Increased Bone Adiposity and Peroxisomal Proliferator-Activated Receptor-2 Expression in Type I Diabetic Mice
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内分泌学杂志 2005年第8期
Departments of Physiology and Radiology (S.B., R.M., L.R.M.), Molecular Imaging Research Center (S.B., R.M., L.R.M.), and Department of Animal Science (M.O.), Michigan State University, East Lansing, Michigan 48824; and Department of Pathology, University of Kentucky (M.-C.F., H.M.), Lexington, Kentucky 40536
Address all correspondence and requests for reprints to: Dr. Laura R. McCabe, Department of Physiology, Michigan State University, 2201 Biomedical Physical Science Building, East Lansing, Michigan 48824. E-mail: mccabel@msu.edu.
Abstract
Decreased bone mass, osteoporosis, and increased fracture rates are common skeletal complications in patients with insulin-dependent diabetes mellitus (IDDM; type I diabetes). IDDM develops from little or no insulin production and is marked by elevated blood glucose levels and weight loss. In this study we use a streptozotocin-induced diabetic mouse model to examine the effect of type I diabetes on bone. Histology and microcomputed tomography demonstrate that adult diabetic mice, exhibiting increased plasma glucose and osmolality, have decreased trabecular bone mineral content compared with controls. Bone resorption could not completely account for this effect, because resorption markers (tartrate-resistant acid phosphatase 5b, urinary deoxypyridinoline excretion, and tartrate-resistant acid phosphatase 5 mRNA) are unchanged or reduced at 2 and/or 4 wk after diabetes induction. However, osteocalcin mRNA (a marker of late-stage osteoblast differentiation) and dynamic parameters of bone formation were decreased in diabetic tibias, whereas osteoblast number and runx2 and alkaline phosphatase mRNA levels did not differ. These findings suggest that the final stages of osteoblast maturation and function are suppressed. We also propose a second mechanism contributing to diabetic bone loss: increased marrow adiposity. This is supported by increased expression of adipocyte markers [peroxisome proliferator-activated receptor 2, resistin, and adipocyte fatty acid binding protein (P2)] and the appearance of lipid-dense adipocytes in diabetic tibias. In contrast to bone marrow, adipose stores at other sites are depleted in diabetic mice, as indicated by decreased body, liver, and peripheral adipose tissue weights. These findings suggest that IDDM contributes to bone loss through changes in marrow composition resulting in decreased mature osteoblasts and increased adipose accumulation.
Introduction
BONE IS A highly specialized and dynamic tissue that undergoes continual remodeling throughout its lifetime. Remodeling processes in bone are a sum of two activities: bone resorption and bone formation. Osteoblasts are responsible for bone formation, whereas osteoclasts resorb bone. An equilibrium between osteoblast and osteoclast activities maintains bone mass and quality. In contrast, alteration of one part of the equilibrium can lead to bone loss (decreased formation and/or increased resorption) or increased bone formation (decreased bone resorption and/or increased bone formation) (1, 2). Changes in this balance, leading to bone loss and osteoporosis, can occur under conditions such as aging, skeletal unloading/disuse, and disease (1, 2, 3, 4, 5).
Insulin-dependent diabetes mellitus (IDDM type I) is a chronic disease characterized by a lack of insulin production. In the absence of insulin, insulin-sensitive cells exhibit markedly reduced glucose uptake, resulting in increased serum glucose levels. Two of the long-term complications of IDDM are diabetic osteopenia and osteoporosis (6, 7, 8, 9, 10, 11, 12, 13, 14, 15, 16, 17, 18), which are associated with increased fracture rate (19, 20, 21, 22) and delayed fracture healing (23, 24, 25). Recent clinical studies found that 67% of men and 57% of women with IDDM suffer from osteopenia of the femoral neck and/or lumbar spine (9), and 14–20% of IDDM patients aged 20–56 yr met the criteria for more extensive bone loss, osteoporosis (9, 18). In diabetic patients, osteoblast function is significantly decreased and is thought to be associated with a maturation defect. This conclusion is based on the fact that serum levels of an early osteoblast marker, peptide of procollagen, remain normal in all types of diabetes, whereas serum levels of a late-stage marker of osteoblast maturation, osteocalcin, are decreased in patients at a range of ages (26). These pathological findings are even more pronounced in diabetic rats (27, 28, 29, 30), which also exhibit decreased bone volume and osteoid surfaces with no reduction in osteoclast number (27, 31) and significantly less osseointegration of bone implants than controls (32). These findings suggest that a decrease in osteoblast number and/or function/maturation could be a major contributor to bone loss in IDDM.
Osteoblasts are derived from mesenchymal stem cells. These pluripotent stem cells can give rise to adipocytes as well as a variety of other cell types (33, 34, 35, 36). During the process of osteogenesis and osteoblast differentiation, the expression of runx2, a transcription factor required for these processes (37, 38), is increased. Late-stage differentiation is marked by elevated osteocalcin expression (39, 40, 41). In contrast, overexpression of peroxisome proliferator-activated receptor 2 (PPAR2), a member of the nuclear receptor transcription factor family (42), induces adipogenesis over osteoblastogenesis in pluripotent cells (43). Interestingly, if PPAR2 is expressed in osteoblasts, it can suppress the mature osteoblast phenotype and induce genes associated with an adipocyte-like phenotype, such as aP2 (also called fatty acid-binding protein 4), fatty acid synthase, and lipoprotein lipase (43). Selection of adipogenesis over osteoblastogenesis is thought to contribute to bone loss associated with a variety of conditions, including age-related and disuse-associated osteoporosis (44, 45, 46, 47, 48).
Although studies have examined the influence of IDDM on human and rat bone, few have directly addressed the influence of diabetes on the skeletal system of mice, an animal model that is amendable to genetic manipulation. In this study we use a streptozotocin-induced IDDM mouse model to examine the influence of this disease on bone. Consistent with human and rat studies, we found a decrease in diabetic bone volume [histologically and by microcomputed tomography (μCT) analyses]. In addition, we found a decrease in osteocalcin mRNA levels, and we are the first to report an increase in the expression of adipocyte markers, PPAR2, resistin, and aP2, and an increase in visible adipocytes in IDDM compared with control tibias. In contrast to bone marrow, peripheral adipose tissue was decreased in size, suggesting that the marrow environment and adipose stores are regulated differently from peripheral sites. These findings support our hypothesis that IDDM causes a shift in skeletal composition marked by increased adipose storage and decreased mature osteoblasts.
Materials and Methods
Diabetic mouse model
Diabetes was induced in adult (15 wk old) male BALB/c mice (Harlem Laboratories, Houston, TX) by daily ip injection of streptozotocin (40 μg/g body weight in 0.1 citrate buffer), a pancreatic ?-cell cytotoxin, for 5 d (49, 50). Controls were injected with buffer alone. Seven days after the last injection, nonfasting blood glucose measurements were made using blood obtained from the lateral saphenous vein and a glucometer (Accu-Check instant, Roche, Indianapolis, IN). This day was considered d 0. Mice with blood glucose levels greater than 300 mg/dl were considered diabetic. Animals were maintained in metabolic cages, kept on a 12-h light, 12-h dark cycle at 23 C, and received food and water ad libitum. After 2 or 4 wk of confirmed diabetic condition, mice (controls and diabetics) were fasted for 6 h, then killed. Tibiae were immediately removed, freed from soft tissue, and either fixed in formalin (for histology and μCT analyses) or snap-frozen in liquid nitrogen and stored at –80 C (for RNA analyses). At 4 wk, the liver and sc femoral fat pads were dissected from the surrounding tissues. Tissues were weighed and snap-frozen in liquid nitrogen or fixed in PBS-buffered formalin. Animal studies were conducted in accordance with the Michigan State University all-university committee on animal use and care.
Plasma measurements
Blood was obtained from 2- and 4-wk-old mice when the animals were killed, and blood serum was prepared from each sample by centrifugation for 5 min at 3000 rpm. Serum was stored frozen at –20 C. The glucose concentration in serum samples was determined using a glucose assay kit (Sigma-Aldrich Corp., St. Louis, MO). Serum osmolality, defined as the expression of the total number of solute particles dissolved in 1 kg solvent (International Federation of Clinical Chemistry), was determined using a vapor pressure osmometer (Wescor, Inc., Logan, UT). Serum osteocalcin levels were measured using a mouse osteocalcin enzyme immunoassay kit (Biomedical Technologies Inc., Stoughton, MA) according to the manufacturer’s instructions. Quantitative determinations of serum glycerol, total triglycerides, and true triglyceride (expressed as equivalent triolein concentration) levels were performed using a serum triglyceride determination kit (Sigma-Aldrich Corp.).
Osteoclast activity-systemic measurements
For measurement of tartrate-resistant acid phosphatase (TRAP), serum from 5 d and 2, 3, and 4 wk postinduction diabetic and control mice was used. Active serum TRAP5b, produced by mouse osteoclasts, was measured according to the manufacturer’s protocol using a solid phase, immunofixed, enzyme activity assay, MouseTRAP (SBA Sciences, Turku, Finland).
For urinary deoxypyridinoline (DPD) measurements, mice were housed in metabolic cages to allow collection of 24-h urine samples (12, 13, and 14 d after confirmation of diabetes induction). Urine specimens were stored at –80 C. Urinary DPD and creatinine levels were determined using a Metra DPD enzyme immunoassay kit and Metra creatinine assay kit, respectively (Quidel Corp., San Diego, CA). DPD levels were expressed relative to creatinine levels.
RNA analysis
Tibiae were crushed under liquid nitrogen conditions using a Bessman tissue pulverizer (Spectrum Laboratories, Inc., Rancho Dominguez, CA), and RNA was extracted using the method of Chomczynski and Sacchi, as previously described (51, 52). RNA integrity was verified by formaldehyde-agarose gel electrophoresis. Synthesis of cDNA was performed by RT with 2 μg total RNA using the SuperScript II kit with oligo(deoxythymidine)12–18 primers as described by the manufacturer (Invitrogen Life Technologies, Inc., Carlsbad, CA). cDNA (1 μl) was amplified by PCR in a final volume of 25 μl using the iQ SYBR Green Supermix (Bio-Rad Laboratories, Hercules, CA) with 10 pmol of each primer (Integrated DNA Technologies, Coralville, IA). Osteocalcin was amplified using 5'-ACG GTA TCA CTA TTT AGG ACC TGT G-3' and 5'-ACT TTA TTT TGG AGC TGC TGT GAC-3' (53). Runx2 was amplified using 5'-GAC AGA AGC TTG ATG ACT CTA AAC C-3' and 5'-TCT GTA ATC TGA CTC TGT CCT TGT G-3' (54). Alkaline phosphatase was amplified using 5'-CGTAATCTACCATGGAGACATTTTC-3' and 5'-GACTGTGGTTACTGCTGATCATTC-3'. PPAR2 was amplified using 5'-TGA AAC TCT GGG AGA TTC TCC TG-3' and CCA TGG TAA TTT CTT GTG AAG TGC-3' (55). Adipocyte fatty acid-binding protein 2 (aP2) was amplified using 5'-GCG TGG AAT TCG ATG AAA TCA-3' and 5'-CCC GCC ATC TAG GGT TAT GA-3' (56). TRAP5 was amplified using 5'-AATGCCTCGACCTGGGA-3' and 5'-CGTAGTCCTCCTTGGCTGCT-3' (57). Resistin was amplified using 5'-GCTGCTGCCAAGGCTGAT-3' and 5'-TCTCCTTCCACCATGTAGTTTCC-3' (58). Cyclophilin, which was not modulated under diabetic conditions, was used as a control for RNA levels; it was amplified using 5'-ATT CAT GTG CCA GGG TGG TGA C-3' and 5'-CCG TTT GTG TTT GGT CCA GCA-3' (59, 60). Real-time PCR was carried out for 40 cycles using the iCycler (Bio-Rad Laboratories), and data were evaluated using iCycler software. Each cycle consisted of 95 C for 15 sec, 60 C for 30 sec (except for runx2 and osteocalcin, which had an annealing temperature of 65 C), and 72 C for 30 sec. RNA-free samples, a negative control, did not produce amplicons. Melting curve and gel analyses were used to verify single products of the appropriate size (in base pairs).
In vitro examination of streptozotocin on pluripotent C3H10T1/2 cells
C3H10T1/2 cells (61) were obtained from American Type Culture Collection (Manassas, VA) and maintained in Eagle’s Basal Medium (Sigma-Aldrich Corp.) with 10% fetal bovine serum (Atlanta Biologicals, Norcross, GA) at 37 C in a humidified atmosphere of 5% CO2/95% air. Cells were plated at a density of 100,000 cells per well of a six-well plate. To induce osteoblast differentiation, bone morphogenic protein 2 (Sigma-Aldrich Corp.) was added at a concentration of 100 ng/ml 24 h after plating (61). Cells were fed every other day with or without an additional 100 ng/ml bone morphogenic protein 2 and were grown for 14 d. Streptozotocin concentrations were 0.01, 0.1, 0.5, and 1.0 mg/ml and was added with every feeding. Cells were fed and treated 24 h before harvesting for RNA analyses or lipid staining.
Tissue and mineralized bone histology and histomorphometry
Proximal tibiae isolated from 4-wk-old control (n = 8) and diabetic (n = 11) mice were fixed in absolute ethanol at room temperature, dehydrated, and embedded in methylmethacrylate as previously described (62). For dynamic measurements, mice were injected ip with 0.2 ml 10 mg/ml calcein (Sigma-Aldrich Corp.) in sterile saline 6 and 2 d before the animals were killed. Sections of 3- and 7-μm thickness were cut with a Microm microtome (model HM360, Carl Zeiss, Inc., Thornwood, NY) and stained with the modified Masson-Goldner Trichrome stain (63). Static and dynamic parameters of bone structure, formation, and bone resorption were measured at standardized sites under the growth plate using the semiautomatic method (Osteoplan II, Kontron, Munich, Germany) at a magnification of x200 (64, 65). For adipose analyses, visible adipocytes (>50 μm) were counted. All parameters comply with the guidelines of the nomenclature committee of the American Society of Bone and Mineral Research (66).
