Endogenous parathyroid hormone–related protein compensates for the absence of parathyroid hormone in promoting bone accrual in vivo in a model of bone marrow ablation

Authors

  • Qi Zhu,

    1. The State Key Laboratory of Reproductive Medicine, The Research Center for Bone and Stem Cells, Department of Anatomy, Histology and Embryology, Nanjing Medical University, Nanjing, China
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    • QZ and XZ contributed equally to this work.
  • Xichao Zhou,

    1. The State Key Laboratory of Reproductive Medicine, The Research Center for Bone and Stem Cells, Department of Anatomy, Histology and Embryology, Nanjing Medical University, Nanjing, China
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    • QZ and XZ contributed equally to this work.
  • Min Zhu,

    1. The State Key Laboratory of Reproductive Medicine, The Research Center for Bone and Stem Cells, Department of Anatomy, Histology and Embryology, Nanjing Medical University, Nanjing, China
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  • Qian Wang,

    1. The State Key Laboratory of Reproductive Medicine, The Research Center for Bone and Stem Cells, Department of Anatomy, Histology and Embryology, Nanjing Medical University, Nanjing, China
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  • David Goltzman,

    1. The Department of Medicine, McGill University, Montreal, Canada
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  • Andrew Karaplis,

    1. The Department of Medicine, McGill University, Montreal, Canada
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  • Dengshun Miao

    Corresponding author
    • The State Key Laboratory of Reproductive Medicine, The Research Center for Bone and Stem Cells, Department of Anatomy, Histology and Embryology, Nanjing Medical University, Nanjing, China
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Address correspondence to: Dengshun Miao, The State Key Laboratory of Reproductive Medicine, The Research Center for Bone and Stem Cells, Department of Anatomy, Histology and Embryology, Nanjing Medical University, Nanjing, Jiangsu, 210029, The People's Republic of China. E-mail: dsmiao@njmu.edu.cn

ABSTRACT

To assess the effect of hypoparathyroidism on osteogenesis and bone turnover in vivo, bone marrow ablation (BMXs) were performed in tibias of 8-week-old wild-type and parathyroid hormone–null (PTH−/−) mice and newly formed bone tissue was analyzed from 5 days to 3 weeks after BMX. At 1 week after BMX, trabecular bone volume, osteoblast numbers, alkaline phosphatase-positive areas, type I collagen-positive areas, PTH receptor–positive areas, calcium sensing receptor–positive areas, and expression of bone formation–related genes were all decreased significantly in the diaphyseal regions of bones of PTH−/− mice compared to wild-type mice. In contrast, by 2 weeks after BMX, all parameters related to osteoblastic bone accrual were increased significantly in PTH−/− mice. At 5 days after BMX, active tartrate-resistant acid phosphatase (TRAP)-positive osteoclasts had appeared in wild-type mice but were undetectable in PTH−/− mice, Both the ratio of mRNA levels of receptor activator of NF-κB ligand (RANKL)/osteoprotegerin (OPG) and TRAP-positive osteoclast surface were still reduced in PTH−/− mice at 1 week but were increased by 2 weeks after BMX. The expression levels of parathyroid hormone–related protein (PTHrP) at both mRNA and protein levels were upregulated significantly at 1 week and more dramatically at 2 weeks after BMX in PTH−/− mice. To determine whether the increased newly formed bones in PTH−/− mice at 2 weeks after BMX resulted from the compensatory action of PTHrP, PTH−/−PTHrP+/− mice were generated and newly formed bone tissue was compared in these mice with PTH−/− and wild-type mice at 2 weeks after BMX. All parameters related to osteoblastic bone formation and osteoclastic bone resorption were reduced significantly in PTH−/−PTHrP+/− mice compared to PTH−/− mice. These results demonstrate that PTH deficiency itself impairs osteogenesis, osteoclastogenesis, and osteoclastic bone resorption, whereas subsequent upregulation of PTHrP in osteogenic cells compensates by increasing bone accrual. © 2013 American Society for Bone and Mineral Research

Introduction

Hypoparathyroidism is a clinical disorder that may present as a result of parathyroid tissue destruction or removal or autoimmune suppression of secretion, or may present as a congenital disorder associated with defined genetic defects, including abnormalities of parathyroid gland development or of parathyroid hormone (PTH) biosynthesis, or secretion. This disease is characterized by low serum calcium levels, elevated serum phosphorus levels, and absent or inappropriately low levels of PTH in the circulation.[1] Regardless of its etiology, the effects of chronic PTH deficiency on the human skeleton are profound. Bone mass is regulated by a delicate balance between bone resorption and formation in a tightly coordinated process termed remodeling. PTH is one of the pivotal modulators of the rate of bone remodeling, and a reduction or absence of circulating PTH leads to a profound reduction accompanied by an increase in bone mass in both cancellous and cortical compartments.[2-4] Results from histomorphometric analysis of iliac crest bone biopsies demonstrated that both the bone formation rate and resorption rate are reduced significantly in trabecular and cortical compartments in hypoparathyroid subjects.[3, 5] However, the mechanisms of the effects of hypoparathyroidism on bone remodeling are unclear.

We previously reported a mouse model deficient in PTH by targeting the Pth gene in embryonic stem cells. Adult Pth-null mice develop hypocalcemia, hyperphosphatemia, and low circulating 1,25-dihydroxyvitamin D3 (1,25(OH)2D3) levels consistent with primary hypoparathyroidism.[6, 7] We also demonstrated that bone turnover was reduced, leading to increased trabecular and cortical bone volume in these adult PTH-deficient mice. When mutant mice were placed on a low-calcium diet, renal 25-hydroxyvitamin D 1 alpha-hydroxylase expression increased despite the absence of PTH, leading to a rise in circulating 1,25(OH)2D3 levels, marked osteoclastogenesis, and profound bone resorption.[7] These results demonstrated the dependence of the skeletal phenotype in animals with genetically depleted PTH on the external environment as well as on internal hormonal and ionic factors.

