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Bone Research Group, Institute of Medical Sciences, Foresterhill, University of Aberdeen, Aberdeen AB25 2ZD, United Kingdom
Address all correspondence and requests for reprints to: Dr. R. J. van’t Hof, Bone Research Group, Institute of Medical Sciences, Department of Medicine and Therapeutics, Foresterhill, Aberdeen AB25 2ZD, United Kingdom. E-mail: r.hof{at}abdn.ac.uk.
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Introduction |
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Most investigators have failed to detect evidence of nNOS expression in normal adult bone or bone cell cultures (13, 14), although the nNOS isoform has been detected during skeletal development (15) and during fracture healing, where nNOS is present in chondral and fibrochondral regions within fracture callus (16).
Until now, the role of nNOS in bone metabolism has been little studied. The importance of NO as a regulator of bone cell function, however (17), coupled with recent evidence that has shown that the central nervous system plays a role in regulating bone turnover and bone mass (18), led us to investigate the role of nNOS in bone metabolism by studying various aspects of bone metabolism in mice with targeted inactivation of the nNOS gene (12).
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Materials and Methods |
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B ligand (RANKL) was obtained from Insight Biotechnology (Middlesex, UK); human recombinant macrophage colony-stimulating factor (M-CSF) was obtained from R&D Systems (Abingdon, UK); and human recombinant TNF
was obtained from Peprotech (London, UK). -MEM, fetal calf serum (FCS), penicillin, and streptomycin were obtained from Life Technologies, Inc. (Paisley, UK), and tissue culture plates were obtained from Costar (Cambridge, MA). All other reagents were from Sigma (Poole, Dorset, UK), unless otherwise indicated in the text.
Animals
Mice with targeted inactivation of the nNOS gene and wild-type controls were obtained from the Jackson Laboratories (Bar Harbor, ME) (strain name: B6;129S4-Nos1tm1Plh; stock no. 002633). The targeting strategy used to generate these mice has been described previously (12, 19). The animals were housed in a designated animal facility and routinely maintained on a 12-h light, 12-h dark cycle and given ad libitum access to food and water. The tibial bones from 10-wk-old female mice (nine nNOS wild-type mice and 12 nNOS KO mice) were dissected and used for bone mineral and histomorphometric analysis (see Bone mineral measurements and Bone histomorphometry).
Bone mineral measurements
Full-body bone mineral density (BMD) and bone mineral content (BMC) were determined by dual-energy x-ray absorptiometry scanning using a PIXImus small animal scanner (GE Medical, Bedford, UK) according to the manufacturer’s instructions.
In addition, volumetric BMC and BMD were measured ex vivo using peripheral quantitative computed tomography at the left proximal tibial metaphysis using an XCT Research M bone densitometer with a voxel size of 70 µm and analysis software version 5.1.4. (Stratec Medizintechnik, Pforzheim, Germany). Daily quality assurance measurements were performed using a plexicoated polyvinyl chloride-fluorinated hydrocarbon phantom (Stratec) according to the manufacturer’s instructions. Scans were performed at the proximal tibial metaphysis 0.9 mm distal to the growth plate, as previously described (2). The precision was 1.19% for total BMD, 3.53% for trabecular BMD, and 1.04% for cortical BMD.
Bone histomorphometry
Bone histomorphometry was performed on left tibiae. The bones were dissected free of soft tissues, fixed in 4% buffered formalin/saline (pH 7.4), and embedded in methyl methacrylate. Longitudinal sections (4 µm) were then prepared and stained with von Kossa and counterstained with Paragon. Histomorphometric measurements were made on sections of the proximal metaphysis distal to the epiphyseal growth plate using a x10 objective lens on a Zeiss Axioskop (Carl Zeiss, Welwyn Garden City, UK) coupled to a Progress C14 digital camera (ISS, Manchester, UK) using a 1 x C-mount adapter. The camera was connected to an image analysis system running in-house-designed software developed using Aphelion ActiveX Objects (Adcis SA, Hérouville-Saint-Clair, France). Bone histomorphometric variables were expressed according to the guidelines of the American Society of Bone and Mineral Research Nomenclature Committee (20).
The mineral apposition rate of bone was assessed on unstained sections using fluorescent microscopy in tibiae obtained from 10-wk-old mice that had received ip injections of 20 mg/kg calcein green (Sigma) 7 and 2 d before they were killed.
