α-1 Antitrypsin Inhibits RANKL-induced Osteoclast Formation and Functions
- 15 Downloads
Osteoporosis is a global public health problem affecting more than 200 million people worldwide. We previously showed that treatment with α-1 antitrypsin (AAT), a multifunctional protein with antiinflammatory properties, mitigated bone loss in an ovariectomized mouse model. However, the underlying mechanisms of the protective effect of AAT on bone tissue are largely unknown. In this study, we investigated the effect of AAT on osteoclast formation and function in vitro. Our results showed that AAT dose-dependently inhibited the formation of receptor activator of nuclear factor κB ligand (RANKL)-induced osteoclasts derived from mouse bone marrow macrophage/monocyte (BMM) lineage cells and the RAW 264.7 murine macrophage cell line. To elucidate the possible mechanisms underlying this inhibition, we tested the effect of AAT on the gene expression of cell surface molecules, transcription factors and cytokines associated with osteoclast formation. We showed that AAT inhibited macrophage colony-stimulating factor (M-CSF)-induced cell surface RANK expression in osteoclast precursor cells. In addition, AAT inhibited RANKL-induced TNF-α production, cell surface CD9 expression and dendritic cell-specific transmembrane protein (DC-STAMP) gene expression. Importantly, AAT treatment significantly inhibited osteoclast-associated mineral resorption. Together, these results uncover new mechanisms for the protective effects of AAT and strongly support the notion that AAT has therapeutic potential for the treatment of osteoporosis.
Bone homeostasis is maintained by the mutual function of bone-resorbing hematopoietic lineage-derived osteoclasts (OCs) and mesenchymal stem cell-derived bone-forming osteoblasts (1). The balance between osteoclasts and osteoblasts is important for normal skeletal formation and function. Therefore, recruitment, proliferation and differentiation of these two types of cells are critical to maintain the normal physiology of bone (2). Osteoclast formation is a normal aspect of skeletal morphogenesis and remodeling; however, disproportionate osteoclast proliferation and activation can lead to excessive bone resorption. This can subsequently lead to chronic systemic bone diseases such as osteoporosis, which is a serious public health problem affecting an estimated 34 million Americans and causing 2 million fractures annually (3,4). Strategies to inhibit excessive osteoclast formation and/or function have proven to have therapeutic usefulness for the treatment of osteoporosis (1). However, the use of currently available drugs is limited due to their side effects, including osteonecrosis of the jaw, which can be caused by nitrogen-containing bisphosphonates, the most commonly used antiresorptive drugs, and denosumab, a monoclonal antibody inhibitor of RANKL (1,5).
OCs are large multinucleated cells. Differentiation of OCs is regulated by receptor activator of nuclear factor κ-B ligand (RANKL) and macrophage colony-stimulating factor (M-CSF) from osteoblasts and stromal cells in the bone marrow environment (6, 7, 8). RANKL plays a critical role in the development, survival and activity of OCs. M-CSF contributes to proliferation, survival, and differentiation of early precursors. Arai et al. (9) identified the early and late stages of osteoclast precursor (OCP) cells and demonstrated that in both the early and late stage of OCP cells, M-CSF stimulates the expression of RANK. RANK on the surface of OCP interacts with RANKL and recruits TNF receptor-associated factor (TRAF) family proteins such as TRAF6, which is an adapter molecule. These TRAF family proteins, especially TRAF6, activate NF-κB and MAP kinases (MAPKs). Activation of NF-κB and MAPKs eventually activates c-Fos, PU.1 and NFATc1, all of which are essential for osteoclast differentiation (10,11). Among these factors, NFATc1 is considered a master switch for regulating terminal differentiation of OCs (12). During OC differentiation, OCP cells fuse with one another to form multinuclear mature osteoclasts. This fusion requires expression of cell fusion-promoting proteins by OCP cells. Several cell fusion-promoting proteins have been identified, including dendritic cell-specific transmembrane protein (DC-STAMP), CD9 and Atp6v0d2, and they are also stimulated by RANKL-RANK signaling pathways (13,14). RANKL also plays an important role in OC activation (15). RANK-RANKL binding on mature OCs triggers internal structural changes and results in secretion of protons and lytic enzymes into a sealed extracellular resorption compartment. Cathepsin K (CatK) is an enzyme responsible for degradation of bone collagen matrices. Acidification of the resorption compartment is important for the activation of CatK. Secretion of protons by the vacuolar H+-ATPase leads to activation of CatK (16,17). Therefore, RANK-RANKL signaling is essential for osteoclast formation and activation (18), and the discovery of the RANK signaling pathway has provided insight into the mechanisms of osteoclastogenesis and activation of bone resorption (17). Inhibition of RANK expression in OCP cells could be one logical approach to inhibit excessive osteoclast formation and activation.
