoy receptor 3 (DcR3),3 also known as TR6 or M68, is a member of the TNFR superfamily and is a decoy receptor for Fas ligand (FasL) and LIGHT (homologous to lymphotoxins, shows inducible expression, and competes with HSV glycoprotein D for herpesvirus entry mediator, a receptor expressed by T lymphocytes) (1, 2, 3). Like osteoprotegerin (4), a member of the TNFR superfamily, DcR3 lacks a transmembrane domain and is regarded as a secreted, rather than a membrane-associated, molecule. Moreover, DcR3 can apparently neutralize the biological effects of FasL and LIGHT by inhibiting the FasL-Fas interaction (1) or by inhibiting LIGHT binding to both the lymphotoxin (LT)-β receptor (LTβR) and the herpes virus entry mediator (2, 5). DcR3 gene expression is increased in malignant tissue (1) and DcR3 protein is overexpressed in human adenocarcinomas of the esophagus, stomach, colon, and rectum (3). Moreover, DcR3 protein was overexpressed in a substantial number of tumors in which gene amplification could not be detected (3). A recent study further demonstrated that DcR3 is amplified and overexpressed in virus (EBV or human T cell leukemia virus-I)-associated lymphomas (6). These results suggest that EBV and human T cell leukemia virus-I may use DcR3 to escape from immune surveillance during lymphomagenesis, or that virus-infected lymphoma cells with DcR3 expression might be selected during multistep tumorigenesis (6). In addition, expression of DcR3 can be detected in malignant glioma cells as well as in human glioblastomas, and its expression correlates with the grade of malignancy (7). Besides tumor cell patients, the DcR3 gene is also overexpressed in silicosis or systemic lupus erythematosus patients (8). Because LIGHT is expressed in dendritic cells (DCs) and acts as a costimulatory factor essential for priming T cell responses (9, 10, 11), we questioned whether DcR3 could suppress immunity by interfering with the maturation and differentiation of DCs.

Growing evidence has demonstrated that members of the TNF superfamily transduce signals after engagement with their receptors (12, 13, 14, 15, 16, 17, 18, 19, 20, 21). In our recent study, we further demonstrated that cross-linking of TNF-related activation-induced cytokine by immobilized soluble receptor activator of NF-κB.Fc fusion protein activated p38 mitogen-activated protein kinase and enhanced IFN-γ secretion via reverse signaling through TNF-related activation-induced cytokine (22), and cross-linking of TNF-related apoptosis-inducing ligand, enhanced proliferation, and IFN-γ secretion of T cells (23).

However, even though “reverse signaling” could be triggered by immobilized receptor.Fc or agonistic mAb, there is no evidence to demonstrate that the soluble receptor.Fc fusion protein can trigger signaling and modulate cell function. In this study, we report that soluble DcR3.Fc binds to CD14+ monocytes and interferes with their differentiation and maturation into DCs. The expression of HLA-DR and other costimulatory molecules, such as CD40 and CD80/B7.1, was suppressed. In contrast, the costimulatory molecule, CD86/B7.2, was up-regulated under the same conditions. Moreover, DcR3.Fc-treated DCs biased T cell differentiation to the Th2 phenotype in allogeneic MLR. Similar results were not observed when Fas.Fc or LTβR.Fc was used in place of DcR3.Fc. Because DcR3.Fc fusion protein has been shown to have the similar binding affinity and specificity as that of DcR3 (24, 25), this raises the argument that DcR3 produced by many human tumor cells might have similar function to DcR3.Fc and could directly suppress host anti-tumor immunity by altering DCs function and skewing the immune response from Th1 to Th2.

LTβR.Fc protein was produced as previously described (10). To generate the DcR3.Fc, the open reading frame of the human DcR3 gene was isolated by RT-PCR using the forward primer: 5′-GGAATTCAAGGACCATGAGGGCGCTG-3′ and the reverse primer: 5′-GGAATTCGTGCACAGGGAGGAAGCGC-3′. The amplified product was ligated in-frame into the EcoRI-cut pUC19-IgG1-Fc vector containing the cDNA of the human IgG1 Fc. The fusion gene was then subcloned into the pBacPAK9 vector (Clontech Laboratories, Palo Alto, CA) and cotransfected with linearized BacPAK6 DNA (Clontech Laboratories) into Sf21 cells. The supernatant from recombinant virus-infected Sf21 cells was filtered and purified on protein A-Sepharose beads. The bound DcR3.Fc protein was then eluted with 0.1 M glycine buffer (pH 3.0) followed by dialysis against PBS.

