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Arteriosclerosis, Thrombosis, and Vascular Biology. 1999;19:2112-2118

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(Arteriosclerosis, Thrombosis, and Vascular Biology. 1999;19:2112-2118.)
© 1999 American Heart Association, Inc.


Vascular Biology

Dexamethasone Enhances In Vitro Vascular Calcification by Promoting Osteoblastic Differentiation of Vascular Smooth Muscle Cells

Katsuhito Mori; Atsushi Shioi; Shuichi Jono; Yoshiki Nishizawa; Hirotoshi Morii

From the Second Department of Internal Medicine, Osaka City University Medical School, Osaka, Japan.

Correspondence and reprint requests to Atsushi Shioi, MD, Second Department of Internal Medicine, Osaka City University Medical School, 1-4-3 Asahi-machi, Abeno-ku, Osaka 545-8585, Japan. E-mail as{at}msic.med.osaka-cu.ac.jp


*    Abstract
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*Abstract
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down arrowDiscussion
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Abstract—Vascular calcification is often associated with atherosclerotic lesions. Moreover, the process of atherosclerotic calcification has several features similar to the mineralization of skeletal tissue. Therefore, we hypothesized that vascular smooth muscle cells might acquire osteoblastic characteristics during the development of atherosclerotic lesions. In the present study, we investigated the effect of dexamethasone (Dex), which is well known to be a potent stimulator of osteoblastic differentiation in vitro, on vascular calcification by using an in vitro calcification model. We demonstrated that Dex increased bovine vascular smooth muscle cell (BVSMC) calcification in a dose- and time-dependent manner. Dex also enhanced several phenotypic markers of osteoblasts, such as alkaline phosphatase activity, procollagen type I carboxy-terminal peptide production, and cAMP responses to parathyroid hormone in BVSMCs. We also examined the effects of Dex on human osteoblast-like (Saos-2) cells and compared its effects on BVSMCs and Saos-2 cells. The effects of Dex on alkaline phosphatase activity and the cAMP response to parathyroid hormone in BVSMCs were less prominent than those in Saos-2 cells. Interestingly, we detected that Osf2/Cbfa1, a key transcription factor in osteoblastic differentiation, was expressed in both BVSMCs and Saos-2 cells and that Dex increased the gene expression of both transcription factors. These findings suggest that Dex may enhance osteoblastic differentiation of BVSMCs in vitro.


Key Words: alkaline phosphatase • core-binding factor-{alpha}1 • atherosclerosis


*    Introduction
up arrowTop
up arrowAbstract
*Introduction
down arrowMethods
down arrowResults
down arrowDiscussion
down arrowReferences
 
Calcification is a common feature of advanced atherosclerotic lesions.1 Atherosclerotic plaque calcification, especially in the coronary arteries, is associated with clinical complications such as myocardial infarction, impaired vascular tone, dissection in angioplasty, poor surgical outcome, and coronary insufficiency due to loss of aortic recoil.2 3 4 5 Although the mechanism of vascular calcification remains to be established, recent evidence suggests that it has several features similar to mineralization in skeletal tissue, including the expression of bone morphogenetic protein-2 (BMP-2), osteocalcin, and osteopontin and the presence of the bone mineral hydroxyapatite and matrix vesicles.6 7 8 9 10 11

To clarify the mechanism of vascular calcification, we developed an in vitro calcification system in which diffuse calcification can be induced by culturing bovine vascular smooth muscle cells (BVSMCs) in the presence of ß-glycerophosphate (ß-GP).12 In this model, alkaline phosphatase (ALP), which is 1 of the markers for osteoblastic differentiation, is critical for vascular calcification and the expression of osteopontin mRNA, which increases during the development of calcification. Furthermore, we identified a local calcium-regulating system in which parathyroid hormone (PTH) –related peptide plays an important role as an autocrine/paracrine regulator of vascular calcification.13 Through recent evidence demonstrated by us and other investigators,5 14 we hypothesized that VSMCs might acquire osteoblastic characteristics during the development of atherosclerotic lesions.

