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Journal of Bone and Mineral Research logoLink to Journal of Bone and Mineral Research
. 2010 May 17;25(11):2438–2446. doi: 10.1002/jbmr.129

Effects of chronic estrogen treatment on modulating age‐related bone loss in female mice

Farhan A Syed 1, Ulrike IL Mödder 1, Matthew Roforth 1, Ira Hensen 1, Daniel G Fraser 1, James M Peterson 1, Merry Jo Oursler 1, Sundeep Khosla 1,✉
PMCID: PMC3119867  NIHMSID: NIHMS301984  PMID: 20499336

Abstract

While female mice do not have the equivalent of a menopause, they do undergo reproductive senescence. Thus, to dissociate the effects of aging versus estrogen deficiency on age‐related bone loss, we sham‐operated, ovariectomized, or ovariectomized and estrogen‐replaced female C57/BL6 mice at 6 months of age and followed them to age 18 to 22 months. Lumbar spines and femurs were excised for analysis, and bone marrow hematopoietic lineage negative (lin–) cells (enriched for osteoprogenitor cells) were isolated for gene expression studies. Six‐month‐old intact control mice were euthanized to define baseline parameters. Compared with young mice, aged/sham‐operated mice had a 42% reduction in lumbar spine bone volume/total volume (BV/TV), and maintaining constant estrogen levels over life in ovariectomized/estrogen‐treated mice did not prevent age‐related trabecular bone loss at this site. By contrast, lifelong estrogen treatment of ovariectomized mice completely prevented the age‐related reduction in cortical volumetric bone mineral density (vBMD) and thickness at the tibial diaphysis present in the aged/sham‐operated mice. As compared with cells from young mice, lin– cells from aged/sham‐operated mice expressed significantly higher mRNA levels for osteoblast differentiation and proliferation marker genes. These data thus demonstrate that, in mice, age‐related loss of cortical bone in the appendicular skeleton, but not loss of trabecular bone in the spine, can be prevented by maintaining constant estrogen levels over life. The observed increase in osteoblastic differentiation and proliferation marker gene expression in progenitor bone marrow cells from aged versus young mice may represent a compensatory mechanism in response to ongoing bone loss. © 2010 American Society for Bone and Mineral Research.

Keywords: aging, estrogen, bone, mice, ovariectomy

Introduction

Aging is associated with significant bone loss in women and in men,1 and considerable work over the past several decades has established the importance of estrogen (E) deficiency following menopause in the pathogenesis of bone loss in postmenopausal women.1 It is now known that E acts directly on osteoblast lineage cells,2 osteoclasts,3, 4 and other bone marrow cells such as T and Bcells5, 6 to inhibit the production of proresorptive cytokines such as tumor necrosis factor α (TNF‐α), interleukin 1 (IL‐1), IL‐6, monocyte colony stimulating factor (M‐CSF), prostaglandin E2 (PGE2), and receptor activator of NF‐κB ligand (RANKL) (for review, see ref. 7) and to enhance the production of antiresorptive factors such as transforming growth factor β (TGF‐β)8 and osteoprotegerin (OPG).9 While bone formation at the tissue level also increases following E deficiency,1 this increase is insufficient to keep pace with the increase in bone resorption, leading to a marked imbalance between bone resorption and bone formation. This is manifested by a rapid phase of bone loss following menopause in women, particularly in trabecular bone.1 However, recent cross‐sectional and longitudinal studies using quantitative computed tomography (QCT) have demonstrated the onset of trabecular bone loss as early as the third decade in women and in men, well before the onset of sex steroid deficiency; by contrast, decreases in cortical bone seem to coincide in women with the onset of menopause.10, 11 These data suggest that in humans, trabecular bone loss, although accelerated by E deficiency, may be largely independent of E, whereas cortical bone loss appears to be linked more closely to E deficiency.

