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Endocrinology, doi:10.1210/en.2006-1404
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Endocrinology Vol. 148, No. 6 2613-2621
Copyright © 2007 by The Endocrine Society

Altered Ovarian Function Affects Skeletal Homeostasis Independent of the Action of Follicle-Stimulating Hormone

Jianjun Gao, Rashmi Tiwari-Pandey, Rana Samadfam, Yinzhi Yang, Dengshun Miao, Andrew C. Karaplis, M. Ram Sairam and David Goltzman

Calcium Research Laboratory and Department of Medicine (J.G., R.S., D.M., D.G.), McGill University Health Centre and McGill University, Montreal, Quebec, Canada H3A 1A1; Molecular Reproduction Research Laboratory (R.T.-P., Y.Y., M.R.S.), Clinical Research Institute of Montreal, Montreal, Quebec, Canada H2W IR7; and Lady Davis Institute for Medical Research and Department of Medicine (A.C.K.), Jewish General Hospital and McGill University, Montreal, Quebec, Canada H3T IE2

Address all correspondence and requests for reprints to: Dr. David Goltzman, Calcium Research Laboratory, McGill University Health Centre, 687 Pine Avenue West, Room H4.67, Montreal, Quebec, Canada H3A 1A1. E-mail: david.goltzman{at}mcgill.ca.


   Abstract
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 References
 
Osteoporosis is a leading public health problem. Although a major cause in women is thought to be a decline in estrogen, it has recently been proposed that FSH or follitropin is required for osteoporotic bone loss. We examined the FSH receptor null mouse (FORKO mouse) to determine whether altered ovarian function could induce bone loss independent of FSH action. By 3 months of age, FORKO mice developed age-dependent declines in bone mineral density and trabecular bone volume of the lumbar spine and femur, which could be partly reversed by ovarian transplantation. Bilateral ovariectomy reduced elevated circulating testosterone levels in FORKO mice and decreased bone mass to levels indistinguishable from those in ovariectomized wild-type controls. Androgen receptor blockade and especially aromatase inhibition each produced bone volume reductions in the FORKO mouse. The results indicate that ovarian secretory products, notably estrogen, and peripheral conversion of ovarian androgen to estrogen can alter bone homeostasis independent of any bone resorptive action of FSH.


   Introduction
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 References
 
DIMINISHED OVARIAN FUNCTION at the time of menopause has long been known to be accompanied by accelerated loss of bone leading to osteoporosis (1, 2). In addition, decreased ovarian function in premenopausal women including adolescents has also been reported to cause osteoporosis (3, 4, 5). Loss of postpubertal ovarian function is associated with decreased inhibition of hypothalamic and pituitary function due to loss of ovarian sex steroids and inhibins, notably inhibin B, resulting in increased circulating levels of follitropin or FSH and LH (6, 7). The assumption has been that the reduction in circulating estrogen facilitates bone loss; however, other ovarian products such as inhibin have also been associated with a role in modulating bone turnover (8). Furthermore increased FSH levels per se have recently been suggested to be required for enhancing osteoclastic bone resorption (9). Estrogen administration for postmenopausal and premenopausal osteoporosis occurring as a result of ovarian failure has been an important mode of therapy (3, 10, 11), and epidemiological studies have confirmed estrogen-induced reductions of osteoporotic fractures in patients with osteoporosis (12). Although estrogen receptor agonists have had major utility in reducing osteoporotic fractures (12, 13), such agonists may (14) or may not (15, 16) have had significant effects on suppression of pituitary gonadotropins. Nevertheless, it may be difficult to distinguish the direct effects of estrogen per se on bone turnover from the effects of reductions in FSH that may be produced by this treatment.

A murine model with targeted ablation of the gene encoding the FSH receptor (the follitropin receptor knockout or FORKO mouse) has provided an important model to attempt to define the role of ovarian vs. nonovarian effects on bone turnover (17). The FORKO mouse fails to develop mature ovarian follicles including granulosa cells and has minimal detectable circulating levels of estrogens. As a result of diminished feedback inhibition, both FSH and LH levels increase (18). As a consequence of the increased circulating LH, androgen production from ovarian theca cells is increased resulting in enhanced testosterone production. Inasmuch as the function of ovarian aromatase (the enzyme converting testosterone to 17ß-estradiol) may be diminished by the resistance to FSH action (19) in the FORKO mouse, ovarian testosterone levels may also accumulate due to decreased metabolism, which may contribute to the increased circulating concentrations of this androgen.

In previous studies, it has been reported that FORKO mice develop an osteoporotic phenotype, including spinal kyphosis by about 4 months of age and vertebral compression at approximately 12 months of age (20). In contrast, more recent studies have indicated that bone loss does not occur in this model (9). In view of the fact that FSH receptors and stimulatory effects in bone have been localized to osteoclasts, the absence of bone loss has been ascribed to the loss of the bone resorptive function of FSH in the FORKO mouse (9). To explore these issues in more detail, we have examined parameters of skeletal function longitudinally in the FORKO mouse, we have assessed the effect of ovariectomy (OVX) on bone metabolism, and we have investigated the role of ovarian androgens on the skeleton of these animals.


   Materials and Methods
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 References
 
Animal studies
The FORKO mice were established as previously described (17) by breeding 129T2/SV EmsJ FSH receptor +/– males and females. These breeding pairs provided +/+ [wild-type (WT)] and –/– (FORKO) female litter mates that allowed a direct comparison. Genotyping by PCR on DNA isolated from tails was as previously described (20) using the FSH receptor primer sequences: 5'-CATGTCAGTAGTACATTAGAG-3' and 5'-AGTTCAATGGCGTTCCG-3' (634 bp) and the Neo+ gene primer sequences: 5'-AGGGACTGGCTGCTATTG-3' and 5'-AGAAAAGCGGCCATTT-TC-3' (348 bp). Transplantation of WT ovaries to FORKO mice was performed at 6 wk of age under anesthesia as previously described (21). Bilateral OVX or sham surgery was performed under anesthesia at 3 wk of age. The androgen receptor antagonist, flutamide (12 mg/pellet), or placebo were administered as slow-release pellets (Innovative Research of America, Sarasota, FL) inserted sc for 60 d. The selective aromatase inhibitor, letrozole (Novartis Pharmaceuticals Canada Inc., Dorval, Quebec, Canada) was dissolved in PBS and was administered orally, as a loading dose of 25 µg/d followed by 12.5 µg/d to FORKO mice from 1–3 months of age. Body weight was unaltered by either flutamide or letrozole administration. In all experiments, tissues were collected at the time the mice were killed for further study. Serum samples were collected, under anesthesia by the intracardiac route just before the mice were killed. Mutant mice and control litter mates were maintained in a virus- and parasite-free barrier facility and exposed to a 12-h light, 12-h dark cycle. All experiments involving animals were performed according to institutionally approved and current animal care guidelines.

