Haptoglobin Acts as a TLR4 Ligand to Suppress Osteoclastogenesis via the TLR4–IFN-β Axis

Haptoglobin (Hp) belongs to a class of acute-phase proteins that are rapidly upregulated upon stimulation by cytokines such as TNF-α, IL-6, and IL-1 (1). Hp is expressed in most tissues and is mainly produced by stromal cells and hepatocytes. The primary function of Hp is to combine with the free hemoglobin (Hb) that is released during hemolysis or healthy RBC turnover, which leads to the elimination of the Hb–Hp complex by macrophages. Free Hb reacts with H₂O₂ to form reactive oxygen species, such as reactive hydroxyl radicals, which cause damage to tissues. Hp prevents oxidative stress by scavenging free Hb (2, 3). In addition to a systemic anti-inflammatory function, recent studies have shown that Hp is involved in the proliferation, function, and cytokine secretion of immune cells. The binding of Hp to CD11b/CD18, which is expressed in various immune cells, including dendritic cells (DCs), monocytes, and macrophages, suppresses the effector functions of macrophages and DCs, such as Ag presentation and phagocytosis and, at the same time, reduces the secretion of proinflammatory cytokines, such as IL-6 (4–6). In the case of T cells, Hp directly binds to activated or resting CD4⁺ and CD8⁺ T cells, thereby inhibiting proliferation (7). These results imply that Hp does not only suppress the inflammatory response but physiologically plays an important role in the biological processes of immune cells.

An osteoclast is a multinuclear giant cell that is formed by the fusion of mononuclear precursor cells of a macrophage-monocyte lineage. Osteoclasts have a unique ability to resorb the bone directly. Two factors are essential for the differentiation from progenitor cells to osteoclasts: M-CSF and the receptor activator of NF-κB ligand (RANKL) (8, 9). These factors induce the expression and activation of c-Fos, a key regulator of osteoclast differentiation. c-Fos raises the level of RANK, the receptor of RANKL, and increases the expression of NF of activated T cells 1 (NFATc1), which is another key transcription factor for osteoclastogenesis (10–12). Hematopoietic progenitor cells derived from c-Fos–deficient mice did not have a significant problem differentiating into macrophages but could not differentiate into osteoclasts (13). These results indicate that c-Fos plays an essential role in the triggering of osteoclast differentiation.

Diseases such as lupus, rheumatoid arthritis (RA), cancer, and osteoarthritis have disease-specific features but commonly have a chronic inflammatory response (1, 14, 15). Persistent and excessive inflammatory responses increase proinflammatory cytokines, such as TNF-α, IL-1, and IL-6, to more than necessary levels, accelerating the recruitment of osteoclast precursor cells and osteoclastogenesis (16–19). Moreover, inflammatory responses increase RANKL expression in fibroblast-like cells and immune cells like T cells and DCs. Thus, in diseases that are accompanied by an inflammatory response, bone loss due to abnormal osteoclast differentiation occurs frequently (20–22). Therefore, various studies have been conducted on inflammation-related molecules to prevent bone loss. Nevertheless, no therapeutic agent has been developed to inhibit bone loss entirely, and there is a continuing demand for therapeutic targets. Previous studies suggest that Hp is involved in various physiological functions of immune cells, indicating that Hp may affect osteoclast differentiation. Results showing that Hp is significantly higher in patients with lupus and RA than in healthy individuals emphasize the importance of studying the effect of Hp on osteoclast differentiation (1, 3). In the current study, we investigated the effect of Hp deficiency on osteoclast differentiation that affects bone phenotype and examined the direct effect of Hp on osteoclastogenesis.

