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Open Access

Peer-reviewed

Research Article

Protection against T1DM-Induced Bone Loss by Zinc Supplementation: Biomechanical, Histomorphometric, and Molecular Analyses in STZ-Induced Diabetic Rats

Abstract

Several studies have established an association between diabetes and alterations in bone metabolism; however, the underlying mechanism is not well established. Although zinc is recognized as a potential preventive agent against diabetes-induced bone loss, there is no evidence demonstrating its effect in chronic diabetic conditions. This study evaluated the effects of zinc supplementation in a chronic (90 days) type 1 diabetes-induced bone-loss model. Male Wistar rats were distributed in three groups: control, type 1 diabetes mellitus (T1DM), and T1DM plus zinc supplementation (T1DMS). Serum biochemical analysis; tibia histomorphometric, biomechanical, and collagen-content analyses; and femur mRNA expression were evaluated. Relative to T1DM, the zinc-supplemented group showed increased histomorphometric parameters such as TbWi and BAr and decreased TbSp, increased biomechanical parameters (maximum load, stiffness, ultimate strain, and Young’s modulus), and increased type I collagen content. Interestingly, similar values for these parameters were observed between the T1DMS and control groups. These results demonstrate the protective effect of zinc on the maintenance of bone strength and flexibility. In addition, downregulation of OPG, COL1A, and MMP-9 genes was observed in T1DMS, and the anabolic effects of zinc were evidenced by increased OC expression and serum ALP activity, both related to osteoblastogenesis, demonstrating a positive effect on bone formation. In contrast, T1DM showed excessive bone loss, observed through reduced histomorphometric and biomechanical parameters, characterizing diabetes-associated bone loss. The bone loss was also observed through upregulation of OPG, COL1A, and MMP-9 genes. In conclusion, zinc showed a positive effect on the maintenance of bone architecture and biomechanical parameters. Indeed, OC upregulation and control of expression of OPG, COL1A, and MMP-9 mRNAs, even in chronic hyperglycemia, support an anabolic and protective effect of zinc under chronic diabetic conditions. Furthermore, these results indicate that zinc supplementation could act as a complementary therapy in chronic T1DM.

Citation: Bortolin RH, da Graça Azevedo Abreu BJ, Abbott Galvão Ururahy M, Costa de Souza KS, Bezerra JF, Bezerra Loureiro M, et al. (2015) Protection against T1DM-Induced Bone Loss by Zinc Supplementation: Biomechanical, Histomorphometric, and Molecular Analyses in STZ-Induced Diabetic Rats. PLoS ONE 10(5): e0125349. https://doi.org/10.1371/journal.pone.0125349

Academic Editor: Dominique Heymann, Faculté de médecine de Nantes, FRANCE

Received: December 1, 2014; Accepted: March 16, 2015; Published: May 1, 2015

Copyright: © 2015 Bortolin et al. This is an open access article distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited

Data Availability: All relevant data are within the paper.

Funding: BJGAA received financial support by National Council of scientific and technological development (protocol 475801/2010). The funders had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript.

Competing interests: The authors have declared that no competing interests exist.

Introduction

Type 1 diabetes mellitus (T1DM) is a chronic disease in which pancreatic beta cells are selectively destroyed, leading to chronic hyperglycemia [1], and several consequential long-term vascular complications such as retinopathy, neuropathy, and nephropathy have been reported [1–3]. Although skeletal abnormalities and bone disease represent an overlooked complication of diabetes, the relationship is well established and recognized as a complex pathogenesis including mechanical, hormonal, and vascular factors involving an imbalance between bone formation and resorption.

Diabetes-induced bone-loss mechanisms are not fully understood [4–7] although a variety of bone-related changes are known to be influenced by hyperglycemia such as bone mineral density, femoral neck geometry, microarchitecture (trabecular, cortical thickness, and bone area), and biomechanical markers of bone turnover (ultimate strain, strength and load, stiffness, and Young’s modulus) [3,8–11]. In addition, formation of the collagen network is affected by an increase in matrix metalloproteinase (MMP) expression in diabetic conditions, especially for MMP-9, which is considered a diabetogenic factor and is upregulated in T1DM, contributing to collagen degradation and resulting in low bone collagen content and poor bone biomechanical integrity [12–15]. One study showed a decrease in bone mineral content (BMC) associated with an increase in urinary calcium excretion in diabetic rats. Moreover, low BMC in diabetic conditions has been associated with low osteocalcin (OC) levels, as this small non-collagenous protein is produced by osteoblasts and is directly involved in bone inorganic matrix development [16].

Several studies have reported controversial alterations in the RANK/RANKL/OPG (receptor activator of nuclear factor kappa β/receptor activator of nuclear factor kappa β ligand/osteoprotegerin) system in hyperglycemic conditions [2,4,8,17–28]. Some studies have shown an increase in RANKL mRNA expression in diabetic bone [2] and decreases in a high-glucose in vitro model [18] and diabetic bone [20]. OPG expression has been shown to increase in young T1DM patients [8], whereas a decrease in gene expression was also observed in T1DM patients [4], in a T1DM animal model [19], and in an in vitro study [17].

As an alternative, investigators have used zinc supplementation for bone-loss prevention in both healthy and hyperglycemic conditions because it is an essential element in bone metabolism, acting as a cofactor for several enzymes and stimulating gene expression of various proteins necessary for bone mineralization and collagenous structure development [10,21–25].

Studies involving in vitro and in vivo models have evaluated the efficacy of zinc supplementation in preventing bone loss [22,23,26–28]. In vitro results have shown a stimulatory effect on osteoblastogenesis through increases in DNA, collagen, calcium, insulin-like growth factor 1 (IGF-I), transforming growth factor beta 1 (TGF-β1), alkaline phosphatase (ALP) activity, and OC [21–22]. Furthermore, an in vivo study involving acute T1DM-induced bone loss and zinc supplementation showed a significant effect of zinc on bone formation associated with an increase in OC mRNA expression [23].

A positive effect of zinc on the recovery of bone architecture has been reported in acute diabetic conditions [23], and bone biomechanical tests in rats showed that zinc maintained overall bone quality and increased fracture resistance [24–25]. Moreover, the effects of zinc on the RANK/RANKL/OPG system has been reported by Yamaguchi [22], who demonstrated the inhibition of RANKL expression in pre-osteoclasts and the stimulation of OPG gene expression in osteoblastic cells, which could act as a decoy receptor by binding to RANKL and preventing RANK signaling.

Although a few reports have shown positive effects of zinc supplementation, all of these studies were performed during an acute period between 7 and 21 days [23,29]. Our aim was to evaluate the effect of zinc under long-term diabetic conditions to provide proof-of-concept for the potential use of zinc supplementation in preventing chronic T1DM-induced bone loss. To the best of our knowledge, this is the first study to evaluate the effects of zinc supplementation over a period of 90 days, which represents a chronic model of diabetes, and provide evidence that zinc ingested as dietary supplement can prevent bone loss through anabolic and osteo-protective effects. We show that this in vivo action results from the stimulation of bone formation and decreased bone resorption as detected by histomorphometric, collagen-content, biomechanical, and quantitative reverse transcription PCR (RT-qPCR) analyses. Together with prior evidence showing a relationship between supplementation and bone metabolism, our data provide compelling evidence for the therapeutic potential of zinc supplementation as a complementary therapy against chronic T1DM-induced bone loss.

