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The Journal of Immunology, 2006, 176: 2197-2207.
Copyright © 2006 by The American Association of Immunologists

Expression and Regulation of the Orphan Receptor RDC1 and Its Putative Ligand in Human Dendritic and B Cells1

Simona Infantino*, Barbara Moepps{dagger} and Marcus Thelen2,*

* Institute for Research in Biomedicine, Bellinzona, Switzerland; and {dagger} Department of Pharmacology and Toxicology, University of Ulm, Ulm, Germany


   Abstract
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 Disclosures
 References
 
Based on phylogenetic analysis and chromosomal mapping, the orphan receptor RDC1 was proposed to be a chemokine receptor. In this study we examined the expression of RDC1 on leukocytes by measuring mRNA levels and receptor expression using a new specific mAb. Both mRNA and protein levels were high in monocytes and B cells, relatively low on immature dendritic cells (DC), and up-regulated during final stages of maturation. Strikingly, in mature plasmacytoid DC the mRNA was up-regulated, but did not correlate with protein surface expression. We indeed report that CpG-activated plasmacytoid DC produce a putative ligand for RDC1, which selectively down-regulates RDC1, but not CXCR4 on primary human B cells. RDC1 expression was found to be tightly regulated during B cell development and differentiation. In blood-derived switch memory B cells, the expression of RDC1 appeared to correlate with the ability to differentiate into plasma cells upon activation, suggesting that RDC1 is a marker for memory B cells, which are competent to become Ab-secreting cells.


   Introduction
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 Disclosures
 References
 
Chemokine receptors are defined by their ability to bind one or more chemokines (1). Together with the chemokines (chemotactic cytokines), chemokine receptors function as the main regulators of leukocyte trafficking under homeostatic and inflammatory conditions (2, 3, 4, 5). Chemokine receptors belong to the rhodopsin subfamily of G protein-coupled receptors containing seven transmembrane domains (6, 7). Upon ligand binding the receptors activate different intracellular signal transduction pathways leading to functional responses: chemotaxis, secretion, and transcriptional activation (8). However, chemokine receptors, such as CXCR4, are also expressed on nonhematopoietic and precursor cells where they also can mediate migration, proliferation, development, and survival (9, 10, 11, 12, 13).

Chemokines are characterized by four structurally conserved cysteines that form two essential disulphide bridges. According to the position of the first two cysteines, members of this family have been classified into four groups: the CXC ({alpha}), CC (beta), CX3C ({delta}), and C ({gamma}) chemokines (14). Based on their function and expression pattern, a classification into inflammatory, homeostatic, and dual-function chemokines was proposed (15). Inflammatory chemokines are induced by proinflammatory stimuli to recruit immune cells to sites of inflammation, whereas homeostatic chemokines are constitutively expressed and are required for leukocyte positioning for immune surveillance (16).

Efficient antimicrobial defense and the generation of a specific immune response critically depend on Ag presentation. Dendritic cells (DC)3 have the capacity to present Ag to naive lymphocytes and to initiate adaptive immune responses. Immature DC take up and process Ag in the periphery and migrate to lymphoid tissues where they complete their maturation program (17). Plasmacytoid DC (pDC) and myeloid DC (mDC) were discovered in human blood by their differential expression of Ig-like transcript (ILT) receptors 1 and 3 (18) and CD11c (19). In vitro monocytes can be differentiated into DC by culturing the cells in the presence of appropriate cytokine combinations (20, 21).

B lymphopoiesis starts in the bone marrow from pluripotent hemopoietic stem cells. Progenitor B cells (pro-B cells) initiate rearrangement of the IgH gene locus leading to DH-JH joints. Further rearrangement of the variable gene regions leads to productive VH-D-JH alleles in precursor B cells (pre-B cells). The development of pre-B cells continues with the L chain gene rearrangement generating relative immature/transitional B cells that are ready to migrate to the periphery where they fully differentiate into mature B cells (22, 23). In secondary lymphoid tissues, B and T lymphocytes separate in two clearly distinct zones. During immune responses a germinal center forms within the B cell area (follicles) where naive B cells become activated, receive costimulatory signals from Th cells, and undergo affinity maturation (24, 25). The proliferating B cells, known as centroblasts further develop into nonproliferating centrocytes. Centroblasts and centrocytes segregate into the dark and light zone of the germinal center, respectively.

RDC1 is an orphan G protein-coupled receptor with seven transmembrane domains. Initially, RDC1 was reported to act as a receptor for vasoactive intestinal peptide (26). Subsequently, RDC1 was defined to be the receptor for calcitonin gene-related peptide and later suggested to bind adrenomedullin (27, 28). However, these observations were dismissed and it was shown that RDC1 cannot account for the calcitonin gene-related peptide and adrenomedullin binding (29). Phylogenetic analyses of over 340 human G protein-coupled receptors indicated that within the rhodopsin family the orphan receptor RDC1 belongs to the subgroup of chemokine receptors (7, 30). In fact, RDC1 possesses high sequence similarity with known chemokine receptors (e.g., CXCR2, 43%). The protein contains typical chemokine receptor signatures, such as a DRY motif at the boundary of third transmembrane helix and the second intracellular loop, a CxNPxxY sequence in the seventh transmembrane domain and four conserved cysteine residues in the extracellular segments. The genes encoding chemokine receptors tend to cluster on chromosomes. In this respect chromosomal mapping revealed that the mouse RDC1 gene is located on chromosome 1 in close proximity to CXCR1, CXCR2, and CXCR4 (31, 32). In humans a similar arrangement of these four receptors genes is found on chromosome 2. Like other chemokine receptors RDC1 can act as a coreceptor for certain HIV and SIV (33, 34, 35, 36, 37). Together these evidences support the hypothesis that RDC1 is a potential chemokine receptor. However, little is known on the expression of RDC1 in mammalian tissues. The mRNA of RDC1 was detected in heart, spleen, and kidney (31). In addition some tumors from brain and peripheral vasculature endothelium express mRNA of RDC1 (38). Recently it was reported that Kaposi sarcoma-associated herpesvirus-induced expression of RDC1 is essential for neoplastic transformation of dermal microvascular endothelial cells (39). A mesangial tumor cell line (MES-13) and neutrophils were also shown to contain RDC1 mRNA (31). However, the expression of the protein, in particular on primary leukocytes, was not investigated.