All animal experiments were performed in accordance with the Animal Care Committee of the Institute of Laboratory Animal Resources of Seoul National University (Seoul, Korea). C57BL/6.Hp-deficient (Hp^−/−) mice were a kind gift from Dr. Lee Ann Garrett-Sinha (23). Eight-week-old male wild-type (WT) or Hp-deficient mice (n = 8 per group) were euthanized, and the femurs were fixed in 4% (w/v) paraformaldehyde for microcomputed tomography (μCT) assessment. The femurs were analyzed by high-resolution μCT (SMX-90CT system; Shimadzu, Kyoto, Japan). Scanning images from μCT were reconstructed by the VGStudio MAX 1.2.1 program (Volume Graphics, Heidelberg, Germany). Each three-dimensional image was analyzed to measure trabecular bone volume, cortical bone volume, trabecular number, and trabecular separation by using TRI/3D-VIE (RATOC System Engineering, Kyoto, Japan).

The samples from animal experiments were decalcified with 12% (w/v) EDTA for 3 wk and embedded in paraffin. After histological sagittal sections (5-μm thickness), calvariae were stained with hematoxylin solution (Sigma-Aldrich, St. Louis, MO) or stained for tartrate-resistant acid phosphatase (TRAP) using a Leukocyte Acid Phosphatase Assay Kit (Sigma-Aldrich) according to manufacturer’s instructions. Femurs were stained with 0.5% methyl green solution (Sigma-Aldrich) and TRAP stain kit. The TRAP-stained cells were analyzed by using the OsteoMeasure XP program (version 1.01; OsteoMetrics, Decatur, GA).

Recombinant human M-CSF and RANKL were obtained from PeproTech (Rocky Hill, NJ). Plasma-derived human Hp was purchased from Sigma-Aldrich. To detect target protein expression, Abs against NFATc1, c-Fos, and TLR4 were purchased from Santa Cruz Biotechnology (Santa Cruz, CA). Abs against TLR4, FLAG, and β-actin were purchased from Sigma-Aldrich. Secondary Abs conjugated with HRP were obtained from Sigma-Aldrich. All other Abs were from Cell Signaling Technology (Beverly, MA). The specific neutralizing Ab against TLR4 was purchased from eBioscience (San Diego, CA). The specific neutralizing Ab against IFN-β was purchased from PBL Biomedical Laboratories (Piscataway, NJ). Normal mouse IgG was purchased from Santa Cruz Biotechnology.

C57BL/6.TLR2– (TLR2^−/−), C57BL/6.TLR4– (TLR4^−/−), and C57BL/6.TLR7 (TLR7^−/−)–deficient mice were a kind gift from Prof. Sung Joong Lee (24). Bone marrow cells were obtained by flushing the bone marrow of the femur and tibia of 6-wk-old male mice. The nonadherent bone marrow cells were further cultured with M-CSF (30 ng/ml) and α-MEM for 3 d to generate bone marrow–derived macrophages (BMMs) as described previously (25). To generate osteoclastogenesis, BMMs (4 × 10⁴ cells/well) were cultured in 48-well plates with complete medium and α-MEM containing 10% (v/v) heat-inactivated FBS and 50 U/ml of penicillin in the presence of M-CSF (30 ng/ml) and RANKL (100 ng/ml) for 4 d. The medium was changed every 3 d. Multinucleated cells were observed on day 4. The cells were fixed in 10% formalin solution for 20 min and permeabilized with 0.1% Triton X-100 for 1 min. After washing twice with PBS, the cells were stained by using a Leukocyte Acid Phosphatase Assay Kit from Sigma-Aldrich following the manufacturer’s instructions. Calvarial osteoblasts were isolated from the calvariae of newborn mice as described previously (26). To generate osteogenic differentiation, osteoblasts were cultured in 48-well tissue culture plates precoated with collagen at a density of 5 × 10⁴ cells/well. After culturing for 24 h, the cells were further cultured in osteogenic medium: α-MEM complete medium containing 10 mM β-glycerophosphate (Sigma-Aldrich), 50 μg/ml ascorbate-2-phosphate (Sigma-Aldrich), and 100 ng/ml of BMP-2. The medium was replaced every 3 d. After culturing, the cells were fixed with 10% of formalin for 20 min. Osteoblast differentiation was measured by alkaline phosphatase or Alizarin Red S staining following manufacturer’s instructions (Sigma-Aldrich).