Methods

Experimental Protocol

All animal experiments and protocols were approved by the Committee on the Ethics of Animal Use and Care of the Federal University of Rio Grande do Norte (permit number 022/2009) and the Committee on the Ethics in Research of the University of Ribeirão Preto (permit number 066/09). All procedures were carried out in strict accordance with the recommendations in the Guide for the Care and Use of Laboratory Animals of the National Institutes of Health [30]. All surgery was performed under thiopental anesthesia, and all efforts were made to minimize suffering.

Fifteen male Wistar rats weighing 220 ± 20 g were obtained from the Laboratory Animal Facility of the University of Ribeirão Preto, Ribeirão Preto, Brazil. During the study period, the animals were housed in standard conditions (12 h light/dark cycle, 22–24°C, and 50–60% humidity) with food and water ad libitum. After one week of acclimatization prior to the experimental procedures, the rats were randomly assigned and equally distributed (five rats per group) to three groups: control, T1DM, and T1DM plus zinc supplementation (T1DMS).

Experimental diabetes was induced by a single intravenous injection of streptozotocin (STZ, Sigma-Aldrich, St. Louis, MO, USA) dissolved in freshly prepared Na citrate buffer (0.1 M, pH 4.5) at a dose of 40 mg/kg of body weight. Equal volumes of vehicle were injected in the control rats. On day 0, i.e., day 5 after induction, blood samples were collected by tail bleeding, and glycemia was assayed using an ACCU-CHEK Advantage glucometer (Roche Diagnostics, Indianapolis, IN, USA). Animals with blood glucose concentrations ≥250 mg/dL were considered diabetic and started receiving a standard (control and T1DM) or supplemented diet (T1DMS). The blood glucose concentrations and body weight were monitored fortnightly for 12 weeks. Clinical diabetic signs such as polyphagia, polydipsia, polyuria, and body weight loss were also monitored [31,32].

Standard diets were formulated in accordance with rodent-specific rules established by the American Institute of Nutrition in 1993 (AIN-93) [33]. Dietary ingredients were provided by Rhoster Industry and Trade Ltd. (São Paulo, Brazil). The control and T1DM groups were fed daily with 20 g of standard diet for 12 weeks. Because the AIN-93 standard diet contains 30 mg zinc/kg diet, we supplemented the diet with 500 mg zinc/kg diet; thus, the T1DMS group was fed a standard diet supplemented with 17-fold geometrically diluted ZnCO3. All animals in the T1DMS group consumed 20 g of the supplemented diet daily during the 12 weeks of study, totaling 10.6 mg ZnCO3 ingestion daily.

Blood and tissue collection and routine biochemical analyses

All animals were euthanized by a lethal dose of thiopental (100 mg/kg), and blood samples were obtained from the abdominal aorta. To prevent possible daily cyclic variations of the measurements, all animals were euthanized between 7:00 am and 9:00 am. The femurs and tibias were harvested and stored for subsequent total RNA extraction and histomorphometric, biomechanical, and collagen-content analyses.

Serum glucose concentration and ALP activity were determined in triplicate using routine methods (BioSystems Reagents and Instruments, Barcelona, Spain) and performed in an RA 50 spectrophotometer (Chemistry System Bayer Diagnostic, Dublin, Ireland). Serum zinc was measured by atomic absorption spectroscopy using a Spectra AA-200 spectrophotometer (Varian Canada, Georgetown, Ontario, Canada).

Histomorphometric analyses

The right tibia of each animal was fixed in a solution of 10% buffered formalin and processed after decalcification in 7.5% nitric acid and embedding in paraffin following standard procedures as described by Duarte et al. [34] with modifications. Longitudinal 7-μm sections were stained with hematoxylin and eosin. The histomorphometric results are presented as the means of four measurements of trabecular separation (TbSp, μm), trabecular width (TbWi, μm) and trabecular bone area (BAr, %) obtained from the metadiaphyseal region using a Nikon Lobophot microscope equipped with a 10× magnification ocular lens (Nikon, Tokyo, Japan). After analysis using ImageJ 1.48v (National Institutes of Health, Bethesda, MD, USA), the results were reported in μm. All parameters complied with the guidelines of the Nomenclature Committee of the American Society of Bone and Mineral Research [35].

Collagen content

Paraffin-embedded samples were also used for collagen quantification by picrosirius red staining according to the procedure described by Rich and Whittaker [36] with modifications. To evaluate the collagen content in sections stained with picrosirius red, four fields of the metadiaphyseal region were observed with a 10× magnification ocular lens in an AxioImager M2 microscope (Carl Zeiss, Jena, Germany). In each field, the percentages of tissue area stained in red and green relative to the total tissue area were calculated according to the formula described by Black et al. [37]. All analyses were performed using ImageJ 1.48v.

Biomechanical testing

Biomechanical analysis was performed on the left tibias previously stored at −80°C using three-point bending mechanical tests according to the procedure described by Korres et al. [12], with modifications. We used a servo hydraulic high-precision universal testing machine, model AG-X 10 kN (Shimadzu Corporation, Tokyo, Japan). Tibias were placed horizontally on the frame with rounded edges at a distance of 30 mm. The load was applied at the mid-shaft of the diaphysis using a punch with a rounded notch. The rate of the imposed displacement was selected as 5 mm/min to simulate static loading conditions. The displacement was imposed continuously until fracture. Failure in the load-displacement curves was defined and observed by the propagation of a nearly vertical fracture starting almost universally at the lower cortical bone surface. Ultimate load, stiffness, ultimate stress, ultimate strain, and Young’s modulus were recorded.

RNA extraction and RT-qPCR

The right femurs of the animals, previously stored at −80°C, were pulverized, and total RNA was extracted using an RNeasy Plus Mini Kit (Qiagen, Valencia, CA, USA). The RNA integrity was assessed by electrophoresis in 1.0% agarose gels with MOPS buffer, concentrations were measured using a Nanodrop ND-1000 spectrophotometer (Thermo Scientific, Wilmington, DE, USA), and RNA was stored at −80°C. Synthesis of cDNA was performed with 1 μg total RNA using a High Capacity cDNA Reverse Transcription Kit (Applied Biosystems, Foster City, CA, USA), according to the manufacturer’s protocol in a MyCycler Thermal Cycler (Bio-Rad, Philadelphia, PA, USA). The cDNA was obtained in a final volume of 50 μL and stored at −20°C until it was used for the RT-qPCR expression assays.

