Normoglycemia partially recovers the disrupted osteoblast differentiation of mesenchymal stem cells induced by type 1 but not type 2 diabetes mellitus

Denise Weffort; Leticia F Adolpho; Alann T P Souza; Gileade P Freitas; Helena B Lopes; Fabiola S Oliveira; Rayana L Bighetti‐Trevisan; Leticia Pitol‐Palin; Doris H Matsushita; Roberta Okamoto; Marcio M Beloti; Adalberto L Rosa

doi:10.1002/jcb.30434

. 2023 Jun 9;124(7):1050–1063. doi: 10.1002/jcb.30434

Normoglycemia partially recovers the disrupted osteoblast differentiation of mesenchymal stem cells induced by type 1 but not type 2 diabetes mellitus

Denise Weffort ¹, Leticia F Adolpho ¹, Alann T P Souza ¹, Gileade P Freitas ^1,², Helena B Lopes ¹, Fabiola S Oliveira ¹, Rayana L Bighetti‐Trevisan ¹, Leticia Pitol‐Palin ³, Doris H Matsushita ³, Roberta Okamoto ³, Marcio M Beloti ¹, Adalberto L Rosa ^1,^✉

PMCID: PMC13484591 PMID: 37293736

Abstract

Type 1 (T1DM) and type 2 (T2DM) diabetes mellitus are characterized by changes in glucose metabolism and cause bone damage via a variety of mechanisms, including effects on osteoblasts. We aimed to evaluate the osteoblast differentiation of mesenchymal stem cells (MSCs) from rats with T1DM or T2DM and the effects of removing the hyperglycemic stimulus on the osteogenic potential of these cells. MSCs from healthy rats were cultured in normoglycemic medium, whereas MSCs from rats with T1DM or T2DM were cultured in hyperglycemic or normoglycemic medium. T1DM and T2DM reduced osteoblast differentiation of MSCs grown in hyperglycemic media, with T1DM having a more pronounced effect, as evidenced by alkaline phosphatase activity, RUNX2 protein expression, and extracellular matrix mineralization, and modulated the gene expression of several components of the bone morphogenetic protein signaling pathway. The restoration of the normoglycemic environment partially recovers the osteogenic potential of MSCs from rats with T1DM but not with T2DM. Our findings highlight the need for specific therapies to treat T1DM‐ or T2DM‐induced bone loss, as both disrupt osteoblast differentiation at distinct levels and likely through different mechanisms.

Keywords: bone, diabetes mellitus, hyperglycemia, mesenchymal stem cells, osteoblasts

1. INTRODUCTION

Diabetes mellitus (DM) is a group of chronic metabolic diseases characterized by hyperglycemia caused by a disruption in insulin secretion, action, or both. The majority of cases are classified as type 1 (T1DM) or type 2 (T2DM). ¹ T1DM is insulin‐dependent because the beta cells are unable to produce insulin due to an autoimmune reaction, whereas T2DM is noninsulin‐dependent and is caused primarily by insulin resistance, accounting for approximately 90% of all DM cases. ² DM has a negative impact on several tissues and organs, including bone tissue, which can be impacted through different mechanisms depending on the disease etiopathology. ¹ , ³

Patients with either T1DM or T2DM exhibit a higher risk of fragility bone fractures; however, although individuals with T1DM clearly exhibit lower bone mineral density (BMD) than nondiabetic age‐matched population, T2DM can either increase, decrease, or have no effect on BMD, as evidenced by discrepancies in literature data. ³ , ⁴ , ⁵ , ⁶ , ⁷ , ⁸ Thus, bone fragility is related not only to decreased BMD but also to bone microarchitecture and composition, which are directly related to bone cell functions, which are regulated by T1DM and T2DM. ⁵ , ⁹

The mechanisms by which DM causes osteoblast dysfunction are not fully understood, but multiple factors are thought to be involved. It has been shown that T1DM reduces osteoblastogenesis, osteoblast proliferation, and the expression of osteoblast protein markers such as RUNX2 and osteocalcin (OC) by inhibiting Wnt signaling and b‐catenin activation in diabetic murine models. ¹⁰ , ¹¹ The osteogenic potential detected by the expression of bone‐related genes, alkaline phosphatase (ALP) activity, and extracellular matrix mineralization is reduced in osteoblasts derived from mice with T1DM compared with nondiabetic mice because of the increased oxidative stress. ¹² Moreover, T1DM‐induced cilia loss in osteoblasts results in impaired bone fracture repair in mice. ¹³

T2DM impairs bone healing in a mouse model by inhibiting osteoblast differentiation of skeletal stem cells and increasing the senescent cell population. ¹⁴ Additionally, the local inflammatory process lasts longer in rats with T2DM, disrupting collagen fiber organization and maturation and delaying alveolar bone repair. ¹⁵ Serum from patients with T2DM regulated the expression of osteoblast‐related genes in mesenchymal stem cells (MSCs) derived from human bone marrow, resulting in osteoblast inhibition and increased adipocyte differentiation. ¹⁴ , ¹⁶ T2DM inhibits osteoblast differentiation by accumulating advanced glycation end products (AGEs). ¹⁷ , ¹⁸ Indeed, cells from individuals with T2DM were more sensitivity to AGE receptor activation and exhibited a higher apoptosis rate, and MSCs from diabetic rats exhibited higher levels of AGEs and apoptosis than cells from nondiabetic rats, which could explain the disruption of osteoblast differentiation. ¹⁷ , ¹⁸

Given the differences in etiopathology between T1DM and T2DM and the fact that hyperglycemia, not just insulin deregulation, disrupts bone healing, ¹⁴ we hypothesized that T1DM and T2DM induce distinct levels of inhibition of osteoblast differentiation of MSCs, which is recovered when the hyperglycemic stimulus is removed. Thus, this study aims to evaluate the osteoblast differentiation of MSCs derived from rats with T1DM or T2DM and the effect of a normoglycemic environment on the osteogenic potential of these MSCs.

2. MATERIALS AND METHODS

2.1. Induction of T1DM and T2DM

Eighteen male Wistar rats aged 10–13 weeks were used with the approval of the Ethics Committee on the Use of Animals of the Araçatuba Dental School, São Paulo State University Júlio de Mesquita Filho (Protocol #2019‐0152). The rats were divided into three groups (n = 6 per group): healthy rats (control), rats with T1DM, and rats with T2DM. Rats were fasted for 12 h before receiving a single intravenous injection of streptozotocin (Sigma‐Aldrich) 60 mg/kg diluted in 0.2 mL of citrate buffer 0.01 M, pH = 4.5 (vehicle). For T2DM induction, rats at 10 weeks of age were fed a high‐fat diet for the duration of the experiment and then received a single intravenous injection of 35‐mg/kg streptozotocin (Sigma‐Aldrich) diluted in a vehicle 4 weeks later. Rats aged 14 weeks received a single intravenous injection of the 0.2‐mL vehicle as a control. Blood glucose levels were measured 1 week after streptozotocin or vehicle administration using the Accu‐Check Active® system (Roche), according to the manufacturer's instructions. T1DM was defined as blood glucose levels greater than 300 mg/dL and T2DM as levels greater than 198 mg/dL. Animals were euthanized 4 weeks after streptozotocin or vehicle administration, and femurs were harvested for morphometry analysis or MSCs isolation. One rat in the T2DM group died 5 days before the established time for euthanasia probably because of diabetic condition, and the morphometry analysis was performed on five animals in this group.

