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Keywords:

  • PARATHYROID HORMONE;
  • ONCOSTATIN M;
  • RANKL;
  • OSTEOCLAST;
  • GP130

Abstract

  1. Top of page
  2. Abstract
  3. Introduction
  4. Materials and Methods
  5. Results
  6. Discussion
  7. Disclosures
  8. Acknowledgements
  9. References

Parathyroid hormone (PTH) is the only approved anabolic agent for osteoporosis treatment. It acts via osteoblasts to stimulate both osteoclast formation and bone formation, with the balance between these two activities determined by the mode of administration. Oncostatin M (OSM), a gp130-dependent cytokine expressed by osteoblast lineage cells, has similar effects and similar gene targets in the osteoblast lineage. In this study, we investigated whether OSM might participate in anabolic effects of PTH. Microarray analysis and quantitative real-time polymerase chain reaction (qPCR) of PTH-treated murine stromal cells and primary calvarial osteoblasts identified significant regulation of gp130 and gp130-dependent coreceptors and ligands, including a significant increase in OSM receptor (OSMR) expression. To determine whether OSMR signaling is required for PTH anabolic action, 6-week-old male Osmr−/− mice and wild-type (WT) littermates were treated with hPTH(1–34) for 3 weeks. In WT mice, PTH increased trabecular bone volume and trabecular thickness. In contrast, the same treatment had a catabolic effect in Osmr−/− mice, reducing both trabecular bone volume and trabecular number. This was not explained by any alteration in the increased osteoblast formation and mineral apposition rate in response to PTH in Osmr−/− compared with WT mice. Rather, PTH treatment doubled osteoclast surface in Osmr−/− mice, an effect not observed in WT mice. Consistent with this finding, when osteoclast precursors were cultured in the presence of osteoblasts, more osteoclasts were formed in response to PTH when Osmr−/− osteoblasts were used. Neither PTH1R mRNA levels nor cAMP response to PTH were modified in Osmr−/− osteoblasts. However, RANKL induction in PTH-treated Osmr−/− osteoblasts was sustained at least until 24 hours after PTH exposure, an effect not observed in WT osteoblasts. These data indicate that the transient RANKL induction by intermittent PTH administration, which is associated with its anabolic action, is changed to a prolonged induction in OSMR-deficient osteoblasts, resulting in bone destruction. © 2012 American Society for Bone and Mineral Research.


Introduction

  1. Top of page
  2. Abstract
  3. Introduction
  4. Materials and Methods
  5. Results
  6. Discussion
  7. Disclosures
  8. Acknowledgements
  9. References

Sustained elevation of parathyroid hormone (PTH), such as during hyperparathyroidism or in experimental infusion, significantly enhances osteoclast formation and bone resorption, yet intermittent treatment increases trabecular bone volume. This property of PTH has been exploited for therapeutic use; intermittent treatment with PTH is the only currently available agent proven to have an anabolic effect on bone.1 The latter effect results from direct action on the osteoblast lineage that promotes differentiation of committed precursors,2 inhibits apoptosis,3 and inhibits production of osteocytic sclerostin.4 There is much interest also in the possibility that a contribution to PTH action comes from products of active osteoclasts.5, 6

PTH stimulates osteoclast formation predominantly by upregulating RANKL production by osteoblast lineage cells. In addition, in vitro studies with a gp130-neutralizing antibody indicated that gp130 signaling is also required for the full osteoclastogenic effect of 1,25-dihydroxyvitamin-D3, interleukin-1, and PTH.7 In addition, precursors derived from gp130 knockout mice were unable to form osteoclasts in response to PTH.8 These findings indicate that PTH-induced osteoclast formation is at least partly dependent on expression of gp130,7 a JAK/STAT activating receptor subunit, which, when bound to ligand-specific receptors, is utilized by a wide range of cytokines including interleukin-11 (IL-11), interleukin-6 (IL-6), ciliary neurotrophic factor (CNTF), oncostatin M (OSM), and cardiotrophin-1 (CT-1). Each of these cytokines is required for normal bone metabolism, as indicated by significant skeletal defects in mouse models with deficiencies in gp130, LIF, LIFR, IL-11R, IL-6, CNTF, OSMR, or CT-1.9–15

PTH and the gp130-signaling cytokine OSM activate similar gene targets and have similar effects on the skeleton; both stimulate RANKL production by osteoblast/stromal cells and stimulate osteoclast formation,16–19 and both stimulate bone formation.1, 13, 20 Despite their different early signaling mechanisms both factors depend on the same distal enhancer region to stimulate expression of the pro-osteoclastogenic factor RANKL.19 Furthermore, OSM appears to enhance the action of PTH on RANKL expression through a cyclic AMP (cAMP)-dependent mechanism.19 It has been suggested that one determining factor of whether PTH has an anabolic or catabolic effect is the dynamics of RANKL production, because in vivo work has shown that intermittently injected PTH transiently increases mRNA for RANKL, whereas continuous treatment resulted in a progressive increase.21

