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Keywords:

  • MECHANICAL STRESS;
  • BONE LOSS;
  • OXIDATIVE STRESS;
  • COPPER/ZINC SUPEROXIDE DISMUTASE;
  • VITAMIN C (VC;
  • ASCORBIC ACID)

ABSTRACT

  1. Top of page
  2. ABSTRACT
  3. Introduction
  4. Materials and Methods
  5. Results
  6. Discussion
  7. Disclosures
  8. Acknowledgments
  9. References
  10. Supporting Information

Oxidative stress contributes to the pathogenesis of age-related diseases as well as bone fragility. Our previous study demonstrated that copper/zinc superoxide dismutase (Sod1)-deficient mice exhibit the induction of intracellular reactive oxygen species (ROS) and bone fragility resulting from low-turnover bone loss and impaired collagen cross-linking (Nojiri et al. J Bone Miner Res. 2011;26:2682–94). Mechanical stress also plays an important role in the maintenance of homeostasis in bone tissue. However, the molecular links between oxidative and mechanical stresses in bone tissue have not been fully elucidated. We herein report that mechanical unloading significantly increased intracellular ROS production and the specific upregulation of Sod1 in bone tissue in a tail-suspension experiment. We also reveal that Sod1 loss exacerbated bone loss via reduced osteoblastic abilities during mechanical unloading. Interestingly, we found that the administration of an antioxidant, vitamin C, significantly attenuated bone loss during unloading. These results indicate that mechanical unloading, in part, regulates bone mass via intracellular ROS generation and the Sod1 expression, suggesting that activating Sod1 may be a preventive strategy for ameliorating mechanical unloading–induced bone loss. © 2013 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. Acknowledgments
  9. References
  10. Supporting Information

Imbalances between oxidation caused by reactive oxygen species (ROS) and reduction elicited by antioxidant systems induce intracellular oxidative stress, leading to the initiation and progression of age-related diseases, including diabetes, hypertension, atherosclerosis, osteoporosis, and neurodegenerative diseases. ROS contain several harmful species such as superoxide anion (O2), hydrogen peroxide (H2O2), and the hydroxyl radical (HO). These species are physiologically generated by mitochondrial respiration as well as cellular enzymatic reactions in response to environmental stimuli. In antioxidant systems, enzymes (dismutase, catalase, and peroxidase) and small molecules (glutathione and vitamins, etc.) detoxify ROS by converting them into nontoxic metabolites such as molecular oxygen and water.[1]

Human studies have reported that serum biomarkers of oxidative stress are upregulated in older patients with decreased bone mineral density as well as those with osteoporosis.[2, 3] Furthermore, animal studies have demonstrated chronological aging-induced bone loss associated with increased ROS production and lipid oxidation of 4-hydroxynonenel (4-HNE) in bone marrow cells in both males and females, indicating that redox imbalances caused by increased ROS production affect bone metabolism as well as other organs.[4, 5] Recently, we also revealed that a deficiency of copper/zinc superoxide dismutase (Sod1), which catalyzes intracellular O2 in the cytoplasm, is associated with increased ROS generation and severe bone fragility occurring via reductions in bone mass and impairment of bone quality in mice, suggesting that cytoplasmic ROS are key molecules of bone metabolism.[6]

Mechanical loading plays an important role in the maintenance of the musculoskeletal function. Reduced mechanical loading causes bone and skeletal muscle atrophy under several conditions, including bed rest, paralysis, and space flight.[7-10] After bed rest for 35 days, mechanical unloading decreases bone mass by 1% to 3% in young people.[11] Such reductions are equivalent to the amount of bone loss induced each year by aging in elderly people.[12] Furthermore, recovery of bone mass requires mechanical loading over a period fourfold longer than the duration of time in space or bed rest.[13, 14] In this context, preventing bone loss induced by mechanical unloading is indispensable for maintaining human quality of life, especially among the elderly. However, there is a lack of effective strategies to prevent common frailty resulting from musculoskeletal atrophy.

In a human study, Smith and colleagues reported that unloading increases the levels of urinary 8-hydroxy-2′-deoxyguanosine (8-OHdG) during long-duration space flight.[15] In addition, Rai and colleagues revealed that head-down bed rest, a valuable ground-based model of mechanical unloading conditions, increases the serum levels of 8-OHdG and malondialdehyde,[16] thus indicating that mechanical unloading induces systemic oxidative stress via ROS production in humans. However, the molecular links between bone metabolism and intracellular ROS signaling during mechanical unloading have not yet been fully elucidated.

In the present study, to clarify the pathophysiological roles of intrinsic ROS in the skeletal response to mechanical unloading, we analyzed the intracellular ROS levels and the expression levels of antioxidant enzymes in bone and bone marrow cells during mechanical unloading. Moreover, we investigated the protective effect of an antioxidant in unloading-induced bone loss.

Materials and Methods

  1. Top of page
  2. ABSTRACT
  3. Introduction
  4. Materials and Methods
  5. Results
  6. Discussion
  7. Disclosures
  8. Acknowledgments
  9. References
  10. Supporting Information

Animals

Wild-type (C57BL/6NCrSlc) and Sod1-deficient mice (Sod1−/−) were purchased from Nihon SLC (Nihon SLC, Shizuoka, Japan) and the Jackson Laboratory (Bar Harbor, ME, USA), respectively. Sod1−/− mice were backcrossed with C57BL/6NCrSlc mice five to six times. To detect osteoblasts and osteocytes in mice, we generated osteoblast- and osteocyte-specific GFP-expressing mice by crossbreeding CAG-CAT-EGFP reporter mice (Center for Animal Resources and Development, Kumamoto University, Kumamoto, Japan)[17] with Dmp1-cre mice.[18] The mice were maintained and studied according to protocols approved by the Animal Care Committee of the Tokyo Metropolitan Institute of Gerontology and Chiba University.