Microtomographic and morphometric analyses of the selected region of the femurs that began at the metaphysis, 0.5 mm from the end of the growth plate, and continued for 2 mm toward the diaphysis were used to evaluate the effect of T1DM and T2DM on bone tissue. The femurs harvested from control rats (n = 6) and rats with T1DM (n = 6) or T2DM (n = 5) were fixed in buffered formalin 10% and analyzed using a microtomography scanner SkyScan 1172 (Bruker‐SkyScan) operating with a 100‐kV X‐ray source detected by an 11‐megapixel camera with a resolution of up to 1 μm. The reconstructions were generated using the NRecon Cluster software (Micro Photonics), and the morphometric parameters bone volume (BV, mm³), percentage of bone volume (BV/TV, %), bone surface (BS, mm²), trabecular number (Tb.N, 1/mm), trabecular separation (Tb.Sp, mm), and bone mineral density (BMD, g/cm³) were evaluated as recommended by the American Society of Bone and Mineral Research. ¹⁹

2.2. MSC harvesting, culture, and osteoblast differentiation

MSCs were isolated from femoral bone marrow and grown in a noninducing culture medium composed of minimum essential medium, modification alpha (α‐MEM, Gibco‐Life Technologies), 10% fetal bovine serum (Gibco‐Life Technologies), 50‐μg/mL gentamicin (Gibco‐Life Technologies), and 0.3‐μg/mL fungizone (Gibco‐Life Technologies). MSCs from control rats and a subset of MSCs from rats with T1DM or T2DM were cultured in normoglycemic (100 mg/dL of glucose or 5 mM) medium (NG). To mimic the in vivo hyperglycemic environment, MSCs from rats with T1DM or T2DM were cultured under hyperglycemic conditions with high glucose concentrations of 600 mg/dL (33 mM, HG1) for MSCs from rats with T1DM (T1DM‐HG1) or 290 mg/dL (16 mM, HG2) for MSCs from rats with T2DM (T2DM‐HG2). To induce osteoblast differentiation, at subconfluence, MSCs were seeded into 24‐well culture plates at a cell density of 2 × 10⁴ cells/well and cultured in osteogenic medium, which is noninducing medium supplemented with 5‐µg/mL ascorbic acid (Gibco‐Life Technologies), 7 mM beta‐glycerophosphate (Sigma‐Aldrich), and dexamethasone 10⁻⁷ M (Sigma‐Aldrich), with the appropriate concentration of glucose for up to 21 days. Thus, the experimental design is based on five groups of MSCs in terms of animal condition and glucose concentration: (a) control—MSCs from healthy rats cultured in NG medium (5 mM glucose), (b) T1DM‐NG—MSCs from rats with T1DM cultured in NG medium, (c) T1DM‐HG1—MSCs from rats with T1DM cultured in HG1 medium (33 mM glucose), (d) T2DM‐NG—MSCs from rats with T2DM cultured in NG medium, and (e) T2DM‐HG2—MSCs from rats with T2DM cultured in HG2 medium (16 mM glucose). These in vitro glucose concentrations were based on the blood glucose levels of the control rats and rats with T1DM or T2DM, and all culture media presented the same osmolality (20 mOsm/kg H₂O) irrespective of the glucose concentration. The cell cultures were kept at 37°C in a humidified atmosphere of 5% CO₂ and 95% air, with the culture media changed every 48 h.

2.3. Analysis of the gene expression of bone markers by real‐time polymerase chain reaction (RT‐qPCR)

On Day 7, RT‐qPCR was used to evaluate the gene expression of Runx2, Osterix, Alp, Oc, and bone sialoprotein (Bsp). The Trizol reagent (Invitrogen) was added to the samples, and total RNA was extracted using the SV Total RNA Isolation System kit (Promega) according to the manufacturer's instructions. The total RNA was quantified using the NanoVue device (GE Healthcare), and a reverse transcription reaction was carried out to synthesize complementary DNA (cDNA) using the High‐Capacity cDNA Reverse Transcription Kit (Thermo Fisher Scientific). The SYBR Green system and primer sequences (Supporting Information: Table S1) were used in a QuantStudio™ 7 Flex System device for RT‐qPCR (Applied Biosystems). The reactions (n = 4) were performed, and gene expression was normalized to the endogenous control Eif2b1 using the $- 2^{∆ ∆ C_{t}}$ method. ²⁰

2.4. Analysis of the cell viability by colorimetric assay

The cell viability was evaluated using a colorimetric assay with 3‐(4,5‐dimethylthiazol‐2‐yl)−2,5‐diphenyltetrazolium bromide (MTT) on days 3, 7, and 10. Briefly, 1 mL of culture medium containing 10% MTT (5 mg/mL) was added in each well, and the cultures were incubated at 37°C for 4 h. The supernatant was then removed from each well, 1 mL of acid isopropanol was added in each well, the plates were shaken for 5 min, and 150 µL of each well was transferred to a 96‐well plate. The absorbance was measured at 570 nm using a μQuant spectrophotometer (BioTek Instruments Inc), and the data (n = 5) were expressed in absorbance.

2.5. Analysis of the ALP activity by colorimetric assay

On Day 10, ALP activity was determined by hydrolysis of the substrate thymolphthalein monophosphate and thymolphthalein release using a commercially available kit (Labtest Diagnostica). The cells were lysed for 30 min at room temperature with 2 mL of 0.1% sodium lauryl sulfate solution (Sigma‐Aldrich). Then, 50 µL of thymolphthalein monophosphate was mixed with 0.5 mL of 0.3 M diethanolamine buffer pH 10.1 and incubated at 37°C for 2 min before adding 50 µL of cell lysate. After 10 min at 37°C, 2 mL of a solution containing Na₂CO₃ (0.09 mM) and NaOH (0.25 mM) was added to stop the reaction, and the absorbance was measured at 590 nm using a μQuant spectrophotometer (BioTek Instruments, Inc.). The data (n = 5) were normalized to total protein content and expressed as μmol of thymolphthalein/h/mg protein in the same cell lysates.