In stimulating bone formation, a mechanism that PTH and OSM have in common is action on the osteocyte, where they both inhibit expression of a Wnt pathway inhibitor, sclerostin.13, 20, 22 Sclerostin is critical for normal bone mass: mutations in sclerostin that lead to its inactivation result in sclerosteosis, a syndrome of greatly increased bone mass.23 Downregulation of sclerostin also plays a role in the enhanced bone formation that occurs in response to mechanical stimulation,24 and consistent with its action to reduce sclerostin expression, upregulation of both OSM and OSM receptor (OSMR) expression in bone is an early effect of mechanical loading of bone.25 In contrast to PTH, the actions of OSM to stimulate bone formation and bone resorption appear to be mediated by two distinct receptor signaling pathways: bone formation and suppression of sclerostin is mediated through the LIF receptor (LIFR), while stimulation of RANKL expression, and osteoclast formation, is mediated through OSMR.13

This study sought to determine how signaling through OSMR might influence the anabolic action of PTH.

Materials and Methods

  1. Top of page
  2. Abstract
  3. Introduction
  4. Materials and Methods
  5. Results
  6. Discussion
  7. Disclosures
  8. Acknowledgements
  9. References

PTH treatment of osteoblasts—microarray and validation

Microarray data obtained from a murine stromal cell line (Kusa 4b10) treated with hPTH(1–34) (Bachem, Bubendorf, Switzerland) is previously described.26 Briefly, Kusa 4b10 cells27 were subcultured at 3000 cells/cm2 in α-MEM (Sigma, St. Louis, MO, USA) + 10% fetal bovine serum (FBS) and after 3 days the medium was replaced with osteoblast differentiating medium [α-MEM + 15% heat-inactivated FBS + ascorbate (50 µg/mL) (Sigma), changed three times per week. FBS was reduced to 2% on day 16. On day 17, cells were treated with 10 nM hPTH(1–34) and were collected at 1, 6, and 24 hours after treatment. Triplicate cultures were used, and independent triplicate cultures were used for qPCR. Full details of the bioinformatics analysis of the microarray data are previously published.26 The same differentiation and treatment protocol was also carried out on primary calvarial osteoblasts obtained from 1–3-day-old C57Bl/6 mice, as previously described.13 Briefly, calvariae were dissected from the animal and placed into ice-cold PBS to remove surrounding tissue and pooled. Sequential digestion of the calvariae was carried out using Collagenase Type II (Worthington, Lakewood, NJ, USA), Dispase (Gibco, Grand Island, NY, USA) in phosphate-buffered saline (PBS) (Sigma) for 4 × 10-minute incubations. The supernatant from each digestion was pooled and plated in complete medium (α-MEM + 10% FBS) and grown at 37°C with 5% CO2 in osteoblast differentiation media (as above) for 7 days.

PTH treatment of WT and Osmr−/− mice

The St. Vincent's Health Animal Ethics Committee approved all animal procedures. Osmr−/− mice backcrossed onto C57Bl/6 were obtained from Dr Atsushi Miyajima (The University of Tokyo).28 WT littermates were used as controls. hPTH(1–34) (Bachem, Bubendorf Switzerland) at a submaximal dose of 30 µg/kg was administered 5 days a week to Osmr−/− and WT male mice beginning at 6 weeks of age for 3 weeks. Double calcein labeling was performed on days 3 and 10 before tissues were collected as described previously.12 One hour after the last PTH injection, animals were bled by cardiac puncture while under inhalation anaesthesia, and tissue samples were collected for analysis.

Specimens for histomorphometry were fixed in 4% paraformaldehyde, embedded in methylmethacrylate as described previously,29 and 5-µm sections were stained with toluidine blue12 or Xylenol orange.30 Histomorphometry was carried out in the secondary spongiosa of the proximal tibia (Osteomeasure, Osteometrics Inc, Decatur GA);11 to ensure that effects on PTH on new trabecular formation were not measured, the region selected was a 1.14-mm deep square, commencing 760 µm below the base of the hypertrophic zone of the growth plate. Whole femoral samples were collected for qPCR analysis as previously described.6 Serum calcium was measured by a modified Arsenazo method.31

MicroCT analysis was performed on femoral specimens, fixed as above, with a Skyscan 1076 microCT (Skyscan, Aartselaar, Belgium). During scanning, femora were enclosed in a plastic tube filled with 70% ethanol. The X-ray source was set at 50 kV and 800 µA and sharpening was set to 40%. Projections were acquired over 180° (step of 0.6°), pixel size of 9 µm, and 2000-millisecond exposure. Image slices were reconstructed by NRecon (Skyscan) with the following settings: beam-hardening correction 30%, ring artifact correction 12, no smoothing, and no defect pixel masking. Reconstructed images were straightened with Dataviewer (Skyscan). A 2-mm cross-section of cortical bone, 4 mm from the distal condyles, was analyzed in the transaxial plane. Global thresholding was used to define mineralized bone; the same threshold (91–252) was used for all samples.