While female mice do not have the equivalent of a menopause, they do undergo reproductive senescence, becoming essentially acyclic by 11 to 16 months of age.12, 13 This is accompanied by significant reductions in circulating estradiol (E2) levels, although in contrast to humans,1 the changes in circulating E2 levels in aging female mice are more difficult to detect. Thus Nelson and colleagues14 found that while serum E2 levels in aging (10 to 12.5 month of age) C57BL/6J mice on day 1 (proestrus) and day 2 (estrus) of the estrus cycle were similar to those found in younger mice (5.5 to 7.5 months of age), E2 levels on day 3 and the preovulatory E2 rise beginning on day 4 were reduced in older mice by 80% and 45%, respectively. Thus female aging mice do develop E deficiency, although this is clearly not as profound as that observed in postmenopausal women. Interestingly, age‐related declines in vertebral and femoral trabecular bone begin in female mice by 2 months of age,15 at a time of sex steroid sufficiency, again suggesting that trabecular bone loss in mice, as in humans, may be relatively independent of sex steroids.

Recognizing that aging female mice develop only mild to moderate E deficiency manifested at certain days of the estrus cycle,14 we sought to dissociate, as definitively as possible, the effects of aging per se versus those of age‐related E deficiency in mice on trabecular versus cortical bone loss over life. To do so, we compared the skeletal phenotype of young control mice (6 months of age) with that of normally aged mice (to 18 to 22 months of age), aged mice with lifelong E deficiency (ovariectomized at 6 months of age and aged to 18 to 22 months), and aged mice who had been ovariectomized at the age of 6 months but continuously replaced with fixed doses of E between the ages of 6 months and 18 to 22 months. Thus, by maintaining relatively constant E levels over life in the latter group, we essentially eliminated the possible effects of even subtle decrements in E levels with aging in female mice and isolated the effects of aging alone on the various skeletal parameters. In addition, in order to better understand possible changes in osteoprogenitor cells with aging, we harvested hematopoietic lineage–negative (lin–) cells from bone marrow of young and aged mice for analysis of osteoblastic genes and pathways. These cells have been shown recently to be highly enriched for osteoprogenitor cells that mineralize in vitro, form bone in vivo, and express bone‐related genes,16 thereby providing a useful cell population for evaluation of the effects of aging on osteoblast progenitor cells.

Methods

Overall study design

We studied five groups of female C57/BL6 mice. Group 1 (Control) consisted of young (6‐month‐old) animals; group 2 (aged/sham‐operated) underwent sham surgery at age 6 months and then were allowed to age to 18 to 22 months; group 3 (aged/OVX) underwent ovariectomy at age 6 months and then were allowed to age to 18 to 22 months; group 4 (aged/OVX/E10) underwent ovariectomy at age 6 months and between the ages of 6 and 18 to 22 months were provided with fresh E2 (90‐day) pellets every 60 days at a dose of 10 µg/kg per day (Innovative Research of America, Sarasota, FL, USA) implanted near the right shoulder blade in order to maintain as constant E2 levels as possible over life; and group 5 (aged/OVX/E40) were identical to group 4 except that they received E2 pellets at a dose of 40 µg/kg per day. The doses of E2 were selected based on a previous dose‐response study from our group17 demonstrating that these doses span the physiologic range for skeletal responses to E in female C57/BL6 mice. However, for every skeletal endpoint examined, changes in the two E groups (relative to young control mice) were virtually identical, so all data are presented for the two E groups combined. For technical and cost reasons, specific analyses were done in random subsets of the control and aged groups; thus the number of animals used in each analysis are indicated in the respective figures and tables.

The animals were housed in a temperature‐controlled room (22 ± 2°C) with a daily 12‐h light/12‐h dark schedule. During the experiments, animals had free access to water and were pair‐fed. At the end of the study, the animals were euthanized by cervical dislocation, and the bones were harvested (see below). The Institutional Animal Care and Use Committee approved all animal procedures.

Body composition and BMD measurements

Mice were anesthetized with Avertin (2,2,2‐tribromoethanol, 720 mg/kg, i.p.) and placed in a prone position on an animal tray. The dual‐energy X‐ray absorptiometry (DXA) measurements for total‐body fat and lean mass were carried out using a Lunar PIXImus densitometer (software Version 1.44.005, Lunar Corp., Madison, WI, USA).