Mechanical bone marrow ablation (BMX) is a very useful experimental model for examining the sequence and mechanism of events involved in osteogenesis and bone turnover within a defined time interval.[8-15] Mechanical BMX provokes a predictable sequence of events, including the formation of a blood clot with inflammation and capillary invasion, mesenchymal cell migration, osteoblast proliferation, and formation of new cancellous medullary bone, followed by osteoclastic bone resorption and bone marrow reconstitution.[16-19] Previous studies have employed this model to investigate the effect of exogenous PTH on the formation of new bone that forms after marrow ablation.[20] Those results showed that the abundance of bone formed in response to PTH1-34 in the marrow cavity after marrow ablation is far more than that formed in response to either the hormone alone without marrow ablation, or to marrow ablation alone.[20] Recently, we employed 1,25-dihydroxyvitamin D (1,25(OH)2 D)-deficient animals with secondary hyperparathyroidism to assess the effect of endogenous PTH on osteogenesis and bone turnover in vivo after mechanical BMX.[21] We showed that endogenous elevated PTH can also increase bone, and does this via enhancing osteoprogenitor differentiation into cells of the osteoblast lineage.

In the present study, we used the mechanical BMX model in 8-week-old wild-type and PTH−/− mice to assess the sequence and mechanisms of osteogenesis in the absence of PTH.

Subjects and Methods

Animals

We previously described the generation, by homologous recombination in embryonic stem cells, of mice carrying a disrupted PTH gene[6] and a disrupted PTHrP gene.[22] To obtain wild-type, PTH−/−, and PTH−/−PTHrP+/− littermates, male and female PTH+/− and PTHrP+/− mice on a C57BL/6J background were crossed. Genomic DNA was isolated from tail clips for genotyping by methods previously described.[23] Wild-type and mutant PTH alleles were detected using a 0.2-kb HindIII/XhoI genomic DNA fragment, and as mutant PTHrP alleles, a 0.62-kb SacI/XhoI genomic fragment was used after digestion of tail tip genomic DNA with PvuII. Mice were maintained in a virus-free and parasite-free barrier facility and exposed to a 12-hour light, 12-hour dark cycle. After weaning, animals were fed normal mouse chow containing 0.97% calcium and 0.85% phosphorus (Charles River Laboratories, St. Constant, Canada) until 2 months of age. The use of animals in this study was approved by the Institutional Animal Care and Use Committee of Nanjing Medical University.

Mechanical bone marrow ablation

BMX of the left tibias was achieved by using a technique we previously reported.[21] Briefly, the mice were given a preemptive dose of buprenorphine (0.05 mg/kg subcutaneously [SC]) and anesthetized with a combination of ketamine (100 mg/kg) and xylazine (10 mg/kg) administered intraperitoneally. The left stifle was shaved and scrubbed with 70% alcohol and povidone–iodine. A 3-mm to 5-mm skin incision was made on the anteromedial surface of the stifle, and the patella was released and retracted laterally. A 23-gauge needle was used in a gently twisting motion to bore a hole through the patellar groove to access the marrow cavity. A 25-gauge needle was inserted into the marrow cavity, which was flushed with 3 mL warm saline. The turbulent motion of the saline, which exits around the needle, disrupts the marrow and removes marrow cells. The patella was returned to its natural position, and the medial ligamentous structures were secured with 5-0 absorbable suture. The skin was closed with 5-0 nylon. Supplemental heat was provided throughout the procedure and recovery, and subcutaneous fluids were given in the immediate postoperative period. Buprenorphine (0.05 mg/kg SC) was administered for analgesia every 12 hours for 2 days. No abnormalities in ambulation, grooming, food and water intake, urination, or defecation were noted postoperatively.

Bone transplantation

Femurs from 1-week old wild-type mice with BMX were transplanted into the back muscle of ROSA26 (B6;129S-Gt(ROSA)26Sor/J) mice, which express lacZ ubiquitously.[24] After 1 week, transplanted bones were harvested and pre-embedding lacZ staining was performed as described.[25]

Micro–computed tomography

Tibias were dissected free of soft tissue, fixed overnight in 70% ethanol, and analyzed by micro–computed tomography (µCT) with a SkyScan 1072 scanner and associated analysis software (SkyScan, Antwerp, Belgium) as described.[26] Briefly, image acquisition was performed at 100 kV and 98 mA with a 0.98 rotation between frames. During scanning, the samples were enclosed in tightly fitting plastic wrap to prevent movement and dehydration. Thresholding was applied to the images to segment the bone from the background. Two-dimensional images were used to generate three-dimensional renderings using the 3D Creator software supplied with the instrument. The resolution of the µCT images is 18.2 μm.

Western blot analysis

Proteins were extracted from diaphyseal regions and quantitated using a kit (Bio-Rad, Mississauga, Ontario, Canada). Thirty-microgram protein samples were fractionated by SDS-PAGE and transferred to nitrocellulose membranes. Immunoblotting was carried out as described[27] using antibodies against insulin-like growth factor 1 (IGF-1) (Santa Cruz Biotechnology, Santa Cruz, CA, USA), Runx2 (Santa Cruz Biotechnology), PTHrP1-34 peptide (Santa Cruz Biotechnology), and PTH/PTHrP receptor (Clone 3D1.1; Upstate, Syracuse, NY, USA). β-tubulin (Santa Cruz Biotechnology) was used as a loading control. Bands were visualized using enhanced chemiluminescence (Amersham, Aylesbury, UK).