To analyze the secreted Hp, osteocalcin (OCN), procollagen type 1 N-terminal propeptide (P1NP) and IFN-β levels, mouse serum, or culture supernatant medium were collected and assessed using IFN-β ELISA kit (PBL Assay Science), OCN ELISA kit (LSBio, Seattle), P1NP ELISA kit (IDS, Boldon, U.K.), or a Hp ELISA kit (Abnova, Taipei City, Taiwan) according to the manufacturer’s instructions.

FITC labeling of Hp was performed by using a FluoroTag FITC Conjugation Kit (Sigma-Aldrich) according to the manufacturer’s instruction. BMMs were cultured in 12-well plates in the presence of M-CSF (30 ng/ml). After 24 h, Hp-FITC was added to the cells and incubated at 37°C for the indicated time. The cells were washed two times with PBS and fixed with 10% formalin for 30 min. After blocking the cells with 1% (w/v) BSA in PBS for 30 min, the Ab against TLR4 (2 μg) in 1% BSA in PBS was added. After 1 h incubation, the cells were washed with PBS for three times on a shaker. Then, anti-rabbit IgG Ab conjugated to Cy3 (Invitrogen) in 1% BSA in PBS was added to cells and incubated for 30 min in the dark room. The immunostained cells were observed under a confocal laser microscope (FV-300; Olympus, Tokyo, Japan).

BMMs (2 × 10⁶ cells per dish) were cultured with M-CSF (30 ng/ml) in a 100-mm diameter cell-culture dish. After 24 h incubation, the cells were treated with PBS, Hp (20 μg/ml), or LPS (1 μg/ml) and incubated for the indicated times. After incubation, the cells were detached by enzyme-free cell dissociation buffer (MilliporeSigma, Billerica, MA). The cells were stained with anti-TLR4 Abs, followed by FITC-conjugated anti-rat IgG, and analyzed by flow cytometry using a FACSCalibur (BD Biosciences, San Diego, CA). Cells stained with isotype control Abs were used as a negative control.

The cells (1 × 10⁴) were cultured on 96-well culture plates. After a 12 h incubation, the medium was exchanged with vehicle or the indicated doses of Hp. After the indicated time of incubation, 10 μl of EZ-Cytox solution (Daeil Lab Services, Seoul, Korea) was added to each well of the plate. After incubation for 2 h, the absorbance was measured with Multiskan (Thermo LabSystems, Philadelphia, PA) at 450 nm.

To detect specific protein expression, the cells were lysed on ice with a buffer containing 20 mM Tris-HCl, 150 mM NaCl, 1% (w/v) Triton X-100, a protease inhibitor, and phosphatase inhibitor (Sigma-Aldrich). After 30 min lysis on ice, the cellular debris was removed by centrifugation at 14,000 rpm × g for 15 min. The supernatant was collected in the tube, and the protein concentration was determined by the use of a DC Protein Assay Kit (Bio-Rad Laboratories, Hercules, CA). The protein samples (20–50 μg) were separated by SDS-PAGE and transferred onto a nitrocellulose membrane (GE Healthcare, Chalfont St. Giles, U.K.). The membrane was blocked with 5% (w/v) skim milk, and target proteins were incubated with the indicated primary Abs overnight. Blots were then incubated with secondary Abs conjugated with HRP and visualized by chemiluminescence reaction using ECL reagents (GE Healthcare).

Gene transduction was performed by using Platinum-E retroviral packaging cells (Cell Biolabs, San Diego, CA) according to the manufacturer’s instruction. Retroviral vectors pMX-c-Fos-FLAG were kindly provided by Prof. Nacksung Kim (27). The retroviral vectors were transiently transfected into Platinum-E cells by using Genjet transfection reagent (SignaGen Laboratories, Gaithersburg, MD). After incubation for 48 h, supernatants containing retrovirus were collected. BMMs were then transduced with the retroviral supernatants in the presence of polybrene (6 μg/ml; Sigma-Aldrich) and M-CSF (30 ng/ml) for 12 h. The cells were further cultured with puromycin (2 μg/ml) and M-CSF (30 ng/ml) for 3 d to remove uninfected cells.