PBMCs were isolated by standard density gradient centrifugation with Ficoll-Paque (Amersham Pharmacia Biotech, Piscataway, NJ) from the heparinized whole blood of normal individuals. Subsequently, CD14+ cells were purified by high-gradient magnetic sorting using the VARIOMACS technique with anti-CD14 microbeads (Miltenyi Biotec, Bergisch Gladbach, Germany). Immature DCs were generated from adherent PBMCs by culture in RPMI 1640 medium (Life Technologies, Gaithersburg, MD) supplemented with 10% FCS (Life Technologies), 800 U/ml human GM-CSF (Leucomax; Schering-Plough, Kenilworth, NJ), and 500 U/ml human IL-4 (R&D Systems, Minneapolis, MN) in the presence or absence of human IgG1 (3 μg/ml; Sigma-Aldrich, St. Louis, MO), LTβR.Fc (3 μg/ml), or DcR3.Fc (3 μg/ml) for 6 days (immature DCs). To prepare mature activated DCs, immature DCs were further incubated with gamma-irradiated (5500 rad) CD40 ligand (CD40L)-expressing L cells (DNAX Research Institute, Palo Alto, CA) at a ratio of 3:1 for 36 h. To purify naive CD4+CD45RA+ T cells, PBMCs were first isolated by Ficoll-Paque centrifugation, and CD4+ cells were then enriched by a CD4+ T cell isolation kit (Miltenyi Biotec). After depletion of non-CD4 T cells, CD4+ T cells were then positively selected by CD45RA microbeads (Miltenyi Biotec) using the VARIOMACS technique. The purity of naive CD4+CD45RA+ T cells was over 95% by flow cytometry analysis.

CD14+ monocyte-derived DCs were harvested and gamma-irradiated (3000 rads) followed by incubation with 5 × 104 allogeneic CD4+CD45RA+ naive T cells/well in U-bottom 96-well microtitration plates (Costar, Cambridge, MA) at ratios of 1:10 to 1:300. After 4 days, [3H]thymidine (Amersham Pharmacia Biotech) was added (0.5 μCi/well) and the cells were incubated for another 16 h. The cells were harvested on a cell harvester (Skatron, Lier, Norway), and the incorporated radioactivity was measured using a beta counter (model LS3801; Beckman Coulter, Brea, CA).

CD4+CD45RA+ naive T cells (2 × 105/well) were plated into U-bottom 96-well microtitration plates (Costar) and cultured with gamma-irradiated DCs (at a DC-T ratio of from 1:10 to 1:300). After 3 days, half of the culture medium was replaced by fresh RPMI 1640 medium (Life Technologies). On day 6, cells were washed with PBS and incubated with PMA (10 ng/ml) and A23187 (1 μg/ml) for 24 h. The supernatants were harvested and stored at −20°C. The concentrations of IL-4 and IFN-γ were measured by OptEIA ELISA (BD PharMingen, San Diego, CA).

Before staining, CD14+ monocyte-derived DCs were harvested and washed twice with FACS staining/washing buffer (1% FCS and 0.1% NaN3 in PBS), followed by incubation with various mAbs or anti-LIGHT polyclonal Ab (11) in staining buffer at 4°C for 20 min. For the samples that were incubated with nonfluorochrome-conjugated Abs, cells were then incubated with appropriate FITC-conjugated secondary Abs at 4°C for 20 min after being washed three times with FACS staining/washing buffer. Cells were fixed with 1% paraformaldehyde in PBS for 30 min at 4°C before the fluorescence was analyzed with a FACSort (BD Biosciences, Mountain View, CA). Alternatively, cells were stained with biotinylated DcR3.Fc or LTβR.Fc as follows: CD14+ monocytes and DCs were first incubated with 100 μl human IgG (100 μg/106 cells; Calbiochem, San Diego, CA) in FACS staining/washing buffer at 4°C for 10 min to prevent nonspecific binding, followed by the addition of 2 μg of biotinylated DcR3.Fc, LTβR.Fc, or IgG1 in 50 μl FACS staining/washing buffer. After washing with FACS staining/washing buffer three times, cells were further incubated at 4°C for 20 min with 50 μl UltraAvidin-PE (Leinco Technologies, Ballwin, MO) or streptavidin-FITC (BD PharMingen) diluted (1/200) in FACS staining/washing buffer. The sources of mAbs are as follows: anti-CD1a-FITC (clone HI149; BD PharMingen), anti-CD11c-PE (B-ly6; BD PharMingen), anti-CD54 (clone 8.4A6; Ancell, Bayport, MN), anti-CD80-PE (clone L307.4; BD PharMingen), anti-CD83-FITC (clone HB15e; BD PharMingen), anti-CD86-PE (clone 2331 FUN-1; BD PharMingen), anti-CD40-FITC (clone LOB716; Serotec, Oxford, U.K.), anti-HLA-DR-FITC (clone B-F1; Serotec), anti-CD14-PE mAb (clone UCHM1; Serotec), biotin-conjugated anti-FasL (clone NOK-1; BD PharMingen), anti-TNF-β/LT-α mAb (clone 9B9; Boehringer Mannheim, Mannheim, Germany), and anti-DC-lysosome-associated membrane glycoprotein mAb (clone 104.G4; Immunotech, Marseille, France).