Immunostaining for nNOS
Immunostaining was performed on primary osteoblasts grown on 13-mm-diameter collagen-coated (0.15 mg/ml) coverslips and on 5-µm cryosections of fresh mouse brain. Specimens were fixed with 1:1 acetone-methanol for 1 min, washed in PBS, and permeabilized for 5 min with Triton X-100 (0.5% in PBS) at room temperature. Nonspecific binding was blocked by a 20-min incubation with 20% FCS in PBS before incubation with anti-nNOS primary antibody (2 µg/ml, anti-nNOS Clone 16; BD Transduction Laboratories, Mississauga, Ontario, Canada) at room temperature for 30 min. Samples were washed three times in PBS for 5 min and subsequently incubated with Alexa Fluor 594 goat antimouse (Molecular Probes, Eugene, OR) at room temperature for 30 min, followed by three more washes in PBS, and mounted in 80% glycerol in PBS. Specimens were examined using a Zeiss Axioskop fluorescence microscope (Carl Zeiss) with a x40 oil objective lens, and images were captured using an Optronics CCD camera (Optronics, Goleta, CA).
Detection of nNOS by RT-PCR
Total RNA was isolated from primary osteoblast cultures and whole mouse brain homogenates using Trizol as described by Helfrich et al. (14). Subsequently, 5 µg of RNA was reverse transcribed using Superscript Reverse Transcriptase (Life Technologies, Inc., Gaithersburg, MD) using a random hexamer as primer, and the resulting cDNA was taken up in 100 µl of sterile water and used to detect nNOS and ß-actin mRNA by PCR. PCRs were performed using a MJ Research Opticon2 Real Time PCR machine (MJ Research, South San Francisco, CA) using the Finnzymes DyNAmo SYBR Green kit (GRI, Braintree, UK) according to the manufacturer’s instructions. The primers used were as follows: nNOS forward, AGC ACC TAC CAG CTC AAG GA; nNOS reverse, ATA GTG ATG GCC GAC CTG AG; ß-actin forward, TCG TGG GGC GCC CCA GGA CC; and ß-actin reverse, GAA ATC GTG CGT GAC ATT AAG GAG. The cycling protocol consisted of an initial step of 94 C for 10 min, followed by 40 cycles of 30 sec at 94 C, 30 sec annealing, 1 min at 72 C, 10 sec at 82 C to melt any primer dimers, and a read of the SYBR Green levels. The annealing temperatures for the nNOS primers and ß-actin primers were 65 and 60 C, respectively. A serial dilution of the whole-brain cDNA was used as a standard curve. The expression of nNOS was calculated relative to this standard and corrected using the ß-actin levels as an internal control. For experiments investigating the effects of cytokines and calciotropic hormones on nNOS expression, osteoblast cultures were treated overnight with 1,25-dihydroxyvitamin D3 (10 nM), parathyroid hormone (1–34 fragment), IL-1 (105 U/ml), or TNF (50 ng/ml) before isolating RNA.
Osteoblast isolation and culture
Osteoblasts were isolated from the calvarial bones of 2-d-old mice by sequential collagenase digestion as described by van’t Hof (21) and cultured in -MEM supplemented with 10% FCS and penicillin/streptomycin cocktail at 37 C in 5% CO2. Osteoblasts were grown overnight, harvested by trypsin digestion, seeded in 96-well plates at 4 x 103 cells/well in 100 µl -MEM supplemented with 10% FCS, penicillin, and streptomycin, and incubated for up to 72 h. After 24, 48, and 72 h of culture, cell numbers were determined by adding 10 µl Alamar Blue reagent (Biosource, Camarillo, CA) per well, incubating the cells for a further 3 h, and measuring viability as fluorescence (excitation 530 nm; emission 590 nm) using a Labtech FL600 plate fluorometer (Biotech Instruments, Winooski, VT).
Alkaline phosphatase (ALP) activity was assessed by measuring the conversion of p-nitrophenyl phosphate to p-nitrophenyl using a Labtech FL600 plate spectrometer. Briefly, after assessing the cell number by the Alamar Blue assay, the cell layer was resuspended in 150 µl of lysis buffer (1 M diethanolamine, 1 mM MgCl2, and 0.05% Triton X-100), and the lysates were stored at –20 C. Fifty microliters of sample and 50 µl of substrate solution (20 mM paranitrophenol phosphate in lysis buffer) were pipetted into 96-well plates, and absorption was measured at 414 nm every 3 min at 37 C.