Collagen sponges that were soaked in the vehicle or Hp (5 mg/kg), in combination with the vehicle or RANKL (1 mg/kg), were implanted onto the calvariae of 8-wk-old mice s.c. Each mixture was injected into the implanted collagen sponge daily for 5 d. The mice were sacrificed 6 d after implantation. The calvariae were collected, fixed in 4% (w/v) paraformaldehyde, and histological analysis was performed as described above.

To measure relative mRNA expression level, total RNA was isolated using TRIzol reagent (Invitrogen). Total RNA (2 μg) was used for first-strand cDNA synthesis using the SuperScript II Preamplification System (Invitrogen). To assess mRNA expression, 20–30 cycles of PCR amplification were performed for each gene. PCR products were separated on a 1–1.2% (v/v) agarose gel, stained with ethidium bromide, and detected by UV light. The relative mRNA expressions were evaluated by ABI Prism 7500 sequence detection system with SYBR Green PCR Master Mix (Applied Biosystems, Foster City, CA). Target mRNA expressions were determined according to the 2-ΔΔ cycle threshold method using HPRT as a reference gene.

To identify a direct interaction between Hp and TLR4, an Hp–TLR4 binding assay was performed as described previously (28). Briefly, high-binding immunoassay plates (Sigma-Aldrich) were coated with BSA, Hp, or LPS for 24 h at 4°C and then blocked in 2% BSA in PBS for 1 h. After three washes with PBS containing 0.05% (v/v) Tween 20, TLR4-Fc fusion protein (2 μg) was added and incubated for 2 h at room temperature. The captured TLR4-Fc was detected by using an HRP-conjugated goat anti-human IgG Ab (Sigma-Aldrich). o-Phenylenediamine dihydrochloride (Sigma-Aldrich) was added for spectrophotometric measurement at 450 nm by using Multiskan (Thermo LabSystems).

All quantitative experiments were performed at least in triplicate. Statistical significance was analyzed by Student t test or one-way ANOVA followed by Tukey–Kramer multiple comparisons test. Values are presented as the mean ± SD. The p values <0.05 were considered significant compared with their respective controls.

To investigate the effect of Hp on bone phenotype, we analyzed the femurs of WT mice and Hp-deficient mice. Computerized tomography analysis of the femurs revealed an osteoporotic phenotype in Hp-deficient mice (Fig. 1A). The bone volume per tissue volume in trabecular bone, trabecular number, and trabecular thickness decreased significantly in Hp-deficient mice compared with WT, and the bone surface per bone volume and trabecular separation increased. Furthermore, the trabecular bone pattern factor (an index of intertrabecular connectivity) was significantly increased in Hp-deficient mice. All these indices indicate a loss of trabecular bone in Hp-deficient mice (Fig. 1B). In contrast, there were no significant differences of cortical bone indices, such as cortical thickness, cortical bone area (Ct.Ar), and cortical area fraction (Ct.Ar/total cross-sectional area (Tt.Ar)) (Fig. 1C).

Next, we performed a TRAP stain to investigate whether the osteoporotic phenotype of Hp-deficient mice was associated with osteoclastogenesis. In comparison with control mice, the number of osteoclasts per bone surface (N.Oc./BS) and osteoclast surface per bone surface (Oc.S/BS) were significantly increased. In contrast, there were no significant differences in the number of osteoblasts per bone surface and osteoblast surface per bone surface (Fig. 1D). Consistently, serum levels of OCN and P1NP, which are bone anabolism markers, showed no difference between WT mice and Hp-deficient mice (Fig. 1E).