Hormone assays
Serum total testosterone, free testosterone, estradiol, and LH were measured with standard techniques in a single assay. Serum total testosterone was measured with a DSL-4100 Testosterone RIA kit (DSL Inc., Webster, TX), which has a detection limit of 0.05 ng/ml and an intraassay coefficient of variation of 8%, and free testosterone was measured with a TKTF2 free testosterone RIA kit (DPC Inc., Los Angeles, CA), which has a 0.55–50 pg/ml calibration range and 0.15 pg/ml analytical sensitivity and less than 8% intraassay coefficient of variation. Serum estradiol was measured with a TKE22 Estradiol RIA kit (DPC Inc.), which has a calibration range of 20–3600 pg/ml, analytical sensitivity of 8 pg/ml, and an intraassay coefficient of variation of less than 8%. Serum LH was measured with an IKLH IRMA kit (DPC Inc.), which has a detection limit of 0.15 mIU/ml and an intraassay coefficient of variation of less than 4%. All samples for each hormone were measured in a single assay.

Bone mineral density (BMD) analysis
Densitometry was performed by PIXImus on the right femur and lumbar vertebrae (L4–L6) under anesthetization as previously described (22). In brief, mice were anesthetized with an im mixture of ketamine (100 mg/kg) and xylazine (0.1 mg/kg) in PBS and placed prone on the platform of a PIXImus densitometer (software version 1.46.007; Lunar Corp., Madison, WI) for BMD measurements of the right femur and lumbar vertebrae (L4–L6) according to the manufacturer’s instruction. In some experiments, the variability in measurements was examined by repeating scans after repositioning the animals. Percent CV of BMD for the repeated scans was 1–3% at all skeletal sites examined.

Microcomputed tomography (microCT)
Femurs obtained from mice were dissected free of soft tissue, fixed overnight in 70% ethanol, and analyzed by microCT with a SkyScan 1072 scanner and associated analysis software (SkyScan, Antwerp, Belgium) as described (23). Scans were performed on the distal part of femur. Image acquisition was performed at 100 kV and 98 µA with a 0.9° rotation between frames. Thresholding was applied to the images to segment the bone from the background. The distal 3.5 mm of the femora was reconstructed using two-dimensional data from scanned slices with the 3D Creator software supplied with the instrument. The trabecular bone region of interest was drawn to include all cancellous bone in the metaphysis, and three-dimensional analysis was performed to calculate bone volume (BV)/tissue volume (TV). The resolution of the microCT images is 18.2 µm.

Histological examination
Hematoxylin and eosin (H&E) and histochemical staining for tartrate-resistant acid phosphatase (TRAP) were performed on decalcified bone sections of femur, tibiae, or vertebrae. The von Kossa staining and double calcein labeling were performed on undecalcified bone sections (24, 25). The left femurs, tibiae, and vertebrae (L1–L3) were removed and fixed in PLP fixative (2% paraformaldehyde containing 0.075 M lysine and 0.01 M sodium periodate solution) overnight at 4 C and processed histologically as described (24). The femur, tibia, and vertebra were decalcified in EDTA glycerol solution for 5–7 d at 4 C. Decalcified bones and other tissues were dehydrated and embedded in paraffin, after which 5-µm sections were cut on a rotary microtome. The sections were stained with H&E or histochemically for TRAP activity or immunohistochemically as described below. The right undecalcified femurs were embedded in LR White acrylic resin (London Resin Company Ltd., London, UK), and 1-µm sections were cut on an ultramicrotome. These sections were stained for mineral with the von Kossa staining procedure and counterstained with toluidine blue.

Enzyme histochemistry for TRAP was performed as previously described (25). De-waxed sections were preincubated for 20 min in buffer containing 50 mM sodium acetate and 40 mM sodium tartrate at pH 5.0. Sections were then incubated for 15 min at room temperature in the same buffer containing 2.5 mg/ml naphthol AS-MX phosphate (Sigma-Aldrich, St. Louis, MO) in dimethylformamide as substrate and 0.5 mg/ml fast garnet GBC (Sigma-Aldrich) 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.

Double calcein labeling was performed by ip injection of mice with 10 µg calcein/g body weight (C-0875; Sigma-Aldrich) at 10 and 3 d before the mice were killed. Bones were harvested and embedded in LR White acrylic resin as described above. Serial sections were cut, and the freshly cut surface of each section was viewed and imaged using fluorescence microscopy. The double calcein interlabel width was measured using Northern Eclipse image analysis software version 6.0 (Empix Imaging Inc., Mississauga, Ontario, Canada), and the mineral apposition rate (MAR; MAR = interlabel width/labeling period) and bone formation rate (BFR) were calculated according to the formula: [BFR = MAR x (dL.Pm + sL.Pm/2)/B.Pm], where dL.Pm is the double-labeled perimeter, sL.Pm is the single-labeled perimeter, and B.Pm is the bone perimeter. Calculations were made on a minimum of duplicate specimens from five mice of each group.