Subcutaneous femoral fat depots were isolated from surrounding tissue and fixed in 10% neutral buffered formalin. Fixed samples were processed on an automated Thermo Electron Excelsior tissue processor for dehydration, clearing, and infiltration using a routine overnight processing schedule. Samples were then embedded in Surgipath embedding paraffin on a Sakura Tissue Tek II embedding center. Paraffin blocks were sectioned at 5 μm on a Reichert Jung 2030 rotary microtome (Leica Instruments GmbH, Nussloch, Germany). Slides were stained with hematoxylin and eosin.
Livers were dissected out, sectioned, and placed in Tissue Tek OCT freezing medium (Miles, Inc., Elkhart, IN) on a sectioned cork; corks were snap-frozen in liquid nitrogen. Frozen tissue corks were stored at –80 C. Tissues were then sectioned on a –20 C Sakura Tissue Tek cryostat at 10 μm. Sections were placed on adhesive slides, air-dried for 30 min, fixed in 37–40% formaldehyde for 1 min, rinsed in running tap water for 5 min, and stained with hematoxylin and eosin.
μCT analysis
Fixed bones and mouse legs were scanned using a GE Explore Locus μCT system (General Electric, Fairfield, CT) at a voxel resolution of 20 μm obtained from 720 views. The beam angle of increment was 0.5, and the beam strength was set at 80 kvp and 450 μA. Each run included control and diabetic bones and a calibration phantom to standardize grayscale values and maintain consistency. Based on autothreshold and isosurface analyses of multiple bone samples, a fixed threshold was used to separate bone from bone marrow. All cortical bone analyses were made in a defined 3-mm3 cube in the middiaphysis 1 mm proximal of the tibial-fibular junction. The polar cross-sectional moment (moment of inertia) was also determined at the tibial-fibular junction using the parallel axis theorem. Trabecular bone analyses were performed in a region of trabecular bone defined at 0.17 mm (1% of the total length) under the growth plate of the proximal tibia extending 2 mm toward the diaphysis and excluding the outer cortical shell. Bone mineral content, mineral density, and volume fraction values were computed with a GE Healthcare MicroView software application for visualization and analysis of volumetric image data.
Statistical analysis
All statistical analyses were performed using the Excel data analysis program (Microsoft Corp., Redmond, WA)) for t test analysis. Values are expressed as the mean ± SE.
Results
IDDM was induced in adult BALB/c mice, using a multiple (5 d) streptozotocin (40 μg/g body weight) injection procedure (49). Streptozotocin selectively destroys pancreatic B cells (50). One week later, nonfasting blood glucose measurements were taken from the saphenous vein to confirm diabetes at levels greater than 300 mg glucose/dl. Control (vehicle-injected) mice tightly maintained blood glucose levels near 180 mg/dl (178 ± 5.3), whereas streptozotocin-injected mice had blood glucose levels averaging 445 ± 27 mg/dl. Serum glucose levels remained elevated throughout the study (Fig. 1). Similarly, blood osmolality, determined by a vapor pressure osmometer, was significantly elevated in diabetic compared with control mice, averaging 325 mmol/kg compared with 302 mmol/kg, respectively (Fig. 1). This suggests that tissues and cells of diabetic mice are exposed to hyperosmotic conditions. In addition to the above analyses, we examined serum triglyceride levels. Diabetic mice exhibited a significant increase in triglyceride levels throughout the study, consistent with previous studies in diabetic mice and humans (67, 68). In contrast, body weights of diabetic mice were decreased throughout the time course of the study (Fig. 1).
FIG. 1. Diabetes increases serum glucose, serum osmolality, and serum triglycerides and is associated with decreased body weight. Serum from 5-, 14-, 21-, and 28-d diabetic and control BALB/c mice was analyzed for glucose concentration, osmolality, and triglyceride levels. Body weight was recorded at every time point. Values are the average ± SE obtained from eight to 11 mice/condition. *, P < 0.05.
To determine the effect of streptozotocin-induced diabetes on the mouse skeleton, μCT analyses were carried out on the tibia, which has regions of predominantly cortical bone (the middiaphysis) and regions of trabecular bone (the metaphysis). Because differences in bone length (as a result of altered development) could add an additional variable to our analyses, we first measured and compared the lengths of control and diabetic tibias. Table 1 demonstrates that tibial length does not significantly differ between control and diabetic (4 wk postinduction) animals, consistent with the animals being on the flat part of the growth curve at the time of diabetes induction. Examination of cortical bone parameters, bone mineral content, bone mineral density, and moment of inertia, demonstrated no significant difference between control and diabetic animals (Table 1). In contrast, all trabecular bone parameters, bone mineral content, bone mineral density, and bone volume, were significantly decreased in diabetic compared with control mice (Table 1). The μCT images in Fig. 2 depict the reduction in trabecular bone within the secondary spongiosa of diabetic tibias in lateral and transverse sections.
TABLE 1. Tibia measurements in control and diabetic animals
FIG. 2. Diabetes reduces trabecular bone volume. A, Representative μCT lateral slices (20 μm) through the proximal tibia from two control and diabetic mice are shown. B, Representative μCT transverse slices (20 μm) through the proximal tibia at a distance of 0.5 or 1 mm from the lowest region of the growth plate.
Consistent with our μCT analyses, histological analyses of diabetic (4 wk after injection) proximal tibia demonstrate a decrease in bone volume marked by decreased trabecular bone volume and thickness (Table 2). Osteoblast number and osteoid parameters (volume, surface, and thickness) did not show a significant difference between control and diabetic animals (Table 2). However, dynamic measurements indicated a trend toward decreased osteoblast function/bone formation, which was significant when label incorporation/surface area was determined (3.5% vs. 1.4% in control and diabetic animals, respectively; Table 2).
TABLE 2. Histological analyses of diabetic and control tibia
Next, we examined systemic and tibial parameters of osteoclast activity. Urinary DPD, a breakdown product of collagen released systemically during osteoclast resorption and readily measured in the urine, indicated that osteoclast activity is not increased in diabetic animals 14 d after induction (Fig. 3A). Serum levels of active TRAP5b (produced by active osteoclasts) were not significantly increased in diabetic animals at any time studied and were actually decreased on d 14 and 21. Histological examination of the proximal tibia also indicated that osteoclast activity was not increased; this was indicated by the lack of a difference in osteoclast number, erosion surface, or erosion depth between control and diabetic animals (4 wk after induction; Fig. 3B). Consistent with this finding, measurement of TRAP5 mRNA levels in control and diabetic tibia did not differ (Fig. 3B).
FIG. 3. Systemic and tibia-specific measures of osteoclast activity in diabetic bone. A, Diabetes decreases serum active TRAP5b levels and has no effect on urinary DPD levels. Urine collected over a 24-h period was obtained from diabetic and control mice in a metabolic cage on d 14. DPD was determined in control and diabetic urine samples and expressed relative to creatinine, to control for differences in urine volume. Serum was obtained from mice at 5, 14, 21, or 28 d after the induction of diabetes. Active TRAP5b levels were determined in control (C) and diabetic (D) mouse serum samples. B, Histomorphometry and TRAP5 mRNA analyses were preformed on tibias obtained from mice 28 d after the induction of diabetes. All values are the average ± SE obtained from four to nine mice per condition. *, P < 0.05.
To assess whether diabetes suppresses osteoblast differentiation, we extracted total RNA from control and diabetic tibias and measured markers of progressive osteoblast differentiation: runx2, alkaline phosphatase, and osteocalcin mRNA levels. Although we did not detect a change in runx2 or alkaline phosphatase, a significant decrease in osteocalcin mRNA levels was evident in diabetic compared with control tibiae (Fig. 4). This was consistent with measurements of serum osteocalcin, which were also decreased in diabetic mice (Fig. 4). This suggests that there is a decrease in mature osteocalcin-expressing osteoblasts in the bones of diabetic animals, which supports our dynamic measurements, indicating a trend toward decreased bone formation. Additional examination of gene expression levels in mouse tibias indicated an increase in PPAR2 mRNA levels, a marker of adipocytes, in diabetic compared with control tibias (Fig. 5). Analysis of two other adipocyte markers, resistin and aP2, also demonstrated increased expression in diabetic tibia (Fig. 5). To confirm that these effects do not result from direct effects of streptozotocin on mesenchymal and osteoblast cells, we chronically treated pluripotent C3H10T1/2 cells (induced toward the osteoblast lineage or uninduced) with 0.01–1 mg/ml streptozotocin. This treatment did not induce adipogenic genes or reduce osteocalcin expression (data not shown), indicating that streptozotocin does not directly influence gene expression in these cells.
FIG. 4. Osteocalcin mRNA and serum levels are decreased in diabetes, in contrast to runx-2 and alkaline phosphatase (Alk Phos) mRNA levels. Total RNA, extracted from tibia isolated from control and diabetic (2 wk) adult BALB/c mice, was used for real-time RT-PCR analysis with SYBR Green dye. Levels of amplified genes were expressed relative to cyclophilin (a housekeeping gene) and relative to control levels, which were set at 1. Serum from 2-wk diabetic and control mice was analyzed for osteocalcin levels. Values are the average ± SE obtained from eight to 11 mice/condition. *, P < 0.05.
FIG. 5. Adipocyte markers, PPAR2, aP2, and resistin are increased in diabetic bone. Total RNA was extracted from tibia isolated from control and diabetic (2 wk) adult BALB/c mice and used for real-time RT-PCR analysis with SYBR Green dye. Levels of PPAR2, aP2, and resistin are expressed relative to cyclophilin (a housekeeping gene). Values are the average ± SE obtained from eight to 11 mice/condition. *, P < 0.05.
In agreement with RNA analyses, histological sections of diabetic tibia demonstrated an increase in marrow adiposity compared with controls (Fig. 6). Quantitation of adipocytes demonstrated that diabetic tibias contain nearly 3-fold more adipocytes (>50 μm in diameter) per area of bone marrow than control tibias (14.4 ± 4.6 vs. 4.8 ± 1.7 adipocytes/mm area of bone marrow, respectively). Interestingly, adipose tissue obtained from control femoral fat pads exhibited lipid-dense adipocytes, in contrast to diabetic adipose tissue, which exhibited lipid-sparse adipocytes (Fig. 6). Although liver architecture appeared somewhat different in diabetic compared with control animals (Fig. 6), it did not exhibit adiposity, as determined by histological examination, Oil Red-O staining (data not shown), and decreased weight (1.31 ± 0.02 vs. 1.46 ± 0.02 for controls; P < 0.001). The reduction in peripheral adiposity was also demonstrated by μCT analysis of transverse sections of mouse legs and by isolating and weighing the fat pads, which were reduced by more than 3-fold in size/weight (Fig. 7).
FIG. 6. Adiposity is increased in diabetic tibia, in contrast to peripheral adipose tissue lipolysis. Tibia (bone) obtained from control and diabetic (4 wk) mice were fixed and processed for plastic embedding and sectioning. Representative Masson-Goldner Trichrome-stained sections demonstrate differences in the amount of mineralized bone (blue stain) and in the number of adipocytes (unstained large circular regions within the marrow) between control and diabetic animals. Representative fat pad and liver sections were obtained from control and diabetic mice. Sections were stained with hematoxylin and eosin and digitally photographed.
FIG. 7. Peripheral adipose tissue is decreased in diabetic mice. Transverse slices of mouse legs were obtained by μCT analysis. Representative images are shown, with medium gray (marked by black arrows) indicating regions of fat. In addition, dissected femoral fat depots were photographed and weighed. The later values were pooled from five to nine mice per condition and are expressed as the average ± SE. *, P < 0.01.
Discussion
Bone loss is a known negative consequence of IDDM. The mechanism accounting for this outcome is unknown, making it difficult to develop optimal therapies. Diabetes is associated with a variety of pathologies, including little or no insulin, decreased IGF-I, hyperglycemia, and increased blood osmolality. An animal model is needed that is amenable to genetic manipulation to allow identification of the mechanisms accounting for diabetes-associated bone loss through knockout and/or transgenic approaches. Therefore, we set out to characterize the bone phenotype in wild-type, BALB/c, streptozotocin-induced diabetic mice.
Our histological analyses demonstrated a significant decrease in mouse trabecular bone volume (by 32%) 4 wk after the induction of diabetes. This amount is similar to bone loss reported in diabetic rats, which ranges from 10–50% (69, 70). Our μCT analyses confirm diabetes-associated bone loss in regions of trabecular bone (density decreased by 24% and volume by 69%), but no statistically significant changes were observed in regions of cortical bone. This is consistent with trabecular bone being more metabolically active than cortical bone and perhaps with an extended time cortical bone loss would become apparent. Alternatively, our results could indicate that trabecular bone (intimately associated with the marrow environment and local cytokine and growth factor stimuli) is more responsive to the diabetic environment than cortical bone (which is thought to be controlled more by systemic factors) (71).
Reports on diabetic osteoclast activity have been variable, but most studies indicate no change or a decrease in activity based on histology and/or secretion of deoxypyridinoline, a breakdown product of bone collagen matrices (69). Herrero et al. (28) demonstrate decreased pyridinium cross-links after 12 wk of diabetes in rats. Similarly, Verhaeghe et al. (29) demonstrate a decrease in urinary DPD secretion in rats diabetic for 8 wk. Our studies also did not demonstrate an increase in diabetic urinary DPD levels; however this measurement can be problematic in diabetics, in who, urine volumes are significantly increased compared with controls, filtration rates may be altered as a result of nephropathy, and collagen breakdown may be occurring in other tissues as well. Therefore, as additional measures of osteoclast activity, we examined TRAP5 mRNA and serum levels. TRAP5 mRNA levels did not differ between control and diabetic tibias at d 14 and 28 (not shown). Active TRAP5b levels in mouse serum, which is a specific systemic measure of osteoclast activity and correlates with other markers of bone turnover (72, 73), did not increase 4 wk after the induction of diabetes (when bone loss was evident), nor were they elevated at time points before this (5, 14, and 21 d). In fact, on d 14 and 21, we observed a significant decrease in active serum TRAP5b levels. A reduction in osteoclast activity could contribute to decreased osteoblast maturation through reduced osteoclast production of osteoblast-enhancing factors (74, 75). Alternatively, the reduction in osteoclast activity could be secondary to reduced osteoblast maturation and its associated decrease in osteoclast signaling (75, 76). It should also be noted that although measurements of osteoclast activity on d 5, 14, 21, and 28 were unchanged or reduced, our data do not exclude the possibility that osteoclast activity is increased at earlier time points. In summary, these results support the hypothesis that osteoclast activity is not increased, and could possibly be decreased, under diabetic conditions at the time points studied.
