Rosiglitazone impacts negatively on bone by promoting osteoblast/osteocyte apoptosis

in Journal of Endocrinology

Thiazolidinediones (TZDs) increase peripheral tissue insulin sensitivity in patients with type 2 diabetes mellitus by activating the nuclear receptor peroxisome proliferator-activated receptor γ (PPARγ). In bone marrow stromal cell cultures and in vivo, activation of PPARγ by high doses (20 mg/kg/day) of TZDs has been reported to alter stem cell differentiation by promoting commitment of progenitor cells to the adipocytic lineage while inhibiting osteoblastogenesis. Here, we have examined the in vivo effects of low-dose rosiglitazone (3 mg/kg/day) on bone, administered to mice by gavage for 90 days. Rosiglitazone-treated mice had increased weight when compared with controls, with no significant alterations in serum levels of glucose, calcium or parathyroid hormone (PTH). Bone mineral density (BMD) at the lumbar vertebrae (L1–L4), ilium/sacrum, and total body was diminished by rosiglitazone treatment. Histologically, bone was characterized by decreased trabecular bone volume and increased marrow space with no significant change in bone marrow adipocity. Decreased osteoblast number and activity due to increased apoptotic death of osteoblasts and osteocytes was apparent while osteoclast parameters and serum levels of osteocalcin, alkaline phosphatase activity, and leptin were unaltered by rosiglitazone treatment. Therefore, the imbalance in bone remodeling that follows rosiglitazone administration arises from increased apoptotic death of osteogenic cells and diminished bone formation leading to the observed decrease in trabecular bone volume and BMD. These novel in vivo effects of TZDs on bone are of clinical relevance as patients with type 2 diabetes mellitus and other insulin resistant states treated with these agents may potentially be at increased risk of osteoporosis.

Abstract

Thiazolidinediones (TZDs) increase peripheral tissue insulin sensitivity in patients with type 2 diabetes mellitus by activating the nuclear receptor peroxisome proliferator-activated receptor γ (PPARγ). In bone marrow stromal cell cultures and in vivo, activation of PPARγ by high doses (20 mg/kg/day) of TZDs has been reported to alter stem cell differentiation by promoting commitment of progenitor cells to the adipocytic lineage while inhibiting osteoblastogenesis. Here, we have examined the in vivo effects of low-dose rosiglitazone (3 mg/kg/day) on bone, administered to mice by gavage for 90 days. Rosiglitazone-treated mice had increased weight when compared with controls, with no significant alterations in serum levels of glucose, calcium or parathyroid hormone (PTH). Bone mineral density (BMD) at the lumbar vertebrae (L1–L4), ilium/sacrum, and total body was diminished by rosiglitazone treatment. Histologically, bone was characterized by decreased trabecular bone volume and increased marrow space with no significant change in bone marrow adipocity. Decreased osteoblast number and activity due to increased apoptotic death of osteoblasts and osteocytes was apparent while osteoclast parameters and serum levels of osteocalcin, alkaline phosphatase activity, and leptin were unaltered by rosiglitazone treatment. Therefore, the imbalance in bone remodeling that follows rosiglitazone administration arises from increased apoptotic death of osteogenic cells and diminished bone formation leading to the observed decrease in trabecular bone volume and BMD. These novel in vivo effects of TZDs on bone are of clinical relevance as patients with type 2 diabetes mellitus and other insulin resistant states treated with these agents may potentially be at increased risk of osteoporosis.

Keywords:

Introduction

Thiazolidinediones (TZDs) are a novel class of insulin-sensitizing antidiabetic agents. In the USA and Canada, two TZDs are indicated for use in type 2 diabetes mellitus, rosiglitazone and pioglitazone. A third, troglitazone, has been associated with significant hepatotoxicity and has been withdrawn from use. In clinical trials, all three TZDs effectively lower blood glucose levels in monotherapy and in combination therapy with sulfonylureas, metformin and insulin. TZDs improve glucose homeostasis by increasing insulin-stimulated glucose disposal in skeletal muscle (Petersen et al. 2000), although the insulin-sensitizing effects of TZDs on muscle appear to be indirect (Norris et al. 2003). The antidiabetic actions of TZDs are likely mediated by their interaction with the nuclear receptor peroxisome proliferator-activated receptor γ (PPARγ) (Lehmann et al. 1995, Hauner 2002). Adipose tissue is the major expression site for PPARγ (Tontonoz et al. 1994), while much lower levels are observed in other tissues, including other major insulin target tissues, such as liver and skeletal muscle.

PPARγ exists in two isoforms, PPARγ1 and PPARγ2, products of alternative promoter usage and different splicing (Zhu et al. 1995). Although the functional significance of these splice variants is not yet clear, PPARγ1 isoform is expressed at low levels in most tissues, whereas PPARγ2 is present selectively in fat tissue, where it plays a key role in regulating adipose tissue differentiation (Tontonoz et al. 1994, Shao & Lazar 1997, Rosen et al. 1999). It is now clear that PPARγ, expressed at or below the levels seen in fat tissue, can convert fibroblastic cells into fully differentiated adipocytes and can cause ‘transdifferentiation’ of cultured myoblasts to adipocytes (Hu et al. 1995). Moreover, the identification of TZDs as ligands for PPARγ has led to the observation that these compounds are potent and effective stimulators of adipogenesis in cells containing endogenous or ectopically expressed PPARγ (Lehmann et al. 1995).

