Decoy Receptor 3 Suppresses TLR2-Mediated B Cell Activation by Targeting NF-κB

Zi-Ming Huang, Jhi-Kai Kang, Chih-Yu Chen, Tz-Hau Tseng, Chien-Wen Chang, Yung-Chi Chang, Shyh-Kuan Tai, Shie-Liang Hsieh and Chuen-Miin Leu
J Immunol June 15, 2012, 188 (12) 5867-5876; DOI: https://doi.org/10.4049/jimmunol.1102516

Abstract

Decoy receptor 3 (DcR3) is a soluble protein in the TNFR superfamily. Its known ligands include Fas ligand, homologous to lymphotoxin, showing inducible expression, and competing with HSV glycoprotein D for herpes virus entry mediator, a receptor expressed by T lymphocytes, TNF-like molecule 1A, and heparan sulfate proteoglycans. DcR3 has been reported to modulate the functions of T cells, dendritic cells, and macrophages; however, its role in regulating B cell activation is largely unknown. In this study, we found that the DcR3.Fc fusion protein bound to human and mouse B cells and suppressed the activation of B cells. DcR3.Fc attenuated Staphylococcus aureus, IgM-, Pam3CSK4-, and LPS-mediated B cell proliferation but did not affect cytokine-induced B cell growth. In the presence of these mitogens, DcR3.Fc did not induce B cell apoptosis, suggesting that DcR3 may inhibit the signal(s) important for B cell activation. Because the combination of Fas.Fc, LT-βR.Fc (homologous to lymphotoxin, showing inducible expression, and competing with HSV glycoprotein D for herpes virus entry mediator, a receptor expressed by T lymphocytes receptor), and DR3.Fc (TNF-like molecule 1A receptor) did not suppress B cell proliferation and because the biological effect of DcR3.Fc on B cells was not blocked by heparin, we hypothesize that a novel ligand(s) of DcR3 mediates its inhibitory activity on B cells. Moreover, we found that TLR2-stimulated NF-κB p65 activation and NF-κB–driven luciferase activity were attenuated by DcR3.Fc. The TLR2-induced cytokine production by B cells was consistently reduced by DcR3. These results imply that DcR3 may regulate B cell activation by suppressing the activation of NF-κB.

Decoy receptor 3 (DcR3, also known as TR6 or M68) is a soluble receptor that belongs to the TNFR superfamily. The gene encoding DcR3 produces a 300-aa polypeptide, and it is found in humans but not in mice. DcR3 is barely detectable in normal tissues; however, it is overexpressed in a variety of tumor cells such as adenocarcinomas of the esophagus, stomach, colon, rectum, and pancreas (1–4) and in lymphomas and gliomas (5, 6). Earlier reports showed that DcR3 binds to Fas ligand (FasL), homologous to lymphotoxin, showing inducible expression, and competing with HSV glycoprotein D for herpes virus entry mediator, a receptor expressed by T lymphocytes (LIGHT), and TNF-like molecule 1A (TL1A) (1, 7, 8). DcR3 prevents FasL- and LIGHT-induced tumor cell apoptosis by neutralizing the interactions of FasL and LIGHT with Fas and lymphotoxin β-receptor (LT-βR), respectively (1, 6, 7). DcR3 also promotes angiogenesis to help tumor growth by binding to TL1A and subsequently stimulating the proliferation of endothelial cells (9).

Accumulating evidence has demonstrated a potent immunomodulatory function of DcR3. DcR3 can neutralize T cell-mediated apoptosis by binding FasL and LIGHT (1, 7). Furthermore, DcR3 may regulate T cell differentiation. The T cell response is biased to a Th2 phenotype with an increased IL-4/IFN-γ ratio in a DcR3-transgenic mouse model (10). Another study reported that the intrathecal injection of the DcR3.Fc protein into mice with experimental autoimmune encephalomyelitis significantly reduced the number of Th17 cells and the production of IL-17 (11). These observations suggest that DcR3 may favor the differentiation of Th2 cells and suppress Th1 and Th17 cells, although the mechanism by which DcR3 controls T cell differentiation remains to be elucidated. In addition to its neutralization activity, DcR3 has been demonstrated to have direct modulatory activities (also called reverse signaling) on various types of immune cells. Through its binding to LIGHT, DcR3 induces signals and inhibits T cell chemotaxis (12). DcR3 has been reported to transduce reverse signaling to modulate the differentiation and maturation of dendritic cells (DCs) and macrophages; intriguingly, these effects are independent of FasL, LIGHT, and TL1A (13–15). The DcR3 ligands in myeloid lineage that mediate the regulation of DcR3 have been identified recently as heparan sulfate proteoglycans (HSPGs), such as CD44v3 and sydecan-2 (16). Although DcR3 is reported to modulate the functions of T cells, DCs, and macrophages, whether it regulates B cell remains to be elucidated.

B cell activation is the first step of the humoral immune response. After encountering T-independent or T-dependent Ags, B cells undergo activation, proliferation, and differentiation to become memory or plasma cells. Several microbial products, including peptidoglycans, LPS, and unmethylated CpG oligonucleotides stimulate polyclonal B cell proliferation and may drive limited plasma cell differentiation without the help of T cells (17–19). In the past decades, extensive studies have revealed the receptors responsible for the response to these microbial B cell mitogens, namely the TLRs.

TLRs are pattern recognition receptors that play a critical role in the regulation of innate and adaptive responses. There are 10 TLRs in humans and 12 TLRs in mice (20). Peptidoglycan and the synthetic agonist Pam3CSK4 are recognized by TLR1/2, LPS binds to TLR4, and CpG binds to TLR9. TLR ligands stimulate B cell activation, the increased expression of activation markers (such as CD25, CD69, CD80, CD86 and MHC class II), and cell proliferation. The production of cytokines, including IL-12p40, IL-12p70, IL-6, IL-10, and TNF-α, is simultaneously induced (21). Upon TLR activation, all TLRs except TLR3 recruit the adaptor molecule MyD88 to the plasma membrane and evoke a series of reactions that lead to the activation to NF-κB and AP-1, two transcription factors that regulate gene expression. TLR signaling also activates MAPK pathways and the PI3K pathway (22–25). Genetic approaches have shown the importance of the NF-κB and PI3K pathways in TLR4-mediated B cell proliferation (26–28); however, the biochemical mechanisms by which other TLRs regulate B cell activation remain to be elucidated.

In the current study, we found an unexpected effect of DcR3.Fc on B cell activation. DcR3.Fc binds to human and mouse B cells, and it is able to suppress the activation and proliferation of B cells. We found that resting B cells do not express FasL, LIGHT, or TL1A. Furthermore, the biological function of DcR3.Fc is not blocked by heparin. The administration of Fas.Fc, LT-βR.Fc (LIGHT receptor), and DR3.Fc (TL1A receptor) does not suppress B cell proliferation, suggesting that its effects are mediated via unknown ligand(s) on the B cells. We discovered that DcR3.Fc attenuates the TLR2 agonist-induced cytokine expression. Finally, we found that the TLR2-stimulated NF-κB activation was suppressed by DcR3. Taken together, our results reveal a regulatory function of DcR3 on B cells.

