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Departments of
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Immunology,
Molecular Biotechnology,
Microbiology, and
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Pathology, University of Washington, Seattle, WA 98195; and
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Molecular Immunology Center, University of Texas Southwestern Medical Center, Dallas, TX 75235
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Abstract |
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 Top Abstract IntroductionMaterials and MethodsResultsDiscussionReferences |
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B ligand (RANKL)/TNF-related activation-induced cytokine
(TRANCE), a primarily T-cell restricted TNF family member that augments
dendritic cell (DC) function. In this report, we demonstrate that
OPG/FDCR-1 is membrane bound on the surface of transfected baby hamster
kidney (BHK) and untransfected FDC-1 cells. We also found a restricted
OPG/FDCR-1 expression pattern in lymphoid cells, specifically in B
cells, DCs and FDC-enriched fractions, which in B cells and DCs is
up-regulated by CD40 stimulation. Because OPG/FDCR-1 shares some
properties with RANK, the first RANKL/TRANCE receptor, we discuss how
the balance between RANK and OPG/FDCR-1 expression could influence
immune responses and, ultimately, germinal center
formation. |
Introduction |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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The molecular signals that direct B cells to live, to proliferate, or
to die are not well understood. The TNFR family members (reviewed in
10 , which include CD95 (Fas) and CD40, have been recently
implicated as critical receptors regulating B cell fate. For example,
CD40, CD154 (CD40 ligand (CD40L)), TNFRI, or lymphotoxin-
-deficient
mice have defects at some stage of GC formation (11, 12, 13, 14). Despite the
overt phenotype of these mice, almost nothing is known about the
regulation of TNF/TNFR family members on FDCs or stromal cells and how
they function in GC creation. Also, members of the TNF/TNFR families
often can affect the expression of other family members. For example,
ligating CD40 on B cells or DCs can up-regulate the expression of
RANK (15), CD95 (Fas) (16, 17) and lymphotoxin- (18).
One major obstacle in trying to characterize molecules involved in GC formation has been the difficulty in isolating FDC. Most of our knowledge about FDCs comes from observations made in situ (19, 20). However, several groups have had some success in purifying FDCs or isolating FDC lines (21, 22, 23, 24, 25, 26, 27, 28). FDCs can associate with B cells via Ag-Ig receptors, and through adhesion molecules such as CD54/CD11a and VCAM/VLA-4. FDC-mediated costimulation can maintain memory B cells and influence the fate of GC B cells. For example, isolated centroblasts or centrocytes undergo apoptosis in culture unless rescued by coculture with FDCs (27, 28, 29, 30).
Previously, our laboratory isolated and characterized a cell line, FDC-1, that has morphologic and functional similarities to FDCs in vivo (22, 23). FDC-1 cells are able to bind B cells, but not T cells, in a CD54-dependent fashion. FDC-1 cells, through soluble and membrane-bound signals, are able to promote B cells to proliferate and secrete IgM and IgG. To define novel molecules that play a role in GC formation, we prepared a cDNA library from FDC-1 cells and sequenced FDC-associated cDNAs. One of these clones encodes a TNFR family member which we termed FDCR-1, for follicular dendritic cell-derived receptor-1. This cDNA is identical to osteoprotegerin (OPG) and osteoclastogenesis inhibitory factor (OCIF), recently reported by Simonet et al. (31) and Yasuda et al. (32), respectively. Currently, this molecule is thought to influence bone metabolism and possibly vascular development (33). OPG/FDCR-1 is a second receptor for RANKL/TNF-related activation-induced cytokine (TRANCE) (34, 35), a T cell restricted molecule that augments DC survival and function (15, 36, 37). Here, we demonstrate that OPG/FDCR-1 is expressed as both a membrane-bound form and secreted form. OPG/FDCR-1 has several properties in common with RANK, the first characterized receptor for RANKL/TRANCE. OPG/FDCR-1 has a restricted expression pattern in cells of the immune system, including DCs, EBV-transformed B cell lines, and tonsillar B cells. Furthermore, it is regulated by CD40 stimulation, suggesting that it could play a role in DC and B cell function.
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Materials and Methods |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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FDC-1 cells were used to prepare an FDC cDNA library using the UNIZAP XR vector (Stratagene, La Jolla, CA). Preparation of the cDNA library was performed according to the manufacturer’s recommendation. The cDNA library was screened by the partial sequencing of a set of 500 cDNA clones through the use of automated fluorescent sequencing, following the recommendation of the manufacturer (Applied Biosystems, Foster City, CA). The sequenced clones were analyzed for relatedness to previously identified genes using public domain programs available through the National Human Genome Project, National Institutes of Health, which can be accessed on the World Wide Web. These programs included Blast-n, Blast-x and Fasta that used BLAST and FASTA sequence comparison algorithms.