Mechanical unloading model

To decrease mechanical loading on the hindlimbs, the male mice (10 weeks of age) were subjected to hindlimb tail suspension. Tape was applied to the surface of the tail to set a metal clip. The end of the clip was fixed to an overhead bar, and the height of the bar was adjusted to maintain the mice at a ∼30° head-down tilt with the hindlimbs elevated above the floor of the cage. The mice were subjected to tail suspension for several days (3, 7, and 14 days). The mice that underwent normal loading were also housed individually under the same conditions, except for tail suspension, for the same duration. After tail suspension, the mice were killed using an overdose of pentobarbital.

Intracellular ROS analysis

For intracellular ROS measurement, bone marrow cells (5–10 × 105 cells/two tibias of a mouse) were collected by flushing tibias with phosphate-buffered saline using 26-G needles and stained with 5 mM of dichlorodihydrofluorescein diacetate (CM-H2DCFDA, Invitrogen, Carlsbad, CA, USA) or 10 µM of dihydroethidium (DHE, Invitrogen) dyes for 30 minutes at 37°C. The stained bone marrow cells were observed using fluorescence microscopy (Leica AF6500, Wetzlar, Germany), and the rate of positive cells stained with CM-H2DCFDA was calculated. The DCF- and DHE-stained bone marrow cells were analyzed for cell number and fluorescence intensity using a flow cytometer (ALTRA, Beckman Coulter, Brea, CA, USA). Data of average intensity were analyzed using the Expo32 software program (ALTRA) and expressed in the figure as the percent increase with respect to normal loading.

Separation of bone marrow cells

Bone marrow cells were flushed from the hindlimbs (tibia) of the mice and washed with staining medium (3% FCS) and resuspended at a concentration of 106 cells/200 µL in staining medium. To separate the bone marrow cells into individual cells, we stained the cells with the following specific antibodies: Ter119–PerCp5.5 (for erythroid cells, eBioscience, San Diego, CA, USA), CD3e-APC (for T lymphocytes, BD Biosciences, San Jose, CA, USA), CD45R-PE-Cy7 (for B lymphocytes, BD Biosciences), Gr1-PE (for granulocytes, BD Biosciences), and F4/80-Ax700 (for macrophages, BD Biosciences).

Oxidative stress markers

The serum was measured with derivatives of reactive-oxygen metabolites (d-ROM) using the Free Radical Analytical System 4 (FRAS4; H&D srl, Parma, Italy), according to the manufacturer's instructions. Protein carbonylation of flushed hindlimb tissue was analyzed with an Oxyblot Protein Oxidation Detection kit (Millipore, Billerica, MA, USA), according to the manufacturer's instructions. For immunohistochemical analyses of oxidative DNA and lipid peroxidation in bone tissue, we prepared fresh frozen sections of femora using the film method developed by Kawamoto and Shimizu.[19] Sections were stained with anti-8-OHdG and anti-4-HNE mouse monoclonal antibodies (NOF Corporation, Tokyo, Japan), followed by a goat anti-mouse biotinylated secondary antibody.

Real-time RT-PCR

To remove the bone marrow cells from bone, femur was thoroughly flushed with saline. Total RNA was extracted from bone marrow cells and flushed-femora using the Trizol reagent (Invitrogen), according to the manufacturer's instructions. cDNA was synthesized from 1 µg of total RNA using reverse transcriptase (Superscript II Preamplification System; Invitrogen). Real-time PCR was performed using an ABI Prism 7500 sequence detection system with the SYBR GREEN PCR Master Mix (Applied Biosystems, Carlsbad, CA, USA), according to the manufacturer's instructions. The detector was programmed with the following PCR conditions: 40 cycles of 15 seconds of denaturation at 95°C and 1 minute of amplification at 60°C. All reactions were performed in triplicate and normalized to the level of the housekeeping gene glyceraldehyde-3-phosphate dehydrogenase (GAPDH). The relative differences in the PCR results were calculated using the comparative cycle threshold method.[20] The following primer sets were used: Sod1 forward, 5′-GCGGTGAACCAGTTGTGTTGTC-3′ and reverse, 5′-CAGTCACATTGCCCAGGTCTCC-3′; Sod2 forward, 5′-ATGTTACAACTCAGGTCGCTCTTC-3′ and reverse, 5′-TGATAGCCTCCAGCAACTCTCC-3′; Cat forward, 5′-ACATGGTCTGGGACTTCTGG-3′ and reverse, 5′-CAAGTTTTTGATGCCCTGGT-3′; Gpx1 forward, 5′-GTCACCGTGTATGCCTT CT-3′ and reverse, 5′-CAAGTTTTTGATGCCCTGGT-3′; Runx2 forward, 5′-TGGCTTGGGTTTCAGGTTAGGG-3′ and reverse, 5′-TCGGTTTCTTAGGGTCTTGGAGTG-3′; Alp forward, 5′-GCTATCTGCCTTGCCTGTATCTG-3′ and reverse, 5′-AGGTGCTTTGGGAATCTGTGC-3′; CatK forward, 5′-GGCTGTGGAGGCGGCTAT-3′ and reverse, 5′-AGAGTCAATGCCTCCGTTCTG-3′; Gapdh forward, 5′-AGAAGGTGGTGAAGCAGGCATC-3′ and reverse, 5′-CGAAGGTGGAAGAGTGGGAGTTG-3′.