To further clarify the inhibitory effect of Hp on osteoclastogenesis, we examined whether Hp could impede osteoclastogenesis in the calvarial RANKL-induced osteoclast formation model. The number of osteoclasts increased by RANKL was decreased by more than 60% in the group treated with Hp compared with the control group (Fig. 2A). To investigate the possibility that osteoblasts affected by Hp are involved in the osteoporotic phenotype of Hp-deficient mice, we examined the effect of Hp on osteoblast differentiation. The osteoblast differentiation was carried out using osteoblast differentiation medium treated with Hp. Experimental results showed no significant difference between the experimental group and the control group in both the alkaline phosphatase stain and the Alizarin red stain (Supplemental Fig. 1A). The mRNA level of BSP and Runx2, which are markers of osteoblast differentiation, did not decrease compared with controls treated with the vehicle. Unexpectedly, the other osteoblastic differentiation markers, Alp and Col1a1, rather marginally increased by Hp treatment compared with vehicle treatment (Supplemental Fig. 1B). Next, to examine whether Hp directly inhibits osteoclastogenesis, we investigated the effect of Hp on osteoclast differentiation by treating BMMs with different concentrations of Hp. Osteoclast differentiation was inhibited by 10 μg/ml of Hp, and osteoclastogenesis was almost blocked by 20 μg/ml of Hp (Fig. 2B). An MTT assay was performed to exclude the effect of Hp on the proliferation of BMMs. Treatment with 50 μg/ml, which is higher than the concentration used in the experiment, did not affect the proliferation of BMMs (Supplemental Fig. 1C). These results suggest that the reduction of bone mass in Hp-deficient mice is closely related to osteoclastogenesis rather than to osteoblast differentiation.

To further investigate the inhibitory effect of Hp on osteoclastogenesis, we examined the effects of Hp on the differentiation phases of osteoclasts. First, we classified the process of osteoclast differentiation as early, middle, and late phases, which correspond to 0–1, 1–2, and 2–3 d after RANKL treatment. In both cases, either treating the osteoclasts with Hp at each phage and then maintaining treatment until mature osteoclasts were generated or treating the osteoclasts with Hp only at each differentiation phage, the inhibitory effect of Hp on osteoclastogenesis was observed only when osteoclasts were treated with Hp in the early differentiation phage (Fig. 2C, 2D). These results suggest that Hp may affect the initial triggering mechanism induced by RANKL.

Based on the finding that Hp inhibits osteoclastogenesis by affecting the early phase of osteoclast differentiation, we investigated whether Hp changes mRNA and protein levels of c-Fos and NFATc1, which are known to be key regulatory factors in the early phase of osteoclast differentiation. Treatment with Hp did not alter c-Fos mRNA levels but decreased c-Fos protein levels and reduced both the mRNA and protein levels of NFATc1 (Fig. 3A, 3B). Because both c-Fos and NFATc1 are essential transcription factors for osteoclast differentiation, the decrease of c-Fos and NFATc1 may inhibit osteoclastogenesis by decreasing the expression of RANK, the RANKL receptor. Thus, we examined whether Hp changed RANK mRNA levels. The mRNA level of RANK was not significantly altered by treatment with Hp when compared with the vehicle (Supplemental Fig. 2A).

It is well known that c-Fos binds to the promoter region of NFATc1, regulating the transcription of NFATc1 (29). Therefore, we hypothesized that Hp reduced the protein level of c-Fos, thereby suppressing the expression of NFATc1. To confirm this, we overexpressed the c-Fos in BMMs and differentiated osteoclasts with or without Hp. The level of NFATc1 reduced by Hp was restored in BMMs transduced with the c-Fos retroviruses, and the inhibitory effect of Hp on osteoclastogenesis also disappeared (Fig. 3C, 3D).

To further investigate mechanisms, we examined whether Hp affects MAPK signaling. Experimental results showed that the ERK pathway and JNK pathway were not affected by Hp, but the NF-κB pathway was affected. The pretreated Hp increased the basal level of p-p65 but suppressed NF-κB activation induced by RANKL stimulation (Supplemental Fig. 2C).