Surface biotinylation
Tissue culture dishes (100 mm2) of near confluent cells (OPG/FDCR-1 transfected OPG/FDCR-1-baby hamster kidney (BHK) cells or untransfected FDC-1 cells) were washed twice with bicarbonate/saline buffer (20 mM bicarbonate (pH 8.0)/150 mM NaCl). The dishes were incubated with 4% of biotinylation reagent (Amersham, Arlington Heights, IL) in bicarbonate/saline buffer at 4°C for 45 min with constant rocking. The cells were washed twice with PBS, then lysed with 1 ml of 1% Nonidet P-40 lysis buffer (250 mM NaCl, 25 mM Tris-HCl (pH 7.5), 5 mM EDTA (pH 8.0), 1% Nonidet P-40, 2 µg/ml aprotinin, and 0.02% NaN3) at 4°C for 15 min. Lysates were centrifuged at 14,000 rpm at 4°C for 10 min. The supernatants were precleared with a pre-immune rabbit serum at a dilution of 1 µl/ml of lysate and 50 µl of packed protein A-Sepharose at 4°C for 4 h with constant rocking. The supernatants were then incubated with anti-FDCR-1 serum at a dilution of 1:1000, or the preimmune serum at a dilution of 1:1000, at 4°C for 30 min with constant rocking. A total of 50 µl of packed protein A-Sepharose was added, and then precipitated at 4°C overnight with constant rocking. The beads were washed three times with lysis buffer and then boiled 5 min. The proteins were separated on a 10% polyacrylamide gel, under reducing or nonreducing conditions, and then transferred to nitrocellulose. The blots were blocked with 1x TBST-B (50 mM Tris-HCl (pH 8.0), 150 mM NaCl, 0.1% Tween-20, and 5% BSA), at 4°C overnight with constant rocking, and then washed five times with TBST (50 mM Tris-HCl (pH 8.0), 150 mM NaCl, and 0.1% Tween-20). Blots were incubated with 0.5 µg/ml of streptavidin/horseradish peroxidase for 1 h at 4°C and washed as before. Biotinylated proteins were visualized by enhanced chemiluminescence (ECL) (Amersham).
Construction of expression vectors and FACS analysis of transfected BHK cells
The full-length OPG/FDCR-1 was cloned into the mammalian
expression vector PCI-neo (Promega, Madison, WI). A portion of FDCR-1
was amplified by PCR using 5'-CCACCCAGGATCCGTTTCCTCCAAAGTA-3' as the
upstream primer and a downstream primer specific for the T3 promoter.
The resulting product was digested with BamHI and
NotI, and then ligated into a modified pSecTag A vector
(Invitrogen, San Diego, CA) where a region encoding a 5'
c-myc epitope had been inserted downstream of the murine
Ig- chain signal peptide. The resulting construct encoded the Ig-
signal peptide, the c-myc epitope and human FDCR-1, residues
24–401. To construct soluble extracellular domain of FDCR-1, amino
acids 24–202 were amplified with a 5' primer
(5'-CCACCCAGGATCCGTTTCCTCCAAAGTA-3') and a 3' primer
(5'-AAGAATGCCTCGAGACACAGGGTAACATCTATTCCAC-3') so that the resulting
product had a BamHI and XhoI site at its 5' and
3' ends, respectively. This product was digested with BamHI
and XhoI and cloned in the BamHI-XhoI
sites of the modified pSecTag A vector. This produced an open reading
frame encoding the murine Ig- chain signal peptide, amino acids
24–202 of FDCR-1, c-myc epitope and 6x His epitope.
The myc-DR3 expression construct was described previously
(38). Briefly, residues 27–418 were cloned in a similar fashion as the
c-myc-OPG/FDCR-1 construct such that the protein had the
Ig- chain signal peptide, c-myc epitope and residues
27–418 of DR3.
