Particular alleles of HLA contribute to disease susceptibility and severity in many autoimmune conditions, but the mechanisms underlying these associations are often unknown. In this study, we demonstrate that the shared epitope (SE), an HLA-DRB1–coded sequence motif that is the single most significant genetic risk factor for erosive rheumatoid arthritis, acts as a signal transduction ligand that potently activates osteoclastogenesis, both in vitro and in vivo. The SE enhanced the production of several pro-osteoclastogenic factors and facilitated osteoclast (OC) differentiation in mouse and human cells in vitro. Transgenic mice expressing a human HLA-DRB1 allele that code the SE motif demonstrated markedly higher propensity for osteoclastogenesis and enhanced bone degradation capacity ex vivo. In addition, the SE enhanced the differentiation of Th17 cells expressing the receptor activator for NF-κB ligand. When the two agents were combined, IL-17 and the SE enhanced OC differentiation synergistically. When administered in vivo to mice with collagen-induced arthritis, the SE ligand significantly increased arthritis severity, synovial tissue OC abundance, and bone erosion. Thus, the SE contributes to arthritis severity by activating an OC-mediated bone-destructive pathway. These findings suggest that besides determining the target specificity of autoimmune responses, HLA molecules may influence disease outcomes by shaping the pathogenic consequences of such responses.
Many autoimmune diseases associate with particular HLA alleles, but the mechanistic basis of these associations is often unknown. Presentation of self-Ags by HLA allele products has been implicated in some cases, but in many others, the mechanism is unclear (1). Based on structural, functional, and evolutionary considerations, we have recently proposed that HLA molecules may contribute to disease pathogenesis through aberrant innate signaling by HLA allele–coded ligands (2). In this study, we have put this hypothesis to the test in an experimental model of an emblematic HLA-associated disease, rheumatoid arthritis (RA).
RA is a crippling disease that afflicts 0.6–1% of the world population. The main manifestation of the disease is chronic joint inflammation and bone erosions, due to overabundance of activated osteoclasts (OCs) in synovial tissues (3–5). Although the pathogenesis of RA is poorly understood, it is clear that genetic factors, particularly the HLA-DRB1 locus (6, 7), play a major role in disease susceptibility. It has been long observed that HLA-DRB1 alleles coding a 5-aa sequence motif called the shared epitope (SE) in the region 70–74 of the DRβ-chain are found in the vast majority of patients with RA (7). The SE not only confers a higher risk for RA, but also increases the likelihood of developing a more severe disease. SE-coding HLA-DRB1 alleles are associated with earlier onset of arthritis and more severe bone erosions (8–11). Furthermore, there is evidence of a gene-dose effect, in which the severity of bone destruction in RA correlates positively with the number of SE-coding HLA-DRB1 alleles (9–11).
The underlying mechanisms by which the SE affects susceptibility to, and severity of, RA are unknown. The prevailing hypothesis postulates that the SE allows presentation of putative self- or foreign arthritogenic Ags (12); however, the identities of such target Ags remain elusive. We have recently demonstrated that the SE functions as a signal transduction ligand that binds to cell-surface calreticulin (CRT) in a strictly allele-specific manner and activates NO-mediated pro-oxidative signaling (13–16). The SE ligand is effective in several different structural formats: in its native conformation as part of cell-surface SE-positive HLA-DR molecules or as SE-expressing HLA-DR tetramers; as cell-free non-HLA recombinant proteins genetically engineered to express the SE motif in its native α helical conformation; or as SE-positive short synthetic peptides. The functional consequences of SE ligand–activated signaling vary, dependent on the cell type. For example, in CD8+CD11c+ dendritic cells, the SE inhibits the activity of IDO, an enzyme known to play an important role in regulatory T cell activation. In CD8−CD11c+ dendritic cells, the SE triggers production of IL-6 and IL-23, cytokines known to be involved in activation and expansion of IL-17–producing T (Th17) cells. The end result of these two complementing effects is a potent SE-activated Th17 polarization, both in vitro and in vivo (17).