For mineralization assays, osteoblasts were seeded in six-well plates at 1 x 105 cells/well in 2 ml of -MEM supplemented with 10% FCS, penicillin, streptomycin, 2 mM glutamine, 10 mM ß-glycerophosphate, and 50 µg/ml L-ascorbic acid. Medium was changed every 3 d, whereas L-ascorbic acid was added daily because of its short half-life. Osteoblasts were cultured for 10, 14, and 17 d. At the end of the culture, osteoblasts were washed five times with PBS and fixed in 70% cold ethanol for 1 h, and mineralized nodules were detected by Alizarin red staining. Briefly, the cultures were washed in deionized water, stained with 40 mM Alizarin red S (pH 4.2) for 20 min at room temperature on an orbital rotator, washed four times with deionized water, and analyzed.
Osteoclast generation
Bone marrow was flushed out of the femora and tibiae of adult mice using Hanks’ balanced salt solution. The resulting cell suspension was spun down at 300 x g for 3 min, resuspended in 10 ml of culture medium (-MEM supplemented with 10% FCS and penicillin/streptomycin), and cultured in one 10-cm petri dish per mouse for 3 d at 37 C in the presence of 100 ng/ml M-CSF. Subsequently, the petri dish was washed three times with PBS, and the adhered cells were harvested using trypsin digestion. The cells were resuspended in culture medium supplemented with 100 ng/ml RANKL and 25 ng/ml M-CSF at a density of 4 x 104 cells/ml, and 125 µl/well of this suspension was seeded onto dentine slices in a 96-well plate. The cells were cultured at 37 C in 5% CO2 for 7 more days, with medium changes after 2 and 4 d. In experiments where cultures were stimulated by TNF, 50 ng/ml of human recombinant TNF (Peprotech) or vehicle (PBS) was present during the last 3 d of culture. At the end of the culture period, osteoclasts were identified by staining for tartrate-resistant acid phosphatase essentially as described by van’t Hof et al. (22). Briefly, dentine slices with adherent cells were fixed in 4% paraformaldehyde, washed with PBS, and incubated with naphthol-ASBI phosphate, pararosanilin, and 30 mM sodium tartrate in acetate buffer at 37 C for 45 min. Tartrate-resistant acid phosphatase-positive cells with three or more nuclei were considered to be osteoclasts. After counting adherent osteoclasts, dentine slices were immersed in 20% sodium hypochlorite to remove the cell layer. Resorption pits were visualized by reflected light microscopy and a Diagnostic Instruments Insight camera (Diagnostic Instruments, Inc., Sterling Heights, MI) coupled to an Image Analysis system (ADCIS, Herouville-Saint-Clair, France). The area resorbed was quantified by Image Analysis using custom software developed using Aphelion ActiveX objects (Adcis) (21).
Statistical analyses
Statistical analyses were performed using SPSS for Windows version 11 (SPSS Inc., Chicago, IL). Significant differences between groups were determined by ANOVA. If more than two groups were compared, significant differences were determined using the Bonferroni correction. All data are presented as means ± SEM, unless stated otherwise. P < 0.05 was considered significant.
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Results |
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). A similar increase was observed in the proximal tibia using peripheral quantitative computed tomography (9.6% in nNOS KO mice and 8.4% in wild-type controls; Fig. 1B). The difference in BMD was particularly pronounced in the trabecular compartment, where BMD was 29% higher in nNOS KO mice than in wild-type controls, with a smaller (3%) increase in the cortical BMD (Fig. 1B). The nNOS KO mice also displayed significant increases in cortical area (7.3 ± 1.6%, P < 0.01), cortical content (10.4 ± 1.5%, P < 0.001), and cortical width (8.4 ± 0.2%, P < 0.01) when compared with control animals. There were no differences in overall length and thickness of the bones (data not shown).
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). Indices of bone formation and bone resorption were also significantly lower in the nNOS KO mice compared with controls. Osteoblast surface and osteoid surface were significantly reduced to between 20 and 30% of the values noted in wild-type mice, whereas indices of bone resorption were reduced by between 30 and 70% (Table 1). No significant differences were observed for osteoid width and mineral apposition rate (Table 1). These differences in BMD and bone histomorphometry were readily apparent on low-power photomicrographs of the tibial metaphysis, where we observed also that the trabecular bone tended to extend deeper into the metaphysis in the nNOS KO mice than in the controls (Fig. 2).
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) and were also able to detect nNOS mRNA in osteoblasts using RT-PCR (Fig. 3E). However, both mRNA and immunolabeling intensities appeared to be much less than those observed in brain cells. When we quantified the levels of nNOS mRNA using real-time PCR, we found the levels in osteoblasts to be 1.4 ± 1.1% of a whole-brain RNA sample used as positive control, and these levels were not affected by overnight treatment of the cells with PTH, vitamin D3, or IL-1 (data not shown).