The post-natal bone marrow of mammals harbors a population of stem/progenitor cells which are adherent, noncirculating and fibroblastic in nature. Stromal stem cells can give rise to all major skeletal tissues, such as cartilage, myelosupportive stroma, and fibrous connective tissue as well as associated adipocytes and osteoblasts, the bone forming cells (Friedenstein et al. 1968, Owen 1988, Kuznetsov et al. 2001). Commitment to a cell lineage depends on activation of transcription factors that simultaneously suppress gene transcription required for expression of the alternate cell lineage phenotype. In this respect, in vitro studies have indicated that activation of PPARγ2 impacts on marrow stem cell differentiation by increasing adipogenesis while decreasing osteoblastogenesis (Lecka-Czernik et al. 1999, 2002). Activated PPARγ2 does this by stimulating adipocyte fatty acid binding protein (aP2) mRNA expression, a marker of fully differentiated adipocytes, while inhibiting Runt-related transcription factor 2 (Runx2) expression, the main regulator of osteoblast differentiation, as well as suppressing the expression of other markers of osteoblast differentiation, such as γ1(I)collagen, alkaline phosphatase, osteocalcin and osteopontin (Lecka-Czernik et al. 1999). PPARγ2 activation may therefore act as a molecular switch between the osteogenic and adipogenic pathways.

Although the in vitro effects of TZDs on osteoblastogenesis are well recognized, the in vivo consequences of these compounds on bone turnover and bone mineral density (BMD) are less well studied and rather controversial. As a consequence of TZD administration, an increase in the number of bone marrow adipocytes at the expense of the number and differentiation potential of osteoblast progenitors could lead to suppression of bone formation and osteoporosis (Schwartz et al. 2002, Rzonca et al. 2004). Alternatively, improvement of glycemic control by TZDs has been associated with decreased bone turnover (Okazaki et al. 1999a) and increased BMD (Watanabe et al. 2003). Interestingly, decrease in bone turnover occurred prior to significant improvement in glucose metabolism, suggesting that TZDs may have direct effects on bone. It remains unclear, therefore, whether TZDs have a beneficial or detrimental effect on bone. Here, we sought to delineate the in vivo effects of administered low-dose rosiglitazone on the murine skeleton. We show that rosiglitazone impacts negatively on bone remodeling by promoting osteoblast and osteocyte apoptosis. Impairment in bone formation ensues leading to decreased trabecular bone volume and BMD.

Materials and Methods

Animals

All animal experiments were reviewed and approved by the institutional animal care committee. Male, non-diabetic C57BL/6 mice were purchased from Charles River (Wilmington, MA, USA) at 4 weeks of age and housed at controlled temperature and humidity with free access to regular chow and water. Following one week of acclimatization, animals were divided in two groups of 27 animals and were gavaged daily for a period of 90 days with either vehicle (0.25% carboxymethyl cellulose (medium viscosity) aqueous solution (5 ml/kg/day)) alone (control group) or vehicle with 3 mg/kg/day rosiglitazone maleate (Avandia, GlaxoSmithKline, King of Prussia, PA, USA) (rosiglitazone-treated group). Four mice from the control group died during the experimental period from causes unrelated to the treatment. Mice were weighed at baseline and at the indicated time intervals.

BMD measurements and microCT analysis

BMD (g/cm2) at the femur, tibia, humerus, ilium and sacrum, lumbar vertebrae (L1–L4), and total body (excluding the head) was measured at 45 and 90 days following initiation of treatment using a Lunar Piximus (GE Lunar, Madison, WI, USA) bone densitometer (variation in repeated measurements calculated to be ~2%). Animals were anesthetized with 240 mg/kg tribromoethanol (Avertin) prior to BMD measurement.

Lumbar vertebrae (L4) were dissected free of soft tissue, fixed overnight in 70% ethanol and analyzed with a SkyScan 1072 micro-CT scanner and associated analysis software (SkyScan, Antwerp, Belgium). Image acquisition was performed at 100 kV and 98 μA with a 0.9 ° rotation between frames. During scanning, the samples were enclosed in a tightly fitting rigid plastic rube to prevent movement. Thresholding was applied to the images to segment the bone from the background and the same threshold setting was used for all samples. 2D images were used to generate 3D reconstructions using the 3D Creator software supplied with the instrument.

Histology and immunohistochemistry

Mice were sacrificed by cervical dislocation. The lumber vertebral bodies were removed and fixed in PLP fixative (2% paraformaldehyde containing 0.075 M lysine and 0.01 M sodium periodate solution) for overnight at 4 °C and processed for histology, as previously described (Miao et al. 2001). Briefly, the lumbar vertebral bodies were decalcified in ethylene-diamine tetra-acetic acid (EDTA) glycerol solution for 5–7 days at 4 °C. Following dehydration and embedding in paraffin, 5 μm sections were cut on a rotary microtome. The sections were stained with hematoxylin and eosin (H&E), and histochemically for total collagen, alkaline phosphatase (ALP) activity and tartrate-resistant acid phosphatase (TRAP) activity. Total collagen was detected in the paraffin sections as described (Lopez-De Leon & Rojkind 1985), (with minor modifications). De-waxed sections were exposed to 1% sirius red in saturated picric acid for 1 h. After washing with distilled water, the sections were dehydrated and mounted with Biomount medium. Immunohistochemical staining for Bcl-2 (Santa Cruz Biotechnology Inc., Santa Cruz, CA, USA) was performed in paraffin-embedded sections of decalcified vertebral bodies using the avidin-biotin-peroxidase complex (ABC) technique.

Enzyme histochemistry for ALP activity was performed as previously described (Miao & Scutt 2002). Briefly, following preincubation overnight in 1% magnesium chloride in 100 mM Tris–maleate buffer (pH 9.2), de-waxed sections were incubated for 2 h at room temperature in a 100 mM Tris–maleate buffer containing naphthol AS-MX phosphate (0.2 mg/ml) (Sigma) dissolved in ethylene glycol monomethyl ether (Sigma) as substrate, and fast red TR (0.4 mg/ml) (Sigma) as stain for the reaction product. After washing with distilled water, the sections were counterstained with Vector methyl green nuclear counterstain (Vector Laboratories, Burlington, Ontario, Canada), and mounted with Kaiser’s glycerol jelly.