During inflammation, several proinflammatory cytokines are produced. These messenger molecules not only perpetuate inflammation but also, in turn, stimulate osteoclast formation and thus bone resorption, leading to osteoporosis and increased fracture rate (19,20). In these scenarios, different proinflammatory cytokines, including TNF-α, IL-1 and IL-6, have been shown to be capable of stimulating increased levels of RANKL/CSF-1-induced osteoclastogenesis (19). Studies have shown that TNF-α induced by RANKL promotes osteoclastogenesis in vitro by modulating RANK signaling pathways.
Human α-1 antitrypsin (AAT) is a protease inhibitor with cytoprotective and antiinflammatory properties. It inhibits lipopolysaccharide-induced secretion of TNF-α and IL-1β, and enhances the production of antiinflammatory IL-10 from human monocytes (21). In inflammation-related disease models, including type 1 diabetes and rheumatoid arthritis, AAT showed therapeutic potential (22, 23, 24, 25, 26). In addition, AAT inhibited the activity of NF-κB, which is important for the gene expression of proinflammatory cytokines (27). Recently, we showed that AAT protein and gene therapies reduced bone loss in an ovariectomized mouse model (28). We also showed that mesenchymal stem cells expressing AAT ameliorate bone loss in osteoporotic mice (29). The goal of this study was to test the effect of AAT on RANKL-induced osteoclast formation and function, and to elucidate the possible underlying mechanism of these effects.
Materials and Methods
Animals and Cells
Six-week-old C57BL/6 mice and TNF-α receptor (TNFR1 and TNFR2) deficient C57BL/6 mice were purchased from Jackson Laboratory (Bar Harbor, ME, USA) and housed in specific pathogen-free conditions under a 12 h light/dark cycle at the University of Florida animal care facility. All procedures were performed according to University of Florida Institutional Animal Care and Use Committee guidelines. Murine leukemic monocyte macrophage cell line RAW 264.7 cells were purchased from American Type Culture Collection (Manassas, VA, USA).
Reagents and Antibodies
Minimum essential medium, α modification (MEM-α) was purchased from Sigma-Aldrich (St. Louis, MO, USA). Fetal bovine serum (FBS), phosphate-buffered saline (PBS) and penicillin/streptomycin were purchased from Corning (Manassas, VA, USA). Recombinant murine RANKL and M-CSF were purchased from Peprotech (Rocky Hill, NJ, USA). For tartrate resistance acid phosphatase (TRAP) staining, a leukocyte acid phosphatase kit was purchased from Sigma-Aldrich. AAT (Prolastin C, Telecris Biotherapeutics, Research Triangle Park, NC, USA) was used. Antimouse CD265 (RANK) phycoerythrin (PE) conjugated antibody, anti-mouse CD9 fluorescein isothiocyanate (FITC) conjugated antibody and 7-amino-actinomycin D (7-AAD) viability staining solution were purchased from eBioscience (San Diego, CA, USA). Anti-DC-STAMP antibody clone 1A2 was purchased from EMD Millipore (Billerica, MA, USA). TNF-α, IL-1β and IL-10 enzyme-linked immunosorbent assay (ELISA) kits were purchased from Peprotech.