Materials and Methods

Cells, Abs, and chemicals

Human and mouse cells were cultured in RPMI 1640 medium containing 100 U/ml penicillin, 100 mg/ml streptomycin, 2 mM l-glutamine, and 10% FCS (Life Technologies, Grand Island, NY). The Ramos, Daudi, and IIA1.6 B cell lines were from Dr. Max D. Cooper (Emory University, Atlanta, GA). The Raji B cell line was from Dr. Cheng-po Hu (Tunghai University, Taichung, Taiwan). The THP-1 cell line was provided by Dr. Shie-Liang Hsieh. The BCL-1 cell line was generously provided by Dr. Kuo-I Lin (Academia Sinica, Taipei, Taiwan). The human tonsils and blood were obtained in accordance with the policies established by the Taipei Veterans General Hospital Institutional Review Board. The mononuclear cells in the tonsils or the buffy coat were isolated by Ficoll-Hypaque gradient centrifugation. The human CD19+ B cells and mouse B220+ splenic B cells were purified using Ab-conjugated microbeads (Miltenyi Biotec, Auburn, CA). The cell purity was determined by staining with either anti-human CD20 or anti-mouse B220 Abs. The PE-conjugated anti-human CD19, CD20, CD69, CD38, IgM, and IgD Abs were purchased from BD Pharmingen (San Diego, CA). The Alexa 488-conjugated anti-mouse B220 and PE-conjugated anti-mouse CD69 Abs were from BioLegend (San Diego, CA). Anti-mouse CD16/32 Ab (clone 2.4G2) was from BD Pharmingen. The plasma-derived human IgG was purchased from Calbiochem (La Jolla, CA). All the animal experiments were conducted in accordance with the guidelines approved by the Animal Care and Usage Committee of National Yang-Ming University. Both the C57B/L6 and BALB/c mouse strains were used in this study, and similar results were obtained. Ly294002 and PD98059 were purchased from Calbiochem. Bay 11-7082, Pam3CSK4, and CpG ODN1826 were obtained from InvivoGen (San Diego, CA). LPS and human IgG1 (hIgG1) were purchased from Sigma-Aldrich (St. Louis, MO). Anti-human CD40 Ab and recombinant human IL-10 were purchased from R&D Systems (Minneapolis, MN). Anti-human IgM F(ab′)2 fragment was from Jackson ImmunoResearch Laboratories (West Grove, PA). Recombinant human IL-2 was from Roche Applied Science (Mannheim, Germany).

The production of recombinant fusion proteins

The recombinant fusion proteins DcR3.Fc, DcR3.Fcmut, Fas.Fc, LT-βR.Fc, and DR3.Fc were expressed by the FreeStyle 293 expression system as described previously (10, 14). Briefly, the expression vector containing the fusion protein was transfected into 293F cells, and the supernatant was collected on days 2, 4, and 6. The purification of the fusion proteins was performed on a Sepharose-protein A column (Amersham Biosciences), and the purity of fusion proteins was determined by silver staining. The bioactivity of DcR3.Fc was checked by THP-1 cell adhesion assay (14) before it was applied to B cells.

The DcR3.Fc binding assay

Before staining, peripheral blood cells were incubated with 1 mg/ml human IgG to block Fc receptors, and mouse B220 cells were preblocked with 100 μg/ml anti-mouse CD16/32 mAb (clone 2.4G2). To assess DcR3.Fc binding, the cells were incubated with biotinylated DcR3.Fc followed by streptavidin-allophycocyanin with the PE-labeled lineage-specific markers and then analyzed by flow cytometry. To test if DcR3.Fc binds to HSPGs, 106 cells were incubated with 10 μg/ml biotinylated DcR3.Fc in the absence or presence of various concentrations of heparin or heparan sulfate on ice for 10 min. After washing with PBS, the cells were incubated with streptavidin-PE for an additional 10 min. The cells were washed twice with PBS and analyzed by flow cytometry.

The analysis of B cell proliferation and activation

The purified human B cells were incubated in 96-well plates (105/well in 200 μl RPMI 1640 supplemented with 10% FCS) for 72 h in the presence or absence of the fixed Staphylococcus aureus Cowan (SAC) or Wood (SAW) strains (Sigma-Aldrich). SAC produces ∼10-fold more protein A than does SAW. The cells were pulsed for an additional 24 h with [3H]thymidine (1 μCi/well; PerkinElmer), harvested, and the [3H]thymidine incorporation was assessed with a liquid scintillation counter. For the mouse splenic B cells, the purified B220+ B cells (1 × 105) were plated and incubated with Pam3CSK4, LPS, or CpG in the presence or absence of hIgG1 (10 μg/ml) or DcR3.Fc (10 μg/ml). After 48 h, [3H]thymidine was added (1 μCi/well), and the cells were incubated for another 24 h. These cells were harvested with a cell harvester, and the incorporated radioactivity was measured using a β counter (PerkinElmer Wallac 1450 Microbeta Trilux Scintillation Counter; PerkinElmer). The stimulation percentage of test samples was calculated as follows: % stimulation = 100% × (cpm of the test sample − cpm of medium control)/(cpm of the hIgG1 control − cpm of medium control). For the cell division assay, the purified B cells were resuspended in PBS/5% FCS, and CFSE (Invitrogen, Carlsbad, CA) was added to a final concentration of 5 μM and incubated for 10 min at room temperature. The cells were then washed once with PBS/5% FCS. The cells were counted and stimulated as described. The data from 10,000 total events were acquired and analyzed using FACSCanto (BD Biosciences, Mountain View, CA). The cell-surface expression of CD69 was assessed 18–24 h after the cells were treated as described above. The percentage of inhibition is calculated as follows: 100% × (percentage of the hIgG1 control − percentage of the test sample)/(percentage of the hIgG1 control − percentage of medium control).

The apoptosis assay

To quantify the apoptotic cells, the cells were incubated with FITC-Annexin V and propidium iodide (PI) according to the manufacturer’s instructions (BD Pharmingen). The apoptotic cells were monitored by flow cytometry (FACSCanto; BD Biosciences).