Cells were grown in 100 mm2 culture dishes to semiconfluency. The cells were washed with PBS, incubated with 0.5 mM EDTA in PBS, and then removed by cell scraping. The harvested cells were centrifuged, the pellet was resuspended in FACS media (2% FCS in RPMI 1640 and 0.1% sodium azide), then divided into 10–20 wells of a 96-well plate for flow cytometry experiments. Cells were incubated with anti-myc (9E20), anti-FDCR-1 sera, at a dilution of 1:1000, or preimmune anti-sera, at a dilution of 1:1000, at 4°C for 40 min. The cells were washed, then incubated with FITC conjugated, F(ab')2 fragments of goat anti-rabbit Ab (BioSource International, Camarillo, CA) at a 1:80 dilution at 4°C for 40 min. The live, unfixed cells were analyzed by flow cytometry. The mean peak fluorescence (MPF) was calculated.
Pulse chase of transfected OPG/FDCR-1-BHK cells and untransfected FDC-1 cells
Adherent cell lines were grown to near confluency. The cells were washed with PBS and then incubated with medium lacking cysteine and methionine for 2 h. The cells were pulsed with media containing 0.2 mCi/ml of [35S]Met,Cys for 1 h and then chased with 5 ml of complete medium. At the indicated times, the medium was removed and the cells lysed with 0.5% Nonidet P-40 lysis buffer. Lysates or cell supernatants were precleared with preimmune sera at a concentration of 1:1000 and 50 µl of packed protein A Sepharose for 4 h at 4°C. Samples were precipitated with anti OPG/FDCR-1 sera (1:1000 dilution) and 50 µl of packed protein A Sepharose overnight at 4°C. Samples were washed with Nonidet P-40 lysis buffer and then separated on a 10% acrylamide gel under nonreducing conditions. Gels were analyzed by autoradiography.
Isolation of primary cells
As described by Grouard et al. (27), to isolate enriched FDC populations, we minced human tonsils in FDC medium (RPMI 1640 with 10% FCS, 25 mM EDTA (pH 7.4), 0.1 mg/ml DNase I, and 2 mg/ml collagenase IV). Tissue fragments were incubated for 12 min at 37°C. The cell suspension was collected after two successive digestions and then separated over Ficoll-Hypaque. The interface cells were removed and washed then rosetted to remove T cells. The Er- cells were washed in FDC medium and then placed on a 15/35% Percoll gradient and spun 800 x g for 20 min. Cells from the interface were recovered and washed in PBS. Cells were resuspended in RPMI 1640 with 10% FCS, 1 mM EDTA, and 1% human serum, and then incubated with phycoerythrin-conjugated anti-CD21 (BioSource International) and FITC-conjugated anti-CD14 (Becton Dickinson, San Jose, CA). Samples were washed and then sorted using flow cytometry. CD21brightCD14bright cells were collected.
As described in Clark et al. (22), tonsillar B cells were isolated by disaggregation of human tonsils with 10% FCS in RPMI and then centrifuged through Ficoll-Hypaque. The mononuclear cells were harvested and then rosetted. The Er- cells were fractionated by a discontinuous percoll gradient. B cells from the following interfaces were collected: <45%, 45/50%, 50/55%, 55/60%, 60/65%, and 65/70%.
DCs were differentiated in vitro from CD14+ peripheral blood monocytes using established methods. Briefly, human blood was separated over Ficoll-Hypaque and the mononuclear cells were collected. PBMCs were washed twice in PBS, 0.5% BSA, and 2 mM EDTA. Cells (20 µl/10 x 106) were sorted by MACS with CD14 conjugated microbeads (Miltenyi Biotec, Sunnyvale, CA). CD14+ PBMCs were washed and resuspended at 2 x 106 cells/ml in RPMI 1640, 10% FCS, 100 ng/ml granulocyte-macrophage CSF, and 15 ng/ml IL-4. Fresh medium with cytokines was added after 3 days. After 7 days of culture, DCs were collected.
Cell stimulations
Tonsillar B cells were suspended 10 x 107 cells/ml or 1 x 107 cells/ml (for cells collected from 60/65% and 65/70% Percoll interfaces) with 1 µg/ml each of goat anti-human IgM F(ab')2 (Jackson ImmunoResearch, West Grove, PA) and anti-CD40 (G28-5) for 18 h at 37°C. DCs were cultured with either anti-CD40 (G28-5) or IgG1 isotype control (MOPC-21) at 10 µg/ml for 72 h at 37°C.
RNA isolation
RNA samples from cell lines, isolated PBMCs, tonsillar B cell Percoll fractions, or DCs were prepared using TRIzol reagent (Life Technologies, Grand Island, NY) following the manufacturer’s instructions.