2.6. Analysis of the RUNX2 protein expression by western blot

The RUNX2 protein expression was evaluated by western blot on Day 10. The cells were lysed with 300 μL of RIPA (radioimmunoprecipitation assay) buffer containing 1× protease inhibitor mixture (Roche Applied Science), 1 mM phenylmethanesulfonyl fluoride (Sigma‐Aldrich), and 25 μM MG132 proteasome inhibitor (Roche Applied Science) in an ultrasound machine (Misonix) with five pulses of 10 s. After 5 min of heating in a water bath at 100°C, 50 μg of protein from each sample were applied to an 8.5% polyacrylamide gel and subject to electrophoresis. The proteins were transferred to a PVDF membrane (Bio‐Rad Laboratories) in a Trans‐Blot SD device (Bio‐Rad Laboratories), and RUNX2 was detected by incubating with a primary anti‐RUNX2 antibody (1:1000, rabbit monoclonal antibody, Cell Signaling), followed by incubation with secondary antibody goat anti‐rabbit IgG‐horseradish peroxidase (HRP; 1:2000, Cell Signaling). The GAPDH protein was detected and used as the endogenous control with anti‐GAPDH primary antibody rabbit polyclonal IgG (1:1000, Santa Cruz Biotechnology) and secondary antibody goat anti‐rabbit IgG‐HRP conjugate (1:2000, Cell Signaling). After incubation with Clarity Western ECL Substrate (Bio‐Rad Laboratories), images were captured in a G:BOX device (Syngene). RUNX2 was quantified (n = 3) using ImageJ Software (NIH) and normalized to GAPDH protein expression.

2.7. Analysis of the extracellular matrix mineralization by Alizarin red staining

The extracellular matrix mineralization was evaluated using Alizarin red staining (Sigma‐Aldrich) on Day 21. The cultures were fixed for 24 h at 4°C with 2 mL of 10% formalin, dehydrated with alcohol, dried at room temperature, and stained with 2% Alizarin red. For qualitative analysis, images of the cultures were captured with a digital camera (Canon EOS Digital Rebel, 6.3 megapixels, with EF100 f/2.8 macrolens). To extract the calcium content, 280 µL of 10% acetic acid was added in each well, and the plates were shaken for 30 min. The solution was vortexed for 30 s, heated at 85°C for 10 min, and then placed on ice for 5 min. The tubes were centrifuged for 15 min at 20,000 rpm, 100 µL of supernatant from each tube was transferred to a 96‐well plate, and 40 µL of 10% ammonium hydroxide was added. The absorbance was measured at 405 nm using a μQuant spectrophotometer (BioTek Instruments Inc), and the data (n = 5) were expressed in absorbance.

2.8. Analysis of the gene expression of components of the BMP signaling pathway by PCR array

The expression profile of genes involved in the BMP signaling pathway was evaluated using a PCR array and the TaqMan Array Plates system (Thermo‐Fisher Scientific) on Day 7. The cDNA was synthesized from 2 µg of RNA using the SuperScript IV VILO Master Mix kit (Thermo Fisher Scientific), according to the manufacturer's instructions. Two cDNAs were synthesized for each experimental group, and the final volume of each reaction was 20 µL. Then, 40 µL of cDNAs was mixed with 400 µL of RNase/DNase free water and 440 µL of TaqMan Advanced Master Mix (Thermo to Fisher Scientific), and 800 µL of this mixture was added in a 384 to well plate containing TaqMan probes and target genes (Supporting Information: Table S2). The reactions were carried out in a QuantStudio™ 7 Flex System device (Applied Biosystems), and the data (n = 3) were analyzed in the Software ExpressionSuite (Thermo to Fisher Scientific). The heatmaps were constructed using the Heatmapper online software at the Wishart Research Group, University of Alberta, Edmonton, Canada. ²¹

2.9. Statistical analysis

The data of the bone morphometric parameters, gene expression of bone markers, ALP activity, protein expression, and extracellular matrix mineralization were analyzed by one‐way analysis of variance (ANOVA) and of the cell viability by two‐way ANOVA, followed by Tukey's posttest. The gene expression of BMP signaling pathway components was analyzed using the Student's t test for each gene in each experimental condition compared with the control. For all comparisons, the level of significance was set at p ≤ 0.05.

3. RESULTS

A detailed section of the results with a description of the charts and p values is presented in the Supporting Information: Data S1.

3.1. Induction of T1DM and T2DM

Because rats with DM were an important part of this study, blood glucose levels were measured over time as a parameter to confirm that the protocols we used could induce DM. The data confirmed that the protocols induced T1DM and T2DM because the normal blood glucose levels in rats range from 50 to 135 mg/dL (Supporting Information: Table S3). Also, it was necessary to evaluate the effect of both T1DM and T2DM on bone tissue. The three‐dimensional reconstructed microtomographic images (Supporting Information: Figure S1A) and morphometric parameters (Supporting Information: Figure S1B) of femurs harvested from control rats and rats with T1DM or T2DM confirmed that both diabetic conditions induced bone loss, with T2DM having a more pronounced effect than T1DM.

3.2. Comparison between MSCs derived from rats with T1DM‐HG1 or T2DM‐HG2

This study compared the effects of T1DM and T2DM on cell viability and osteoblast differentiation in MSCs grown in hyperglycemic conditions. MSCs from rats with T1DM‐HG1 had lower gene expression of the bone markers Runx2, Osterix, Alp, Oc, and Bsp than MSCs from control rats or rats with T2DM‐HG2 (Figure 1), and the phenotypic markers ALP activity, RUNX2 protein expression, extracellular matrix mineralization, and cell viability (Figure 2). In line with this, the modulation of BMP signaling pathway components was more pronounced in MSCs from rats with T1DM‐HG1 than T2DM‐HG2 (Figure 3). These findings suggest that T1DM and T2DM reduce cell viability and osteogenic potential in MSCs, with T1DM having a more pronounced effect.

Gene expression of the bone markers, *Runx2*, *Osterix*, alkaline phosphatase (*Alp*), osteocalcin (Oc), and bone sialoprotein (*Bsp*) of MSCs from healthy rats (control) grown in normoglycemic osteogenic medium and MSCs from rats with type 1 (T1DM‐HG1) or type 2 (T2DM‐HG2) diabetes mellitus grown in hyperglycemic osteogenic medium on Day 7. The data are presented as mean ± standard deviation (n = 4). *p ≤ 0.05.

Cell viability on Days 3, 7, and 10, alkaline phosphatase (ALP) activity on day 10, RUNX2 protein expression on Day 10, and extracellular matrix mineralization on Day 21 of MSCs from healthy rats (control) grown in normoglycemic osteogenic medium and MSCs from rats with type 1 (T1DM‐HG1) or type 2 (T2DM‐HG2) diabetes mellitus grown in hyperglycemic osteogenic medium. The data are presented as mean ± standard deviation (n = 5). *p ≤ 0.05.

Heatmap (A) and Venn diagram (B) of the gene expression of components of the bone morphogenetic protein (BMP) signaling pathway of MSCs from healthy rats (control) grown in normoglycemic osteogenic medium and MSCs from rats with type 1 (T1DM‐HG1) or type 2 (T2DM‐HG2) diabetes mellitus grown in hyperglycemic osteogenic medium on Day 7.

3.3. Comparison between MSCs derived from rats with T1DM‐HG1 or T1DM‐NG

Based on the reduced cell viability and osteoblast differentiation of MSCs induced by T1DM, we evaluated the effect of hyperglycemic stimulus withdrawal on the same cellular parameters. MSCs from rats with T1DM‐HG1 had lower Runx2 and Bsp gene expression (Figure 4) and RUNX2 protein expression, extracellular matrix mineralization, and cell viability (Figure 5), both of which were lower than MSCs from control rats. The expression profile of BMP signaling pathway components in MSCs from rats with T1DM‐NG tended to be more similar to that of MSCs from control rats than to that of MSCs from rats with T1DM‐HG1 (Figure 6). These data show that normoglycemia partially restored the disrupted cell viability and osteoblast differentiation of MSCs induced by T1DM.