Murine osteoclasts were generated from BMM lineage cells as described previously (30). Briefly, femurs and tibiae were removed aseptically from 6- to 7-wk-old C57BL/6 male mice and dissected free of adhering tissues. The bone ends were cut off with scissors and the marrow cavities were flushed with 3 mL of MEM-α through one end of the bone using a sterile 27-gauge needle. The bone marrows were filtered with 70 µm nylon mesh filter (Fisher Scientific, Pittsburgh, PA, USA), centrifuged to collect the pellet and treated with 1–2 mL of NH4Cl solution (STEMCELL Technologies, Vancouver, BC, Canada) to lysis of red blood cells. The bone marrow cells were then washed once with MEM-α, suspended in MEM-α supplemented with 10% FBS and 1% penicillin/streptomycin, and cultured in 20 × 106 cells/100 mm diameter cell culture dish with M-CSF (100 ng/mL) in a humidified atmosphere of 5% CO2 for 16 h. During that time, BMMs and their precursors can survive as nonadherent cells (31), which are called early-stage OCP cells. Nonadherent cells were harvested and cultured for another 3 d in medium containing M-CSF (100 ng/mL). Then, floating cells were removed by pipetting, and attached cells, which we considered late-stage OCP cells, were collected by scraping. To generate osteoclasts, late-stage OCP cells were cultured with RANKL (100 ng/mL) and M-CSF (50 ng/mL) for an additional 3 d in 96-well cell culture plate (2 × 104 cells/0.25 mL/well) or 24-well plate (1 × 105 cells/0.5 mL/well). Since generating osteoclasts from BMM cells requires 7 d, we added different concentrations of AAT (0.5, 1 and 2 mg/mL) at different time points to investigate its effect on osteoclast formation and function. We named our studies Experiments 1–3. In Exp-1, AAT was added from d 0–7; in Exp-2, AAT was added from d 4–7; and in Exp-3, AAT was added from d 0–4. A procedure similar to that mentioned above was used to generate osteoclasts from TNF-α receptor (TNFR1 and TNFR2) deficient C57BL/6 mice, and in this case, AAT was added according to Exp-2 (d 4–7). To generate osteoclasts from the RAW 264.7 cell line, cells were cultured in MEM-α medium supplemented with 10% FBS and 1% penicillin/streptomycin with RANKL (100 ng/mL) in 96-well cell culture plate (8 × 103 cells/0.25 mL/well) or 24-well plate (30 × 103 cells/0.5 mL/well) with or without AAT in different concentrations (0.5, 1 and 2 mg/mL) for 6 d. Old media was replaced with fresh media containing RANKL (100 ng/mL) on d 3 (31).
Osteoclasts were generated as described above. To determine the TRAP+ osteoclasts, cells were washed with PBS, fixed with cold 4% paraformaldehyde and permeabilized with 0.5% Triton X-100. TRAP+ cells were detected using a leukocyte acid phosphatase kit following the manufacturer’s instructions. The positive cells for TRAP staining contain red granular material in cells. TRAP+ multinuclear cells containing ≥3 nuclei were considered multinuclear osteoclast cells (MNCs). The cells were examined under a microscope (Zeiss Axiovert 200 inverted fluorescence microscope using Axicam MRc5) and counted.
Resorption Pit Assay
For resorption pit assays, we performed two different experiments. First, we generated the osteoclasts using M-CSF and RANKL on a 96-well osteo assay surface plate (Corning, NY, USA) as described above and treated them with different concentrations of AAT (0.5, 1 and 2 mg/mL) from d 0–7. Cells were removed using 10% bleach and resorption pits were photographed with a microscope (Zeiss Axiovert 200 inverted fluorescence microscope using Axicam MRc5) and analyzed with Image J version 1.50b software. In another experiment, we first generated osteoclast cells on 96-well tissue culture plate using M-CSF and RANKL as described above. At d 6 of osteoclast induction, the cells were then plated on the 96-well osteo assay surface plate and allowed to settle for 2 h, then incubated with different concentrations of AAT (0.5, 1 and 2 mg/mL) for an additional 3 d. Cells were removed using 10% bleach and resorption pits were photographed and analyzed with Image J software.