COS7 cells were transfected with the pFLAG-FasL, pFLAG-LIGHT, or pFLAG-CMV2 vector by the calcium phosphate method. Three days after transfection, cells were harvested and resuspended by lysis buffer (1% Nonidet P-40, 150 mM NaCl, 50 mM Tris-HCl (pH 8), 1 mM PMSF, 2 μg/ml aprotinin, and 2 μg/ml leupeptin), followed by incubation with anti-LIGHT polyclonal Ab (11), DcR3.Fc, or LTβR.Fc. Immunoprecipitates were collected on protein A beads (Amersham Pharmacia Biotech). Samples were fractionated by SDS-PAGE and then probed with anti-FLAG M2 Ab (Sigma-Aldrich) by Western blot analysis.

HT-29 cells were seeded in flat-bottom 96-well microtitration plates (Costar) at a density of 5 × 103/well overnight at 37°C, followed by incubation with recombinant soluble LIGHT (sLIGHT) (25 ng/ml) and IFN-γ (10 U/ml; Boehringer Mannheim), in conjunction with IgG1 (1 μg/ml), TNFRI.Fc (1 μg/ml), LTβR.Fc (1 μg/ml), or DcR3.Fc (1 μg/ml) for 4 days. To quantitate FasL-induced apoptosis, cells were cultured in flat-bottom 96-well microtitration plates (Costar) at a density of 105/well in the presence of FasL (Upstate Biotechnology, Lake Placid, NY) and IgG1, Fas.Fc, DcR3.Fc, or LTβR.Fc in the concentration of 1 μg/ml for 16 h. The survival rate was determined by MTT assay as previously described (26). Survival rate was determined by OD570 of cells treated with FasL vs OD570 of cells cultured in medium only.

It has been reported that DcR3 is up-regulated in certain tumors (1, 2, 3, 6, 7) and tumor-associated DCs usually have a low T cell stimulatory capacity (reviewed by Banchereau et al.; Ref. 27). Therefore, we asked if DcR3 could modulate the functions of DCs. We first tested the binding specificity of the DcR3.Fc to FasL and LIGHT. DcR3.Fc precipitated both FasL and LIGHT (Fig. 1,A), and both DcR3.Fc and Fas.Fc inhibited activation-induced apoptosis of Jurkat cells and CD4+ T cells (Fig. 1, B and C). Furthermore, DcR3.Fc inhibited LIGHT and IFN-γ-mediated apoptosis of HT-29 cells to a similar degree as LTβR.Fc did (Fig. 1,D). To further confirm the binding specificity between DcR3.Fc and FasL, Jurkat cells were treated with rFasL in the presence of receptor.Fc fusion proteins. As shown in Fig. 1 E, DcR3.Fc has a similar effect as that of Fas.Fc to inhibit FasL-induced Jurkat cell apoptosis. These results indicate that DcR3.Fc interacted with FasL and LIGHT, and that it possessed the same binding specificity as previously described (1, 2, 24, 25).

FIGURE 1.

Specific interaction between DcR3. Fc and its ligands. A, Immunoprecipitation of FasL and LIGHT by DcR3.Fc. COS7 cells were transfected with pFLAG-FasL, pFLAG-LIGHT, or pFLAG-CMV2 by the calcium phosphate method. After 3 days, cell lysates were incubated with anti-LIGHT polyclonal Ab, DcR3.Fc or LTβR.Fc, followed by incubation with protein A-conjugated Sepharose beads. Finally, the immunoprecipitates were fractionated on SDS-PAGE and probed with anti-FLAG M2 mAb. B and C, Inhibition of activation-induced apoptosis in Jurkat cells and CD3+ T cells by DcR3.Fc. Jurkat cells (106/well) were stimulated with PMA (20 ng/ml) and ionomycin (2 μg/ml) for 18 h (B), while CD4+ T cells (5 × 105/well) were activated with immobilized anti-CD3 (20 μg/ml) for 5 days (C), in 96-well microtitration plates, followed by restimulation with OKT-3 (20 μg/ml) for 24 h in the presence of Fas.Fc (10 μg/ml), DcR3.Fc (10 μg/ml), or human IgG1 (10 μg/ml). Percentages of cell death were determined by PI staining. Statistical analysis by two-tailed Student’s t test revealed significant differences between IgG1- and DcR3.Fc-treated samples (∗, p < 0.05). The data represent the mean ± SD for three experiments. D, Inhibition of LIGHT-mediated apoptosis in HT-29 cells. HT-29 cells were incubated with sLIGHT (25 ng/ml) and IFN-γ (10 U/ml) in the presence of IgG1 (1 μg/ml) or Fc fusion proteins (1 μg/ml) for 4 days, and the survival rate was determined as described in Materials and Methods. One representative experiment of three is shown. ∗, p < 0.05 when compared with IFN-γ treated samples; ∗∗, p < 0.05 when compared with IFN-γ and sLIGHT treated samples. E, Neutralization of FasL-induced apoptosis in Jurkat cells. Jurkat cells were treated with different concentrations of FasL in the presence of IgG1 (1 μg/ml), or Fc fusion proteins (1 μg/ml) for 16 h, and the cell survival rate was determined by MTT assay.