3.4. Comparison between MSCs derived from rats with T2DM‐HG2 or T2DM‐NG

Because the normoglycemic environment had a positive effect on MSCs derived from rats with T1DM, we used the same rationale and performed the same assays for T2DM. In contrast with T1DM, all evaluated cellular parameters (Figures 7 and 8) indicated that the hyperglycemic stimulus withdrawal had a negative effect on MSCs from rats with T2DM. The expression profile of BMP signaling pathway components in MSCs from rats with T2DM‐HG2 and T2DM‐NG differed when compared with MSCs from control rats (Figure 9). This data set shows that, unlike T1DM, normoglycemia was unable to restore the reduced cell viability and osteoblast differentiation of MSCs induced by T2DM.

4. DISCUSSION

The deleterious effects of T1DM and T2DM on bone involve damages to tissue microarchitecture and composition, which are directly related to bone cell functions. ⁵ , ⁹ The findings of this study revealed that T1DM and T2DM disrupt osteoblast differentiation, with T1DM having a more pronounced effect than T2DM. We found that when MSCs derived from rats with T1DM but not T2DM were removed from a hyperglycemic environment, they recovered some of their osteogenic potential. Our findings also suggest that the inhibitory effects of T1DM and T2DM on osteoblast differentiation are due, at least in part, to BMP signaling pathway regulation.

The use of a single high dose injection of streptozotocin to induce T1DM and high fat diet feeding followed by a low dose injection of streptozotocin to induce T2DM are well established protocols that resulted in hyperglycemic conditions as measured by blood glucose levels in the animals. ²² , ²³ , ²⁴ , ²⁵ , ²⁶ The deleterious effects of T1DM and T2DM on bone tissue were clearly noticed by observing the morphometric parameters of rat femurs, which revealed that T2DM had a greater negative impact than T1DM. Data from the literature show that patients with T1DM have lower BMD than nondiabetic patients, whereas T2DM can either increase, decrease, or have no effect on BMD. ³ , ⁴ , ⁵ , ⁶ , ⁷ , ⁸ In rodents, experimental T1DM induced by streptozotocin decreased BMD, BV/TV, and Tb.Sp of femurs when diabetic rats were compared with nondiabetic rats. ²⁷ , ²⁸ The femurs of Zucker diabetic fatty rats that develop T2DM spontaneously exhibit lower BMD, BV/TV, and Tb.N than those of nondiabetic animals. ²⁹ , ³⁰ Our findings are consistent with these studies because we observed a negative impact of T1DM and T2DM on bone tissue; however, none of them compared the effects of both diabetes types, as we did here. High‐fat diet feeding associated with streptozotocin may modulate other cellular mechanisms than streptozotocin alone, which could explain why T2DM has more severe effects on bone tissue than T1DM, as a high‐fat diet is associated with bone loss. ³¹ , ³²

Despite the fact that bone loss was more evident in rats with T2DM than T1DM, the osteogenic potential of MSCs from rats with T1DM was lower than that of MSCs from rats with T2DM when both were cultured under hyperglycemic conditions. In vitro studies have shown that high glucose concentrations in the range we used here have a negative impact on osteoblast differentiation of MSCs. ³³ , ³⁴ , ³⁵ , ³⁶ , ³⁷ MSCs from rats with T1DM exhibited lower expression of bone marker genes than control, whereas MSCs from rats with T2DM exhibited a more complex response. Despite these differences in gene expressions between MSCs from rats with T1DM and T2DM, both resulted in less differentiated osteoblasts, as revealed by inhibition of phenotypic markers, with T1DM having a more pronounced effect. Because osteoblast differentiation is sensitive to increased glucose doses, the more deleterious effect of T1DM than T2DM on the osteogenic potential of MSCs may be explained by the higher glucose concentration of HG1 (33 mM) than HG2 (16 mM), both selected to mimic the blood glucose levels of the animals with T1DM and T2DM, respectively. ³⁴ One limitation of our study is that the effect of a high‐fat diet on bone tissue of animals with T2DM was not simulated in the in vitro environment, reducing the harmful effect of T2DM on osteoblast differentiation. The higher inhibitory effect of T1DM on osteoblast differentiation of MSCs than T2DM could be attributed, at least in part, to modulation of BMP signal transduction. Although T1DM and T2DM both regulated the gene expression of several key components of the BMP signaling pathway in MSCs grown under hyperglycemic conditions, only T1DM downregulated the expression of the osteogenic factors Bmp1, Bmp2, Bmp4, Bmp5, Gdf5, and Sox4, whereas T2DM only downregulated secondary BMP targets, such as Id1 and Smad 4. ³⁸ , ³⁹ , ⁴⁰ , ⁴¹ , ⁴² , ⁴³ In contrast to our hypothesis that hyperglycemia inhibits osteoblast differentiation by downregulating the BMP signaling pathway, it has been shown that high glucose promotes the formation of mineralized extracellular matrix in MC3T3‐E1 cells via the BMP‐4/Smad signaling pathway. ⁴⁴ Such differences could be explained by the fact that we used primary MSCs derived from rats with T1DM or T2DM, whereas they used a single mouse cell line.

To investigate if removing the hyperglycemic stimulus could restore the osteogenic potential of MSCs derived from rats with T1DM and T2DM, the cells were cultured under normoglycemic conditions. Indeed, the recovery of the normoglycemic environment partially restored the ability of MSCs derived from animals with T1DM to differentiate into osteoblasts. Insulin therapy has been shown to reduce the deleterious effects of T1DM on rodent bone tissue. ⁴⁵ The use of cinaciguat, a nitric oxide‐independent activator of oxidized guanylate cyclase, reduced the oxidative stress induced by hyperglycemia and restored the osteoblast differentiation capacity of MSCs derived from mice with T1DM. ¹² Clinically, it is possible to speculate that controlling glucose levels in patients with T1DM may help to reverse the bone damage induced by this metabolic disorder. In contrast to T1DM, the recovering effect of the osteogenic potential induced by removing the hyperglycemic stimulus was not observed in MSCs derived from animals with T2DM. In fact, most osteoblast markers were reduced in normoglycemic conditions, indicating that the cellular mechanisms triggered by T1DM and T2DM to modulate osteoblast differentiation are distinct and may be related to different signaling pathways regulated to induce hyperglycemia. Furthermore, the glucose concentration of the HG1 medium (33 mM) may be slightly toxic, as suggested by cell viability data and previously observed for glucose concentrations higher than 25 mM, which could explain why the normoglycemic environment partially recovered the osteogenic potential of MSCs derived from rats with T1DM. ³⁴ However, in the absence of the high‐fat diet stimulus, the lower glucose concentration in HG2 medium (16 mM) may have some beneficial effects on osteoblast differentiation, as the deletion of the hyperglycemia reduced the osteoblast differentiation of MSCs derived from rats with T2DM. In MC3T3‐E1 cells, glucose at a concentration of 16.5 mM was found to increase extracellular matrix mineralization compared with a normoglycemic condition. ⁴⁴ The restoration of the normoglycemic environment changed the gene expression profile of BMP signaling pathway components in MSCs from animals with T1DM and T2DM, reversing the downregulation of osteogenic factors, such as Bmp1, Gdf5, and Sox4 induced by T1DM and Gsc and Igf1 induced by T2DM. Although our findings suggest that the BMP signaling pathway is involved in the bone loss induced by DM, further studies are needed to clarify the cellular mechanisms involved in the hyperglycemia–BMP–osteoblast differentiation circuit and the actual in vivo implications of this circuit in clinical findings and therapies to treat bone defects of patients with diabetes.