For TRAP enzyme histochemistry, de-waxed sections were pre-incubated 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) in dimethylformamide as substrate, and 0.5 mg/ml fast garnet GBC (Sigma) as 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 (Miao et al. 2002).

Computer-assisted image analysis

Following histochemical staining of sections, images of fields were photographed using a Sony digital camera. Images of micrographs from single sections were digitally recorded using a rectangular template, and recordings were processed using Northern Eclipse image analysis software, version 5.0 (Empix Imaging Inc., Mississauga, Ontario, Canada). For determining trabecular bone volume/total volume (BV/TV) in collagen stained sections, ALP positive area, and number and size of osteoclasts in TRAP histochemical stained sections, thresholds were set using green and red channels (Miao et al. 2001). Trabecular bone volume was measured in the lumbar vertebral bodies at x 25 magnification while ALP and TRAP parameters were measured in the trabecular region of lumbar vertebral bodies at x 100 magnification.

Detection of apoptotic cells

De-waxed paraffin sections were stained using an in situ cell death detection kit (Boehringer Mannheim, Indianapolis, IN, USA). Following treatment with 3 μg/ml of proteinase K for 20 min at room temperature, bone sections were incubated with a reaction mixture for terminal deoxynucleotidyltransferase-mediated dUTP nick-end-labeling of DNA strand breaks (TUNEL) method for 60 min at 37 °C. Sections were then incubated with Converter-AP (Boehringer Mannheim) for 30 min at 37 °C and alkaline phosphatase activity was visualized after 10–15 min of treatment with Fast Red TR/Naphthol AS-MX phosphate (Sigma), containing 1 mM levamisole to inhibit endogenous alkaline phosphatase activity. Sections were counterstained with methyl green and mounted with Kaiser’s glycerol jelly. The number of total apoptotic osteoblasts and osteocytes was counted manually using microscopy and the percentage of apoptotic cells was calculated.

Bone marrow cell cultures

Tibiae and femurs from mice treated with vehicle or rosiglitazone were removed under aseptic conditions and bone marrow cells were flushed out with DMEM containing 10% FCS, 50 μg/ml ascorbic acid, 10 mM β-glycerophosphate and 10−8 M dexamethasone. Cells were dispersed by repeated pipetting and a single cell suspension was achieved by forcefully expelling the cells through a 22-gauge syringe needle. Total bone marrow cells (106) were cultured in 36-cm2 dishes in 5 ml of the above-mentioned medium. The medium was changed every 4 days and cultures were maintained for 18 days. At the end of the culture period, cells were washed with PBS, fixed with PLP fixative and stained cytochemically for alkaline phosphatase (ALP). Following staining, ALP positive colony forming units (CFU-fALP) were counted manually.

Blood and serum biochemistry Following collection, blood glucose was measured using a Glucometer Elite kit (Bayer Canada, Inc., Toronto, Ontario, Canada) and serum samples were aliquoted and stored at −70 °C until assayed. Serum levels of osteocalcin were determined using an immunoradiometric assay (Immutopics, Inc., San Clemente, CA, USA), ALP activity using an absorbtivity kit (Sigma Diagnostics), calcium using a colorimetric determination kit (Sigma Diagnostics), intact parathyroid hormone (PTH) using an enzyme-linked immunosorbent assay (Immutopics, Inc., San Clemente, CA, USA), and leptin using an enzyme immunometric assay (Assay Designs Inc., Ann Arbor, MI, USA).

Statistics

BMD and histochemical results were analyzed by Graph-Pad Prism (GraphPad Software Inc., San Diego, CA, USA) using an unpaired t-test. Results were expressed as mean±s.e.m. The difference was considered statistically significant when P<0.05.

Results

Rosiglitazone increases body weight

For the entire duration of treatment, mice from both control- and rosiglitazone-treated groups gained weight (Fig 1A). At the beginning of the experiment, both groups were similar weight (19.70±0.21 g for the control group, 19.95±0.20 g for the rosiglitazone-gavaged group). However, following the fourth week of treatment, a weight difference became apparent, as the rosiglitazone-gavaged group gained more weight than control animals. By the sixth week of treatment the difference in weight became statistically significant (24.17±0.38 g and 25.23±0.25 g for the control group and the rosiglitazone treated group, respectively, P<0.05) and was maintained until the end of the 12 week treatment period (26.35±0.51 g and 27.31±0.33 g). The increase in weight for the rosiglitazone-treated group compared with the control group was 3.5% of body weight.

Rosiglitazone does not alter serum biochemistry

Blood and serum samples were collected following 90 days of treatment and assayed for glucose, calcium and PTH. As seen in Fig. 1B, no significant difference was observed in blood glucose levels (261.4±9.0 mg/dL and 255.5±5.9 mg/dL), levels of serum calcium (9.09±0.16 mg/dL and 8.91±0.15 mg/dL) and PTH (70.30±4.65 pg/mL and 74.39±4.45 pg/mL) between the control group and rosiglitazone-treated group, respectively.

Rosiglitazone treatment decreases BMD

BMD was measured at 45 and 90 days following initiation of treatment. At 45 days, no statistically significant difference in BMD between the two groups was observed. However, as shown in Fig. 2A, following 90 days of treatment a significant decrease in BMD measurements was observed at the lumbar vertebrae, ilium/sacrum and total body in the rosiglitazone-gavaged group compared with the controls. BMD at lumbar vertebrae (L1–L4) of rosiglitazone-treated mice was 7.08% lower than that of controls (0.05608±0.00093 g/cm2 vs 0.06035± 0.00132 g/cm2, respectively), 4.87% lower at the ilium/ sacrum (0.05505±0.00062 g/cm2 vs 0.05787± 0.00103 g/cm2) and 4.72% lower for total body (0.04823±0.00098 g/cm2 vs 0.05062±0.00112 g/cm2). Differences in measured BMD were also observed between the two groups at the tibia, femur and humerus, although they did not reach statistical significance.