RT-qPCR was performed on the following genes using the TaqMan Assay: RANKL (Rn00589289_m1), OPG (Rn00563499_m1), OC (Rn00566386_g1), COL1A1 (Rn01463848_m1), MMP-2 (Rn01538170_m1), MMP-9 (Rn00579162_m1), and glyceraldehyde-3-phosphate dehydrogenase (GAPDH, Rn00579162_m1) (Applied Biosystems). PCR assays were carried out in 96-well plates using a 7500 Fast Real-time PCR System (Applied Biosystems). Relative expression was calculated using the 2-ΔΔCT method [38], and results are presented as fold-change versus the control group mean values, normalized to GAPDH; Ct did not show significant variation between the control and T1DM groups.

Statistical analyses

Statistical analyses were performed with GraphPad PRISM version 5.0 (GraphPad Software Inc., San Diego, CA, USA). In all data, the normality test failed, and we therefore used the nonparametric Kruskal-Wallis ANOVA on Ranks and Dunn’s post-hoc method of multiple comparisons versus control. p-values < 0.05 were considered statistically significant.

Results

Biochemical analyses and body weight

Biochemical analyses and body weight results are shown in Table 1. As expected, blood glucose concentrations in T1DM and T1DMS rats were than those in control rats (p < 0.001). Hyperglycemia was associated with polyphagia, polydipsia, and polyuria (data not shown) in the diabetic rats, indicating that experimental diabetes was successfully induced. The baseline body weight at the beginning of the study was similar in control and diabetic groups (average 220 ± 20 g). However, after 90 days of the experimental period, which represents a chronic condition, T1DM and T1DMS groups showed significantly reduced body weight (p < 0.001).

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Table 1. Biochemical analyses and body weight of control, diabetic, and diabetic plus zinc supplementation groups.

https://doi.org/10.1371/journal.pone.0125349.t001

No significant difference was observed in serum ALP activity between the T1DM and control groups. However, ALP activity was significantly higher in the T1DMS group than in the control and T1DM groups (p < 0.001).

Serum zinc concentration was decreased in T1DM rats compared to control rats (p < 0.05), and we observed the expected increase in serum zinc concentration in T1DMS rats compared to the control and T1DM groups (p < 0.01).

Histological and histomorphometric analyses

Histological and histomorphometric results are shown in Figs 1 and 2, respectively. Increased TbSp (Fig 2A) and decreased TbWi (Fig 2B) and BAr (Fig 2C) were observed in the T1DM group compared to the control group (p < 0.001, p < 0.001, and p < 0.01, respectively). The T1DMS group showed reduced TbSp and increased TbWi and BAr compared to the T1DM group (Fig 1, p < 0.001, p < 0.001, and p < 0.05, respectively). In addition, T1DMS rats exhibited similar results when compared to control rats. Representative histological data are shown in Fig 1.

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Fig 1. Histological analyses of the right tibias.

Hematoxylin and eosin staining of longitudinal sections of tibias of the control (A), type 1 diabetes mellitus (T1DM) (B), and T1DM plus zinc supplementation (T1DMS) (C) groups. TB, trabecular bone; MS, medullary space; magnification 20X, scale bar: 50 μm.

https://doi.org/10.1371/journal.pone.0125349.g001

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Fig 2. Histomorphometric analyses of structural bone architecture.

Trabecular separation (TbSP, μm) (A), trabecular width (TbWi, μm) (B), and trabecular bone area (BAr, %) (C) of control, type 1 diabetes mellitus (T1DM), and T1DM plus zinc supplementation (T1DMS) rats. All data are shown as means ± SEM. Comparisons between groups were analyzed with Kruskal-Wallis ANOVA on Ranks and Dunn’s post-hoc. p < 0.01*/## vs. control group; p < 0.001*/### vs. control group; p < 0.05 **/# vs. T1DM group; p < 0.001**/### vs. T1DM group.

https://doi.org/10.1371/journal.pone.0125349.g002

Collagen content

The collagen analysis results are shown in Fig 3. We observed a decrease in total collagen in T1DM rats compared to control rats (Fig 3A, p < 0.05), whereas there was no difference between T1DMS rats and control rats. We also found a decrease in type I collagen in T1DM rats compared to controls (Fig 3B, p < 0.05), and T1DMS results were increased 1.2-fold compared to the T1DM group; however, this increase was not statistically significant. No differences were observed for type III collagen (Fig 3C).

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Fig 3. Assessment of collagen deposition by picrosirius red staining.

Tibia staining for collagen content (picrosirius red). Total collagen (A), collagen type I (B), and collagen type III (C) contents of the control, type 1 diabetes mellitus (T1DM), and T1DM plus zinc supplementation (T1DMS) groups. All data are shown as means ± SEM. Comparisons between groups were analyzed with Kruskal-Wallis ANOVA on Ranks and Dunn’s post-hoc. p < 0.05 */# vs. control group.

https://doi.org/10.1371/journal.pone.0125349.g003

Biomechanical testing data

Biomechanical parameters are shown in Table 2. Significantly decreased values for ultimate load, stiffness, ultimate strain, and Young’s modulus in the T1DM group was observed relative to control values (p < 0.01, p < 0.01, p < 0.05, and p < 0.05, respectively). Interestingly, the values of these parameters were higher (2-, 1.5-, 1.2-, and 1.2-fold, respectively) in the T1DMS group than in the T1DM group. In addition, we observed that ultimate load, ultimate strain, and Young’s modulus values in the T1DMS group were similar to the control values. The stiffness was uniquely decreased in T1DMS rats compared to controls (p < 0.05) but showed a 1.2-fold increase compared to T1DM rats. No significant difference was observed in the ultimate stress parameter.

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Table 2. Tibia biomechanical parameters of control, diabetic, and diabetic plus zinc supplementation groups.

https://doi.org/10.1371/journal.pone.0125349.t002

mRNA expression data

Molecular bone metabolic parameters are summarized in Fig 4. The mRNA expression levels of OPG, COL1A, and MMP-9 (Fig 4B, 4D, and 4F, respectively) were increased 37.8-, 17.3-, and 344.4-fold, respectively (p < 0.05, p < 0.01, and p < 0.01, respectively) in T1DM rats compared with control rats. Interestingly, the T1DMS group showed decreased expression of these genes (31.1-, 8.3-, and 313.9-fold, respectively) in comparison to T1DM rats, although this result was not statistically significant. In addition, no alterations were observed in these genes between the control and T1DMS groups (Fig 4B, 4D and 4F) except the OC mRNA expression, which showed a 17.9-fold increase (p < 0.05) compared with the control group (Fig 4C). No significant difference was found for MMP-2 mRNA expression between the groups.

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Fig 4. Relative mRNA expression quantification.

RANKL (A), OPG (B), OC (C), COL1A (D), MMP-2 (E), and MMP-9 (F) mRNA expression in bone tissue of control, type 1 diabetes mellitus (T1DM), and T1DM plus zinc supplementation (T1DMS) rats. All data are expressed as fold-change vs. control group values, normalized to GAPDH. Comparisons between groups were analyzed with Kruskal-Wallis ANOVA on Ranks and Dunn’s post-hoc. p < 0.05*/# vs. control group; p < 0.01*/## vs. control group.

https://doi.org/10.1371/journal.pone.0125349.g004

Discussion

Several studies have shown that bone turnover and skeletal integrity are affected by diabetes; however, the underlying mechanism of diabetes-induced bone loss remains elusive, as does the influence of disease stage in its development [3,4,17,34].