RT-PCR and quantitative RT-PCR

PCR primers for detection of human FasL, LIGHT, and TL1A are human FasL: sense, 5′-CAGCTCTTCCACCTACAGAAGG-3′ and antisense, 5′-CTCTTAGAGCTTATATAAGCCG-3′; human LIGHT: sense, 5′-AGATCTTGACGGACCTGCAGGCTCC-3′ and antisense, 5′-CTTCACACCATGAAAGCCC-3′; human TL1A: sense, 5′-GGAATTCCATGGCCGAGGATCTGG-3′ and antisense, 5′-GTCTTCCGACTCTGGGATCAG-3′; mouse FasL: sense, 5′-ACCACTACCACCGCCATCACAA-3′ and antisense, 5′-CCAGAGATCAGAGCGGTTCCATA-3′; mouse LIGHT: sense, 5′-ATGGAGAGTGTGGTACAGCCTTC-3′ and antisense, 5′-GACCATGAAAGCTCCGAAATAGG-3′; and mouse TL1A: sense, 5′-AGTCCCAGTGGAAGTGCTG-3′ and antisense, 5′-GTGCTAAGTCCTGCGAGGAT-3′. To perform quantitative RT-PCR, the B220+ cells cultured for 8 h with Pam3CSK4 in the presence or absence of 10 μg/ml hIgG1 or DcR3.Fc were harvested and lysed by TRIzol reagent (Invitrogen Life Technologies). The RNA was purified, and the cDNA was generated by the SuperScript III First-Strand Synthesis System (Invitrogen) according to the manufacturer’s instructions. The PCR was performed on the LightCycler thermal cycler system (Roche Diagnositics, Mannhein, Germany) using the LightCycler FirstStart DNA Master SYBR Green I system (Roche). The following primers were used in quantitative RT-PCR: 18S rRNA (5′-ACAATACAGGACTCTTTCGAG-3′ and 5′-GAGCTTTTTAACTGCAGCAAC-3′); IL-6 (5′-GCCTTCCCTACTTCACAAGT-3′ and 5′-GAATTGCCATTGCACAACTCT-3′); TNF-α (5′-CCTCACACTCAGATCATCTTC-3′ and 5′-CGGCTGGCACCACTAGTTG-3′); IL-12p40 (5′-ACATGGTGAAGACGGCCAG-3′ and 5′-GAAGTCTCTCTAGTAGCCAG-3′); IL-10 (5′-ATGCAGGACTTTAAGGGTTAC-3′ and 5′-CCTGAGGGTCTTCAGCTTC-3′); and IκB (5′-AACCTGCAGCAGACTCCAC-3′ and 5′-GCTACACTGGCCAGGCAG-3′).

The DNA-binding assay for NF-κB

The purified splenic B cells were treated with 0.3 μg/ml Pam3CSK4 with 10 μg/ml hIgG1 or DcR3.Fc and incubated at 37°C for 60 min. The cells were then harvested, and the nuclear extracts were collected by the Nuclear Extract kit (Active Motif, Carlsbad, CA) according to the manufacturer’s instructions. The DNA binding activity of the NF-κB subunit p65 was assessed by the TransAM NF-κB kit (Active Motif). Briefly, 4 μg nuclear extract was incubated in wells coated with oligonucleotides with the NF-κB consensus sequence for 1 h. After washing, an anti-p65–specific Ab was added to each well and incubated for 1 h. The secondary HRP-conjugated Ab was then added and incubated for another hour. After the colorimetric reaction with the substrate, the absorbance was measured at 450 nm.

The preparation of the lentiviral vector and the luciferase reporter assay

To generate a lentiviral reporter vector, a DNA fragment containing an NF-κB response element that drives the luciferase reporter gene luc2P was amplified by PCR (forward primer, 5′-CATAGGATCCTACCTGAGCTCGCTAG-3′; reverse primer, 5′-GAAACTGCAGTTAGACGTTGATCCT-3′) using pGL4.32[luc2P/NF-κB-RE/Hygro] as a template. The PCR product was cleaved and cloned into a BamHI- and PstI-digested lentiviral vector, pLKOAS3.puro (generously provided by Dr. Lih-Hwa Hwang, Institute of Microbiology and Immunology, National Yang-Ming University, Taipei, Taiwan). This lentiviral reporter vector was named pLKOAS3-NF-κB-RE-luc2P.puro. To prepare the lentiviruses, the pLKOAS3-NF-κB-RE-luc2P.puro, pCMV-Δ R8.91, and pMD.G vectors (gifts from Dr. Lih-Hwa Hwang) were cotransfected into 293T cells. Fresh medium was added after 6 h, and the supernatant containing the viral particles was collected 40 and 64 h posttransfection. The viral supernatant was filtered through a 0.22-μm filter and kept at −70°C until use. The splenic B220+ cells were purified and stimulated with 10 μg/ml LPS for 24 h before transduction. The virions were added to the stimulated B cells in the presence of 10 μg/ml LPS and 8 μg/ml polybrene and centrifuged for 50 min at 450 × g. The viral supernatant was removed after an overnight incubation, and fresh medium containing 10 μg/ml LPS was added for another 24 h. The cells were moved to a medium without LPS for 2 h and then plated and stimulated under various conditions for 6 h. The cells were lysed, and the firefly luciferase activity was measured with the reagents from the Luciferase Assay System (Promega) according to the manufacturer’s instructions.

The detection of the active forms of Erk and Akt

The purified splenic B cells were cultured for 30 min with Pam3CSK4 in the presence or absence of hIgG1 (10 μg/ml) or DcR3.Fc (10 μg/ml), harvested, and lysed with RIPA buffer containing protease inhibitors and phosphatase inhibitors. The phosphorylation of Erk and Akt were detected with either the phospho-ERK1/2 Enzyme Immunometric Assay Kit (Assay Designs, Ann Arbor, MI) or the phospho-Akt (Ser473) STAR ELISA Kit (Upstate, Millipore) according to the manufacturer’s instructions. Briefly, the cell lysate was added to microtiter plates precoated with monoclonal anti-ERK or anti-Akt Abs. After washing, the active forms of ERK or Akt were detected by the incubation with specific anti–phospho-ERK or anti–phospho-Akt Abs conjugated with HRP. The concentrations of phospho-ERK or phospho-Akt were evaluated by OD proportional to standards.

Statistical analysis

The two-tailed Student t test was used for the statistical analysis of the results. The differences were considered to be significant when p < 0.05. *p < 0.05, **p < 0.01, ***p < 0.005.

Results

DcR3.Fc binds to human and mouse B cells

In addition to DCs and macrophages, the DcR3.Fc fusion protein also bound to the B cells in the peripheral blood (Fig. 1A) and to mature human B cell lines (Raji and Daudi) (Fig. 1B). To test if DcR3.Fc could bind to all the differentiation stages of B cells, we purified the CD19+ B cells from the tonsils and performed the binding assay. Biotinylated hIgG1 was used as negative control. Based on the expression of IgD and CD38, tonsillar B cells can be divided into five populations: naive cells, pregerminal center cells, germinal center cells, plasma cells, and memory cells (29). We found that all tonsillar B cells were able to bind DcR3.Fc to various degrees (Fig. 1C). The binding of DcR3.Fc to naive cells was strong and the highest in the pregerminal center cells. Furthermore, DcR3.Fc also bound to the mouse spleen B cells (B220+) and two mouse B cell lines (BCL-1 and IIA1.6) (Fig. 1B). Although the DcR3.Fc binding to splenic B cells was weak, it was consistently observed in all of the experiments (n = 6). The binding of DcR3.Fc was specific because DcR3.Fc did not bind to the peripheral blood T cells (Fig. 1A). Based on these results, we speculated that DcR3 might regulate the function of human and mouse B cells.