Rosiglitazone impairs BMD accretion

A different evolution of BMD over time was apparent in the rosiglitazone-treated mice compared with controls. As shown in Fig. 2B, BMD increase over time had a steeper slope for the control group than the treated animals at the lumbar vertebrae (L1–L4) and ilium/sacrum. Measured at 45 and 90 days following treatment initiation, lumbar vertebral BMD increased from 0.05477 g/cm2 to 0.06035 g/cm2, an increase of 0.00558±0.00149 g/cm2, or 10.18% over 45 days, whereas for the rosiglitazone-treated group the lumbar vertebrae BMD progressed from 0.05401 g/cm2 to 0.05608 g/cm2, an increase of only 0.00207±0.00078 g/cm2 or 3.83% over the same time period. Similarly, ilium/sacrum BMD measured at 45 days and 90 days following treatment initiation went from 0.05245 g/cm2 to 0.05787 g/cm2 in the control group, which represents an increase of 0.00542±0.00105 g/cm2 or 10.33%, while in the case of the rosiglitazone-treated animals the change was less pronounced, increasing from 0.05224 g/cm2 to 0.05505 g/cm2; an increase of only 0.00281±0.00072 g/cm2 or 5.38%. These findings indicate that rosiglitazone administration negatively influences BMD accretion. Although similar changes were observed for BMD evolution at the femur, tibia, humerus and total body between the two groups, they did not reach statistical significance.

Rosiglitazone treatment decreases trabecular bone volume

To further investigate the observed BMD changes that result from 90 days of rosiglitazone treatment, histological sections were prepared from lumbar vertebrae. Shown in Fig 3A, histological sections of lumbar vertebra (L4) illustrate a loss in trabecular bone volume in the rosiglitazone-treated mice at three months when compared with the control group. In rosiglitazone- treated mice the trabeculae were thinner and an increase in trabecular separation was observed. In parallel, an increase in the bone marrow space was apparent. When quantified by histomorphometry, a statistically significant decrease of 22.6% in trabecular bone volume BV/TV was observed in the rosiglitazone-treated bone specimens compared with controls (Fig. 3B). Therefore, the observed BMD changes following rosiglitazone administration are in part due to a decrease in trabecular bone volume. Interestingly, a trend toward a corresponding increase in bone marrow fat content was apparent, although statistical significance was not attained (Fig. 3C). The observed alterations in trabecular bone content and architecture (trabecular number, thickness and separation) following rosiglitazone treatment were further confirmed by micro CT analysis (Table 1). Decreases in cortical bone volume were also apparent but did not attain statistical significance.

Rosiglitazone negatively influences bone formation

Decreases in bone volume can arise either from an increase in bone resorption or a decrease in bone formation, processes that are regulated primarily by osteoclasts and osteoblasts, respectively. We therefore sought to determine which of these processes was altered by rosiglitazone treatment. TRAP activity which is expressed by active osteoclasts and therefore serves as a marker of osteoclast activity, was similar in histochemically-stained bone sections from rosiglitazone-treated and control animals (Fig. 4A). Moreover, using computer-assisted histomorphometric analysis, no difference was observed between the two groups in the number of TRAP positive osteoclasts/field (58.2±3.2 for controls vs 62.3±3.3 for the rosiglitazone-treated group) and in the average size of TRAP positive osteoclasts (37.8±2.0 μm2 for control vs 39±0.9 μm2 for the rosiglitazone-treated group) (Fig. 4B).

In sharp contrast, histochemical staining for ALP activity, which localizes on the surface of osteoblasts and serves as marker of osteoblast activity, was decreased in the rosiglitazone-treated bone specimens when compared with the control group (Fig. 5A). In addition, the number of osteoblasts per tissue area was 51.6% lower in the rosiglitazone-treated group (45±4.6/mm2 vs 93±9.6/ mm2 for the control group), an observation that was corroborated by quantification of the total number of ALP-positive area/tissue area (1.61±0.10% for controls vs 0.34±0.05% for the rosiglitazone-gavaged group, P<0.001). Finally, in vivo labeling of bones with calcein demonstrated a marked decrease in mineral apposition rate (MAR), a parameter of bone formation, following rosigli-tazone treatment (0.81±0.03 for controls vs 0.59±0.02 for rosiglitazone-treated group, P<0.05) (Fig. 5B). These findings demonstrate that the observed decrease in trabecular bone volume and BMD following rosiglitazone administration arises primarily from its negative impact on parameters of bone formation, namely osteoblast number and function, rather than from an increase in bone resorption.

Low-dose rosiglitazone does not alter bone marrow stem cell commitment

To elucidate whether the observed decrease in bone formation arises in part from impaired commitment of bone marrow stem cells to the osteogenic lineage, ex vivo bone marrow cultures were prepared from control and rosiglitazone-treated mice. Compared with controls, rosiglitazone cultures repeatedly demonstrated an equivalent capacity to differentiate to CFU-fALP (~30% of total CFU-f) (results not shown). Therefore, low-dose rosiglitazone does not negatively influence bone formation by decreasing the commitment of bone marrow precursors toward osteogenic differentiation.

Increased osteoblast/osteocyte apoptosis due to rosiglitazone treatment

Next, we sought to identify an alternate mechanism by which osteoblast biology is deranged by low-dose rosiglitazone administration. Rosiglitazone could influence osteoblast/osteocyte survival, leading to the observed decrease in osteogenic cell number. Using TUNEL staining, an increased number of apoptotic osteoblasts and osteocytes were indeed observed in bone specimens from rosiglitazone-treated mice (Fig. 6A) and the percentage of apoptotic osteoblasts/osteocytes was increased nearly 5-fold compared with controls (Fig. 6B). Moreover, Bcl-2 immunoreactivity in osteogenic cells was diminished following treatment with rosiglitazone (Fig. 6C). Therefore, one way that rosiglitazone impacts negatively on bone formation is by decreasing Bcl-2 expression in osteoblasts and osteocytes thereby promoting their apoptotic death.