Western blot analysis

Bone marrow cells were collected from the hindlimbs of the mice and sonicated in radioimmunoprecipitation assay (RIPA) buffer containing a mixture of protease and phosphatase inhibitors (Roche Diagnostics, Mannheim, Germany) and centrifuged at 12,000 g for 30 minutes. The supernatant was assayed for the protein concentration using the DC Protein Assay Kit (BioRad, Hercules, CA, USA). Equal amounts (7.5 µg) of total protein were subjected to 15% SDS-PAGE and electroblotted onto a PVDF membrane (0.2 µm pore size, Immobilon-PSQ, Millipore). The membranes were blocked in 2% blocking reagent (ECL Advance Blocking Agent, GE Healthcare, Piscataway, NJ, USA) and probed with antibodies against Sod1 (1:1500; SOD-100; StressGen, Enzo Life Sciences, Farmingdale, NY, USA), Sod2 (1:5000; SOD-110; StressGen, Enzo Life Sciences), and Gapdh (1:2000; Sigma, St. Louis, MO, USA). The signals were detected using the ECL system (ECL plus, GE Healthcare) and a luminoimage analyzer LAS-3000 mini (Fuji Film, Tokyo, Japan).

Dual-energy X-ray absorptiometry and microcomputed tomography analysis

We measured the bone mineral density (BMD) of the isolated femora using a PIXImus instrument (Lunar Corp., Madison, WI, USA). CT scanning was performed with a ScanXmate-A090S Scanner (Comscantecno, Co., Ltd., Kanagawa, Japan). Three-dimensional microstructural image data were reconstructed and structural indices were calculated using TRI/3D-BON software (RATOC System Engineering, Kyoto, Japan). Bone morphometric analyses were performed at a region 0.3 to 0.6 mm above the distal growth plates of the femora. For cancellous bone, the bone volume/tissue volume (BV/TV) value, trabecular number (Tb. N), trabecular thickness (Tb. Th), and trabecular separation (Tb. Sp) were measured.

Histomorphometric analysis

The bone formation rates (BFRs) in the cancellous regions of the femora were measured in the undecalcified sections, as previously described.[21] Briefly, to measure mineral apposition rate (MAR), we labeled mineralized bone by calcein (Dojindo, Kumamoto, Japan) double injection. We analyzed calcein fluorescence-labeled double parallel at five or more locations in cancellous bone and calculated the average value. In the decalcified sections, tartrate-resistant acid phosphatase (TRAP)-positive multinucleated cells were counted as osteoclasts to evaluate osteoclast surface/bone surface (Oc.S/BS) and osteoclast number/bone surface (N.Oc/B.Pm).

Administration of the antioxidant vitamin C

We previously demonstrated that antioxidant vitamin C (VC) significantly attenuated oxidative stress and age-related changes, such as skin atrophy and osteoporosis, in Sod1−/− mice.[6, 22] In this context, we here investigated the protective effect of VC on unloading-induced bone loss in Sod1−/− and wild-type mice. VC (2 mmol/kg/day) or vehicle was administered intraperitoneally twice a day at 8:00 a.m. and 7:00 p.m. each day in the Sod1−/− and wild-type mice (five per group) subjected to normal loading or tail suspension. To prepare the VC, ascorbate (sodium L-ascorbate, Sigma) was dissolved immediately before use in ice-cold distilled water, as described previously.[23] After 14 days, the animals were killed to prepare the hindlimbs for assessment of bone mass. There were no significant changes in the weights of the mice during the experimental period.

Statistical analysis

The statistical analyses were performed using Student's t test for comparisons between two groups and ANOVA with Tukey's test for multiple comparisons of more than three groups. All data are expressed as the mean ± standard deviation (SD).

Results

  1. Top of page
  2. ABSTRACT
  3. Introduction
  4. Materials and Methods
  5. Results
  6. Discussion
  7. Disclosures
  8. Acknowledgments
  9. References
  10. Supporting Information

Mechanical unloading increases the intracellular ROS level in bone marrow and bone-forming cells

To investigate the pathophysiological role of ROS in reduced mechanical stimulation, we performed tail-suspension experiments using C57BL/6N mice and analyzed the intracellular ROS levels in hindlimb bone marrow cells. A ROS fluorescence analysis using CM-H2DCFDA showed that tail suspension lasting for 7 days significantly increases the rate of DCF-positive bone marrow cells compared with normal loading (Fig. 1A–D). A flow cytometer analysis also revealed that unloading significantly increases the average level of intracellular ROS in bone marrow cells for 7 days after tail suspension compared with normal loading (Fig. 1E, F). Moreover, a serum analysis showed that the level of ROS is significantly higher under conditions of reduced loading than under conditions of normal loading (Fig. 1G). To further analyze the ROS generation in cells, we separated bone marrow cells using a flow cytometer and cell surface markers. A flow cytometer analysis showed that unloading for 7 days significantly increases the intracellular ROS levels in a large population of cells, including erythroid cells, granulocytes, and B lymphocytes, in bone marrow cells (Supplemental Fig. S1A, B). Interestingly, unloading significantly decreased the percentage of erythroid cells and B lymphocytes while increasing that of granulocytes in the bone marrow cells (Supplemental Fig. S1C).