Northern blot analysis of human cell lines, isolated cells, and human tissue
Total cellular RNA (5 µg) was resolved by electrophoresis in an 0.8% formaldehyde agarose gel in 1x MOPS buffer and then transferred to a nylon membrane (Nytran, Schleicher & Schuell, Keene, NH) by capillary blotting. RNA was cross-linked by UV irradiation. 32P-labeled probe was generated with random hexamer primers. Hybridization was performed at 42°C in hybridization buffer (50% formamide, 1x Denhardt’s solution, 5x SSC, and 250 µg/ml denatured salmon sperm DNA). Blots were washed four times with 2x SSC and 0.2% SDS for 5 min at room temperature and then washed at high stringency, twice in 0.1x SSC and 0.2% (w/v) SDS for 30 min at 50°C. Membranes were exposed to autoradiographic film at -70°C.
For the multiple tissue Northern blot of lymphoid tissues (see Fig. 5
D) (Clontech, Palo Alto, CA), 32P-labeled probe
was generated using the PCR radioactive labeling system (Life
Technologies) and following the manufacturer’s instructions.
Hybridization and washing were performed following the manufacturer’s
instruction.
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RNA from isolated CD21brightCD14bright FDC was reverse transcribed and amplified using the Smart cDNA synthesis kit (Clontech) following the manufacturer’s instructions. The amplified cDNA was used in the following RT-PCR experiments.
RT-PCR analysis was performed using the titan one-tube RT-PCR system
(Boehringer Mannheim, Indianapolis, IN) following the manufacturer’s
instructions. Briefly, 0.75–1 µg of total RNA from cells was
subjected to reverse transcription with AMV-RT at 55°C
for 30 min. The PCR was performed using 400 nM each of the
following primers: 5'-GTTCTGCTTGAAACATAGGAGCTGCCCTCCTGG-3' or
5'-GCGGCCCCGGGGAGGTTTGCTGTTCCTACAAAGTTT-3' ("upstream
primers"), and 5'-GCGGTCGACTAAGCAGCTTATTTTTACTGA-3' ("downstream
primer"). For RNA loading controls, oligonucleotides specific for the
glyceraldehyde-3-phosphate dehydrogenase (GAPDH) transcript were used:
5'-TGAAGGTCGGAGTCAACGGATTTGGT-3' and
5'-CATGTGGGCCATGAGGTCCACCAC-3'. For tonsillar B cells analysis, the
PCR products were transferred to Genescreen membrane (NEN Life
Sciences, Boston, MA) then probed with an oligonucleotide with the
following sequence: 5'-GGTTTGCTGTTCCTACAAAGTT-3'. A total of 100 ng of
the oligonucleotide was end labeled with 0.02 mCi of
[32P-
]dATP using DNA polynucleotide kinase. The probe
was added to 50 ml of 5x SSPE plus 1% SDS. After overnight
hybridization at 57°C, the membranes were washed with 2x SSPE for 30
min and then 2x SSPE plus 1% SDS for 1 min. Specific hybridization of
the oligonucleotide probe to DNA fragments was visualized by
autoradiography.
Representational difference analysis (RDA) was performed as described (39, 40). The cell substrates used in RDA were kindly provided by Dr. Francine Briere (Schering-Plough, Dardilly, France). Bm2 cells were purified from human tonsils based on the expression of follicular mantle markers, such as surface IgD and CD23, and the absence of GC markers, such as CD38, as described (8, 41). Purified cells were resuspended at 5 x 105 cells/ml in RPMI 1640 medium with 10% FCS. A total of 100–200 ml of the suspension was dispensed in microwells. SAC particles (0.001–0.005%, v/v) or sCD40L (16) were added to the cultures and the cells were harvested after 48 h. Amplicons were constructed by digesting double-stranded cDNA from either SAC or sCD40L-activated Bm2 cells with DpnII for 4 h at 37°C. After digestion R12 (5'-GATCTGCGGTGA-3') and R24 (5'-AGCACTCTCCAGCCTCTCACCGCA-3') linkers were ligated to the double-stranded cDNA with T4 DNA ligase. Amplicons were obtained after 20 cycles of PCR amplification using as template 1/10 of the purified double-stranded cDNA plus linkers ligation reactions. The products of these amplifications were then pooled and purified by phenol extraction and ethanol precipitation, and 3–10 µg of each amplicon product were electrophoresed through a 1% agarose gel, the immobilized on Zeta probe membranes (Bio-Rad, Hercules, CA). After capillary transfer and UV cross-linking, the blots were hybridized with a 32P-labeled double-stranded cDNA probe generated by the PCR radioactive labeling system (Life Technologies), followed by stringent washing, and developed by using a PhosphoImager and the Imagequant software package (Molecular Dynamics, Sunnyvale, CA).