Cell culture of Osmr−/− and WT cells and cAMP assay

Osteoclasts were generated from WT and Osmr−/− bone marrow macrophages in the presence of WT and Osmr−/− primary calvarial osteoblasts as previously described.13 Cultures were stimulated with hPTH(1–34) (100 ng/mL) with a change of medium at day 3 and fixed at day 7 with 4% formaldehyde, incubated with acetone:ethanol (50:50), and air dried before TRAP+ staining. TRAP+ cells with two to four or more than five nuclei were counted. Primary calvarial osteoblasts from WT and Osmr−/− mice, differentiated for 7 days were assayed for cAMP formation in response to hPTH(1–34) as previously described.32

PCR method

RNA was isolated from homogenized bone specimens, including bone marrow, prepared as described previously,33 and from cell digests using Trizol (Invitrogen, Carlsbad, CA, USA) and concentration determined by spectrophotometer (Nanodrop ND1000). cDNA was synthesized (Random primers, 10 mM dNTP, 5× First-Strand buffer, 0.1 M DTT, RNaseOUT, Superscript III RT (200 U/µL)) using 1 µg material as follows: 5 minutes at 65°C, 1 minute at 4°C; 5 minutes at 25°C, 60 minutes at 50°C, 15 minutes at 70°C, held at 4°C (Px2 Thermal Cycler, Thermo Electron Corporation, Pittsburgh, PA, USA). Amplification was carried out with AmpliTaq Gold (Perkin-Elmer, Norwalk, CT, USA), SYBR Green (Invitrogen) and specific oligonucleotide primers as shown in Table 1 (Sigma-Genosys, St. Louis, MO, USA). qPCR conditions used were: 1 cycle 10 minutes 95°C, 40 cycles 30 seconds at 95°C, 1 minute at 60°C, 30 seconds at 72°C; 1 cycle 1 minute at 95°C, 30 seconds at 55°C, 30 seconds at 95°C (Stratagene Mx3000P). PTH did not regulate hypoxanthine phosphoribosyltransferase 1 (HPRT1) or β2-microglobulin, the housekeeping genes used for cell culture and skeletal samples, respectively.

Table 1. Primers for qPCR Reactions
GeneForward primer (5′–3′)Reverse primer (3′–5′)
CLCF1CTTAGCTGGGACCTACCTGAACCACACTTCCAAGTTGACCGT
CNTFTCTGTAGCCGCTCTATCTGGGGTACACCATCCACTGAGTCAA
CRLF1CTCCCTGCAAGCTACCTGCAGGGTGGAGGTGTTAAGGAGG
gp130ACGCAGTCAAAGTCCGTCTCCTGATTTGCCCACCTTGTTT
HPRT1TGATTAGCGATGATGAACCAGAGAGGGCCACAATGTGATG
IL-6TAGTCCTTCCTACCCCAATTTCCTTGGTCCTTAGCCACTCCTTC
IL-11TGCTGACAAGGCTTCGAGTAGGATCACAGGTTGGTCTGG
LIFCTCTTCCCATCACCCCTGTATGGTCTTCTCTGTCCCGTTG
LIFRCTTGCAATGTGCCACTCACTCGAGCACCACTTTGTCTTGA
OSMRAAACATGATATTTCAGATAGAGATCAGTAGACTCTTATGAAATGTTTGACACACTCCAA
PTHR1TTCCAGGGATTTTTTGTTGCAGTCCAATGCCAGTGTCCAG

Statistical methods

Statistically significant effects were determined by one- or two-way analysis of variance (ANOVA) followed by Tukey post hoc test using Prism 5. All data is presented as means ± SEM. p < 0.05 was considered statistically significant.

Results

  1. Top of page
  2. Abstract
  3. Introduction
  4. Materials and Methods
  5. Results
  6. Discussion
  7. Disclosures
  8. Acknowledgements
  9. References