Examination of osteoblast maturation revealed a decrease in osteocalcin expression, a marker of late-stage osteoblast differentiation. Both tibial osteocalcin mRNA and systemic serum osteocalcin levels were reduced by more than 40%. This reduction correlates with the decrease in mineral apposition rate and label incorporation in diabetic histological sections. Given the high metabolic rate of mice, in which a 4-wk period is roughly equivalent to 1 yr in human terms, it is possible that this suppression can alone account for the bone loss we observed. Our findings of suppressed osteocalcin expression are supported by nearly all studies measuring serum osteocalcin in diabetic rats and humans, which report, on the average, a 50% decrease in osteocalcin levels compared with controls (27, 28, 77). This decrease is independent of the level of exercise the animals receive (29). Diabetes-associated suppression of osteoblast maturation is also suggested in bone implant/healing studies in rats, which indicate that diabetic animals exhibit decreased calvarial defect healing (30) and decreased bone formation around titanium or hydroxyapatite bone implants (78, 79, 80). In marrow-ablated mice, both runx2 and osteocalcin mRNA levels were decreased in diabetic relative to control bones 4 and 6 d after ablation, a time when bone formation was dramatically induced in controls (81). In our studies we did not detect a difference in runx2 mRNA levels between control and diabetic tibias. Our mRNA analyses were performed at 2 wk to examine changes in gene expression that occur before major bone loss, so the animals may not have been diabetic long enough to detect a suppression in runx2 mRNA levels in whole bone. The suppression of osteoblast differentiation in diabetes is also consistent with our in vitro analyses examining the role of hyperglycemia in the regulation of osteoblast phenotype. Specifically, we demonstrated that acute hyperglycemic as well as hyperosmotic conditions can suppress late-stage markers of osteoblast differentiation in vitro, such as osteocalcin (82). In this study we show that diabetic mice do indeed exhibit elevated plasma osmolality, which could contribute to a modulation in gene expression.
The above data support the hypothesis that diabetics lose bone through the suppression of late-stage osteoblast differentiation, whereas progression through earlier stages appears unaffected (as indicated by unaltered runx2 and alkaline phosphatase mRNA levels). However, we also propose a second pathway that contributes to diabetic bone loss, which involves the regulation of marrow adiposity. Specifically, our studies demonstrate for the first time that the number of lipid-dense adipocytes and the expression of adipocyte markers (PPAR2, resistin, and aP2) are increased in diabetic type I bone. From our experiments we cannot distinguish whether lipid-sparse adipocytes that are always present in the marrow are accumulating lipid and becoming visible or whether mesenchymal pluripotent cells are becoming adipocytes. The later could occur at the expense of osteoblast lineage selection (eventually leading to decreased osteoblast number at more extended experimental time points) or in addition to selecting the osteoblast pathway (thereby not reducing the number of immature osteoblast cell number).
Selection of adipogenesis over osteoblastogenesis is a common theme that has been reported in other conditions of bone loss, including age-related osteoporosis and disuse (44, 45, 46, 48, 83). PPAR can play a role in this selection, based on studies demonstrating that PPAR insufficiency results in the enhancement of osteogenesis and the suppression of adipogenesis in mice (84) and on studies demonstrating that elevation of PPAR2 levels can promote adipogenesis in pluripotent mesenchymal cells in vitro (43, 85). Thus, PPAR2 levels have a dominant suppressive influence on osteogenesis. In our studies, at least two possibilities can explain the function of PPAR2 elevation: either it functions as an inducer of adipogenesis or it represents a marker of increased adipocyte number and/or adipogenesis. It is known that PPAR levels and adipogenesis can be regulated by a variety of factors, including bone morphogenic protein (86, 87, 88), estrogen (89, 90), PTHrP (91), and TGF-? (92). In addition, low serum IGF-I levels can contribute to reduced bone mineral density and osteoblast differentiation and increase the number of adipocytes in bone (93, 94). The role of IGF-I is of particular interest to our IDDM animal studies, because diabetic animals and humans exhibit reduced IGF-I levels (95, 96). It is interesting that reduced IGF-I and insulin may promote marrow adipogenesis in vivo, in contrast to cell culture studies in which the lack of these factors reduces the probability of adipogenesis to occur (97).
Although elevation of PPAR2 is clearly critical for allowing PPAR2 signaling to occur, it is also important to have PPAR2 ligands available to bind to this receptor and activate its function (98). Our studies demonstrate that triglycerides are elevated in diabetic mice, and it is known that fatty acids can activate PPAR2 (98, 99) and promote differentiation of osteoblast-like cells into adipocyte-like cells (100). Thus, we hypothesize that diabetes-associated hypertriglyceridemia can compound and additional direct bone loss through promoting PPAR activity and adiposity in bone.
In summary, we demonstrate that IDDM mice exhibit bone loss. Interestingly, although animals are losing weight, under conditions of low serum insulin and IGF-I, adipogenesis or lipogenesis is still active in bone. This suggests that pluripotent cells and adipocytes in bone are somehow different or receive different signals compared with other adipose-precursor cells and adipocytes in other areas of the body. We also demonstrate that IDDM conditions suppress osteoblast maturation/function without increasing osteoclast activity. We hypothesize that an increase in bone adiposity and a decrease in mature osteoblasts contribute to the bone loss in diabetes. Understanding the mechanisms promoting bone adiposity and the importance of elevated PPAR2 expression under IDDM conditions will contribute to the development of better treatments to prevent bone loss in IDDM.
Acknowledgments
We thank Regina Irwin for her technical expertise, Drs. Laurie McCauley and Joseph Bidwell for their helpful comments, and the Investigative Histology Laboratory, Department of Physiology, Division of Human Pathology, Michigan State University.
References
Seeman E 2003 Invited review: pathogenesis of osteoporosis. J Appl Physiol 95:2142–2151
Carmeliet G, Vico L, Bouillon R 2001 Space flight: a challenge for normal bone homeostasis. Crit Rev Eukaryotic Gene Expression 11:131–144
Harpavat M, Keljo DJ, Regueiro MD 2004 Metabolic bone disease in inflammatory bowel disease. J Clin Gastroenterol 38:218–224
Boling EP 2004 Secondary osteoporosis: underlying disease and the risk for glucocorticoid-induced osteoporosis. Clin Ther 26:1–14
Diamond TH, Higano CS, Smith MR, Guise TA, Singer FR 2004 Osteoporosis in men with prostate carcinoma receiving androgen-deprivation therapy: recommendations for diagnosis and therapies. Cancer 100:892–899
Hui SL, Epstein S, Johnston Jr CC 1985 A prospective study of bone mass in patients with type I diabetes. J Clin Endocrinol Metab 60:74–80
Levin ME, Boisseau VC, Avioli LV 1976 Effects of diabetes mellitus on bone mass in juvenile and adult-onset diabetes. N Engl J Med 294:241–245
Piepkorn B, Kann P, Forst T, Andreas J, Pfutzner A, Beyer J 1997 Bone mineral density and bone metabolism in diabetes mellitus. Horm Metab Res 29:584–591
Kemink SA, Hermus AR, Swinkels LM, Lutterman JA, Smals AG 2000 Osteopenia in insulin-dependent diabetes mellitus; prevalence and aspects of pathophysiology. J Endocrinol Invest 23:295–303
Gunczler P, Lanes R, Paz-Martinez V, Martins R, Esaa S, Colmenares V, Weisinger JR 1998 Decreased lumbar spine bone mass and low bone turnover in children and adolescents with insulin dependent diabetes mellitus followed longitudinally. J Pediatr Endocrinol Metab 11:413–419
Buysschaert M, Cauwe F, Jamart J, Brichant C, De Coster P, Magnan A, Donckier J 1992 Proximal femur density in type 1 and 2 diabetic patients. Diabetes Metab 18:32–37
Tuominen JT, Impivaara O, Puukka P, Ronnemaa T 1999 Bone mineral density in patients with type 1 and type 2 diabetes. Diabetes Care 22:1196–1200
Hampson G, Evans C, Petitt RJ, Evans WD, Woodhead SJ, Peters JR, Ralston SH 1998 Bone mineral density, collagen type 1 1 genotypes and bone turnover in premenopausal women with diabetes mellitus. Diabetologia 41:1314–1320
Wiske PS, Wentworth SM, Norton Jr JA, Epstein S, Johnston Jr CC 1982 Evaluation of bone mass and growth in young diabetics. Metabolism 31:848–854
McNair P, Christiansen C, Christensen MS, Madsbad S, Faber OK, Binder C, Transbol I 1981 Development of bone mineral loss in insulin-treated diabetes: a 1 1/2 years follow-up study in sixty patients. Eur J Clin Invest 11:55–59
Auwerx J, Dequeker J, Bouillon R, Geusens P, Nijs J 1988 Mineral metabolism and bone mass at peripheral and axial skeleton in diabetes mellitus. Diabetes 37:8–12
Krakauer JC, McKenna MJ, Buderer NF, Rao DS, Whitehouse FW, Parfitt AM 1995 Bone loss and bone turnover in diabetes. Diabetes 44:775–782
Munoz-Torres M, Jodar E, Escobar-Jimenez F, Lopez-Ibarra PJ, Luna JD 1996 Bone mineral density measured by dual x-ray absorptiometry in Spanish patients with insulin-dependent diabetes mellitus. Calcif Tissue Int 58:316–319
Bouillon R 1991 Diabetic bone disease. Calcif Tissue Int 49:155–160
Meyer HE, Tverdal A, Falch JA 1993 Risk factors for hip fracture in middle-aged Norwegian women and men. Am J Epidemiol 137:1203–1211
Schwartz AV, Sellmeyer DE, Ensrud KE, Cauley JA, Tabor HK, Schreiner PJ, Jamal SA, Black DM, Cummings SR 2001 Older women with diabetes have an increased risk of fracture: a prospective study. J Clin Endocrinol Metab 86:32–38
Forsen L, Meyer HE, Midthjell K, Edna TH 1999 Diabetes mellitus and the incidence of hip fracture: results from the Nord-Trondelag Health Survey. Diabetologia 42:920–925
White CB, Turner NS, Lee GC, Haidukewych GJ 2003 Open ankle fractures in patients with diabetes mellitus. Clin Orthop 414:37–44
Folk JW, Starr AJ, Early JS 1999 Early wound complications of operative treatment of calcaneus fractures: analysis of 190 fractures. J Orthop Trauma 13:369–372
Herskind AM, Christensen K, Norgaard-Andersen K, Andersen JF 1992 Diabetes mellitus and healing of closed fractures. Diabetes Metab 18:63–64
Bouillon R, Bex M, Van Herck E, Laureys J, Dooms L, Lesaffre E, Ravussin E 1995 Influence of age, sex, and insulin on osteoblast function: osteoblast dysfunction in diabetes mellitus. J Clin Endocrinol Metab 80:1194–1202
Verhaeghe J, van Herck E, Visser WJ, Suiker AM, Thomasset M, Einhorn TA, Faierman E, Bouillon R 1990 Bone and mineral metabolism in BB rats with long-term diabetes. Decreased bone turnover and osteoporosis. Diabetes 39:477–482
Herrero S, Calvo OM, Garcia-Moreno C, Martin E, San Roman JI, Martin M, Garcia-Talavera JR, Calvo JJ, del Pino-Montes J 1998 Low bone density with normal bone turnover in ovariectomized and streptozotocin-induced diabetic rats. Calcif Tissue Int 62:260–265
Verhaeghe J, Thomsen JS, van Bree R, van Herck E, Bouillon R, Mosekilde L 2000 Effects of exercise and disuse on bone remodeling, bone mass, and biomechanical competence in spontaneously diabetic female rats. Bone 27:249–256
Shyng YC, Devlin H, Sloan P 2001 The effect of streptozotocin-induced experimental diabetes mellitus on calvarial defect healing and bone turnover in the rat. Int J Oral Maxillofac Surg 30:70–74
Sasaki T, Kaneko H, Ramamurthy NS, Golub LM 1991 Tetracycline administration restores osteoblast structure and function during experimental diabetes. Anat Rec 231:25–34
McCracken M, Lemons JE, Rahemtulla F, Prince CW, Feldman D 2000 Bone response to titanium alloy implants placed in diabetic rats. Int J Oral Maxillofac Implants 15:345–354
Ferrari G, Cusella-De Angelis G, Coletta M, Paolucci E, Stornaiuolo A, Cossu G, Mavilio F 1998 Muscle regeneration by bone marrow-derived myogenic progenitors. Science 279:1528–1530
Krause DS, Theise ND, Collector MI, Henegariu O, Hwang S, Gardner R, Neutzel S, Sharkis SJ 2001 Multi-organ, multi-lineage engraftment by a single bone marrow-derived stem cell. Cell 105:369–377
Petersen BE, Bowen WC, Patrene KD, Mars WM, Sullivan AK, Murase N, Boggs SS, Greenberger JS, Goff JP 1999 Bone marrow as a potential source of hepatic oval cells. Science 284:1168–1170
Sanchez-Ramos J, Song S, Cardozo-Pelaez F, Hazzi C, Stedeford T, Willing A, Freeman TB, Saporta S, Janssen W, Patel N, Cooper DR, Sanberg PR 2000 Adult bone marrow stromal cells differentiate into neural cells in vitro. Exp Neurol 164:247–256
Karsenty G 2001 Minireview: transcriptional control of osteoblast differentiation. Endocrinology 142:2731–2733