The STZ-induced diabetes model has been extensively used, making it particularly useful for building upon and comparing study results [2,10,23,29,37,39–42]. The benefits of the STZ-induced diabetic model include the ability to induce diabetes in a genetically altered animal, maintain the model in a controlled environment, regularly monitor and directly measure serum and bone factors, obtain bone samples for high-resolution analyses, and choose the time of diabetic induction (compared to waiting for diabetes to occur in spontaneous models) [2]. Indeed, STZ induction of diabetes causes a bone phenotype consistent with human studies [8,43–49] and with spontaneous mouse models such as NOD mice [50], confirming the utility of the STZ model for studying mechanisms of T1-diabetes-induced bone loss.

Several studies have used zinc supplementation to preserve bone structure and metabolism, as zinc is an essential nutrient for human and animal growth [22,25,26,51,52]. Zinc deficiency during adolescence may increase the risk of bone disease later in life due to reduced mineralization during the consolidation phase of bone mineral acquisition [53]. The protective effect of zinc on bone is suggested primarily by its stimulatory effect on cell proliferation, differentiation, and mineralization in osteoblasts, thereby promoting bone formation [21,22]. Additionally, zinc may stimulate the expression of various cellular proteins, including Runx2/Cbfa1 (Runt-related transcription factor 2/Core binding factor alpha 1), type I collagen, ALP, and OC. Zinc also increases cellular production of IGF-I and TGF-β1. Moreover, this ion also suppresses the osteoclast-like cell formation induced by various bone-resorbing factors in bone marrow culture (e.g., RANKL in pre-osteoclasts) and stimulates OPG gene expression in osteoblastic cells, which can inhibit the binding of RANKL to RANK in pre-osteoclastic cells [21].

Herein, we chose to evaluate the zinc protective effect for a period of 90 days of T1DM because chronic high glucose exposure may present an extreme condition for studying key aspects of bone alteration such as architecture, biomechanics, and gene regulation.

Although the beneficial influence of zinc is recognized in T1DM-related bone loss, few studies have evaluated the effect of zinc supplementation in chronic conditions [23,27,29,42,54,55]. The majority of studies have shown a zinc protective effect in short-term T1DM animal models, restoring calcium content, ALP activity, and DNA content after 14 days of T1DM onset [29,42]. Furthermore, zinc prevented diabetes-induced osteoclastogenesis and decreased osteoblastogenesis two weeks after diabetes onset, as evaluated by histomorphometric, biochemical, and molecular parameters [23].

Our results demonstrate significant bone loss associated with long-term T1DM and, because zinc supplementation was initiated following diabetes onset, suggest an important protective effect of zinc against excessive bone loss in chronic T1DM.

The protective effect of zinc was supported through histomorphometric parameters, which showed decreased TbSp and increased TbWi and BAr in T1DMS rats relative to T1DM rats. These results emphasize the importance of supplementation as an anabolic and protective agent against reduction in bone architecture. Similarly, in a T1DM-induced bone loss model over 14 days, Iitsuka et al. [23] showed recovery of BAr, TbWi, and the number of osteoclasts and osteoblasts in the supplemented group, suggesting that zinc restored diabetes-induced osteopenia by acting on both bone formation and resorption through regulation of osteoclast and osteoblast numbers. Furthermore, in normal rats, Ovesen et al. [24] found that alimentary zinc deficiency in growing rats reduced distal femoral metaphysis and femoral diaphysis during four weeks of study.

The protective effect of zinc supplementation was highlighted by altered trabecular structures (increased TbSp and diminished TbWi and BAr) in the T1DM group, evidencing negative effects of hyperglycemia on bone structure and leading to significant bone loss. These results corroborate a previous 120-day study in our laboratory [34] in which we observed a significant increase in TbSp, a decrease in TbWi, and a progressive ~77% reduction in BAr accompanied by a proportional expansion of the marrow space. Other investigators observed alterations in TbWi and TbSp two and eight weeks after the onset of experimental diabetes, respectively [39,56]. Moreover, our results are consistent with μ-CT analyses by Thrailkill et al. [3] and Martin and McCabe [57], even though these studies employed a short-term diabetes model.

In addition to histomorphometric analyses, the biomechanical integrity of bone is an important factor affecting the risk of fracture. Diabetes-induced structural abnormalities that predispose bone to fractures may occur spontaneously or with minimal trauma in patients [58].

In the present study, zinc supplementation prevented diabetes-induced alterations: the T1DMS group showed increased mineralization content (stiffness parameter) and bone strength (ultimate stress parameter), which may reflect the bone resistance to fracture, compared to the T1DM group, and values were similar between the T1DMS and control groups [59]. Furthermore, maintenance of the inorganic matrix may preserve the bone flexibility observed through increased ultimate strain and Young’s modulus in T1DMS rats compared to T1DM rats. Young’s modulus is a basic material property that is independent of geometry, represents the ability of bone to resist deformation, and is associated with ultimate strain in reflecting important parameters related to bone flexural conditions [59].

Only few studies have investigated the effect of zinc on bone biomechanical parameters, however in a non-diabetic rodent model [24,25]. To the best of our knowledge, the present study is the first to evaluate the effect of zinc supplementation on chronic T1DM-induced bone loss through bone biomechanical parameters.

The zinc protective effect was supported through reduced values for biomechanical parameters (stiffness, ultimate stress, ultimate strain, and Young’s modulus) in the T1DM group. These results suggest that the bone integrity changes observed under conditions of high glucose exposure may be attributed to interrelated factors such as macroscopic structure (size and shape), architecture (cortical and tissue), and bone substance (organic and inorganic components), all of which may influence mechanical strength.

Additionally, the reduction in biomechanical parameters in T1DM rats agrees with studies using similar tests at seven [11] and eight weeks after diabetes confirmation [10,12].

Interestingly, the increase in biomechanical parameters was associated with increased collagen content in the zinc supplementation group, further suggesting an important role for zinc in maintenance of the organic matrix during this long-term (90 days) study. However, for the T1DM group, the reduction in type I collagen content supports the low biomechanical properties found after 90 days of study.

Finally, we analyzed mRNA expression levels of key genes associated with bone metabolism. To the best of our knowledge, this is the first study to evaluate the effects of zinc supplementation in a chronic model of diabetes-induced bone loss (90 days) through the analysis of RANKL, OPG, OC, COL1A, MMP-2, and MMP-9 mRNA expression.

MMP-9 was downregulated in the T1DMS group relative to the T1DM group, indicating maintenance of bone type I collagen and correlating with the greater flexural strength observed in zinc-supplemented rats. On the other hand, the upregulation of MMP-9 and the low biomechanical and histomorphometric properties of the T1DM group suggest bone loss under the hyperglycemic condition. This hypothesis is supported by the association between MMP-9 and degradation of bone collagens in the subosteoclastic microenvironment and its potential role in normal bone remodeling and pathologic bone resorption [60].