Regulation of gp130-binding cytokines and receptors by PTH

PTH treatment of Kusa 4b10 cells led to a dramatic increase in expression levels of a range of cytokines that signal through gp130 and their receptor complex components detected by microarray (Table 2). These results were validated by qPCR on independent samples from Kusa 4b10 cells treated with PTH (Fig. 1) and in primary calvarial osteoblasts (Table 2). The receptor gp130 itself was increased approximately four- to sixfold by PTH at 6 and 24 hours, and OSMR levels were increased 12-fold at the same time points (Fig. 1). The strongest responses observed were a 30–60-fold increase in IL-6 mRNA levels at 1 and 6 hours after PTH administration. IL-11 levels were also dramatically increased by PTH 6 hours after commencement of treatment (Fig. 1). LIF levels were increased approximately fourfold by PTH at 1 and 6 hours after administration (Fig. 1), whereas LIFR was inhibited to a similar extent at 6 and 24 hours (Fig. 1). Cardiotrophin-like cytokine factor 1 (CLCF1) and cytokine receptor-like factor 1 (CRFL1), two components of a composite cytokine that signals through a gp130:LIFR:CNTFR complex were mildly elevated (Fig. 1). mRNA levels of the following IL-6 superfamily members were below the level of detection for analysis by microarray: IL-31, GPL (gp130-like receptor), IL-27, CT-1, and CT-2. mRNA levels of the following IL-6 superfamily members were not regulated by PTH or PTHrP in the RNA prepared for microarray: CNTFR, CNTF, IL-11R, IL-6R, OSM, sortilin 1, and WSX-1.

Table 2. Effects of PTH Treatment on Expression of gp130 Family Members in Kusa 4b10 Cells (Microarray Results) and in Primary Calvarial Osteoblasts (Real-Time PCR)
Protein1 Hour6 Hours24 Hour
Kusa 4b10 (Microarray)Osteoblasts (PCR)Kusa 4b10 (Microarray)Osteoblasts (PCR)Kusa 4b10 (Microarray)Osteoblasts (PCR)
  1. Values are fold change in gene expression. For microarray data, where multiple probesets were available, the value is the mean fold change for all probesets. For real-time PCR data, data is mean ± SEM from three independent experiments. For clarity, only statistically significant changes (p < 0.05) are shown; —, no statistically significant change.

Glycoprotein 130 (gp130)3.81.7 ± 0.11.6 ± 0.4
Interleukin-6 (IL-6)30.016 ± 4.361.410 ± 3.6
Interleukin-11 (IL-11)4.3 ± 1.649.87.1 ± 2.3
Leukemia inhibitory factor (LIF)4.74.4 ± 0.93.9
LIF receptor (LIFR)−3.50.26 ± 0.07
Oncostatin M receptor (OSMR)4.52.4 ± 0.82.23.2 ± 0.6
Cytokine receptor-like factor-1 (CRLF1)1.72.4
Cardiotrophin-like cytokine factor-1 (CLCF1)1.82.0
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Figure 1. Effect of PTH on gp130 cytokines and receptor mRNA levels in Kusa 4b10 cells. Kusa 4b10 cells were differentiated for 17 days in osteoblast differentiation media then treated with hPTH(1–34) at 10 nM (black squares) or untreated (white squares). Shown are mRNA levels for glycoprotein 130 (gp130), oncostatin M receptor (OSMR), interleukin-6 (IL-6), interleukin-11 (IL-11), leukemia inhibitory factor (LIF), LIF receptor (LIFR), cytokine receptor-like factor 1 (CRLF1), and cardiotrophin-like cytokine factor 1 (CLCF1). Data are means ± SEM fold change of each gene relative to HPRT1 from three independent experiments. *p < 0.05, **p < 0.01, ***p < 0.001 versus untreated control cells at the same time point.

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Anabolic PTH treatment reduced trabecular bone volume in Osmr−/− mice

Osmr−/− mice exhibit a higher than usual bone mass at 12 weeks of age, including a low level of osteoclast formation and a low level of bone formation.13 Treatment of 6-week-old WT mice with 30 µg/kg/day of hPTH(1–34) for 3 weeks provided a significant increase in both trabecular bone volume and trabecular thickness within the secondary spongiosa of WT mice (Fig. 2A, B). PTH treatment did not significantly alter cortical thickness in mice of either genotype (mean Ct.Th ± SEM: WT vehicle: 138.6 ± 4.4 µm; WT + PTH: 139.6 ± 2.8 µm; Osmr−/− vehicle: 138.5 ± 4.1 µm; Osmr−/−+ PTH: 137.3 ± 3.7 µm). Osmr−/− mice of the same age and sex demonstrated a significantly higher trabecular bone volume, confirming our previous report.13 In these mice, the same PTH treatment resulted in a significant reduction in trabecular bone volume, trabecular number, and trabecular separation (Fig. 2B) indicating a catabolic effect of intermittent PTH administration in Osmr−/− mice.

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Figure 2. The response of trabecular bone to intermittent low dose PTH treatment is significantly modified in Osmr−/− mice. (A) Von Kossa stained sections show the extent of trabecular bone in each group as noted on the figure; white boxes show approximate location of the region measured for histomorphometry. (B) Trabecular bone volume (BV/TV) and trabecular thickness (Tb.Th) were both significantly increased by 30 µg/kg/day hPTH(1–34) five times per week for 3 weeks in WT mice. In contrast, in Osmr−/− mice, the same treatment significantly reduced BV/TV, trabecular number (Tb.N), and increased trabecular separation (Tb.Sp). Data are mean ± SEM from 6–13 mice per group. *p < 0.05; **p < 0.01 versus saline-treated mice of the same genotype (effect of PTH); +p < 0.05 versus saline-treated WT (effect of OSMR deletion).