FIGURE 1.

Specific interaction between DcR3. Fc and its ligands. A, Immunoprecipitation of FasL and LIGHT by DcR3.Fc. COS7 cells were transfected with pFLAG-FasL, pFLAG-LIGHT, or pFLAG-CMV2 by the calcium phosphate method. After 3 days, cell lysates were incubated with anti-LIGHT polyclonal Ab, DcR3.Fc or LTβR.Fc, followed by incubation with protein A-conjugated Sepharose beads. Finally, the immunoprecipitates were fractionated on SDS-PAGE and probed with anti-FLAG M2 mAb. B and C, Inhibition of activation-induced apoptosis in Jurkat cells and CD3+ T cells by DcR3.Fc. Jurkat cells (106/well) were stimulated with PMA (20 ng/ml) and ionomycin (2 μg/ml) for 18 h (B), while CD4+ T cells (5 × 105/well) were activated with immobilized anti-CD3 (20 μg/ml) for 5 days (C), in 96-well microtitration plates, followed by restimulation with OKT-3 (20 μg/ml) for 24 h in the presence of Fas.Fc (10 μg/ml), DcR3.Fc (10 μg/ml), or human IgG1 (10 μg/ml). Percentages of cell death were determined by PI staining. Statistical analysis by two-tailed Student’s t test revealed significant differences between IgG1- and DcR3.Fc-treated samples (∗, p < 0.05). The data represent the mean ± SD for three experiments. D, Inhibition of LIGHT-mediated apoptosis in HT-29 cells. HT-29 cells were incubated with sLIGHT (25 ng/ml) and IFN-γ (10 U/ml) in the presence of IgG1 (1 μg/ml) or Fc fusion proteins (1 μg/ml) for 4 days, and the survival rate was determined as described in Materials and Methods. One representative experiment of three is shown. ∗, p < 0.05 when compared with IFN-γ treated samples; ∗∗, p < 0.05 when compared with IFN-γ and sLIGHT treated samples. E, Neutralization of FasL-induced apoptosis in Jurkat cells. Jurkat cells were treated with different concentrations of FasL in the presence of IgG1 (1 μg/ml), or Fc fusion proteins (1 μg/ml) for 16 h, and the cell survival rate was determined by MTT assay.

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We then tested the functions of DcR3 in modulating DC maturation and differentiation. CD14+ monocytes were differentiated into immature DCs by culture with GM-CSF and IL-4 in conjunction with DcR3.Fc, LTβR.Fc, Fas.Fc, or human IgG1 for 6 days. The cells were then stimulated with CD40L to induce DC maturation. We found that DcR3.Fc, but not LTβR.Fc, Fas.Fc (data not shown) or human IgG1 modulated DC differentiation and maturation (Fig. 2). Before CD40L stimulation, it was apparent that the expression of CD1a, a hallmark of the DC phenotype, was suppressed by DcR3.Fc in immature DCs in a dose-dependent manner (Fig. 2,A, upper panel), while CD86/B7.2 was up-regulated in a dose-dependent manner (Fig. 2,A, lower panel). The modulatory effect of DcR3.Fc on the expression of CD1a and CD86/B7.2 reaches a plateau at 3 μg/ml (Fig. 2 A).

FIGURE 2.

Modulation of surface marker expression and cytokine secretion of DCs by DcR3.Fc. A, CD14+ monocytes were cultured with GM-CSF (800 U/ml) and IL-4 (500 U/ml) for 6 days to differentiate them into immature DCs in the presence of various concentrations of IgG1 and DcR3.Fc. Surface marker expression was analyzed by flow cytometry analysis and the mean fluorescence intensity was calculated by CellQuest software (BD Biosciences). ∗, p < 0.05 when compared with IgG1-treated samples. The data represent the mean ± SD for three experiments. B and C, Human IgG1 (3 μg/ml), DcR3.Fc (3 μg/ml), or LTβR.Fc (3 μg/ml) were added to the culture of DCs in the presence of GM-CSF (800 U/ml) and IL-4 (500 U/ml) (denoted as GMIL4). Surface markers expressed by DCs before CD40L stimulation (B) or after CD40L stimulation (C) were analyzed by flow cytometry analysis (shaded histograms, isotype control Abs; open histograms, specific Abs). Dead cells were gated out by propidium iodide staining. One representative experiment of three is shown.

FIGURE 2.

Modulation of surface marker expression and cytokine secretion of DCs by DcR3.Fc. A, CD14+ monocytes were cultured with GM-CSF (800 U/ml) and IL-4 (500 U/ml) for 6 days to differentiate them into immature DCs in the presence of various concentrations of IgG1 and DcR3.Fc. Surface marker expression was analyzed by flow cytometry analysis and the mean fluorescence intensity was calculated by CellQuest software (BD Biosciences). ∗, p < 0.05 when compared with IgG1-treated samples. The data represent the mean ± SD for three experiments. B and C, Human IgG1 (3 μg/ml), DcR3.Fc (3 μg/ml), or LTβR.Fc (3 μg/ml) were added to the culture of DCs in the presence of GM-CSF (800 U/ml) and IL-4 (500 U/ml) (denoted as GMIL4). Surface markers expressed by DCs before CD40L stimulation (B) or after CD40L stimulation (C) were analyzed by flow cytometry analysis (shaded histograms, isotype control Abs; open histograms, specific Abs). Dead cells were gated out by propidium iodide staining. One representative experiment of three is shown.