MMP-9, a proteolytic member of the metalloproteinase family also named 92-kD type IV collagenase (gelatinase B), can degrade the components of the bone organic matrix and process both helical and denatured forms of type I collagen [13,60]. It is expressed at high levels in rabbit and human osteoclasts and in multinucleated cells of giant-cell tumors of bone [14]. Okada et al. [61] reported that MMP-9 has relatively broad substrate specificity, hydrolyzing collagen types I, III, IV, and V, and gelatins, and 50–80% of its full activity is retained at acidic pH. Grigoriadis et al. [62] showed that the expression of this enzyme is altered by bone-resorption activity.

Studies in non-diabetic osteoporotic bone demonstrated that osteoclasts can synthesize MMP-9 and exocytose it to degrade bone matrix components, especially type I collagen. Interestingly, some cells lining the surface of the bone matrix also express MMP-9, which is indirectly involved in bone resorption by removing the osteoid layer in this context, thereby exposing mineralized bone matrix and facilitating the adhesion of osteoclasts [63]. Although the hyperexpression of MMP-9 may contribute to collagen degradation, an important role for MMP-9 in healthy bone was reported by Nyman et al. [13] using MMP-9−/− mice, which showed changes in the trabecular architecture and cortical structure compared with wild-type mice, suggesting that bone quality is influenced by the MMPs expressed by osteoblasts and osteoclasts.

Thus, the reduced expression of MMP-9 associated with maintenance of histomorphometric and biomechanical parameters in the zinc-supplemented group suggests that zinc prevents bone resorption. In comparison, the high-level of MMP-9 expression in bone tissues of T1DM rats and reduced collagen content reinforce the association of MMPs with collagen in osteoporotic bone.

Furthermore, the increases in collagen density, trabecular spaces, Young’s modulus, and ultimate strain parameters and the reduced MMP-9 gene expression in the T1DMS group compared to the T1DM group and the similarity of these values to the control group suggest that zinc supplementation protects against organic matrix degradation and bone loss.

The expression of COL1A in bone tissues of T1DMS rats was lower than that in T1DM rats, suggesting a positive effect on zinc on maintenance of bone metabolism through bone architecture (histomorphometric parameters) and bone strength and flexibility (biomechanical parameters) similar to the control group. Several studies have previously observed similar results regarding the protective effect of zinc supplementation against collagen degradation in in vitro studies [28,64], clinical studies in patients [55,65], and animal models [23,51].

COL1A expression in bone tissues of T1DM rats was upregulated, possibly indicating an increase in bone loss associated with a chronic diabetic condition. These results indicate high bone turnover as an alternative mechanism to maintenance of bone homeostasis that was not necessary for the T1DMS group, providing further support for the protective effect of zinc on the bone metabolism. Similar to our T1DM results, the COL1A response was also observed in in vitro studies, suggesting that hyperglycemia can affect bone tissue by inducing excessive production of osteoid matrix [17]. This observation may also be explained by the increase in MMP-9 mRNA that possibly leads to degradation of the organic matrix in hyperglycemic conditions. Moreover, McCabe et al. [66] reported that the chronic exposure of osteoblasts to hyperglycemic cell culture media increased collagen production and altered the cellular phenotype toward that of osteocytes [67].

The discovery of RANK, RANKL, and OPG, factors involved in the control of osteoclast differentiation and osteoporosis, has advanced bone research into a new era. The RANK/RANKL/OPG system is an important signal transduction pathway that regulates bone resorption, modeling, and remodeling. The binding of OPG to RANKL inhibits binding between RANKL and RANK, thereby preventing osteoclast precursor differentiation and fusion to form mature osteoclasts. Thus, the relative concentrations of RANKL and OPG in bone are a major determinant of bone mass and strength.

OPG expression was downregulated in the T1DMS group, suggesting that bone homeostasis is maintained by zinc supplementation even in chronic hyperglycemic conditions. Moreover, the zinc effect on OPG mRNA expression is controversial. Iitsuka et al. [23] showed unaltered OPG mRNA expression during zinc supplementation for 14 days in a T1DM-induced bone loss animal model, and Fong et al. [68] reported similar OPG mRNA expression between postnatal and control groups. However, a review by Yamaguchi [22] showed that zinc plays an important role in bone growth by stimulating the expression of OPG mRNA in osteoblastic cells.

We observed upregulation of OPG in a chronic hyperglycemic condition, suggesting an attempted protective response against excessive bone loss induced by diabetic conditions. When OPG expression is increased relative to RANKL expression, the latter is expected to become unavailable to bind RANK in pre-osteoclasts, resulting in reduced bone resorption [17]. OPG upregulation in T1DM was also reported in T1DM patients in a previous study in our laboratory, indicating that diabetes during pubertal growth, which is associated with proinflammatory processes, may cause deficient bone-mass gain [8]. Moreover, an in vitro study has demonstrated excessive OPG synthesis at different glucose concentrations [17]. Thus, the upregulation of OPG expression in T1DM rats suggests that this increase protects the bone against resorption by inhibiting osteoclast differentiation mediated by RANK-RANKL binding. Although the present results show that the upregulation of OPG in T1DM rats did not result in detectable protection against trabecular structures, the loss of bone mass may have been more pronounced without this increased expression.

These results suggest an important role for zinc on bone protection in chronic T1DM and are supported by maintenance of bone architecture (histomorphometric and collagen content) and biomechanical proprieties and downregulation of genes involved in organic matrix degradation (MMP-9 and COL1A).

Zinc supplementation also showed an anabolic effect, as evidenced by OC mRNA upregulation and increased serum ALP activity after the 90-day experimental period. OC mRNA is primarily expressed by post-proliferating and terminally mature osteoblasts [69–72] and regulates mineralization of the extracellular matrix [73–75]. ALP is an important serum marker associated with osteoblast activity and bone formation and is necessary for bone mineralization and development of collagenous structures [76]. Thus, the upregulation of OC and increased serum ALP activity are consistent with the hypothesis that zinc regulates osteoblastogenesis, suggesting a possible induction of bone formation and mineralization [22]. Furthermore, these results are consistent with Young’s modulus values showing a high resistance to fracture, supporting the suggestion that zinc supplementation may protect the bone architecture in both mineral and non-mineral content, leading to greater biomechanical strength.

In conclusion, zinc supplementation prevented bone loss in chronic T1DM rats as demonstrated by the maintenance of bone homeostasis and bone architecture, strength, and flexibility. In addition, zinc-induced OC upregulation and RANKL, OPG, COL1A, and MMP-9 downregulation, even in chronic hyperglycemia, support a protective role for zinc in long-term diabetic conditions through stimulating expression of the mineralizing phenotype in osteoblasts and reducing expression of the resorptive phenotype in osteoclasts. Moreover, the protective zinc effect is supported after chronic hyperglycemia in T1DM leads to bone loss, as evidenced by alterations in bone structures associated with poor bone quality.