FIGURE 1.
FIGURE 1.

DcR3.Fc binds to B cells in humans and mice. (A) DcR3.Fc binds to blood B cells. PBMCs were incubated with 1 mg/ml human IgGs to block Fc receptors. The cells were then incubated with biotin-labeled DcR3.Fc or hIgG1 followed by streptavidin-conjugated allophycocyanin. The Abs to the lineage-specific markers (CD3, CD20, and CD14) were added to identify each cell type. Filled peak, hIgG1-biotin; thin line, DcR3.Fc-biotin. One representative result of four independent experiments is shown. (B) DcR3.Fc binds to B cell lines. Raji and Daudi are human B cell lines; THP-1 is a human monocytic cell line; B220+ indicates the purified mouse splenic B220+ B cells. BCL-1 is a surface IgM+ murine B cell line. IIA1.6 is a surface IgG+ mouse B cell line. Filled peak, hIgG1-biotin; thin line, DcR3.Fc-biotin. One representative result of at least three independent experiments is shown. (C) DcR3.Fc binds to all stages of B cells. CD19+ tonsillar B cells were purified by magnetic beads and used to perform the DcR3.Fc binding assay as described in (A), except that anti-IgD and CD38 Abs were used to dissect the five stages of B cells. One representative result of five independent experiments is shown. GC, Germinal center cells.

DcR3.Fc suppresses human B cell proliferation and activation

To test if DcR3 could regulate the function of B cells, we first examined its effects on B cell activation and proliferation. CD19+ B cells were purified from the buffy coat of human peripheral blood and stimulated with fixed S. aureus in the presence or absence of DcR3.Fc ex vivo. The purified human IgG1 was used as the negative control. S. aureus can stimulate B cell division and activation in a T cell-independent manner. Two strains of S. aureus were used in this study: SAW and SAC. We found that DcR3.Fc inhibited SAW- or SAC-induced proliferation as measured by the [3H]thymidine incorporation assay (Fig. 2A and data not shown) in human B cells. DcR3.Fc suppressed B cell proliferation in 12 out of 13 donors in >10 independent experiments with an average inhibition of 50.7 ± 11.9% (Fig. 2B, right panel). The effect of DcR3 was not limited to mitogen-stimulated proliferation. When we used anti-CD40 plus anti-IgM Abs to mimic T-dependent stimulation, DcR3.Fc was also able to reduce B cell proliferation (Fig. 2B; average inhibition 37.3 ± 14.1%; n = 4). The inhibitory effect of DcR3 on SAW- or BCR-induced proliferation was specific because DcR3.Fc did not affect IL-2– plus IL-10–mediated human B cell proliferation (Fig. 2B; average inhibition −2.7 ± 19.7%; n = 3). The DcR3.Fcmut fusion protein cannot bind FcγR and showed a similar inhibitory effect on human B cell proliferation, indicating that this suppression was attributed to the DcR3 portion (Fig. 2B, left panel).

FIGURE 2.
FIGURE 2.

DcR3.Fc suppresses the proliferation and activation of human B cells. (A) DcR3.Fc attenuates S. aureus (Wood strain, marked as SAW)-induced cell proliferation. The CD19+ B cells were purified to ∼99% purity from the peripheral blood using magnetic beads. The cells (2 × 105) were seeded in 96-well plates and cultured with the indicated reagents for 3 d. After adding 1 μCi [3H]thymidine to each well, the cells were cultured for additional 24 h and harvested to detect the incorporated isotope. The results are presented as the means ± SD of triplicate samples. The experiments were performed >10 times, and a set of representative data is presented. (B) DcR3.Fc inhibits anti-IgM and anti-CD40 Ab-stimulated cell proliferation. The B cells were stimulated either with anti-IgM (5 μg/ml) and anti-CD40 (1 μg/ml) or with anti-CD40 (1 μg/ml) plus IL-2 (100 U/ml) and IL-10 (25 ng/ml) in the presence of 10 μg/ml hIgG1, DcR3.Fc, or DcR3.Fcmut. Left panel, One representative of three independent experiments is shown. Right panel is a summary of all proliferation results in human B cells. Percentage of stimulation was calculated as described in Materials and Methods. (C) DcR3.Fc suppresses SAC-stimulated cell activation. The peripheral B cells were purified and stimulated as described in (A), and the cells were harvested 24 h later to stain CD69 and analyzed by flow cytometry. Dashed line, isotype control; filled peak and thin line, anti-CD69-PE. A set of representative data of four independent experiments is shown. (D) DcR3.Fc decreases the SAC- and BCR-induced B cell division. Filled peak, labeled cells without stimulation; thin line, cells after in vitro culture for 4 d. One representative of three to four independent experiments is shown. *p < 0.05, ***p < 0.005.

Similarly, the SAC-induced B cell activation was suppressed by DcR3.Fc as evaluated by the upregulation of activation marker CD69 (Fig. 2C). The B cells from four out of six donors responded to DcR3.Fc, and the percentage of CD69+ cells was reduced by 27.5% (hIgG1, 29.1 ± 10.5% versus DcR3.Fc, 21.1 ± 10.3%; p < 0.005). To confirm the antiproliferation activity of DcR3, we used the CFSE assay to address the same question. Consistent with the [3H]thymidine uptake results, we observed that DcR3.Fc reduced the SAC- or BCR-induced cell division (Fig. 2D). DcR3.Fc decreased cell division stimulated by SAC by 38% (hIgG1, 22 ± 8.6% versus DcR3.Fc, 13.6 ± 9%; p < 0.005, n = 5) and by the anti-IgM plus anti-CD40 Abs by 20% (hIgG1, 32.4 ± 18.1% versus DcR3.Fc, 25.8 ± 16.3%; p < 0.05, n = 3). In contrast, DcR3.Fc slightly increased cell division induced by IL-2 plus IL-10 by 8% (hIgG1, 40.8 ± 10.9% versus DcR3.Fc, 44.4 ± 12.7%; n = 4). Together, these results demonstrate that DcR3 specifically suppresses the SAW- and BCR-induced cell proliferation and activation in human B cells.