Quantitative real-time RT-PCR

RNA was isolated from diaphyseal regions, using Trizol reagent (Invitrogen, Carlsbad, CA, USA) according to the manufacturer's protocol. Reverse transcription reactions were performed using the SuperScript First-Strand Synthesis System (Invitrogen) as described.[26] To determine the number of cDNA molecules in the reverse transcribed samples, real-time PCR analyses were performed using the LightCycler system (Roche, Indianapolis, IN, USA). PCR was performed using 2 μL LightCycler DNA Master SYBR Green I (Roche), 0.25 μM of each 5′ and 3′ primer, and 2-μL samples or H2O to a final volume of 20 µL. The MgCl2 concentration was adjusted to 3 mM. Samples were denatured at 95°C for 10 seconds, with a temperature transition rate of 20°C per second. Amplification and fluorescence determination were carried out in four steps: denaturation at 95°C for 10 seconds, with a temperature transition rate of 20°C/second; annealing for 5 seconds, with a temperature transition rate of 8°C/s; extension at 72°C for 20 seconds, with a temperature transition rate of 4°C/s; and detection of SYBR Green fluorescence, which reflects the amount of double-stranded DNA, at 86°C for 3 seconds. The amplification cycle number was 35. To discriminate specific from nonspecific cDNA products, a melting curve was obtained at the end of each run. Products were denatured at 95°C for 3 seconds, and the temperature was then decreased to 58°C for 15 seconds and raised slowly from 58°C to 95°C using a temperature transition rate of 0.1°C/s. To determine the number of copies of the targeted DNA in the samples, purified PCR fragments of known concentrations were serially diluted and served as external standards that were measured in each experiment. Data were normalized with GAPDH levels in the samples. The primer sequences used for the real-time PCR were the same as described previously.[21]

Histology

Tibias were removed and fixed in PLP fixative (2% paraformaldehyde containing 0.075 M lysine and 0.01 M sodium periodate) overnight at 4°C and processed histologically as described.[28] Tibias were decalcified in ethylene-diamine tetraacetic acid (EDTA)-glycerol solution for 5 to 7 days at 4°C. Decalcified tibias were dehydrated and embedded in paraffin, after which 5-μm sections were cut on a rotary microtome. The sections were stained with hematoxylin and eosin (HE) or histochemically for total collagen and alkaline phosphatase (ALP) activity and tartrate-resistant acid phosphatase (TRAP) or immunohistochemical staining as described in the next section.

Histochemical staining for collagen, ALP, and TRAP

Total collagen was detected in paraffin-embedded sections using a modified method of Lopez-De Leon and Rojkind.[29] Dewaxed sections were exposed to 1% Sirius Red in saturated picric acid for 1 hour. After washing with distilled water, the sections were counterstained with hematoxylin and mounted with Biomount medium.

Enzyme histochemistry for ALP activity was performed as described.[30] Briefly, following preincubation overnight in 1% magnesium chloride in 100 mm Tris–maleate buffer (pH 9.2), dewaxed sections were incubated for 2 hours at room temperature in a 100 mM Tris–maleate buffer containing naphthol AS-MX phosphate (0.2 mg/mL; Sigma, St. Louis, MO, USA) dissolved in ethylene glycol monomethyl ether (Sigma) as substrate and Fast Red TR (0.4 mg/mL; Sigma) as a stain for the reaction product. After washing with distilled water, the sections were counterstained with Vector methyl green nuclear counterstain (Vector Laboratories Inc., Ontario, Canada) and mounted with Kaiser's glycerol jelly.

Enzyme histochemistry for TRAP was performed using a modification of a previously described protocol.[31] Dewaxed sections were preincubated for 20 minutes in buffer containing 50 mM sodium acetate and 40 mM sodium tartrate at pH 5.0. Sections were then incubated for 15 minutes at room temperature in the same buffer containing 2.5 mg/mL naphthol AS-MX phosphate (Sigma) in dimethylformamide as substrate and 0.5 mg/mL Fast Garnet GBC (Sigma) as a color indicator for the reaction product. After washing with distilled water, the sections were counterstained with methyl green and mounted in Kaiser's glycerol jelly.

Immunohistochemical staining

Immunohistochemical staining for type I collagen, the type 1 PTH/PTHrP receptor (PTHR) and the calcium sensing receptor (CaSR) was performed using the avidin-biotin-peroxidase complex technique with affinity-purified goat anti-human type I collagen antibody (Southern Biotechnology Associates, Birmingham, AL, USA), affinity-purified mouse anti-PTHR antibody (Clone 3D1.1), and rabbit anti-CaSR polyclonal antibody (Abcam, Cambridge, MA, USA). Briefly, dewaxed and rehydrated paraffin-embedded sections were incubated with methanol-hydrogen peroxide (1:10) to block endogenous peroxidase activity and then washed in Tris-buffered saline (pH 7.6). The slides were then incubated with the primary antibody overnight at room temperature. After rinsing with Tris-buffered saline for 15 minutes, tissues were incubated with biotinylated secondary antibody (Sigma). Sections were then washed and incubated with the Vectastain Elite ABC reagent (Vector Laboratories) for 45 minutes. After washing, brown pigmentation was likewise produced using 3,3-diaminobenzidine (2.5 mg/mL). After washing with distilled water, the sections were counterstained with Mayer's hematoxylin, dehydrated in graded ethanol and xylene, and mounted with Biomount medium.

In situ cell death detection

Dewaxed paraffin sections were stained with an In Situ Cell Death Detection Kit (Roche, Germany). The slides were first incubated with 10 µg/mL proteinase K for 15 minutes, rinsed with phosphate buffered saline (PBS), incubated with 3% H2O2 and methanol to block the endogenous peroxidase activity, and rinsed with PBS. The slides were then incubated with terminal deoxynucleotidyl transferase mediated dUTP nick-end labeling (TUNEL) at 37°C for 1 hour. All slides were counterstained with 4′,6-diamidino-2-phenylindole (DAPI). Immunofluorescence staining was performed using the immunofluorescence marker included in the kit (using an excitation wavelength in the range of 450–500 nm and detection in the range of 515–565 nm [green]). As a negative control, the terminal transferase was omitted.

Computer-assisted image analysis

After HE staining or histochemical/immunohistochemical staining of sections from 6 mice of each genotype, images of fields were photographed with a Sony digital camera. Images of micrographs from single sections were digitally recorded using a rectangular template, and recordings were processed and analyzed using Northern Eclipse image analysis software as described.[26, 28] For determining the ALP-positive, type I collagen–positive, PTHR-positive, and CaSR-positive areas in stained newly formed bone sections, thresholds were set using green and red channels. The thresholds were determined interactively and empirically on the basis of three different images. Subsequently, this set threshold was used to automatically analyze all recorded images of all sections that were stained in the same staining session under exactly the same conditions. The ratio of the areas of cytochemical-stained or immunohistochemical-stained regions was calculated automatically by the software in each microscopic field.