image

Figure 1. Mechanical unloading increased intracellular ROS production in bone marrow cells. The intracellular ROS levels were measured using CM-H2DCFDA dye. A fluorescence microscopy analysis of the stained bone marrow cells isolated from the hindlimbs of mice that underwent normal loading (A) and unloading for 3 (B) and 7 days (C). Scale bars = 100 µm. (D) The frequency of DCF-positive cells. (E) An analysis of fluorescence intensity of bone marrow cells using a flow cytometer. (F) The relative ROS levels in bone marrow cells obtained from mice that underwent normal loading and unloading for 3 and 7 days using a flow cytometer. n = 6–8 each group. (G) The serum levels of reactive oxygen metabolites (d-ROMs). n = 5 each group. *p < 0.05, **p < 0.01. The error bars indicate the SD.

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Next, to clarify intracellular ROS accumulation in bone-forming cells during unloading, we generated osteoblast- and osteocyte-specific GFP-expressing mice by crossbreeding CAG-CAT-EGFP reporter mice with Dmp1-cre mice. Because Dmp1-cre mice selectively express cre recombinase in mature osteoblasts and osteocytes, Dmp1-cre;CAG-CAT-EGFP double transgenic mice show GFP-labeled mature osteoblasts and osteocytes in bone tissue under a cre-loxp system. After unloading for 7 days, we measured intracellular ROS levels of Dmp1-GFP–positive cells in bone marrow cells of the double transgenic mice using a DHE staining. A flow cytometer analysis revealed that unloading significantly increases the average level of intracellular ROS in Dmp1-GFP–positive cells compared with normal loading (Fig. 2A). Interestingly, we observed that unloading also significantly enhanced intracellular ROS levels in Dmp1-GFP–negative cells (Fig. 2A), confirming the increased ROS levels of bone marrow cells (Fig. 1). These results indicated that unloading increases intracellular ROS levels in bone-forming cells, including mature osteoblasts and osteocytes. Furthermore, to evaluate the oxidation state in bone tissue, we analyzed the oxidative stress markers in bone tissue during unloading. Oxyblot analysis revealed that unloading significantly increased protein carbonylation in flushed hindlimb tissue compared with normal loading (Fig. 2B, C). Similarly, immunohistochemical staining also revealed that 8-OHdG, oxidative DNA, and 4-HNE, lipid peroxidation, were positively stained in cells on bone surface and bone tissue during unloading compared with normal loading (Fig. 2D, E). Taken together, these findings strongly indicated that mechanical unloading increased the intracellular ROS levels, resulting in redox imbalance in bone marrow cells and bone tissue.

image

Figure 2. Mechanical unloading increased intracellular ROS production in bone-forming cells and oxidative stress markers in bone tissue. (A) An analysis of fluorescence intensity of bone-forming and non-bone-forming cells using a flow cytometer. The intracellular ROS levels of Dmp1-GFP–positive and –negative cells in bone marrow cells obtained from hindlimb of Dmp1-cre;CAG-CAT-EGFP double transgenic mice that underwent normal loading and unloading for 7 days. n = 8 each group. (B) Protein carbonylation in flushed hindlimb tissue from mice underwent normal loading and unloading for 14 days. (C) Quantification of protein carbonylation in (B). n = 5 each group. *p < 0.05, ***p < 0.001. The error bars indicate the SD. (D, E) Immunohistochemical analyses of oxidative DNA and lipid peroxidation in hindlimb tissue from mice underwent normal loading and unloading for 7 days. Frozen sections of femora were stained with anti-8-OHdG (D) and anti-4-HNE (E) mouse monoclonal antibodies followed by a goat anti-mouse biotinylated secondary antibody. Arrowheads show antibody-positive osteoblast-like cells on bone surface. Scale bars = 20 µm.

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Mechanical unloading specifically upregulates the Sod1 expression in bone marrow cells and bone tissue

Next, to explore the role of the antioxidant defense system during mechanical unloading, we analyzed the expression levels of antioxidant enzymes such as Sod1, manganese superoxide dismutase (Sod2), catalase (Cat), and glutathione peroxidase 1 (Gpx1) in bone marrow cells. Real-time PCR analyses confirmed that mechanical unloading significantly downregulates the expression levels of the bone formation–related genes alkaline phosphatase (Alp) and runt-related transcription factor 2 (Runx2) and upregulates the expression level of Cathepsin K (CatK), a bone resorption marker gene (Supplemental Fig. S2A–C). Interestingly, real-time PCR analyses also showed that the Sod1 expression is specifically upregulated in bone marrow cells obtained from hindlimbs after unloading for 7 days (Fig. 3A), although the expressions of antioxidant enzymes such as Sod2, Cat, and Gpx1 are not altered during unloading (Fig. 3B–D). A Western blot analysis revealed that the Sod1 protein level, but not that of Sod2, is significantly increased in bone marrow cells after unloading for 7 days (Fig. 3E–G). Furthermore, we analyzed the expression levels of antioxidant enzymes in flushed tibia to clarify the expression pattern in bone tissue itself without bone marrow cells. As expected, the Sod1 expression is specifically upregulated in hindlimb bone during unloading for 14 days (Fig. 3H), although the expression levels of antioxidant enzymes such as Sod2, Cat, and Gpx1 are not altered during unloading (Fig. 3I–K). These data indicate that the Sod1 expression is selectively upregulated during mechanical unloading in bone marrow cells and bone tissue.