Peripheral QCT measurements were performed with the mice placed in a supine position on a gantry using the Stratec XCT Research SA Plus using software Version 5.40 (Norland Medical Systems, Inc., Fort Atkinson, WI, USA). Slice images were measured 1 mm proximal to the tibia‐fibula junction. For cortical bone, the threshold was set at 710 mg/cm3; the coefficient of variation (CV) was 3.9%.

Micro–computed tomography (µCT)

The quantitative analysis of trabecular bone at the spine and distal femoral metaphysis was done by µCT (µCT20, Scanco Medical AG, Basserdorf, Switzerland) at a resolution of 7 µm per slice, and at least 100 slices were morphed (segmentation: 0.8/1/220). The femoral scans were carried out in an area corresponding to the secondary spongiosa in the metaphyseal region, 0.5 mm proximal to the growth plate. Three‐dimensional analysis was conducted to calculate morphometric parameters defining trabecular bone mass and microarchitecture, including bone volume/total volume (BV/TV), trabecular number (Tb.N), thickness (Tb.Th), and separation (Tb.Sp). Cortical porosity at the femur was calculated using the formula 1 – BV/TV, after extracting out all the trabecular elements using appropriate thresholds, and values were normalized to those in the control mice.

Bone histomorphometry

The femurs and lumbar spines (L1–L4) were processed for histomorphometry as described previously.18, 19 Briefly, dehydrated specimens were embedded in glycol methyl methacrylate, sectioned, and stained with the Goldner's stain. For the femoral bone, the region of interest (ROI) was located in the distal femoral metaphysis 250 µm away from the growth plate, spanning a total area of 1.25 mm2, which included both trabecular bone and bone marrow. For the vertebral sections, the measurements were carried out in fields situated in the center of the bone, at least 250 µm away from the cranial and caudal growth plates. The total area scanned was 1.25 mm2. The areas measured for both the femurs and vertebrae were kept consistent between samples, and the sections were read by a single investigator in a blinded fashion. Osteoblast number was defined as the number of osteoblasts per millimeter of cancellous perimeter and the osteoclast number as the number of multinucleated osteoclasts on eroded surfaces per millimeter of cancellous perimeter. All histomorphometric measurements were performed with the Osteomeasure Analysis System (Osteometrics, Atlanta, GA, USA).

Gene expression analysis

Bone marrow cells were flushed from the femurs and processed as follows for gene expression analyses: Following Ficoll extraction, cells were magnetically labeled with a mouse hematopoietic lineage cell depletion kit (Miltenyi Biotec GmbH, Bergish Gladbach, Germany) containing antibodies to CD5, CD45R (B220), CD11b, Gr‐1 (Ly‐6G/C), 7‐4, and Ter‐119, according to the manufacturer's instructions. After one wash, the cell suspension was loaded onto an autoMACS cell sorter (Miltenyi Biotec GmbH). MACS‐sorted lin– cells from bone marrow then were stored in RNAlater (RLT) buffer (Qiagen, Hilden, Germany) at −80°C for later extraction of RNA, as described below. Total RNA was isolated using spin columns (Microcolumns, Qiagen, Valencia, CA, USA). DNase treatment to digest all genomic DNA that could lead to false‐positive gene expression results was done following RNA isolation using Turbo DNA‐free DNase (Ambion, Austin, TX, USA). The RNA quantity and purity were measured with a Nanodrop spectrophotometer (Thermo Scientific, Wilmingon, DE, USA), and RNA integrity was determined using the Agilent 2100 Bioanalyzer (Santa Clara, CA, USA). Because the overall number of lin– cells was low and therefore the yield of total RNA was limited to run in‐depth gene expression analyses, we used the WT‐Ovation Pico RNA amplification system (NuGEN, Technologies, Inc., San Carlos, CA, USA) to synthesize microgram quantities of amplified cDNA starting with total RNA input amounts of approximately 50 ng. In this linear amplification system, the relative representation of each transcript species in the original sample is maintained during and after amplification.20, 21 For the QPCR analyses, we designed primers using the Primer Express Program (Applied Biosystems, Foster City, CA, USA). Primer sequences for any of the genes analyzed in this article are available on request. The primers (1.8 pmol per reaction, MWG‐Biotech AG, Ebersberg, Germany) were applied to 384‐well PCR plates in a sucrose solution (30% sucrose, 1 mM Tris, pH 8.5) and dried onto the plates for 1 hour in a heated speed vacuum. The PCR reactions then were run in the ABI Prism 7900HT Real Time System (Applied Biosystems) using SYBR Green (BioRad, Hercules, CA, USA) as the detection method. Five nanograms of the amplified cDNA was used per reaction, and each gene was run in triplicate in a total volume of 10 µL. Negative reverse‐transcriptase controls were included to check for genomic DNA contamination. Normalization for variations in input RNA was performed using a panel of 10 housekeeping genes (18S, G6PDH, GAPDH, HPRT, L13a, RPII, TBP, α‐tubulin, β2‐microglobulin, and β‐actin), with the geNorm algorithm22, 23 used to select the 3 to 4 most stable housekeeping genes from the 10 on the plate. The PCR Miner algorithm24 was used to correct for variations in amplification efficiencies.