Stein GS, Lian JB, van Wijnen AJ, Stein JL, Montecino M, Javed A, Zaidi SK, Young DW, Choi JY, Pockwinse SM 2004 Runx2 control of organization, assembly and activity of the regulatory machinery for skeletal gene expression. Oncogene 23:4315–4329
Lian JB, Stein GS, Stein JL, van Wijnen AJ 1998 Osteocalcin gene promoter: unlocking the secrets for regulation of osteoblast growth and differentiation. J Cell Biochem 30–31(Suppl):62–72
Winchester SK, Selvamurugan N, D’Alonzo RC, Partridge NC 2000 Developmental regulation of collagenase-3 mRNA in normal, differentiating osteoblasts through the activator protein-1 and the runt domain binding sites. J Biol Chem 275:23310–23318
Sasano Y, Zhu JX, Tsubota M, Takahashi I, Onodera K, Mizoguchi I, Kagayama M 2002 Gene expression of MMP8 and MMP13 during embryonic development of bone and cartilage in the rat mandible and hind limb. J Histochem Cytochem 50:325–332
Rosen ED, Spiegelman BM 2001 PPAR: a nuclear regulator of metabolism, differentiation, and cell growth. J Biol Chem 276:37731–37734[Free Full Text]
Lecka-Czernik B, Gubrij I, Moerman EJ, Kajkenova O, Lipschitz DA, Manolagas SC, Jilka RL 1999 Inhibition of Osf2/Cbfa1 expression and terminal osteoblast differentiation by PPAR2. J Cell Biochem 74:357–371
Jilka RL, Weinstein RS, Takahashi K, Parfitt AM, Manolagas SC 1996 Linkage of decreased bone mass with impaired osteoblastogenesis in a murine model of accelerated senescence. J Clin Invest 97:1732–1740
Kajkenova O, Lecka-Czernik B, Gubrij I, Hauser SP, Takahashi K, Parfitt AM, Jilka RL, Manolagas SC, Lipschitz DA 1997 Increased adipogenesis and myelopoiesis in the bone marrow of SAMP6, a murine model of defective osteoblastogenesis and low turnover osteopenia. J Bone Miner Res 12:1772–1779
Verma S, Rajaratnam JH, Denton J, Hoyland JA, Byers RJ 2002 Adipocytic proportion of bone marrow is inversely related to bone formation in osteoporosis. J Clin Pathol 55:693–698
Moerman EJ, Teng K, Lipschitz DA, Lecka-Czernik B 2004 Aging activates adipogenic and suppresses osteogenic programs in mesenchymal marrow stroma/stem cells: the role of PPAR-2 transcription factor and TGF-?/BMP signaling pathways. Aging Cell 3:379–389
Nuttall ME, Gimble JM 2004 Controlling the balance between osteoblastogenesis and adipogenesis and the consequent therapeutic implications. Curr Opin Pharmacol 4:290–294
Pechhold K, Patterson NB, Blum C, Fleischacker CL, Boehm BO, Harlan DM 2001 Low dose streptozotocin-induced diabetes in rat insulin promoter-mCD80-transgenic mice is T cell autoantigen-specific and CD28 dependent. J Immunol 166:2531–2539
Szkudelski T 2001 The mechanism of alloxan and streptozotocin action in B cells of the rat pancreas. Physiol Res 50:537–546
McCabe LR, Kockx M, Lian J, Stein J, Stein G 1995 Selective expression of fos- and jun-related genes during osteoblast proliferation and differentiation. Exp Cell Res 218:255–262
Chomczynski P, Sacchi N 1987 Single-step method of RNA isolation by acid guanidinium thiocyanate-phenol-chloroform extraction. Anal Biochem 162:156–159
Ontiveros C, McCabe LR 2003 Simulated microgravity suppresses osteoblast phenotype, Runx2 levels and AP-1 transactivation. J Cell Biochem 88:427–437
Ontiveros C, Irwin R, Wiseman RW, McCabe LR 2004 Hypoxia suppresses runx2 independent of modeled microgravity. J Cell Physiol 200:169–176
Kast-Woelbern HR, Dana SL, Cesario RM, Sun L, de Grandpre LY, Brooks ME, Osburn DL, Reifel-Miller A, Klausing K, Leibowitz MD 2004 Rosiglitazone induction of Insig-1 in white adipose tissue reveals a novel interplay of peroxisome proliferator-activated receptor and sterol regulatory element-binding protein in the regulation of adipogenesis. J Biol Chem 279:23908–23915
Li J, Takaishi K, Cook W, McCorkle SK, Unger RH 2003 Insig-1 "brakes" lipogenesis in adipocytes and inhibits differentiation of preadipocytes. Proc Natl Acad Sci USA 100:9476–9481
Wiren KM, Zhang XW, Toombs AR, Kasparcova V, Gentile MA, Harada S, Jepsen KJ 2004 Targeted overexpression of androgen receptor in osteoblasts: unexpected complex bone phenotype in growing animals. Endocrinology 145:3507–3522
Harada K, Shen WJ, Patel S, Natu V, Wang J, Osuga J, Ishibashi S, Kraemer FB 2003 Resistance to high-fat diet-induced obesity and altered expression of adipose-specific genes in HSL-deficient mice. Am J Physiol 285:E1182–E1195
Lewis ML, Hughes-Fulford M 2000 Regulation of heat shock protein message in Jurkat cells cultured under serum-starved and gravity-altered conditions. J Cell Biochem 77:127–134
Trogan E, Choudhury RP, Dansky HM, Rong JX, Breslow JL, Fisher EA 2002 Laser capture microdissection analysis of gene expression in macrophages from atherosclerotic lesions of apolipoprotein E-deficient mice. Proc Natl Acad Sci USA 99:2234–2239
Date T, Doiguchi Y, Nobuta M, Shindo H 2004 Bone morphogenetic protein-2 induces differentiation of multipotent C3H10T1/2 cells into osteoblasts, chondrocytes, and adipocytes in vivo and in vitro. J Orthop Sci 9:503–508
Malluche HH, Faugere MC 1986 Atlas of mineralized bone histology. New York: Karger
Goldner J 1938 A modification of the Masson Trichrome technique for routine laboratory purposes. Am J Pathol 14:237–243
Manaka RC, Malluche HH 1981 A program package for quantitative analysis of histologic structure and remodeling dynamics of bone. Comput Programs Biomed 13:191–202
Malluche HH, Monier-Faugere MC 1994 The role of bone biopsy in the management of patients with renal osteodystrophy. J Am Soc Nephrol 4:1631–1642
Parfitt AM, Drezner MK, Glorieux FH, Kanis JA, Malluche HH, Meunier PJ, Ott SM, Recker RR 1987 Bone histomorphometry: standardization of nomenclature, symbols and units. J Bone Miner Res 6:595–610
Miljanovic B, Glynn RJ, Nathan DM, Manson JE, Schaumberg DA 2004 A prospective study of serum lipids and risk of diabetic macular edema in type 1 diabetes. Diabetes 53:2883–2892
Greenwalt DE, Scheck SH, Rhinehart-Jones T 1995 Heart CD36 expression is increased in murine models of diabetes and in mice fed a high fat diet. J Clin Invest 96:1382–1388
Locatto ME, Abranzon H, Caferra D, Fernandez MC, Alloatti R, Puche RC 1993 Growth and development of bone mass in untreated alloxan diabetic rats. Effects of collagen glycosylation and parathyroid activity on bone turnover. Bone Miner 23:129–144
Waud CE, Marks Jr SC, Lew R, Baran DT 1994 Bone mineral density in the femur and lumbar vertebrae decreases after twelve weeks of diabetes in spontaneously diabetic-prone BB/Worcester rats. Calcif Tissue Int 54:237–240
Mundy GR, Chen D, Oyajobi BO 2003 Bone Remodeling. In: Favus M, ed. Primer on the metabolic bone diseases and disorders of mineral metabolism, 5th Ed. Washington DC: American Society of Bone and Mineral Research; 46–58
Halleen JM, Alatalo SL, Suominen H, Cheng S, Janckila AJ, Vaananen HK 2000 Tartrate-resistant acid phosphatase 5b: a novel serum marker of bone resorption. J Bone Miner Res 15:1337–1345
Halleen JM, Ylipahkala H, Alatalo SL, Janckila AJ, Heikkinen JE, Suominen H, Cheng S, Vaananen HK 2002 Serum tartrate-resistant acid phosphatase 5b, but not 5a, correlates with other markers of bone turnover and bone mineral density. Calcif Tissue Int 71:20–25
Dai XM, Zong XH, Akhter MP, Stanley ER 2004 Osteoclast deficiency results in disorganized matrix, reduced mineralization, and abnormal osteoblast behavior in developing bone. J Bone Miner Res 19:1441–1451
Phan TC, Xu J, Zheng MH 2004 Interaction between osteoblast and osteoclast: impact in bone disease. Histol Histopathol 19:1325–1344
Kwan Tat S, Padrines M, Theoleyre S, Heymann D, Fortun Y 2004 IL-6, RANKL, TNF-/IL-1: interrelations in bone resorption pathophysiology. Cytokine Growth Factor Rev 15:49–60
Epstein S, Takizawa M, Stein B, Katz IA, Joffe, II, Romero DF, Liang XG, Li M, Ke HZ, Jee WS 1994 Effect of cyclosporin A on bone mineral metabolism in experimental diabetes mellitus in the rat. J Bone Miner Res 9:557–566
Siqueira JT, Cavalher-Machado SC, Arana-Chavez VE, Sannomiya P 2003 Bone formation around titanium implants in the rat tibia: role of insulin. Implant Dent 12:242–251
Giglio MJ, Giannunzio G, Olmedo D, Guglielmotti MB 2000 Histomorphometric study of bone healing around laminar implants in experimental diabetes. Implant Dent 9:143–149
Takeshita F, Iyama S, Ayukawa Y, Kido MA, Murai K, Suetsugu T 1997 The effects of diabetes on the interface between hydroxyapatite implants and bone in rat tibia. J Periodontol 68:180–185
Lu H, Kraut D, Gerstenfeld LC, Graves DT 2003 Diabetes interferes with the bone formation by affecting the expression of transcription factors that regulate osteoblast differentiation. Endocrinology 144:346–352
Zayzafoon M, Stell C, Irwin R, McCabe LR 2000 Extracellular glucose influences osteoblast differentiation and c-Jun expression. J Cell Biochem 79:301–310
Meunier P, Aaron J, Edouard C, Vignon G 1971 Osteoporosis and the replacement of cell populations of the marrow by adipose tissue. A quantitative study of 84 iliac bone biopsies. Clin Orthop 80:147–154
Akune T, Ohba S, Kamekura S, Yamaguchi M, Chung UI, Kubota N, Terauchi Y, Harada Y, Azuma Y, Nakamura K, Kadowaki T, Kawaguchi H 2004 PPAR insufficiency enhances osteogenesis through osteoblast formation from bone marrow progenitors. J Clin Invest 113:846–855
Rzonca SO, Suva LJ, Gaddy D, Montague DC, Lecka-Czernik B 2004 Bone is a target for the antidiabetic compound rosiglitazone. Endocrinology 145:401–406
Skillington J, Choy L, Derynck R 2002 Bone morphogenetic protein and retinoic acid signaling cooperate to induce osteoblast differentiation of preadipocytes. J Cell Biol 159:135–146
Fux C, Mitta B, Kramer BP, Fussenegger M 2004 Dual-regulated expression of C/EBP- and BMP-2 enables differential differentiation of C2C12 cells into adipocytes and osteoblasts. Nucleic Acids Res 32:e1
Chen D, Ji X, Harris MA, Feng JQ, Karsenty G, Celeste AJ, Rosen V, Mundy GR, Harris SE 1998 Differential roles for bone morphogenetic protein (BMP) receptor type IB and IA in differentiation and specification of mesenchymal precursor cells to osteoblast and adipocyte lineages. J Cell Biol 142:295–305
Okazaki R, Inoue D, Shibata M, Saika M, Kido S, Ooka H, Tomiyama H, Sakamoto Y, Matsumoto T 2002 Estrogen promotes early osteoblast differentiation and inhibits adipocyte differentiation in mouse bone marrow stromal cell lines that express estrogen receptor (ER) or ?. Endocrinology 143:2349–2356
Dang Z, Lowik CW 2004 The balance between concurrent activation of ERs and PPARs determines daidzein-induced osteogenesis and adipogenesis. J Bone Miner Res 19:853–861
Chan GK, Miao D, Deckelbaum R, Bolivar I, Karaplis A, Goltzman D 2003 Parathyroid hormone-related peptide interacts with bone morphogenetic protein 2 to increase osteoblastogenesis and decrease adipogenesis in pluripotent C3H10T 1/2 mesenchymal cells. Endocrinology 144:5511–5520
Ahdjoudj S, Lasmoles F, Holy X, Zerath E, Marie PJ 2002 Transforming growth factor ?2 inhibits adipocyte differentiation induced by skeletal unloading in rat bone marrow stroma. J Bone Miner Res 17:668–677
Rosen CJ, Ackert-Bicknell CL, Adamo ML, Shultz KL, Rubin J, Donahue LR, Horton LG, Delahunty KM, Beamer WG, Sipos J, Clemmons D, Nelson T, Bouxsein ML, Horowitz M 2004 Congenic mice with low serum IGF-I have increased body fat, reduced bone mineral density, and an altered osteoblast differentiation program. Bone 35:1046–1058
Zhang M, Xuan S, Bouxsein ML, von Stechow D, Akeno N, Faugere MC, Malluche H, Zhao G, Rosen CJ, Efstratiadis A, Clemens TL 2002 Osteoblast-specific knockout of the insulin-like growth factor (IGF) receptor gene reveals an essential role of IGF signaling in bone matrix mineralization. J Biol Chem 277:44005–44012
Verhaeghe J, Oloumi G, van Herck E, van Bree R, Dequeker J, Einhorn TA, Bouillon R 1997 Effects of long-term diabetes and/or high-dose 17 ?-estradiol on bone formation, bone mineral density, and strength in ovariectomized rats. Bone 20:421–428
Jehle PM, Jehle DR, Mohan S, Bohm BO 1998 Serum levels of insulin-like growth factor system components and relationship to bone metabolism in type 1 and type 2 diabetes mellitus patients. J Endocrinol 159:297–306
MacDougald OA, Lane MD 1995 Transcriptional regulation of gene expression during adipocyte differentiation. Annu Rev Biochem 64:345–373
Lecka-Czernik B, Moerman EJ, Grant DF, Lehmann JM, Manolagas SC, Jilka RL 2002 Divergent effects of selective peroxisome proliferator-activated receptor-2 ligands on adipocyte versus osteoblast differentiation. Endocrinology 143:2376–2384
Forman BM, Tontonoz P, Chen J, Brun RP, Spiegelman BM, Evans RM 1995 15-Deoxy-12,14-prostaglandin J2 is a ligand for the adipocyte determination factor PPAR. Cell 83:803–812
Diascro Jr DD, Vogel RL, Johnson TE, Witherup KM, Pitzenberger SM, Rutledge SJ, Prescott DJ, Rodan GA, Schmidt A 1998 High fatty acid content in rabbit serum is responsible for the differentiation of osteoblasts into adipocyte-like cells. J Bone Miner Res 13:96–106(Sergiu Botolin, Marie-Cla)
Address all correspondence and requests for reprints to: Dr. Laura R. McCabe, Department of Physiology, Michigan State University, 2201 Biomedical Physical Science Building, East Lansing, Michigan 48824. E-mail: mccabel@msu.edu.