Statistical analysis

Data from image analysis are presented as mean ± SEM. Statistical comparisons were made using a two-way ANOVA, and for two group comparison, Bonferroni test was used after ANOVA. A p value of <0.05 was considered significant.

Results

Effect of BMX on new trabecular bone accrual in wild-type and PTH−/− mice

To assess whether BMX produced a different effect on trabecular bone accrual in wild-type and PTH−/− mice, bone tissue development was analyzed at 5 days, and at 1, 2, and 3 weeks after BMX, by histochemical staining for total collagen and µCT. Before BMX, no bone tissue development was detected in the diaphyseal region in either wild-type or PTH−/− mice (Fig. 1A). In wild-type mice, BMX induced apparent new bone formation in the proximal tibia marrow cavity at 5 days after BMX, whereas fibroblasts filled in the distal marrow cavity. At 1 week after BMX, newly formed bone tissues were clearly evident in the marrow cavity and bone marrow cells began to be seen between the newly formed trabeculae. Most of the newly formed bone disappeared at 2 weeks after BMX and the bone marrow was regenerated. At 3 weeks after BMX, bone marrow had recovered to presurgery levels in wild-type mice, with only a few newly formed trabecular bones remaining in the bone marrow (Fig. 1A). In contrast, in PTH−/− mice, the proximal tibia marrow cavity was filled with fibroblasts at 5 days after BMX, whereas the distal marrow cavity was filled with clots and capillaries. Newly formed bone tissue began to appear in the proximal tibia marrow cavity at 1 week after BMX with fibroblasts filling in the distal marrow cavity. The new bone reached maximal levels only 2 weeks after BMX and bone marrow cells began to appear between newly formed trabeculae. In addition, more remnants of trabecular bone remained in the bone marrow of PTH−/− mice than of wild-type mice at 3 weeks after BMX (Fig. 1A). When trabecular bone volume was quantitated, the bone volume was significantly lower in PTH−/− mice than in wild-type mice at 5 days and 1 week after BMX; however, it was significantly higher at 2 and 3 weeks after BMX (Fig. 1B). The rate of appearance and quantity of new trabecular bone observed by µCT scanning images were consistent with those observed by histology in both wild-type and PTH−/− mice at 1 and 2 weeks after BMX (Fig. 1C).

Figure 1.

Effect of BMX on new trabecular bone formation in wild-type and PTH−/− mice. (A) Representative micrographs of paraffin sections of tibias from 8-week-old wild-type (WT) and PTH−/− mice stained with Sirius Red for total collagen before BMX (Cont) and at 5 days (5D), 1 week (1W), 2 weeks (2W), and 3 weeks (3W) after bone marrow ablation (BMX). Magnification, ×50. (B) Quantitation of newly formed trabecular bone volume relative to tissue volume (BV/TV, %) in diaphyseal regions. (C) Representative three-dimensional reconstructed longitudinal sections of µCT scanning images of the tibias from 8-week-old wild-type (WT) and PTH−/− mice at 1 week (1W) and 2 weeks (2W) after BMX. Each value is the mean ± SEM of determinations in 5 mice of each genotype. *p < 0.05; **p < 0.01; ***p < 0.001 compared with WT mice.

Effect of BMX on osteoblasts in wild-type and PTH−/− mice

To determine whether alterations of new trabecular bone volume induced by BMX were associated with altered osteoblastic function, osteoblast number and activity were examined at 1, 2, and 3 weeks after BMX by histology (Fig. 2A), histochemistry for ALP (Fig. 2C), and immunohistochemistry for type I collagen (Fig. 2E). At 1 week after BMX, osteoblast numbers (Fig. 2B), ALP-positive areas (Fig. 2D), and type I collagen–positive areas (Fig. 2F) were all decreased significantly in PTH−/− mice compared to wild-type mice. In contrast, all osteoblast parameters were increased significantly at 2 and 3 weeks after BMX in PTH−/− mice compared to wild-type mice. The maximum increase in osteoblast numbers, ALP-positive areas, and type I collagen–positive areas in PTH−/− mice at 2 weeks after BMX were, however, significantly lower than the corresponding increase in those parameters in wild-type mice at 1 week after BMX (Fig. 2B, D, F).

Figure 2.

Effect of BMX on osteoblasts in wild-type and PTH−/− mice. Representative micrographs of paraffin-embedded sections of tibial diaphyseal regions from 8-week-old wild-type (WT) and PTH−/− mice at 1 week (1W), 2 weeks (2W), and 3 weeks (3W) after BMX stained with (A) hematoxylin and eosin (HE), (C) histochemically for ALP, and (E) immunohistochemically for type I collagen (Col I). Magnification, ×400. (B) Osteoblast number relative to tissue area (N.Ob/T.Ar, #/mm2), (D) ALP-positive areas, and (F) Col I immunopositive areas were measured by computer-assisted image analysis. Each value is the mean ± SEM of determinations in 5 mice of each genotype. *p < 0.05; ***p < 0.001 compared with WT mice. #p < 0.05 compared with WT mice at 1W after BMX.

To assess whether alterations of osteoblast number were associated with altered apoptosis of osteoblast precursors or osteoblasts, apoptotic osteoblast precursors or osteoblasts were examined at 5 days, and at 1 and 2 weeks after BMX by TUNEL staining. The percentage of apoptotic osteoblast precursors or osteoblasts was increased at 1 week after BMX in wild-type mice, but was decreased at this time in PTH−/− mice compared to wild-type mice (Fig. 3A, B).

Figure 3.

Effect of BMX on osteoblast apoptosis in wild-type and PTH−/− mice and the recruitment of osteoprogenitor cells. (A) Representative micrographs of paraffin-embedded sections of tibial diaphyseal regions from 8-week-old wild-type (WT) and PTH−/− mice at 5 days (5D), 1 week (1W), and 2 weeks (2W) after BMX stained with DAPI for nuclei (upper panel) or TUNEL (middle panel) or merged images (third panel). Magnification, ×400. (B) The percentage of apoptotic osteoblasts relative to total cells. (C) Femurs from 1-week old wild-type mice with BMX were (D) transplanted into the back muscle of β-galactosidase transgenic mice for 1 week. (E) Representative micrograph of a Lac-Z histochemically-stained diaphyseal region section of the transplanted bone. Each value is the mean ± SEM of determinations in 5 mice of each genotype. *p < 0.05; **p < 0.01; ***p < 0.001 compared with WT mice.