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Histomorphometric parameters indicated a significant increase in osteoblast generation and osteoid deposition in both WT and Osmr−/− mice treated with PTH compared with saline-treated littermate controls (Fig. 3). Although these parameters were lower in the Osmr−/− mice, osteoid and osteoblast markers were increased to the same extent by PTH treatment in both wild-type (WT) and Osmr−/− mice. Furthermore, dynamic histomorphometry revealed that, whereas there was no increase in the mineralizing surface, mineral appositional rate (MAR), a parameter that indicates osteoblast function, was significantly elevated in both WT and Osmr−/− mice; as observed with the osteoid and osteoblast parameters, the stimulatory effect of PTH on MAR was not significantly different between the two genotypes by two-way ANOVA.

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Figure 3. The enhanced osteoblast formation in response to intermittent low-dose PTH treatment is not significantly modified in Osmr−/− mice. Osteoid volume (OV/BV), osteoid surface (OS/BS), osteoblast surface (ObS/BS), number of osteoblasts (NOb/BPm), and mineral appositional rate (MAR) were all significantly increased in mice of both genotypes treated with 30 µg/kg/day hPTH(1–34) five times per week for 3 weeks. Mineralizing surface (MS/BS) was not significantly modified. Data are mean ± SEM from 6–13 mice per group. *p < 0.05; **p < 0.01 versus saline-treated mice of the same genotype.

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In WT mice, PTH treatment did not modify osteoclast surface or osteoclast numbers (Fig. 4A). In contrast, in Osmr−/− mice, both osteoclast numbers and surface were almost doubled by PTH treatment, which also caused a mild, but significant increase in the size of the Osmr−/− osteoclasts (Fig. 4A). Furthermore, 1 hour after the final PTH injection, serum calcium levels were significantly elevated in Osmr−/− but not in WT mice, compared with saline-treated controls (Fig. 4A), consistent with elevated osteoclast activity at this stage.

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Figure 4. The response of osteoclasts to PTH treatment is enhanced in Osmr−/− mice and in cocultures of cells from Osmr−/− mice. (A) Osteoblast surface (OcS/BS), number of osteoclasts (NOc/BPm), and osteoclast length (OcL) were significantly increased in PTH-treated Osmr−/− mice, but not in WT mice treated with 30 µg/kg/day hPTH(1–34) five times per week for 3 weeks. Serum calcium levels were significantly increased, 1 hour after the last PTH injection in Osmr−/− mice, but not in WT mice. Data are mean ± SEM from 6–13 mice per group. *p < 0.05; **p < 0.01 versus saline-treated mice of the same genotype (effect of PTH); ++p < 0.01, ++ + p < 0.001 versus saline-treated WT (effect of OSMR deletion). (B) Number of TRAcP+ multinucleated cells generated from PTH-treated cocultures was significantly increased when bone marrow and osteoblasts from Osmr−/− mice were used; the left column in each pair shows TRAcP+ cells with two to four nuclei, whereas the right column shows cells with 5+ nuclei. Data is mean ± SD from two independent experiments. *p < 0.05, ***p < 0.001 versus culture with WT osteoblasts (O'BLAST) and the same genotype of bone marrow (MARROW) (effect of Osmr−/− osteoblasts); ++ + p < 0.001 versus culture with WT marrow and the same genotype of osteoblasts (effect of Osmr−/− marrow). Images below graph are TRAcP stained representative images for each column, all taken at the same magnification.

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We next carried out coculture experiments to determine whether there was a difference in the ability of Osmr−/− osteoblasts to support osteoclast formation in response to PTH. We have previously shown that osteoclast precursors from Osmr−/− mice have a normal osteoclastogenic response to recombinant RANKL and macrophase-colony stimulating factor (M-CSF).13 Consistent with the results observed in vivo, more osteoclasts were formed by Osmr−/− osteoclast precursors in response to PTH in coculture, and this effect was enhanced in the presence of Osmr−/− osteoblasts (Fig. 4B). The osteoclasts formed in the presence of Osmr−/− osteoblasts were also larger than those formed in the presence of WT osteoblasts, and there were more osteoclasts with more than five nuclei (Fig. 4B). Surprisingly the number of osteoclasts formed in the presence of Osmr−/− osteoblasts was enhanced further in the presence of Osmr−/− bone marrow macrophages. Previously, we have shown that OSMR cannot be detected in osteoclasts by immunohistochemistry of mouse bone. To confirm this, and check for expression in precursors, we carried out real-time PCR on bone marrow macrophages treated with RANKL and M-CSF, but OSMR was below the limit of detection at all time points, confirming that the receptor is not expressed in osteoclasts or their precursors.