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We further tested the modulatory effect of DcR3.Fc on the expression of other activation and differentiation markers of DCs (Fig. 2, B and C and Table I). It is interesting to note that the expression of DC-lysosome-associated membrane glycoprotein (a marker of mature DCs) was up-regulated by DcR3.Fc, but not by LTβR.Fc or IgG1 before CD40L stimulation (Fig. 2,B). In contrast, the expression of HLA-DR was down-regulated by DcR3.Fc in immature DCs (mean fluorescence intensity = 55 ± 18) compared with those treated with IgG1 (mean fluorescence intensity = 132 ± 36) and LTβR.Fc (mean fluorescence intensity = 127 ± 18) (Fig. 2,B and Table I). Compared with IgG1, it is interesting to note that DcR3.Fc increased the expression of CD86/B7.2 on immature DCs up to 2-fold when compared with control proteins. By contrast, the expression of CD11c, a lineage marker of myeloid cells, was unaffected under the same condition (data not shown). When DcR3.Fc treated-immature DCs were stimulated with CD40L for 36 h, the expression of CD54/ICAM-1, CD80/B7.1 and CD86/B7.2 were all up-regulated, while the expression of CD1a was not affected by CD40L stimulation. The expression of CD40 is only slightly up-regulated before and after CD40L stimulation (mean fluorescence intensity = 17 ± 11 vs 31 ± 23). Compared with IgG1- and LTβR.Fc-treated mature DCs, the expression of CD1a, CD54/ICAM-1, HLA-DR, and CD80/B7.1 is still suppressed, but the expression of CD86/B7.2 is further up-regulated by DcR3.Fc (Fig. 2,C and Table I). Therefore, DcR3.Fc, but not LTβR.Fc, profoundly affected the differentiation and maturation of DCs.

Table I.

Mean fluorescence intensity of surface markers on receptor.Fc-treated DCsa

Mean Fluorescence IntensityCD1aCD40CD54HLA-DRCD80CD86
GMIL4 73 ± 16 31 ± 9 75 ± 16 105 ± 23 20 ± 5 76 ± 15 
GMIL4/IgG1 80 ± 24 29 ± 10 83 ± 18 132 ± 36 21 ± 3 72 ± 14 
GMIL4/DcR3.Fc 10 ± 4 17 ± 11 67 ± 15 55 ± 18 10 ± 4 162 ± 43 
GMIL4/LTβR.Fc 57 ± 18 27 ± 11 76 ± 19 127 ± 18 21 ± 3 84 ± 17 
GMIL4/CD40L 40 ± 14 47 ± 17 210 ± 62 343 ± 31 54 ± 15 113 ± 52 
GMIL4/IgG1/CD40L 38 ± 18 46 ± 14 238 ± 66 287 ± 27 43 ± 16 117 ± 48 
GMIL4/DcR3.Fc/CD40L 9 ± 3 31 ± 23 118 ± 36 75 ± 8 22 ± 9 237 ± 52 
GMIL4/LTβR.Fc/CD40L 31 ± 13 40 ± 14 219 ± 61 298 ± 39 46 ± 4 112 ± 42 
Mean Fluorescence IntensityCD1aCD40CD54HLA-DRCD80CD86
GMIL4 73 ± 16 31 ± 9 75 ± 16 105 ± 23 20 ± 5 76 ± 15 
GMIL4/IgG1 80 ± 24 29 ± 10 83 ± 18 132 ± 36 21 ± 3 72 ± 14 
GMIL4/DcR3.Fc 10 ± 4 17 ± 11 67 ± 15 55 ± 18 10 ± 4 162 ± 43 
GMIL4/LTβR.Fc 57 ± 18 27 ± 11 76 ± 19 127 ± 18 21 ± 3 84 ± 17 
GMIL4/CD40L 40 ± 14 47 ± 17 210 ± 62 343 ± 31 54 ± 15 113 ± 52 
GMIL4/IgG1/CD40L 38 ± 18 46 ± 14 238 ± 66 287 ± 27 43 ± 16 117 ± 48 
GMIL4/DcR3.Fc/CD40L 9 ± 3 31 ± 23 118 ± 36 75 ± 8 22 ± 9 237 ± 52 
GMIL4/LTβR.Fc/CD40L 31 ± 13 40 ± 14 219 ± 61 298 ± 39 46 ± 4 112 ± 42 
a

CD14+ monocytes were cultured in the presence of GM-CSF and IL-4 (GMIL4), or in conjunction with 3 μg/ml of DcR3.Fc, or LTβR.Fc. After 6 days, the developed immature DCs were further stimulated with CD40L-expressing L cells in the presence of Fc fusion proteins for 36 h. The expression of surface markers was analyzed by flow cytometry and the mean fluorescence intensity was calculated by CellQuest software (BD Biosciences). The data represents the mean ± SD from six independent experiments.