Osteoblastic differentiation is a multistep process, proceeding through defined stages of maturation from a committed progenitor cell of mesenchymal origin capable of proliferation to a postproliferative osteoblast expressing bone phenotypic markers.15 However, the molecular basis of osteoblast-specific gene expression and differentiation remains unclear. Recently, a key regulatory transcription factor in osteoblastic differentiation, osteoblast-specific transcription factor-2/core-binding factor-{alpha} subunit 1 (Osf2/Cbfa1), has been identified. The Osf2/Cbfa1 gene generates 2 types of transcripts, osteoblast-specific and T cell–specific isoforms.16 17 18 In the mouse, the osteoblast isoform is different from the T-cell isoform in that the former contains a unique 87–amino acid sequence at its amino-terminal end. However, the precise roles of the 2 transcripts in osteoblastic differentiation still remain unclear. The homozygous Osf2/Cbfa1 (–/–) mouse shows a total lack of bone and a retention of the partially calcified cartilaginous skeleton.18 In humans, mutations of this gene cause cleidocranial dysplasia, an autosomal dominant skeletal disorder.17 19 Moreover, overexpression of the osteoblast isoform in nonosteoblastic cells induces expression of the principal osteoblast-specific genes, such as {alpha}1(I) procollagen, osteopontin, bone sialoprotein, and osteocalcin.16 Therefore, Osf2/Cbfa1 (osteoblast isoform) is thought to be 1 of the "master genes" for osteoblastic differentiation.

Dexamethasone (Dex), a potent, synthetic glucocorticoid, is well known to promote differentiation of progenitor cells, such as bone marrow stromal cells, to the osteoblastic phenotype.20 Chronic treatment of osteoblast cultures with Dex increases the number of mineralized bone nodules in primary fetal rat calvarial osteoblast cultures.21 22 Glucocorticoid promotes phenotypic markers of osteoblast differentiation, such as ALP, cAMP responses to PTH, osteopontin, bone sialoprotein, and osteocalcin, while it depresses production of insulin-like growth factor-I and type I collagen by osteoblasts.20 23 24 25 26 27 28 29 30

Although glucocorticoids have been shown to inhibit in vitro proliferation of VSMCs and prevent the development of atherosclerosis in experimental animals,31 32 33 34 35 retrospective studies including pathological findings obtained at autopsy have suggested that glucocorticoids adversely affect atherogenesis in humans.36 Moreover, the involvement of glucocorticoids in atherogenesis is supported by the strong correlation between an increased serum cortisol level in humans and the extent of coronary artery disease.37 Furthermore, long-term administration of glucocorticoids induces several metabolic and pathophysiological complications, such as insulin resistance, hypertension, and hyperlipidemia, which are thought to be coronary risk factors. However, the mechanism of glucocorticoid action in atherogenesis is poorly understood. Considering the fact that calcification is a common feature of advanced atherosclerotic lesions, it is important to evaluate the effect of glucocorticoids on vascular calcification.

In this study, we investigated the effect of Dex on vascular calcification by using an in vitro calcification model. We first demonstrated that Dex increased calcium deposition in a time- and dose-dependent manner. In this process, Dex increased ALP activity, its mRNA expression, and procollagen type I C-peptide (PICP) production and influenced cAMP responses to PTH. Finally, we demonstrated that Dex promoted expression of the Osf2/Cbfa1 gene. These results suggest that Dex may stimulate vascular calcification by promoting osteoblastic differentiation of VSMCs.


*    Methods
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up arrowIntroduction
*Methods
down arrowResults
down arrowDiscussion
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Reagents
Media, FCS, and sodium pyruvate were purchased from GIBCO. ß-GP and Dex were obtained from Sigma Chemical Co. Unless otherwise mentioned, all other reagents were obtained from Wako Pure Chemical Industries, Ltd.