Th17 cells are central players in RA pathogenesis (18). Relevant to the focus of this study, these cells have been previously shown to activate osteoclastogenesis by several mechanisms. In addition to their direct pro-osteoclastogenic effect through IL-17 production, Th17 cells express high levels of the receptor activator for NF-κB (RANK) ligand (RANKL), a key factor in osteoclastogenesis (19). Concurrently, Th17 cells activate local inflammation that involves cytokines, such as IL-6, IL-1, and TNF-α, which further increase RANKL expression and synergistically promote osteoclastogenesis (20). Finally, IL-17 can increase RANK expression on the surface of OC precursor cells and thereby sensitize them to the osteoclastogenic effect of RANKL (21).
Given the fact that the SE activates Th17 polarization, the key role of these cells in osteoclastogenesis, and the known association between the SE and erosive disease, we have undertaken to investigate whether the SE has a direct pro-osteoclastogenic effect. The rationale for this study is further strengthened by the fact that the SE is a potent activator of NO and reactive oxygen species (ROS) (13–16), signaling molecules that have been previously shown to affect the recruitment, differentiation, and activation of OCs (22–25).
In this study, we demonstrate that the SE ligand has a dual effect on OC differentiation. It directly enhanced production of pro-osteoclastogenic factors and facilitated OC differentiation and functional activation. In addition, the SE ligand enhanced Th17+RANKL+ cell differentiation. When administered in vivo to mice with collagen-induced arthritis (CIA), the SE ligand increased disease severity, synovial OC abundance, and bone destruction.
Materials and Methods
Mice and reagents
Experiments were carried out in 6–10-wk-old DBA/1 mice as well as a DBA/1 mouse line carrying transgenic (Tg) collagen type II (CII)-specific TCR (D1Lac.Cg-Tg [TCRa,TCRb]24Efro/J), designated in this study as DBA/1 CII-TCR Tg mice, kindly provided by Dr. Steven Lundy (University of Michigan Medical School). All mice were from The Jackson Laboratory (Bar Harbor, ME). Tg mice, which endogenously express SE-positive HLA-DR4 (0401) or SE-negative HLA-DR4 (0402) alleles, were also used. These animals were kindly provided by Dr. Chella David [Mayo Clinic (26)] and are referred to as DRB1*0401 Tg and DRB1*0402 Tg mice, respectively. All mice were maintained and housed in the University of Michigan Unit for Laboratory Animal Medicine facility, and all experiments were performed in accordance with protocols approved by University of Michigan Committee on Use and Care of Animals.
M-CSF, RANKL, human rTGF-β, and rIL-1β, as well as murine rIL-4, rIL-23, rGM-CSF, rIL-17, rIL-6, rM-CSF, and rRANKL, were purchased from PeproTech (Rocky Hill, NJ). Denatured chicken CII was purchased from Chondrex (Redmond, WA). CFA containing Mycobacterium tuberculosis H37Ra was purchased from BD Difco (Franklin Lakes, NJ). Rabbit anti-mouse CRT Ab was purchased from Thermo Scientific. Rabbit BCL–X s/l (S-18) polyclonal Ig was purchased from Santa Cruz Biotechnology. Rat anti-mouse IL-6 and anti–TNF-α Abs, along with isotype-matched control Abs, were purchased from R&D Systems (Minneapolis, MN). PE-conjugated anti-mouse RANK (clone R12.31) was purchased from BioLegend (San Diego, CA). mAbs against mouse CD3 (clone 2C11), IL-4 (clone 11B11), IFN-γ (clone R46A2), and IL-2 (clone S4B6) were purified from the supernatants of hybridomas obtained from the University of Michigan Hybridoma Core Facility. Purified anti-mouse CD28 (clone 37.51) and murine rIL-23 were purchased from eBioscience (San Diego, CA). FITC anti-mouse CD4 (clone GK 1.5), PE, or APC-conjugated anti-mouse IL-17A mAb (clone TC11-18H 10.1) and PE anti-mouse CD254 (TRANCE, RANKL; clone IKK22/5) were purchased from BioLegend. All other commercial reagents were purchased from Sigma-Aldrich (St. Louis, MO)
Measurement of NO, ROS, and cytokine production
NO and ROS production was determined as previously described (16) using the NO probe 4,5-diaminofluorescein diacetate or the ROS probe 2′,7′-dichlorodihydrofluorescein. The fluorescence level was recorded every 5 min over a period of 500 min using a Fusion αHT system (PerkinElmer Life Sciences) at an excitation wavelength of 488 nm and emission wavelength of 515 nm. NO and ROS levels are shown as fluorescence units (FU); their production rates are expressed as FU per minute. To measure cytokine production, cell-culture supernatants were collected every 24 h for 4 d and assayed for IL-6, IL-1α, IL-1β, TNF-α, IL-17, and RANKL using commercial ELISA kits (R&D Systems, Minneapolis, MN), following the manufacturer’s instruction.