Immunohistochemical staining
The PTH receptor (PTHR) (26) and aromatase were assessed by immunohistochemistry using the avidin-biotin complex techniques as previously described followed by digital recording and image analysis as described (27). Briefly, affinity-purified mouse monoclonal anti-PTHR antibody (05-517, Upstate, Lake Placid, NY) (1:200) and polyclonal antihuman aromatase antibody (kindly donated by Dr. N. Harada, Fujita Health University, Toyoake, Japan) (1:200) were applied to dewaxed paraffin sections overnight at room temperature, respectively. For the aromatase studies, specimens were pretreated with bovine testicular hyaluronidase for 30 min at 37 C followed by 3% H2O2 for 10 min at 37 C. As a negative control for both PTHR and aromatase staining, preimmune serum was substituted for the primary antibody. After washing with high-salt buffer [50 mM Tris-HCl, 2.5% NaCl, 0.05% Tween 20 (pH 7.6)] for 10 min at room temperature followed by two 10-min washes with Tris-buffered saline, secondary antibody was added, and substrate staining was performed as per the manufacturer’s instructions in the Vectastain avidin-biotin complex kit (Vector Laboratories, Peterborough, UK). Sections were developed with either diaminobenzidine or alkaline phosphatase, producing red pigmentation for PTHR and brown for aromatase, respectively. The sections were counterstained with methyl green and mounted with Kaiser’s glycerol jelly after washing with distilled water. The immunostaining reactions were photographed using a Leica digital camera (Leica, Wetzlar, Germany) and processed for image analysis. Thus, after immunohistochemical staining, all digital images were captured at a magnification of x200 that allowed inclusion of the metaphysial area of interest in a rectangular inclusion frame. All recorded images were then analyzed using Northern Eclipse image analysis software, version 5.0 (Empix Imaging, Inc.). Quantitative data for PTHR was obtained using standard thresholding methods as described previously (24). Data obtained from thresholding approaches was expressed by the term summary total gray, the term used as a predetermined output measure by the manufacturer of the software. This term refers to the total signal derived from a given area using an 8-bit digital storage intensity of gray scale levels defined by integers ranging from 0–255 for the level of gray (gray scale pixel brightness) associated with each pixel within the given area. Comparisons were made on a minimum of duplicate specimens from five mice of each genotype.

Histomorphometry
After H&E staining or histochemical or immunohistochemical staining, histomorphometric measurements were performed as previously described (28) in the secondary spongiosa in the metaphyseal area (0.5 mm below the growth plate) at the distal end of femur. The parameter measured for bone mass was the trabecular BV/TV (percentage). The parameters obtained for bone formation were the osteoblast surface per bone surface (Ob.S/BS; percentage), MAR (micrometers per day), and BFR (micrometers per day). The parameter measured for bone resorption was the osteoclast surface per bone surface (Oc.S/BS; percentage). After H&E staining or histochemical staining of sections from five mice of each genotype, images of fields were photographed with a Leica 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 (22, 23, 24, 25, 27).

Quantitative real-time PCR
Total RNA was extracted from whole humeri, using Trizol reagent (Invitrogen, Carlsbad, CA) according to the manufacturer’s protocol. Total RNA was reverse transcribed to cDNA using the Expand Reverse Transcriptase (Roche Applied Science, Mannheim, Germany). The number of cDNA molecules in the reverse-transcribed samples was determined by real-time PCR analyses using the LightCycler system (Roche Applied Science) with the following primers: receptor activator of nuclear factor {kappa} B ligand (RANKL), 5'-GGTCGGGCAATTCTG AATT-3' and 5'-GGGGAATTACAAAGTGCACCAG (813 bp); osteoprotegerin (OPG), 5'-TGGAGATCGAATTCTGCTTG-3' and 5'-TCAAGTGCTTGAGGGCATAC-3' (719 bp); and glyceraldehyde-3-phosphate dehydrogenase (GAPDH), 5'-GGTCGGTGTGAACGGATTTG-3' and 5'-ATGAGCCCTTCCACAATG-3' (508 bp). The conditions were 2 µl LightCycler DNA master SYBR Green I, 0.25 µM each 5' and 3' primer, and 2 µl sample and/or H2O to a final volume of 20 µl. The MgCl2 concentration was adjusted to 3 mM. Samples were amplified for 45 cycles with a temperature transition rate of 20 C/sec for all three steps, which included denaturation at 95 C for 10 sec, annealing at 60 C for 5 sec, and extension at 72 C for the proper time. A melting curve was obtained at the end of each run to discriminate specific from nonspecific cDNA products, by denaturing at 95 C for 3 sec, then decreasing the temperature to 70 C for 30 sec and then raising the temperature slowly from 70 to 95 C using a temperature transition rate of 0.2 C/sec. The cDNA content was normalized by subtracting the cycle numbers [cycle time (Ct)] of GAPDH from those of the target gene ({Delta}Ct = Ct of target gene – Ct of GAPDH), and the gene expression level was calculated using the formula of 2–({Delta}Ct). The normalized RANKL levels relative to the normalized OPG levels in the FORKO mice were expressed as a percentage of this ratio in WT mice.

Statistical analysis
Data were presented as the mean ± SEM. Statistical comparisons were made using two-way ANOVA with Bonferroni adjustment or with a one-way nonparametric ANOVA, and P < 0.05 was considered significant.


   Results
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 References
 
Age-related bone loss in the FORKO mouse and rescue by ovarian transplantation
BMD increased with age in both WT and FORKO mice; however, by 3 months of age, the BMD of the FORKO mouse was significantly lower than that of the WT, and these differences were slightly but not significantly greater in vertebral (lumbar) than in long bone (femur) (Fig. 1AGo). Thus, at 3 months of age, in the FORKO mouse compared with WT mice, femoral spine BMD was reduced by 4.9% and lumbar spine BMD by 5.6%. By 6 months of age, femoral spine BMD was reduced by 6.1% and lumbar spine by 7.7%.


Figure 1
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  		FIG. 1. BMD decreases in FORKO mice but increases after ovarian transplantation. Temporal changes in BMD of the femur and lumbar spine in FORKO vs. WT mice (A). Each bar represents the mean of duplicate determinations in eight mice. Serum estradiol (B), LH (C), and free testosterone concentrations (D) in sham-operated and FORKO mice in 5 months old after transplantation. D, Detection limits of the assays. Each bar represents the mean of duplicate determinations in five mice. BMD and trabecular BV/TV changes at 5 months of age in sham-operated (SHAM) mice vs. transplanted (TRANS) FORKO mice (E and F). Each bar represents the mean of duplicate determinations in five mice. Error bars, SEM. *, P < 0.05; **, P < 0.01; ***, P < 0.001.