Thus, these results suggest the therapeutic potential of zinc supplementation as a complementary therapy to prevent bone loss in patients with diabetes or other related chronic diseases, resulting in better bone protection during growth as well as in adult and advanced ages.

Acknowledgments

We acknowledge UNAERP for supporting this experimental study, including animal maintenance and biochemical analyses. We are also grateful to the students and technicians from LABMULT/UFRN/RN.

Author Contributions

Conceived and designed the experiments: AAR RHB LAR. Performed the experiments: RHB FSS DESM CESM AASB GO JFB MAGU MBL KSCS. Analyzed the data: AAR RHB LAR BJGAA VMGDML MGA FSS ADL. Contributed reagents/materials/analysis tools: AAR LAR BJGAA MVLF CESM. Wrote the paper: AAR RHB LAR BJGAA MAGU.

References

  1. 1. American Diabetes Association. Standards of medical care in diabetes—2014. Diabetes Care. 2014;37 Suppl 1: S14–S80. pmid:24357209
  2. 2. Motyl K, McCabe LR. Streptozotocin, type I diabetes severity and bone. Biol Proced Online. 2009;11: 296–315. pmid:19495918
  3. 3. Thrailkill KM, Liu L, Wahl EC, Bunn RC, Perrien DS, Cockrell GE, et al. Bone formation is impaired in a model of type 1 diabetes. Diabetes. 2005;54: 2875–2881. pmid:16186388
  4. 4. Starup-Linde J. Diabetes, biochemical markers of bone turnover, diabetes control, and bone. Front Endocrinol (Lausanne). 2013;4:21. pmid:23482417
  5. 5. Pietschmann P, Skalicky M, Kneissel M, Rauner M, Hofbauer G, Stupphann D, et al. Bone structure and metabolism in a rodent model of male senile osteoporosis. Exp Gerontol. 2007;42: 1099–1108. pmid:17949933
  6. 6. Hofbauer LC, Brueck CC, Singh SK, Dobnig H. Osteoporosis in patients with diabetes mellitus. J Bone Miner Res. 2007;22: 1317–1328. pmid:17501667
  7. 7. Prisby RD, Swift JM, Bloomfield SA, Hogan HA, Delp MD. Altered bone mass, geometry and mechanical properties during the development and progression of type 2 diabetes in the Zucker diabetic fatty rat. J Endocrinol. 2008;199: 379–388. pmid:18755885
  8. 8. Loureiro MB, Ururahy MA, Freire-Neto FP, Duarte VM, Oliveira GH, Hirata RDC, et al. Low bone mineral density is associated to poor glycemic control and increased OPG expression in children and adolescents with type 1 diabetes. Diabetes Res Clin Pract. 2014;103: 452–457. pmid:24529565
  9. 9. McCabe LR. Understanding the pathology and mechanisms of type I diabetic bone loss. J Cell Biochem. 2007;102: 1343–1357. pmid:17975793
  10. 10. Erdal N, Gürgül S, Demirel C, Yildiz A. The effect of insulin therapy on biomechanical deterioration of bone in streptozotocin (STZ)-induced type 1 diabetes mellitus in rats. Diabetes Res Clin Pract. 2012;97: 461–467. pmid:22483749
  11. 11. Reddy GK, Stehno-Bittel L, Hamade S, Enwemeka CS. The biomechanical integrity of bone in experimental diabetes. Diabetes Res Clin Pract. 2001;54: 1–8. pmid:11532324
  12. 12. Korres N, Tsiridis E, Pavlou G, Mitsoudis A, Perrea DN, Zoumbos AB, et al. Biomechanical characteristics of bone in streptozotocin-induced diabetic rats: An in-vivo randomized controlled experimental study. World J Orthop. 2013;4: 124–129. pmid:23878780
  13. 13. Nyman JS, Lynch CC, Perrien DS, Thiolloy S, O’Quinn EC, Patil CA, et al. Differential effects between the loss of MMP-2 and MMP-9 on structural and tissue-level properties of bone. J Bone Miner Res. 2011;26: 1252–1260. pmid:21611966
  14. 14. Mohtai M, Smith RL, Schurman DJ, Tsuji Y, Torti FM, Hutchinson NI, et al. Expression of 92-kD type IV collagenase/gelatinase (gelatinase B) in osteoarthritic cartilage and its induction in normal human articular cartilage by interleukin 1. J Clin Invest. 1993;92: 179–185. pmid:8325982
  15. 15. Vandooren J, Van den Steen PE, Opdenakker G. Biochemistry and molecular biology of gelatinase B or matrix metalloproteinase-9 (MMP-9): the next decade. Crit Rev Biochem Mol Biol. 2013;48: 222–272. pmid:23547785
  16. 16. Zhang SQ, Chen GH, Lu WL, Zhang Q. Effects on the bones of vanadyl acetylacetonate by oral administration: a comparison study in diabetic rats. J Bone Miner Metab. 2007;25: 293–301. pmid:17704994
  17. 17. Cunha JS, Ferreira VM, Maquigussa E, Naves MA, Boim MA. Effects of high glucose and high insulin concentrations on osteoblast function in vitro. Cell Tissue Res. 2014;358: 249–256. pmid:24859221
  18. 18. Wittrant Y, Gorin Y, Woodruff K, Horn D, Abboud HE, Mohan S, et al. High d(+)glucose concentration inhibits RANKL-induced osteoclastogenesis. Bone. 2008;42: 1122–1130. pmid:18378205
  19. 19. Motyl KJ, Botolin S, Irwin R, Appledorn DM, Kadakia T, Amalfitano A, et al. Bone inflammation and altered gene expression with type I diabetes early onset. J Cell Physiol. 2009;218: 575–583. pmid:19006181
  20. 20. Amorim FP, Ornelas SS, Diniz SF, Batista AC, da Silva TA. Imbalance of RANK, RANKL and OPG expression during tibial fracture repair in diabetic rats. J Mol Histol. 2008;39: 401–408. pmid:18592139
  21. 21. Yamaguchi M. Role of nutritional zinc in the prevention of osteoporosis. Mol Cell Biochem. 2010;338: 241–254. pmid:20035439
  22. 22. Yamaguchi M. Nutritional factors and bone homeostasis: synergistic effect with zinc and genistein in osteogenesis. Mol Cell Biochem. 2012;366: 201–221. pmid:22476903
  23. 23. Iitsuka N, Hie M, Tsukamoto I. Zinc supplementation inhibits the increase in osteoclastogenesis and decrease in osteoblastogenesis in streptozotocin-induced diabetic rats. Eur J Pharmacol. 2013;714: 41–47. pmid:23735664
  24. 24. Ovesen J, Møller-Madsen B, Thomsen JS, Danscher G, Mosekilde L. The positive effects of zinc on skeletal strength in growing rats. Bone. 2001;29: 565–570. pmid:11728928