In vitro assay for OC differentiation
Murine OCs were generated from RAW 264.7 cells or primary bone marrow cells (BMCs) isolated from femurs and tibias as previously described (27, 28). Briefly, BMCs were cultured in 48-well plates (2 × 105/well) in anti-MEM medium supplemented with 10% FBS, 100 U/ml penicillin, and 100 μg/ml streptomycin in the presence of 10 ng/ml M-CSF alone during the first 2 d, followed by 4 additional d in the presence of 10 ng/ml of M-CSF, plus 20 ng/ml RANKL. To differentiate OC from RAW 264.7, cells were cultured in the same way (2 × 104/well), except that RANKL was added at a concentration of 20 ng/ml, for 5 to 6 d. Human OCs were differentiated from PBMCs isolated from healthy blood donors as previously described (29). PBMCs were cultured for 7 d in 100 ng/ml M-CSF and 100 ng/ml RANKL-supplemented in 10% FCS DMEM. To quantify the number of OCs, cultures were fixed and stained for tartrate-resistant acid phosphatase (TRAP) activity using an acid phosphatase kit (Kamiya Biomedical Company, Seattle, WA) according to the manufacturer’s instructions. TRAP-positive multinucleated OCs (more than three nuclei) were counted using a tissue-culture inverted microscope (29).
In vitro bone resorption assays
Degradation of osteoblast-derived bone matrix was quantified as previously described (30), with some modifications. Briefly, 12,000 osteosarcoma cells (SaOS-2) per well were cultured in McCoy’s 5A medium supplemented with 15% FBS in 48-well polystyrene culture plates. When cultures reached 80–90% confluence, the medium was changed to osteoblast differentiation medium (anti-MEM; Life Technologies) containing 10% FBS, 2 mM glutamine, 300 mM ascorbic acid, and 10 mM β-glycerol phosphate. After 20–25 d, osteoblasts were removed using 15 mM NH4OH. Mouse BMCs (200,000 cells/well in 48-well plates) were plated on the matrix in an OC differentiation medium as above. At different time points, cells were removed using 15 mM NH4OH, and matrix was stained with Von Kossa dye. Photographs of individual wells were taken using a transmitted light microscope, and matrix abundance was quantified by Image J software (National Institutes of Health).
To determine ex vivo bone degradation, 5-mm–diameter bovine cortical bone disks were prepared and studied as described (31), with some modifications. Briefly, disks were washed and sonicated in distilled water and stored dry at room temperature. Before use, bone disks were sterilized by immersion in ethanol and placed under UV light for 30 min. Single disks were placed in individual wells with 0.5 ml anti-MEM plus 10 ng/ml M-CSF plus 20 ng/ml RANKL. Mouse BMCs, 400,000 cells/well, were incubated for 10 d with replenishment of fresh media every other day. At the end of incubation, bone disks were removed and stained for TRAP, and the number of OCs per disk was determined as above. Cells and debris were then removed by two bursts of 15-s sonication in concentrated ammonium hydroxide. Disks were stained with 1% toluidine blue for 30 s, and resorption pits were counted by scanning the entire surface of each disk with a reflected light microscope.