We demonstrated that T1DM and T2DM disrupt osteoblast differentiation of MSCs, with T1DM having a more pronounced effect. To the best of our knowledge, this is the first study to show that removing the hyperglycemic stimulus partially restores the osteoblast differentiation capacity of MSCs derived from rats with T1DM but not with T2DM. Our findings also indicate that the decreased osteogenic potential of MSCs induced by both, but mainly, T1DM is due, at least in part, to the modulation of the BMP signaling pathway. These findings highlight the need for specific therapeutic approaches to treat T1DM‐ or T2DM‐induced bone loss, as both disrupt osteoblast differentiation at distinct levels and most likely through different mechanisms.

AUTHOR CONTRIBUTIONS

Denise Weffort, Leticia F. Adolpho, Alann T. P. Souza, Gileade P. Freitas, Helena B. Lopes, Fabiola S. Oliveira, Rayana L. Bighetti‐Trevisan, and Leticia Pitol‐Palin: data curation, formal analysis, investigation, methodology. Leticia F. Adolpho: writing (original draft), data curation, formal analysis, investigation, methodology. Doris H. Matsushita, Roberta Okamoto: conceptualization, investigation, methodology, writing (review and editing). Marcio M. Beloti: conceptualization, data curation, formal analysis, investigation, writing (original draft, review, and editing). Adalberto L. Rosa: conceptualization, data curation, formal analysis, funding acquisition, investigation, project administration, resources, writing (original draft, review, and editing).

CONFLICT OF INTEREST STATEMENT

The authors declare no conflict of interest.

ETHICS STATEMENT

This study was approved by the Ethics Committee on the Use of Animals of the Araçatuba Dental School, São Paulo State University Júlio de Mesquita Filho (Protocol #2019‐0152).

Supporting information

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ACKNOWLEDGMENTS

The authors would like to thank Roger Rodrigo Fernandes and Adriana Luísa Gonçalves for their technical assistance during the execution of the study, and ENAGO (www.enago.com) for the English language review. This work was supported by the São Paulo Research Foundation (FAPESP, Grants 2019/18711‐7 and 2017/12622‐7) and the National Council for Scientific and Technological Development (CNPq, Grant 307698/2021‐1).

Weffort D, Adolpho LF, Souza ATP, et al. Normoglycemia partially recovers the disrupted osteoblast differentiation of mesenchymal stem cells induced by type 1 but not type 2 diabetes mellitus. J Cell Biochem. 2023;124:1050‐1063. 10.1002/jcb.30434

Denise Weffort and Leticia Faustino Adolpho contributed equally to the study.

DATA AVAILABILITY STATEMENT

The data that support the findings of this study are available from the corresponding author upon reasonable request.