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Results |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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To characterize novel FDC-associated molecules, we constructed a
size selected library from FDC-1 cells and randomly sequenced the cDNA
inserts. One of these clones encoded a TNFR family member, which we
called FDCR-1. Concurrent with our finding, others reported an
identical amino acid sequence for OPG/OCIF, a molecule found to
influence osteoclastogenesis (31, 42). When we compared the
cysteine-rich amino terminus, FDCR-1 had 46% amino acid similarity to
CD40, a key molecule regulating B cell maturation and GC formation (43, 44). FDCR-1 also had high similarity to TNFRII. An alignment with
RANK, CD40, and TNFRII is shown in Fig. 1A. When we compared the
carboxy terminus, we did not find high homology with any known protein;
however, FDCR-1 had some residues at corresponding positions that are
commonly found in death domains of other TNFR family members (Fig. 1B). Interestingly, there is no long region in this sequence
that corresponds to a classical transmembrane region. However,
Yamaguchi et al. (34) have found that the putative cytoplasmic death
domain homology region of OPG/OCIF can transduce an apoptotic signal,
strongly suggesting that OPG/OCIF may be a transmembrane protein. There
are several possibilities which might explain these observations (see
Discussion).
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To test the possibility that OPG/FDCR-1 was expressed on cell
membranes, we generated rabbit heteroantisera to recombinant FDCR-1 and
expressed recombinant, myc epitope-tagged FDCR-1 in BHK
cells. Using these reagents, we found that FDCR-1 is expressed on cell
membranes using flow cytometry (Fig. 2),
surface biotinylation (Fig. 3), and
surface iodination (data not shown). We transfected BHK cells with a
myc epitope-tagged FDCR-1 or a similar construct of a
related molecule myc epitope-tagged DR3, a known membrane
bound member of the TNFR family (45). These cells were indirectly
stained with either an anti-myc Ab or the rabbit
anti-FDCR-1 serum, followed by goat anti-rabbit
F(ab')2-conjugated to FITC. Samples were analyzed by flow
cytometry (Fig. 2). The anti-myc epitope Ab clearly
detected the full-length myc/FDCR-1 construct, the
myc/DR3 construct, but not a truncated version of
myc/FDCR-1 containing only the amino terminus. The MPF fold
increase for the epitope-specific Ab over the negative control was 2.8
times and 3.2 times for the myc/FDCR-1 and
myc/DR3 constructs, respectively. The anti-FDCR-1 serum
detected the FDCR-1 molecule on the surface of the transfected cells
but not the truncated version or myc/DR3. The MPF fold
increase for the specific antisera over the negative control was 2.1.
Neither Ab reacted against cells transfected with vector alone. Thus,
the full-length FDCR-1, including the carboxy region, is required for
expression on the cell membrane. Also, the antiserum is able to detect
FDCR-1 on the surface of FDC-1 cells, the cell line from which we
isolated the cDNA (Fig. 2B). The MPF fold increases for the
epitope specific Ab over the normal rabbit serum (NRS) or secondary Ab
alone negative controls were 2.1 times and 2.3 times, respectively.
Thus, OPG/FDCR-1 is expressed on the surface of some untransfected
cells.
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Under reducing conditions, this protein migrated at 
60 kDa, slightly
higher than a nonspecific protein. Under nonreducing conditions, the
anti-FDCR-1 serum precipitated a protein of 120 kDa, suggesting
that OPG/FDCR-1 is a dimer on cell membranes.
Simonet et al. (31) and Tsuda et al. (42) found that OPG/FDCR-1 was
present in supernatants of transfected cells or untransfected human
embryonic lung fibroblasts. We tested whether FDCR-1 was expressed as a
soluble form in our hands. Using the FDCR-1-BHK cells or nontransfected
FDC-1 cells, we performed a pulse-chase analysis. In Fig. 4, FDC-1 cells were metabolically labeled
with [35S]Met,Cys. Using the anti-FDCR-1 serum, we
immunoprecipitated proteins from either the cell supernatants or whole
cellular lysates at several time points after addition of cold medium.
FDCR-1 was detected in the supernatant within 30 min after addition of
cold medium, and persisted in the supernatant for 10 h (Fig. 4).
Similar results were obtained with FDCR-1-BHK cells. The m.w. of the
soluble form was 60–65 kDa, confirming the results of Simonet et al.