To determine whether osteogenic cells induced by BMX were recruited from circulating osteoprogenitor cells, femurs from 1-week-old wild-type mice with BMX (Fig. 3C) were transplanted into the muscle of β-galactosidase–expressing transgenic mice (Fig. 3D). After 1 week, transplanted bones were harvested and recipient-derived osteogenic cells were identified in diaphyseal regions by Lac-Z histochemical staining. The results showed that Lac-Z–positive osteoblasts and fibroblastic cells were detected in the newly formed bone surface and bone marrow cavity, respectively (Fig. 3E).

To further determine whether new trabecular bone induced by BMX was associated with osteoblastic bone formation-related gene and protein expression, the expression of genes and proteins were measured by real-time RT-PCR and Western blots, respectively, in long bone diaphyseal extracts from wild-type and PTH−/− mice at 1 and 2 weeks after BMX. Consistent with the histomorphometric observations, gene expression levels of Runx2, ALP, type I collagen, and osteocalcin (Fig. 4AD), and protein expression levels of Runx2 and IGF-1 were also downregulated significantly in PTH−/− mice at 1 week after BMX, but were upregulated dramatically at 2 weeks after BMX (Fig. 4EG).

Figure 4.

Effect of BMX on the expression of genes and proteins related to osteoblastic activity and osteoclastic bone resorption in wild-type and PTH−/− mice. Real-time RT-PCR was performed on long bone diaphyseal extracts from WT and PTH−/− mice at 1 week and 2 weeks after BMX for the gene expression of (A) Runx2, (B) ALP, (C) type I collagen (Col I), and (D) osteocalcin (OCN). Messenger RNA expression is calculated as a ratio to the GAPDH mRNA level and expressed relative to levels of WT mice at 1 week after BMX. (E) Western blots of long bone diaphyseal extracts from WT and PTH−/− mice at 1 week and 2 weeks after BMX for expression of Runx2 and IGF-1. β-tubulin was used as loading control for Western blots. (F) Runx2 and (G) IGF-1 protein levels relative to β-tubulin protein level were assessed by densitometric analysis and expressed relative to levels of WT mice at 1 week after BMX. (H) Representative micrographs of sections of the tibial diaphyseal regions from WT and PTH−/− mice before BMX (Cont) and at 5 days (5D), 1 week (1W), 2 weeks (2W), and 3 weeks (3W) after BMX, stained histochemically for TRAP. (I) Osteoclast surface relative to bone surface (Oc.S/BS, %) was quantitated by computer-assisted image analysis. (J) Real-time RT-PCR was performed on long bone diaphyseal extracts from WT and PTH−/− mice at 5 days, 1 week, 2 weeks, and 3 weeks after BMX for RANKL and OPG mRNA. Messenger RNA expression is calculated as a ratio relative to the GAPDH mRNA level and expressed relative to levels of wild-type mice at 1 week after BMX. Ratio of RANKL/OPG relative mRNA levels was calculated. Each value is the mean ± SEM of determinations in 5 mice of each group. *p < 0.05; **p < 0.01; ***p < 0.001 compared with WT mice.

Effect of BMX on osteoclastic bone resorption in wild-type and PTH−/− mice

To determine whether alterations of new trabecular bone induced by BMX were associated with osteoclastic bone resorption, paraffin sections were stained histochemically for TRAP (Fig. 4H). Osteoclast surface (Fig. 4I) was determined by image analysis. Before BMX, no TRAP-positive osteoclasts were detected in the diaphyseal region in either wild-type or PTH−/− mice (Fig. 4H). In wild-type mice active osteoclasts were seen at 5 days, peaked at 1 week, and then progressively fell as the marrow cavity matured and marrow cells became evident. At 5 days after BMX, however, no active TRAP-positive osteoclasts were detectable in PTH−/− mice. The TRAP-positive osteoclast surface peaked at 2 weeks after BMX in PTH−/− mice before falling again at 3 weeks as the marrow cavity matured. The osteoclast surface was reduced significantly at all time points; ie, at 1, 2, and 3 weeks in PTH−/− mice compared with their wild-type littermates (Fig. 4H, I). We also examined alterations in the expression of genes related to bone resorption. Gene expression levels of RANKL and OPG were determined by real-time RT-PCR, and the ratio of RANKL/OPG mRNA levels was calculated. Results revealed that alterations in the ratios of mRNA levels of RANKL/OPG were consistent with those of the TRAP-positive osteoclast surfaces (Fig. 4J).

Effect of BMX on the expression of PTHrP, PTHR, and CaSR in wild-type and PTH−/− mice

To assess whether alterations of new trabecular bone formation induced by BMX were associated with those of PTHrP and PTHR expression, mRNA and proteins were extracted from long bone diaphysis at 1 and 2 weeks after BMX and real-time RT-PCR and Western blots were performed. The results showed that the expression levels of PTHrP at both mRNA and protein levels were upregulated significantly at 1 week and more dramatically at 2 weeks after BMX in PTH−/− mice relative to their wild-type littermates (Fig. 5AC). In contrast, the expression levels of PTHR protein were downregulated at 1 week, but were upregulated at 2 weeks after BMX in PTH−/− mice relative to their wild-type littermates (Fig. 5B, C). We also examined the localization and relative expression levels of PTHR and CaSR in newly formed bony tissue at 5 days, 1 week, and 2 weeks after BMX by immunohistochemical staining and image analysis. Results showed that PTHR was not only localized in osteoblast precursors and osteoblasts, but also localized in newly embedded osteocytes and their processes (Fig. 5E). CaSR was localized in osteoblast precursors and osteoblasts and some newly embedded osteocytes (Fig. 5G). Both PTHR and CaSR positive areas were decreased significantly at 5 days and 1 week after BMX, but increased significantly at 2 weeks after BMX in PTH−/− mice relative to their wild-type littermates (Fig. 5EH).