Rosiglitazone treatment does not alter serum markers

Serum samples collected following 90 days of treatment with rosiglitazone were assayed for ALP activity (57.84 ±3.16 U/L and 54.83±3.47 U/L), and osteocalcin (19.65±1.11 ng/mL and 20.50±0.54 ng/mL), which were similar in the two groups (Fig. 7). Serum levels of leptin, a marker for adipocyte function/differentiation, was also measured. Again, no significant difference was observed between control and rosiglitazone-treated animals (815.10±57.94 pg/mL and 801.90±63.48 pg/mL, respectively).

Discussion

In this study, we sought to delineate the in vivo consequences of low-dose rosiglitazone (3 mg/kg/day) administration on BMD and parameters of bone turnover in the murine skeleton. We demonstrate that a 3-month oral treatment with rosiglitazone in mice profoundly diminishes osteoblast/osteocyte number and activity by decreasing Bcl-2 expression and thereby promoting their apoptotic death. This leads to diminished bone formation and the observed decrease in trabecular bone volume and BMD. Our findings therefore raise the potential concern that long-term use of TZDs in patients with type 2 diabetes mellitus and other insulin-resistant states may have negative consequences on skeletal homeostasis, leading to decreased bone strength and propensity for fragility fractures.

In our study, blood glucose and biochemical parameters of calcium homeostasis were unaltered by TZD treatment. Interestingly, the weight of mice in both control and rosiglitazone-treated groups increased over time. This advocates that the animals were healthy and experiencing a normal growth pattern. Therefore, the changes that arose in bone as a consequence of rosiglitazone treatment were not due to nutritional deficiency or other health problems. In fact, following 90 days of treatment, the rosiglitazone-gavaged group had an increase in body weight 3.65% higher than the control group, concordant with what has been the clinical experience in patients with type 2 diabetes mellitus treated with TZDs. Indeed, administration of rosiglitazone has been associated with a weight gain of 0.7–3.5 kg in treated patients (Malinowski & Bolesta 2000). The weight gain may be due to increased deposition of subcutaneous fat (Kawai et al. 1999, Kelly et al. 1999), water retention (Niemeyer & Janney 2002) and increased food intake (Shimizu et al. 1998).

In vitro, TZDs promote bone marrow stromal cell adipogenesis at the expense of osteogenesis (Gimble et al. 1996, Lecka-Czernik et al. 1999). It was therefore expected that the compound’s capacity to activate PPARγ would diminish osteoblast differentiation while increasing bone marrow adipogenesis, as substantiated recently in vivo (Rzonca et al. 2004). Indeed, in our studies, osteoblast number and parameters of cell function were profoundly decreased by TZD administration. However, the anticipated increase in bone marrow adiposity was not apparent, although a trend toward it was evident. Moreover, we did not observe altered differentiation in ex vivo cultured bone marrow stromal cells from mice treated with low-dose rosiglitazone. Several explanations could account for our findings. First, reciprocal changes in adipogenesis and osteoblastogenesis are not an inevitable consequence of PPARγ2 activation. For example, rosiglitazone, the TZD that binds most efficiently to PPARγ2 (its affinity being nearly 10 times higher than that of troglitazone) (Camp et al. 2000), activates both proadipogenic and antiosteoblastogenic pathways. However, the thiazolidine acetamide ligand GW0072 [(±)-(2S,5S)-4-(4-(4-carboxyphenyl)butyl)-2-heptyl-4-oxo-5-thaizolidine N,N-dibenzyl-acetamide] is antiosteogenic without stimulating adiposity differentiation. Troglitazone on the other hand, induces proliferation and differentiation of committed adipocyte precursors without affecting the mesenchymal stem cells and osteoblast differentiation (Tornvig et al. 2001). Therefore, activation of PPARγ2 in vitro may stimulate adipocyte differentiation, suppress osteoblast differentiation or exhibit both activities, depending on the activating ligand (Lecka-Czernik et al. 2002). Second, adipose tissue at different sites may have distinct responses to TZDs. For instance, treatment with TZDs increases subcutaneous adipose tissue while decreasing intraabdominal fat accumulation (Adams et al. 1997, Mori et al. 1999). Third, the dose of rosiglitazone administered here may have been too low to significantly increase adipocyte differentiation in bone marrow stem cells but sufficiently high to induce osteoblast/osteocyte apoptosis. Indeed, the 3 mg/kg/day dose of rosiglitazone used here is much lower than the 20 mg/kg/day employed by Rzonca and co-workers (Rzonca et al. 2004) in their studies that demonstrated an increase in bone marrow adiposity. Therefore, the effects of TZDs on bone marrow adiposity may be dose-dependent. Finally, adipogenesis and osteo-blastogenesis may be independent processes, as previously suggested (Tornvig et al. 2001), whereby the increase in bone marrow adipogenesis is a passive process where adipocytes occupy the space left by the absent trabecular bone elements (Nuttall & Gimble 2000).