The inhibition of RANKL-induced c-Fos protein upregulated by IFN-β, which is independent of the mRNA expression level of c-Fos, was reported by Takanayagi group (30). Therefore, we hypothesized that Hp increased the expression of IFN-β, and the increased IFN-β acted autocrine, inhibiting osteoclastogenesis. To investigate this possibility, we examined the expression of IFN-β in BMMs after treatment with Hp. Treatment with Hp increased the mRNA level of IFN-β, and the amount of secreted IFN-β from BMMs was increased in proportion to the concentration of Hp (Fig. 4A, 4B). In addition, Hp promoted the phosphorylation of STAT 1 and 2, which are known to be downstream regulators of IFN-β signaling (Supplemental Fig. 3A). These results indicate that the autocrine loop of IFN-β was formed by Hp.

To investigate whether the inhibitory effect of Hp was associated with IFN-β, we examined the effect of Hp on osteoclastogenesis when IFN-β was neutralized with IFN-β Ab. The IFN-β Ab significantly restored osteoclastogenesis suppressed by Hp (Fig. 4C). In addition, the blockade of IFN-β restored the protein level of c-Fos, which was reduced by Hp (Fig. 4D). However, despite the complete restoration of the c-Fos protein level, the inhibition of osteoclastogenesis had not entirely disappeared. To exclude the possibility that Hp inhibited osteoclastogenesis through other mechanisms, the inhibitory effect of Hp on osteoclastogenesis was examined using BMMs from IFNAR1-deficient mice (Fig. 4E, Supplemental Fig. 3C). The inhibitory effect of Hp was rescued entirely in IFNAR1-deficient BMMs, and no effect of Hp on osteoclastogenesis was observed (Fig. 4F). Also, the phosphorylation of STAT1 and 2 was wholly suppressed (Supplemental Fig. 3B). These results suggest that IFN-β increased by Hp acts autocrine, inhibiting osteoclast differentiation by reducing the c-Fos protein level.

In previous studies, TLR signaling was shown to regulate IFN-β expression (31). Among TLRs expressed in BMMs, we have focused on TLR2, 4, and 7, which are important in regulating type 1 IFN signaling and have a high rate of expression in BMMs. First, we investigated whether they act as a receptor for Hp. When BMMs derived from TLR2- and TLR7-deficient mice were treated with a combination of RANKL and Hp, the inhibitory effect of Hp on osteoclastogenesis was maintained. However, the osteoclastogenesis inhibitory effect of Hp was not observed in BMMs derived from TLR4-deficient mice (Fig. 5A, Supplemental Fig. 4A). In line with the result of Fig. 5A, Hp treatment to BMMs derived from TLR2- and TLR7-deficient mice increased IFN-β mRNA levels without a significant difference when compared with WT mice. In addition, osteoclast differentiation was restored by blocking IFN-β (Fig. 5B, Supplemental Fig. 4B). However, in the BMMs derived from TLR4-deficient mice, the expression of IFN-β was not promoted with Hp treatment (Fig. 5B). Consistently, the deletion of TLR4 restored the protein levels of c-Fos and NFATc1, which were reduced by Hp (Fig. 5C). Moreover, it was observed that the inhibitory effect of Hp on osteoclastogenesis entirely disappeared by blocking the TLR4 receptor with TLR4 Ab (Supplemental Fig. 4C). These results suggest that Hp promoted IFN-β expression through TLR4.