Figure 5.

Effect of BMX on the expression of PTHrP, PTHR and CaSR in wild-type and PTH−/− mice. (A) Real-time RT-PCR was performed on long bone diaphyseal extracts from WT and PTH−/− mice at 1 week and 2 weeks after the BMX for gene expression of PTHrP. Messenger RNA expression is calculated as a ratio relative to the GAPDH mRNA level and expressed relative to levels of WT mice at 1 week after BMX. (B) Western blots of long bone diaphyseal extracts from WT and PTH−/− mice at 1 week and 2 weeks after BMX for the expression of PTHrP and PTHR. β-tubulin was used as loading control for Western blots. (C) PTHrP and (D) PTHR protein levels relative to β-tubulin protein level were assessed by densitometric analysis and expressed relative to levels of WT mice at 1 week after BMX. Representative micrographs of sections of the tibial diaphyseal regions from WT and PTH−/− mice at 5 days (5D), 1 week (1W), and 2 weeks (2W) after BMX stained immunohistochemically for (E) PTHR and (G) CaSR. (F) PTHR immunopositive areas and (H) CaSR immunopositive areas were measured by computer-assisted image analysis. Each value is the mean ± SEM of determinations in 5 mice of each genotype. *p < 0.05; **p < 0.01; ***p < 0.001 compared with WT mice.

Effect of PTHrP haploinsufficiency on newly formed trabecular bone turnover induced by BMX in PTH−/− mice

Next, we examined if PTHrP haploinsufficiency affected trabecular bone turnover induced by BMX in PTH−/− mice. PTH−/−PTHrP+/− mice were generated and BMX was performed in wild-type, PTHrP+/−, PTH−/−, and PTH−/−PTHrP+/− mice. Parameters for newly formed trabecular bone turnover were compared in wild-type PTHrP+/−, PTH−/−, and PTH−/−PTHrP+/− mice at 2 weeks after BMX. The results showed that the volume of newly formed bones and type I collagen–positive areas were increased significantly, whereas osteoblast number relative to tissue area was decreased significantly in PTHrP+/− mice compared with their wild-type littermates (Fig. 6AF). The volume of newly formed bone, osteoblast numbers, and type I collagen–positive areas were decreased significantly in PTH−/−PTHrP+/− mice compared with PTH−/− mice, although these parameters were still higher than in the wild-type mice (Fig. 6AF). TRAP-positive osteoclast surface was reduced in PTHrP+/− and PTH−/− mice and further reduced in PTH−/−PTHrP+/− mice relative to their wild-type littermates at 2 weeks after BMX (Fig. 6G, H).

Figure 6.

Effect of PTHrP haploinsufficiency on newly formed trabecular bone turnover induced by BMX in PTH−/− mice. (A) Representative micrographs of paraffin sections of tibiae from 8-week-old wild-type (WT), PTHrP+/−, PTH−/−, and PTH−/−PTHrP+/− mice at 2 weeks (2W) after BMX, stained with Sirius Red for total collagen. (B) Quantitation of newly formed trabecular bone volume relative to tissue volume (BV/TV, %) in diaphyseal regions. Paraffin-embedded sections of tibial diaphyseal regions from 8-week-old wild-type (WT), PTHrP+/−, PTH−/−, and PTH−/−PTHrP+/− mice at 2 weeks (2W) after BMX, stained with (C) hematoxylin and eosin (HE), (E) immunohistochemically for type I collagen (Col-I), and (G) histochemically for TRAP. (D) Osteoblast number relative to tissue area (N.Ob/T.Ar, #/mm2), (F) Col I immunopositive areas, (H) osteoclast surface relative to bone surface (Oc.S/BS, %) were quantitated by computer-assisted image analysis. Each value is the mean ± SEM of determinations in 5 mice of each genotype. *p < 0.05; **p < 0.01; ***p < 0.001 compared with WT mice. #p < 0.05; ##p < 0.01 compared with PTH−/− mice.

Consistent with the histomorphometric observations, gene expression levels of Runx2 (Fig. 7A), ALP (Fig. 7B), type I collagen (Fig. 7C), osteocalcin (Fig. 7D), and the ratio of RANKL/OPG (Fig. 7E) were decreased significantly at 2 weeks after BMX in PTH−/−PTHrP+/− mice compared with PTH−/− mice. We also examined the alterations of PTHrP expression at both mRNA and protein levels and PTHR expression at protein levels by real-time RT-PCR and Western blots, respectively. Our results confirmed that PTHrP expression at both mRNA and protein levels and PTHR expression at protein levels were upregulated significantly at 2 weeks after BMX in PTH−/− mice, and further demonstrated that they were downregulated at 2 weeks after BMX in PTH−/−PTHrP+/− mice compared with both wild-type and PTH−/− mice (Fig. 7FI).

Figure 7.

The effect of PTHrP haploinsufficiency on the expression of genes and proteins related to newly formed trabecular bone turnover induced by BMX in PTH−/− mice. Real-time RT-PCR was performed on long bone diaphyseal extracts from WT, PTH−/−, and PTH−/−PTHrP+/− mice at 2 weeks after BMX for the gene expression of (A) Runx2, (B) ALP, (C) type I collagen (Col I), (D) osteocalcin (OCN), (E) RANKL and OPG mRNA, and (F) PTHrP. Messenger RNA expression is calculated as a ratio relative to the GAPDH mRNA level and expressed relative to levels of WT mice at 2 weeks after BMX. (G) Western blots of long bone diaphyseal extracts from WT, PTH−/−, and PTH−/−PTHrP+/− mice at 2 weeks after BMX for the expression of PTHrP and PTHR. β-tubulin was used as loading control for Western blots. (H) PTHrP and (I) PTHR protein levels relative to β-tubulin protein level were assessed by densitometric analysis and expressed relative to levels of WT mice at 2 weeks after BMX. Each value is the mean ± SEM of determinations in 5 mice of each genotype. *p < 0.05; **p < 0.01; ***p < 0.001 compared with WT mice. #p < 0.05; ##p < 0.01 compared with PTH−/− mice.