DcR3.Fc suppresses the TLR2- and TLR4-mediated mouse B cell activation and proliferation

In addition to human B cells, DcR3.Fc bound to mouse splenic B cells and B cell lines (Fig. 1B). Therefore, we tested the effect of DcR3.Fc on mouse B cell proliferation and activation. The B220+ spleen B cells from C57BL/6 mice were purified and incubated with various stimuli. In accordance with its effect on human B cells, DcR3.Fc suppressed the SAW- or SAC-induced proliferation of mouse spleen B cells (data not shown). A similar inhibitory effect of DcR3 on the splenic B cells from BALB/c mice was also observed (data not shown). To study how DcR3 reduces B cell proliferation and activation, we examined its activity on TLR-mediated stimulation because the signaling pathways transduced by TLRs are relatively defined. TLR2, TLR4, and TLR9 agonists were used to stimulate mouse B cells because all of them have been reported to induce B cell proliferation and activation (17, 19). We found that DcR3.Fc reduced the cell proliferation mediated by Pam3CSK4 and LPS but not by CpG (Fig. 3A). At a concentration of 10 μg/ml, DcR3.Fc suppressed the TLR2-induced B cell proliferation in 9 out of 10 mice with an average inhibition of 38 ± 6% (Fig. 3A, bottom right panel). Similarly, DcR3.Fc reduced the LPS-stimulated [3H]thymidine uptake in seven out of eight mice by an average of 49.2 ± 13.1%. The antiproliferative activity of DcR3.Fc on the TLR2 and TLR4 ligands was further evaluated by the CFSE assay, and similar inhibitory results were observed (Fig. 3C). The percentage of Pam3CSK4-stimulated cell division was reduced from 36 ± 9.4% to 23.2 ± 5.7% (36% decrease; n = 3, p < 0.05) by DcR3.Fc. Similarly, LPS-induced cell division was also suppressed by DcR3.Fc (59.1 ± 8.6% to 46.2 ± 9.4%; n = 3, p < 0.005) Consistent with the cell proliferation, the CD69 upregulation by Pam3CSK4 and LPS was also substantially suppressed by DcR3 (Fig. 3B). DcR3.Fc treatment led to a 30.9% decrease of the Pam3CSK4- stimulated CD69+ cells (n = 5; p < 0.005) and a 37% decrease of the LPS-stimulated cells (n = 3; p < 0.005). Together, our observations indicate that DcR3.Fc selectively attenuates the TLR2- and TLR4-mediated cell proliferation and activation in mouse B cells.

FIGURE 3.
FIGURE 3.

DcR3.Fc suppresses the TLR-mediated proliferation and activation of mouse B cells. (A) DcR3.Fc suppresses the Pam3CSK4- and LPS-induced B cell proliferation. Spleen B220+ cells (105) were seeded in 96-well plates and treated with the indicated reagents (concentration unit is μg/ml). A representative dataset of three to seven independent experiments is shown. The bottom right panel is a summary of proliferation results in mouse B cells. Pam3CSK4 (n = 9), LPS (n = 7), and CpG (n = 3). (B) DcR3.Fc suppresses the Pam3CSK4- and LPS-induced B cell activation. CD69 expression in the cells treated with 0.1 μg/ml Pam3CSK4 or 0.1 μg/ml LPS in the presence of 10 μg/ml hIgG1 or 10 μg/ml DcR3.Fc was measured 24 h after incubation. A representative dataset of three to five independent experiments is shown. (C) DcR3.Fc attenuates the Pam3CSK4- or LPS-induced cell division. Filled peak, labeled cells without stimulation; thin line, cells after in vitro culture for 3 d. One representative experiment of three is shown. **p < 0.01, ***p < 0.005.

DcR3.Fc does not affect B cell apoptosis in vitro

Recently, DcR3 has been reported to induce the apoptosis of human DCs (30). This raises the possibility that the suppression of proliferation by DcR3.Fc may be due to the induction of B cell death. To test this, we treated human CD19+ B cells with or without DcR3.Fc in the presence of SAC and measured the percentage of apoptotic cells. We found that the SAC treatment slightly reduced the Annexin V-positive cells compared with the medium control (Fig. 4A; the dot plots show one representative result, and the bar graph is a summary of three donors). In the presence of SAC, DcR3.Fc marginally augmented the percentage of apoptotic cells (by an average of ∼5% at 24, 48, or 72 h) when compared with hIgG1. To further examine the direct effect of DcR3 on human B cell apoptosis, we cultured CD19+ B cells with DcR3.Fc or hIgG1 and measured cell apoptosis. Compared to the control, DcR3.Fc did not increase human B cell apoptosis at 24, 48, or 72 h (Fig. 4C, left panel), although 10 μg/ml DcR3.Fc was reported to dramatically induce DC apoptosis (30). Because DcR3.Fc only showed a marginal effect on human B cell apoptosis in the presence of SAC, and because it did not induce apoptosis alone, we conclude that DcR3.Fc does not affect B cell apoptosis in humans.

FIGURE 4.
FIGURE 4.

DcR3.Fc does not induce B cell apoptosis. The human peripheral blood CD19+ B cells (A) and mouse splenic B220+ B cells (B) were purified and incubated with the indicated reagents in culture for various time periods. The cells were harvested at indicated time point and stained with Annexin V and PI and then analyzed by flow cytometry. A total of 0.3 μg/ml Pam3CSK4, 10 μg/ml hIgG1, and 10 μg/ml DcR3.Fc was used. Dot plots are representatives of three (A) or five (B) independent experiments. Bar graphs represent the mean Annexin V+ percentages ± SD of data. n = 3 (A); n = 7 (B). (C) DcR3.Fc does not induce B cell apoptosis by itself. Human blood B cells (left panel, n = 3) and mouse splenic B220+ B cells (right panel, n = 4) were purified and incubated with 1, 3, and 10 μg/ml DcR3.Fc, and the cells were stained with Annexin V and PI at indicated time points. The data are shown as the mean percentages ± SD of all the independent experiments.

We have also examined the effect of DcR3.Fc on mouse B cell apoptosis. As shown in Fig. 4B, >60% of the purified splenic B cells died when they were cultured in vitro for 24 or 48 h, and Pam3CSK4 treatment significantly reduced the number of apoptotic cells (medium, 70.5 ± 13.5% versus Pam3CSK4, 49.2 ± 14% at 24 h and 83.5 ± 8.1% versus 40.1 ± 16.4% at 48 h; n = 7). In the presence of Pam3CSK4, DcR3.Fc did not increase mouse B cell apoptosis at 24 h. At 48 h, the DcR3.Fc treatment slightly increased the number of Annexin V+ cells (Fig. 4B, bar graph; hIgG1, 42.2 ± 17.3% versus DcR3.Fc, 48.2 ± 19.4%; n = 7). To further assess the role of DcR3.Fc on mouse B cell apoptosis, we incubated splenic B cells with hIgG1 or DcR3.Fc and measured cell apoptosis. We did not observe an increase of apoptotic cells by the DcR3.Fc treatment at 6, 18, 24, and 48 h (Fig. 4C, right panel and data not shown; n = 2–4). Taken together, these data show that DcR3.Fc seems to have little effect on the apoptosis of mouse B cells.