image

Figure 3. The Sod1 expression was specifically upregulated during unloading in bone marrow cells and bone tissue. Real-time PCR analyses of the mRNA expression in bone marrow cells of mice that underwent normal loading and unloading for 3 and 7 days. (A) Sod1: superoxide dismutase 1; (B) Sod2: superoxide dismutase 2; (C) Cat: catalase; (D) Gpx1: glutathione peroxidase 1. (E) A Western blot analysis of the Sod1 and Sod2 levels in bone morrow lysate obtained from the hindlimbs of mice that underwent normal loading and unloading for 7 days. The relative expressions of Sod1 (F) and Sod2 (G). n = 6–8 each group. (H–K) Real-time PCR analyses of the mRNA expression in flushed tibias of mice that underwent normal loading and unloading for 14 days. (A) Sod1: superoxide dismutase 1; (I) Sod2: superoxide dismutase 2; (J) Cat: catalase; (K) Gpx1: glutathione peroxidase 1. n = 6–10 each group. *p < 0.05. The error bars indicate the SD.

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Sod1 deficiency exacerbates bone loss during mechanical unloading

To clarify the protective role of Sod1 during mechanical unloading in bone, we compared the rates of change among bone parameters on day 14 after unloading in 10-week-old Sod1-deficient (Sod1−/−) and wild-type (Sod1+/+) male mice. A dual-energy X-ray absorptiometry (DXA) analysis revealed that loss of BMD was enhanced in the Sod1−/− mice compared with that observed in the Sod1+/+ mice during mechanical unloading (Fig. 4A). A micro-computed tomography (µCT) analysis also showed that loss of trabecular bone volume (BV/TV) was significantly exacerbated in the Sod1−/− mice compared with that observed in the Sod1+/+ mice (Fig. 4B, C; Supplemental Fig. S3A). The percent change of trabecular thickness (Tb.Th) was decreased (Fig. 4D; Supplemental Fig. S3B) and that of trabecular separation (Tb.Sp) was increased in the Sod1−/− mice compared with that observed in the Sod1+/+ mice (Fig. 4F; Supplemental Fig. S3D). The percent change of trabecular number (Tb.N) tended to decrease in the Sod1−/− mice compared with that observed in the Sod1+/+ mice (Fig. 4E; Supplemental Fig. S3C). These results indicate that Sod1 deficiency exacerbates bone loss during reduced mechanical stimulation.

image

Figure 4. Sod1 deficiency enhanced unloading-induced bone loss. A morphological study using DXA and µCT scanning of the femora of 10-week-old Sod1+/+ and Sod1−/− mice subjected to normal loading or unloading for 14 days. (A) The bone mineral density (BMD) of whole femurs was measured using DXA. (B) µCT reconstruction of the trabecular region above the distal femur growth plate. (C–F) The morphological parameters of the trabecular regions measured with µCT. The percent changes of morphological parameters in the hindlimbs of mice that underwent unloading for 14 days compared with those observed in mice that underwent normal loading: (C) bone volume fraction (BV/TV), (D) trabecular thickness (Tb.Th), (E) trabecular number (Tb.N). (F) Increased rates of trabecular separation (Tb.Sp) were observed in the hindlimbs of mice that underwent unloading for 14 days compared with those observed in mice that underwent normal loading. n = 8 each group. *p < 0.05, ***p < 0.001. The error bars indicate the SD.

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Sod1 deficiency enhances suppression of bone formation during mechanical unloading

To clarify the mechanisms underlying the exacerbation of bone loss during mechanical unloading in the Sod1−/− mice, we conducted a dynamic histomorphometric analysis of femora using a calcein double-labeling method. Sod1 deficiency significantly enhanced the percent changes of the MAR (Fig. 5A, B; Supplemental Fig. S4A) and mineralizing surface (MS/BS; Fig. 5A, C; Supplemental Fig. S4B) during mechanical unloading. The percent change of the BFR during tail suspension, calculated by multiplying the MAR by the MS/BS, was significantly increased in the Sod1−/− mice compared with that observed in the Sod1+/+ mice (Fig. 5D; Supplemental Fig. S4C).

image

Figure 5. Sod1 deficiency reduced bone formation parameters under mechanical unloading conditions. (A) Fluorescent micrographs of representative calcein labels. Scale bars = 100 µm. The percent changes of bone formation parameters in the hindlimbs of mice that underwent unloading for 14 days compared with those observed in mice that underwent normal loading: (B) mineral apposition rate (MAR), (C) mineralized surface per bone surface (MS/BS), (D) bone formation rate (BFR). n = 8 each group. *p < 0.05, **p < 0.01, ***p < 0.001. The error bars indicate the SD.