The mechanism by which TZDs alter osteoblast number and function is likely multifactorial and mediated through activation of PPARγ-dependent pathways, although mechanisms that are PPARγ-independent need also be considered. Previous studies have suggested that TZDs affect the early steps of osteogenic differentiation (Paulik & Lenhard 1997, Jackson & Demer 2000, Rzonca et al. 2004). Our finding that rosiglitazone treatment increases osteogenic cell apoptosis provides a novel explanation for the observed decrease in bone formation that follows administration of this compound in mice. Moreover, it substantiates previous in vivo evidence for the proapoptotic effect of TZDs on tumor cell growth (Elstner et al. 1998, Heaney et al. 2003) and in vitro reports showing that TZDs can induce apoptosis in diverse cell types including endothelial cells (Bishop-Bailey & Hla 1999), vascular smooth muscle cells (Okura et al. 2000), monocyte-derived macrophages (Chinetti et al. 1998) and various human cancer cells (Elstner et al. 1998, Takahashi et al. 1999, Chang & Szabo 2000, Eibl et al. 2001). Interestingly, the in vivo proapoptotic effects of rosiglitazone on pituitary tumor cells required the oral administration of high doses of rosiglitazone (20–50 mg/kg/day), while lower doses (5 mg/kg/day) were ineffective (Heaney et al. 2003). On the other hand, a significant reduction in the in vivo growth of MKN45 gastric cells in nude mice was reported after treatment with 6 mg/kg/day of rosiglitazone (Leung et al. 2004). This compares favorably with our observation that an oral dose of 3 mg/kg/day induced pronounced proapoptotic action on osteoblasts, indicating perhaps a higher sensitivity of these skeletal cells to the cellular actions of rosiglitazone. Although the molecular mechanisms by which TZDs induce apoptosis remain to be defined, our findings would indicate that activation of PPARγ leads to cell growth arrest and apoptosis, at least in part, by reducing Bcl-2 expression. This is corroborated by the recent observation that TZDs induce G0-G1 cell-cycle arrest and apoptosis (Heaney et al. 2003) by decreasing Bcl-2 and increasing Bax expression (Heaney et al. 2003, Yokoyama et al. 2003), and inducing transcription of the growth arrest and DNA damage-inducible 45 (GADD45) gene (Bruemmer et al. 2003). Further studies will be required to substantiate whether these additional molecular mechanisms underlie the proapoptotic effects of rosiglitazone on osteoblasts.

It is noteworthy that our findings with respect to serum leptin levels do not differ from what has been reported in other studies. TZDs transcriptionally inhibit leptin expression in adipocytes (Zhang et al. 1996) and serum leptin levels decrease (Boden et al. 2003, Watanabe et al. 2003) or remain unchanged (Satoh et al. 2003) following TZD administration in patients with type 2 diabetes mellitus. The change of BMD in these patients correlated negatively with that of serum leptin, a potent inhibitor of bone mass accretion (Ducy et al. 2000). Yet, percent changes in BMD were not significantly higher in nondiabetic subjects (Watanabe et al. 2003). It is possible that the dose of rosiglitazone administered here might have been too low to show a change in leptin expression. In rats, administration of rosiglitazone at doses of 2–5 mg/kg/day or higher decreased leptin mRNA levels in adipose tissue (De Vos et al. 1996). Yet, doses of 20 mg/kg/day in mice also failed to increase serum leptin levels, despite increases in bone marrow and brown fat content (Rzonca et al. 2004). Therefore, it is likely that a change in leptin expression does occur in adipose tissue, but is not reflected in circulating serum leptin levels.

Finally, our results do not support published in vitro findings suggesting that TZDs inhibit osteoclast formation (Okazaki et al. 1999b, 2000). As illustrated by TRAP staining and histomorphometric assessment of osteoclast number and surface area, rosiglitazone failed to alter these parameters under the in vivo conditions of our study. These observations could be explained in part by the type of TZD used (troglitazone vs rosiglitazone) and/or the effective TZD concentration reached at the level of the osteoclast. It would be anticipated that the observed changes in osteoblast parameters would produce parallel compensatory changes in osteoclast activity. Maintenance of appropriate bone mass depends upon the precise balance of bone formation and bone resorption. Balancing bone resorption and formation is centered on the ability of osteoblastic cells not only to form bone but to also regulate the rate of osteoclast differentiation (Suda et al. 1992). Here, the apparent unaltered nature of osteoclastic parameters may indeed be indicative of a persistent bone resorptive activity contributing to the observed decrease in bone mass. It is likely therefore, that rosiglitazone uncouples osteoblast–osteoclast cross talk during the bone remodeling process.

Controversy exists with regard to BMD changes in patients with type 2 diabetes mellitus (reviewed in Leidig-Bruckner & Ziegler 2001). On one hand, hyperinsulinemia and relatively high BMI are protective against bone loss in type 2 diabetes mellitus. On the other, increased calciuria and decreased osteoblastic function due to hyperglycemia may lead to deterioration of bone mass (Terada et al. 1998). Considering the likelihood that patients with type 2 diabetes mellitus may potentially be at a somewhat higher risk of developing osteoporosis, treatment with TZDs could increase this risk further. The risk of deleterious effects by these agents on bone would require further investigation in the clinical setting (Quesada-Gomez & Serrano-Alferez 2001). Nevertheless, our findings would indicate that more awareness of the risks could help in the prevention of osteoporosis in diabetic patients treated with TZDs. This could be accomplished by instituting timely and appropriate management for the prevention and treatment of osteoporosis in these patients. Of greater concern may be the advocated use of TZDs in young women with polycystic ovary syndrome and in individuals with other insulin resistance states (Day 1999) at a time of peak bone mass accretion. As illustrated by our study, the anticipated increase of BMD over time was profoundly hindered by rosiglitazone, suggesting that caution should be exercised when TZDs are administered in these groups of patients.

In summary, rosiglitazone administration in vivo leads to an imbalance in bone remodeling, in part by promoting osteoblast and osteocyte apoptosis. This in turn, has deleterious consequences on bone formation and ultimately on trabecular bone volume and BMD. These findings and the ever-increasing widespread use of TZDs in patients with type 2 diabetes mellitus and other insulin-resistant states, strongly argues for prospective clinical studies that will address the complex effects of this class of oral antidiabetic agents on skeletal homeostasis.