Because DcR3.Fc modulated the expression of surface molecules important for Ag presentation, we asked the effect of DcR3.Fc-treated DC to modulate T cell proliferation and differentiation. As shown in Fig. 3,A, DcR3.Fc suppressed T cell proliferation when the T cells were incubated with DcR3.Fc-treated mature DCs at DC-T ratios of 1:10 and 1:30 (Fig. 3,A, upper panel), or with DcR3.Fc-treated immature DCs at DC-T ratios of 1:10 (Fig. 3 A, lower panel).

FIGURE 3.

Modulatory effects of DcR3.Fc on T cell proliferation in MLR. A, Allogeneic CD4+ T cells were cultured with gamma-irradiated DCs in 96-well microtitration plates for 4 days at different DC-T ratios of 1:10 (□) and 1:30 (▪), and the T cell proliferation was measured as described in Materials and Methods.B and C, The concentrations of IFN-γ and IL-4 secreted by CD4+CD45RA+ naive T cells which cocultured with immature DCs (B) or with mature DCs (C) were determined by ELISA. One representative experiment of three is shown. GMIL4: GM-CSF (800 U/ml) and IL-4 (500 U/ml); ∗, p < 0.05 when compared with untreated or IgG1-treated DCs.

FIGURE 3.

Modulatory effects of DcR3.Fc on T cell proliferation in MLR. A, Allogeneic CD4+ T cells were cultured with gamma-irradiated DCs in 96-well microtitration plates for 4 days at different DC-T ratios of 1:10 (□) and 1:30 (▪), and the T cell proliferation was measured as described in Materials and Methods.B and C, The concentrations of IFN-γ and IL-4 secreted by CD4+CD45RA+ naive T cells which cocultured with immature DCs (B) or with mature DCs (C) were determined by ELISA. One representative experiment of three is shown. GMIL4: GM-CSF (800 U/ml) and IL-4 (500 U/ml); ∗, p < 0.05 when compared with untreated or IgG1-treated DCs.

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In addition to its suppressive effect on T cell proliferation, DcR3.Fc also modulated the secretion of IFN-γ and IL-4 of CD4+CD45RA+ naive T cells. We found that DcR3.Fc-treated immature DCs enhanced IL-4 secretion (1.5-fold) compared with IgG1- or LTβR.Fc-treated immature DCs (Fig. 3,B, lowerpanel). In contrast, DcR3.Fc-treated immature DCs did not significantly affect IFN-γ secretion (Fig. 3 B, upper panel). Therefore, DcR3.Fc-treated immature DCs increased the IL-4-IFN-γ ratio via the enhancement of IL-4 production in CD4+CD45RA+ naive T cells.

We then tested the effect of DcR3-treated mature DCs on IFN-γ and IL-4 secretion from CD4+CD45RA+ naive T cells. To address this aspect, immature DCs were stimulated with CD40L for 36 h to induce DC maturation, followed by incubation with allogeneic CD4+CD45RA+ naive T cells at various DC-T ratios. We found that DcR3.Fc-treated mature DCs did not affect the secretion of IFN-γ at a DC-T ratio of 1:30 (Fig. 3,C, upper panel). Consistently, DcR3.Fc-treated mature DCs enhanced IL-4 secretion at a DC-T ratio of 1:30 (Fig. 3 C, lower panel) to 2-fold compared with IgG1- or LTβR.Fc-treated mature DCs. Therefore, DcR3-treated mature DCs also increased the IL-4-IFN-γ ratio by enhancing IL-4 secretion. According to this data, it is suggested that DcR3.Fc might cause both immature and mature DCs to skew the immune response toward Th2 development.

In previous experiments, we demonstrated that DcR3.Fc, but not LTβR.Fc, could modulate the expression of several surface molecules and enhance IL-4 secretion of CD4+CD45RA+ naive T cells. This suggested that DcR3.Fc and LTβR.Fc might bind to distinct molecules to execute their respective functions. To address this aspect, we used anti-LIGHT, anti-LTα, and anti-FasL Abs, as well as DcR3.Fc and LTβR.Fc fusion proteins, to stain CD14+ monocytes and immature and mature DCs to define the molecule that interacts with DcR3.Fc. As shown in Fig. 4 A, DcR3.Fc, but not LTβR.Fc, bound freshly isolated CD14+ monocytes. In contrast, anti-LIGHT, anti-LTα, and anti-FasL Abs did not bind CD14+ monocytes. This suggested that DcR3.Fc could bind to a surface molecule distinct from LIGHT, the membrane form of LT, and FasL on CD14+ monocytes.

FIGURE 4.