In this study we characterize the expression of RDC1 in human leukocytes using a new mAb. Marked expression of RDC1 was found in monocytes, which somehow decreased during differentiation into immature DC, but was up-regulated during maturation of DC. Mature pDC expressed high levels of receptor mRNA, but not of the protein. In agreement, we observed that CpG-activated pDC produced an activity that selectively down-regulates RDC1 on primary human B cells. In the B cell compartment, RDC1 appears to be tightly regulated during development. In peripheral blood switch memory B cells, RDC1 expression correlates with the capacity to differentiate into plasma cells upon polyclonal activation. The results suggest that RDC1+ memory B cells are competent to become Ab-secreting cells (ASC).


   Materials and Methods
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 Disclosures
 References
 
Abs and streptavidin

The following Abs were used: anti-CD4-PE (13B8.2), anti-CD3-FITC (UCHT1), anti-CD3-PC5 (UCHT1), anti-CD14-FITC (RM052), anti-CD14-PC5 (RM052), anti-CD27-PE (1A4CD27), anti-CD24-PE (M-L233), anti-CD38-PE (T16), anti-CD38-PC5 (LS198-4-3), anti-CD45RA-FITC (ALB11), anti-CD56-PC5 (N901), anti-CD77-FITC (5B5), and anti-ILT3-PE (ZM3.8) (Immunotech); anti-CD19-FITC (4G7), anti-CD19-PE (HIB19), anti-CD19-PE Cy7 (SJ25C1), anti-CD19-allophycocyanin (4G7), anti-human CD123 (IL-3R{alpha}; 9F5), anti-CXCR4-PE (12G5), and streptavidin-allophycocyanin Cy7 (BD Biosciences); anti-CD1c-FITC (ADA-8E7), mAb blood DC Ag (BDCA)-4-PE (ADA-17F6), magnetic microbeads (anti-PE, anti-FITC, anti-CD1, anti-CD9, anti-CD14) (Miltenyi Biotec); goat anti-human IgM-Cy5, IgA-Cy5, or IgG-Cy5 (Jackson ImmunoResearch Laboratories); anti-CD10-FITC (SJ5-1B4) and anti-CD19 TRI-COLOR (SJ25-C1) (Caltag Laboratories); anti-CD11c-FITC (3.9) and CD27-allophycocyanin (O323) (eBioscience); mouse anti-CXCL12 (79018), biotinylated goat anti-CXCL12, and streptavidin-HRP (R&D Systems); Alexa Fluor 594-F(ab')2 of goat anti-mouse IgG, Alexa Fluor 594-F(ab')2 of goat anti-rabbit IgG, Alexa Fluor 488-streptavidin, and streptavidin-allophycocyanin (Molecular Probes); anti-CD20cy (L26), anti-CD3 (T3-4B5), rabbit anti-CD3, biotinylated F(ab')2 of rabbit anti-mouse Ig, and streptavidin-ABC complex/alkaline phosphatase (DakoCytomation) and mAb 97A6-PE (40).

Preparation of anti-RDC1 mAb 9C4

The N terminus of human RDC1 (first 27 aa) was fused with three different carrier proteins: GST, human IgG1, and maltose-binding protein. Recombinant fusion proteins were expressed in bacteria (GST and maltose-binding protein) or S2 insect cells (human IgG1) and purified by affinity chromatography. The recombinant proteins were used for immunization and two booster injections of mice. Hybridoma cells were prepared by standard protocols and positive clones were screened by ELISA using the synthetic peptide (DYSEPGNFSDISWPC) corresponding to aa 7–21 of the N terminus of RDC1. Clone 9C4 expressing IgG1 subclass anti-human RDC1 reacts with 300.19 mouse pre-B cells expressing human RDC1, but not with the corresponding parental cells (41). RDC1 expression in 300.19 cells was confirmed by RT-PCR and staining for a receptor-associated hemagglutinin tag. Abs were purified from culture supernatants using Gamma-Bind Plus Sepharose (GE Healthcare). Monoclonal Ab 9C4 was biotinylated with biotin pentafluorophenyl ester (biotin PFP; Perbio Science) according to the manufacturer’s instructions.

Cell lines

The B cell lines Reh (pro-B), 207 (pre-B) (42), Raji (IgM+), and Daudi (IgM+) were obtained from DSMZ, and Fc4M (IgG+) was provided by A. Lanzavecchia (Institute for Research in Biomedicine, Bellinzona, Switzerland). Cells were cultured in B cell medium (RPMI 1640 supplement with 10% FBS, 1% Glutamax, 1% nonessential amino acids, 1 mM sodium pyruvate, 50 U/ml penicillin, 50 µg/ml streptomycin, and 50 µM 2-ME (all from Invitrogen Life Technologies)).

Cell preparation and culture

PBMC were prepared from buffy coats of healthy donors as described elsewhere (43). Leukocytes were isolated from PBMC using a combination of magnetic and FACS sorting. Briefly, PBMC were incubated with mAb anti-CD4-PE and anti-CD8-FITC for T cell, anti-CD56-PE and anti-CD19-FITC for NK and B cells, and mAb 97A6-PE and anti-CD14-FITC for basophil and monocyte separation. Cells were then enriched with anti-PE and anti-FITC magnetic microbeads on MACS separation columns (LS column; Miltenyi Biotec) and were further sorted by a FACSAria (BD Biosciences) to >99% purity (data not shown).

Monocyte-derived DC were generated as previously described (44). Maturation was induced by the addition of 200 ng/ml LPS from Salmonella abortus equi (Sigma-Aldrich). Both pDC and mDC were sorted by a FACSAria to >99% purity and cultured in 96-well flat-bottom plates (2.5 x 105 cells in 200 µl medium/well) as previously described (43). The pDC were activated with the following maturation stimuli: CpG-A (ODN-2216) 5'-G*G*GGGACGATCGTCG*G*G*G*G*G-3' (5 µg/ml; asterisk represents phosphorothioate linkage) (45), influenza virus (40 hemagglutinin U/ml strain A/Beijing/353/89) a gift from I. Julkunen (National Public Health Institute, Helsinki, Finland), and mDC with LPS (100 ng/ml) for 24 h. PBMC were depleted of pDC by labeling with mAb BDCA-4-PE followed by anti-PE magnetic microbead sorting. Depleted (PBMC of pDC) or nondepleted PBMC (2 x 106) were cultured in the presence of LPS (100 ng/ml) or CpG-A.