Abstract
Decreased bone mass, osteoporosis, and increased fracture rates are common skeletal complications in patients with insulin-dependent diabetes mellitus (IDDM; type I diabetes). IDDM develops from little or no insulin production and is marked by elevated blood glucose levels and weight loss. In this study we use a streptozotocin-induced diabetic mouse model to examine the effect of type I diabetes on bone. Histology and microcomputed tomography demonstrate that adult diabetic mice, exhibiting increased plasma glucose and osmolality, have decreased trabecular bone mineral content compared with controls. Bone resorption could not completely account for this effect, because resorption markers (tartrate-resistant acid phosphatase 5b, urinary deoxypyridinoline excretion, and tartrate-resistant acid phosphatase 5 mRNA) are unchanged or reduced at 2 and/or 4 wk after diabetes induction. However, osteocalcin mRNA (a marker of late-stage osteoblast differentiation) and dynamic parameters of bone formation were decreased in diabetic tibias, whereas osteoblast number and runx2 and alkaline phosphatase mRNA levels did not differ. These findings suggest that the final stages of osteoblast maturation and function are suppressed. We also propose a second mechanism contributing to diabetic bone loss: increased marrow adiposity. This is supported by increased expression of adipocyte markers [peroxisome proliferator-activated receptor 2, resistin, and adipocyte fatty acid binding protein (P2)] and the appearance of lipid-dense adipocytes in diabetic tibias. In contrast to bone marrow, adipose stores at other sites are depleted in diabetic mice, as indicated by decreased body, liver, and peripheral adipose tissue weights. These findings suggest that IDDM contributes to bone loss through changes in marrow composition resulting in decreased mature osteoblasts and increased adipose accumulation.
Introduction
BONE IS A highly specialized and dynamic tissue that undergoes continual remodeling throughout its lifetime. Remodeling processes in bone are a sum of two activities: bone resorption and bone formation. Osteoblasts are responsible for bone formation, whereas osteoclasts resorb bone. An equilibrium between osteoblast and osteoclast activities maintains bone mass and quality. In contrast, alteration of one part of the equilibrium can lead to bone loss (decreased formation and/or increased resorption) or increased bone formation (decreased bone resorption and/or increased bone formation) (1, 2). Changes in this balance, leading to bone loss and osteoporosis, can occur under conditions such as aging, skeletal unloading/disuse, and disease (1, 2, 3, 4, 5).
Insulin-dependent diabetes mellitus (IDDM type I) is a chronic disease characterized by a lack of insulin production. In the absence of insulin, insulin-sensitive cells exhibit markedly reduced glucose uptake, resulting in increased serum glucose levels. Two of the long-term complications of IDDM are diabetic osteopenia and osteoporosis (6, 7, 8, 9, 10, 11, 12, 13, 14, 15, 16, 17, 18), which are associated with increased fracture rate (19, 20, 21, 22) and delayed fracture healing (23, 24, 25). Recent clinical studies found that 67% of men and 57% of women with IDDM suffer from osteopenia of the femoral neck and/or lumbar spine (9), and 14–20% of IDDM patients aged 20–56 yr met the criteria for more extensive bone loss, osteoporosis (9, 18). In diabetic patients, osteoblast function is significantly decreased and is thought to be associated with a maturation defect. This conclusion is based on the fact that serum levels of an early osteoblast marker, peptide of procollagen, remain normal in all types of diabetes, whereas serum levels of a late-stage marker of osteoblast maturation, osteocalcin, are decreased in patients at a range of ages (26). These pathological findings are even more pronounced in diabetic rats (27, 28, 29, 30), which also exhibit decreased bone volume and osteoid surfaces with no reduction in osteoclast number (27, 31) and significantly less osseointegration of bone implants than controls (32). These findings suggest that a decrease in osteoblast number and/or function/maturation could be a major contributor to bone loss in IDDM.
Osteoblasts are derived from mesenchymal stem cells. These pluripotent stem cells can give rise to adipocytes as well as a variety of other cell types (33, 34, 35, 36). During the process of osteogenesis and osteoblast differentiation, the expression of runx2, a transcription factor required for these processes (37, 38), is increased. Late-stage differentiation is marked by elevated osteocalcin expression (39, 40, 41). In contrast, overexpression of peroxisome proliferator-activated receptor 2 (PPAR2), a member of the nuclear receptor transcription factor family (42), induces adipogenesis over osteoblastogenesis in pluripotent cells (43). Interestingly, if PPAR2 is expressed in osteoblasts, it can suppress the mature osteoblast phenotype and induce genes associated with an adipocyte-like phenotype, such as aP2 (also called fatty acid-binding protein 4), fatty acid synthase, and lipoprotein lipase (43). Selection of adipogenesis over osteoblastogenesis is thought to contribute to bone loss associated with a variety of conditions, including age-related and disuse-associated osteoporosis (44, 45, 46, 47, 48).
Although studies have examined the influence of IDDM on human and rat bone, few have directly addressed the influence of diabetes on the skeletal system of mice, an animal model that is amendable to genetic manipulation. In this study we use a streptozotocin-induced IDDM mouse model to examine the influence of this disease on bone. Consistent with human and rat studies, we found a decrease in diabetic bone volume [histologically and by microcomputed tomography (μCT) analyses]. In addition, we found a decrease in osteocalcin mRNA levels, and we are the first to report an increase in the expression of adipocyte markers, PPAR2, resistin, and aP2, and an increase in visible adipocytes in IDDM compared with control tibias. In contrast to bone marrow, peripheral adipose tissue was decreased in size, suggesting that the marrow environment and adipose stores are regulated differently from peripheral sites. These findings support our hypothesis that IDDM causes a shift in skeletal composition marked by increased adipose storage and decreased mature osteoblasts.
Materials and Methods
Diabetic mouse model
Diabetes was induced in adult (15 wk old) male BALB/c mice (Harlem Laboratories, Houston, TX) by daily ip injection of streptozotocin (40 μg/g body weight in 0.1 citrate buffer), a pancreatic ?-cell cytotoxin, for 5 d (49, 50). Controls were injected with buffer alone. Seven days after the last injection, nonfasting blood glucose measurements were made using blood obtained from the lateral saphenous vein and a glucometer (Accu-Check instant, Roche, Indianapolis, IN). This day was considered d 0. Mice with blood glucose levels greater than 300 mg/dl were considered diabetic. Animals were maintained in metabolic cages, kept on a 12-h light, 12-h dark cycle at 23 C, and received food and water ad libitum. After 2 or 4 wk of confirmed diabetic condition, mice (controls and diabetics) were fasted for 6 h, then killed. Tibiae were immediately removed, freed from soft tissue, and either fixed in formalin (for histology and μCT analyses) or snap-frozen in liquid nitrogen and stored at –80 C (for RNA analyses). At 4 wk, the liver and sc femoral fat pads were dissected from the surrounding tissues. Tissues were weighed and snap-frozen in liquid nitrogen or fixed in PBS-buffered formalin. Animal studies were conducted in accordance with the Michigan State University all-university committee on animal use and care.
Plasma measurements
Blood was obtained from 2- and 4-wk-old mice when the animals were killed, and blood serum was prepared from each sample by centrifugation for 5 min at 3000 rpm. Serum was stored frozen at –20 C. The glucose concentration in serum samples was determined using a glucose assay kit (Sigma-Aldrich Corp., St. Louis, MO). Serum osmolality, defined as the expression of the total number of solute particles dissolved in 1 kg solvent (International Federation of Clinical Chemistry), was determined using a vapor pressure osmometer (Wescor, Inc., Logan, UT). Serum osteocalcin levels were measured using a mouse osteocalcin enzyme immunoassay kit (Biomedical Technologies Inc., Stoughton, MA) according to the manufacturer’s instructions. Quantitative determinations of serum glycerol, total triglycerides, and true triglyceride (expressed as equivalent triolein concentration) levels were performed using a serum triglyceride determination kit (Sigma-Aldrich Corp.).
Osteoclast activity-systemic measurements
For measurement of tartrate-resistant acid phosphatase (TRAP), serum from 5 d and 2, 3, and 4 wk postinduction diabetic and control mice was used. Active serum TRAP5b, produced by mouse osteoclasts, was measured according to the manufacturer’s protocol using a solid phase, immunofixed, enzyme activity assay, MouseTRAP (SBA Sciences, Turku, Finland).
For urinary deoxypyridinoline (DPD) measurements, mice were housed in metabolic cages to allow collection of 24-h urine samples (12, 13, and 14 d after confirmation of diabetes induction). Urine specimens were stored at –80 C. Urinary DPD and creatinine levels were determined using a Metra DPD enzyme immunoassay kit and Metra creatinine assay kit, respectively (Quidel Corp., San Diego, CA). DPD levels were expressed relative to creatinine levels.
RNA analysis
Tibiae were crushed under liquid nitrogen conditions using a Bessman tissue pulverizer (Spectrum Laboratories, Inc., Rancho Dominguez, CA), and RNA was extracted using the method of Chomczynski and Sacchi, as previously described (51, 52). RNA integrity was verified by formaldehyde-agarose gel electrophoresis. Synthesis of cDNA was performed by RT with 2 μg total RNA using the SuperScript II kit with oligo(deoxythymidine)12–18 primers as described by the manufacturer (Invitrogen Life Technologies, Inc., Carlsbad, CA). cDNA (1 μl) was amplified by PCR in a final volume of 25 μl using the iQ SYBR Green Supermix (Bio-Rad Laboratories, Hercules, CA) with 10 pmol of each primer (Integrated DNA Technologies, Coralville, IA). Osteocalcin was amplified using 5'-ACG GTA TCA CTA TTT AGG ACC TGT G-3' and 5'-ACT TTA TTT TGG AGC TGC TGT GAC-3' (53). Runx2 was amplified using 5'-GAC AGA AGC TTG ATG ACT CTA AAC C-3' and 5'-TCT GTA ATC TGA CTC TGT CCT TGT G-3' (54). Alkaline phosphatase was amplified using 5'-CGTAATCTACCATGGAGACATTTTC-3' and 5'-GACTGTGGTTACTGCTGATCATTC-3'. PPAR2 was amplified using 5'-TGA AAC TCT GGG AGA TTC TCC TG-3' and CCA TGG TAA TTT CTT GTG AAG TGC-3' (55). Adipocyte fatty acid-binding protein 2 (aP2) was amplified using 5'-GCG TGG AAT TCG ATG AAA TCA-3' and 5'-CCC GCC ATC TAG GGT TAT GA-3' (56). TRAP5 was amplified using 5'-AATGCCTCGACCTGGGA-3' and 5'-CGTAGTCCTCCTTGGCTGCT-3' (57). Resistin was amplified using 5'-GCTGCTGCCAAGGCTGAT-3' and 5'-TCTCCTTCCACCATGTAGTTTCC-3' (58). Cyclophilin, which was not modulated under diabetic conditions, was used as a control for RNA levels; it was amplified using 5'-ATT CAT GTG CCA GGG TGG TGA C-3' and 5'-CCG TTT GTG TTT GGT CCA GCA-3' (59, 60). Real-time PCR was carried out for 40 cycles using the iCycler (Bio-Rad Laboratories), and data were evaluated using iCycler software. Each cycle consisted of 95 C for 15 sec, 60 C for 30 sec (except for runx2 and osteocalcin, which had an annealing temperature of 65 C), and 72 C for 30 sec. RNA-free samples, a negative control, did not produce amplicons. Melting curve and gel analyses were used to verify single products of the appropriate size (in base pairs).
In vitro examination of streptozotocin on pluripotent C3H10T1/2 cells
C3H10T1/2 cells (61) were obtained from American Type Culture Collection (Manassas, VA) and maintained in Eagle’s Basal Medium (Sigma-Aldrich Corp.) with 10% fetal bovine serum (Atlanta Biologicals, Norcross, GA) at 37 C in a humidified atmosphere of 5% CO2/95% air. Cells were plated at a density of 100,000 cells per well of a six-well plate. To induce osteoblast differentiation, bone morphogenic protein 2 (Sigma-Aldrich Corp.) was added at a concentration of 100 ng/ml 24 h after plating (61). Cells were fed every other day with or without an additional 100 ng/ml bone morphogenic protein 2 and were grown for 14 d. Streptozotocin concentrations were 0.01, 0.1, 0.5, and 1.0 mg/ml and was added with every feeding. Cells were fed and treated 24 h before harvesting for RNA analyses or lipid staining.
Tissue and mineralized bone histology and histomorphometry
Proximal tibiae isolated from 4-wk-old control (n = 8) and diabetic (n = 11) mice were fixed in absolute ethanol at room temperature, dehydrated, and embedded in methylmethacrylate as previously described (62). For dynamic measurements, mice were injected ip with 0.2 ml 10 mg/ml calcein (Sigma-Aldrich Corp.) in sterile saline 6 and 2 d before the animals were killed. Sections of 3- and 7-μm thickness were cut with a Microm microtome (model HM360, Carl Zeiss, Inc., Thornwood, NY) and stained with the modified Masson-Goldner Trichrome stain (63). Static and dynamic parameters of bone structure, formation, and bone resorption were measured at standardized sites under the growth plate using the semiautomatic method (Osteoplan II, Kontron, Munich, Germany) at a magnification of x200 (64, 65). For adipose analyses, visible adipocytes (>50 μm) were counted. All parameters comply with the guidelines of the nomenclature committee of the American Society of Bone and Mineral Research (66).
Subcutaneous femoral fat depots were isolated from surrounding tissue and fixed in 10% neutral buffered formalin. Fixed samples were processed on an automated Thermo Electron Excelsior tissue processor for dehydration, clearing, and infiltration using a routine overnight processing schedule. Samples were then embedded in Surgipath embedding paraffin on a Sakura Tissue Tek II embedding center. Paraffin blocks were sectioned at 5 μm on a Reichert Jung 2030 rotary microtome (Leica Instruments GmbH, Nussloch, Germany). Slides were stained with hematoxylin and eosin.