  25. 25. Brzóska MM, Galazyn-Sidorczuk M, Rogalska J, Roszczenko A, Jurczuk M, Majewska K, et al. Beneficial effect of zinc supplementation on biomechanical properties of femoral distal end and femoral diaphysis of male rats chronically exposed to cadmium. Chem Biol Interact. 2008;171: 312–324. pmid:18164699
  26. 26. Sunar F, Baltaci AK, Ergene N, Mogulkoc R. Zinc deficiency and supplementation in ovariectomized rats: their effect on serum estrogen and progesterone levels and their relation to calcium and phosphorus. Pak J Pharm Sci. 2009;22: 150–154. pmid:19339224
  27. 27. Abo-El-Asrar M, Farid SM, El Maraghy MO, Mohamedeen AK. Serum osteocalcin, zinc nutritive status and bone turnover in children and adolescents with type1 diabetes mellitus. J Diabetes Metab. 2011;2: 128.
  28. 28. Yamaguchi M, Goto M, Uchiyama S, Nakagawa T. Effect of zinc on gene expression in osteoblastic MC3T3-E1 cells: enhancement of Runx2, OPG, and regucalcin mRNA expressions. Mol Cell Biochem. 2008;312: 157–166. pmid:18327666
  29. 29. Yamaguchi M, Uchiyama S. Preventive effect of zinc acexamate administration in streptozotocin-diabetic rats: restoration of bone loss. Int J Mol Med. 2003;12: 755–761. pmid:14533005
  30. 30. National Research Council (US) Committee for the Update of the Guide for the Care and Use of Laboratory Animals. Guide for the care and use of laboratory animals. 2011.
  31. 31. Melo MGD, Dória GAA, Serafini MR, Araújo AAS. Valores de referência Hematológicos e Bioquímicos de Ratos (Rattus novergicus linhagem Wistar) provenientes do biotério central da Universidade Federal de Sergipe. Scientia Plena. 2012;8: 1–6.
  32. 32. Dantas JA, Ambiel CR, Cuman RKN, Baroni S, Bersani-Amado CA. Valores de referência de alguns parâmetros fisiológicos de ratos do Biotério Central da Universidade Estadual de Maringá, Estado do Paraná. Acta Sci Heal Sci. 2006;28: 165–170.
  33. 33. Reeves PG, Nielsen FH, Fahey GC Jr. AIN-93 purified diets for laboratory rodents: final report of the American Institute of Nutrition ad hoc writing committee on the reformulation of the AIN-76A rodent diet. J Nutr. 1993;123: 1939–1951. pmid:8229312
  34. 34. Duarte VM, Ramos AM, Rezende LA, Macedo UB, Brandão-Neto J, Almeida MG, et al. Osteopenia: a bone disorder associated with diabetes mellitus. J Bone Miner Metab. 2005;23: 58–68. pmid:15616896
  35. 35. Dempster DW, Compston JE, Drezner MK, Glorieux FH, Kanis JA, Malluche H, et al. Standardized nomenclature, symbols, and units for bone histomorphometry: a 2012 update of the report of the ASBMR Histomorphometry Nomenclature Committee. J Bone Miner Res. 2013;28: 2–17. pmid:23197339
  36. 36. Rich L, Whittaker P. Collagen and picrosirius red staining: a polarized light assessment of fibrillar hue and spatial distribution. Brazilian J Morphol Sci. 2005;22: 97–104.
  37. 37. Black MJ, D’Amore A, Auden A, Stamp L, Osicka T, Panagiotopoulos S, et al. Chronic type 1 diabetes in spontaneously hypertensive rats leads to exacerbated cardiac fibrosis. Cardiovasc Pathol. 2010;19: 361–370. pmid:19747849
  38. 38. Livak KJ, Schmittgen TD. Analysis of relative gene expression data using real-time quantitative PCR and the 2(-Delta Delta C(T)) Method. Methods. 2001;25: 402–408. pmid:11846609
  39. 39. Goodman WG, Hori MT. Diminished bone formation in experimental diabetes. Relationship to osteoid maturation and mineralization. Diabetes. 1984;33: 825–831. pmid:6381178
  40. 40. Hamada Y, Kitazawa S, Kitazawa R, Fujii H, Kasuga M, Fukagawa M, et al. Histomorphometric analysis of diabetic osteopenia in streptozotocin-induced diabetic mice: a possible role of oxidative stress. Bone. 2007;40: 1408–1414. pmid:17251074
  41. 41. Hie M, Iitsuka N, Otsuka T, Tsukamoto I. Insulin-dependent diabetes mellitus decreases osteoblastogenesis associated with the inhibition of Wnt signaling through increased expression of Sost and Dkk1 and inhibition of Akt activation. Int J Mol Med. 2011;28: 455–462. pmid:21567076
  42. 42. Uchiyama S, Yamaguchi M. Alteration in serum and bone component findings induced in streptozotocin-diabetic rats is restored by zinc acexamate. Int J Mol Med. 2003;12: 949–954. pmid:14612972
  43. 43. Bouillon R, Bex M, Van Herck E, Laureys J, Dooms L, Lesaffre E, et al. Influence of age, sex, and insulin on osteoblast function: osteoblast dysfunction in diabetes mellitus. J Clin Endocrinol Metab. 1995;80: 1194–1202. pmid:7714089
  44. 44. Kemink SA, Hermus AR, Swinkels LM, Lutterman JA, Smals AG. Osteopenia in insulin-dependent diabetes mellitus; prevalence and aspects of pathophysiology. J Endocrinol Invest. 2000;23: 295–303. pmid:10882147
  45. 45. Auwerx J, Dequeker J, Bouillon R, Geusens P, Nijs J. Mineral metabolism and bone mass at peripheral and axial skeleton in diabetes mellitus. Diabetes. 1988;37: 8–12. pmid:3335279
  46. 46. Levin ME, Boisseau VC, Avioli LV. Effects of diabetes mellitus on bone mass in juvenile and adult-onset diabetes. N Engl J Med. 1976;294: 241–245. pmid:1244549
  47. 47. Tuominen JT, Impivaara O, Puukka P, Rönnemaa T. Bone mineral density in patients with type 1 and type 2 diabetes. Diabetes Care. 1999;22: 1196–1200. pmid:10388989
  48. 48. Muñoz-Torres M, Jódar E, Escobar-Jiménez F, López-Ibarra PJ, Luna JD. Bone mineral density measured by dual X-ray absorptiometry in Spanish patients with insulin-dependent diabetes mellitus. Calcif Tissue Int. 1996;58: 316–319. pmid:8661964
  49. 49. Buysschaert M, Cauwe F, Jamart J, Brichant C, De Coster P, Magnan A, et al. Proximal femur density in type 1 and 2 diabetic patients. Diabete Metab. 1992;18: 32–37. pmid:1563534
  50. 50. Botolin S, McCabe LR. Bone loss and increased bone adiposity in spontaneous and pharmacologically induced diabetic mice. Endocrinology. 2007;148: 198–205. pmid:17053023