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9. Rubin MR. Bone cells and bone turnover in diabetes mellitus. Curr Osteoporos Rep. 2015;13(3):186‐191. 10.1007/s11914-015-0265-0 [DOI] [PubMed] [Google Scholar]
10. Chen S, Yang L, He S, et al. Preactivation of β‐catenin in osteoblasts improves the osteoanabolic effect of PTH in type 1 diabetic mice. J Cell Physiol. 2020;235(2):1480‐1493. 10.1002/jcp.29068 [DOI] [PubMed] [Google Scholar]
11. 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(3):455‐462. 10.3892/ijmm.2011.697 [DOI] [PubMed] [Google Scholar]
12. Kalyanaraman H, Schwaerzer G, Ramdani G, et al. Protein kinase G activation reverses oxidative stress and restores osteoblast function and bone formation in male mice with type 1 diabetes. Diabetes. 2018;67(4):607‐623. 10.2337/db17-0965 [DOI] [PMC free article] [PubMed] [Google Scholar]
13. Chinipardaz Z, Liu M, Graves D, Yang S. Diabetes impairs fracture healing through disruption of cilia formation in osteoblasts. Bone. 2021;153:116176. 10.1016/j.bone.2021.116176 [DOI] [PMC free article] [PubMed] [Google Scholar]
14. Figeac F, Tencerova M, Ali D, et al. Impaired bone fracture healing in type 2 diabetes is caused by defective functions of skeletal progenitor cells. Stem Cells. 2022;40(2):149‐164. 10.1093/stmcls/sxab011 [DOI] [PubMed] [Google Scholar]
15. Pitol‐Palin L, de Souza Batista FR, Gomes‐Ferreira PHS, et al. Different stages of alveolar bone repair process are compromised in the type 2 diabetes condition: an experimental study in rats. Biology. 2020;9(12):471. 10.3390/biology9120471 [DOI] [PMC free article] [PubMed] [Google Scholar]
16. Deng X, Xu M, Shen M, Cheng J. Effects of type 2 diabetic serum on proliferation and osteogenic differentiation of mesenchymal stem cells. J Diabetes Res. 2018;2018:1‐9. 10.1155/2018/5765478 [DOI] [PMC free article] [PubMed] [Google Scholar]
17. Phimphilai M, Pothacharoen P, Kongtawelert P, Chattipakorn N. Impaired osteogenic differentiation and enhanced cellular receptor of advanced glycation end products sensitivity in patients with type 2 diabetes. J Bone Miner Metab. 2017;35(6):631‐641. 10.1007/s00774-016-0800-9 [DOI] [PubMed] [Google Scholar]
18. Stolzing A, Sellers D, Llewelyn O, Scutt A. Diabetes induced changes in rat mesenchymal stem cells. Cells Tissues Organs. 2010;191(6):453‐465. 10.1159/000281826 [DOI] [PubMed] [Google Scholar]
19. Bouxsein ML, Boyd SK, Christiansen BA, Guldberg RE, Jepsen KJ, Müller R. Guidelines for assessment of bone microstructure in rodents using microcomputed tomography. J Bone Miner Res. 2010;25(7):1468‐1486. 10.1002/jbmr.141 [DOI] [PubMed] [Google Scholar]
20. Livak KJ, Schmittgen TD. Analysis of relative gene expression data using real‐time quantitative PCR and the 2−ΔΔCT method. Methods. 2001;25(4):402‐408. 10.1006/meth.2001.1262 [DOI] [PubMed] [Google Scholar]
21. Babicki S, Arndt D, Marcu A, et al. Heatmapper: web‐enabled heat mapping for all. Nucleic Acids Res. 2016;44(W1):W147‐W153. 10.1093/nar/gkw419 [DOI] [PMC free article] [PubMed] [Google Scholar]
22. Silva JC, Sampaio P, Fernandes MH, Gomes PS. The osteogenic priming of mesenchymal stem cells is impaired in experimental diabetes. J Cell Biochem. 2015;116(8):1658‐1667. 10.1002/jcb.25126 [DOI] [PubMed] [Google Scholar]
23. Skovsø S. Modeling type 2 diabetes in rats using high fat diet and streptozotocin. J Diabetes Investig. 2014;5(4):349‐358. 10.1111/jdi.12235 [DOI] [PMC free article] [PubMed] [Google Scholar]
24. Yin D, Tao J, Lee DD, et al. Recovery of islet β‐Cell function in streptozotocin‐induced diabetic mice. Diabetes. 2006;55(12):3256‐3263. 10.2337/db05-1275 [DOI] [PubMed] [Google Scholar]
25. Srinivasan K, Viswanad B, Asrat L, Kaul CL, Ramarao P. Combination of high‐fat diet‐fed and low‐dose streptozotocin‐treated rat: a model for type 2 diabetes and pharmacological screening. Pharmacol Res. 2005;52(4):313‐320. 10.1016/j.phrs.2005.05.004 [DOI] [PubMed] [Google Scholar]
26. Junod A, Lambert AE, Stauffacher W, Renold AE. Diabetogenicaction of streptozotocin: relationship of dose to metabolic response. J Clin Invest. 1969;48(11):2129‐2139. 10.1172/JCI106180 [DOI] [PMC free article] [PubMed] [Google Scholar]
27. Londzin P, Kisiel‐Nawrot E, Kocik S, et al. Effects of diosgenin on the skeletal system in rats with experimental type 1 diabetes. Biomed Pharmacother. 2020;129:110342. 10.1016/j.biopha.2020.110342 [DOI] [PubMed] [Google Scholar]
28. 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(3):461‐467. 10.1016/j.diabres.2012.03.005 [DOI] [PubMed] [Google Scholar]
29. Picke AK, Gordaliza Alaguero I, Campbell GM, et al. Bone defect regeneration and cortical bone parameters of type 2 diabetic rats are improved by insulin therapy. Bone. 2016;82:108‐115. 10.1016/j.bone.2015.06.001 [DOI] [PubMed] [Google Scholar]
30. Hamann C, Goettsch C, Mettelsiefen J, et al. Delayed bone regeneration and low bone mass in a rat model of insulin‐resistant type 2 diabetes mellitus is due to impaired osteoblast function. Am J Physiol‐Endocrinol Metabol. 2011;301(6):E1220‐E1228. 10.1152/ajpendo.00378.2011 [DOI] [PubMed] [Google Scholar]
31. Dai F, Zhang Y, Xu D, et al. Effects of long term diabetogenic high fat diet on bone in ovariectomized female rats. Biotech Histochem. 2023;98(1):20‐28. 10.1080/10520295.2022.2083685 [DOI] [PubMed] [Google Scholar]
32. Chen F, Wang Y, Wang H, et al. Flaxseed oil ameliorated high‐fat‐diet‐induced bone loss in rats by promoting osteoblastic function in rat primary osteoblasts. Nutr Metab. 2019;16:71. 10.1186/s12986-019-0393-0 [DOI] [PMC free article] [PubMed] [Google Scholar]
33. Gong F, Gao L, Ma L, Li G, Yang J. Uncarboxylated osteocalcin alleviates the inhibitory effect of high glucose on osteogenic differentiation of mouse bone marrow‐derived mesenchymal stem cells by regulating TP63. BMC Mol Cell Biol. 2021;22(1):24. 10.1186/s12860-021-00365-7 [DOI] [PMC free article] [PubMed] [Google Scholar]
34. Aswamenakul K, Klabklai P, Pannengpetch S, et al. Proteomic study of in vitro osteogenic differentiation of mesenchymal stem cells in high glucose condition. Mol Biol Rep. 2020;47(10):7505‐7516. 10.1007/s11033-020-05811-x [DOI] [PubMed] [Google Scholar]
35. Huang KC, Chuang PY, Yang TY, Huang TW, Chang SF. Hyperglycemia inhibits osteoblastogenesis of rat bone marrow stromal cells via activation of the Notch2 signaling pathway. Int J Med Sci. 2019;16(5):696‐703. 10.7150/ijms.32707 [DOI] [PMC free article] [PubMed] [Google Scholar]
36. Wang R, Zhang Y, Jin F, Li G, Sun Y, Wang X. High‐glucose‐induced miR‐214‐3p inhibits BMSCs osteogenic differentiation in type 1 diabetes mellitus. Cell Death Discov. 2019;5:143. 10.1038/s41420-019-0223-1 [DOI] [PMC free article] [PubMed] [Google Scholar]
37. Cao B, Liu N, Wang W. High glucose prevents osteogenic differentiation of mesenchymal stem cells via lncRNA AK028326/CXCL13 pathway. Biomed Pharmacother. 2016;84:544‐551. 10.1016/j.biopha.2016.09.058 [DOI] [PubMed] [Google Scholar]
38. Caddy JC, Luoma LM, Berry FB. FOXC1 negatively regulates BMP‐SMAD activity and Id1 expression during osteoblast differentiation. J Cell Biochem. 2020;121(5‐6):3266‐3277. 10.1002/jcb.29595 [DOI] [PubMed] [Google Scholar]
39. Asharani PV, Keupp K, Semler O, et al. Attenuated BMP1 function compromises osteogenesis, leading to bone fragility in humans and zebrafish. Am J Human Genetics. 2012;90(4):661‐674. 10.1016/j.ajhg.2012.02.026 [DOI] [PMC free article] [PubMed] [Google Scholar]
40. Wutzl A, Rauner M, Seemann R, et al. Bone morphogenetic proteins 2, 5, and 6 in combination stimulate osteoblasts but not osteoclasts in vitro. J Orthop Res. 2010;28(11):1431‐1439. 10.1002/jor.21144 [DOI] [PubMed] [Google Scholar]
41. Nissen‐Meyer LSH, Jemtland R, Gautvik VT, et al. Osteopenia, decreased bone formation and impaired osteoblast development in Sox4 heterozygous mice. J Cell Sci. 2007;120(16):2785‐2795. 10.1242/jcs.003855 [DOI] [PubMed] [Google Scholar]
42. Zeng Q, Li X, Beck G, Balian G, Shen FH. Growth and differentiation factor‐5 (GDF‐5) stimulates osteogenic differentiation and increases vascular endothelial growth factor (VEGF) levels in fat‐derived stromal cells in vitro. Bone. 2007;40(2):374‐381. 10.1016/j.bone.2006.09.022 [DOI] [PubMed] [Google Scholar]
43. Ahrens M, Ankenbauer T, Schröder D, Hollnagel A, Mayer H, Gross G. Expression of human bone morphogenetic Proteins‐2 or ‐4 in murine mesenchymal progenitor C3H10T½ cells induces differentiation into distinct mesenchymal cell lineages. DNA Cell Biol. 1993;12(10):871‐880. 10.1089/dna.1993.12.871 [DOI] [PubMed] [Google Scholar]
44. Takeno A, Kanazawa I, Tanaka K, et al. High glucose promotes mineralization via bone morphogenetic protein 4‐Smad signals in early stage of osteoblast differentiation. Diabetol Int. 2020;12(2):171‐180. 10.1007/s13340-020-00463-5 [DOI] [PMC free article] [PubMed] [Google Scholar]
45. Limirio PHJO, Soares PBF, Venâncio JF, Rabelo GD, Soares CJ, Dechichi P. Type I diabetes mellitus and insulin therapy on bone microarchitecture, composition and mechanical properties. Curr Diabetes Rev. 2022;18(8):e301121198427. 10.2174/1573399818666211130142153 [DOI] [PubMed] [Google Scholar]

Associated Data

This section collects any data citations, data availability statements, or supplementary materials included in this article.