Discussion

In the present study, we employed combined genetically modified mouse models with mechanical BMX approaches to assess the effect of endogenous PTH deficiency on osteogenesis and bone turnover in vivo. Our results demonstrated that PTH-deficient mice following BMX displayed delayed osteogenesis and osteoblastic activity, with delayed osteoclastogenesis and reduced osteoclastic activity. At 5 days and 1 week after BMX, new trabecular bone volume was decreased significantly in PTH-deficient mice as a result of delayed osteogenesis and osteoblastic activity, and osteoclastogenesis was also reduced in this phase. In contrast, at 2 weeks after BMX, trabecular bone volume was increased significantly in PTH-deficient mice associated with increased osteoblastic activity.

Although previous studies have demonstrated that bone mass is increased, with a reduced bone formation rate in hypoparathyroid adult subjects,[3] the underlying mechanisms for the increased bone accrual are unknown. Our studies in adult PTH−/− animals also demonstrated an increased bone mass with a decrease in mineral apposition rate.[7] We previously reported, however, that perinatal PTH−/− mice have reduced trabecular bone compared with wild-type littermates.[6, 26, 32, 33] This reduction was not due to increased osteoclastic activity, which was actually diminished. The reduction in trabecular bone mass was, in fact, a result of decreased osteoblasts, and demonstrated the anabolic activity of endogenous PTH in bone. A number of in vitro studies have corroborated the stimulatory effect of PTH on osteoblasts. PTH has been reported to activate adhesion in bone marrow stromal precursor cells[34] and to promote proliferation and differentiation and also to inhibit apoptosis in osteoblasts and osteocytes.[35] The expression of a constitutively active PTHR in osteogenic cells resulted in an enhanced ex vivo expansion of osteogenic cells.[36] It has also been reported that some of these effects of PTH may be mediated by activation of the promoter of Runx2, an essential and early transcriptional regulator in osteoblast differentiation, leading to increased Runx2 mRNA and protein in osteoblasts. Runx2 has also been implicated in the antiapoptotic effect of PTH.[37] Previous studies have also demonstrated that IGF-1 is required for the skeletal anabolic actions of PTH.[38, 39] In the present study, our results with BMX in vivo demonstrated that endogenous PTH deficiency caused an obvious delay in osteogenesis and osteoblastic bone formation consistent with our previous findings in perinatal mice. Thus, at 1 week after BMX, trabecular bone volume, osteoblast numbers, ALP-positive areas, PTHR-positive areas, and type I collagen deposition in bone matrix were all decreased significantly, whereas the percentage of apoptotic cells was increased significantly in the diaphyseal regions of bones of PTH−/− mice compared to wild-type mice. Gene expression levels of Runx2, ALP, type I collagen and OCN, and protein expression levels of Runx2, PTHR, and IGF-1 were also downregulated significantly in bones of PTH−/− mice compared to wild-type mice at this time. We previously reported a mouse model with targeted ablation of the enzyme 25-hydroxyvitamin D-1α-hydroxylase [1α(OH)ase−/− mice], which are deficient in the active form of vitamin D, 1,25(OH)2D.[40] After weaning, the 1α(OH)ase−/− mice that were fed a diet of regular mouse chow developed hypocalcemia, hypophosphatemia, secondary hyperparathyroidism, and the skeletal abnormalities characteristic of rickets and osteomalacia, with increased osteoblast numbers and trabecular bone volume. These increases in bone volume most likely also reflected the anabolic activity of the increased endogenous PTH. Recently, we used this animal model to assess the effect of endogenous PTH on osteogenesis and bone turnover in vivo after mechanical BMX. We found that trabecular volume, and indices of osteoblast number and activity all increased significantly after BMX in the 1,25(OH)2D-deficient animals with secondary hyperparathyroidism. Results from these studies indicated that increased endogenous PTH can stimulate the recruitment of osteoprogenitor cells and the differentiation of osteoblastic cells, and finally promote bone formation. Conversely, results from our current study suggest that endogenous PTH deficiency impaired osteogenesis by inhibiting the recruitment of osteoprogenitor cells and their differentiation into active osteoblastic cells.

In contrast to the reduced osteogenesis and osteoblastic bone formation observed at the first week after BMX, we unexpectedly found that all parameters related to osteoblastic activity were increased significantly from 2 weeks after BMX in PTH−/− mice compared to wild-type mice. Thus, most new trabecular bone had been resorbed at 2 weeks after BMX in wild-type mice, whereas trabecular bone volume was increased in PTH−/− mice over that of wild-type mice at 2 weeks after BMX. This marked alteration resulted from increased osteoblastic bone activity, although osteoclasts were also increased at this time. Consequently, we sought to determine the mechanism that might be responsible for this relative increase in bone formation at 2 weeks after BMX relative to the reduction in bone formation at 1 week after BMX.