Statistical analysis

For comparisons between groups, we used an ANOVA; where this was significant, the Fisher protected least significant difference test was used for pairwise comparisons between groups. All data are presented as mean ± SEM, with the exception of the gene expression data, which were not normally distributed. These are shown as the median, interquartile (25th to 75th percentile) range (IQR), and pairwise comparisons for gene expression between the control and aged/sham‐operated groups were done using the Wilcoxon rank sum test. We also used the O'Brien umbrella test25 to assess whether changes in gene expression occurred along a priori defined pathways (osteoblast differentiation, proliferation, and apoptosis markers, Wnt target genes, and bone morphogenetic protein target genes) based on knowledge of cellular signaling. This method provides a more robust means to analyze gene expression data and increases the power of detecting changes in genes occurring in prespecified clusters rather than in isolation. p < .05 was considered significant.

Results

Body weights and composition

Figure 1 shows the body weight and composition data in the various groups. Body weights were higher in all the aged groups compared with the young control mice, but the differences were not statistically significant (Fig. 1 A, ANOVA p = .245). Lean mass by DXA, however, was significantly greater in all the aged groups compared with the young control group (Fig. 1 B). Interestingly, while the aged/sham‐operated mice had significantly greater total‐body fat mass than the young controls (with a similar trend in the aged/OVX mice), the aged/OVX/E group had a total‐body fat mass that was identical to that of the young control mice (Fig. 1 C), indicating that the presence of constant E levels over life prevented age‐related increases in fat mass.

Figure 1.

Figure 1

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(A) Body weight, (B) lean mass by DXA, and (C) fat mass by DXA in the study groups. ANOVA p values were .245, <.001, and .024 for body weight, lean mass, and fat mass, respectively; when the ANOVA was significant, pairwise comparisons between groups were done using the Fishers protected least significant difference test, and p values are as noted; n = 8 to 12 per group.

Trabecular bone at the spine

Figure 2 shows the structural trabecular bone data at the spine analyzed by µCT. As compared with the young control mice, BV/TV was 42% lower in the aged/sham‐operated mice and 61% lower in the aged/OVX mice (Fig. 2 A), confirming previous findings of significant trabecular bone loss over life in mice.15 Interestingly, however, BV/TV was identical in the aged/OVX/E group to that in the aged/sham‐operated group and significantly lower than in the young control mice. Similar findings were noted for changes in Tb.N, Tb.Th, and Tb.Sp (Fig. 2 B–D); in each case, the aged/OVX/E group was identical to the aged/sham‐operated group and significantly different from the young control animals. These data thus clearly demonstrate that age‐related trabecular bone loss at the spine present in the aged/sham‐operated group was unaltered by maintaining constant E levels over life.

Figure 2.

Figure 2

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Trabecular parameters in the study groups analyzed using µCT at the lumbar spine. (A) BV/TV, (B) Tb.N, (C) Tb.Th, and (D) Tb.Sp. ANOVA p values were .003, .002, .048, and .036 for BV/TV, Tb.N, Tb.Th, and Tb.Sp, respectively; pairwise comparisons between groups were done using the Fishers protected least significant difference test, and p values are as noted; n = 5 to 10 per group.