The ability of Osmr−/− osteoblasts to respond to PTH was then assessed. Osmr−/− calvarial osteoblasts, differentiated for 7 days in osteoblast differentiation media, expressed the same level of PTH receptor (PTH1R) mRNA as WT osteoblasts prepared in the same manner (Fig. 5A), and the cAMP dose–response and time course of response to PTH was identical in Osmr−/− and WT calvarial osteoblasts (Fig. 5A). OSM treatment at 2.5 ng/mL did not induce cAMP activity (data not shown). The effects of PTH on expression of mRNA for gp130, IL-6, IL-11, LIF, and LIFR were not significantly different in Osmr−/− and WT primary calvarial osteoblasts (data not shown).

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Figure 5. PTH-induced increase in RANKL mRNA levels is sustained in osteoblasts from Osmr−/− mice, despite a normal cAMP response and normal PTH receptor levels. (A) Real-time PCR for PTH receptor (PTH1R) levels in primary calvarial osteoblasts from WT and Osmr−/− mice, each differentiated in osteoblast differentiation media for 7 days. Dose–response curve and time course of cAMP response of WT and Osmr−/− calvarial osteoblasts, differentiated in osteoblast differentiation media for 7 days, to hPTH(1–34). (B) mRNA levels of osteoprotegerin (OPG) and RANKL in whole bones collected from WT and Osmr−/− mice treated with 30 µg/kg/day hPTH(1–34) five times per week for 3 weeks; samples were collected 1 hour after the last PTH injection. Data are mean ± SEM from six to nine mice per group. *p < 0.05; ***p < 0.001 versus saline-treated mice of the same genotype. (C) OPG and RANKL mRNA levels, relative to HPRT1, were assessed in osteoblasts derived from WT and Osmr−/− mice with PTH treatment (100 ng/mL). Samples were collected at 1, 6, 8, and 24 hours after PTH treatment was commenced. Data shown are means ± SEM from four independent experiments. *p < 0.05 versus cells of the same genotype that were not treated with PTH. (D) ICER mRNA levels, relative to HPRT1 in osteoblasts derived from WT and Osmr−/− mice with PTH treatment (100 ng/mL). Data shown are mean ± SEM from four independent experiments.

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Levels of mRNA transcripts for key factors that regulate osteoclast formation were assessed, both in primary calvarial osteoblasts, and in bone specimens from the WT and Osmr−/− mice treated with PTH, collected 1 hour after the last PTH injection. Osteoprotegerin (OPG) was significantly reduced and RANKL significantly increased by PTH treatment to a similar extent in the bone specimens from both WT and Osmr−/− mice treated with PTH (Fig. 5B); the reduction in OPG was slightly less in Osmr−/− mice, but this difference was not statistically significant. In calvarial osteoblast preparations, PTH treatment did not significantly modify OPG mRNA levels compared with untreated cells in WT or Osmr−/− osteoblasts at any time point studied, nor was it significantly different between the two genotypes (Fig. 5C). RANKL expression was equal in WT and Osmr−/− osteoblasts, and PTH treatment increased RANKL mRNA levels significantly at 6 and 8 hours after treatment was commenced in both populations (Fig. 5C). After 24 hours, in four independent experiments, although the high level of RANKL induced by PTH had returned to baseline levels in the WT calvarial osteoblasts, the high level of RANKL expression was sustained in osteoblasts from Osmr−/− mice, so that at 24 hours it was still 17-fold higher than baseline, and threefold higher than WT treated with PTH (Fig. 5C). In later separate experiments using the same dose of PTH, RANKL mRNA levels after PTH treatment were also elevated in osteoblasts from Osmr−/− but not WT at 16 hours (mean RANKL:HPRT1 of two independent experiments ± SD: WT untreated: 0.021 ± 0.014; WT PTH: 0.029 ± 0.010; Osmr−/− untreated: 0.036 ± 0.001; Osmr-/- PTH: 0.071 ± 0.012) and 24 hours (WT untreated: 0.014 ± 0.010; WT PTH: 0.036 ± 0.012; Osmr−/− untreated: 0.037 ± 0.013; Osmr−/− PTH: 0.088 ± 0.015). Because enhanced RANKL response to PTH has also been described in mice lacking the inducible cAMP early repressor (ICER),34 we assessed expression of this factor in WT and OSMR-deficient osteoblasts treated with PTH, but observed no difference in response (Fig. 5E). This consistent finding of sustained elevation in RANKL mRNA levels may explain the enhanced osteoclast formation and bone resorption in Osmr−/− mice treated with PTH.