(31). Under nonreducing conditions, the m.w. of soluble OPG/FDCR-1 was
120 kDa, consistent with it being a homodimer.
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Next we probed isolated RNA from either human tissues or cell
lines for their ability to express OPG/FDCR-1 (Fig. 5). In lymphoid tissues, we detected
strong OPG/FDCR-1 expression lymph node, fetal liver, bone marrow, and,
to a lesser extent, in spleen and thymus (Fig. 5D). It is
also expressed in thyroid and spinal cord (data not shown). We detected
OPG/FDCR-1 transcripts in human heart, lung, liver, kidney, and
pancreas. Simonet et al. (31) did not detect OPG expression in human
liver, which is the only difference from this report. The reason for
this difference is not clear. Although the poly(A)+ RNA was
obtained from the same source, we are not certain whether the tissues
were isolated from the same individual or whether technical reasons can
account for this observation. OPG/FDCR-1 does not appear to be
expressed in brain, skeletal muscle, thymocytes, or peripheral blood
lymphocytes (Fig. 5, B–D). Expression was not
detected after activation of peripheral blood lymphocytes with PMA or
after separation into unstimulated T cell or B cell populations (data
not shown).
We also evaluated whether OPG/FDCR-1 was expressed by various cell
lines. We were unable to detect OPG/FDCR-1 transcripts in a number of
hematopoietic lineages or epithelial cell lines (Fig. 5, A
and B). In independent experiments with a different isolate
of HeLa RNA, we detected expression of OPG/FDCR-1, suggesting subline
differences in HeLa cells. OPG/FDCR-1 was expressed in all stages of
smooth muscle cells evaluated (Fig. 5E). The strong
OPG/FDCR-1 expression by heart and smooth muscle cells supports the
possibility that OPG/FDCR-1 has a role in vascular development. In the
developing mouse embryo (E15) Simonet et al. (31) demonstrated that
OPG/FDCR-1 was highly expressed in several regions of the developing
vasculature, and Bucay et al. (33) demonstrated that
opg-/- mice have arterial calcification.
We then extended our expression analysis by using RT-PCR, a method more
efficient and sensitive than Northern blot analysis. Because we
isolated OPG/FDCR-1 from an FDC-like cell line, we wanted to test
whether it is expressed in normal human FDCs. Using Percoll gradient
fractionation and cell sorting for
CD21highCD14+ cells, we isolated highly
enriched FDCs (27). OPG/FDCR-1 transcription was detected in enriched
FDCs (Fig. 6A). Theoretically,
some B cells may still be associated with the FDC clusters (27), so we
cannot be entirely sure whether the FDCs, B cells associated with FDCs,
or both cell types express OPG/FDCR-1. Nevertheless, we can conclude
OPG/FDCR-1 is expressed in FDC clusters.
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A), and EBV B lymphomas, such
as BJAB (Fig. 6B), Ramos and B104 (data not shown), do not
express OPG/FDCR-1. Interestingly, we were able to detect OPG/FDCR-1
expression in EBV-transformed lymphoblastoid cell lines, CESS, T5-1,
and MP-1 (Fig. 6B). Detection of OPG/FDCR-1 expression in
CESS using RT-PCR but not by Northern blot analysis probably reflects
the difference in sensitivity between these two assays. Thus, B lineage
cell lines do express OPG/FDCR-1, and its expression could be
differentially regulated during the B cell activation and maturation
process.
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To test whether activated normal B cells express OPG/FDCR-1, we
isolated tonsillar B cells and fractionated them on a Percoll gradient.
We stimulated each fraction with anti-IgM and anti-CD40 and
then compared changes in OPG/FDCR-1 transcripts by RT-PCR.
Amplification products were probed with an OPG/FDCR-1-specific
oligonucleotide to assure specificity. After activation, OPG/FDCR-1
transcription was up-regulated in all B cell fractions (Fig. 7A). The less dense, in vivo-activated B cells had higher
levels of constitutive OPG/FDCR-1 transcription than the dense
fractions. In the dense fractions, ligation with anti-IgM and
anti-CD40 clearly increased expression.
Because EBV-transformed cell lines generally resemble cells that are
activated through the CD40 pathway, this raised the possibility that
OPG/FDCR-1 expression was regulated by CD40 ligation of B cells. To
test this, we performed representational difference analysis on primary
tonsillar B cells. Using defined markers, we isolated naive follicular
mantle B cells (Bm2) (39). These cells were stimulated with either SAC
or sCD40L and then assayed for expression of OPG/FDCR-1. As shown in
Fig. 7B, we were able to detect a high level of OPG/FDCR-1
expression in CD40-stimulated naive B cells compared to
SAC-activated cells.