  51. 51. Hadley KB, Newman SM, Hunt JR. Dietary zinc reduces osteoclast resorption activities and increases markers of osteoblast differentiation, matrix maturation, and mineralization in the long bones of growing rats. J Nutr Biochem. 2010;21: 297–303. pmid:19369052
  52. 52. Kwun IS, Cho YE, Lomeda RA, Shin HI, Choi JY, Kang YH, et al. Zinc deficiency suppresses matrix mineralization and retards osteogenesis transiently with catch-up possibly through Runx 2 modulation. Bone. 2010;46: 732–741. pmid:19913120
  53. 53. Ryz NR, Weiler HA, Taylor CG. Zinc deficiency reduces bone mineral density in the spine of young adult rats: a pilot study. Ann Nutr Metab. 2009;54: 218–226. pmid:19506366
  54. 54. Arreola F, Paniagua R, Díaz-Bensussen S, Urquieta B, López-Montaño E, Partida-Hernández G, et al. Bone mineral content, 25-hydroxycalciferol and zinc serum levels in insulin-dependent (type I) diabetic patients. Arch Invest Med (Mex). 1990;21: 195–199. pmid:2103709
  55. 55. Maser RE, Stabley JN, Lenhard MJ, Owusu-Griffin P, Provost-Craig MA, Farquhar WB, et al. Zinc intake and biochemical markers of bone turnover in type 1 diabetes. Diabetes Care. 2008;31: 2279–2280. pmid:18809628
  56. 56. Tsuchida T, Sato K, Miyakoshi N, Abe T, Kudo T, Tamura Y, et al. Histomorphometric evaluation of the recovering effect of human parathyroid hormone (1–34) on bone structure and turnover in streptozotocin-induced diabetic rats. Calcif Tissue Int. 2000;66: 229–233. pmid:10666500
  57. 57. Martin LM, McCabe LR. Type I diabetic bone phenotype is location but not gender dependent. Histochem Cell Biol. 2007;128: 125–133. pmid:17609971
  58. 58. Seino Y, Ishida H. Diabetic osteopenia: pathophysiology and clinical aspects. Diabetes Metab Rev. 1995;11: 21–35. pmid:7600905
  59. 59. Turner CH. Bone strength: current concepts. Ann N Y Acad Sci. 2006;1068: 429–446. pmid:16831941
  60. 60. Zhao H, Xia Z, Cai G, Du J, Zhu T, Shen L. Expression of type-I collagen and matrix metalloproteinase-9 mRNA in bone of castrated adult female rats: effects of estrogen. Chin Med J (Engl). 1998;111: 551–555. pmid:11245078
  61. 61. Okada Y, Naka K, Kawamura K, Matsumoto T, Nakanishi I, Fujimoto N, et al. Localization of matrix metalloproteinase 9 (92-kilodalton gelatinase/type IV collagenase = gelatinase B) in osteoclasts: implications for bone resorption. Lab Invest. 1995;72: 311–322. pmid:7898050
  62. 62. Grigoriadis AE, Wang ZQ, Cecchini MG, Hofstetter W, Felix R, Fleisch HA et al. c-Fos: a key regulator of osteoclast-macrophage lineage determination and bone remodeling. Science. 1994;266: 443–448. pmid:7939685
  63. 63. Lorenzo JA, Pilbeam CC, Kalinowski JF, Hibbs MS. Production of both 92- and 72-kDa gelatinases by bone cells. Matrix. 1992;12: 282–290. pmid:1435512
  64. 64. Uchiyama S, Yamaguchi M. Anabolic effect of beta-cryptoxanthin in osteoblastic MC3T3-E1 cells is enhanced with 17beta-estradiol, genistein, or zinc sulfate in vitro: the unique effect with zinc on Runx2 and alpha1(I) collagen mRNA expressions. Mol Cell Biochem. 2008;307: 209–219. pmid:17899321
  65. 65. Gur A, Colpan L, Cevik R, Nas K, Jale Sarac A. Comparison of zinc excretion and biochemical markers of bone remodelling in the assessment of the effects of alendronate and calcitonin on bone in postmenopausal osteoporosis. Clin Biochem. 2005;38: 66–72. pmid:15607319
  66. 66. Botolin S, McCabe LR. Chronic hyperglycemia modulates osteoblast gene expression through osmotic and non-osmotic pathways. J Cell Biochem. 2006;99: 411–424. pmid:16619259
  67. 67. Mizuno A, Amizuka N, Irie K, Murakami A, Fujise N, Kanno T, et al. Severe osteoporosis in mice lacking osteoclastogenesis inhibitory factor/osteoprotegerin. Biochem Biophys Res Commun. 1998;247: 610–615. pmid:9647741
  68. 68. Fong L, Tan K, Tran C, Cool J, Scherer MA, Elovaris R, et al. Interaction of dietary zinc and intracellular binding protein metallothionein in postnatal bone growth. Bone. 2009;44: 1151–1162. pmid:19254786
  69. 69. Owen TA, Aronow M, Shalhoub V, Barone LM, Wilming L, Tassinari MS, et al. Progressive development of the rat osteoblast phenotype in vitro: reciprocal relationships in expression of genes associated with osteoblast proliferation and differentiation during formation of the bone extracellular matrix. J Cell Physiol. 1990;143: 420–430. pmid:1694181
  70. 70. Stein GS, Lian JB, Owen TA. Relationship of cell growth to the regulation of tissue-specific gene expression during osteoblast differentiation. FASEB J. 1990;4: 3111–3123. pmid:2210157
  71. 71. Weinreb M, Shinar D, Rodan GA. Different pattern of alkaline phosphatase, osteopontin, and osteocalcin expression in developing rat bone visualized by in situ hybridization. J Bone Miner Res. 1990;5: 831–842. pmid:2239367
  72. 72. McCabe LR, Kockx M, Lian J, Stein J, Stein G. Selective expression of fos- and jun-related genes during osteoblast proliferation and differentiation. Exp Cell Res. 1995;218: 255–262. pmid:7737363
  73. 73. Ducy P, Desbois C, Boyce B, Pinero G, Story B, Dunstan C, et al. Increased bone formation in osteocalcin-deficient mice. Nature. 1996;382: 448–452. pmid:8684484
  74. 74. Fedde KN, Blair L, Silverstein J, Coburn SP, Ryan LM, Weinstein RS, et al. Alkaline phosphatase knock-out mice recapitulate the metabolic and skeletal defects of infantile hypophosphatasia. J Bone Miner Res. 1999;14: 2015–2026. pmid:10620060
  75. 75. Anderson HC, Sipe JB, Hessle L, Dhanyamraju R, Atti E, Camacho NP, et al. Impaired calcification around matrix vesicles of growth plate and bone in alkaline phosphatase-deficient mice. Am J Pathol. 2004;164: 841–847. pmid:14982838
  76. 76. Beattie JH, Avenell A. Trace element nutrition and bone metabolism. Nutr Res Rev. 1992;5: 167–188. pmid:19094319