It is known that IFN regulatory factor (IRF) 3 and 7 are involved in the regulation of IFN-β (32). Therefore, we investigated whether Hp affects the phosphorylation of IRF3 and 7. The phospho-IRF3 was not detected by stimulation (data not shown). In contrast, the phosphorylated form of IRF7 was significantly increased within 12 h after Hp stimulation. However, when BMMs derived from TLR4-deficient mice were stimulated by Hp, the phosphorylated form of IRF7 was not significantly increased (Fig. 5D). In previous studies, the MyD88-dependent pathway is known to be a common pathway activated by TLR4 signaling. Therefore, there is a possibility that TLR4 signaling activated by Hp regulates the osteoclast differentiation through the MyD88-dependent pathway. However, the inhibitory effect of Hp on osteoclastogenesis was maintained in BMMs derived from MyD88-deficient mice (Supplemental Fig. 4D).

We showed that TLR4 is involved in the IFN-β expression induced by Hp, but there is no report that Hp works as a ligand for TLR4. To investigate whether Hp binds directly to TLR4, we performed a binding assay. LPS increased the absorbance, which indicates the attachment of the TLR4 recombinant protein, depending on the plate-coating dose. A similar pattern occurred for Hp (Fig. 6A). According to previous reports, TLR4 binds with ligands to induce endocytosis, resulting in a TLR4-ligand complex entering the cell. To examine the endocytosis induced by Hp, the amount of the Hp–TLR4 complex introduced to cells was detected by confocal microscopy (33). It was observed that the colocalized regions of FITC-labeled Hp and Cy3-labeled TLR4 gradually increased in the cells depending on the length of the Hp treatment time (Fig. 6B). To investigate whether colocalized regions were increased by endocytosis of the TLR4–Hp complex, we counted the cells expressing TLR4 on the cell surface using FACS. It was observed that TLR4-positive cells were decreased in a time-dependent manner after treatment with Hp and were finally decreased by 80% (Fig. 6C). These results indicated that Hp binds directly to TLR4 and acts as a ligand.

To investigate whether Hp inhibits osteoclastogenesis through TLR4 in vivo, we examined osteoclast differentiation by applying calvarial RANKL-induced osteoclast formation model to WT and TLR4-deficient mice. The osteoclast formation by RANKL was observed in the calvariae in both WT and TLR4-deficient mice by TRAP staining. However, in the case of treatment with both RANKL and Hp, the osteoclast formation was suppressed to the vehicle-treated level in the WT mice. In the TLR4-deficient mice, the osteoclast formation was reduced by Hp in RANKL-treated mice, but the extent of reduction was much lower in TLR4^−/− mice than in WT mice (Fig. 7A). In addition, osteoclast formation parameters such as Oc.S/BS and N.Oc./BS indicated that the inhibitory effect of Hp on osteoclastogenesis was decreased in TLR4-deficient mice (Fig. 7B).

Previous studies have shown that the binding of Hp to CD11b/CD18 in macrophages, which are one of the osteoclast precursors, activates the NF-κB pathway (4, 34). Therefore, our initial expectation was that Hp promotes osteoclastogenesis. However, unlike this expectation, the osteoporotic phenotype was observed in the femur of the Hp-deficient mice and Hp showed a significant inhibitory effect on osteoclast differentiation in proportion to the treatment concentration (Figs. 1A, 2B). The activation of the NF-κB pathway induced by RANKL was suppressed by Hp, and the protein levels of c-Fos and NFATc1 were also decreased (Fig. 3B, Supplemental Fig. 2C). In addition, we observed that the osteoclast inhibitory effect of Hp was maintained in CD11b knock down BMMs (data not shown). These results indicate that Hp inhibited osteoclast differentiation in a way independent on its binding to CD11b/CD18. Our study rather showed that TLR4 is the important Hp receptor in mediating the antiosteoclastogenic effect of Hp. The lack of inhibitory effects of Hp on TLR4^−/− cells in vitro (Fig. 5C) and TLR4^−/− calvariae in vivo (Fig. 7A) clearly demonstrate that Hp inhibits osteoclasts via TLR4. To our knowledge, we are the first to report that TLR4 acts as a receptor for Hp, and Hp inhibits osteoclast differentiation through the TLR4–IFN-β axis (Fig. 7C).