Detection of Cytokines
Osteoclast cells were generated as described above. During osteoclast generation, the culture medium was collected and centrifuged at 1000 rpm for 5 min at 25°C to remove any dead cells. The concentrations of TNF-α, IL-1β and IL-10 were determined using murine ELISA development kits following the manufacturer’s instructions.
Flow Cytometry Analysis
Flow cytometry analysis was carried out with FACSCalibur CellQuest Pro version 5.2.1 (BD Biosciences, San Jose, CA, USA) and data were analyzed using FCS Express version 4 software (Denovo) at the University of Florida Flow Cytometry Core. Antibodies used in this study were PE-conjugated anti-mouse CD265 (RANK) antibody, FITC conjugated anti-mouse CD9 antibody and anti-mouse DC-STAMP antibody. For staining of intracellular DC-STAMP, cells were permeabilized and stained using the cytofix/cytoperm kit (BD Bioscience). Dead cells stained with 7-AAD viability staining solution were excluded from the analysis. A gate was set of living cells and mean fluorescence intensity (MFI) was compared with unstained cells.
Real-time Polymerase Chain Reaction
To quantify gene expression levels, total RNA was extracted from cultured cells with TRIzol reagent (Invitrogen, Carlsbad, CA, USA). Complementary DNA was synthesized from total RNA using reverse transcriptase (Qiagen, Hilden, Germany) and subjected to real-time polymerase chain reaction (PCR). Results were normalized to the gene expression levels of glyceral-dehyde-3-phosphate dehydrogenase (GAPDH) in the same sample. The fold-change ratios between test and control samples were calculated. The following primers were used: for NFATc1, 5′-CCG TCC AAG TCA GTT TCT ATG T-3′ (forward) and 5′-GTC CGT GGG TTC TGT CTT TAT-3′ (reverse); for cFos, 5′-GAA TCC GAA GGG AAC GGA ATA A-3′ (forward) and 5′-TCT CCG CTT GGA GTG TAT CT-3′ (reverse); for GAPDH, 5′-TGC ACC ACC AAC TGC TTA G-3′ (forward), and 5′- GGA TGC AGG GAT GAT GTT C-3′ (reverse); for NFκB, 5′-TACAAGCTGGCTGGTGGGGA-3′ (forward) and 5′-GTCGCGGGTCTCAGGACCTT-3′ (reverse); for RANK, 5′-CAC AGA CAA ATG CAA ACC TT G-3′ (forward) and 5′-GTG TTC TGGAAC CAT CTT CCT CC-3′ (reverse); for DC-STAMP, 5′-TCCTCCATGAACAAACAGTTCCAA-3′ (forward) and 5’ AGACGTGGTTTAGGAATGCAGCTC-3′ (reverse); and for cathepsin K, 5′TCAGAAGATGACGGGACTCA-3′ (Forward) and 5′-TCTTGAGTTGGCCCTCCA-3′ (reverse).
Determination of Cathepsin K Activity
Cathepsin K activity was determined by using a cathepsin K drug discovery kit (BML-AK430; Enzo Life Sciences, Farmingdale, NY, USA) according to the manufacturer’s protocol.
Data were analyzed using one-way analysis of variance with GraphPad Prism5 software, followed by Dunnett’s multiple comparison test. Student t test was used to compare two samples. The data are presented as mean ± standard error of the mean (SEM), and values of P <0.05 were considered statistically significant.
All supplementary materials are available online at https://doi.org/www.molmed.org.