Livers were dissected out, sectioned, and placed in Tissue Tek OCT freezing medium (Miles, Inc., Elkhart, IN) on a sectioned cork; corks were snap-frozen in liquid nitrogen. Frozen tissue corks were stored at –80 C. Tissues were then sectioned on a –20 C Sakura Tissue Tek cryostat at 10 μm. Sections were placed on adhesive slides, air-dried for 30 min, fixed in 37–40% formaldehyde for 1 min, rinsed in running tap water for 5 min, and stained with hematoxylin and eosin.
μCT analysis
Fixed bones and mouse legs were scanned using a GE Explore Locus μCT system (General Electric, Fairfield, CT) at a voxel resolution of 20 μm obtained from 720 views. The beam angle of increment was 0.5, and the beam strength was set at 80 kvp and 450 μA. Each run included control and diabetic bones and a calibration phantom to standardize grayscale values and maintain consistency. Based on autothreshold and isosurface analyses of multiple bone samples, a fixed threshold was used to separate bone from bone marrow. All cortical bone analyses were made in a defined 3-mm3 cube in the middiaphysis 1 mm proximal of the tibial-fibular junction. The polar cross-sectional moment (moment of inertia) was also determined at the tibial-fibular junction using the parallel axis theorem. Trabecular bone analyses were performed in a region of trabecular bone defined at 0.17 mm (1% of the total length) under the growth plate of the proximal tibia extending 2 mm toward the diaphysis and excluding the outer cortical shell. Bone mineral content, mineral density, and volume fraction values were computed with a GE Healthcare MicroView software application for visualization and analysis of volumetric image data.
Statistical analysis
All statistical analyses were performed using the Excel data analysis program (Microsoft Corp., Redmond, WA)) for t test analysis. Values are expressed as the mean ± SE.
Results
IDDM was induced in adult BALB/c mice, using a multiple (5 d) streptozotocin (40 μg/g body weight) injection procedure (49). Streptozotocin selectively destroys pancreatic B cells (50). One week later, nonfasting blood glucose measurements were taken from the saphenous vein to confirm diabetes at levels greater than 300 mg glucose/dl. Control (vehicle-injected) mice tightly maintained blood glucose levels near 180 mg/dl (178 ± 5.3), whereas streptozotocin-injected mice had blood glucose levels averaging 445 ± 27 mg/dl. Serum glucose levels remained elevated throughout the study (Fig. 1). Similarly, blood osmolality, determined by a vapor pressure osmometer, was significantly elevated in diabetic compared with control mice, averaging 325 mmol/kg compared with 302 mmol/kg, respectively (Fig. 1). This suggests that tissues and cells of diabetic mice are exposed to hyperosmotic conditions. In addition to the above analyses, we examined serum triglyceride levels. Diabetic mice exhibited a significant increase in triglyceride levels throughout the study, consistent with previous studies in diabetic mice and humans (67, 68). In contrast, body weights of diabetic mice were decreased throughout the time course of the study (Fig. 1).
FIG. 1. Diabetes increases serum glucose, serum osmolality, and serum triglycerides and is associated with decreased body weight. Serum from 5-, 14-, 21-, and 28-d diabetic and control BALB/c mice was analyzed for glucose concentration, osmolality, and triglyceride levels. Body weight was recorded at every time point. Values are the average ± SE obtained from eight to 11 mice/condition. *, P < 0.05.
To determine the effect of streptozotocin-induced diabetes on the mouse skeleton, μCT analyses were carried out on the tibia, which has regions of predominantly cortical bone (the middiaphysis) and regions of trabecular bone (the metaphysis). Because differences in bone length (as a result of altered development) could add an additional variable to our analyses, we first measured and compared the lengths of control and diabetic tibias. Table 1 demonstrates that tibial length does not significantly differ between control and diabetic (4 wk postinduction) animals, consistent with the animals being on the flat part of the growth curve at the time of diabetes induction. Examination of cortical bone parameters, bone mineral content, bone mineral density, and moment of inertia, demonstrated no significant difference between control and diabetic animals (Table 1). In contrast, all trabecular bone parameters, bone mineral content, bone mineral density, and bone volume, were significantly decreased in diabetic compared with control mice (Table 1). The μCT images in Fig. 2 depict the reduction in trabecular bone within the secondary spongiosa of diabetic tibias in lateral and transverse sections.
TABLE 1. Tibia measurements in control and diabetic animals
FIG. 2. Diabetes reduces trabecular bone volume. A, Representative μCT lateral slices (20 μm) through the proximal tibia from two control and diabetic mice are shown. B, Representative μCT transverse slices (20 μm) through the proximal tibia at a distance of 0.5 or 1 mm from the lowest region of the growth plate.
Consistent with our μCT analyses, histological analyses of diabetic (4 wk after injection) proximal tibia demonstrate a decrease in bone volume marked by decreased trabecular bone volume and thickness (Table 2). Osteoblast number and osteoid parameters (volume, surface, and thickness) did not show a significant difference between control and diabetic animals (Table 2). However, dynamic measurements indicated a trend toward decreased osteoblast function/bone formation, which was significant when label incorporation/surface area was determined (3.5% vs. 1.4% in control and diabetic animals, respectively; Table 2).
TABLE 2. Histological analyses of diabetic and control tibia
Next, we examined systemic and tibial parameters of osteoclast activity. Urinary DPD, a breakdown product of collagen released systemically during osteoclast resorption and readily measured in the urine, indicated that osteoclast activity is not increased in diabetic animals 14 d after induction (Fig. 3A). Serum levels of active TRAP5b (produced by active osteoclasts) were not significantly increased in diabetic animals at any time studied and were actually decreased on d 14 and 21. Histological examination of the proximal tibia also indicated that osteoclast activity was not increased; this was indicated by the lack of a difference in osteoclast number, erosion surface, or erosion depth between control and diabetic animals (4 wk after induction; Fig. 3B). Consistent with this finding, measurement of TRAP5 mRNA levels in control and diabetic tibia did not differ (Fig. 3B).
FIG. 3. Systemic and tibia-specific measures of osteoclast activity in diabetic bone. A, Diabetes decreases serum active TRAP5b levels and has no effect on urinary DPD levels. Urine collected over a 24-h period was obtained from diabetic and control mice in a metabolic cage on d 14. DPD was determined in control and diabetic urine samples and expressed relative to creatinine, to control for differences in urine volume. Serum was obtained from mice at 5, 14, 21, or 28 d after the induction of diabetes. Active TRAP5b levels were determined in control (C) and diabetic (D) mouse serum samples. B, Histomorphometry and TRAP5 mRNA analyses were preformed on tibias obtained from mice 28 d after the induction of diabetes. All values are the average ± SE obtained from four to nine mice per condition. *, P < 0.05.
To assess whether diabetes suppresses osteoblast differentiation, we extracted total RNA from control and diabetic tibias and measured markers of progressive osteoblast differentiation: runx2, alkaline phosphatase, and osteocalcin mRNA levels. Although we did not detect a change in runx2 or alkaline phosphatase, a significant decrease in osteocalcin mRNA levels was evident in diabetic compared with control tibiae (Fig. 4). This was consistent with measurements of serum osteocalcin, which were also decreased in diabetic mice (Fig. 4). This suggests that there is a decrease in mature osteocalcin-expressing osteoblasts in the bones of diabetic animals, which supports our dynamic measurements, indicating a trend toward decreased bone formation. Additional examination of gene expression levels in mouse tibias indicated an increase in PPAR2 mRNA levels, a marker of adipocytes, in diabetic compared with control tibias (Fig. 5). Analysis of two other adipocyte markers, resistin and aP2, also demonstrated increased expression in diabetic tibia (Fig. 5). To confirm that these effects do not result from direct effects of streptozotocin on mesenchymal and osteoblast cells, we chronically treated pluripotent C3H10T1/2 cells (induced toward the osteoblast lineage or uninduced) with 0.01–1 mg/ml streptozotocin. This treatment did not induce adipogenic genes or reduce osteocalcin expression (data not shown), indicating that streptozotocin does not directly influence gene expression in these cells.
FIG. 4. Osteocalcin mRNA and serum levels are decreased in diabetes, in contrast to runx-2 and alkaline phosphatase (Alk Phos) mRNA levels. Total RNA, extracted from tibia isolated from control and diabetic (2 wk) adult BALB/c mice, was used for real-time RT-PCR analysis with SYBR Green dye. Levels of amplified genes were expressed relative to cyclophilin (a housekeeping gene) and relative to control levels, which were set at 1. Serum from 2-wk diabetic and control mice was analyzed for osteocalcin levels. Values are the average ± SE obtained from eight to 11 mice/condition. *, P < 0.05.
FIG. 5. Adipocyte markers, PPAR2, aP2, and resistin are increased in diabetic bone. Total RNA was extracted from tibia isolated from control and diabetic (2 wk) adult BALB/c mice and used for real-time RT-PCR analysis with SYBR Green dye. Levels of PPAR2, aP2, and resistin are expressed relative to cyclophilin (a housekeeping gene). Values are the average ± SE obtained from eight to 11 mice/condition. *, P < 0.05.
In agreement with RNA analyses, histological sections of diabetic tibia demonstrated an increase in marrow adiposity compared with controls (Fig. 6). Quantitation of adipocytes demonstrated that diabetic tibias contain nearly 3-fold more adipocytes (>50 μm in diameter) per area of bone marrow than control tibias (14.4 ± 4.6 vs. 4.8 ± 1.7 adipocytes/mm area of bone marrow, respectively). Interestingly, adipose tissue obtained from control femoral fat pads exhibited lipid-dense adipocytes, in contrast to diabetic adipose tissue, which exhibited lipid-sparse adipocytes (Fig. 6). Although liver architecture appeared somewhat different in diabetic compared with control animals (Fig. 6), it did not exhibit adiposity, as determined by histological examination, Oil Red-O staining (data not shown), and decreased weight (1.31 ± 0.02 vs. 1.46 ± 0.02 for controls; P < 0.001). The reduction in peripheral adiposity was also demonstrated by μCT analysis of transverse sections of mouse legs and by isolating and weighing the fat pads, which were reduced by more than 3-fold in size/weight (Fig. 7).
FIG. 6. Adiposity is increased in diabetic tibia, in contrast to peripheral adipose tissue lipolysis. Tibia (bone) obtained from control and diabetic (4 wk) mice were fixed and processed for plastic embedding and sectioning. Representative Masson-Goldner Trichrome-stained sections demonstrate differences in the amount of mineralized bone (blue stain) and in the number of adipocytes (unstained large circular regions within the marrow) between control and diabetic animals. Representative fat pad and liver sections were obtained from control and diabetic mice. Sections were stained with hematoxylin and eosin and digitally photographed.
FIG. 7. Peripheral adipose tissue is decreased in diabetic mice. Transverse slices of mouse legs were obtained by μCT analysis. Representative images are shown, with medium gray (marked by black arrows) indicating regions of fat. In addition, dissected femoral fat depots were photographed and weighed. The later values were pooled from five to nine mice per condition and are expressed as the average ± SE. *, P < 0.01.
Discussion
Bone loss is a known negative consequence of IDDM. The mechanism accounting for this outcome is unknown, making it difficult to develop optimal therapies. Diabetes is associated with a variety of pathologies, including little or no insulin, decreased IGF-I, hyperglycemia, and increased blood osmolality. An animal model is needed that is amenable to genetic manipulation to allow identification of the mechanisms accounting for diabetes-associated bone loss through knockout and/or transgenic approaches. Therefore, we set out to characterize the bone phenotype in wild-type, BALB/c, streptozotocin-induced diabetic mice.
Our histological analyses demonstrated a significant decrease in mouse trabecular bone volume (by 32%) 4 wk after the induction of diabetes. This amount is similar to bone loss reported in diabetic rats, which ranges from 10–50% (69, 70). Our μCT analyses confirm diabetes-associated bone loss in regions of trabecular bone (density decreased by 24% and volume by 69%), but no statistically significant changes were observed in regions of cortical bone. This is consistent with trabecular bone being more metabolically active than cortical bone and perhaps with an extended time cortical bone loss would become apparent. Alternatively, our results could indicate that trabecular bone (intimately associated with the marrow environment and local cytokine and growth factor stimuli) is more responsive to the diabetic environment than cortical bone (which is thought to be controlled more by systemic factors) (71).
Reports on diabetic osteoclast activity have been variable, but most studies indicate no change or a decrease in activity based on histology and/or secretion of deoxypyridinoline, a breakdown product of bone collagen matrices (69). Herrero et al. (28) demonstrate decreased pyridinium cross-links after 12 wk of diabetes in rats. Similarly, Verhaeghe et al. (29) demonstrate a decrease in urinary DPD secretion in rats diabetic for 8 wk. Our studies also did not demonstrate an increase in diabetic urinary DPD levels; however this measurement can be problematic in diabetics, in who, urine volumes are significantly increased compared with controls, filtration rates may be altered as a result of nephropathy, and collagen breakdown may be occurring in other tissues as well. Therefore, as additional measures of osteoclast activity, we examined TRAP5 mRNA and serum levels. TRAP5 mRNA levels did not differ between control and diabetic tibias at d 14 and 28 (not shown). Active TRAP5b levels in mouse serum, which is a specific systemic measure of osteoclast activity and correlates with other markers of bone turnover (72, 73), did not increase 4 wk after the induction of diabetes (when bone loss was evident), nor were they elevated at time points before this (5, 14, and 21 d). In fact, on d 14 and 21, we observed a significant decrease in active serum TRAP5b levels. A reduction in osteoclast activity could contribute to decreased osteoblast maturation through reduced osteoclast production of osteoblast-enhancing factors (74, 75). Alternatively, the reduction in osteoclast activity could be secondary to reduced osteoblast maturation and its associated decrease in osteoclast signaling (75, 76). It should also be noted that although measurements of osteoclast activity on d 5, 14, 21, and 28 were unchanged or reduced, our data do not exclude the possibility that osteoclast activity is increased at earlier time points. In summary, these results support the hypothesis that osteoclast activity is not increased, and could possibly be decreased, under diabetic conditions at the time points studied.