Differentiation of Th17 cells in vitro
Splenocytes (2 × 105/well) were cultured in 96-well plates in IMDM (Life Technologies, Carlsbad, CA) supplemented with 2 mM l-glutamine, 10% FBS, 1% penicillin-streptomycin, 10 mM HEPES buffer solution, and 50 mM 2-ME in the presence of Th17-polarizing mixture containing: anti–IL-4 (2 μg/ml), anti–IFN-γ (2 μg/ml), rhTGF-β (1 ng/ml), recombinant murine (rm)IL-6 (20 ng/ml), rmIL-23 (10 ng/ml), recombinant human IL-1β (10 ng/ml), anti-CD3 (2 μg/ml), and anti-CD28 (5 μg/ml). Cells were stimulated with 50 μg/ml SE-positive 65-79*0401, SE-negative 65-79*0402, or PBS and cultured for 5 d. After 5 d, cells were stimulated with PMA (5 ng/ml) and ionomycin (500 ng/ml) for the last 6 h of culture. Brefeldin A (10 μg/ml) was added to the culture for the last 5 h. Cells were then harvested and stained for surface marker using FITC anti-mouse CD4 and isotype controls followed by fixation and permeabilization using a Cytofix/Cytoperm kit (eBioscience). Intracellular staining was performed using APC-conjugated anti-mouse IL-17A mAb and PE-conjugated anti-mouse RANKL. Mean florescence intensity and percentages of stained cells were determined by FACS analysis.
In other experiments, lymph node or splenic cells isolated from CIA mice were restimulated ex vivo for 5 d in the presence or absence of denatured chicken CII (100 μg/ml) before Th17 abundance was determined by flow cytometry as above.
CIA induction and in vivo peptide administration
DBA/1 CII-TCR Tg mice (6–10 wk old) were immunized with chicken CII in CFA. In brief, 50 μl emulsion containing 100 μg CII in 25 μl 0.05 M acetic acid and 25 μl CFA was injected intradermally at the base of the tail. In addition, mice were injected twice per week i.p. with 100 μg either SE-positive peptide 65-79*0401 or SE-negative peptide 65-79*0402 in 50 μl PBS. Other groups were injected with 50 μl PBS alone. Mice were monitored, and paw swelling was determined as previously described (32) using a visual scoring system on a four-point scale for each paw: 0, no arthritis; 1, swelling and redness confined to digits; 2, minor swelling and redness spreading from the digits to the distal paw; and 3, major swelling and redness extending proximally from the paw.
Joint tissue studies
Limbs were dissected and decalcified in 10% EDTA for 14 d at 4°C. After decalcification, the specimens were processed for paraffin embedding and serial sectioned. The histological sections were deparaffinized, rehydrated, and stained with H&E or for TRAP activity using an acid phosphatase kit (Kamiya Biomedical Company, Seattle, WA). To determine OC abundance, TRAP-positive multinucleated cells were counted. Data, shown as OC count, represent mean ± SEM of the total number of OCs in front and rear paws plus knees.
Bone damage was evaluated by radiography and microcomputed tomography (micro-CT). Front and hind limbs from arthritic mice were dissected, fixed in 10% formalin, and stored in 70% ethanol. Limbs were scanned ex vivo by a micro-CT system (eXplore Locus SP; GE Healthcare Pre-Clinical Imaging, London, ON, Canada) in distilled water. The protocol included the source powered at 80 kV and 80 μA. In addition to a 0.508-mm Al filter, an acrylic beam flattener was used to reduce beam-hardening artifact (33). Exposure time was defined at 1600 ms/frame with 400 views taken at increments of 0.5°. With four frames averaged and binning at 2 × 2, the images were reconstructed with an 18-μm isotropic voxel size.
Regions of interest were defined through a spline-fitting algorithm to create separate masks for the carpals, tarsals, calcaneus, talus, or phalanges. The global threshold for cortical bone was defined at 2000 HU and 1200 HU for trabecular bone. Image analysis was run using MicroView 2.2 Gold (GE Healthcare Pre-Clinical Imaging). Hounsfield values were mapped using a color gradient to qualitatively analyze density patterns. Tissue mineral content, tissue mineral density, and bone volume fraction were quantified for each region as previously described (33).
Radiographs were taken using a microradiography system (Faxitron X-ray Corporation, Wheeling, IL) with the following operating settings: 27 peak voltage, 2.5 mA anode current, and an exposure time of 2.6 s. Coded radiographs were evaluated by an experienced rheumatologist who was blinded to the treatment. The presence of bone destruction was assessed separately for front and rear paws and scored on a scale of 0–5, ranging from no damage to complete destruction of the joints, as previously described (34).