Cortical bone measurements

As noted earlier, since data from human studies have suggested that trabecular bone loss over life, although accelerated by E deficiency, may be largely independent of sex steroids, whereas the onset of cortical bone loss is closely associated with E deficiency following menopause,10, 11 we contrasted the preceding changes in trabecular bone at the spine with those present in cortical bone at the tibial diaphysis. As shown in Fig. 3, cortical volumetric bone mineral density (vBMD; Fig. 3 A), area (Fig. 3 B), and thickness (Fig. 3 C) at the tibial diaphysis were significantly lower in the aged/sham‐operated and aged/OVX mice than in the young control mice. In contrast to the spine trabecular bone findings, however, maintaining constant E levels over life in the aged/OVX/E mice resulted in all the cortical bone parameters being identical to those present in the young control mice and significantly greater than those in the aged/sham‐operated (or aged/OVX) mice (Fig. 3), demonstrating that maintaining constant E levels over life can prevent “age‐related” cortical bone loss in mice. We further corroborated these findings at the tibial diaphysis by examining changes in cortical porosity by µCT at the femur. This increased by 23% (p = .023) and 27% (p = .008) in the aged/sham‐operated and aged/OVX mice compared with the young control mice, respectively, but only by 8% (p = .316 versus young controls) in the aged/OVX/E mice.

Figure 3.

Figure 3

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Cortical parameters in the study groups analyzed by pQCT at the tibial diaphysis. (A) Cortical vBMD, (B) cortical area, and (C) cortical thickness. ANOVA p values were .002, .002, and .001 for cortical vBMD, area, and thickness, respectively; pairwise comparisons between groups were done using the Fishers protected least significant difference test, and p values are as noted; n = 7 to 12 per group.

Trabecular bone at the femoral metaphysis

While our primary aim was to compare changes in trabecular bone at the spine with those present in cortical bone at the tibial diaphysis, we also examined trabecular bone at the femoral metaphysis in the four groups. As shown in Table 1, while the changes in trabecular bone volume and structural parameters at this site were similar to those observed at the lumbar spine in the aged/sham‐operated and aged/OVX mice compared with the young control animals, maintaining constant E levels over life in the aged/OVX/E mice resulted in marked increases in BV/TV, Tb.N, and Tb.Th at this site. Given this aberrant response to E in trabecular bone at the femoral metaphysis, it was not possible to use our model to dissociate possible effects of aging versus E deficiency on age‐related trabecular bone loss at this site.

Table 1.

Trabecular Parameters in the Study Groups Analyzed Using µCT at the Femoral Metaphysis

Control Aged/sham‐operated Aged/OVX Aged/OVX/E ANOVA
BV/TV, % 2.42 ± 0.89 0.38 ± 0.18 0.35 ± 0.20 23.3 ± 4.6*,**,*** <.001
Tb.N, /mm 1.75 ± 0.09 1.23 ± 0.22 0.69 ± 0.24 4.74 ± 0.80*,**,*** .002
Tb.Th, mm 0.05 ± 0.003 0.05 ± 0.01 0.04 ± 0.01 0.08 ± 0.01*,**,*** .002
Tb.Sp, mm 0.59 ± 0.03 0.89 ± 0.15 0.83 ± 0.28 0.30 ± 0.06**,*** .009
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ANOVA p values are indicated; when the ANOVA was significant, pairwise comparisons between groups were done using the Fishers protected least significant difference test.

*

p < .01 versus the control group.

**

p < .01 versus the aged/sham‐operated group.

***

p < .01 versus the aged/OVX group; n = 5 to 10 per group.