AAT Inhibited RANKL-Induced Osteoclast Formation in a Dose-Dependent Manner
Effect of AAT on Early- and Late-stage OCP Cells
AAT Inhibited RANKL-Induced TNF-α Secretion During Late-stage Osteoclast Formation
A recent study has shown that AAT can significantly reduce the binding of TNF-α to TNF-α receptors (TNFR1 and TNFR2) (32). It has also been reported that RANKL induces TNF-α production (33), which stimulates RANKL-induced osteoclastogenesis by an autocrine mechanism in vitro. Based on this information, we tested whether inhibition of TNF-α played a critical role in AAT-mediated inhibition of osteoclast formation. We generated osteoclasts using BMM cells from a TNF-α receptor (TNFR1 and TNFR2) deficient mouse. Our results show that inhibition of osteoclast formation required a higher dose of AAT (2 mg/mL) (Figure 3E) and that AAT significantly decreased the TNF-α level (Figure 3F). These results suggest that other pathways may be involved in the inhibition of RANKL-induced osteoclast formation.
AAT Inhibited M-CSF-Induced RANK and Related Gene Expression in Early-stage and Late OCP Cells
AAT Inhibited RANKL-Induced CD9 Expression and DC-STAMP Gene Expression in OCP Cells
AAT Inhibited RANKL-Induced Bone Resorption by Osteoclasts
AAT Inhibited CatK Activity and RANKL-Induced CatK Gene Expression
In this study, we have shown for the first time that AAT efficiently inhibits RANKL-induced osteoclast formation and bone resorption. We have demonstrated that AAT reduces M-CSF-induced RANK receptor expression and downregulates M-CSF-induced regulatory gene expression (NF-κB and cFos). We have also shown that AAT inhibits RANKL-induced TNF-α production, CD9 expression and DC-STAMP gene expression, and Cat K gene expression and activity. Together, our results uncover novel mechanisms underlying the protective effect of AAT on bone loss and indicate that AAT has therapeutic potential for the treatment of osteoporosis. AAT is a Food and Drug Administration-approved drug and is generally considered safe for the treatment of α-1 antitrypsin deficiency disease (35,36). Considering that all currently used antiresorptive drugs for the treatment of osteoporosis have side effects (5,37), the safety profile of AAT could make it an appealing candidate for the treatment of osteoporosis.
We investigated the effect of AAT on osteoclast formation and found that AAT treatment (from d 0–7) inhibited osteoclast formation efficiently. Further analysis showed that late AAT treatment (d 4–7) inhibited osteoclast formation and RANKL-induced TNF-α secretion, while early AAT treatment (d 0–4) inhibited osteoclast formation by inhibiting M-CSF-induced RANK expression. We showed by MTT assay that this efficient inhibition was not due to cell apoptosis (data not shown). These findings clearly demonstrate that AAT inhibited osteoclast formation by multiple mechanisms.
TNF-α is produced by many types of cells, including monocytes/macrophages, osteoblasts and various cancer cells, and is involved in inflammatory tissue destruction, particularly bone resorption (5). RANKL induction of osteoclastogenesis is accompanied by a rapid and transient increase in TNF-α mRNA and TNF-α release in the precursor cell, which can act as an autocrine factor in osteoclastogenesis (33,38). In the present study, we showed that AAT reduced RANKL-induced TNF-α production. One possible mechanism is that AAT inhibits ADAM17, also known as a TNF-α-converting enzyme, which cleaves and releases soluble TNF-α (39). A recent study by Bergin et al. (32) showed that AAT can reduce the binding of TNF-α to its receptors (TNFR1 and TNFR2). Since TNF-α can self-regulate its gene expression (33), blocking the binding of TNF-α to its receptors by AAT may also contribute to inhibition of TNF-α production. We also tested the effect of AAT in cells without TNF-α receptors (TNFR1 and TNFR2) and showed that a higher dose of AAT effectively inhibited osteoclast formation, indicating that blocking TNF-α receptors is not the only mechanism for the function of AAT. In fact, AAT can enter the target cells and directly interact with cellular proteins (40).