PTHrP shares important amino acid sequence homology with PTH within the biologically important NH2-terminal domain,[41, 42] which permits the NH2-terminal sequence of the two peptides to bind to a common receptor, termed the type I PTHR.[43] This interaction is known to be the molecular basis of the analogous physiological actions of PTHrP and PTH. PTH and PTHrP appear to bind to receptors on both osteoblastic stromal cells and osteoblasts,[44, 45] through which they stimulate new bone formation and, after activating the RANKL signaling system, also stimulate osteoclastic bone resorption. We previously demonstrated that mice heterozygous for PTHrP gene deletion (PTHrP+/− mice) exhibit haploinsufficiency.[44] Although these animals appear normal at birth, they develop progressive osteopenia with age. We also previously demonstrated that osteoblast-specific ablation of Pthrp in mice results in osteoporosis and impaired bone formation both in vivo and ex vivo.[46] Consequently, PTHrP seems essential for the maintenance of normal trabecular bone mass in postnatal growing animals. In previous studies we also demonstrated that PTHrP haploinsufficiency reduced trabecular bone of the PTH−/− mice to levels below those in wild-type animals by decreasing osteoprogenitor cell recruitment, enhancing osteoblast apoptosis, and diminishing bone formation.[23] Consequently, in the current study, we assessed whether alterations of new trabecular bone formation induced by BMX were associated with those of PTHrP and PTHR expression. We found that the expression levels of PTHrP at both mRNA and protein levels were upregulated significantly at 1 week and more dramatically at 2 weeks after BMX in PTH−/− mice relative to their wild-type littermates, whereas the expression levels of PTHR protein was downregulated at 1 week, but upregulated at 2 weeks after BMX in PTH−/− mice relative to their wild-type littermates. When we examined if PTHrP haploinsufficiency affected newly formed trabecular bone turnover induced by BMX in PTH−/− mice, we found that the volume of newly formed bone, osteoblast numbers, and indices of osteoblast activity were all decreased in PTH−/−PTHrP+/− mice compared to PTH−/− mice. Our findings suggest a sequence of alterations for osteogenic events induced by BMX in PTH-deficient mice, which involves delay of the recruitment of osteoprogenitor cells expressing PTHrP in the absence of PTH, a gradual increase in the action of PTHrP locally with stimulation of osteogenic cell proliferation and differentiation into osteoblasts, and subsequent enhanced osteoblastic bone formation. This sequence of events, therefore, appeared to have led to a PTHrP–induced compensatory response for reduced osteogenesis and osteoblastic bone formation caused by PTH deficiency. Interestingly, there was not good concordance of the increase of PTHrP and of the cell surface PTHR at 2 weeks after BMX. We previously reported, using a “knockin” mouse model in which the nuclear localization signal and the carboxyl-terminus of PTHrP had been deleted, that PTHrP may also exert intracrine effects in enhancing bone formation.[47] Furthermore recent large genomewide association studies in humans have associated polymorphisms of the gene encoding PTHrP (PTHLH)[48] with reduced bone mineral density. Similar associations with the gene encoding PTHR (or PTH) were not noted. Consequently the contribution of the NH2-domain of PTHrP via the PTHR relative to the contribution of the mid-region and C-terminus to the increased bone-forming actions of PTHrP observed in the current studies remain to be elucidated. It was noted that 2 weeks after BMX, in PTH−/− mice which express both alleles of PTHrP, the maximal levels of osteoblast numbers and activity and of trabecular bone volume had not quite reached the maximum levels observed in the wild-type mice. This suggests that both active peptides are required for a complete anabolic effect. Evidence in support of this was provided by our previous studies in which compound PTH−/−PTHrP−/− newborn mice were found to have reduced trabecular bone volume and osteoblast numbers relative to PTH−/− mice.[6]

We[49] and others[50] have previously shown that 1,25 dihydroxyvitamin D is a potent transcriptional inhibitor of PTHrP. The initial increase in PTHrP, observed at the 1 week time interval, may therefore have been guided by the low levels of the active form of vitamin D, which we have previously reported in PTH−/− mice.[7] In addition, the low expression levels of CaSR in osteoblastic cells at this time may have contributed to the initial increase. Thus, global haploinsufficiency of CaSR[51] or mammary-specific deletion of CaSR[52] have both been reported to increase PTHrP gene expression in the mammary gland, resulting in higher milk PTHrP concentrations. By 2 weeks after BMX, however, CaSR levels were increased in osteogenic cells and were presumably capable of transducing signals mediated by extracellular calcium. Reductions in extracellular free calcium levels in wild-type mice[52] have also been reported to increase PTHrP gene expression in the mammary gland, leading to higher milk PTHrP concentrations. The ambient hypocalcemia in the PTH−/− mice may therefore have now resulted in further osteoblastic PTHrP production in the presence of low circulating 1,25dihydroxyvitamin D.

Our previous studies in postnatal PTH−/− mice demonstrated that in the presence of a normal-calcium diet, there was decreased bone turnover associated with a marked decrease in resorption.[7] In the present study, we confirmed that osteoclastic numbers and activity were compromised. At 5 days after BMX, TRAP-positive osteoclasts were undetectable in PTH−/− mice, whereas active TRAP-positive osteoclasts had already appeared in wild-type mice at that time. The ratio of mRNA levels of RANKL/OPG and TRAP-positive osteoclast surface were reduced significantly in PTH−/− mice compared to wild-type mice at all time periods after BMX. These results suggest that impaired osteoclastogenesis in PTH-deficient mice with BMX may have been due to a reduced ratio of RANKL to OPG expressed by osteoblastic supporting cells that were delayed in their recruitment to the bone regeneration site and to a consequent delayed osteoclastic progenitor recruitment, resulting in retarded bone marrow regeneration. In the current study, we found also that the osteoclast surface was reduced at 2 weeks after BMX in PTHrP+/− mice compared with wild-type mice and was further reduced in PTH−/−PTHrP+/− mice relative to PTH−/− mice. These results are consistent with the known role of PTHrP in promoting bone resorption. In the PTH−/− mice, the peak in both RANKL/OPG expression and in TRAP-positive osteoclasts was seen at 2 weeks after BMX, when PTHrP levels were increased, consistent therefore with the action of PTHrP to enhance RANKL production by PTHR-positive preosteoblasts in the hypoparathyroid mice.[53]

In summary, PTH deficiency appears to result in a sequence of delay of osteoprogenitor cell recruitment and differentiation accompanied by a delay in osteoclastogenesis and osteoclastic bone resorption. Ultimately there appears to be compensatory increase in osteoblastic activity and osteoclastic resorption induced by upregulating PTHrP and PTHR in osteogenic cells. New trabecular bone is therefore formed followed by a marrow cavity. Results from this study therefore shed new light on the mechanisms of endogenous PTH as an anabolic agent and its interaction with endogenous PTHrP in bone.

Disclosures

All authors state that they have no conflicts of interest.

Acknowledgments

This work was supported by the National Basic Research Program of China (2012CB966902) and by the Key Project grants from the National Natural Science Foundation of China (No. 81230009) (to DM), and by a grant from the Canadian Institutes for Health Research (to DG and AK).

Authors' roles: Conceived and designed the experiments: DM. Performed the experiments: QZ, XZ, MZ, QW. Analyzed the data: QZ, XZ. Contributed reagents/materials/analysis tools: AK, DG. Wrote the paper: DM, QZ, DG.

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