Osteoblast and osteoclast parameters

Table 2 shows histologic data for osteoblast and osteoclast parameters in the four groups in trabecular bone at the lumbar spine and femoral metaphysis. Overall, these showed reductions in osteoblast surface/bone surface (ObS/BS) and in osteoblast numbers/bone perimeter (NOb/BPm) in all the aged groups compared with the young control mice; no consistent changes were noted for osteoclast surface/bone surface (OcS/BS) or osteoclast numbers/bone perimeter (NOc/BPm). Thus, while E treatment is known to reduce osteoclast numbers in mice,26 we did not observe such a reduction in our model. The reasons for this discrepancy are unclear but may have to do with the duration of E treatment. Most previous studies of E effects on osteoclast numbers have been done using 1 to 2 months of E treatment26; by contrast, the essentially lifelong treatment with E in the aged/OVX/E group may have had different effects on osteoclast numbers in our study.

Table 2.

Osteoblast and Osteoclast Numbers at the Lumbar Spine and Femoral Metaphysis in the Study Groups

Control Aged/sham‐operated Aged/OVX Aged/OVX/E ANOVA
Lumbar spine
 Osteoblast surface/bone surface (ObS/BS), % 13.4 ± 2.1 6.5 ± 0.9** 8.6 ± 1.6 8.0 ± 1.4* 0.029
 Osteoblast number/bone perimeter (NOb/BPm), /mm 14.8 ± 2.4 7.9 ± 1.0* 11.0 ± 1.7 9.5 ± 1.7* 0.068
 Osteoclast surface/bone surface (OcS/BS), % 9.0 ± 1.0 8.3 ± 1.2 5.3 ± 1.0 7.5 ± 1.8 0.468
 Osteoclast number/bone perimeter (NOc/BPm), /mm 3.6 ± 0.4 3.3 ± 0.5 2.4 ± 0.5 2.5 ± 0.4 0.177
Femoral metaphysis
 Osteoblast surface/bone surface (ObS/BS), % 29.2 ± 4.8 6.7 ± 2.1*** 7.8 ± 2.7*** 13.5 ± 2.4*** <0.001
 Osteoblast number/bone perimeter (NOb/BPm), /mm 30.4 ± 3.7 8.0 ± 2.9*** 10.4 ± 3.7*** 15.6 ± 2.6** <0.001
 Osteoclast surface/bone surface (OcS/BS), % 7.3 ± 1.1 8.5 ± 3.3 9.4 ± 4.2 7.3 ± 2.1 0.936
 Osteoclast number/bone perimeter (NOc/BPm), /mm 3.1 ± 0.5 4.6 ± 1.4 3.3 ± 1.6 3.0 ± 0.6 0.616
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ANOVA p values are indicated; when the ANOVA was significant, pairwise comparisons between groups were done using the Fisher protected least significant difference test.

*

p < .05.

**

p < .01.

***

p < .001 versus the control group; n = 6 to 12 per group.

Comparison of gene expression in lin– cells from aged/sham‐operated versus young control mice

Since all the aged mice had reductions in osteoblast numbers on bone surfaces compared with the young control mice (Table 2), we harvested hematopoietic lin– cells, which have been shown recently to be highly enriched for osteoprogenitor cells,16 from the young control and aged/sham‐operated mice and examined the expression of key genes/gene pathways involved in bone metabolism in order to gain further insights into the possible mechanisms underlying this age‐related osteoblast deficit. As shown in Fig. 4, when analyzed as a group, markers for osteoblast differentiation (Fig. 4 A) and proliferation (Fig. 4 B), but not apoptosis (Fig. 4 C), overall were significantly higher in the aged/sham‐operated mice compared with the young control mice. This was accompanied by significantly higher levels of Wnt (Fig. 5 A) and bone morphogenetic protein (BMP; Fig. 5 B) target genes in the cells from the aged/sham‐operated versus young control mice. The cells from aged/sham‐operated mice also expressed higher levels of mRNA for ERα, ERβ, BMP receptors type IA and type II, and LRP5, but not LRP6 (Fig. 5 C). mRNA levels for BMP‐2 (data not shown), BMP‐4 (Fig. 5 A), and BMP‐6 (data not shown) were similar in the two groups of mice. However, there was a trend (p = .059) for mRNA levels of BMP‐7 to be higher in the aged/sham‐operated [median (IQR), 2.40 (1.54, 2.82)] than in the young control mice [1.00 (0.51, 1.28)]. We also tested for possible differences in expression of a number of Wnt genes (Wnt3a, ‐4, ‐7a, ‐7b, and ‐10b), but none of these mRNAs was detectable in the majority of the samples (data not shown).