Supplementary Materials

Supporting information.

JCB-124--s005.docx^{(20.3KB, docx)}

Supporting information.

JCB-124--s003.tif^{(4.6MB, tif)}

Supporting information.

JCB-124--s001.docx^{(16.4KB, docx)}

Supporting information.

JCB-124--s002.docx^{(20.6KB, docx)}

Supporting information.

JCB-124--s004.docx^{(15KB, docx)}

Data Availability Statement

The data that support the findings of this study are available from the corresponding author upon reasonable request.

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[jcb30434-bib-0015] 15. Pitol‐Palin L, de Souza Batista FR, Gomes‐Ferreira PHS, et al. Different stages of alveolar bone repair process are compromised in the type 2 diabetes condition: an experimental study in rats. Biology. 2020;9(12):471. 10.3390/biology9120471 [DOI] [PMC free article] [PubMed] [Google Scholar]

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[jcb30434-bib-0017] 17. Phimphilai M, Pothacharoen P, Kongtawelert P, Chattipakorn N. Impaired osteogenic differentiation and enhanced cellular receptor of advanced glycation end products sensitivity in patients with type 2 diabetes. J Bone Miner Metab. 2017;35(6):631‐641. 10.1007/s00774-016-0800-9 [DOI] [PubMed] [Google Scholar]

[jcb30434-bib-0018] 18. Stolzing A, Sellers D, Llewelyn O, Scutt A. Diabetes induced changes in rat mesenchymal stem cells. Cells Tissues Organs. 2010;191(6):453‐465. 10.1159/000281826 [DOI] [PubMed] [Google Scholar]

[jcb30434-bib-0019] 19. Bouxsein ML, Boyd SK, Christiansen BA, Guldberg RE, Jepsen KJ, Müller R. Guidelines for assessment of bone microstructure in rodents using microcomputed tomography. J Bone Miner Res. 2010;25(7):1468‐1486. 10.1002/jbmr.141 [DOI] [PubMed] [Google Scholar]

[jcb30434-bib-0020] 20. Livak KJ, Schmittgen TD. Analysis of relative gene expression data using real‐time quantitative PCR and the 2−ΔΔCT method. Methods. 2001;25(4):402‐408. 10.1006/meth.2001.1262 [DOI] [PubMed] [Google Scholar]

[jcb30434-bib-0021] 21. Babicki S, Arndt D, Marcu A, et al. Heatmapper: web‐enabled heat mapping for all. Nucleic Acids Res. 2016;44(W1):W147‐W153. 10.1093/nar/gkw419 [DOI] [PMC free article] [PubMed] [Google Scholar]

[jcb30434-bib-0022] 22. Silva JC, Sampaio P, Fernandes MH, Gomes PS. The osteogenic priming of mesenchymal stem cells is impaired in experimental diabetes. J Cell Biochem. 2015;116(8):1658‐1667. 10.1002/jcb.25126 [DOI] [PubMed] [Google Scholar]

[jcb30434-bib-0023] 23. Skovsø S. Modeling type 2 diabetes in rats using high fat diet and streptozotocin. J Diabetes Investig. 2014;5(4):349‐358. 10.1111/jdi.12235 [DOI] [PMC free article] [PubMed] [Google Scholar]

[jcb30434-bib-0024] 24. Yin D, Tao J, Lee DD, et al. Recovery of islet β‐Cell function in streptozotocin‐induced diabetic mice. Diabetes. 2006;55(12):3256‐3263. 10.2337/db05-1275 [DOI] [PubMed] [Google Scholar]

[jcb30434-bib-0025] 25. Srinivasan K, Viswanad B, Asrat L, Kaul CL, Ramarao P. Combination of high‐fat diet‐fed and low‐dose streptozotocin‐treated rat: a model for type 2 diabetes and pharmacological screening. Pharmacol Res. 2005;52(4):313‐320. 10.1016/j.phrs.2005.05.004 [DOI] [PubMed] [Google Scholar]

[jcb30434-bib-0026] 26. Junod A, Lambert AE, Stauffacher W, Renold AE. Diabetogenicaction of streptozotocin: relationship of dose to metabolic response. J Clin Invest. 1969;48(11):2129‐2139. 10.1172/JCI106180 [DOI] [PMC free article] [PubMed] [Google Scholar]

[jcb30434-bib-0027] 27. Londzin P, Kisiel‐Nawrot E, Kocik S, et al. Effects of diosgenin on the skeletal system in rats with experimental type 1 diabetes. Biomed Pharmacother. 2020;129:110342. 10.1016/j.biopha.2020.110342 [DOI] [PubMed] [Google Scholar]

[jcb30434-bib-0028] 28. 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(3):461‐467. 10.1016/j.diabres.2012.03.005 [DOI] [PubMed] [Google Scholar]

[jcb30434-bib-0029] 29. Picke AK, Gordaliza Alaguero I, Campbell GM, et al. Bone defect regeneration and cortical bone parameters of type 2 diabetic rats are improved by insulin therapy. Bone. 2016;82:108‐115. 10.1016/j.bone.2015.06.001 [DOI] [PubMed] [Google Scholar]

[jcb30434-bib-0030] 30. Hamann C, Goettsch C, Mettelsiefen J, et al. Delayed bone regeneration and low bone mass in a rat model of insulin‐resistant type 2 diabetes mellitus is due to impaired osteoblast function. Am J Physiol‐Endocrinol Metabol. 2011;301(6):E1220‐E1228. 10.1152/ajpendo.00378.2011 [DOI] [PubMed] [Google Scholar]

[jcb30434-bib-0031] 31. Dai F, Zhang Y, Xu D, et al. Effects of long term diabetogenic high fat diet on bone in ovariectomized female rats. Biotech Histochem. 2023;98(1):20‐28. 10.1080/10520295.2022.2083685 [DOI] [PubMed] [Google Scholar]

[jcb30434-bib-0032] 32. Chen F, Wang Y, Wang H, et al. Flaxseed oil ameliorated high‐fat‐diet‐induced bone loss in rats by promoting osteoblastic function in rat primary osteoblasts. Nutr Metab. 2019;16:71. 10.1186/s12986-019-0393-0 [DOI] [PMC free article] [PubMed] [Google Scholar]