Characterization of the interaction of DcR3.Fc with cell surface molecules. Monocytes (A) and CD14+ monocyte-derived immature and mature DCs (B) were stained with the following Abs: anti-LIGHT polyclonal Ab, anti-LTα mAb, and biotinylated anti-FasL mAb, followed by FITC-conjugated secondary Abs or PE-conjugated UltraAvidin. Alternatively, cells were stained with biotinylated receptor. Fc fusion proteins (LTβR.Fc and DcR3.Fc) or biotinylated IgG1, followed by incubation with either PE-conjugated UltraAvidin or FITC-conjugated streptavidin. The mean fluorescence intensity of cells was analyzed by FACSort and by CellQuest software (BD Biosciences). Shaded histograms, control staining; open histograms, specific staining. One representative experiment of three is shown.

FIGURE 4.

Characterization of the interaction of DcR3.Fc with cell surface molecules. Monocytes (A) and CD14+ monocyte-derived immature and mature DCs (B) were stained with the following Abs: anti-LIGHT polyclonal Ab, anti-LTα mAb, and biotinylated anti-FasL mAb, followed by FITC-conjugated secondary Abs or PE-conjugated UltraAvidin. Alternatively, cells were stained with biotinylated receptor. Fc fusion proteins (LTβR.Fc and DcR3.Fc) or biotinylated IgG1, followed by incubation with either PE-conjugated UltraAvidin or FITC-conjugated streptavidin. The mean fluorescence intensity of cells was analyzed by FACSort and by CellQuest software (BD Biosciences). Shaded histograms, control staining; open histograms, specific staining. One representative experiment of three is shown.

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DcR3.Fc, LTβR.Fc, and anti-LIGHT polyclonal Ab all bound to immature DCs. In contrast, only DcR3.Fc bound to mature DCs (Fig. 4 B). In a previous study, we showed that LIGHT is expressed on the surface of immature DCs, but is shed from mature DCs and is barely detectable by LTβR.Fc (9). DcR3.Fc detected strong fluorescence signals in both immature and mature DCs, while LTβR.Fc did not detect any signal in mature DCs, and only a weak signal could be detected in immature DCs. From this, we conclude that the molecule detected by DcR3.Fc on mature DCs is distinct from LIGHT, indicating that DcR3 binds to both LIGHT and an unidentified novel ligand in immature DCs. Because FasL was not expressed on freshly isolated monocytes and CD14+ monocyte-derived DCs cultured with GM-CSF and IL-4, we conclude that the specific signal detected by DcR3.Fc on monocytes and mature DCs must differ from those of LIGHT and FasL.

The information obtained in this study suggests that DcR3.Fc, when cultured with CD14+ monocyte-derived DCs in vitro, can modulate the expression of surface molecules, reduce T cell proliferation, and enhance IL-4 production of CD4+CD45RA+ naive T cells. In contrast, neither LTβR.Fc nor Fas.Fc had a similar effect, so DcR3.Fc might modulate DC function by interacting with a non-FasL non-LIGHT molecule expressed, at least, on monocytes and DCs. Because DcR3 is up-regulated in cancer patients, DcR3 might be one of the factors responsible for immunosuppression found in cancer patients.

Tumor cells can produce factors that promote blood vessel growth (neovascularization) to meet their increasing demand for oxygen and nutrients. In addition, tumor cells express many immunosuppressive factors, such as TGF-β, IL-10, DF3/MUC1 (28), and RCAS1 (29), to suppress the host immune response and facilitate tumor growth. DcR3, a decoy receptor capable of neutralizing FasL and LIGHT, has been reported to be overexpressed in tumor cells originating from the gastrointestinal tract and the pulmonary system (1, 3). Thus, it has been speculated that DcR3 might inhibit host immune responses by neutralizing the cytotoxic effects of FasL and LIGHT.

A recent study demonstrated that both DcR3.Fc and rDcR3 have the same binding affinity to LIGHT and have the same inhibitory effect on the development of CTL in mice (24) indicating that the biological effect of DcR3.Fc is equivalent to rDcR3 protein. Therefore, the modulatory effect of DcR3.Fc observed in this study should be able to reflect the function of DcR3. In this study, we demonstrated that DcR3.Fc could modulate DC differentiation and activation via a ligand distinct from FasL and LIGHT. Because FasL is not expressed on CD14+ monocyte-derived DCs cultured with GM-CSF and IL-4, and LTβR.Fc did not affect CD14+ monocyte-derived DCs in a similar fashion as DcR3.Fc, the modulatory effects of DcR3 cannot be attributed to neutralization of LIGHT or FasL. Therefore, our observations suggest that DcR3 may act as an effector molecule to modulate DC functions via its binding to surface molecules to trigger “reverse signaling” as found for other members of the TNF superfamily (12, 13, 14, 15, 16, 17, 18, 19, 20, 21, 22, 23). However, we cannot rule out the possibility that DcR3 might interact with an unidentified molecule capable of triggering signals to induce DC differentiation, thus the addition of DcR3 blocks this signaling pathway and results in the failure to follow the normal process of DC differentiation and maturation in the in vitro culture system.