As the RANK-RANKL interaction is indispensable for osteoclast formation (18) and M-CSF induces RANK expression in early-stage osteoclast precursors, we focused on AAT effects on RANK expression. We found a significant inhibitory effect of AAT on M-CSF-induced RANK expression in early-stage OCP cells. A recent study has shown that cFos, a transcription factor, is essential for upregulation of RANK expression in osteoclast precursor cells (34). Another study showed that M-CSF upregulated cFos expression in mature osteoclasts, at least in part via transcriptional activation of the fos gene (41). Therefore, M-CSF-induced cFos expression in OCP cells is believed to play a critical role in RANKL-induced osteoclastogenesis. In our study, we showed that AAT inhibited M-CSF-induced cFos mRNA expression. It is possible that downregulation of cFos gene expression by AAT leads to a reduction of RANK expression on the cell surface. It has also been reported that RANKL-induced expression of NFATc1, a master regulator of osteoclast differentiation, is tightly regulated by cFos (42). In the present study, we observed a reduction of NFATc1 gene expression. Together, these data suggest that AAT inhibits osteoclast differentiation by reducing cell surface RANK expression via downregulation of the cFos gene and NF-κB gene expression.
Since the RANKL-RANK interaction plays an important role in the expression of a set of cell fusion-related cell surface proteins, including CD9 and DC-STAMP, AAT-mediated reduction of RANK can consequently lead to a reduction of CD9 and DC-STAMP. Indeed, we showed that early AAT treatment significantly reduced CD9 cells (Figure 5A). Similarly, we showed that early AAT treatment (without RANKL) also inhibited cell surface DC-STAMP levels (Figure 5C). However, we observed no or a minor effect of AAT in the condition that RANKL presents (Figures 5B, D). It is possible that RANKL-induced cell surface DC-STAMP quickly internalized as the cells fused and underwent degradation, while our methods of detection were not able to show the complex dynamics. Nonetheless, our results show that AAT clearly inhibited RANKL-induced DC-STAMP gene expression (Figure 5E). These results support our hypothesis.
To test the effects of AAT on osteoclast function, we performed pit formation studies. We generated osteoclasts on osteo assay surface plates and found a significant reduction of pit formation with AAT treatment. It is possible that the inhibitory effect of AAT on osteoclast formation led to smaller areas of pit formation. To rule out this possibility, we first generated osteoclasts and then tested the effect of AAT on their mineral resorption. Our studies show a significant reduction of pit formation in the AAT-treated group. In addition, we show that AAT inhibited the activity and gene expression of CatK. These results clearly demonstrate that AAT has an inhibitory effect on bone resorption by mature osteoclasts.
In summary, our studies provide the following novel findings: (1) AAT efficiently inhibits osteoclast formation; (2) AAT reduces M-CSF-induced expression of regulatory genes (NF-κB and cFos); (3) AAT inhibits M-CSF-induced cell surface RANK receptor expression; (4) AAT inhibits RANKL-induced TNF-α production, CD9 expression and DC-STAMP gene expression; (5) AAT inhibits osteoclast-associated bone mineral resorption; and (6) AAT inhibits the enzymatic activity and gene expression of CatK. These findings provide novel mechanisms for the protective effect of AAT on bone and strongly support that AAT has therapeutic potential for the treatment of osteoporosis.
The authors declare they have no competing interests as defined by Molecular Medicine or other interests that might be perceived to influence the results and discussion reported in this paper.
EME is a visiting scholar from Zagazig University and is supported by a scholarship from the Egyptian government. We thank Dr. Jay Cao (US Department of Agriculture, Agriculture Research Service’s Grand Forks Human Nutrition Research Center) for his assistance with part of the gene expression studies and suggestions regarding induction of osteoclast formation.
This work was supported by a grant from the University of Florida.
Open Access This article is licensed under a Creative Commons Attribution-NonCommercial-NoDerivatives 4.0 International License, which permits any non-commercial use, sharing, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, and provide a link to the Creative Commons license. You do not have permission under this license to share adapted material derived from this article or parts of it.
The images or other third party material in this article are included in the article’s Creative Commons license, unless indicated otherwise in a credit line to the material. If material is not included in the article’s Creative Commons license and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder.
To view a copy of this license, visit (https://doi.org/creativecommons.org/licenses/by-nc-nd/4.0/)