Table 1

Quantitative assessment by microCT of trabecular architecture and cortical bone thickness in vertebrae from control and rosiglitazone-treated mice

BV/TV (%)Tb.N (1/mm)Tb.Th (mm)Tb.Sp (mm)Cortical thickness (mm)
Tb.N, trabecular number; Tb.Th, trabecular thickness; Tb.Sp, trabecular spacing. The data are mean± s.e.m. after 90 days of treatment; *P<0.05 for rosiglitazone vs control (n=7 per group).
Control0.127±0.0153.89±0.120.056±0.0010.221±0.0090.175±0.004
Rosiglitazone0.091±0.013*3.12±0.11*0.047±0.003*0.275±0.008*0.169±0.003
Figure 1
Figure 1

Effects of rosiglitazone on body weight, blood glucose, and serum parameters of calcium homeostasis. (A) Rosiglitazone treatment increases body weight. The weight shown is the mean weight (g)± s.e.m., n=23 for the control group, n=27 for rosiglitazone-treated group; * P<0.05 at 45 and 90 days of treatment. (B) Effect of rosiglitazone on blood and serum biochemistry. There was no difference in blood glucose and serum levels of calcium and PTH between vehicle-treated control (open bars) and rosiglitazone-treated (solid bars) mice (n=12–22 and 14–22, respectively, depending on the assay).

Citation: Journal of Endocrinology 183, 1; 10.1677/joe.1.05723

Figure 2
Figure 2

Effect of rosiglitazone treatment on BMD. (A) BMD at the lumbar vertebrae (L1–L4), the ilium/sacrum, and total body BMD. The data shown are the mean BMD (g/cm2s.e.m., n=23 for control group and n=27 for rosiglitazone-treated group measured following 90 days of treatment; *P<0.05, ** P<0.01. (B) Evolution of BMD over time at the lumbar vertebrae (L1–L4) and ilium/sacrum. The data are the mean± s.e.m. BMD after 45 and 90 days of treatment; *P<0.05.□, control;•, rosiglitazone.

Citation: Journal of Endocrinology 183, 1; 10.1677/joe.1.05723

Figure 3
Figure 3

Vertebrae histology and histomorphometry (A) Histological sections of lumbar vertebra (L4) from control and rosiglitazone-gavaged animals. Sections were stained with H&E and are representative of 3–6 sections obtained from all lumbar vertebrae. (B) Histomorphometric analysis of bone volume/total volume (BV/TV), a measure of trabecular bone content, at L4. The data shown are the mean± s.e.m., n=6; * P<0.05. (C) Bone marrow fat content (%), as determined by densitometric analysis. The data shown are the mean± s.e.m., n=23 for control group and n=27 for the rosiglitazone group; P=0.12.

Citation: Journal of Endocrinology 183, 1; 10.1677/joe.1.05723

Figure 4
Figure 4

Rosiglitazone and osteoclasts. (A) Coloration of TRAP activity, a marker of osteoclast activity for control and rosiglitazone-gavaged groups. Upper panels, x 100; lower panels, x 400. Stained sections of lumbar vertebra (L4) are representative of eight determinations. (B) Computer-assisted image analysis of the number and size of TRAP positive osteoclasts per field. The data shown are the mean± s.e.m. of 12 readings.

Citation: Journal of Endocrinology 183, 1; 10.1677/joe.1.05723

Figure 5
Figure 5

Effects of rosiglitazone on osteoblast parameters. (A) Immunoenzymatic detection of alkaline phosphatase (ALP) activity for vehicle- and rosiglitazone-gavaged mice. Left panels, x 100; middle panels, x 400. Stained sections of lumbar vertebra (L4) are representative of six determinations. Right panels, in vivo calcein labeling of bones; distance between arrowheads represents the MAR. (B) Computer-assisted image analysis of osteoblasts per tissue area, ALP positive area per tissue area, and MAR. The data shown are the mean± s.e.m. of 11 readings. * P<0.05, ** P<0.01, *** P<0.001.

Citation: Journal of Endocrinology 183, 1; 10.1677/joe.1.05723

Figure 6
Figure 6

Osteogenic cell apoptosis following rosiglitazone administration. (A) Increased numbers of apoptotic osteoblasts and osteocytes were detected (arrows) following TUNEL staining in bone sections of lumbar vertebra (L4) from rosiglitazone-treated animals compared with control specimens. (B) The number of apoptotic osteogenic cells in rosiglitazone-treated and control bone sections was quantified. Twelve samples from each group have been used for image analysis, ***P<0.001. (C) Bcl-2 immunoreactivity in trabecular bone of lumbar vertebra (L4) from control and rosiglitazone-treated mice. Shown are micrographs from representative stained sections from three different animals in each group.

Citation: Journal of Endocrinology 183, 1; 10.1677/joe.1.05723

Figure 7
Figure 7

Serum markers of bone and adipose tissue metabolism. Serum levels of ALP activity and osteocalcin (bone formation markers), and leptin (adipocity marker) in vehicle- and rosiglitazone-treated mice. Shown are the mean± s.e.m., n=20–22 for control groups and rosiglitazone-treated groups, depending on the assay.

Citation: Journal of Endocrinology 183, 1; 10.1677/joe.1.05723

The technical assistance of H Li with the analysis of bones is greatly appreciated. This work was supported by grants from the Canadian Institutes of Health Research (CIHR). Support for MAS has been provided by the Medical Student Summer Research Bursary Award Program, Faculty of Medicine, McGill University. ACK is a recipient of the CIHR Scientist Award.

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    Effects of rosiglitazone on body weight, blood glucose, and serum parameters of calcium homeostasis. (A) Rosiglitazone treatment increases body weight. The weight shown is the mean weight (g)± s.e.m., n=23 for the control group, n=27 for rosiglitazone-treated group; * P<0.05 at 45 and 90 days of treatment. (B) Effect of rosiglitazone on blood and serum biochemistry. There was no difference in blood glucose and serum levels of calcium and PTH between vehicle-treated control (open bars) and rosiglitazone-treated (solid bars) mice (n=12–22 and 14–22, respectively, depending on the assay).