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Sod1 deficiency does not alter bone resorption parameters during mechanical unloading

To investigate the bone resorption activity in Sod1−/− mice during mechanical unloading, we performed tartrate-resistant acid phosphatase (TRAP) staining using femora under both normal loading and mechanical unloading conditions in Sod1−/− and Sod1+/+ mice. A histomorphometric analysis showed no differences in bone resorption parameters, including both osteoclast surface (Oc.S/BS; Fig. 6A, B; Supplemental Fig. S5A) and osteoclast number (N.Oc/B.Pm; Fig. 6A, C; Supplemental Fig. S5B), during mechanical unloading between the Sod1−/− and Sod1+/+ mice.

image

Figure 6. Sod1 deficiency did not alter bone resorption parameters under unloading conditions. (A) Tartrate-resistant acid phosphatases (TRAP) staining of the decalcified sections at the distal ends of the femora (red signals). Scale bars = 100 µm. (B, C) The percent changes of bone resorption parameters were observed in the hindlimbs of mice that underwent unloading for 14 days compared with those observed in mice that underwent normal loading: (B) osteoclast surface per bone surface (Oc.S/BS) and (C) osteoclast number per bone surface (N.Oc/B.Pm). n = 8 each group. The error bars indicate the SD.

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VC administration attenuates bone loss during mechanical unloading in Sod1−/− mice

Next, to clarify the involvement of enhanced ROS in the exacerbation of bone loss during mechanical unloading in the Sod1−/− mice, we administrated VC in the mice subjected to tail suspension for 14 days. A µCT analysis revealed that VC administration significantly mitigated the loss of trabecular BV/TV during mechanical unloading (Fig. 7A, B; Supplemental Fig. S6A). Similarly, VC administration alleviated the reduction of trabecular thickness (Tb.Th) and the increase of trabecular separation (Tb.Sp) and tended to attenuate the reduction of trabecular number (Tb.N) (Fig. 7C–E; Supplemental Fig. S6BD). These data indicate that the administration of VC attenuated bone loss during mechanical unloading, suggesting that intrinsic ROS accumulation was involved in the exacerbation of bone loss during unloading in loss in Sod1−/− mice.

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Figure 7. VC administration attenuated bone loss during mechanical unloading in Sod1−/− mice. A morphological study of VC administration (2 mmol/kg/day) for Sod1−/− mice using µCT scanning of the femora after normal loading or unloading for 14 days. (A) µCT reconstruction of the trabecular region above the distal femur growth plate. (B–E) The percent changes of morphological parameters in the hindlimbs of Sod1−/− mice that underwent unloading for 14 days compared with those observed in the mice that underwent normal loading. The morphological parameters of the trabecular regions measured with µCT: (B) bone volume fraction (BV/TV), (C) trabecular thickness (Tb.Th), (D) trabecular number (Tb.N), (E) trabecular separation (Tb.Sp). n = 5 each group. *p < 0.05, **p < 0.01, ***p < 0.001. The error bars indicate the SD.

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VC administration attenuated bone loss during unloading in wild-type mice

Finally, to investigate the effects of reducing the levels of intracellular ROS on bone loss during mechanical unloading, we administrated VC as an antioxidant in wild-type mice subjected to tail suspension for 14 days. Interestingly, a DXA analysis showed that daily administration of VC significantly represses the loss of BMD that occurs during mechanical unloading (Fig. 8A). A µCT analysis also revealed that VC administration significantly mitigated the loss of trabecular BV/TV (Fig. 8B, C; Supplemental Fig. S7A). Similarly, other bone morphometric parameters, including trabecular thickness (Tb.Th), number (Tb.N), and separation (Tb.Sp), were alleviated by the administration of VC (Fig. 8D–F; Supplemental Fig. S7B–D). These data strongly indicate that the administration of VC attenuates bone loss during mechanical unloading.

image

Figure 8. VC administration attenuated bone loss during unloading in wild-type mice. A morphological study of VC administration (2 mmol/kg/day) for wild-type mice using DXA and µCT scanning of the femora after normal loading or unloading for 14 days. (A) The bone mineral density (BMD) of whole femurs was measured using DXA. (B) µCT reconstruction of the trabecular region above the distal femur growth plate. (C–F) The percent changes of morphological parameters in the hindlimbs of wild-type mice that underwent unloading compared with those observed in the mice that underwent normal loading. The morphological parameters of the trabecular regions measured with µCT: (C) bone volume fraction (BV/TV), (D) trabecular thickness (Tb.Th), (E) trabecular number (Tb.N), (F) trabecular separation (Tb.Sp). n = 5 each group. *p < 0.05, **p < 0.01. The error bars indicate the SD.

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Discussion

  1. Top of page
  2. ABSTRACT
  3. Introduction
  4. Materials and Methods
  5. Results
  6. Discussion
  7. Disclosures
  8. Acknowledgments
  9. References
  10. Supporting Information

Mechanical unloading increases the intracellular ROS levels and oxidative damages in bone tissue

In the present study, we revealed that mechanical unloading results in increased levels of intracellular DCF-positive ROS in bone marrow cells (Fig. 1A–F) and intracellular DHE-positive ROS in bone-forming cells (Fig. 2A). Furthermore, we confirmed that unloading also increased oxidative damage markers, such as protein carbonylation, oxidative DNA, and lipid peroxidation, in femora (Fig. 2B–E). To our knowledge, this is the first study to report that mechanical unloading increases the intracellular ROS levels and oxidative damages in bone tissue. We also identified erythroid cells, granulocytes, B lymphocytes, and Dmp1-expressing cells that were associated with increased intracellular ROS production in response to tail suspension (Fig. 2A; Supplemental Fig. S1A, B), suggesting that these cells are mechanical-sensing cells localized in bone marrow and bone. These data suggest that the redox imbalance induced by intrinsic ROS accumulation in these cells may alter bone metabolism during unloading stimulation.