Tonsils were obtained from routine tonsillectomies performed at the "Ospedale San Giovanni." After mincing, tonsils were treated with 1 mg/ml collagenase D (Roche Diagnostics) for 45 min at 37°C and tonsillar mononuclear cells were prepared by Ficoll-Paque density centrifugation. Bone marrow cells were prepared by Ficoll-Paque density centrifugation from specimens obtained from "Ospedale San Giovanni."

Crude CD19+ B cell fractions obtained by magnetic microbead sorting as described earlier were stained with anti-CD27-PE, anti-IgA-Cy5, anti-IgM-Cy5, anti-IgG-Cy5, Lineage (anti-CD3-PC5, anti-CD14-PC5, and anti-CD56-PC5), and biotinylated mAb 9C4 followed by Alexa Fluor 488-streptavidin. Naive (CD19+, CD27–, IgA–, IgG–, Lineage–), IgM memory (CD19+, CD27+, IgA–, IgG–, Lineage–), IgA memory (CD19+, CD27+, IgG–, IgM–, Lineage–, RDC1+, RDC1–), and IgG memory (CD19+, CD27+, IgA–, IgM–, Lineage–, RDC1+, RDC1–) B cells were sorted by FACSAria. All B cell subpopulations were >99% pure (data not shown). To determine cell growth, B cell subpopulations were labeled for 7 min at room temperature with 1 µM CFSE (Molecular Probes). Cells were cultured in 96-well flat-bottom plates (2.5 x 104 cells in 200 µl medium/well) in B cell medium. Stimulation of B cells was induced by BCR cross-linking with goat anti-human Ig F(ab')2 (2.5 µg/ml; Jackson ImmunoResearch Laboratories) and addition of 2.5 µg/ml CpG-B (ODN 2006) 5'-T*C*G*T*C*G*T*T*T*T*G*T*C*G*T*T*T*T*G*T*C*G*T*T*-3' (46).

RT-PCR

Total RNA was isolated with TRIzol Reagent (Invitrogen Life Technologies). As a control, human fetal RNA (Stratagene) was used. cDNA was prepared with Superscript III reverse transcriptase (Invitrogen Life Technologies) using oligo(dT) as primer. The cDNA was PCR amplified with Taq polymerase (Euroclone) using the following oligonucleotide primers: RDC1 (5'-TGGTCAGTCTCGTGCAGCAC-3' and 5'-GCCAGCAGACAAGGAAGACC-3'), CXCL12 (5'-CTGTGCACGTTGGAACTTTTATTACTGGGG-3' and 5'-CAAACCCAGTCACTCAAGCGAGCT-3'), and beta-actin (5'-GGCCAACCGCGAGAAGATGACCC-3' and 5'-CAGCGGAACCGCTCATTGCCAATGG-3'). PCR products were resolved by gel electrophoresis.

FACS analysis

Leukocytes were resuspended in PBS, 1% FBS, 0.01% NaN3 and incubated with the appropriate Abs for 15 min on ice. When indicated, staining with secondary Abs or color-conjugated streptavidin was performed for 15 min on ice in the same buffer. Dead cells were excluded by staining with 10 µg/ml propidium iodide.

Cytospins

Mouse pre-B cells (300.19) stably transfected with human RDC1 were centrifuged onto microscope slides (500 rpm for 3 min), fixed with methanol/acetone (1:1), and stained with biotinylated mAb 9C4 (20 µg/ml) followed by Alexa Fluor 488-streptavidin. Wide-field microscopic images were deconvoluted to obtain optical sections of the cells (Open Lab; Improvision).

Immunohistological analysis

Frozen sections (5 µm) were fixed with methanol/acetone (1:1), decorated with the indicated mouse Abs, and stained with biotinylated F(ab')2 of rabbit anti-mouse Ig and streptavidin-ABC complex/alkaline phosphatase according to the manufacturer’s instructions. Nuclei were counterstained with hematoxylin. For fluorescence analysis, sections were stained with rabbit anti-human CD3, mouse anti-human CD20cy, and biotinylated mAb 9C4 followed by Alexa Fluor 594-F(ab')2 goat anti-rabbit IgG or Alexa Fluor 594-F(ab')2 goat anti-mouse IgG (Molecular Probes) and Alexa Fluor 488-streptavidin.

ELISA

At day 7 supernatants from B cell cultures were tested for IgM, IgA, and IgG. ELISA plates (Greiner Bio-One) were coated overnight at 4°C with 10 µg/ml goat anti-human IgM, IgA, or IgG (Southern Biotechnology Associates) and then blocked with PBS containing 10% FCS for 2 h at room temperature. Serial dilutions of culture supernatants (1/10 initial dilution) were added to the wells and incubated for 2 h at 37°C. After washing, alkaline phosphatase goat anti-human IgM, IgA, or IgG was added to the plates. After 2 h at 37°C, plates were developed with FAST pNPP (p-nitrophenylphosphate; Sigma-Aldrich) according to the manufacturer’s instructions. Calibrated human serum was used as standard.

CXCL12 in 24-h culture supernatants derived from pDC was determined on plates precoated with mouse anti-CXCL12 (2 µg/ml). Supernatants and CXCL12 standards (0.75–20 ng/ml) were incubated for 2 h at room temperature. Bound CXCL12 was detected with biotinylated goat anti-CXCL12 (0.2 µg/ml) and streptavidin-HRP. Plates were developed with 0.03% H2O2 in ABTS solution in 0.1 M sodium-citrate (pH 4.35).