One of the most striking effects of DcR3.Fc on DC differentiation was the up-regulation of CD86/B7.2 and down-regulation of CD80/B7.1. In addition to activation through TCR by the Ag-MHC complex, the costimulatory signal induced by interaction of CD28 or CTLA-4 on the T cell surface with either CD80/B7.1 or CD86/B7.2 on APC is required for T cell proliferation and cytokine secretion. It has been reported that IFN-γ could up-regulate, while IL-10 could down-regulate, both CD80/B7.1 and CD86/B7.2 expression (30, 31). Thus, the unique feature of DcR3.Fc up-regulation of CD86/B7.2 with simultaneous down-regulation of CD80/B7.1 might be an invaluable tool to dissect the underlying mechanism for the regulation of CD80/B7.1 and CD86/B7.2 expression. Furthermore, it has been demonstrated that even though CD80/B7.1 and CD86/B7.2 equivalently costimulate IL-2 and IFN-γ production, CD86/B7.2 preferentially induced more IL-4 production than CD80/B7.1 (32, 33). In this study, we clearly demonstrated that DcR3 up-regulated CD86/B7.2 and down-regulated CD80/B7.1 expression on DCs. In addition, DcR3.Fc-treated DCs dramatically enhanced IL-4 secretion by CD4+CD45RA+ naive T cells. Thus, our observation is in accordance with previous reports that CD86/B7.2 preferentially activates IL-4 expression and Th2 development (32, 33, 34). In addition to CD80/B7.1and CD86/B7.2, it has been reported that blocking or absence of LFA-1 (CD11a/CD18)/ICAM-1 (CD54) interaction promotes IL-4 secretion (35, 36, 37). In DcR3.Fc-treated mature DC, we found that the expression level of CD54/ICAM-1 is suppressed (mean fluorescence intensity = 118 ± 36), compared with IgG1- (mean fluorescence intensity = 238 ± 66) or LTβR.Fc- (mean fluorescence intensity = 219 ± 61) treated mature DCs (Table I). Therefore, another mechanism of DcR3.Fc-treated DC to induce Th2 differentiation might be through the down-regulation of CD54/ICAM-1, resulting in the enhanced secretion of Th2 cytokine. However, we cannot rule out the possibility that other molecules are also responsible for the Th2 polarization effect of DcR3.Fc-treated DCs.

Recent work suggests that different DC subsets contribute significant polarizing influences on Th differentiation. In the murine system, lymphoid-derived DCs induce high levels of the Th1 cytokines IFN-γ and IL-2 but little or no Th2 cytokines, while the myeloid-derived DCs induce large amounts of the Th2 cytokines IL-4 and IL-10, in addition to IFN-γ and IL-2 (38). In contrast, human monocyte (pDC1)-derived DCs (DC1) induce Th1 differentiation, whereas DC2 derived from CD4+CD3CD11c plasmacytoid cells (pDC2) induce Th2 differentiation (39). However, DcR3 appears to suppress CD4+ T cell proliferation and influence CD4+ T cell differentiation via modulating the expression of CD80/B7.1, CD86/B7.2, and other yet-defined molecules in DC1, thus altering their ability to polarize the immune response from Th1 to Th2. This suggests that the ability of the DC1 subset to induce Th1 cell polarization could be altered by other environmental factors.

In our preliminary study, we found that DcR3 is up-regulated in the serum of certain cancer patients (unpublished observations), therefore tumor cells might suppress immune reaction by secreting DcR3 to modulate DC activation and differentiation, in addition to neutralizing the cytotoxic effect of FasL and LIGHT. However, the nature of the novel ligand for DcR3 expressed on CD14+ monocyte and DC is still unclear. It will be very interesting to ask whether DcR3 could modulate the differentiation of other cell lineages, and to identify the novel ligand interacting with DcR3 in the future.

We thank Dr. Steve Roffler for critical comments on this work and Drs. Nien-Jung Chen, Hsian-Guey Hsieh, and Norimitsu Kadowaki for their technical assistance. We owe much to Dr. Hugh McDevitt for his kindness in facilitating this work at Stanford University.

1

This work was mainly supported by Grant NHRI-CN-BP-8902S from the National Health Research Institute, Taiwan; National Sciences Council Grants NSC90-2320-B-010-109 and NSC90-2318-B-010-009-M51; and Grant GMYM 8902 from the Chi-Mei Foundational Hospital (Tainan, Taiwan).

3

Abbreviations used in this paper: DcR3, decoy receptor 3; FasL, Fas ligand; LT, lymphotoxin; LIGHT, homologous to LTs, shows inducible expression, and competes with HSV glycoprotein D for herpesvirus entry mediator, a receptor expressed by T lymphocytes; LTβR, LTβ receptor; DC, dendritic cell; sLIGHT, soluble LIGHT; CD40L, CD40 ligand.

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