The relationship between mechanical unloading and intracellular ROS production in skeletal muscles has been discussed. Mechanical unloading in skeletal muscles causes increased intracellular ROS production and metabolic imbalances between protein synthesis and degradation, resulting in muscle atrophy.[24-26] Furthermore, alleviation of intracellular ROS production by antioxidants effectively prevents muscle atrophy induced by mechanical unloading.[24, 27-29] Interestingly, Powers and Jackson reported that excess mechanical loading caused by heavy exercise results in increased intracellular ROS production in skeletal muscles.[30] Moreover, physical exercise of moderate intensity induces appropriate ROS generation and positive adaptations in molecular signaling in muscles.[31] Taken together with our results, these findings suggest that suitable mechanical loading is required to maintain adaptable cellular ROS levels and physiological function in tissues such as bone and muscles.

The Sod1 expression is selectively upregulated in bone tissue during mechanical unloading

In the present study, the Sod1 expression was selectively upregulated in bone marrow cells and bone tissue in response to unloading (Fig. 3). With respect to muscles, Brocca and colleagues reported that unloading selectively upregulates the Sod1 expression in the soleus (a slow oxidative muscle) but not the gastrocnemius (a fast glycolytic muscle).[25, 26] The oxidative soleus muscle is more sensitive than the glycolytic gastrocnemius muscle to unloading-induced atrophy,[25, 26] suggesting that the selective expression of Sod1 plays a major role in ROS metabolism in the soleus. Chen and colleagues reported that differentiated osteoblasts show increased oxygen consumption and decreased glycolysis compared with that observed in undifferentiated osteoblasts, indicating that mature osteoblasts are similar to the soleus in terms of energy metabolism.[32] These data suggest that oxygen availability may selectively regulate the ROS levels and the Sod1 expression in these tissues during unloading.

During transcriptional regulation of the Sod1 gene, transcriptional factors such as nuclear factor kappa B (NF-κB) activator protein-1 (Ap-1) and specificity protein-1 (Sp-1) bind to transcriptional regulatory elements in the promoter region of the Sod1 gene.[33] Ishijima and colleagues reported that unloading increases the NF-κB expression in bone marrow cells.[34] Furthermore, ROS directly regulate Ap-1 and Sp-1 binding in their transcriptional regulatory elements.[35] In this context, unloading may induce selective upregulation of the Sod1 gene by activating these transcriptional factors. Further analyses are therefore needed to clarify the molecular mechanism of the selective Sod1 expression in bone under unloading conditions.

Sod1 deficiency enhances unloading-induced bone loss

In this study, we showed that Sod1 loss enhances bone loss during mechanical unloading (Fig. 4), suggesting that Sod1 protects against unloading-induced bone loss by regulating the levels of intracellular ROS. It has been reported that Sod1 deficiency causes several age-related diseases, including retinal degeneration, lacrimal gland dysfunction, fatty liver, luteal degeneration, skin thinning, and acceleration of Alzheimer's-like phenotypes.[22, 36-40] These data imply that Sod1 plays an important role in protecting against the progression of aging, based on the free-radical theory.[41] Moreover, our group previously reported that Sod1−/− mice exhibit significant bone fragility owing to low bone mass and impaired bone quality.[6] Sod1 deficiency also reduces bone formation and impairs RANKL/M-CSF signaling by reducing osteoblastic viability, thereby leading to decreased bone resorption with a reduced number of osteoclasts. On the other hand, Sod1 deficiency does not affect osteoclast viability and function. These data indicate that Sod1 is a key molecule in bone metabolism that is specifically involved in bone formation rather than bone resorption. Indeed, we revealed that Sod1 deficiency enhances reductions in the levels of bone formation parameters (MAR, MS/BS, and BFR) rather than increasing the levels of bone resorption parameters (Oc.S/BS and N.Oc/B.Pm) during unloading (Figs. 6).

Almeida and colleagues reported that oxidative stress was accumulated in bone tissue during chronological aging and by estrogen deficiency, resulting in the impairment of bone metabolism.[4] They also demonstrated that antioxidant therapy prevented estrogen deficiency–induced bone loss.[4] Taken together with our results, these findings suggest that ROS accumulation induced by several conditions, such as aging, estrogen deficiency, or SOD1 depletion, may be a pivotal regulator of bone metabolism.

Administration of vitamin C prevents bone loss during mechanical unloading

VC possesses two major biological functions: it is a scavenger of ROS such as O2 and H2O2[42] and a cofactor for collagen synthesis.[43] In this study, we revealed that VC treatment significantly suppresses unloading-induced bone loss in Sod1−/− and wild-type mice (Figs. 8). The administration of VC for 2 weeks did not increase bone mass in wild-type mice under normal loading compared with that observed with saline treatment. These results indicate that the protective effects of VC are explained by its scavenger actions against ROS rather than its actions as a cofactor for collagen synthesis. Furthermore, recent studies have reported that resveratrol and hydrogen, which exhibit multiple actions, including antioxidative stress,[44, 45] effectively prevent bone loss in rats during mechanical unloading.[46, 47] Taken together with our results, these findings suggest that antioxidants prevent bone loss by repressing enhanced ROS production during mechanical unloading.

In conclusion, we demonstrated that mechanical unloading increases both intracellular ROS production and the Sod1 expression in bone tissue including bone marrow cells. We also showed that amelioration of intracellular ROS production prevents bone loss during mechanical unloading. Regulating the redox balance by specifically activating Sod1 may prevent the bone loss that occurs during decreased mechanical stimulation.