Cell Culture and In Vitro Calcification
BVSMCs were obtained from the media of aortas by an explant method as previously described.12 Cells that had migrated from the explants were collected and maintained in Dulbecco's modified Eagle's medium (DMEM; high glucose [4.5 g/L]) containing 15% FCS and 10 mmol/L sodium pyruvate supplemented with 100 U/mL penicillin and 100 µg/mL streptomycin (growing medium) at 37°C in a humidified atmosphere containing 5% CO2. The cells up to passage 8 were used for experiments. Human osteoblast-like cells (Saos-2) and Jurkat cells were obtained from the American Type Culture Collection (Manassas, Va) and maintained in DMEM supplemented with 15% FCS. Calcification of BVSMCs and Saos-2 cells was induced as previously described.12 After reaching confluence, the cells were incubated in DMEM containing 15% FCS in the presence of 10 mmol/L ß-GP. The medium was replaced with fresh medium every 2 days. In the time-course experiments, the beginning day of culture in the calcification medium was defined as day 0.

Quantification of Calcium Deposition
The cells were decalcified with 0.6N HCl for 24 hours. The calcium content was determined by measuring the concentrations of calcium in the HCl supernatant by the o-cresolphthalein complexone method (Calcium C-test Wako; Wako Pure Chemical Industries). After decalcification, the cells were washed 3 times with PBS and solubilized with 0.1N NaOH/0.1% SDS. The protein content was measured with a bicinchoninic acid protein assay kit (Pierce). The calcium content of the cell layer was normalized to protein content.

ALP Assay
After the cells were washed twice with PBS, the cellular proteins were solubilized with 1% Triton X-100 in 0.9% NaCl and centrifuged, and the supernatants were assayed for ALP activity as described previously.12 One unit was defined as the activity producing 1 nmol of p-nitrophenol for 30 minutes. Protein concentrations were determined with a bicinchoninic acid protein assay kit (Pierce).

PICP Assay
The cells were plated into 24-well plates and grown to confluence. The medium was replaced with phenol red–free DMEM containing 0.2% FCS in the presence of Dex. After the indicated period of incubation, the supernatants were collected and stored at -20°C until assay. PICP secreted into the culture medium by BVSMCs and Saos-2 cells was assessed by measuring the PICP content of the culture supernatant with an enzyme immunosorbent assay kit (PIP EIA kit, Takara). The data were normalized to the protein content of the cell layer.

Measurement of cAMP
cAMP responses to PTH were assessed by measuring intracellular cAMP. The cells were plated into 24-well plates and grown to confluence before treatment with either vehicle or various concentrations of Dex (10-10 to 10-7 mol/L). After 4 days of treatment with Dex, the cells were washed twice with PBS and preincubated for 10 minutes with DMEM containing 0.1% BSA and 1.0 mmol/L 3-isobutyl-1-methylxanthine at 37°C. Human PTH(1–34) (10-7 mol/L) was then added to the medium, and the cells were incubated for an additional 15 minutes at 37°C. Thereafter the medium was removed, and the cell layer containing cAMP was extracted with 500 µL of 5% trichloroacetic acid. One-hundred-microliter aliquots of these samples were washed 3 times with 5 volumes of water-saturated ethyl ether and then dried. The extract was analyzed for cAMP by utilizing a cAMP radioimmunoassay kit (Yamasa Shoyu).

Preparation of cDNA Probes
The human ALP (liver/bone/kidney type) cDNA probe was obtained from the Japanese Cancer Research Resources Bank, Osaka, Japan. Human Osf2/Cbfa1 cDNA probes (585-bp fragments) were obtained by reverse transcription of an mRNA from Saos-2 cells, followed by polymerase chain reaction and subcloning into the TA cloning vector (Invitrogen). Sequences of the obtained cDNA were confirmed by the dideoxy sequencing method. The Osf2/Cbfa1 cDNA probe, which contains the carboxy terminus of the coding region (684 to 1268 bp of human Osf2/Cbfa1 cDNA), can recognize both isoforms of Osf2/Cbfa1 gene transcripts, ie, osteoblast-specific and T cell–specific isoforms, as confirmed by Northern blot analysis with polyA+ RNA of Saos-2 and of Jurkat (human T-cell line) cells, respectively.