RDC1 internalization assay

At day 4 after activation (described earlier), PBMC and PBMC depleted of pDC were stained with CD19-FITC and biotinylated mAb 9C4 followed by streptavidin-allophycocyanin. In coculture experiments, freshly isolated B cells were labeled with CFSE and incubated for 1 h in the presence of pDC. Otherwise B cells were treated for 1 h with supernatants derived from pDC cultures in the presence or absence of 1 µM CXCL12 or with the chemokine alone. Receptor re-expression was measured following treatment of B cells with pDC-derived supernatants for 1 h. Cells were washed twice with medium and incubated at 37°C for different times. When indicated cells were exposed for 90 s on ice to low pH (acidic wash, 100 mM NaCl, 50 mM glycine/HCl (pH 3)), diluted with PBS, 1% FBS, and washed twice (47). RDC1 and CXCR4 surface expression was detected with the respective Abs.


   Results
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 Disclosures
 References
 
Expression of RDC1 in leukocytes

To determine the expression of RDC1 at the protein level, we generated a new mouse mAb that specifically recognizes the N terminus of the human protein (see Materials and Methods). Hybridoma supernatants were screened against a peptide corresponding to aa 7–21 of the N terminus of human RDC1. Clone 9C4 produced an IgG1 that specifically recognizes ectopically expressed human RDC1 on mouse pre-B cells 300.19, Chinese hamster ovary cell and HEK293 cells (data not shown). The anti-RDC1 mAb 9C4 was used to determine receptor expression on PBMC. Fig. 1A shows a marked expression of RDC1 on basophils, monocytes, and B cells, whereas in CD8+ T cells the receptor was barely detectable (data not shown). In CD4+ T cells only a subpopulation (~5%) of CD45RA– (~2%) and CD45RA+ (~2.3%) stained positive. This population stained also heterogeneous for other T cell markers (data not shown), such as CXCR5 (follicular T cells), CD56 (NKT cell), and CD25 (regulatory T cells). On freshly isolated DC, RDC1 was homogeneously expressed on mDC (CD1c+) and pDC (BDCA-4+). However, the level of RDC1 was more pronounced in mDC. RT-PCR analysis of RDC1 mRNA in human PBMC (Fig. 1B) revealed a similar expression pattern being most abundant in B cells, at high levels in monocytes and basophils, and somewhat lower in CD4+ T lymphocytes and NK cells. No messenger could be detected in CD8+ T cells.


Figure 1
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  		FIGURE 1. Analysis of RDC1 expression in human PBMC. A, RDC1 surface expression in PBMC was measured by flow cytometry using biotinylated mAb 9C4. Basophils (97A6+; gate basophils, and DC), CD4+ T cells (CD45RA–/+; gate CD4+, CD14–, CD56–, lymphocytes), and B cells (CD19+) are shown in the upper row. Monocytes (CD14+), mDC (CD1c+; gate CD14–, CD19–, basophils, and DC) and pDC (BDCA4+; gate CD14–, basophils, and DC) are shown in the bottom row. CD4+ T cells and B cells were analyzed using a primary lymphocyte-selective gate in forward and side light scatter. Basophils and DC were measured in an appropriate scatter gate. B, RT-PCR analysis of RDC1 mRNA in PBMC. Total RNA was isolated from sorted PBMC. One representative donor of five is shown.

 
Expression of RDC1 in secondary lymphoid organs

In the light of the distinct expression of RDC1 in blood we investigated the distribution of the receptor in leukocytes from a lymphoid organ. FACS analysis of leukocytes derived from human tonsils revealed a differential distribution of RDC1 in follicle-derived B cells. Fig. 2A shows expression of RDC1 on B cells of the germinal center, the site of Ag-driven proliferation, affinity maturation, and memory B cell generation (24, 25). Both, centroblasts and centrocytes express low levels of RDC1 (Fig. 2A). Conversely, tonsil-derived CD19+ B cells displayed overall a heterogeneous RDC1 expression. In fact, it was possible to detect RDC1high and RDC1low populations (Fig. 2A). However, the heterogeneous expression of RDC1 was similar in naive and memory B cells (data not shown). As in PBMC, only a small fraction (~2–5%) of CD4+ T cells express RDC1.


Figure 2
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  		FIGURE 2. Expression of RDC1 on tonsil-derived leukocytes. A, Surface expression of RDC1 was measured by flow cytometry on gated germinal center centroblasts (CB; CD19+, CD38high, CD77+), centrocytes (CC; CD19+, CD38high, CD77–), CD19+ B cells, mDC (CD11c+, ILT3+, CD3–, CD14–, CD19–, CD56–), pDC (CD11c–, ILT3+, CD3–, CD14–, CD19–, CD56–), and CD4+ T cells (CD4+, CD14–, CD56–). Gray tinted RDC1 (mAb 9C4) (gray-filled histogram) and control (black line histogram) are shown. B, Immunohistochemical analysis of frozen sections from human tonsils. Upper panel images of sections stained for T cells with anti-CD3 (red, left), for B cells with anti-CD20 (red, right), and for RDC1 with 9C4 (green). Images are merged fluorescence, and colocalization of RDC1 with CD3 and CD20 appears yellow. Middle and lower panel images show RDC1 (mAb 9C4), B cells (anti-CD20), T cells (anti-CD3), and pDCs (anti-CD123) in red, and nuclei are counterstained with hematoxylin.

 
Immunohistochemical analysis of frozen sections from tonsils showed a prominent localization of RDC1 in B cell follicles (Fig. 2B). In agreement with the results previously shown, a strong expression of RDC1 on CD20+ cells could be demonstrated by double immunofluorescence staining (Fig. 2B, upper panel, right). RDC1 was predominantly expressed inside the follicle, in the B cell area. In particular, a ring of RDC1+ cells was visible in the internal part of the follicle, an area very close to the mantle zone (Fig. 2B, upper panel, left). Only few T cells were RDC1+. In Fig. 2B (middle and lower panels), two sets of serial sections counterstained with hematoxylin are shown. Again, RDC1 was found in B cell area (CD20+), inside the follicle.

Determination of RDC1 expression in tonsil-derived DC revealed a peculiarity. FACS analysis showed a marked difference between the few mDC that can be isolated from tonsils and the pDC (Fig. 2A). RDC1 was highly expressed on mDC (CD11c+, ILT3+, CD3–, CD14–, CD19–, CD56–), but could not be detected on pDC (CD11c–, ILT3+, CD3–, CD14–, CD19–, CD56–). Similarly, immunohistochemistry indicates that CD123-positive pDC surround the follicles in a region that is clearly negative for RDC1+ cells (Fig. 2B). The observations suggest that under ongoing or late immune responses, as normally found in specimens from tonsillectomies, RDC1 is down-regulated on these cells compared with pDC isolated from the circulation.