Acknowledgments

  1. Top of page
  2. ABSTRACT
  3. Introduction
  4. Materials and Methods
  5. Results
  6. Discussion
  7. Disclosures
  8. Acknowledgments
  9. References
  10. Supporting Information

We thank Prof Lynda Bonewald (Department of Oral Biology, UMKC School of Dentistry, University of Missouri–Kansas City) for providing Dmp1-cre mice, Atsushi Furuhata (Department of Host Defense and Biochemical Researcher, Juntendo University Graduate School of Medicine), Takako Ikegami, Tomoko Ikeda (Laboratory of Molecular and Biochemical Research, Research Support Center, Juntendo University Graduate School of Medicine), Shuichi Shibuya, and Toshihiko Toda (Department of Advanced Aging Medicine, Chiba University Graduate School of Medicine) for their technical assistance, and Drs Muneaki Ishijima and Masashi Nagao (Department of Orthopaedics, Juntendo University) for their helpful discussions.

This study was supported by the Program for the Promotion of Basic Research Activities for Innovative Biosciences (TS) and by a Grant-in-Aid for Young Scientists (B; No. 21791415 and 24791569 to HN, and No. 21791416 and 24791568 to YS) from the Ministry of Education, Science, Culture, Sports, and Technology.

Authors' roles: DM carried out major biochemical and biological experiments and wrote the article. KKo, YO, and MK carried out mouse work. KW carried out the FACS analyses. DM, HN, YS, and TS designed the experiments. DM, HN, YS, YA, TT, KKa, and TS discussed the hypothesis and interpreted the data. HN, YS, TT, and KKa edited the article. TS coordinated and directed the project and wrote the manuscript.

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  3. Introduction
  4. Materials and Methods
  5. Results
  6. Discussion
  7. Disclosures
  8. Acknowledgments
  9. References
  10. Supporting Information
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Supporting Information

  1. Top of page
  2. ABSTRACT
  3. Introduction
  4. Materials and Methods
  5. Results
  6. Discussion
  7. Disclosures
  8. Acknowledgments
  9. References
  10. Supporting Information

Additional Supporting Information may be found in the online version of this article.

FilenameFormatSizeDescription
jbmr1981-sm-0001-SuppFigs-S1.pdf273K

Supplemental Figure S1. Unloading increased ROS production in erythroid cells, granulocytes and B lymphocytes. Bone marrow cells were separated using a flow cytometer and cell surface markers: anti-Ter119 (erythroid cells), CD3e (T lymphocytes), CD45R (B lymphocytes), Gr1 (granulocytes) and F4/80 (macrophages) antibodies. (A) Analyses of fluorescence intensity of the cells using a flow cytometer. (B) The relative fluorescence intensity of the cells. Normal loads are shown as 100% in the bar graphs. (C) The total percentage of the cell number with or without unloading. n = 5 each group. *P <0.05, ***P <0.001. The error bars indicate the SD.

Supplemental Figure S2. Real-time PCR analyses of the mRNA expression in bone marrow cells of mice that underwent normal loading and unloading for 3 and 7 days. (A) Alp: alkaline phosphatase, (B) Runx2: runt-related gene 2, (C) CatK: cathepsin K. n=6–8 each group. *P <0.05. The error bars indicate the SD.

Supplemental Figure S3. Morphological parameters of μCT scanning of the femora of 10-week-old Sod1+/+ and Sod1/ mice subjected to normal loading or unloading for 14 days: (A) bone volume fraction (BV/TV, %), (B) trabecular thickness (Tb.Th, μm), (C) trabecular number (Tb.N, 1/mm), (D) trabecular separation (Tb.Sp, μm). n = 8 each group. The error bars indicate the SD.

Supplemental Figure S4. Bone formation parameters in the hind-limbs of Sod1+/+ and Sod1/ mice that underwent unloading for 14 days compared with those observed in mice that underwent normal loading: (A) mineral apposition rate (MAR, μm/day), (B) mineralized surface per bone surface (MS/BS, %), (C) bone formation rate (BFR, μm3/μm2/day). n = 8 each group. The error bars indicate the SD.

Supplemental Figure S5. Bone resorption parameters in the hind-limbs of Sod1+/+ and Sod1/ mice that underwent unloading for 14 days compared with those observed in mice that underwent normal loading: (A) osteoclast surface per bone surface (Oc.S/BS, %) and (B) osteoclast number per bone surface (N.Oc/B.Pm, 1/mm). n = 8 each group. The error bars indicate the SD.

Supplemental Figure S6. Morphological parameters of μCT scanning of the femora of 10-week-old Sod1/ mice subjected to normal loading or unloading for 14 days with administration of vehicle and VC: (A) bone volume fraction (BV/TV, %), (B) trabecular thickness (Tb.Th, μm), (C) trabecular number (Tb.N, 1/mm), (D) trabecular separation (Tb.Sp, μm). n = 5 each group. The error bars indicate the SD.

Supplemental Figure S7. Morphological parameters of μCT scanning of the femora of 10-week-old wild-type mice subjected to normal loading or unloading for 14 days with administration of vehicle and VC: (A) bone volume fraction (BV/TV, %), (B) trabecular thickness (Tb.Th, μm), (C) trabecular number (Tb.N, 1/mm), (D) trabecular separation (Tb.Sp, μm). n = 5 each group. The error bars indicate the SD.

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