RNA Isolation and Northern Blot Analysis
Total RNA was prepared using the acid-guanidinium-isothiocyanate-phenol-chloroform extraction method. PolyA+ RNA was obtained by use of an mRNA isolation kit (Microfast Track kit, Invitrogen) by using oligo(dT) cellulose for adsorption. Twenty micrograms of total RNA and 1 microgram of polyA+ RNA were denatured and separated by electrophoresis on 1% agarose gels containing formaldehyde and transferred to a nylon filter (Hybond N, Amersham). Blots were prehybridized for 24 hours at 37°C in a buffer containing 50% formamide, 3x SSC (1x SSC is 0.15 mol/L NaCl and 15 mmol/L sodium citrate, pH 7.4), 50 mmol/L Tris-HCl (pH 7.5), 0.1% SDS, 20 µg/mL denatured salmon sperm DNA, and 1x Denhardt's solution and then hybridized at 37°C for 48 hours with cDNA probes for human ALP and Osf2/Cbfa1, which had been labeled with [{alpha}-32P]dCTP (3000 Ci/mL, New England Nuclear) by use of a random priming method (Megaprime cDNA labeling system, Amersham). Blots were washed and autoradiographed with x-ray film at -70°C. The amounts of mRNA were quantified by densitometric scanning and normalized by comparison with glyceraldehyde-3-phosphate dehydrogenase (GAPDH).

Measurements of DNA Synthesis
DNA synthesis of BVSMCs was evaluated by [3H]thymidine incorporation assays. The cells were grown in 24-well plates until confluent and then incubated in serum-free DMEM for 48 hours. After 48 hours, the medium was changed to DMEM containing 15% FCS or platelet-derived growth factor (PDGF)-BB (10 ng/mL) in the presence or absence of the indicated concentrations of Dex. The cells were subsequently incubated for 21 hours and then labeled with 1 µCi/mL [3H]thymidine (6.7 Ci/mmol, New England Nuclear) for an additional 3 hours. [3H]Thymidine incorporated into DNA was evaluated by trichloroacetic acid precipitation and counting in a scintillation counter (Beckman Instruments).

Statistics
In certain experiments, data were analyzed for statistical significance by ANOVA with post hoc analysis, unless otherwise stated. These analyses were performed with the assistance of a computer program (StatView version 4.11, Abacus Concepts).


*    Results
up arrowTop
up arrowAbstract
up arrowIntroduction
up arrowMethods
*Results
down arrowDiscussion
down arrowReferences
 
We first examined the effect of Dex on BVSMC calcification. As previously described, ß-GP induced calcium deposition in a time-dependent manner. In the presence of ß-GP, Dex (10-7 mol/L) significantly increased calcium deposition compared with calcified controls at each time point (Figure 1ADown). The calcium deposition in the Dex-treated group increased to 175% of the calcified control value on day 6. Likewise, Dex promoted calcium deposition in a dose-dependent manner on day 4, and the calcium deposition increased to 322% of the calcified control value at 10-7 mol/L (Figure 1BDown). These results suggest that Dex increases BVSMC calcification.



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Figure 1. Effects of Dex on BVSMC calcification. A, Time-dependent effects of Dex. Cells were cultured in calcification medium for the indicated time periods in the presence of 10-7 mol/L Dex as described in Methods. Calcium contents of the cell layers were measured by the o-cresolphthalein complexone method, normalized to cellular protein content, and are presented as mean±SEM (n=3). Differences compared with calcified controls at each time point were statistically significant (*P<0.05, Fisher's protected least significant difference test [PLSD]). Uncalcified control group is indicated by open circles, calcified control group by open squares, and Dex-treated group by closed squares. ß-GP (+) and (-) indicate the presence and absence of ß-GP, respectively. B, Dose-dependent effect of Dex. Cells were cultured in calcification medium for 4 days in the presence of the indicated doses of Dex. Calcium contents of the cell layers were assessed as described above and are presented as mean±SEM (n=3). Differences compared with calcified controls were statistically significant (*P<0.05, Fisher's PLSD). ß-GP (+) and (-) indicate the presence and absence of ß-GP, respectively.