Data are expressed as mean ± SEM from triplicate samples. All experiments were repeated at least three times. Unless otherwise stated, all statistical analyses were performed using a two-tailed Student t test (*p < 0.05, **p < 0.01).
The SE activates NO and ROS production and facilitates OC differentiation in pre-OCs
We have previously demonstrated that the SE ligand triggers NO and ROS production in several cell types (13–16). Given the known effect of NO and ROS on OCs (22–25), we examined whether the SE could activate NO and ROS signaling in the OC precursor cell line RAW 264.7. To this end, cells were treated with the SE ligand 65-79*0401, an SE-negative control peptide 65-79*0402, or PBS, and NO and ROS production was measured. As seen in Fig. 1A and 1B, the SE ligand 65-79*0401 activated markedly higher NO and ROS production compared with the control peptide or PBS. Activation of NO and ROS signaling in RAW 264.7 cells by the SE ligand, similar to its effect in fibroblasts (16), depended on cell-surface CRT (Fig. 1C, 1D).
Because NO and ROS possess pro-osteoclastogenic effects (22–25), we sought to determine whether the SE could affect OC differentiation. To this end, RAW 264.7 cells were cultured with different concentrations of the SE-positive ligand 65-79*0401, a control SE-negative peptide 65-79*0402, or PBS. OC abundance was determined by quantification of multinucleated, TRAP-positive cells. The SE ligand 65-79*0401 activated increasing OC differentiation over time (Fig. 1E), with an optimum concentration of 50 μg/ml (Fig. 1F). Under these conditions, the SE ligand could activate significantly higher differentiation rates even in the absence of exogenous RANKL; however, a more potent effect was seen in the presence of the recombinant protein (Fig. 1G, 1H). To determine whether the effect was specifically dependent on the SE sequence motif (QKRAA, QRRAA, or RRRAA), we tested a panel of SE-positive or -negative synthetic peptides. As can be seen in Table I, peptide-activated osteoclastogenesis was strictly dependent on the SE sequence motif.
The SE facilitates primary cell OC differentiation and bone degradation in vitro
To examine the effect of the SE ligand on primary cells, OC differentiation was induced in BMCs derived from DBA/1 mice as previously described (27, 28). Differentiation of OCs from BMCs was carried out in the presence of the SE ligand 65-79*0401, an SE-negative control peptide 65-79*0402, or PBS as above. After 6 d, cells were stained for TRAP, and OCs were counted. As seen in Fig. 2A, the SE significantly increased OC differentiation. The SE ligand facilitated OC differentiation in healthy human PBMCs as well. To date, 11 such blood samples were tested. In all but two cases, the SE-positive peptide 65-79*0401 produced strong pro-osteoclastogenic effects. A representative experiment is shown in Fig. 2B.
An endogenously expressed SE had a similar effect to that seen with an exogenously added synthetic ligand. Fresh BMCs isolated from naive DRB1*0401 Tg [Tg mice expressing human HLA-DR molecules coded by the SE-positive HLA-DRB1*04:01 allele (26)] showed higher constitutive abundance of TRAP-positive mononuclear (pre-OC) cells compared with BMCs from DRB1*0402 Tg, expressing an SE-negative HLA-DR molecule (Fig. 2C). Furthermore, after 6 d in OC-differentiating culture conditions, BMCs from DRB1*0401 Tg mice showed a significantly higher number of fully differentiated OCs (Fig. 2D). The osteoclastogenic propensity of DRB1*0401 Tg BMCs was accompanied by a higher capacity to degrade artificial bone matrix (Fig. 2E) and bovine bone disks (Fig. 2F) ex vivo. Thus, the SE ligand facilitates in vitro differentiation of OCs and enhances their bone-degrading functional activity.
The SE activates production of pro-osteoclastogenic factors
RA synovial fluids and tissues express high levels of inflammatory cytokines, such as IL-1, IL-6, IL-17, and TNF-α, which play important roles in bone destruction (20, 35). To determine the effect of the SE on pro-osteoclastogenic cytokine production, we cultured RAW 264.7 (Fig. 3A) or primary mouse BMCs (Fig. 3B) in OC-differentiating conditions in the presence of the SE ligand 65-79*0401, an SE-negative control peptide 65-79*0402, or PBS. Cytokine levels were measured in culture supernatants by ELISA. As seen in Fig. 3A and 3B, the SE ligand significantly augmented IL-6 and TNF-α production by both RAW 264.7 and BMCs. There was no increased production of IL-1α or IL-1β in either RAW 264.7 cells or BMCs. Additionally, the SE ligand did not increase IL-17 levels in BMC cultures or RANKL in RAW 264.7 cells.