Regulated expression of RDC1 in DC

DC exhibit a specific chemokine receptor expression profile that is tightly regulated during differentiation and maturation (48). Although immature DC express receptors such as CXCR1, CCR1, CCR2, and CCR5, which bind chemokines produced at sites of inflammation, maturation of the cells induces reprogramming leading to the up-regulation of receptors for homing chemokines (44). We therefore tested whether maturation of DC results in an alteration of RDC1 expression. FACS analysis of cultured blood-derived immature DC revealed RDC1 protein expression as found on freshly isolated cells (Fig. 3A). In vitro monocyte-derived DC also express RDC1 like their progenitors. Maturation of DC unveiled interesting differences. Treatment of mDC and monocyte-derived DC with LPS induced an up-regulation of RDC1, which was most prominent after 24 h on matured monocyte-derived DC (Fig. 3A). Similarly, treatment of pDC with influenza virus, a TLR7 ligand, induced a moderate up-regulation of RDC1. By contrast, stimulation of pDC maturation with the TLR9 agonist CpG-A (ODN 2216) resulted in a complete loss of RDC1 surface expression, resembling the situation in tonsillar pDC.


Figure 3
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  		FIGURE 3. RDC1 expression on activated DC. RDC1 surface expression was measured by flow cytometry. A, pDC (middle row) and mDC (lower row) were cultured for 24 h in medium or in the presence of influenza virus (Flu, 40 hemagglutinin U/ml), CpG-A (ODN 2216, 5 µg/ml), or LPS (100 ng/ml). Monocyte-derived DC (moDC; upper row) were treated for the indicated times with LPS (200 ng/ml). RDC1 (mAb 9C4) (gray-filled histogram) and control (black line histogram) without primary Ab are shown. B, Monocyte-derived DC were activated as indicated. Medium control was measured after 20 h (top panel). RDC1 mRNA levels were determined in freshly isolated pDC and mDC and after 24 h of culture as described (bottom panels).

 
RT-PCR analysis in DC revealed a marked up-regulation of RDC1 mRNA under all conditions of maturation (Fig. 3B). LPS-induced maturation of mDC mirrored the results obtained by FACS analysis, showing an increased mRNA level after 24 h. Over time a continuous raise in messenger content was observed in monocyte-derived DC following treatment with 200 ng/ml LPS. In pDC TLR7-mediated maturation induced a marginal elevation of RDC1 mRNA, which is in agreement with the moderate increase in RDC1 surface expression measured by FACS analysis. However, treatment of pDC with CpG-A resulted in a marked up-regulation of RDC1 mRNA, contrasting the observed loss of surface expression of the receptor under these conditions. Stimulation of TLR9 with CpG-A is known to induce the production of type I IFNs and chemokines in pDC (43, 49). Thus, it is conceivable that pDC may produce an activity that can trigger RDC1 internalization.

pDC-released factors induce internalization of RDC1

Chemokine receptors become rapidly internalized upon exposure to the appropriate chemokines (8, 50). We reasoned that the loss of receptor surface expression on pDC in the presence of up-regulated mRNA levels could be due to an autocrine secretion of a specific ligand for RDC1. To substantiate this hypothesis, we treated PBMC and PBMC depleted of pDC with LPS or CpG-A. Stimulation with CpG-A was chosen because this ligand selectively activates TLR9 in pDC inducing the secretion of IFN-{alpha}, but does not stimulate TLR9 expressed in B cells (51). In cultures of PBMC that were treated for 4 days with CpG-A, a marked down-regulation of RDC1 on CD19+ B cells was observed by flow cytometry (Fig. 4A). Conversely, in cultures of PBMC that were depleted of pDC, only a minor reduction of RDC1 expression on CD19+ cells was found. These data support the view that pDC secrete ligands that induce RDC1 internalization. Addition of LPS to either culture condition had no effect on RDC1 expression on B cells (Fig. 4A). Furthermore, CpG-A treatment of leukocytes other than pDC appears not to cause the production of an activity that induced RDC1 internalization.


Figure 4
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  		FIGURE 4. Activated pDC release factors that induce internalization of RDC1. A, PBMC (left) and PBMC depleted of pDC (PBMC-pDC, right) were cultured for 4 days in medium alone ({square}) or in the presence of 100 ng/ml LPS ({cjs2108}) or of 5 µg/ml CpG-A ({blacksquare}). RDC1 expression as a percentage was measured by flow cytometry on CD19+ cells. Expression in cells cultured with medium alone was set to 100% expression. Mean values (±SD) represent duplicate determinations in cells from three donors (p < 0.02). B, pDC were treated for 24 h with medium (NA), influenza virus (Flu, 40 hemagglutinin U/ml), and CpG-A (5 µg/ml). CD19+ cells isolated by magnetic sorting were added to the pDC cultures for 1 h (left) or incubated for 1 h with supernatants (sup) harvested from the pDC cultures (right). RDC1 expression was measured on CD19+ B cells by flow cytometry. Control is designated without primary Ab. One representative experiment of three is shown. C, B cells were treated with supernatants of CpG-activated pDC (CpG-A, 5 µg/ml) for 1 h. After washing the expression of RDC1 was measured at different times (0, 15, and 60 min) on CD19+ cells by flow cytometry. RDC1 expression following treatment with control cell culture medium obtained in the absence of CpG-A was set to 100%. D, Cytospins of the 300.19 mouse pre-B cells stably expressing human RDC1 incubated with medium (left) or treated with supernatants from CpG-A-activated pDC (right). Cytospins were stained with mAb 9C4 to reveal RDC1 expression (green). Nuclei were stained with DAPI (blue). Insets depict enlargements of the corresponding areas of cytoplasms.