ALP is known to be 1 of the phenotypic markers of osteoblastic differentiation. Because we reported that ALP plays an important role in this calcification system, we next examined the effect of Dex on ALP activity in BVSMCs. As a positive control of Dex's effect, we utilized human osteoblast-like (Saos-2) cells. In the absence of ß-GP, Dex (10-7 mol/L) enhanced ALP activity in a time-dependent manner, and ALP activity had increased to 222% of controls on day 6 (Figure 2ADown). On day 4, Dex dose-dependently increased ALP activity in the absence of ß-GP, and the maximal effect (236% of control) was observed at 10-7 mol/L (Figure 2BDown). In the presence of ß-GP, Dex also increased ALP activity in BVSMCs, but the response to Dex was less prominent than that in its absence (Table 1Down). Furthermore, a greater response was observed in Saos-2 cells, both in the absence and presence of ß-GP (Table 1Down). Next, we examined the effect of Dex on expression of the ALP gene in BVSMCs. Dex dose-dependently promoted the expression of ALP mRNA at 48 hours, and the maximal effect was observed at 10-7 mmol/L (180% increase of control; Figure 2CDown). Taken together, these results suggest that Dex may accelerate BVSMC calcification partially through enhancing expression of the ALP gene and its activity.



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Figure 2. Effects of Dex on ALP activity and expression of ALP mRNA in BVSMCs. A, Time-dependent effect of Dex. Cells were cultured for the indicated periods of time in the presence of 10-7 mol/L Dex as described in Methods. ALP activities were measured, normalized to cellular protein contents, and are presented as mean±SEM (n=3). Differences compared with controls (CTL) at each time point were statistically significant (*P<0.05, Fisher's PLSD). Control group is indicated by open circles and the Dex-treated group (Dex) by closed squares. B, Dose-dependent effect of Dex. Cells were cultured for 4 days in the presence of the indicated doses of Dex as described. ALP activities were measured, normalized to cellular protein contents, and are presented as mean±SEM (n=3). Differences compared with controls were statistically significant (*P<0.05, Fisher's PLSD). C, Dose-dependent effect of Dex on the expression of ALP mRNA in BVSMCs. Cells were cultured for 48 hours in the presence of the indicated doses of Dex as described. After 48-hour treatment, the cells were harvested for isolation of total RNA. Twenty micrograms of total RNA from BVSMCs was electrophoresed, blotted, and probed with the cDNA of human (liver/bone/kidney type) ALP. Top, Autoradiograph of Northern blot of ALP. Bottom, Densitometric analysis of autoradiograms was performed and results are presented as the ratio of ALP to GAPDH (mean±SEM, n=3). The differences compared with control (CTL) were statistically significant (*P<0.05, Fisher's PLSD).


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Table 1. Effects of Dex on ALP Activities and PICP Production in BVSMCs and Saos-2 Cells

Type I collagen is 1 of the early phenotypic markers for osteoblastic differentiation and may play an important role in this process. We utilized PICP as an index for type I collagen synthesis. We measured PICP contents of the culture supernatants to investigate whether Dex affects type I collagen synthesis by BVSMCs. In the absence of ß-GP, Dex (10-7 mol/L) increased PICP secretion by BVSMCs in a time-dependent manner up to day 6, and on day 6, PICP production by Dex-treated BVSMCs had reached 241% of untreated BVSMCs (Figure 3ADown). On day 4, Dex dose-dependently increased PICP production, and at 10-7 mol/L the maximal effect was observed (313% increase of control; Figure 3BDown). Interestingly, Dex exerted no stimulatory effect on PICP production by BVSMCs in the presence of ß-GP (Table 1Up). Furthermore, Dex did not affect PICP production by Saos-2 cells, irrespective of ß-GP (Table 1Up).



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Figure 3. Effects of Dex on PICP production by BVSMCs. A, Time-dependent effect of Dex. Cells were cultured for the indicated periods of time in the presence of 10-7 mol/L Dex as described in Methods. PICP contents of culture supernatants were measured by an enzyme immunosorbent assay and normalized to cellular protein contents. Data are presented as mean±SEM (n=3). Differences compared with control (CTL) at each time point were statistically significant (*P<0.05, Fisher's PLSD). Control group is indicated by open circles and Dex-treated group by closed squares. B, Dose-dependent effect of Dex. Cells were cultured for 4 days in the presence of the indicated doses of Dex as described. PICP contents of culture supernatants were measured by enzyme immunosorbent assay and normalized to cellular protein contents. Data are presented as mean±SEM (n=3). Differences compared with control were statistically significant (*P<0.05, Fisher's PLSD).