The suppression of B cell proliferation by DcR3 is not correlated with FasL, LIGHT, TL1A, and HSPGs

Next, we examined what receptor mediates the suppressive effect of DcR3 in B cells. DcR3 has been reported to bind FasL, LIGHT, TL1A, and HSPGs (such as CD44v3 and sydecan-2). Using RT-PCR, we were unable to detect the expression of FasL, LIGHT, or TL1A in the human peripheral blood CD19+ cells, the naive tonsillar B cells, and the mouse splenic B220+ B cells (Fig. 5A and data not shown).

FIGURE 5.
FIGURE 5.

The antiproliferation activity of DcR3 is not related to any known ligands of DcR3. (A) RT-PCR of FasL, LIGHT, and TL1A in human B cells. −, H2O control; +, positive control; Ramos, cDNA from Ramos B cell line; PBMC-B, cDNA from purified human blood B cells; β2m, β2-microglobulin. One representative of six experiments is shown. (B) Heparin does not block the binding of DcR3.Fc to human B cells. The THP-1 or PBMC cells were incubated with 10 μg/ml biotin-labeled DcR3.Fc in the absence or presence of various concentrations of heparin (1–100 μg/ml). After washing, streptavidin-PE was added before flow cytometry analysis. Additional anti-CD20–APC Ab was used to mark the B cells in PBMCs, and the analysis was gated on the CD20+ cells. The DcR3.Fc-bound cells were gated to determine the percentage of positive-staining cells. A representative experiment of five is shown. (C and D) The inhibitory effect of DcR3.Fc is not mediated by any known ligands. The proliferation of peripheral human B cells was performed as described in Fig. 2, except that 100 μg/ml heparin or 10 μg/ml Fas.Fc, LT-βR.Fc, or DR3.Fc was used in the assay. One representative dataset of three independent experiments is shown.

To test if DcR3 can bind to B cells via HSPGs, we used heparin, heparan sulfate, and other polysaccharides to determine if they could prevent the DcR3 binding to human and mouse B cells. Our results show that none of the tested carbohydrates could block the DcR3.Fc binding to B cells (Fig. 5B and data not shown), including heparin and heparan sulfate, two HSPGs reported to dramatically inhibit the binding and function of DcR3.Fc in THP-1 cells (16). We found that heparin significantly reduced the DcR3.Fc binding to THP-1 cells in a dose-dependent manner, and the inhibition reached ∼100% at the concentration of 100 μg/ml (Fig. 5B). However, heparin could not block the DcR3.Fc binding to the human blood CD19+ B cells (Fig. 5B). Similarly, the binding of DcR3.Fc to the mouse splenic B220+ B cells was not changed by the addition of heparin, heparan sulfate, chondroitin sulfate, or dermatin sulfate (data not shown). Consistent with these results, heparin did not neutralize the inhibitory activity of DcR3.Fc in human B cell (Fig. 5C).

Although we could not detect the expression of FasL, LIGHT, or TL1A in the resting B cells from humans and mice, these DcR3 ligands might be upregulated upon B cell activation and subsequently mediate the suppressive effect. To further test the involvement of FasL, LIGHT, and TL1A in the suppression of B cell proliferation, we used the recombinant fusion proteins Fas.Fc, LT-βR.Fc, and DR3.Fc, which are receptors for FasL, LIGHT, or TL1A, respectively, to test if they have the same suppressive effects. Our results showed that neither the addition of each one of these receptor fusion proteins nor the combination of all three receptors could reproduce the antiproliferative activity of DcR3.Fc (Fig. 5D and data not shown). Based on these observations, we assume that the attenuation of DcR3 on B cells is mainly mediated by unknown ligand(s) and not by any of these identified ligands.

DcR3.Fc suppresses the TLR2-induced NF-κB activation and cytokine expression

To elucidate the signaling pathway affected by DcR3, we firstdissected the signaling pathway critical for the TLR2- and TLR4-mediated B cell proliferation. In addition to the activation of NF-κB and IFN regulatory factors, TLRs also activate ERK1/2 and PI3K/Akt pathways (22–25). Because the major function of IFN regulatory factors is to regulate the expression of type I IFNs, we focused on the role of the NF-κB, ERK, or PI3K pathways in the TLR-mediated B cell proliferation. As shown in Fig. 6A, the cell proliferation induced either by Pam3CSK4 or LPS was dramatically suppressed by NF-κB or PI3K inhibitors (BAY11-7082 and LY294002, respectively) in a dose-dependent manner. At a concentration of 0.5 μM, the NF-κB inhibitor blocked the TLR2- or TLR4-stimulated cell growth almost completely. The PI3K inhibitor Ly294002 was less potent than the NF-κB inhibitor; however, it also completely suppressed cell proliferation at 10 μM. By contrast, the MEK inhibitor PD98059 had very little effect on the TLR2 or TLR4-mediated B lymphocyte proliferation. At a concentration of 20 μM, PD98059 did not suppress the Pam3CSK4- or LPS-induced cell growth (data not shown). These observations indicate that the NF-κB and PI3K pathways play major roles in the TLR2- and TLR4-mediated B cell proliferation. We hypothesized that these two pathways may be the targets for DcR3.

FIGURE 6.
FIGURE 6.

DcR3.Fc reduces NF-κB activity in the mouse B220+ B cells. (A) The NF-κB and PI3K inhibitors block the TLR2- or TLR4-induced B cell proliferation. The purified splenic B220+ cells were incubated with Pam3CSK4 or LPS in the absence or presence of various concentrations of the inhibitor of PI3-K (LY294002) and NF-κB (BAY11-7082) for 72 h in the [3H]thymidine uptake assay. One representative of three experiments is shown. (B) DcR3.Fc does not affect the activation of ERK or Akt. The B220+ cells were stimulated with 0.25–1 μg/ml Pam3CSK4 in the presence or absence of DcR3.Fc, and the cell lysate was harvested after 30 min to detect the active forms of ERK or Akt (pERK or pAkt). The levels of pERK and pAkt were measured by specific ELISA assays and calculated according to standard controls. One representative of two experiments is shown. (C) DcR3 reduces the DNA-binding activity of the NF-κB p65 subunit. The purified mouse B220+ B cells were either left untreated (0), or treated with 0.3 μg/ml Pam3CSK4 in the presence of 30 μg/ml hIgG1, 10 μg/ml DcR3.Fc, or 30 μg/ml DcR3.Fc for 60 min. The nuclear extract was prepared and used to measure the DNA-binding activity of p65 by an ELISA assay. The result is shown as the OD readout of the samples with the blank subtracted. One representative result from three independent experiments is shown. (D) DcR3.Fc reduces the TLR2-induced NF-κB reporter activity. The purified mouse B220+ B cells were stimulated with 10 μg/ml LPS for 24 h and transduced with lentiviruses containing the NF-κB reporter. After resting in fresh medium for 2 h, the cells were plated and treated with the indicated reagents for 6 h. The cell lysates were harvested and used to measure luciferase activity. One representative of five independent experiments is shown. (E) DcR3.Fc suppresses the Pam3CSK4-induced cytokine expression. B220+ cells (106) were seeded and treated as follows: lane 1, medium only; lane 2, 0.3 μg/ml Pam3CSK4; lane 3, Pam3CSK4 plus 10 μg/ml hIgG1; and lane 4, Pam3CSK4 plus 10 μg/ml DcR3.Fc. Eight hours later, the RNA from the treated cells was extracted for the measurement of cytokine mRNA levels by real-time RT-PCR. The data were normalized with the amount of 18S rRNA in each sample. One representative of three experiments is shown. *p < 0.05, **p < 0.01, ***p < 0.005.