 
To determine whether the bone loss in the FORKO mouse was associated with the alteration in ovarian function, allogeneic ovarian transplantation was performed. After ovarian transplantation, circulating estradiol concentrations in the transplanted FORKO mice increased (Fig. 1BGo), LH and testosterone concentrations fell (Fig. 1Go, C and D), and, by 5 months of age, femoral and lumbar spine BMD losses and trabecular bone loss as determined by histomorphometry were greatly reduced in the transplanted FORKO mice (Fig. 1Go, E and F).

MicroCT analysis in the FORKO mouse at 3 months of age revealed a marked reduction of trabecular bone (Fig. 2AGo) (45.9% reduction of BV/TV from 15.53–8.4%, P < 0.01), which was confirmed by the reduction in trabecular BV (BV/TV) observed by histomorphometry (Fig. 2BGo). The magnitude of the BV/TV in the WT mice was in line with that previously reported in several strains of mice (29, 30). The reduction in BV/TV was associated with significant decreases in osteoblasts (Fig. 2CGo), in the MAR (Fig. 2DGo), and in the BFR (which was reduced by 37% from 0.48–0.30 µm/d; P < 0.01) (Fig. 2EGo). A significant increase in the osteoclast surface was noted from H&E sections (Fig. 2FGo), and these increases in Oc/BS levels were confirmed on TRAP-stained sections. Furthermore, the ratio of the gene expression of RANKL, the major activator of osteoclasts, to OPG, a decoy receptor for RANKL (31), was increased (Fig. 2Go, F and G, respectively).


Figure 2
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  		FIG. 2. Reduced trabecular bone, decreased bone formation, and increased bone resorption in FORKO mice at 3 months of age. MicroCT image of distal femur (A) and trabecular BV/TV (B). Both show a reduction in trabecular bone in the FORKO relative to the WT mice. Ob.S/BS (C), MAR (D), and BFR (E) were all reduced, but the Oc.S/BS (F) was increased. Each bar represents the mean of duplicate determinations in five animals per group. G, Ratio of RANKL to OPG mRNA levels as determined by real-time PCR. Each bar shows the ratio of RANKL to OPG relative to the ratio of RANKL to OPG in the WT mice and represents a mean of four determinations. Error bars, SEM. **, P < 0.01.

 
Effect of OVX on bone metabolism in the FORKO mouse
To determine whether other ovarian products might contribute to the maintenance of bone mass in the FORKO mouse, we performed bilateral OVX at 3 wk of age and compared ovariectomized FORKO and WT mice. By 3 months of age, marked losses of BMD were observed in both WT and FORKO mice, which progressed with age (Fig. 3Go, A and B). These losses were again more prominent in the lumbar spine (lumbar vertebrae) of the ovariectomized FORKO mice compared with intact sham-operated FORKO mice [4.2% (P < 0.05) in the femur and 7.8% (P < 0.01) in the lumbar spine at 3 months of age (difference between femoral and lumbar spine BMD losses was significant at P < 0.05) and 6.9% (P < 0.01) in the femur and 15.9% (P < 0.001) in the lumbar spine at 6 months of age (difference between femoral and lumbar spine BMD losses was significant at P < 0.01)]. The BMD losses were also slightly more prominent in the lumbar spine of the WT after OVX compared with intact sham-operated WT mice [8% (P < 0.001) in the femur and 9.5% (P < 0.01) in the lumbar spine at 3 months of age and 13% (P < 0.001) in the femur and 19% (P < 0.001) in the lumbar spine at 6 months of age (difference between femoral and lumbar spine losses was significant at P < 0.05)]. Of major importance, there was no significant difference in the BMD of the femur or of the lumbar spine between FORKO ovariectomized and WT ovariectomized animals at either 3 or 6 months of age, i.e. the losses in BMD after OVX in the FORKO mice reduced BMD to the levels observed after OVX in the WT mice. MicroCT analysis demonstrated a reduction in trabecular bone of 39.7% in the FORKO ovariectomized mouse and of 59.1% in the WT ovariectomized mouse, and von Kossa staining of long bones confirmed marked reductions in trabecular bone after OVX in the FORKO as well as the WT animals (Fig. 3Go, C and D). By histomorphometry, the trabecular BV in FORKO-OVX was reduced by 42% compared with FORKO-sham at 3 months of age such that, as with microCT analysis, there was no significant difference between WT-OVX and FORKO-OVX in BV/TV (Fig. 3EGo).


Figure 3
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  		FIG. 3. OVX reduces BMD and trabecular bone in FORKO mice. BMD of the femur (A) and lumbar spine (B) in WT and FORKO mice after OVX and after sham operation (SHAM) in FORKO and WT mice. BMD was performed in duplicate in eight mice in each group. MicroCT images (C) of distal femurs at 3 months of age. Von Kossa staining (D) of distal femurs at 3 months of age. Trabecular BV relative to TV (BV/TV) (E) of WT and FORKO sham-operated and ovariectomized mice at 3 months of age. Each bar represents the mean of duplicate determinations from five mice per group. Error bars, SEM. *, P < 0.05; **, P < 0.01; ***, P < 0.001. NS, Not significant.

 
By 3 months of age, in the ovariectomized WT mice, the staining intensity of osteoblasts for the PTHR (Fig. 4BGo), the osteoblast surface (Fig. 4CGo), the MAR (Fig. 4DGo), and the BFR (Fig. 4EGo) were all elevated. In contrast, no significant increase in any of these parameters was observed in the ovariectomized FORKO mice (Fig. 4Go). Thus, the BFRs for the WT sham, WT ovariectomized, FORKO sham, and FORKO ovariectomized were 0.45, 0.57, 0.31, and 0.28 µm/d, respectively (Fig. 4EGo); thus, there was a 27% increase after OVX in WT mice (P < 0.05) but no significant increase in the ovariectomized FORKO compared with the FORKO sham. Nevertheless, the osteoclast surface relative to bone surface was significantly increased after OVX in both the WT and the FORKO mice from H&E-stained sections, and the Oc.S/BS levels were confirmed on TRAP-stained sections. (Fig. 4FGo). Furthermore, the ratio of the gene expression levels in bone of RANKL to OPG was significantly increased after OVX in both WT and FORKO mice (Fig. 4GGo).