 
To confirm the ability of pDC to produce RDC1 ligands, pDCs were isolated and treated separately for 24 h with influenza virus (TLR7) and CpG-A (TLR9). B cells were added to unstimulated and stimulated pDC cultures and after 1 h analyzed for RDC1 expression (Fig. 4B, left). When the pDC were pretreated with CpG-A a strong internalization of RDC1 on B cells was observed. By contrast, RDC1 surface expression was not affected when pDC were preincubated with medium or with influenza virus. We next tested whether a secreted ligand from pDC induces RDC1 internalization. Thus, supernatants from cultures of pDC treated with medium or CpG-A were harvested after 24 h and added to B cells for 1 h. Fig. 4B shows that supernatants from CpG-A-treated cell cultures retained full activity in internalizing RDC1 on B cells, indicating that a soluble factor released from pDC triggers receptor endocytosis. Treatment of pDC with CpG-B (ODN 2006), which is considered to be a stronger agonist of TLR9 (52), but does not induce type I IFN in pDC, also lead to the production of a RDC1 internalizing activity (data not shown).

Internalized chemokine receptors are either recycled to the surface or become lysosomally degraded (8, 53, 54). We therefore investigated the endocytic trafficking of RDC1 in primary human B cells after treatment with culture supernatants derived from CpG-A-treated pDC (Fig. 4C). After 1 h of exposure to cell culture supernatants, CD19+ B cells were washed and subsequently incubated at 37°C to follow receptor recycling. Cell surface RDC1 was measured at different times by flow cytometry. Fig. 4C shows that the pDC-derived supernatant caused the internalization of ~50% of RDC1. Surface re-expression of the receptor was detected within the first 15 min and only a minor further increase in RDC1 density was observed for up to 60 min. The observation indicates that only a small part (15–20%) becomes degraded upon internalization of the receptors and that recycle is the predominant pathway.

Intracellular localization of RDC1 after treatment with supernatants derived from CpG-activated pDC was also confirmed by fluorescence microscopy. Cytospins of mouse 300.19 pre-B cells expressing human RDC1 were stained with biotinylated mAb 9C4 (Fig. 4D). Following 30 min exposure to pDC-derived supernatants, a strong increase in intracellular RDC1 could be observed. Immunofluorescence corresponding to RDC1 was detected on endosomal structures throughout the cytoplasm.

Recently, Balabanian et al. (41) reported that the chemokine CXCL12 binds RDC1 expressed on T cells. Therefore, we tested whether activated pDC may produce CXCL12. mRNA was prepared from resting and TLR7- and TLR9-activated pDC and analyzed for CXCL12 content. Fig. 5A reveals that under either condition no mRNA for CXCL12 could be detected. Similarly, ELISA did not indicate any possible secretion of CXCL12 by CpG-treated pDCs (Fig. 5B), indicating that the RDC1-internalizing activity in supernatants of pDC is not CXCL12. Fig. 5C further confirms this conclusion. Supernatants from CpG-treated pDC induced a marked internalization of RDC1, which was not increased when added together with 1 µM CXCL12. By contrast, CXCL12 alone or in the presence of supernatants from untreated pDC induced only borderline RDC1 internalization. To confirm the lack of activity of the chemokine on RDC1 internalization in B cells, we determined the effect of CXCL12 on both RDC1 and CXCR4. Double staining for CXCR4 and RDC1 on B cells demonstrates that CXCL12 triggers selectively the internalization of CXCR4, but has no effect on RDC1 expression (Fig. 5D). Finally, we investigated whether supernatants from CpG-treated pDC may induce the internalization of CXCR4. To exclude the possibility that potential ligands in supernatants from pDC could interfere with Ab binding, cells were submitted to a brief acidic wash to remove bound ligands (47). Fig. 5E shows that culture supernatants from pDC have no affect on CXCR4 expression on CD19+ B cells, whereas a marked internalization of RDC1 was obtained with supernatants derived from CpG-treated pDC.


Figure 5
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  		FIGURE 5. Activated pDC release factors that are distinct from CXCL12. A, RT-PCR analysis of CXCL12 mRNA expression in control and activated pDC. Total RNA was isolated from sorted pDC incubated in medium alone and in the presence of TLR7 influenza virus (Flu, 40 hemagglutinin U/ml) or TLR9 (CpG-A, 5 µg/ml) ligands. Total fetal human cDNA was used as control. One representative donor of three is shown. B, CXCL12 production in culture supernatants of control and CpG-A (5 µg/ml, 24 h) treated pDC was measured by ELISA. Standards ({cjs2108}) were obtained with synthetic CXCL12 (0.75–20 ng/ml). Medium from control pDC (NA, {square}) and CpG-A-treated cells ({blacksquare}) are shown. Mean values are from two independent experiments. C, CD19+ cells were incubated for 1 h with supernatants derived from pDC cultures (NA or CpG-A) in the presence or absence of 1 µM CXCL12. RDC1 expression was measured by flow cytometry. Black line histogram represents untreated cells (NT). D, B cells were incubated for 1 h in the presence or absence 1 µM CXCL12 at 37°C. RDC1 and CXCR4 expression were determined with biotinylated 9C4 and 12G5-PE, respectively. E, CD19+ B cells were incubated with supernatants from untreated (NA) and pDC activated with CpG-A (5 µg/ml). Surface expression of RDC1 (mAb 9C4) and CXCR4 (mAb 12G5) was measured after a brief acidic wash (47 ) to remove potential ligands that could interfere with Ab binding.

 
RDC1 expression during B cell development

The marked levels of RDC1 on blood- and tonsil-derived CD19+ B cells (Figs. 1 and 2) prompted us to investigate the expression of the receptor during B cell development. First, we tested a panel of B cell lines representing different stages of B cell development to measure a potential regulation of RDC1 expression. FACS analysis of RDC1 surface expression in pro-B (Reh), pre-B (207), IgM+ (Raji, Daudi), and IgG+ (Fc4M) B cell lines shows an increase in mean fluorescence intensity as the cells represent more mature stages of B cell development (Fig. 6A). Similarly, we found that the levels of RDC1 mRNA were augmented during B cell maturation. Fig. 6B illustrates that the mRNA is almost absent in pro-B (Reh) and pre-B (207) cells, but becomes strongly up-regulated at later stages when the cells express membrane IgM (Raji, Daudi) and IgG (Fc4M).