Discussion

  1. Top of page
  2. Abstract
  3. Introduction
  4. Materials and Methods
  5. Results
  6. Discussion
  7. Disclosures
  8. Acknowledgements
  9. References

The anabolic effect of PTH remains poorly understood, although a number of key candidates play significant roles in this effect.35 In this work, we show that PTH modifies osteoblastic expression of a range of gp130-interacting cytokines and receptors. Furthermore, when PTH was administered to mice deficient in OSMR, the anabolic effect was converted to a catabolic effect. Even though PTH stimulated bone formation in these mice, osteoclast numbers were increased, with the imbalance leading to a reduction in trabecular bone volume and trabecular number. The enhanced osteoclast formation caused by PTH treatment in the absence of OSMR was also observed in cultured cells from the Osmr−/− mice and this may relate to a conversion of the normally transient increase in RANKL expression associated with intermittent PTH treatment21 to a persistently high level of RANKL in the absence of OSMR.

The stimulation of IL-11 mRNA levels by PTH confirmed earlier work showing increased IL-11 bioactivity in primary osteoblasts treated with PTH.7 The PTH-induced increases in gp130, IL-6, and LIF mRNA levels have also been reported previously.7, 36 We now report stimulation of two newer members of this family, CLCF-1, and one of its receptor components, CRLF-1. These two factors form a signaling complex with gp130, LIFR, and CNTFR,37–39 and we have recently shown that CLCF-1 (also known as CLC) inhibits osteoblast differentiation in vitro.11 In addition, expression of OSMR, a receptor subunit that heterodimerizes with gp130 and is capable of intracellular signaling, was also significantly enhanced by PTH, an effect that has not previously been reported. We have recently shown that both OSM and OSMR are expressed by osteoblasts and osteocytes, but not osteoclasts13 and that OSMR mediates the stimulatory effect of OSM on RANKL and osteoclast formation.13

In contrast to enhanced OSMR expression, LIFR expression was inhibited by PTH treatment of osteoblasts. Despite its name, LIFR is not utilized only by LIF, but is a common signaling subunit that also mediates the actions of CT-1, CNTF, CLCF-1, cardiotrophin-2, and OSM.38, 40, 41 OSM, CT-1, CNTF, and CLCF-1, like LIF, also regulate osteoblast and osteoclast formation.11–13, 42 Although the binding affinity of murine OSM to LIFR is much lower than its affinity for OSMR,43 we found recently that murine OSM can act through LIFR.13 Furthermore, through this path, OSM stimulates bone formation and inhibits expression of sclerostin, an osteocytic inhibitor of bone formation.13 Sclerostin is also strongly downregulated by PTH, and involved in its anabolic action.4 The similar increase in bone formation and osteoblast indices with PTH treatment in WT and Osmr−/− mice, despite the lower level of bone formation in the latter, indicates that even though OSMR is required for normal bone formation in the process of physiological bone remodeling, it does not play a critical role in the effect of PTH on osteoblast differentiation or activity. This does not negate the possibility that the effect of PTH on bone formation may involve OSM:LIFR signaling and its regulation of sclerostin.

The RANKL gene contains five distal enhancer regions, shown to be direct targets of the vitamin D receptor and/or cAMP. OSM and PTH both enhance RANKL expression through an additional region −88 kb upstream of the transcription start site.19, 44, 45 STAT3 binds to this region, where it alters histone H4 acetylation and recruits RNA polymerase II.45 We have previously shown that STAT3 is phosphorylated by OSM in murine primary osteoblasts, and RANKL expression induced by OSM is impaired in the absence of OSMR.13 In addition, we previously reported that osteoclast formation in response to 1,25-dihydroxyvitamin-D3 is impaired in the presence of Osmr−/− osteoblasts.13 Although stimulation of RANKL expression by both PTH and OSM depends on cAMP and PKA expression, OSM does not enhance CRE-luciferase reporter activity,46 nor did it stimulate cAMP activity itself. In this context, enhanced osteoclast formation in the absence of OSMR is surprising. Although we have no evidence that RANKL expression levels per se are increased in the absence of OSMR, the PTH-induced expression of RANKL by OSMR-deficient osteoblasts was sustained.

Sustained stimulation of RANKL expression in Osmr−/− osteoblasts, and the enhanced osteoclast formation in response to PTH that we observe here is consistent with data showing that catabolic effects of PTH results from a sustained stimulation of RANKL while anabolic effects of intermittent PTH administration in WT rats only transiently increased RANKL expression.21 Our observation of OSMR deletion converting an anabolic dosage of PTH to a catabolic effect is consistent with the anabolic dose of PTH requiring a transient stimulation of RANKL.5, 6 Although the enhanced osteoblast activity does not depend on this stimulation to be transient, because we observe here an unchanged enhancement in osteoblast generation, osteoid production and mineral apposition rate in response to PTH, this increase in osteoblast activity cannot have an anabolic effect in the presence of the high level of osteoclast formation that occurs in the absence of OSMR signaling.