Both IL-6 and TNF-α enhance osteoclastogenesis, and their levels are substantially elevated in the synovial fluid of RA patients (36, 37). We therefore examined whether these cytokines contribute to SE-activated osteoclastogenesis using neutralizing Abs. Both anti–IL-6 (Fig. 3C) and anti–TNF-α (Fig. 3D) neutralizing Abs inhibited SE-activated mouse bone marrow–derived OC generation. Thus, we conclude that IL-6 and TNF-α play a role in SE-activated osteoclastogenesis.
OC differentiation depends largely on an interaction between RANKL and its signaling receptor RANK (38). OC precursor cells can be sensitized to RANKL by increasing RANK expression, with resultant increased OC generation. We therefore undertook to determine whether the SE increases the expression of RANK on OC precursor cells. RAW 264.7 cells were treated with the SE ligand 65-79*0401, SE-negative peptide 65-79*0402, or PBS in the presence of low concentration (10 ng/ml) of recombinant RANKL. RANK expression was significantly higher on RAW 264.7 cell-differentiated OCs when they were cultured in the presence of the SE ligand (Fig. 3E).
Th17 cells possess a pro-osteoclastogenic effect, which is attributed partly to their own RANKL expression and partly to IL-17 production. We have previously demonstrated that the SE ligand enhances Th17 cell differentiation (17). Accordingly, experiments to determine whether the SE could activate Th17-dependent osteoclastogenesis were performed. SE-stimulated DBA/1 splenocytes demonstrated significantly higher abundance of RANKL-expressing Th17 cells compared with splenic cells stimulated with the control peptide or PBS (Fig. 3F). Furthermore, under suboptimal osteoclastogenic tissue-culture conditions, the SE ligand interacted with IL-17 (Fig. 3G). As can be seen, in limiting concentrations of RANKL (5 ng/ml) and IL-17 (0.1–1.0 ng/ml), the SE ligand and IL-17 had a synergistic pro-osteoclastogenic effect on mouse BMCs. Thus, our results suggest that in addition to its direct T cell–independent effect on OCs, the SE exerts a Th17 cell–mediated pro-osteoclastogenic effect by enhancing the differentiation of RANKL-expressing IL-17–producing T cells.
Arthritogenic effects of the SE in vivo
To determine the effect of the SE in vivo, we studied CIA. CII-immunized DBA/1 CII-TCR Tg mice were injected intraperitoneally with SE-positive peptide 65-79*0401, SE-negative control peptide 65-79*0402, or PBS. As can be seen in Fig. 4, SE-treated CIA mice showed higher Th17 abundance in regional lymph nodes compared with mice treated with the control peptide (Fig. 4A, 4B). Ex vivo lymph node cell expansion under Th17-differentiating conditions in the presence or absence of CII showed selective expansion of CII-specific Th17 cells in mice treated with the SE compared with mice treated with the control peptide (Fig. 4C, 4D).
Importantly, treatment with the SE ligand significantly facilitated disease onset (Fig. 5A) and increased joint swelling (Fig. 5B) compared with mice injected with the control peptide or PBS. The tissue swelling effect was transient, yet clearly significant, particularly during days 21–35 after immunization (p = 8.7 × 10−5, paired Student t test). Radiologic analysis showed that mice treated with the SE ligand had more severe erosive bone damage when compared with mice treated with the control peptide (Fig. 5C, 5D). Micro-CT–based bone mineral density imaging showed reduced bone mass in SE-treated mice, particularly in the ankles, with the most significant loss found in the calcaneus (Figs. 5E, 5F).
Histologic examination of arthritic joints revealed significantly higher abundance of OCs in SE-treated mice (Fig. 6A, 6B). Additionally, there was a higher abundance of TRAP-positive mononuclear (pre-OC) cells in the bone marrow of SE-treated mice (Fig. 6C). Upon ex vivo differentiation for 6 d, BMCs harvested from SE-treated CIA mice formed significantly higher numbers of OCs (Fig. 6D).