Figure 4.

Figure 4

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Relative expression of mRNA levels for (A) osteoblast differentiation markers, (B) proliferation markers, and (C) apoptosis markers in the young control (C) and aged/sham‐operated (A/S) mice. The numerical p values are from the O'Brien umbrella test25 assessing changes in the genes within a cluster as a group. Data are shown as the median, interquartile (25th to 75th percentile) range. Note that for the panels to the right of the double‐dashed lines in panel A, the right‐hand scale should be used. *p < .05; **p < .01; ***p < .001 for pairwise comparisons versus the control group, n = 10 per group. AP = alkaline phosphatase; ON = osteonectin; OP = osteopontin; BSP = bone sialoprotein; PS = periostin; OCN = osteocalcin; cyc = cyclin.

Figure 5.

Figure 5

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Relative expression of mRNA levels for (A) Wnt target genes, (B) BMP target genes, and (C) ERα, ERβ, BMP receptor type IA, BMP receptor type II, LRP5, and LRP6. In panels A and B, the numerical p values are from the O'Brien umbrella test25 assessing changes in the genes within a cluster as a group. Data are shown as the median, interquartile (25th to 75th percentile) range. Note that for the panels to the right of the double‐dashed lines in panel A, the right‐hand scale should be used. *p < .05; **p < .01; ***p < .001 for pairwise comparisons versus the control group, n = 10 per group.

Discussion

While mice are a very useful model to study effects of age and E deficiency on the skeleton, the absence of a clear menopause and yet evidence for reproductive senescence with aging,12, 13 along with reductions in serum E2 levels during diestrus,14 leave open the question of whether age‐related bone loss in female mice is due to aging alone or to concomitant effects of some degree of age‐related E deficiency. Moreover, recent data in humans indicate that age‐related trabecular bone loss, while accentuated by E deficiency, may be largely independent of sex steroids, whereas cortical bone loss seems to be at least temporally closely associated with the onset of menopause.10, 11 Thus it is also important to define whether changes in trabecular versus cortical bone over life in mice have similar divergent relationships to E deficiency. We chose an approach that maintained relatively constant E levels over life to address this question and found that doing so in female mice failed to prevent age‐related trabecular bone loss and structural changes at the spine. By contrast, cortical vBMD, area, and thickness at the tibial diaphysis remained at levels found in young control mice in aged mice given lifelong E treatment. One interpretation of these findings is that in mice, as appears to be the case in humans,10, 11 age‐related trabecular bone loss occurs largely independent of E deficiency, whereas age‐related cortical bone loss depends on reductions in E levels.

While this is clearly the interpretation we favor, we recognize potential limitations of our model. Using subcutaneous E2 pellets we could not, for example, mimic the normally fluctuating E2 levels present in female mice over the course of the estrus cycle. While this limitation is virtually impossible to circumvent, previous work from our group17 has shown that over the dose range for slow‐release pellets of 10 to 40 µg/kg per day used in this study, ovariectomy‐induced bone loss is effectively prevented at multiple sites, arguing that the skeleton does not require the fluctuating E2 levels of the estrus cycle in order to respond to E. Moreover, both trabecular bone at the spine and cortical bone at the tibial diaphysis were exposed to the same E2 levels in the aged/OVX/E mice and yet had very different responses in terms of age‐related changes.