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) or in the ability of these cells to form mineralized bone nodules (Fig. 5). However, ALP levels were significantly higher in nNOS KO cells under basal and PTH-stimulated conditions. Surprisingly, we also found that RANKL- and M-CSF-induced osteoclast formation was increased in bone marrow cultures derived from nNOS KO mice, both in the absence and presence of TNF stimulation (Fig. 4C).
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Discussion |
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The mechanisms responsible for the reduction in bone turnover and increased bone mass in nNOS KO mice are unclear at present. Expression of nNOS has been detected in osteoblasts and osteoclasts during bone development in rats (15) and during endochondral bone formation in healing fracture callus (16), which raises the possibility that expression of nNOS during fetal life could play a role in regulating skeletal development. Although this could partly explain the difference in BMD between nNOS KO mice and controls, it is unlikely to explain the reduction in bone turnover in adult mice. Furthermore, a local effect of nNOS on bone cell activity seems unlikely because the nNOS isoform is expressed at very low levels by bone cells. The in vitro studies of osteoblast proliferation and bone nodule formation that we conducted also argue against a cell-autonomous effect of nNOS because cell growth was almost identical in nNOS KO and wild-type cultures, and no qualitative differences in bone nodule formation were observed. Surprisingly, however, ALP activity was increased in nNOS KO osteoblasts when compared with wild-type osteoblasts, as was RANKL- and TNF-stimulated osteoclast formation. A possible explanation for the suppressed bone turnover that we observed in vivo and the normal or increased activity of nNOS KO cells that we observed in vitro is that nNOS acts through a systemic endocrine or neurogenic pathway to regulate bone turnover. Two scenarios can be envisaged, one in which nNOS stimulates the systemic release of factors that enhance bone turnover and another in which nNOS blocks the release of factors that inhibit bone turnover. The observation that ALP activity was increased and osteoclast formation enhanced in nNOS KO-derived cells in vitro fits best with the notion that there is a non-cell-autonomous, systemic inhibition of bone turnover in nNOS KO mice and that the cells are released from that inhibition when cultured in vitro. Of course, we cannot exclude the possibility that the low levels of nNOS that are expressed by bone cells exert direct effects on osteoclast and osteoblast activity, in an opposite direction to the effects that are observed in vivo.
Because nNOS is strongly expressed in the central nervous system, it is tempting to speculate that nNOS might influence bone metabolism by a neurogenic relay, as has been suggested for the effects of leptin (26, 27, 28) and neuropeptide Y (29) on bone metabolism. It is unlikely that the leptin signaling pathway is involved because leptin inhibits nNOS mRNA expression in the hypothalamus (30) and, at the same time, reduces bone mass (26). Alterations in circulating levels of sex hormones are also unlikely because the strain of mice we studied has been reported to have normal fertility (11). However, a different strain of nNOS KO mice created by deletion of the heme-binding region in exon 6 showed evidence of hypogonadism (31), which if anything, would be expected to reduce bone mass. High levels of nNOS expression are also observed in skeletal muscle, but an effect mediated through skeletal muscle seems unlikely because previous investigators have found no evidence of muscle pathology in the same strain of nNOS KO mice as we studied here (32).
In conclusion, this study has shown that nNOS KO mice have increased bone mass and decreased bone turnover, indicating that the nNOS isoform plays an important role in the regulation of bone mass and bone turnover in mice. The low levels of nNOS expression in bone and the contrast between the suppressed bone turnover in vivo and the normal or increased activity of nNOS KO-derived bone cells in vitro indicate that the actions of nNOS on bone remodeling are not cell autonomous and probably are mediated by a hitherto unidentified endocrine pathway or neurogenic relay that regulates bone turnover. Further studies are now warranted to identify and characterize this pathway and to define the underlying mechanisms by which nNOS regulates bone turnover.
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Acknowledgments |
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Footnotes |
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Abbreviations: ALP, Alkaline phosphatase; BMC, bone mineral content; BMD, bone mineral density; eNOS, endothelial nitric oxide synthase; FCS, fetal calf serum; iNOS, inducible nitric oxide synthase; KO, knockout; M-CSF, macrophage colony-stimulating factor; nNOS, neuronal nitric oxide synthase; NO, nitric oxide; NOS, nitric oxide synthase; RANKL, receptor activator of nuclear factor B ligand.
Received February 17, 2004.
Accepted for publication July 30, 2004.
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