RA, an emblematic HLA-associated autoimmune disease, is characterized by extensive bone damage, which for unknown reasons is more severe in patients carrying SE-coding HLA-DRB1 alleles (9–11). This study identified the SE as a signal transduction ligand that can potently activate a bone-destructive pathway, both in vitro and in vivo. Our previous studies have demonstrated that the SE acts as a signal transduction ligand that interacts with cell-surface CRT and activates immune dysregulatory events both in vitro and in vivo (13–17). In this study, we demonstrate, for the first time to our knowledge, that the SE ligand has direct arthritogenic effects. These findings have several important implications for our understanding of the mechanisms governing autoimmune arthritis, as discussed below.
Our results demonstrated that similar to its effect in fibroblasts (16), SE signaling in OCs depended on interaction with its receptor, CRT. We have previously demonstrated that the SE ligand binds to a particular site on cell-surface CRT (39) in an allele-specific manner, activates NO and ROS signaling, and leads to distinct functional consequences, dependent on the cell type with which it interacts. For example, in CD8+ dendritic cells, the SE signal inhibits IDO activation, whereas in CD8− dendritic cells, it activates production of Th17-polarizing cytokines. The end result of this dual effect is immune dysregulation with Th17 polarization (17). In this study, we demonstrated that similar to fibroblasts, lymphocytes, and dendritic cells, the SE ligand interacted with cell-surface CRT on OCs and activated NO and ROS production. Different from other cell types, however, in OCs, these signaling events resulted in a distinct lineage-specific functional effect (osteoclastogenesis). Thus, consistent with our previous observations, cell-surface CRT is the SE signal-transducing receptor in many cell types. Although activation of the CRT-mediated pathway by the SE triggers lineage-invariant signaling events, the functional consequences are lineage specific.
Recent studies have implicated Th17 cells in the pathogenesis of RA and CIA and demonstrated that the signature cytokine of Th17 cells, IL-17, can activate OC differentiation and activity (19, 20, 40). Based on this prior knowledge and our own data (17), it would have been reasonable to hypothesize that the SE ligand may increase bone damage secondary to its effect on Th17 polarization. Unexpectedly, however, we discovered in this study that the SE had two distinct effects: enhancement of RANKL-expressing Th17 cell differentiation on the one hand and direct effect on OC differentiation in the absence of T cells (i.e., in RAW 264.7 cells) on the other. It is noteworthy that SE effect was enhanced in the presence of IL-17, suggesting that although the SE had two distinct effects, the two mechanisms operate synergistically. Importantly, the SE ligand stimulated osteoclastogenesis in human cell cultures as well, indicating the effect is species unrestricted.
SE-activated osteoclastogenesis was found to be mediated by previously identified pathways. For example, SE effect involved production of intracellular signaling molecules (NO and ROS) and cytokines (IL-6 and TNF-α) that have been previously found to participate in osteoclastogenesis. The unifying effector mechanism behind SE pro-osteoclastogenic effects is unknown, but IL-6 is a plausible candidate because it has been previously found to mediate SE-activated Th17 polarization (17) and, in the presence of soluble IL-6R, was shown to activate osteoclastogenesis (41). Thus, the data in Fig. 3 suggest that the seemingly unconnected SE-induced events may all be mediated by IL-6. Obviously, deciphering cause–effect relationships in a multilineage, multifactorial system such as SE-induced osteoclastogenesis requires further studies.
Accumulating evidence indicates that bone damage in RA affects both local and remote skeletal tissues (42). In this study, SE bone effects were seen in articular, peri-articular, and extra-articular bone sites. Radiological data confirmed that SE-activated OC accumulation in vivo was associated with greater degree of articular erosion. In addition, imaging studies demonstrated diffuse bone resorption in peri-articular (e.g., carpal bones) and extra-articular (e.g., calcaneus) tissues as well. In addition, the SE ligand potently activated in vitro osteoclastogenesis in BMCs collected from naive mice. Furthermore, freshly isolated BMCs from SE-treated CIA mice showed higher abundance of pre-OCs and higher susceptibility to osteoclastogenesis when cultured ex vivo in OC-differentiating conditions. When taken together, these findings indicate that SE ligand-activated bone-damaging effects in vivo extended beyond the joint compartment, resembling the bone damage distribution seen in RA.