To test our hypothesis, we measured the activation of Akt and NF-κB upon TLR2 stimulation with or without DcR3.Fc. An ELISA-based analysis was used to detect the active forms of Akt (pAkt), ERKs (pERKs), and NF-κB after the B220+ B cells were stimulated in vitro by Pam3CSK4. The TLR2 agonist induced a 2.5-fold increase of pERKs and a 3–5-fold increase of pAkt (Fig. 6B); however, DcR3.Fc did not attenuate the activation of ERKs or Akt. In contrast, the DNA-binding activity of NF-κB p65 subunit was substantially reduced by DcR3.Fc upon Pam3CSK4 treatment in a dose-dependent manner (Fig. 6C). In three independent experiments, DcR3.Fc suppressed the NF-κB DNA-binding activity by 28.3 ± 5.5% or 54.3 ± 16.8% at a concentration of 10 or 30 μg/ml, respectively. To ascertain that DcR3 suppressed NF-κB activation, an NF-κB–driven luciferase reporter system was used. The purified splenic B220+ cells were transduced with a lentiviral NF-κB reporter and stimulated with Pam3CSK4 in the presence of hIgG1 or DcR3.Fc. As shown in Fig. 6D, Pam3CSK4 increased NF-κB promoter activity in the presence of hIgG1 by ∼5-fold; however, DcR3.Fc suppressed the induction of NF-κB activity by 2.8-fold (10 μg/ml) or 1.9-fold (30 μg/ml). The average inhibition of the NF-κB reporter activity by 10 μg/ml DcR3.Fc was 58.5 ± 9.7% (n = 3). A similar inhibitory effect of DcR3 was also observed in the LPS-stimulated reporter assay (data not shown). These results indicate that DcR3 suppressed the TLR2- and TLR4-induced activation of NF-κB. To further support this idea, we examined the expression of proinflammatory cytokines, such as IL-6, TNF-α, and the p40 subunit of IL-12, following TLR2 stimulation. We found that DcR3.Fc reduced the TLR2-induced cytokine expressions by 50% (Fig. 6E, compare lane 4 to lane 3). Two other NK-κB target genes, IL-10 and IκB, were also suppressed by DcR3.Fc. The levels of TLR2 and PTEN mRNA remained unchanged by the treatment of DcR3.Fc (data not shown). Taken together, our results demonstrate that DcR3.Fc partially suppresses the activation of the NF-κB pathway and subsequently reduces B cell activation and proliferation.

Discussion

DcR3 has been reported to regulate the differentiation and functions of T cells, macrophages, and DCs; however, its effect on B cells remains largely unknown. In this study, we demonstrated that DcR3.Fc suppressed B cell activation and proliferation, and this effect was not due to the induction of cell death. Furthermore, the NF-κB activation and cytokine expression stimulated by TLR agonists were substantially reduced by DcR3. The inhibition of B cells by DcR3 might be mediated by a novel ligand(s) of DcR3. In summary, our results indicate a very complex role for DcR3 in the regulation of the immune system. In addition to neutralizing or inducing reverse signaling in T cells and myeloid cells, DcR3 may also modulate B cell activation. When this article was under review, Cheng et al. (31) reported that DcR3.Fc suppresses Pam3CKS4-induced mouse B cell proliferation in vitro and collagen-specific IgG2a production in a collagen-induced arthritis model. Altogether, their and our findings support the idea that DcR3 may regulate B cell proliferation.

DcR3 was found to prevent the FasL- or LIGHT-mediated apoptosis of tumor cells (1, 6, 7) and the activation-induced cell death of T cells (10). On the contrary, a high concentration of DcR3.Fc (10 μg/ml) activates DC apoptosis: it induced a 3.2-fold and 2.4-fold increase of apoptotic DCs at 36 and 72 h, respectively (30). However, we did not observe any significant anti- or proapoptotic activity of DcR3.Fc on the B cells. This phenomenon is well demonstrated in human B cells (Fig. 4A). In the case of mouse B cells, we consistently observed a marked decrease in the number of apoptotic mouse B cells upon the addition of Pam3CSK4 (Fig. 4B). This phenomenon seems to be in contrast with the dogma that activation induces apoptosis in lymphocytes. Under our experimental conditions, mouse B cells undergo spontaneous apoptosis without stimulation. The addition of the TLR2 agonist induces massive cell division, thereby increasing the total cell number, whereas the nondividing cells keep dying. When the speed of cell division exceeds that of cell death, an increase in the number of live cells after stimulation should be detected. In the literature, activation-induced cell death is triggered by BCR or CD40 signaling alone (32, 33). Although CpG does not regulate mouse splenic B cell death, LPS is able to suppress cell apoptosis (33, 34). TLR2 has been reported to mediate apoptosis in the human THP-1 monocytic cell line and in mouse microglia cells (35, 36), but its role in B cell apoptosis remains largely unknown. Our observations suggest that the activation of the TLR2 signaling pathway may not induce B cell apoptosis. Whether Pam3CSK4 inhibits apoptosis requires further investigation. In the presence of Pam3CSK4, the DcR3.Fc treatment did not affect mouse B cell apoptosis at 24 and 48 h. We also did not observe an induction of B cell apoptosis by the addition of DcR3.Fc alone. Compared to its significant effect on DC apoptosis, we conclude that DcR3.Fc may not regulate spontaneous cell death of primary B cells, although we cannot rule out the possibilities that DcR3 may block Fas- or activation-induced B cell apoptosis.