Figure 4
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  		FIG. 4. Changes in bone formation and bone resorption 3 months after OVX in WT and FORKO mice. Double calcein labeling (A; scale bar, 50 µm), immunohistochemistry of the PTHR (B; scale bar, 50 µm), Ob.S/BS (C), MAR (D), and BFR (E) in WT and FORKO mice after either sham operation (SHAM) or OVX. Although staining intensity of the PTHR, reflecting osteoblasts (B), was increased after OVX in WT mice, no such increase was observed after OVX in FORKO mice. Ob.S/BS (C) and BFR (E) were significantly increased in ovariectomized WT mice but not in ovariectomied FORKO mice. Osteoclast surface relative to bone surface (Oc.S/B.S) was increased after OVX in both WT and FORKO mice (F). Each bar represents the mean of duplicate determinations in five mice of each group. Ratio of mRNA levels of RANKL to OPG from femurs of sham-operated or ovariectomized WT and FORKO mice (G). Each bar shows the ratio of RANKL to OPG relative to the ratio of RANKL to OPG in the sham ovariectomized WT mice and represents the mean of four determinations. Error bars represent the SEM. *, P < 0.05; **, P < 0.01; ***, P < 0.001.

 
Effect of ovarian androgens on bone in the FORKO mice
In view of the high reported circulating concentrations of androgens in the FORKO mouse (20), we next examined the possible impact of these hormones on the FORKO skeleton. Circulating testosterone concentrations were 7.8-fold increased in FORKO mice relative to WT controls. After OVX, testosterone concentrations were undetectable in both WT and FORKO mice (Fig. 5AGo). To determine whether androgens were directly altering bone via the androgen receptor, FORKO mice were treated with the nonsteroidal androgen receptor antagonist, flutamide. At 3 months of age, flutamide treatment slightly, but not significantly, reduced BMD in the femur (by 3.0% compared with a 4.7% reduction by OVX) and in the lumbar spine (by 2.3% compared with a reduction of 9.3% by OVX) (Fig. 5BGo). To determine whether androgens were altering bone indirectly after conversion to estrogen, FORKO mice were treated with the aromatase inhibitor, letrozole. Aromatase has previously been shown to be expressed in skeletal cells (32); therefore, we assessed whether it was present in skeletal tissue of the FORKO mouse. Aromatase protein expression was indeed evident predominantly in osteoblastic cells of FORKO as well as WT mice (Fig. 5CGo). At 3 months of age, letrozole treatment produced a significant reduction of BMD both in the femur (by 6.8%) and in the lumbar spine (by 12.1%) (Fig. 5DGo). Analysis by microCT and von Kossa staining demonstrated a reduction of trabecular bone in the FORKO mice after treatment with flutamide but a more prominent reduction with letrozole (Fig. 6Go, A and B). Histomorphometric analysis confirmed both a significant flutamide-induced reduction of trabecular BV (10.4% reduction, P < 0.05) and a more marked letrozole-induced decrease (20.4% reduction, P < 0.05) (Fig. 6CGo). Flutamide treatment did not significantly alter osteoblastic surface (Fig. 6DGo) and the BFR (Fig. 6EGo) but significantly increased the osteoclastic surface relative to the bone surface (14.6% increase, P < 0.05) (Fig. 6FGo). Letrozole treatment reduced the osteoblastic surface (15.6% reduction, P < 0.05) and the BFR (15.1% reduction, P < 0.05) (Fig. 6Go, D and E, respectively), whereas it increased the osteoclastic surface (16.7% increase, P < 0.05) (Fig. 6FGo).


Figure 5
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  		FIG. 5. Role of testosterone in the FORKO mouse. Serum testosterone concentrations were elevated in the sham-operated (SHAM) FORKO mice relative to sham-operated WT mice and became undetectable after OVX. Each bar represents the mean level of duplicate determinations in five mice. D, Detection limits of the assays (A). BMD in the femur and lumbar spine of the flutamide-treated relative to the vehicle-treated FORKO mouse and relative to the OVX FORKO mouse (B). Each bar represents the mean of duplicate determinations in six animals. Expression of aromatase by immunohistochemistry in osteoblastic cells of the FORKO mouse relative to the WT mouse (C; scale bar, 100 µm). Reductions of BMD of the femur and lumbar spine in FORKO mice treated with letrozole or vehicle (D). Each bar represents the mean of duplicate determinations in six mice. *, P < 0.05; ***, P < 0.001. Error bars represent the SEM.

 

Figure 6
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  		FIG. 6. BV and indices of bone formation and resorption in flutamide-treated and letrozole-treated FORKO mice. MicroCT images (A) and von Kossa stains (B) of femurs from FORKO mice treated from left to right, respectively, with flutamide vehicle, flutamide (flu), OVX (ovx), letrozole (let), or letrozole vehicle (pbs). Trabecular BV (BV/TV) was significantly reduced by flutamide, OVX and letrozole relative to untreated FORKO controls (C). The Ob.S/BS (D) was increased by OVX relative to FORKO controls and reduced by letrozole. The BFR (E) was significantly reduced by letrozole but not by flutamide. Oc.S/BS (F) was increased by flutamide and letrozole as well as by OVX. Each bar represents the mean of duplicate determinations from five animals. Error bars, SEM. *, P < 0.05; ***, P < 0.001.

 

   Discussion
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
References
 
In our studies, transplantation of ovaries with functional FSH receptors at 6 wk of age partly restored the diminished estrogen levels in the FORKO mouse and also partly abrogated the loss of bone that had clearly occurred in the absence of FSH action in this model. To determine whether ovarian products other than estrogen could also influence bone homeostasis in the FORKO model, we examined the effect of removing endogenous ovarian secretion products via bilateral OVX at puberty, at which time ovarian function is markedly augmented. The timing of the OVX we performed in both FORKO and WT mice is a limitation of our study in its analogy to postmenopausal osteoporosis in that these mice do not represent models of postmenopausal osteoporosis. Nevertheless, because of the seemingly important role of estrogens in maintaining bone mass and preventing bone loss after puberty, developmental models in which estrogen deficiency occurs at an even earlier age than 3 wk, and which include the double null mutant estrogen receptor {alpha} and ß (ER{alpha}ß–/–) (33, 34), the aromatase-deficient (ArKO) mouse (35, 36, 37) and indeed the FSH receptor knockout mouse (9, 18, 20) have all been used to explore the pathophysiology of osteoporosis. Furthermore, in humans, reduced ovarian function in growing adolescents (3, 4, 5) has been associated with reduced bone mass, which has been termed osteoporosis; the attainment of peak bone mass is at least partly dependent on optimal estrogen levels (38); and peak bone mass, in addition to accelerated postmenopausal bone loss, is regarded as a highly significant cause of postmenopausal osteoporosis (2, 39). Our results are therefore relevant to the contribution of the pituitary-ovarian axis to the development of osteoporosis.