DCs are also known to become activated and enhance their T cell
stimulatory capabilities upon CD40 ligation (46, 47). We tested whether
DCs could also up-regulate FDCR-1 expression after CD40 ligation. DCs
were isolated from human peripheral blood mononuclear cells and
differentiated in culture using granylocyte-macrophage CSF and IL-4
(46, 48). We stimulated DCs for 72 h with anti-CD40. By
RT-PCR, we observed a clear up-regulation of FDCR-1 transcription (Fig. 7C).
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Discussion |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Currently, OPG/FDCR-1 has been reported to function in bone metabolism. OPG/FDCR-1 has been demonstrated by two groups to inhibit osteoclastogenesis. Yasuda et al. (32, 49) and Lacey et al. (35) proposed that OPG/FDCR-1 acts as a soluble inhibitor that prevents RANKL/TRANCE interaction with its receptor. In this respect, OPG/FDCR-1 would behave in an analogous way that TRAIL-R3 (TRID) (50, 51) or soluble CD95 (Fas) do in blocking nonlocal interactions between their soluble ligands, i.e., TRAIL or Fas ligand (FasL), respectively, and their membrane bound receptors, TRAIL-R1, -R2, -R4 (52) and membrane CD95 (53), respectively. Another possibility is that RANK and OPG/FDCR-1 may have different abilities to signal through surface bound RANKL/TRANCE. There is evidence that transmembrane forms of TNF family molecules are capable of transducing an activation signal, such as FasL and CD154 (CD40L) (54, 55).
Another model that has not been fully explored is the possibility that OPG/FDCR-1 exists as a transmembrane protein. We have demonstrated that OPG/FDCR-1 is expressed on the surface of normal or transfected cells. The inhibitory activity of the molecule on osteoclastogenesis resides in its amino half, from residues 1 to 198 (31, 34). The function of the remaining carboxy half and the reason for its high evolutionary conservation need to be elucidated. OPG/FDCR-1 may exist on the membrane either bound by the extracellular matrix or as a transmembrane protein. By comparing elution rates from a heparin column of mutant vs intact forms of the molecule, Yamaguchi et al. (34) suggest that OPG/FDCR-1 can bind to the extracellular matrix protein, heparin. The carboxy-terminal region, which we found necessary for membrane expression on transfected BHK and nontransfected FDC-1 cells, is necessary for OPG/FDCR-1 binding to the heparin column. Thus, it is possible that membrane OPG/FDCR-1 functions analogous to extracellular matrix trapped cytokines (56).
However, Yamaguchi et al. (34) also found that the carboxy region of
OPG/FDCR-1, when anchored to a classic transmembrane domain, can
transduce an apoptotic signal, thus suggesting that OPG/FDCR-1 is a
transmembrane protein. We have transfected OPG/FDCR-1 into several cell
types but have detected neither increases in cell death nor NF-B
activation (data not shown). Our biochemical results are consistent
with either a matrix-bound and/or a transmembrane form of the protein.
Interestingly, while OPG/FDCR-1 does not have a classical hydrophobic
region separating its NH2 and COOH regions, when we plotted
one region on a helical wheel, we noted that the charged residues
aligned on one face of an -helix. Thus, transmembrane OPG/FDCR-1
molecules might form homodimers that could possibly associate via these
amphipathic helices and then insert into the membrane. Further
biochemical studies will be necessary to test this possibility. Another
explanation is that an alternatively spliced transcript may encode an
OPG/FDCR-1 transmembrane form, analogous to the related family member,
CD95 (Fas), which is produced in two forms by alternate spliced
transcripts (57). In any case, the function of membrane associated
OPG/FDCR-1 is not yet clear. Possibly, a transmembrane form of
OPG/FDCR-1 could transduce a signal that imparts a differentiation
program to the cell which expresses it. The secreted form of OPG/FDCR-1
prevents systemic effects of the ligand, effectively making the local
interactions between ligand and the membrane form the relevant ones.