We have shown four lines of evidence indicating the existence of a novel DcR3 receptor(s) on B cells. First, we were unable to detect the expression of any known ligands of DcR3 in the resting B cells. Secondly, heparin, heparan sulfate, and all of the carbohydrates tested failed to block the binding of DcR3.Fc to the B cells. Third, heparin failed to reverse the inhibitory effect of DcR3 on the human B cells. Fourth, Fas.Fc, LT-βR.Fc, and DR3.Fc did not show a similar suppression effect as DcR3 did. The last result was especially critical because the upexpression of FasL and LIGHT in B cells under various conditions has been reported before (37–39). We have detected low levels of LIGHT mRNA in the CD38+ pregerminal and germinal center cells from two tonsil samples (C.-Y. Chen and C.-M. Leu, unpublished observations), which raises the possibility that the activated B cells may express known ligands of DcR3. Consistent with this observation, we found that the DcR3.Fc binding to the pregerminal center cells subsequently increased (Fig. 1C). Whether the enhanced binding of DcR3 can be attributed to LIGHT, FasL, or TL1A requires further investigation. However, Fas.Fc, DR3.Fc, and LT-βR.Fc were unable to inhibit B cell proliferation (Fig. 5D). Collectively, our data indicate the existence of a novel DcR3 ligand on B cells, and we favor the hypothesis that DcR3 binds to a new receptor(s) and suppresses B cell activation. We do not yet know the identity of this novel receptor.

The data presented in this study reveal differences between the myeloid cells and B cells in response to DcR3. Most obviously, apoptosis and MHC class II expression (Y.-C. Chang, S.-L. Hsieh, and C.-M. Leu, unpublished observations) in B cells were not regulated by exogenous DcR3. This may be due to the differential ligand expression in these two lineages. Monocytes and DCs express HSPGs (including CD44v3 and Syndecan-2) and TL1A (40–42), and HSPGs are important in the induction of apoptosis of DCs (30) and reduction of MHC class II expression in monocytes. TL1A can be secreted or expressed on the surface of monocytes and DCs. Because the membrane-bound form of TL1A contains a cytoplasmic region, DcR3 binding to the surface TL1A may transduce a signal. The influence of DcR3 on monocytes and DCs may be caused by the combination of all of the signals evoked by HSPGs and TL1A. However, there is no TL1A, CD44v3, or Sydecan-2 expression in B cells, and our results demonstrated that the antiproliferative effect of DcR3 is independent of HSPGs. This suggests that there are distinct regulatory roles for DcR3 on the different APCs. The different subsets of DcR3 ligands expressed on the APCs might help fine-tune the functions of the APCs.

In this study, we demonstrated that DcR3 suppressed NF-κB activation in B cells.

In addition, our unpublished observations (M.-H. Chen, Z.-M. Huang, C.-M. Leu, and H.-Y. Lin) indicate that a high level of DcR3 in the synovial fluid from rheumatoid arthritis (RA) patients is associated with low RA disease activity (Disease Activity Score 28). Together with the previous findings that DcR3 promotes Th2- but suppresses Th1- and Th17-type responses (10, 11), these data lead us to propose that the transient upregulation of DcR3 may help attenuate inflammation. The DcR3 level in healthy donors is extremely low. The elevated expression of DcR3 was reported in the tissues from patients with Crohn’s diseases (43) and in the serum and/or synovial fluid of patients with systemic lupus erythematosus (SLE) and RA (44–46 and M.-H. Chen, Z.-M. Huang, C.-M. Leu, and H.-Y. Lin, unpublished observations). The induction of DcR3 in autoimmune patients seems to be stimulated by TNF-α (or by other inflammatory cytokines) via the NF-κB pathway (47). Therefore, DcR3 might be a negative-feedback regulator of inflammation. Although the level of DcR3 is increased in the serum of SLE patients, its local concentration may not be high enough to attenuate inflammation. This might be the reason why DcR3 fails to suppress active inflammation despite its increased circulation levels. In contrast, sustained elevated levels of DcR3 may protect activated lymphocytes from Fas-mediated apoptosis via binding FasL, leading to the accumulation of self-reactive T and B cells and to the pathogenesis of autoimmune diseases. Our findings in this study support the notion that DcR3 may impair B cell proliferation and inflammation; however, the long-term effect of continuous increase of DcR3 in the pathogenesis of autoimmune diseases remains to be investigated. Administration of DcR3.Fc fusion protein into autoimmune animal models with arthritis or SLE-like syndromes may help clarify the role of DcR3 in the regulation of inflammation.

In this study, we used peripheral blood B cells and found an inhibitory effect of DcR3 on cell activation. Peripheral blood B cells consist of naive B and memory B cells. We are not sure if both populations of B cells respond to DcR3.Fc because we did not separate these two populations to perform the experiments. Because most of the mouse splenic B cells are at the naive stage, we assume that at least naive B cells are influenced by DcR3. In addition to naive B cells, DcR3.Fc binds to all other stages of B cells, including germinal center cells, memory cells, and plasma cells. It is possible that DcR3 might modulate B cell differentiation or Ab secretion. We have tested the effect of DcR3.Fc on the LPS-stimulated Ab secretion by mouse B cells in vitro and found that LPS-induced IgM production per cell was blocked (J.-K. Kang and C.-M. Leu, unpublished observations). We found that the B cell activation in vitro was suppressed and speculated similar effects in vivo. As expected, we found a general reduction of specific IgG secretion in the DcR3 transgenic mice immunized with T-dependent or -independent Ags (W.-J. Hsu and C.-M. Leu, unpublished observations). In accordance with our observations, the injection of DcR3 plasmid into collagen-induced arthritis mice decreases the serum level of collagen-specific IgG2a (31). Therefore, DcR3 is able to suppress B cells both in vitro and in vivo. Our data support the role of DcR3 as a suppressor of the TLR- or BCR-mediated B cell activation/proliferation.

Disclosures

The authors have no financial conflicts of interest.

Acknowledgments

We thank Drs. Kuo-I Lin, Chi-Ju Chen, and Lih-Hwa Hwang for helpful discussions. We also thank Dr. Ming-Zong Lai for careful review of the manuscript, Drs. Lih-Hwa Hwang and Nien-Jung Chen for providing reagents, and Dr. Yueh-Hsuan Chan, Po-Chun Liu, and Chin-Wen Wei for technical assistance.

Footnotes

  • This work was supported by National Science Council Grants NSC96-2320-B-010-011-MY3 and NSC99-2320-B-010-005-MY3 and Taipei Veterans General Hospital Grants V95S5-008, V96S5-005, V97S5-001, and V98S5-005.

  • Abbreviations used in this article:

    DC
    dendritic cell
    DcR3
    decoy receptor 3
    FasL
    Fas ligand
    hIgG1
    human IgG1
    HSPG
    heparan sulfate proteoglycan
    LIGHT
    homologous to lymphotoxin, showing inducible expression, and competing with HSV glycoprotein D for herpes virus entry mediator, a receptor expressed by T lymphocytes
    LT-βR
    lymphotoxin β receptor
    PI
    propidium iodide
    RA
    rheumatoid arthritis
    SAC
    Staphylococcus aureus Cowan strain
    SAW
    Staphylococcus aureus Wood strain
    SLE
    systemic lupus erythematosus
    TL1A
    TNF-like molecule 1A.

  • Received August 31, 2011.
  • Accepted April 9, 2012.

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