A number of other factors have been shown to be required for the anabolic effect of PTH on the skeleton.35 These include transcription factors ATF447 and CREM,34 signaling molecules such as β-arrestin,48 cytokines such as FGF2,49 IL-18,50 and IGF-1,51 matrix proteins including osteonectin,52 and factors that regulate Wnt pathway signaling including sFRP-153, 54 and sclerostin.4 In most of these studies, when the changes in cell number and behavior have been reported, it is the action of PTH on osteoblast generation or activity that has been modified. There are two significant exceptions. The modified effect of anabolic PTH in CREM and β-arrestin null mice is similar to our observations here in OSMR-deficient mice. That is, the increase in osteoblast number and surface induced by PTH is unchanged in these mice, but in all three knockout models, osteoclast generation in response to PTH is enhanced. Although a direct comparison between these studies cannot be made, because of differences in treatment protocols and analyses performed, the increased in osteoclast formation in response to PTH is striking, particularly given the involvement of both β-arrestin and CREM in intracellular cAMP-dependent signaling of PTH1R. Furthermore, in cell culture studies, osteoclast formation in response to PTH treatment was enhanced in cultures when both osteoblast and osteoclast precursors were derived from either β-arrestin55 or CREM34 null mice.

Although the mechanism by which intermittent PTH administration induced a sustained increase in RANKL in OSMR null osteoblasts remains unclear, our observation is similar to previous work in osteoblasts null for β-arrestin55 or the inducible cAMP early repressor (ICER).34 β-Arrestin inhibits cAMP accumulation in COS-7 cells,56 but because cAMP accumulation in OSMR deficient osteoblasts treated with PTH was normal, altered β-arrestin signaling is unlikely to be the cause of the sustained RANKL elevation we report. ICER, a truncated form of the CREM family of transcription factors inhibits cAMP-dependent transcription,57 and its expression is stimulated in MC3T3-E1 cells and in calvarial samples treated with PTH.58 OSMR null osteoblasts did not demonstrate altered basal levels of ICER or modified ICER transcriptional response to PTH compared with WT osteoblasts. This indicates that the role of OSMR in shortening the osteoblastic RANKL response to PTH is not upstream of ICER or β-arrestin.

In the coculture system, OSMR deficient osteoclast precursors had an enhanced ability to differentiate in response to PTH in the presence of Osmr−/− osteoblasts, even though osteoclast precursors lack PTHR1, and neither osteoclasts nor their precursors express OSMR. There is no inherent difference in the ability of Osmr−/− osteoclast precursors to form osteoclasts; we have previously shown that osteoclast formation in response to RANKL is not altered in Osmr−/− osteoclast precursors,13 and others have reported no difference in macrophage colony forming units from Osmr−/− bone marrow.28 Furthermore, osteoclast formation by Osmr−/− osteoclast precursors in response to PTH was normal in the presence of WT osteoblasts. Thus, the microenvironment provided by Osmr−/− osteoblasts enhances the ability of Osmr−/− osteoclast precursors to form osteoclasts in response to PTH, although the mechanism is not yet resolved. The increased responsiveness of both of these populations would both contribute to the increased osteoclast formation observed in response to PTH in vivo.

In conclusion, OSMR expression is induced by PTH, suggesting an amplification of OSM effects in osteoblasts in the presence of PTH. Signaling through OSMR limits the duration of RANKL production in osteoblast lineage cells, thus allowing a transient rise in RANKL production, which is required for anabolic action of PTH.

Acknowledgements

  1. Top of page
  2. Abstract
  3. Introduction
  4. Materials and Methods
  5. Results
  6. Discussion
  7. Disclosures
  8. Acknowledgements
  9. References

This work was supported by the NHMRC (Australia) Program Grant 345401 and Project Grant 1002728 to NAS and TJM; NAS is an NHMRC (Australia) Senior Research Fellow.

We thank Dr. Atsushi Miyajima for providing Osmr−/− breeders. We also thank the staff at the Bioresources Centre, St. Vincent's Health, for excellent animal care. The Victorian State Government Operational Infrastructure Support Scheme provides support to St. Vincent's Institute.

Authors' roles: Study design: NAS, TJM. Study conduct: ECW, NEM, IJP, PWMH, EHA, JMQ. Data collection: ECW, NEM, IJP, PWMH. Data analysis: ECW, IJP, NEM, PWMH, NAS. Drafting manuscript: ECW, NAS. Revising manuscript: ECW, NAS, TJM. Approving final version of manuscript: all authors. NAS takes responsibility for the integrity of the data analysis.

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  3. Introduction
  4. Materials and Methods
  5. Results
  6. Discussion
  7. Disclosures
  8. Acknowledgements
  9. References
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