It is at present unclear why trabecular versus cortical bone might have different responses to E, but one explanation may come from studies by Bord and colleagues27 in developing human bone showing that whereas osteocytes and osteoblasts in cortical bone predominantly expressed estrogen receptor α (ERα), the same cells in trabecular bone expressed not only ERα but also significantly higher levels of ERβ than were present in cortical bone. While ERβ by itself may have little or no direct role in regulating bone turnover,28 its main role in fact may be to modulate the action of ERα because ERα/β heterdimers appear to be less sensitive to E than ERα homodimers.29, 30 Thus, if bone cells (ie, osteocytes, osteoblasts, and osteoclasts) in trabecular bone express more ERβ than present in these cells in cortical bone, trabecular bone cells would be less sensitive to all the actions of E on bone. As a consequence, even “young normal” E levels would not be able to fully prevent bone loss in this compartment, thereby leading to significant bone loss over life, independent of E levels. Consistent with this, Windahl and colleagues31 have found that female ERβ knockout mice are partially protected against age‐related trabecular bone loss. By contrast, if cells in cortical bone contained predominantly ERα, then bone loss in this compartment may be much more dependent on even the mild to moderate E deficiency present in aging female mice,12, 13, 14 and maintaining E levels throughout life would be sufficient to prevent cortical bone loss, as we observed at the tibial diaphysis. Clearly, however, further studies are needed to test this hypothesis. In addition, it is also possible that differential effects of E in trabecular versus cortical bone could be due to differential expression of another E receptor, distinct from ERα and ERβ. For example, recent studies have implicated a role for the G protein–coupled receptor 30 (GPR30) in nongenomic E signaling in human bone.32, 33 but it is at present unclear whether expression of GPR30 is different in trabecular versus cortical bone.

An alternative explanation for our findings could be that estrogen is more efficacious at preventing age‐related bone loss at a site of greater load bearing (ie, cortical bone at the tibial diaphysis) compared with the relatively non‐weight‐bearing trabecular bone in the spine. There is certainly considerable evidence supporting interactions between ER signaling and the effects of mechanical loading,34 so our observations could be related to differences in loading at the two sites examined.

In addition to prevention of cortical bone loss at the tibia, we also found that maintaining constant E levels over life prevented age‐related increases in fat mass. These findings are consistent with increasing evidence for a role for E in regulating peripheral fat depots. Thus female mice lacking either ERα or aromatase have increased total‐body and gonadal fat relative to wild‐type controls.35, 36 In addition, male ERα knockout mice have a near doubling of epididymal, perirenal, and inguinal white adipose tissue compared with their wild‐type litter mates by 1 year of age.37

Our gene expression data in bone marrow lin– cells showing an increase in expression of osteoblastic differentiation and proliferation markers as well as in BMP and Wnt target genes in the aged/sham‐operated mice compared with the young control mice are perhaps somewhat surprising in light of the marked reduction in osteoblast numbers we found on bone surfaces. We used methods identical to those used recently by the Aubin group16 to isolate bone marrow lin– cells, and these investigators have shown clearly that the bone marrow lin– population of cells is highly enriched for osteoprogenitor cells expressing bone‐related genes and capable of mineralization in vitro and bone formation in vivo in transplantation assays. The most plausible explanation for our findings is that there are compensatory changes in aging mice leading to increased BMP and Wnt signaling and osteoblast marker expression as well as proliferation in this marrow progenitor population. The fact that this does not translate into an increase in mature osteoblasts on bone surfaces may be due to some type of block in late differentiation of these cells and/or increased apoptosis of more mature cells with aging. This intriguing hypothesis clearly warrants further evaluation because it may help to identify the specific defect(s) leading to reduced osteoblast numbers and bone formation with aging.

In summary, using a model where we maintained relatively constant E levels over life, our data clearly demonstrate that age‐related trabecular bone loss at the lumbar spine is independent of any effects of E deficiency in female mice. Conversely, our findings are consistent with a role for E in maintaining cortical bone over life in this mouse model. We recognize potential limitations of our model, but our conclusions are further supported by the consistency of our findings in mice with data in humans on the patterns of trabecular versus cortical bone loss over life.10, 11 Finally, our gene expression data suggest additional hypotheses regarding a possible block in late differentiation or reduced survival of osteoblastic cells with aging that require further evaluation in mice and humans.

Disclosures

FS is currently an employee of Abbott Laboratories; however, this work was done while he was employed by Mayo Foundation. All the authors state that they have no conflicts of interest.

Acknowledgements

We would like to thank Elizabeth Atkinson, MS, for help with the statistical analyses using the O'Brien umbrella test, Kelly Hoey for technical and logistic support, and Glenda Evans for bone µCT scanning. This work was supported by NIH Grants AG028936 and AG004875.

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