Our observation that OPG/FDCR-1 is regulated by CD40 in B cells and DCs further suggests a role for OPG/FDCR-1 in immune functions. The importance of CD40 in various stages of GC formation, B cell differentiation, and memory B cell development is well documented (3, 8, 44, 58, 59). RANK and OPG/FDCR-1 share the features of both being receptors for RANKL/TRANCE and both being up-regulated by CD40 ligation. Also, the expression pattern of OPG/FDCR-1 in hematopoietic tissues somewhat parallels that of RANK; i.e., it can be observed in EBV lymphoblastoid cells, DCs, and certain stromal cells. In DCs, OPG/FDCR-1 expression, like RANK expression, is also up-regulated by CD40 stimulation. In terms of the immune system, the significance of RANK and OPG/FDCR-1 coexpression and coregulation will likely require understanding how the interplay between the two receptors and their ligand, RANKL is regulated.
How OPG/FDCR-1 functions in the regulation of the immune response is unknown. However, OPG/FDCR-1 may function in an analogous manner as it does in bone metabolism. During osteoclastogenesis, RANKL/TRANCE binding to osteoclast progenitors (possibly through RANK) may be an essential signal for their maturation. OPG/FDCR-1 may also block development of progenitor osteoclasts (31, 32). Simonet et al. (31) demonstrated that overexpression of the soluble form of OPG can inhibit osteoclastogenesis, and Yasuda et al. (49) found that soluble OPG can inhibit binding of RANKL/TRANCE to progenitor osteoclasts. According to this model, osteoclast production is balanced by relative amounts of RANK and OPG/FDCR-1. Bone-resorbing signals stimulate osteoblasts or stromal cells to express more RANKL/TRANCE, which results in increased osteoclast differentiation.
From this model, we can propose a similar and testable model for the
function of OPG/FDCR-1 in the immune response. In the immune system,
RANKL/TRANCE expression is relative to T cell restricted (15, 35, 36, 37), and RANKL/TRANCE can enhance DC aggregation and
allostimulation in vitro (15, 37). CD40L+
RANKL+ T cells may interact with cells that express
OPG/FDCR-1; i.e., DCs, B cells, and FDCs. DCs and B lineage cells can
express both RANK and OPG/FDCR-1, and both molecules are up-regulated
after CD40 stimulation. On T cells, RANKL/TRANCE can activate JNK (36)
and NF-B, which may play a role in T cell viability (15). Analogous
to the bone metabolism model OPG/FDCR-1 could be a negative regulator
of RANKL/TRANCE by interfering with RANKL/TRANCE signaling to DCs or T
cells. In this model, OPG/FDCR-1 would attenuate the RANKL/TRANCE
signal given by T cells. Another possibility is that OPG/FDCR-1 exists
as a transmembrane receptor and directly transduces a negative signal
via its death domain (34).
Recently OPG/FDCR-1 was reported to be a fifth receptor for TRAIL (60). TRAIL-R1 and -R2 have active death domain regions (38), and TRAIL-R3, -R4, and soluble OPG/FDCR-1 may serve as decoy receptors for TRAIL (52, 60). However, how TRAIL and its five receptors function in the in vivo immune response is by no means clear. Whether there is a complex cross regulation between ligands RANKL and TRAIL with OPG/FDCR-1 and the other TRAIL receptors will be subject of further investigation.
In summary, the expression of OPG/FDCR-1, the second receptor for RANKL/TRANCE, in lymphoid tissues and by cells of the immune system implicates its involvement in immune responses, which adds to its characterized role in bone metabolism and vascular development. The fact that it is has a very similar expression pattern to the first RANKL/TRANCE receptor, RANK, opens the possibility that OPG/FDCR-1 plays a role in activation of DCs and B cells during the GC response.
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Acknowledgments |
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Footnotes |
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2 Address correspondence and reprint requests to Dr. Edward A. Clark, Department of Microbiology, University of Washington Medical School, 1959 NE Pacific, Box 357242, Seattle, WA 98195. E-mail address:
3 Abbreviations used in this paper: GC, germinal center; DC, dendritic cell; FDC, follicular dendritic cell; OPG, osteoprotegerin; OCIF, osteoclastogenesis inhibitory factor; RDA, representational difference analysis; SAC, staphylococcus aureus Cowan I; TRANCE, TNF-related activation-induced cytokine; MPF, mean peak fluorescence; RANK, receptor activator of NF-B; RANKL, RANK ligand; MACS, magnetic cell sorting; AMV-RT, avian myeloblastosis virus-reverse transcriptase.
4 This work was supported by grants DE08229, RR00166, GM37905. P.M.C. is supported by a fellowship from the Cancer Research Fund of the Damon Runyon-Walter Winchell Foundation.
Received for publication May 21, 1998. Accepted for publication August 3, 1998.
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