Examination of osteoblast maturation revealed a decrease in osteocalcin expression, a marker of late-stage osteoblast differentiation. Both tibial osteocalcin mRNA and systemic serum osteocalcin levels were reduced by more than 40%. This reduction correlates with the decrease in mineral apposition rate and label incorporation in diabetic histological sections. Given the high metabolic rate of mice, in which a 4-wk period is roughly equivalent to 1 yr in human terms, it is possible that this suppression can alone account for the bone loss we observed. Our findings of suppressed osteocalcin expression are supported by nearly all studies measuring serum osteocalcin in diabetic rats and humans, which report, on the average, a 50% decrease in osteocalcin levels compared with controls (27, 28, 77). This decrease is independent of the level of exercise the animals receive (29). Diabetes-associated suppression of osteoblast maturation is also suggested in bone implant/healing studies in rats, which indicate that diabetic animals exhibit decreased calvarial defect healing (30) and decreased bone formation around titanium or hydroxyapatite bone implants (78, 79, 80). In marrow-ablated mice, both runx2 and osteocalcin mRNA levels were decreased in diabetic relative to control bones 4 and 6 d after ablation, a time when bone formation was dramatically induced in controls (81). In our studies we did not detect a difference in runx2 mRNA levels between control and diabetic tibias. Our mRNA analyses were performed at 2 wk to examine changes in gene expression that occur before major bone loss, so the animals may not have been diabetic long enough to detect a suppression in runx2 mRNA levels in whole bone. The suppression of osteoblast differentiation in diabetes is also consistent with our in vitro analyses examining the role of hyperglycemia in the regulation of osteoblast phenotype. Specifically, we demonstrated that acute hyperglycemic as well as hyperosmotic conditions can suppress late-stage markers of osteoblast differentiation in vitro, such as osteocalcin (82). In this study we show that diabetic mice do indeed exhibit elevated plasma osmolality, which could contribute to a modulation in gene expression.
The above data support the hypothesis that diabetics lose bone through the suppression of late-stage osteoblast differentiation, whereas progression through earlier stages appears unaffected (as indicated by unaltered runx2 and alkaline phosphatase mRNA levels). However, we also propose a second pathway that contributes to diabetic bone loss, which involves the regulation of marrow adiposity. Specifically, our studies demonstrate for the first time that the number of lipid-dense adipocytes and the expression of adipocyte markers (PPAR2, resistin, and aP2) are increased in diabetic type I bone. From our experiments we cannot distinguish whether lipid-sparse adipocytes that are always present in the marrow are accumulating lipid and becoming visible or whether mesenchymal pluripotent cells are becoming adipocytes. The later could occur at the expense of osteoblast lineage selection (eventually leading to decreased osteoblast number at more extended experimental time points) or in addition to selecting the osteoblast pathway (thereby not reducing the number of immature osteoblast cell number).
Selection of adipogenesis over osteoblastogenesis is a common theme that has been reported in other conditions of bone loss, including age-related osteoporosis and disuse (44, 45, 46, 48, 83). PPAR can play a role in this selection, based on studies demonstrating that PPAR insufficiency results in the enhancement of osteogenesis and the suppression of adipogenesis in mice (84) and on studies demonstrating that elevation of PPAR2 levels can promote adipogenesis in pluripotent mesenchymal cells in vitro (43, 85). Thus, PPAR2 levels have a dominant suppressive influence on osteogenesis. In our studies, at least two possibilities can explain the function of PPAR2 elevation: either it functions as an inducer of adipogenesis or it represents a marker of increased adipocyte number and/or adipogenesis. It is known that PPAR levels and adipogenesis can be regulated by a variety of factors, including bone morphogenic protein (86, 87, 88), estrogen (89, 90), PTHrP (91), and TGF-? (92). In addition, low serum IGF-I levels can contribute to reduced bone mineral density and osteoblast differentiation and increase the number of adipocytes in bone (93, 94). The role of IGF-I is of particular interest to our IDDM animal studies, because diabetic animals and humans exhibit reduced IGF-I levels (95, 96). It is interesting that reduced IGF-I and insulin may promote marrow adipogenesis in vivo, in contrast to cell culture studies in which the lack of these factors reduces the probability of adipogenesis to occur (97).
Although elevation of PPAR2 is clearly critical for allowing PPAR2 signaling to occur, it is also important to have PPAR2 ligands available to bind to this receptor and activate its function (98). Our studies demonstrate that triglycerides are elevated in diabetic mice, and it is known that fatty acids can activate PPAR2 (98, 99) and promote differentiation of osteoblast-like cells into adipocyte-like cells (100). Thus, we hypothesize that diabetes-associated hypertriglyceridemia can compound and additional direct bone loss through promoting PPAR activity and adiposity in bone.
In summary, we demonstrate that IDDM mice exhibit bone loss. Interestingly, although animals are losing weight, under conditions of low serum insulin and IGF-I, adipogenesis or lipogenesis is still active in bone. This suggests that pluripotent cells and adipocytes in bone are somehow different or receive different signals compared with other adipose-precursor cells and adipocytes in other areas of the body. We also demonstrate that IDDM conditions suppress osteoblast maturation/function without increasing osteoclast activity. We hypothesize that an increase in bone adiposity and a decrease in mature osteoblasts contribute to the bone loss in diabetes. Understanding the mechanisms promoting bone adiposity and the importance of elevated PPAR2 expression under IDDM conditions will contribute to the development of better treatments to prevent bone loss in IDDM.
Acknowledgments
We thank Regina Irwin for her technical expertise, Drs. Laurie McCauley and Joseph Bidwell for their helpful comments, and the Investigative Histology Laboratory, Department of Physiology, Division of Human Pathology, Michigan State University.
References
Seeman E 2003 Invited review: pathogenesis of osteoporosis. J Appl Physiol 95:2142–2151
Carmeliet G, Vico L, Bouillon R 2001 Space flight: a challenge for normal bone homeostasis. Crit Rev Eukaryotic Gene Expression 11:131–144
Harpavat M, Keljo DJ, Regueiro MD 2004 Metabolic bone disease in inflammatory bowel disease. J Clin Gastroenterol 38:218–224
Boling EP 2004 Secondary osteoporosis: underlying disease and the risk for glucocorticoid-induced osteoporosis. Clin Ther 26:1–14
Diamond TH, Higano CS, Smith MR, Guise TA, Singer FR 2004 Osteoporosis in men with prostate carcinoma receiving androgen-deprivation therapy: recommendations for diagnosis and therapies. Cancer 100:892–899
Hui SL, Epstein S, Johnston Jr CC 1985 A prospective study of bone mass in patients with type I diabetes. J Clin Endocrinol Metab 60:74–80
Levin ME, Boisseau VC, Avioli LV 1976 Effects of diabetes mellitus on bone mass in juvenile and adult-onset diabetes. N Engl J Med 294:241–245
Piepkorn B, Kann P, Forst T, Andreas J, Pfutzner A, Beyer J 1997 Bone mineral density and bone metabolism in diabetes mellitus. Horm Metab Res 29:584–591
Kemink SA, Hermus AR, Swinkels LM, Lutterman JA, Smals AG 2000 Osteopenia in insulin-dependent diabetes mellitus; prevalence and aspects of pathophysiology. J Endocrinol Invest 23:295–303
Gunczler P, Lanes R, Paz-Martinez V, Martins R, Esaa S, Colmenares V, Weisinger JR 1998 Decreased lumbar spine bone mass and low bone turnover in children and adolescents with insulin dependent diabetes mellitus followed longitudinally. J Pediatr Endocrinol Metab 11:413–419
Buysschaert M, Cauwe F, Jamart J, Brichant C, De Coster P, Magnan A, Donckier J 1992 Proximal femur density in type 1 and 2 diabetic patients. Diabetes Metab 18:32–37
Tuominen JT, Impivaara O, Puukka P, Ronnemaa T 1999 Bone mineral density in patients with type 1 and type 2 diabetes. Diabetes Care 22:1196–1200
Hampson G, Evans C, Petitt RJ, Evans WD, Woodhead SJ, Peters JR, Ralston SH 1998 Bone mineral density, collagen type 1 1 genotypes and bone turnover in premenopausal women with diabetes mellitus. Diabetologia 41:1314–1320
Wiske PS, Wentworth SM, Norton Jr JA, Epstein S, Johnston Jr CC 1982 Evaluation of bone mass and growth in young diabetics. Metabolism 31:848–854
McNair P, Christiansen C, Christensen MS, Madsbad S, Faber OK, Binder C, Transbol I 1981 Development of bone mineral loss in insulin-treated diabetes: a 1 1/2 years follow-up study in sixty patients. Eur J Clin Invest 11:55–59
Auwerx J, Dequeker J, Bouillon R, Geusens P, Nijs J 1988 Mineral metabolism and bone mass at peripheral and axial skeleton in diabetes mellitus. Diabetes 37:8–12
Krakauer JC, McKenna MJ, Buderer NF, Rao DS, Whitehouse FW, Parfitt AM 1995 Bone loss and bone turnover in diabetes. Diabetes 44:775–782
Munoz-Torres M, Jodar E, Escobar-Jimenez F, Lopez-Ibarra PJ, Luna JD 1996 Bone mineral density measured by dual x-ray absorptiometry in Spanish patients with insulin-dependent diabetes mellitus. Calcif Tissue Int 58:316–319
Bouillon R 1991 Diabetic bone disease. Calcif Tissue Int 49:155–160
Meyer HE, Tverdal A, Falch JA 1993 Risk factors for hip fracture in middle-aged Norwegian women and men. Am J Epidemiol 137:1203–1211
Schwartz AV, Sellmeyer DE, Ensrud KE, Cauley JA, Tabor HK, Schreiner PJ, Jamal SA, Black DM, Cummings SR 2001 Older women with diabetes have an increased risk of fracture: a prospective study. J Clin Endocrinol Metab 86:32–38
Forsen L, Meyer HE, Midthjell K, Edna TH 1999 Diabetes mellitus and the incidence of hip fracture: results from the Nord-Trondelag Health Survey. Diabetologia 42:920–925
White CB, Turner NS, Lee GC, Haidukewych GJ 2003 Open ankle fractures in patients with diabetes mellitus. Clin Orthop 414:37–44
Folk JW, Starr AJ, Early JS 1999 Early wound complications of operative treatment of calcaneus fractures: analysis of 190 fractures. J Orthop Trauma 13:369–372
Herskind AM, Christensen K, Norgaard-Andersen K, Andersen JF 1992 Diabetes mellitus and healing of closed fractures. Diabetes Metab 18:63–64
Bouillon R, Bex M, Van Herck E, Laureys J, Dooms L, Lesaffre E, Ravussin E 1995 Influence of age, sex, and insulin on osteoblast function: osteoblast dysfunction in diabetes mellitus. J Clin Endocrinol Metab 80:1194–1202
Verhaeghe J, van Herck E, Visser WJ, Suiker AM, Thomasset M, Einhorn TA, Faierman E, Bouillon R 1990 Bone and mineral metabolism in BB rats with long-term diabetes. Decreased bone turnover and osteoporosis. Diabetes 39:477–482
Herrero S, Calvo OM, Garcia-Moreno C, Martin E, San Roman JI, Martin M, Garcia-Talavera JR, Calvo JJ, del Pino-Montes J 1998 Low bone density with normal bone turnover in ovariectomized and streptozotocin-induced diabetic rats. Calcif Tissue Int 62:260–265
Verhaeghe J, Thomsen JS, van Bree R, van Herck E, Bouillon R, Mosekilde L 2000 Effects of exercise and disuse on bone remodeling, bone mass, and biomechanical competence in spontaneously diabetic female rats. Bone 27:249–256
Shyng YC, Devlin H, Sloan P 2001 The effect of streptozotocin-induced experimental diabetes mellitus on calvarial defect healing and bone turnover in the rat. Int J Oral Maxillofac Surg 30:70–74
Sasaki T, Kaneko H, Ramamurthy NS, Golub LM 1991 Tetracycline administration restores osteoblast structure and function during experimental diabetes. Anat Rec 231:25–34
McCracken M, Lemons JE, Rahemtulla F, Prince CW, Feldman D 2000 Bone response to titanium alloy implants placed in diabetic rats. Int J Oral Maxillofac Implants 15:345–354
Ferrari G, Cusella-De Angelis G, Coletta M, Paolucci E, Stornaiuolo A, Cossu G, Mavilio F 1998 Muscle regeneration by bone marrow-derived myogenic progenitors. Science 279:1528–1530
Krause DS, Theise ND, Collector MI, Henegariu O, Hwang S, Gardner R, Neutzel S, Sharkis SJ 2001 Multi-organ, multi-lineage engraftment by a single bone marrow-derived stem cell. Cell 105:369–377
Petersen BE, Bowen WC, Patrene KD, Mars WM, Sullivan AK, Murase N, Boggs SS, Greenberger JS, Goff JP 1999 Bone marrow as a potential source of hepatic oval cells. Science 284:1168–1170
Sanchez-Ramos J, Song S, Cardozo-Pelaez F, Hazzi C, Stedeford T, Willing A, Freeman TB, Saporta S, Janssen W, Patel N, Cooper DR, Sanberg PR 2000 Adult bone marrow stromal cells differentiate into neural cells in vitro. Exp Neurol 164:247–256
Karsenty G 2001 Minireview: transcriptional control of osteoblast differentiation. Endocrinology 142:2731–2733