OVX in our animals dramatically and progressively reduced BMD to the level of that seen in WT litter mates after bilateral OVX. Consequently, it would appear that bone loss occurring in the nonovariectomized FORKO mouse plus bone loss occurring due to OVX in the FORKO mouse could account for the majority if not all of the bone loss that is observed after OVX in WT animals. These results therefore suggest that the predominant if not sole influence of FSH in modulating skeletal mass in this model was via the ovary.

The temporal association of reduced serum testosterone and increased bone loss after OVX in our model prompted us to explore further a possible role for testosterone in this process. Testosterone is converted to 17ß-estradiol via the aromatase enzyme that is a product of the cyp19 gene and a member of the cytochrome P450 family (40) and is found in several tissues including ovary, fat, and bone. Previous studies in rat granulosa cells in vitro have reported that aromatase action may be dependent on FSH stimulation (19), although recent studies have suggested that gene expression and protein expression of ovarian aromatase were not substantially reduced in the young FORKO mouse (20). We first localized the presence of aromatase on osteoblastic cells in the skeleton of the FORKO mouse. In view of the fact that circulating estrogen levels are virtually undetectable in the FORKO mouse, the demonstration of skeletal aromatase suggested that testosterone conversion might be occurring locally in bone rather than in extraosseous tissues. We then demonstrated that aromatase inhibition could indeed produce a substantial reduction in bone mass, indicating that local skeletal production of estrogen was an important regulator of bone mass in the FORKO mouse and appears to play a substantial role in maintaining bone mass. Use of aromatase inhibition in postmenopausal women has been reported to further decrease bone mass above that seen in the postmenopausal state, suggesting that extraovarian conversion of androgens to estrogen also plays an important role in maintaining bone mass in humans (41). Further studies to elucidate the regulation of skeletal aromatase may therefore be indicated to develop mechanisms for enhancing skeletal conversion of testosterone to estrogen.

Our results therefore show that both aromatase inhibition and androgen blockade reduce trabecular bone as previously reported in other models (34, 36, 37), although the effects in the FORKO mouse were considerably stronger with aromatase inhibition. Consequently, ovarian androgens play a major role in protecting the skeleton of the FORKO mouse from the bone loss of ovarian failure. Further consideration may therefore have to be given to the role of androgen and particularly to its skeletal conversion to estrogen in assessing normal and abnormal fluctuations in bone turnover both during reproductive life and after menopause.

Although skeletal maintenance of the FORKO mouse is therefore clearly ovary-dependent, differences were observed in the mechanisms of bone loss in the ovariectomized FORKO mice and in the ovariectomized WT animals. Thus, although increased bone resorption was the predominant mechanism of bone loss in both FORKO and WT animals, this was accompanied by increased bone formation, i.e. increased bone turnover in the WT ovariectomized mice we studied. Although markedly increased bone formation after OVX has not always been reported in WT mice, bone turnover appeared to be clearly uncoupled in the FORKO mice, and increased resorption was associated with either no change in parameters of bone formation or actual reductions. Such effects on bone formation have been observed in the (ER{alpha}ß) mice (33, 34) and are therefore compatible with long-term reduced estrogen effects in these developmental models. Consequently, this apparent uncoupling in the FORKO mouse may indicate that lifelong exposure to reduced ovarian estrogens can alter the bone microenvironment and blunt the mechanisms that normally link bone resorption to formation, many of which are still not well understood. Alternatively, it may indicate that other ovarian products, either by direct or indirect mechanisms, inhibit increases in bone formation. Although the absence of FSH action on osteoclasts could theoretically diminish coupling, it has been reported that FSH per se has no effect on osteoblastic bone formation (9). These subtle but potentially important mechanistic issues therefore clearly require further exploration.

In summary, our studies point to the critical role of ovarian secretory products in maintaining skeletal integrity in the FSH receptor-deficient mouse. Whether ovarian products other than estrogen and testosterone modify the activity of these sex steroids will need to be evaluated in the future. Our studies suggest that a continued focus on identifying and maximizing the actions of ovarian-derived agonists on bone may be a fruitful avenue to develop improved agents for treatment of osteoporosis.


   Acknowledgments
 
We acknowledge the excellent facilities of the Centre for Bone and Periodontal Research at McGill where imaging and histology were performed.


   Footnotes
 
First Published Online March 1, 2007

Abbreviations: BFR, Bone formation rate; BMD, bone mineral density; BV, bone volume; Ct, cycle time; GAPDH, glyceraldehyde-3-phosphate dehydrogenase; H&E, hematoxylin and eosin; MAR, mineral apposition rate; microCT, microcomputed tomography; Ob.S/BS, osteoblast surface per bone surface; Oc.S/BS, osteoclast surface per bone surface; OPG, osteoprotegerin; OVX, ovariectomy; PTHR, PTH receptor; RANKL, receptor activator of nuclear factor {kappa} B ligand; TRAP, tartrate-resistant acid phosphatase; TV, tissue volume; WT, wild type.

This work was supported by the Canadian Institutes for Health Research (grants to A.C.K., M.R.S., and D.G.). R.S. was the recipient of a fellowship award from the Skeletal Health Training Program of the Canadian Institutes for Health Research.

J.G., R.T.-P., R.S., Y.Y., D.M., A.C.K., M.R.S., and D.G. have nothing to declare.

Received October 17, 2006.

Accepted for publication February 22, 2007.