Osteoclastogenesis, increased NO and ROS levels, and Th17 overabundance have all been implicated in many autoimmune conditions. Then how does the SE lead to the development of RA rather than other diseases? Our model (2, 13, 43, 44) proposes that RA disease onset depends on a constellation of events involving particular cell lineages and target tissues that are affected by SE signaling, as well as the relative potency of the signals and cross-talk with other pathways. Additionally, RA development depends on additional genes besides SE-coding HLA-DRB1 alleles. Finally, nongenetic influences (e.g., environmental factors) play important roles in RA development. Importantly, it has been previously shown that in RA, the SE and cigarette smoking interact synergistically to increase disease risk (45). Thus, many factors besides the presence of a SE are required to jointly trigger disease onset in RA.
The findings of this study lend important support to the SE Ligand hypothesis (2, 13, 43, 44). As mentioned above, the underlying mechanism of SE–RA association is unknown. Our in vitro and in vivo studies to date (13–17) have indirectly supported the SE Ligand hypothesis. However, direct evidence to conclusively implicate the SE as a signal transduction ligand in the pathogenesis of arthritis has been missing. By demonstrating the effect of the SE in CIA and identifying the pathophysiologic basis of that effect, this study helps to determine the functional role of the SE. Our data demonstrated that the SE ligand had potent pathogenically relevant effects at micromolar-range concentrations of a synthetic cell–free molecule that does not possess Ag-presentation properties. Consistent with this, we have recently developed a biostable peptidomimetic SE ligand that is ∼100,000 times more active on a molar basis than the SE ligand used in this study both in signaling (46) and osteoclastogenesis (J. Fu, S. Ling, Y. Liu, S. Naveh, C. Gilon, and J. Holoshitz, manuscript in preparation).
The newly identified pathogenic mechanism of SE may provide answers to several questions that Ag presentation cannot clearly explain. For example, RA concordance rate in monozygotic twins (47) and bone damage severity (9–11) both correlate directly with the number of SE-coding alleles. Such dose-dependent patterns are more consistent with signal transduction than Ag presentation. Further, in addition to its well-known effect on RA, the SE has been found to associate with bone erosions in non-RA conditions, such as psoriatic arthritis (48), systemic lupus erythematosus (49), and periodontal disease (50). Those conditions do not share a common pathogenesis or a putative target Ag. It is therefore conceivable that it is the SE ligand activity, rather than Ag presentation, that renders individuals carrying certain HLA-DRB1 alleles susceptible to excessive bone damage in a variety of clinical settings. Our finding that naive Tg mice carrying the SE-coding HLA-DRB1 allele have higher inherent propensity for osteoclastogenesis is consistent with this scenario.
In conclusion, we identified SE, a sequence motif best known for its association with severe RA, as a signaling ligand that facilitates OC-mediated bone damage. Beyond uncovering a functional effect of the single most significant genetic risk factor in RA, for which a mechanism of action has eluded the field for over two decades (6, 7), these findings introduce a novel paradigm that could provide a plausible mechanistic context to the enigmatic association of HLA alleles with a wide range of diseases (1).
The authors have no financial conflicts of interest.
We thank Dr. Chella David for providing HLA-DR4 Tg mice, Hahyung Kim for technical assistance, and Gail Quaderer for administrative support.
This work was supported by grants from the National Institutes of Health (GM088560, AR059085, AR056786, and AR55170 to J.H.), an Innovative Basic Science Award from the American College of Rheumatology, and a Johnson & Johnson Diversity Support grant.
Abbreviations used in this article:
- bone marrow cell
- collagen-induced arthritis
- collagen type II
- fluorescence unit
- microcomputed tomography
- rheumatoid arthritis
- receptor activator for NF-κB
- receptor activator for NF-κB ligand
- reactive oxygen species
- shared epitope
- tartrate-resistant acid phosphatase.
- Received August 2, 2012.
- Accepted October 22, 2012.
- Copyright © 2012 by The American Association of Immunologists, Inc.