[jcb30434-bib-0033] 33. Gong F, Gao L, Ma L, Li G, Yang J. Uncarboxylated osteocalcin alleviates the inhibitory effect of high glucose on osteogenic differentiation of mouse bone marrow‐derived mesenchymal stem cells by regulating TP63. BMC Mol Cell Biol. 2021;22(1):24. 10.1186/s12860-021-00365-7 [DOI] [PMC free article] [PubMed] [Google Scholar]

[jcb30434-bib-0034] 34. Aswamenakul K, Klabklai P, Pannengpetch S, et al. Proteomic study of in vitro osteogenic differentiation of mesenchymal stem cells in high glucose condition. Mol Biol Rep. 2020;47(10):7505‐7516. 10.1007/s11033-020-05811-x [DOI] [PubMed] [Google Scholar]

[jcb30434-bib-0035] 35. Huang KC, Chuang PY, Yang TY, Huang TW, Chang SF. Hyperglycemia inhibits osteoblastogenesis of rat bone marrow stromal cells via activation of the Notch2 signaling pathway. Int J Med Sci. 2019;16(5):696‐703. 10.7150/ijms.32707 [DOI] [PMC free article] [PubMed] [Google Scholar]

[jcb30434-bib-0036] 36. Wang R, Zhang Y, Jin F, Li G, Sun Y, Wang X. High‐glucose‐induced miR‐214‐3p inhibits BMSCs osteogenic differentiation in type 1 diabetes mellitus. Cell Death Discov. 2019;5:143. 10.1038/s41420-019-0223-1 [DOI] [PMC free article] [PubMed] [Google Scholar]

[jcb30434-bib-0037] 37. Cao B, Liu N, Wang W. High glucose prevents osteogenic differentiation of mesenchymal stem cells via lncRNA AK028326/CXCL13 pathway. Biomed Pharmacother. 2016;84:544‐551. 10.1016/j.biopha.2016.09.058 [DOI] [PubMed] [Google Scholar]

[jcb30434-bib-0038] 38. Caddy JC, Luoma LM, Berry FB. FOXC1 negatively regulates BMP‐SMAD activity and Id1 expression during osteoblast differentiation. J Cell Biochem. 2020;121(5‐6):3266‐3277. 10.1002/jcb.29595 [DOI] [PubMed] [Google Scholar]

[jcb30434-bib-0039] 39. Asharani PV, Keupp K, Semler O, et al. Attenuated BMP1 function compromises osteogenesis, leading to bone fragility in humans and zebrafish. Am J Human Genetics. 2012;90(4):661‐674. 10.1016/j.ajhg.2012.02.026 [DOI] [PMC free article] [PubMed] [Google Scholar]

[jcb30434-bib-0040] 40. Wutzl A, Rauner M, Seemann R, et al. Bone morphogenetic proteins 2, 5, and 6 in combination stimulate osteoblasts but not osteoclasts in vitro. J Orthop Res. 2010;28(11):1431‐1439. 10.1002/jor.21144 [DOI] [PubMed] [Google Scholar]

[jcb30434-bib-0041] 41. Nissen‐Meyer LSH, Jemtland R, Gautvik VT, et al. Osteopenia, decreased bone formation and impaired osteoblast development in Sox4 heterozygous mice. J Cell Sci. 2007;120(16):2785‐2795. 10.1242/jcs.003855 [DOI] [PubMed] [Google Scholar]

[jcb30434-bib-0042] 42. Zeng Q, Li X, Beck G, Balian G, Shen FH. Growth and differentiation factor‐5 (GDF‐5) stimulates osteogenic differentiation and increases vascular endothelial growth factor (VEGF) levels in fat‐derived stromal cells in vitro. Bone. 2007;40(2):374‐381. 10.1016/j.bone.2006.09.022 [DOI] [PubMed] [Google Scholar]

[jcb30434-bib-0043] 43. Ahrens M, Ankenbauer T, Schröder D, Hollnagel A, Mayer H, Gross G. Expression of human bone morphogenetic Proteins‐2 or ‐4 in murine mesenchymal progenitor C3H10T½ cells induces differentiation into distinct mesenchymal cell lineages. DNA Cell Biol. 1993;12(10):871‐880. 10.1089/dna.1993.12.871 [DOI] [PubMed] [Google Scholar]

[jcb30434-bib-0044] 44. Takeno A, Kanazawa I, Tanaka K, et al. High glucose promotes mineralization via bone morphogenetic protein 4‐Smad signals in early stage of osteoblast differentiation. Diabetol Int. 2020;12(2):171‐180. 10.1007/s13340-020-00463-5 [DOI] [PMC free article] [PubMed] [Google Scholar]

[jcb30434-bib-0045] 45. Limirio PHJO, Soares PBF, Venâncio JF, Rabelo GD, Soares CJ, Dechichi P. Type I diabetes mellitus and insulin therapy on bone microarchitecture, composition and mechanical properties. Curr Diabetes Rev. 2022;18(8):e301121198427. 10.2174/1573399818666211130142153 [DOI] [PubMed] [Google Scholar]

PERMALINK

Normoglycemia partially recovers the disrupted osteoblast differentiation of mesenchymal stem cells induced by type 1 but not type 2 diabetes mellitus

Denise Weffort

Leticia F Adolpho

Alann T P Souza

Gileade P Freitas

Helena B Lopes

Fabiola S Oliveira

Rayana L Bighetti‐Trevisan

Leticia Pitol‐Palin

Doris H Matsushita

Roberta Okamoto

Marcio M Beloti

Adalberto L Rosa

Abstract

1. INTRODUCTION

2. MATERIALS AND METHODS

2.1. Induction of T1DM and T2DM

2.2. MSC harvesting, culture, and osteoblast differentiation

2.3. Analysis of the gene expression of bone markers by real‐time polymerase chain reaction (RT‐qPCR)

2.4. Analysis of the cell viability by colorimetric assay

2.5. Analysis of the ALP activity by colorimetric assay

2.6. Analysis of the RUNX2 protein expression by western blot

2.7. Analysis of the extracellular matrix mineralization by Alizarin red staining

2.8. Analysis of the gene expression of components of the BMP signaling pathway by PCR array

2.9. Statistical analysis

3. RESULTS

3.1. Induction of T1DM and T2DM

3.2. Comparison between MSCs derived from rats with T1DM‐HG1 or T2DM‐HG2

Figure 1.

Figure 2.

Figure 3.

3.3. Comparison between MSCs derived from rats with T1DM‐HG1 or T1DM‐NG

Figure 4.

Figure 5.

Figure 6.

3.4. Comparison between MSCs derived from rats with T2DM‐HG2 or T2DM‐NG

Figure 7.

Figure 8.

Figure 9.

4. DISCUSSION

AUTHOR CONTRIBUTIONS

CONFLICT OF INTEREST STATEMENT

ETHICS STATEMENT

Supporting information

ACKNOWLEDGMENTS

DATA AVAILABILITY STATEMENT

REFERENCES

Associated Data

Supplementary Materials

Data Availability Statement

ACTIONS

PERMALINK

RESOURCES

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