   References
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 References
 

  1. Albright FS, Smith PH, Richardson AM 1941 Postmenopausal osteoporosis. JAMA 116:2465–2474[Abstract/Free Full Text]
  2. Riggs BL, Khosla S, Melton 3rd LJ 2002 Sex steroids and the construction and conservation of the adult skeleton. Endocr Rev 23:279–302[Abstract/Free Full Text]
  3. Warren MP, Chua A 2006 Appropriate use of estrogen replacement therapy in adolescents and young adults with Turner syndrome and hypopituitarism in light of the Women’s Health Initiative. Growth Horm IGF Res Suppl A:S98–S102
  4. Bondy CA, Bakalov VK 2006 Investigation of cardiac status and bone mineral density in Turner syndrome. Growth Horm IGF Res Suppl A:S103–S108
  5. Tauchmanova L De Simone G, Musella T, Orio F, Ricci P, Nappi C, Lombardi G, Colao A, Rotoli B, Selleri C 2006 Effects of various antiresorptive treatments on bone mineral density in hypogonadal young women after allogeneic stem cell transplantation. Bone Marrow Transplant 37:81–88[Medline]
  6. Martin TJ, Gaddy D 2006 Bone loss goes beyond estrogen. Nat Med 12:612–613[CrossRef][Medline]
  7. Baron R 2006 FSH versus estrogen: who’s guilty of breaking bones? Cell Metab 3:302–305[CrossRef][Medline]
  8. Gaddy-Kurten D, Coker JK, Abe E, Jilka RL, Manolagas SC 2002 Inhibin suppresses and activin stimulates osteoblastogenesis and osteoclastogenesis in murine bone marrow cultures. Endocrinology 143:74–83[Abstract/Free Full Text]
  9. Sun L, Peng Y, Sharrow AC, Iqbal J, Zhang Z, Papachristou DJ, Zaidi S, Zhu LL, Yaroslavskiy BB, Zhou H, Zallone A, Sairam MR, Kumar TR, Bo W, Braun J, Cardoso-Landa L, Schaffler MB, Moonga BS, Blair HC, Zaidi M 2006 FSH directly regulates bone mass. Cell 125:247–260[CrossRef][Medline]
  10. Goltzman D 2002 Discoveries, drugs and skeletal disorders. Nat Rev Drug Discov 1:784–796[CrossRef][Medline]
  11. Lindsay R 2004 Hormones and bone health in postmenopausal women. Endocrine 24:223–230[CrossRef][Medline]
  12. Anderson GL, Limacher M, Assaf AR, Bassford T, Beresford SA, Black H, Bonds D, Brunner R, Brzyski R, Caan B, Chlebowski R, Curb D, Gass M, Hays J, Heiss G, Hendrix S, Howard BV, Hsia J, Hubbell A, Jackson R, Johnson KC, Judd H, Kotchen JM, Kuller L, LaCroix AZ, Lane D, Langer RD, Lasser N, Lewis CE, Manson J, Margolis K, Ockene J, O’Sullivan MJ, Phillips L, Prentice RL, Ritenbaugh C, Robbins J, Rossouw JE, Sarto G, Stefanick ML, Van Horn L, Wactawski-Wende J, Wallace R, Wassertheil-Smoller S; Women’s Health Initiative Steering Committee 2004 Effects of conjugated equine estrogen in postmenopausal women with hysterectomy: the Women’s Health Initiative randomized controlled trial. JAMA 291:1701–1712[Abstract/Free Full Text]
  13. Ettinger B, Black DM, Mitlak BH, Knickerbocker RK, Nickelsen T, Genant HK, Christiansen C, Delmas PD, Zanchetta JR, Stakkestad J, Gluer CC, Krueger K, Cohen FJ, Eckert S, Ensrud KE, Avioli LV, Lips P, Cummings SR 1999 Reduction of vertebral fracture risk in postmenopausal women with osteoporosis treated with raloxifene: results from a 3-year randomized clinical trial. Multiple Outcomes of Raloxifene Evaluation (MORE) Investigators. JAMA 282:637–645[Abstract/Free Full Text]
  14. Schlegel W, Petersdorf LI, Junker R, Schulte H, Ebert C, Von Eckardstein A 1999 The effects of six months of treatment with a low-dose of conjugated oestrogens in menopausal women. Clin Endocrinol (Oxf) 51:643–651[CrossRef][Medline]
  15. Cheng WC, Yen ML, Hsu SH, Chen KH, Tsai KS 2004 Effects of raloxifene, one of the selective estrogen receptor modulators, on pituitary-ovary axis and prolactin in postmenopausal women. Endocrine 23:215–218[CrossRef][Medline]
  16. Lasco A, Cannavo S, Gaudio A, Morabito N, Basile G, Nicita-Mauro V, Frisina N 2002 Effects of long-lasting raloxifene treatment on serum prolactin and gonadotropin levels in postmenopausal women. Eur J Endocrinol 147:461–465[Abstract]
  17. Dierich A, Sairam MR, Monaco L, Fimia GM, Gansmuller A, LeMeur M, Sassone-Corsi P 1998 Impairing follicle-stimulating hormone (FSH) signaling in vivo: targeted disruption of the FSH receptor leads to aberrant gametogenesis and hormonal imbalance. Proc Natl Acad Sci USA 95:13612–13617[Abstract/Free Full Text]
  18. Danilovich N, Javeshghani D, Xing W, Sairam MR 2002 Endocrine alterations and signaling changes associated with declining ovarian function and advanced biological aging in follicle-stimulating hormone receptor haploinsufficient mice. Biol Reprod 67:370–378[Abstract/Free Full Text]
  19. Fitzpatrick SL, Richards JS 1991 Regulation of cytochrome P450 aromatase messenger ribonucleic acid and activity by steroids and gonadotropins in rat granulosa cells. Endocrinology 129:1452–1462[Abstract/Free Full Text]
  20. Danilovich N, Babu PS, Xing W, Gerdes M, Krishnamurthy H, Sairam MR 2000 Estrogen deficiency, obesity, and skeletal abnormalities in follicle-stimulating hormone receptor knockout (FORKO) female mice. Endocrinology 141:4295–4308[Abstract/Free Full Text]
  21. Weissman A, Gotlieb L, Colgan T, Jurisicova A, Greenblatt EM, Casper RF 1999 Preliminary experience with subcutaneous human ovarian cortex transplantation in the NOD-SCID mouse. Biol Reprod 60:1462–1467[Abstract/Free Full Text]