In this study, when BMMs were stimulated by Hp, phosphorylation of IRF7 was significantly increased, whereas the phosphorylation of IRF3 was undetected. The lack of IRF3 phosphorylation, which has been shown to be necessary for the expression of IFN-β, was unexpected. However, these previous results were obtained by using LPS as a TLR4 signaling stimulator (35). Therefore, in the current study, using Hp as a TLR4 signaling stimulator, there is a possibility that IRF7 regulates IFN-β expression independently of IRF3. Besides, unlike the previous studies, which reported that the expression level of IRF7 was very low in most cells and induced by type 1 IFN–mediated signaling, the basal protein level of IRF7 in BMMs was fairly high and did not alter even after starvation for 6 h in our study (data not shown). These results suggest that TLR4-mediated signaling induced by Hp in BMMs may be different from previous studies. Although the regulatory mechanism involved in IFN-β induction by Hp has not been clearly elucidated, our results suggest that IRF7 activation is crucial for the Hp-stimulated IFN-β induction.

Increased proinflammatory cytokines in response to TLR4 signaling have been shown to increase RANKL expression in osteoblast-like stromal cells and to activate immune cells, thereby promoting bone loss due to indirect stimulation of osteoclastogenesis (16, 36–38). However, in the current study, we observed that Hp reduced osteoclastogenesis in a calvarial RANKL-injection model. It was also observed that trabecular bones of the femurs were decreased in Hp-deficient mice compared with WT mice. Therefore, our results were unexpected when considering that the activation of TLR4 by Hp promotes TNF-α expression and NF-κB activity in a similar manner to LPS (Supplemental Fig. 2B, 2C). These unexpected results may be not only because of the inhibitory effect of Hp on osteoclast differentiation observed in this study but also the inflammatory response alleviatory function through the regulation of immune cells or the elimination of reactive oxygen species. In previous studies, Hp reduces T cell proliferation, which is considered to be one of the major source cells of RANKL (7). It is also known that binding to CD11b/CD18-expressing cells, such as macrophages and DCs, suppresses Ag-presenting function and IL-6 expression, thereby alleviating inflammatory conditions (5, 6). These results suggest that even though TNF-α induced by Hp causes an inflammatory response in the body, it can be compensated and masked by the anti-inflammatory effect of Hp. This notion may explain the reason for our observation that the inhibitory effect of Hp on osteoclastogenesis in TLR4-deficient mice did not completely disappear (Fig. 7A, 7B). Hp may function directly via other TLR4 and also indirectly via other receptors in suppressing osteoclastogenesis. However, because the receptors of Hp have not been completely identified and the effects of Hp on other cell types such as stromal cells have not been studied, further studies are needed to clarify the physiological function of Hp.

Currently, therapeutics such as bisphosphonates and denosumab that suppress osteoclast differentiation and function are widely used for the treatment of osteoporosis and have shown significant positive results in patients (39). The majority of patients receiving these therapies also suffer from persistent inflammatory diseases such as RA or cancer, which often involves pathologic osteoclastogenesis (9, 15, 40). Therefore, they receive additional treatment for the suppression of inflammation. As the use of multiple medicines increases the potential to cause unwanted side effects, there is a continuing demand for effective and safe therapeutic agents that cure both bone loss and inflammation for inflammatory bone diseases. In addition to anti-inflammatory effects of Hp previously reported, our results that Hp-induced IFN-β expression, inhibiting osteoclastogenesis, suggest that Hp is a good candidate. In particular, IFN-β has an anti-tumor effect and has been studied for the application to cancer therapy (41–43). It has also been reported that IFN-β injection significantly alleviated paw swelling and severity of disease in animal models of RA (44, 45). In further studies, we will investigate the value of Hp as a therapeutic molecule for diseases such as cancer and RA, which cause chronic inflammation and bone loss.

We thank LeeAnn Sinha (Department of Biochemistry, State University of New York, Buffalo) for kindly providing Hp-deficient mice.

The authors have no financial conflicts of interest.