Figure 6
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  		FIGURE 6. Regulation of RDC1 expression during B cell development. RDC1 was measured in B cell lines (A and B) and bone marrow-derived leukocytes (C). FACS analysis of protein (A) and RT-PCR analysis of mRNA (B) levels of RDC1 were determined in B cell lines representing different stages of development: Reh (pro-B), 207 (pre-B), Raji (IgM+), Daudi (IgM+), and Fc4M (IgG+). C, Bone marrow B cells were stained for CD10, CD24, CD38, IgM, and RDC1. Three populations of B cells (left) can be identified in the CD24+CD38high gate: pro-pre-B (CD10+, IgM–), immature B cell (CD10+/low, IgMlow), and transitional B cell (CD10low/–, IgMhigh). Histograms of RDC1 expression (right) are shown for the B cell populations at the left. Dead cells stained with propidium iodide were excluded from analysis.

 
Hemopoietic precursors differentiate in the bone marrow into immature and transitional B cells. The latter migrate to the periphery and complete differentiation into mature B cells (23). To corroborate the observations made in B cell lines we measured RDC1 surface expression in developing B cells from human bone marrow. Fig. 6C shows that pro-pre B cells (IgM–) were weakly positive for the receptor, but during development the levels of RDC1 increased from the relative immature (IgMlow) to the transitional (IgMhigh) B cells. The findings essentially confirm the differential regulation of RDC1 expression observed in the B cell lines.

RDC1 expression on switch memory B cells correlates with the ability to become plasma cells

Analysis of mature CD19+ B cell subtypes reveals a heterogeneous appearance of RDC1 (Fig. 7A). IgM+ memory (CD27+) and naive (CD27–) cells strongly express RDC1, whereas switch memory B cells (IgA+ and IgG+, CD27+) contain two apparent populations displaying high (RDC1+) and low (RDC1–) levels of the receptor. It is known that chemokine receptors play a critical role during differentiation of B cells into plasma cells (55, 56). We reasoned that a potential correlation may exist between RDC1 expression and the ability of the different B cell populations to differentiate into ASC. To this end we investigated B cell differentiation in response to polyclonal activation in vitro. B cells were isolated from PBMC with anti-CD19 magnetic beads and subsequently sorted into naive (CD27–) and memory (CD27+) IgM+ cells. Switch memory B cells (IgA+ CD27+ and IgG+ CD27+) were sorted into RDC1+ and RDC1– populations. The efficiency of the FACS separation of RDC– and RDC1+ was verified by reanalyzing receptor expression in the sorted populations (Fig. 7A). The cells were then labeled with CFSE to monitor cell division and stimulated via BCR cross-linking in the presence of CpG-B, which triggers TLR9 in B cells (51). At day 5 the efficiency of CD38high CD27+ plasma cell generation for each condition was determined (Fig. 7B). In agreement with previous findings (57), BCR cross-linking in the presence of CpG-B is not sufficient to differentiate naive B cells into plasma cells, however the same treatment induces a marked differentiation of IgM+ memory B cells into ASC (29%). Inspection of the switch memory pools revealed an interesting novel correlation between the initial level of RDC1 expression and the ability of cells to differentiate into ASC. Essentially, IgA+ and IgG+ switch memory cells were able to differentiate into plasma cells (34 and 14.5%, respectively) only when the cells were RDC1-positive. By contrast, RDC1– switch memory cells poorly became ASC (2.6% of IgA+ and 0.5% of IgG+).


Figure 7
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  		FIGURE 7. RDC1+ switch memory B cells can differentiate into plasma cells. A, FACS analysis of RDC1 expression on PBMC-derived B cells: naive B cells (CD19+, IgM+, CD27–, Lineage–), memory B cells (CD19+, IgM+, CD27+, Lineage–), and switch memory (CD19+, IgA+, CD27+, Lineage–) and (CD19+, IgG+, CD27+, Lineage–). B, Plasma cell differentiation of B cell cultures was induced via BCR cross-linking with 2.5 µg/ml goat anti-human Ig F(ab')2 in the presence of 2.5 µg/ml CpG-B (ODN 2006). Sorted (gates indicated at right in A) naive, IgM+ memory, IgA+ RDC1+, IgA+ RDC1–, IgG+ RDC1+, and IgG+ RDC1– B cells were labeled with CFSE. Following 5 days of stimulation CD38 and CD27 expression was determined. CD38highCD27+ plasma cells in each B cell population are indicated as a percentage. C, Cell proliferation (CFSE) and cell number (#) were measured after 5 days on matched culture aliquots. Histograms of switch memory B cells (IgA+, left and IgG+, right), which were at day 0 for RDC1+ (gray-filled histogram) and RDC1– (black line histogram), are shown. Events were acquired for a fixed time. D, After 7 days of culture as in C, Ig production was measured in supernatants with ELISA: IgM production (left) by memory ({cjs2108}) and naive ({blacksquare}) IgM+ cells; IgA production (center) by IgA+ RDC1+ ({cjs2108}) and IgA+ RDC1– ({blacksquare}), and IgG production (right) by IgG+ RDC1+ ({cjs2108}) and IgG+ RDC1– ({blacksquare}) switch memory B cells. Representative data shown are from one experiment of four.

 
Comparing proliferation and survival of RDC1+ and RDC1– switch memory B cells revealed another interesting difference. During polyclonal activation for 5 days of culture both populations proliferated almost equally. However, survival of RDC1– switch memory (IgA+ and IgG+) was strongly reduced as disclosed by >10 times lower cell numbers with respect to RDC1+ cells (Fig. 7C). Similarly, determination of Ig production 7 days following polyclonal stimulation confirmed that the RDC1+ switch memory B cells were able to differentiate into ASC and secrete Ig (Fig. 7D). Consistent with the frequency to develop into plasma cells, the highest concentrations of IgA or IgG were found in supernatants of RDC1+ B cell subsets. Similarly, following activation IgM was strongly secreted by IgM+ memory but not by naive B cells. Taken together these findings suggest that RDC1 is a potential marker for the ability of memory cells to differentiate into plasma cells.