  22. Miao D, He B, Jiang Y, Kobayashi T, Soroceanu MA, Zhao J, Su H, Tong X, Amizuka N, Gupta A, Genant HK, Kronenberg HM, Goltzman D, Karaplis AC 2005 Osteoblast-derived PTHrP is a potent endogenous bone anabolic agent that modifies the therapeutic efficacy of administered PTH 1–34. J Clin Invest 115:2402–2411[CrossRef][Medline]
  23. Miao D, Li J, Xue Y, Su H, Karaplis AC, Goltzman D 2004 Parathyroid hormone-related peptide is required for increased trabecular bone volume in parathyroid hormone-null mice. Endocrinology 145:3554–3562[CrossRef][Medline]
  24. Miao D, Bai X, Panda D, McKee M, Karaplis A, Goltzman D 2001 Osteomalacia in hyp mice is associated with abnormal phex expression and with altered bone matrix protein expression and deposition. Endocrinology 142:926–939[Abstract/Free Full Text]
  25. Miao D, Scutt A 2002 Recruitment, augmentation and apoptosis of rat osteoclasts in 1,25-(OH)2D3 response to short-term treatment with 1,25-dihydroxyvitamin D3 in vivo. BMC Musculoskelet Disord 3:16[CrossRef][Medline]
  26. Samadfam R, Xia Q, Goltzman D 2007 Cotreatment of parathyroid hormone with osteoprotegerin or alendronate increases its anabolic effect on the skeleton of oophorectomized mice. J Bone Miner Res 22:55–63[CrossRef][Medline]
  27. Panda DK, Al Kawas S, Seldin MF, Hendy GN, Goltzman D 2004 Inactivation of the 25-hydroxyvitamin D 1{alpha}-hydroxylase and vitamin D receptor demonstrates independent and interdependent effects of calcium and vitamin D on skeletal and mineral homeostasis. J Biol Chem 279:16754–16766[Abstract/Free Full Text]
  28. Parfitt AM, Drezner MK, Glorieux FH, Kanis JA, Malluche H, Meunier PJ, Ott SM, Recker RR 1987 Bone histomorphometry: standardization of nomenclature, symbols, and units. Report of the ASBMR Histomorphometry Nomenclature Committee. J Bone Miner Res 2:595–610[Medline]
  29. Westbroek I, Waarsing JH, van Leeuwen JP, Waldum H, Reseland JE, Weinans H, Syversen U, Gustafsson BI 12 December 2006 Long-term fluoxetine administration does not result in major changes in bone architecture and strength in growing rats. J Cell Biochem Epub ahead of print, PMID 17163489
  30. Pierroz DD, Bouxsein ML, Rizzoli R, Pierroz DD, Bouxsein ML, Rizzoli R, Ferrari SL 2006 Combined treatment with a ß-blocker and intermittent PTH improves bone mass and microarchitecture in ovariectomized mice. Bone 39:260–267[Medline]
  31. Boyle WJ SW, Lacey DL 2003 Osteoclast differentiation and activation. Nature 423:337–342[CrossRef][Medline]
  32. van der Eerden BC, Lowik CW, Wit JM, Karperien M 2004 Expression of estrogen receptors and enzymes involved in sex steroid metabolism in the rat tibia during sexual maturation. J Endocrinol 180:457–467[Abstract]
  33. Sims NA, Dupont S, Krust A, Clement-Lacroix P, Minet D, Resche-Rigon M, Gaillard-Kelly M, Baron R 2002 Deletion of estrogen receptors reveals a regulatory role for estrogen receptors-ß in bone remodeling in females but not in males. Bone 30:18–25[Medline]
  34. Sims NA, Clement-Lacroix P, Minet D, Fraslon-Vanhulle C, Gaillard-Kelly M, Resche-Rigon M, Baron R 2003 A functional androgen receptor is not sufficient to allow estradiol to protect bone after gonadectomy in estradiol receptor-deficient mice. J Clin Invest 111:1319–1327[CrossRef][Medline]
  35. Miyaura C, Toda K, Inada M, Ohshiba T, Matsumoto C, Okada T, Ito M, Shizuta Y, Ito A 2001 Sex- and age-related response to aromatase deficiency in bone. Biochem Biophys Res Commun 280:1062–1068[CrossRef][Medline]
  36. Yeh S, Tsai MY, Xu Q, Mu XM, Lardy H, Huang KE, Lin H, Yeh SD, Altuwaijri S, Zhou X, Xing L, Boyce BF, Hung MC, Zhang S, Gan L, Chang C 2002 Generation and characterization of androgen receptor knockout (ARKO) mice: an in vivo model for the study of androgen functions in selective tissues. Proc Natl Acad Sci USA 99:13498–13503[Abstract/Free Full Text]
  37. Oz OK, Hirasawa G, Lawson J, Nanu L, Constantinescu A, Antich PP, Mason RP, Tsyganov E, Parkey RW, Zerwekh JE, Simpson ER 2001 Bone phenotype of the aromatase deficient mouse. J Steroid Biochem Mol Biol 79:49–59[CrossRef][Medline]
  38. Yilmaz D EB, Bilgin E, Gumuser G, Onur E, Pinar ED 2005 Bone mineral density in girls and boys at different pubertal stages: relation with gonadal steroids, bone formation markers, and growth parameters. J Bone Miner Metab 23:476–482[CrossRef][Medline]
  39. Misra M, Klibanski A 2006 Anorexia nervosa and osteoporosis. Rev Endocr Metab Disord 7:91–99[CrossRef][Medline]
  40. Nelson DR, Kamataki T, Waxman DJ, Nelson DR, Kamataki T, Waxman DJ, Guengerich FP, Estabrook RW, Feyereisen R, Gonzalez FJ, Coon MJ, Gunsalus IC, Gotoh O, Okuda K, Nebert DW 1993 The P450 superfamily: update on new sequences, gene mapping, accession numbers, early trivial names of enzymes, and nomenclature. DNA Cell Biol 12:1–51[Medline]
  41. Eastell R, Hannon R 2005 Long-term effects of aromatase inhibitors on bone. J Steroid Biochem Mol Biol 95:151–154[CrossRef][Medline]



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Hypogonadal Bone Loss: Sex Steroids or Gonadotropins?
Endocrinology, June 1, 2007; 148(6): 2610 - 2612.
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