Stein GS, Lian JB, van Wijnen AJ, Stein JL, Montecino M, Javed A, Zaidi SK, Young DW, Choi JY, Pockwinse SM 2004 Runx2 control of organization, assembly and activity of the regulatory machinery for skeletal gene expression. Oncogene 23:4315–4329
Lian JB, Stein GS, Stein JL, van Wijnen AJ 1998 Osteocalcin gene promoter: unlocking the secrets for regulation of osteoblast growth and differentiation. J Cell Biochem 30–31(Suppl):62–72
Winchester SK, Selvamurugan N, D’Alonzo RC, Partridge NC 2000 Developmental regulation of collagenase-3 mRNA in normal, differentiating osteoblasts through the activator protein-1 and the runt domain binding sites. J Biol Chem 275:23310–23318
Sasano Y, Zhu JX, Tsubota M, Takahashi I, Onodera K, Mizoguchi I, Kagayama M 2002 Gene expression of MMP8 and MMP13 during embryonic development of bone and cartilage in the rat mandible and hind limb. J Histochem Cytochem 50:325–332
Rosen ED, Spiegelman BM 2001 PPAR: a nuclear regulator of metabolism, differentiation, and cell growth. J Biol Chem 276:37731–37734[Free Full Text]
Lecka-Czernik B, Gubrij I, Moerman EJ, Kajkenova O, Lipschitz DA, Manolagas SC, Jilka RL 1999 Inhibition of Osf2/Cbfa1 expression and terminal osteoblast differentiation by PPAR2. J Cell Biochem 74:357–371
Jilka RL, Weinstein RS, Takahashi K, Parfitt AM, Manolagas SC 1996 Linkage of decreased bone mass with impaired osteoblastogenesis in a murine model of accelerated senescence. J Clin Invest 97:1732–1740
Kajkenova O, Lecka-Czernik B, Gubrij I, Hauser SP, Takahashi K, Parfitt AM, Jilka RL, Manolagas SC, Lipschitz DA 1997 Increased adipogenesis and myelopoiesis in the bone marrow of SAMP6, a murine model of defective osteoblastogenesis and low turnover osteopenia. J Bone Miner Res 12:1772–1779
Verma S, Rajaratnam JH, Denton J, Hoyland JA, Byers RJ 2002 Adipocytic proportion of bone marrow is inversely related to bone formation in osteoporosis. J Clin Pathol 55:693–698
Moerman EJ, Teng K, Lipschitz DA, Lecka-Czernik B 2004 Aging activates adipogenic and suppresses osteogenic programs in mesenchymal marrow stroma/stem cells: the role of PPAR-2 transcription factor and TGF-?/BMP signaling pathways. Aging Cell 3:379–389
Nuttall ME, Gimble JM 2004 Controlling the balance between osteoblastogenesis and adipogenesis and the consequent therapeutic implications. Curr Opin Pharmacol 4:290–294
Pechhold K, Patterson NB, Blum C, Fleischacker CL, Boehm BO, Harlan DM 2001 Low dose streptozotocin-induced diabetes in rat insulin promoter-mCD80-transgenic mice is T cell autoantigen-specific and CD28 dependent. J Immunol 166:2531–2539
Szkudelski T 2001 The mechanism of alloxan and streptozotocin action in B cells of the rat pancreas. Physiol Res 50:537–546
McCabe LR, Kockx M, Lian J, Stein J, Stein G 1995 Selective expression of fos- and jun-related genes during osteoblast proliferation and differentiation. Exp Cell Res 218:255–262
Chomczynski P, Sacchi N 1987 Single-step method of RNA isolation by acid guanidinium thiocyanate-phenol-chloroform extraction. Anal Biochem 162:156–159
Ontiveros C, McCabe LR 2003 Simulated microgravity suppresses osteoblast phenotype, Runx2 levels and AP-1 transactivation. J Cell Biochem 88:427–437
Ontiveros C, Irwin R, Wiseman RW, McCabe LR 2004 Hypoxia suppresses runx2 independent of modeled microgravity. J Cell Physiol 200:169–176
Kast-Woelbern HR, Dana SL, Cesario RM, Sun L, de Grandpre LY, Brooks ME, Osburn DL, Reifel-Miller A, Klausing K, Leibowitz MD 2004 Rosiglitazone induction of Insig-1 in white adipose tissue reveals a novel interplay of peroxisome proliferator-activated receptor and sterol regulatory element-binding protein in the regulation of adipogenesis. J Biol Chem 279:23908–23915
Li J, Takaishi K, Cook W, McCorkle SK, Unger RH 2003 Insig-1 "brakes" lipogenesis in adipocytes and inhibits differentiation of preadipocytes. Proc Natl Acad Sci USA 100:9476–9481
Wiren KM, Zhang XW, Toombs AR, Kasparcova V, Gentile MA, Harada S, Jepsen KJ 2004 Targeted overexpression of androgen receptor in osteoblasts: unexpected complex bone phenotype in growing animals. Endocrinology 145:3507–3522
Harada K, Shen WJ, Patel S, Natu V, Wang J, Osuga J, Ishibashi S, Kraemer FB 2003 Resistance to high-fat diet-induced obesity and altered expression of adipose-specific genes in HSL-deficient mice. Am J Physiol 285:E1182–E1195
Lewis ML, Hughes-Fulford M 2000 Regulation of heat shock protein message in Jurkat cells cultured under serum-starved and gravity-altered conditions. J Cell Biochem 77:127–134
Trogan E, Choudhury RP, Dansky HM, Rong JX, Breslow JL, Fisher EA 2002 Laser capture microdissection analysis of gene expression in macrophages from atherosclerotic lesions of apolipoprotein E-deficient mice. Proc Natl Acad Sci USA 99:2234–2239
Date T, Doiguchi Y, Nobuta M, Shindo H 2004 Bone morphogenetic protein-2 induces differentiation of multipotent C3H10T1/2 cells into osteoblasts, chondrocytes, and adipocytes in vivo and in vitro. J Orthop Sci 9:503–508
Malluche HH, Faugere MC 1986 Atlas of mineralized bone histology. New York: Karger
Goldner J 1938 A modification of the Masson Trichrome technique for routine laboratory purposes. Am J Pathol 14:237–243
Manaka RC, Malluche HH 1981 A program package for quantitative analysis of histologic structure and remodeling dynamics of bone. Comput Programs Biomed 13:191–202
Malluche HH, Monier-Faugere MC 1994 The role of bone biopsy in the management of patients with renal osteodystrophy. J Am Soc Nephrol 4:1631–1642
Parfitt AM, Drezner MK, Glorieux FH, Kanis JA, Malluche HH, Meunier PJ, Ott SM, Recker RR 1987 Bone histomorphometry: standardization of nomenclature, symbols and units. J Bone Miner Res 6:595–610
Miljanovic B, Glynn RJ, Nathan DM, Manson JE, Schaumberg DA 2004 A prospective study of serum lipids and risk of diabetic macular edema in type 1 diabetes. Diabetes 53:2883–2892
Greenwalt DE, Scheck SH, Rhinehart-Jones T 1995 Heart CD36 expression is increased in murine models of diabetes and in mice fed a high fat diet. J Clin Invest 96:1382–1388
Locatto ME, Abranzon H, Caferra D, Fernandez MC, Alloatti R, Puche RC 1993 Growth and development of bone mass in untreated alloxan diabetic rats. Effects of collagen glycosylation and parathyroid activity on bone turnover. Bone Miner 23:129–144
Waud CE, Marks Jr SC, Lew R, Baran DT 1994 Bone mineral density in the femur and lumbar vertebrae decreases after twelve weeks of diabetes in spontaneously diabetic-prone BB/Worcester rats. Calcif Tissue Int 54:237–240
Mundy GR, Chen D, Oyajobi BO 2003 Bone Remodeling. In: Favus M, ed. Primer on the metabolic bone diseases and disorders of mineral metabolism, 5th Ed. Washington DC: American Society of Bone and Mineral Research; 46–58
Halleen JM, Alatalo SL, Suominen H, Cheng S, Janckila AJ, Vaananen HK 2000 Tartrate-resistant acid phosphatase 5b: a novel serum marker of bone resorption. J Bone Miner Res 15:1337–1345
Halleen JM, Ylipahkala H, Alatalo SL, Janckila AJ, Heikkinen JE, Suominen H, Cheng S, Vaananen HK 2002 Serum tartrate-resistant acid phosphatase 5b, but not 5a, correlates with other markers of bone turnover and bone mineral density. Calcif Tissue Int 71:20–25
Dai XM, Zong XH, Akhter MP, Stanley ER 2004 Osteoclast deficiency results in disorganized matrix, reduced mineralization, and abnormal osteoblast behavior in developing bone. J Bone Miner Res 19:1441–1451
Phan TC, Xu J, Zheng MH 2004 Interaction between osteoblast and osteoclast: impact in bone disease. Histol Histopathol 19:1325–1344
Kwan Tat S, Padrines M, Theoleyre S, Heymann D, Fortun Y 2004 IL-6, RANKL, TNF-/IL-1: interrelations in bone resorption pathophysiology. Cytokine Growth Factor Rev 15:49–60
Epstein S, Takizawa M, Stein B, Katz IA, Joffe, II, Romero DF, Liang XG, Li M, Ke HZ, Jee WS 1994 Effect of cyclosporin A on bone mineral metabolism in experimental diabetes mellitus in the rat. J Bone Miner Res 9:557–566
Siqueira JT, Cavalher-Machado SC, Arana-Chavez VE, Sannomiya P 2003 Bone formation around titanium implants in the rat tibia: role of insulin. Implant Dent 12:242–251
Giglio MJ, Giannunzio G, Olmedo D, Guglielmotti MB 2000 Histomorphometric study of bone healing around laminar implants in experimental diabetes. Implant Dent 9:143–149
Takeshita F, Iyama S, Ayukawa Y, Kido MA, Murai K, Suetsugu T 1997 The effects of diabetes on the interface between hydroxyapatite implants and bone in rat tibia. J Periodontol 68:180–185
Lu H, Kraut D, Gerstenfeld LC, Graves DT 2003 Diabetes interferes with the bone formation by affecting the expression of transcription factors that regulate osteoblast differentiation. Endocrinology 144:346–352
Zayzafoon M, Stell C, Irwin R, McCabe LR 2000 Extracellular glucose influences osteoblast differentiation and c-Jun expression. J Cell Biochem 79:301–310
Meunier P, Aaron J, Edouard C, Vignon G 1971 Osteoporosis and the replacement of cell populations of the marrow by adipose tissue. A quantitative study of 84 iliac bone biopsies. Clin Orthop 80:147–154
Akune T, Ohba S, Kamekura S, Yamaguchi M, Chung UI, Kubota N, Terauchi Y, Harada Y, Azuma Y, Nakamura K, Kadowaki T, Kawaguchi H 2004 PPAR insufficiency enhances osteogenesis through osteoblast formation from bone marrow progenitors. J Clin Invest 113:846–855
Rzonca SO, Suva LJ, Gaddy D, Montague DC, Lecka-Czernik B 2004 Bone is a target for the antidiabetic compound rosiglitazone. Endocrinology 145:401–406
Skillington J, Choy L, Derynck R 2002 Bone morphogenetic protein and retinoic acid signaling cooperate to induce osteoblast differentiation of preadipocytes. J Cell Biol 159:135–146
Fux C, Mitta B, Kramer BP, Fussenegger M 2004 Dual-regulated expression of C/EBP- and BMP-2 enables differential differentiation of C2C12 cells into adipocytes and osteoblasts. Nucleic Acids Res 32:e1
Chen D, Ji X, Harris MA, Feng JQ, Karsenty G, Celeste AJ, Rosen V, Mundy GR, Harris SE 1998 Differential roles for bone morphogenetic protein (BMP) receptor type IB and IA in differentiation and specification of mesenchymal precursor cells to osteoblast and adipocyte lineages. J Cell Biol 142:295–305
Okazaki R, Inoue D, Shibata M, Saika M, Kido S, Ooka H, Tomiyama H, Sakamoto Y, Matsumoto T 2002 Estrogen promotes early osteoblast differentiation and inhibits adipocyte differentiation in mouse bone marrow stromal cell lines that express estrogen receptor (ER) or ?. Endocrinology 143:2349–2356
Dang Z, Lowik CW 2004 The balance between concurrent activation of ERs and PPARs determines daidzein-induced osteogenesis and adipogenesis. J Bone Miner Res 19:853–861
Chan GK, Miao D, Deckelbaum R, Bolivar I, Karaplis A, Goltzman D 2003 Parathyroid hormone-related peptide interacts with bone morphogenetic protein 2 to increase osteoblastogenesis and decrease adipogenesis in pluripotent C3H10T 1/2 mesenchymal cells. Endocrinology 144:5511–5520
Ahdjoudj S, Lasmoles F, Holy X, Zerath E, Marie PJ 2002 Transforming growth factor ?2 inhibits adipocyte differentiation induced by skeletal unloading in rat bone marrow stroma. J Bone Miner Res 17:668–677
Rosen CJ, Ackert-Bicknell CL, Adamo ML, Shultz KL, Rubin J, Donahue LR, Horton LG, Delahunty KM, Beamer WG, Sipos J, Clemmons D, Nelson T, Bouxsein ML, Horowitz M 2004 Congenic mice with low serum IGF-I have increased body fat, reduced bone mineral density, and an altered osteoblast differentiation program. Bone 35:1046–1058
Zhang M, Xuan S, Bouxsein ML, von Stechow D, Akeno N, Faugere MC, Malluche H, Zhao G, Rosen CJ, Efstratiadis A, Clemens TL 2002 Osteoblast-specific knockout of the insulin-like growth factor (IGF) receptor gene reveals an essential role of IGF signaling in bone matrix mineralization. J Biol Chem 277:44005–44012
Verhaeghe J, Oloumi G, van Herck E, van Bree R, Dequeker J, Einhorn TA, Bouillon R 1997 Effects of long-term diabetes and/or high-dose 17 ?-estradiol on bone formation, bone mineral density, and strength in ovariectomized rats. Bone 20:421–428
Jehle PM, Jehle DR, Mohan S, Bohm BO 1998 Serum levels of insulin-like growth factor system components and relationship to bone metabolism in type 1 and type 2 diabetes mellitus patients. J Endocrinol 159:297–306
MacDougald OA, Lane MD 1995 Transcriptional regulation of gene expression during adipocyte differentiation. Annu Rev Biochem 64:345–373
Lecka-Czernik B, Moerman EJ, Grant DF, Lehmann JM, Manolagas SC, Jilka RL 2002 Divergent effects of selective peroxisome proliferator-activated receptor-2 ligands on adipocyte versus osteoblast differentiation. Endocrinology 143:2376–2384
Forman BM, Tontonoz P, Chen J, Brun RP, Spiegelman BM, Evans RM 1995 15-Deoxy-12,14-prostaglandin J2 is a ligand for the adipocyte determination factor PPAR. Cell 83:803–812
Diascro Jr DD, Vogel RL, Johnson TE, Witherup KM, Pitzenberger SM, Rutledge SJ, Prescott DJ, Rodan GA, Schmidt A 1998 High fatty acid content in rabbit serum is responsible for the differentiation of osteoblasts into adipocyte-like cells. J Bone Miner Res 13:96–106(Sergiu Botolin, Marie-Cla)