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    Effect of rosiglitazone treatment on BMD. (A) BMD at the lumbar vertebrae (L1–L4), the ilium/sacrum, and total body BMD. The data shown are the mean BMD (g/cm2s.e.m., n=23 for control group and n=27 for rosiglitazone-treated group measured following 90 days of treatment; *P<0.05, ** P<0.01. (B) Evolution of BMD over time at the lumbar vertebrae (L1–L4) and ilium/sacrum. The data are the mean± s.e.m. BMD after 45 and 90 days of treatment; *P<0.05.□, control;•, rosiglitazone.

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    Vertebrae histology and histomorphometry (A) Histological sections of lumbar vertebra (L4) from control and rosiglitazone-gavaged animals. Sections were stained with H&E and are representative of 3–6 sections obtained from all lumbar vertebrae. (B) Histomorphometric analysis of bone volume/total volume (BV/TV), a measure of trabecular bone content, at L4. The data shown are the mean± s.e.m., n=6; * P<0.05. (C) Bone marrow fat content (%), as determined by densitometric analysis. The data shown are the mean± s.e.m., n=23 for control group and n=27 for the rosiglitazone group; P=0.12.

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    Rosiglitazone and osteoclasts. (A) Coloration of TRAP activity, a marker of osteoclast activity for control and rosiglitazone-gavaged groups. Upper panels, x 100; lower panels, x 400. Stained sections of lumbar vertebra (L4) are representative of eight determinations. (B) Computer-assisted image analysis of the number and size of TRAP positive osteoclasts per field. The data shown are the mean± s.e.m. of 12 readings.

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    Effects of rosiglitazone on osteoblast parameters. (A) Immunoenzymatic detection of alkaline phosphatase (ALP) activity for vehicle- and rosiglitazone-gavaged mice. Left panels, x 100; middle panels, x 400. Stained sections of lumbar vertebra (L4) are representative of six determinations. Right panels, in vivo calcein labeling of bones; distance between arrowheads represents the MAR. (B) Computer-assisted image analysis of osteoblasts per tissue area, ALP positive area per tissue area, and MAR. The data shown are the mean± s.e.m. of 11 readings. * P<0.05, ** P<0.01, *** P<0.001.

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    Osteogenic cell apoptosis following rosiglitazone administration. (A) Increased numbers of apoptotic osteoblasts and osteocytes were detected (arrows) following TUNEL staining in bone sections of lumbar vertebra (L4) from rosiglitazone-treated animals compared with control specimens. (B) The number of apoptotic osteogenic cells in rosiglitazone-treated and control bone sections was quantified. Twelve samples from each group have been used for image analysis, ***P<0.001. (C) Bcl-2 immunoreactivity in trabecular bone of lumbar vertebra (L4) from control and rosiglitazone-treated mice. Shown are micrographs from representative stained sections from three different animals in each group.

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    Serum markers of bone and adipose tissue metabolism. Serum levels of ALP activity and osteocalcin (bone formation markers), and leptin (adipocity marker) in vehicle- and rosiglitazone-treated mice. Shown are the mean± s.e.m., n=20–22 for control groups and rosiglitazone-treated groups, depending on the assay.

  • AdamsM Montague CT Prins JB Holder JC Smith SA Sanders L Digby JE Sewter CP Lazar MA Chatterjee VK & O’Rahilly S 1997 Activators of peroxisome proliferator-activated receptor gamma have depot-specific effects on human preadipocyte differentiation. Journal of Clinical Investigation1003149–3153.

    • Search Google Scholar
    • Export Citation
  • Bishop-BaileyD & Hla T 1999 Endothelial cell apoptosis induced by the peroxisome proliferator-activated receptor (PPAR) ligand 15-deoxy-Δ1214-prostaglandin J2. Journal of Biological Chemistry27417042–17048.

    • Search Google Scholar
    • Export Citation
  • BodenG Cheung P Mozzoli M & Fried SK 2003 Effect of thiazolidinediones on glucose and fatty acid metabolism in patients with type 2 diabetes. Metabolism52753–759.

    • Search Google Scholar
    • Export Citation
  • BruemmerD Yin F Liu J Berger JP Sakai T Blaschke F Fleck E Van Herle AJ Forman BM & Law RE 2003 Regulation of the growth arrest and DNA damage-inducible gene 45 (GADD45) by peroxisome proliferator-activated receptor gamma in vascular smooth muscle cells. Circulation Research9338–47.

    • Search Google Scholar
    • Export Citation
  • CampHS Li O Wise SC Hong YH Frankowski CL Shen X Vanbogelen R & Leff T 2000 Differential activation of peroxisome proliferator-activated receptor-gamma by troglitazone and rosiglitazone. Diabetes49539–547.

    • Search Google Scholar
    • Export Citation
  • ChangTH & Szabo E 2000 Induction of differentiation and apoptosis by ligands of peroxisome proliferator-activated receptor γ in non-small cell lung cancer. Cancer Research601129–1138.

    • Search Google Scholar
    • Export Citation
  • ChinettiG Griglio S Antonucci M Torra IP Delerive P Majd Z Fruchart JC Chapman J Najib J & Staels B 1998 Activation of proliferator-activated receptors α and β induces apoptosis of human monocyte-derived macrophages. Journal of Biological Chemistry27325573–25580.

    • Search Google Scholar
    • Export Citation
  • DayC1999 Thiazolidinediones: a new class of antidiabetic drugs. Diabetic Medicine16179–192.

  • De VosP Lefebvre AM Miller SG Guerre-Millo M Wong K Saladin R Hamann LG Staels B Briggs MR & Auwerx J 1996 Thiazolidinediones repress ob gene expression in rodents via activation of peroxisome proliferator-activated receptor gamma. Journal of Clinical Investigation981004–1009.

    • Search Google Scholar
    • Export Citation
  • DucyP Amling M Takeda S Priemel M Schilling AF Beil FT Shen J Vinson C Rueger JM & Karsenty G 2000 Leptin inhibits bone formation through a hypothalamic relay: a central control of bone mass. Cell100197–207.

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