   Discussion
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
Disclosures
 References
 
In this study, we characterized the expression pattern of the orphan receptor RDC1 in leukocytes. We observed a clear up-regulation of RDC1 during DC maturation. In CpG-matured pDC the mRNA was markedly increased, but did not correlate with surface expression of RDC1. Indeed we report that these pDC secrete a ligand that specifically induces the down-regulation of RDC1. A prominent expression of the receptor at the mRNA and at the protein level was also found in circulating B cells. RDC1 expression, however, appears to be tightly regulated during B cell development and differentiation. Moderate RDC1 expression is observed on pro-B and pre-B cells and becomes gradually up-regulated during development into the relative immature/transitional B cell state, supporting the view that the receptor is required for mature B cell function. Accordingly, naive and IgM+ memory cells express uniformly substantial levels of RDC1. In contrast, in switch memory B cells, receptor expression correlates with the ability to produce Ig in vitro, suggesting that RDC1 might be essential for survival and differentiation of the switch memory cohort. Accordingly, after 5 days of activation, RDC1+ switch memory B cells displayed an >10-fold higher cell number with respect to RDC1– B cells. Thus, it is conceivable that RDC1 might support antiapoptotic signals reminiscent to CXCR4 in CD34+ cells (11).

The expression of RDC1 on DC is intriguing. In pDC treatment with a TLR9 ligand leads to a marked up-regulation of the mRNA but not of the protein. This apparent paradox could be explained by our finding that CpG treatment of pDC leads to the secretion of an activity, which strongly down-regulates RDC1 on B cells and monocytes (data not shown). Consistent with such view, we found no RDC1 expression on pDC isolated from tonsils. The lack of detection of RDC1 even on pDC in frozen sections argues that during persistent immune responses the receptor becomes completely down-regulated and degraded. However, in vitro activation of pDC, mDC, or monocyte-derived DC with inflammatory stimuli leads to increased mRNA expression, which resembles the expression pattern of homing receptors, such as CCR7 and CXCR4, during primary response phase of DC (44, 58).

The observation that treatment of pDC with TLR9-activating CpG induces the release of an activity that efficiently internalizes RDC1 on peripheral B cells is interesting. Generally, chemokine receptor internalization is ligand-induced and depends on receptor-mediated signal transduction leading to its phosphorylation, binding of arrestins, and endocytosis via clathrin-coated vesicles (53). Thus, internalization of RDC1 by pDC-derived supernatants suggests the presence of a specific receptor-activating ligand. We observed that CpG-A, CpG-B, and CpG-C (data not shown), which produce high, none, and intermediate levels of type I IFNs (52, 59), respectively, were almost equally efficient in stimulating pDC to secrete a RDC1 internalizing activity. The lack of correlation with type I IFN production suggests that TLR9 activates different pathways in pDC for the secretion of the cytokine and for the potential RDC1 ligand and that type I IFN may be dispensable for the latter process. Recently, Balabanian et al. (41) reported that CXCL12 (SDF-1) binds to RDC1 when expressed on T cells. However, in agreement with a report previous (49) we could detect neither CXCL12 mRNA in pDC nor the protein in culture supernatants of activated cells. Moreover, addition of 1 µM CXCL12 only marginally induced RDC1 internalization on B cells; conversely, pDC-derived culture supernatants did not affect CXCR4 expression. We have tested over 40 human chemokines including several derivates of CXCL12 (60, 61) for their ability to internalize RDC1 expressed on B cells without success. Together our observations strongly support the hypothesis that pDC release a selective ligand for RDC1 that is distinct from known chemokines. Characterization of the potential ligand of RDC1 will provide more insights. However, purification of the activity from cultures supernatants of CpG-treated pDC or identification by comparative gene expression studies will require some time.

The finding that pDC release an RDC1 internalizing activity suggests that the receptor participates in the communication between pDC and B cells. During inflammation pDC infiltrate lymph nodes settling near high endothelial venules (18) and near follicles (Fig. 2B). Although we have no direct evidence for RDC1-dependent B cell migration as shown for T cells (41), it is conceivable that pDC recruit or contribute to the activation of B cells through this receptor. pDC produce large amounts of type I IFNs and IL-6 that stimulate memory B cell differentiation into plasma blasts and ASC (18, 62). Our findings that in vitro RDC1+ memory and switch memory cells have a stronger propensity to survive and to differentiate into ASC backs the hypothesis that RDC1 is part of a pDC-B cell interaction program. However, a direct contribution of RDC1 to the B cell differentiation into ASC remains speculative. We also noticed a decrease in RDC1 expression in proliferating IgM+ memory B cells in vitro (data not shown) as well as in centroblasts and centrocytes (Fig. 3). Thus, it can be envisaged that at advanced stages of B cell differentiation into ASC, the cells may not require functional RDC1 expression. Accordingly, bone marrow plasma cells stained only slightly positive for RDC1 (data not shown). Thus, RDC1 expression on circulating memory B cells appears to be a critical marker for plasma cell differentiation potential and correlates with the ability of the cells to secrete Ig.


   Acknowledgments
 
We thank Dr. L. Leoncini, Bellinzona, Switzerland, for providing bone marrow samples, Dr. D. Jarossay for assistance in cells sorting, Dr. K. Karjalainen for expression of fusion proteins in S2 cells, and Dr. A. Lanzavecchia and Dr. S. Wirths for critical reading of the manuscript.


   Disclosures
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 Disclosures
References
 
The authors have no financial conflict of interest.


   Footnotes
 
The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked advertisement in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

1 This work was supported by the Helmut Horten Foundation, the Swiss National Science Foundation (to M.T.), and the European Community Framework Programme 6-Network of Excellence Grant LSHG-CT-2003–502935. Back

2 Address correspondence and reprint requests to Dr. Marcus Thelen, Institute for Research in Biomedicine, Via Vincenzo Vela 6, CH-6500 Bellinzona, Switzerland. E-mail address: marcus.thelen{at}irb.unisi.ch Back

3 Abbreviations used in this paper: DC, dendritic cell; pDC, plasmacytoid DC; mDC, myeloid DC; ILT, Ig-like transcript; BDCA, blood DC Ag; ASC, Ab-secreting cell. Back

Received for publication August 24, 2005. Accepted for publication December 9, 2005.


   References
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 Disclosures
 References
 

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