As another marker of osteoblastic differentiation, we investigated the effect of Dex on cAMP production in response to PTH stimulation. Dex stimulated cAMP responses to PTH in BVSMCs both in the presence and absence of ß-GP (Table 2Down). The stimulatory effect of Dex in the absence of ß-GP was greater than that in its presence. Moreover, Dex exerted greater responses in Saos-2 cells compared with BVSMCs (Table 2Down). These data suggest that Dex promotes cAMP responsiveness to PTH in BVSMCs as well as in Saos-2 cells.


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Table 2. Effects of Dex on cAMP Production Stimulated by PTH in BVSMCs and Saos-2 Cells

We next assessed the effect of Dex on gene expression of Osf2/Cbfa1 in BVSMCs. The mRNA of Osf2/Cbfa1 expressed in BVSMCs was compared with that in Saos-2 and Jurkat cells (Figure 4ADown). The transcript in BVSMCs was larger than that in Saos-2 and Jurkat cells. The transcripts detected in Saos-2 and Jurkat cells are thought to be human osteoblast– and T cell–specific isoforms, respectively. Additionally, Dex (10-7 mol/L) increased Osf2/Cbfa1 mRNA expression 24 hours after treatment in Saos-2, but the presence of ß-GP did not affect its expression (Figure 4BDown). In BVSMCs, Dex also enhanced mRNA expression in a time-dependent manner (Figure 4CDown). Furthermore, the presence of ß-GP exerted no apparent effect on expression at 24 hours (data not shown).



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Figure 4. Expression of Osf2/Cbfa1 mRNA. A, Northern blot analysis of Osf2/Cbfa1 in various cells. One microgram of polyA+ RNA from BVSMCs, Jurkat cells, and Saos-2 cells was electrophoresed, blotted, and probed with cDNAs of human Osf2/Cbfa1. B, Northern blot analysis of Osf2/Cbfa1in Saos-2 cells. Saos-2 cells were incubated for 24 hours with vehicle or 10-7 mol/L Dex in the presence or absence of ß-GP. After treatment with Dex, cells were harvested for isolation of polyA+ RNA. One microgram of polyA+ RNA from Saos-2 cells was electrophoresed, blotted, and probed with cDNAs of human Osf2/Cbfa1. Autoradiograph of Northern blot of Osf2/Cbfa1. C, Northern blot analysis of Osf2/Cbfa1in BVSMCs. Cells were cultured for the indicated periods of time in the presence of 10-7 mol/L Dex as described. After treatment with Dex, cells were harvested for isolation of polyA+ RNA. One microgram of polyA+ RNA from BVSMCs was electrophoresed, blotted, and probed with cDNA of human Osf2/Cbfa1. Top, Autoradiograph of Northern blot of Osf2/Cbfa1. Bottom, Densitometric analysis of autoradiograms was performed and results are presented as the ratio of Osf2/AML3/Cbfa1 to GAPDH (mean±SEM, n=3). Differences compared with control (CTL) at each time were statistically significant (*P<0.05, Fisher's PLSD).

Because it is likely that Dex may promote expression of osteoblastic markers in BVSMCs by inhibiting their proliferative capacity, we finally examined the effect of Dex on DNA synthesis in BVSMCs. The cells were incubated in DMEM containing 15% FCS or 10 ng/mL PDGF-BB in the presence or absence of the indicated concentrations of Dex for 24 hours. Both FCS and PDGF-BB stimulated DNA synthesis in BVSMCs, and Dex inhibited the stimulatory effect of PDGF-BB on DNA synthesis (Figure 5ADown). However, Dex did not affect DNA synthesis stimulated by FCS (Figure 5BDown), suggesting that Dex may not exert a direct effect on BVSMC proliferation in this calcification model.



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Figure 5. Effects of Dex on PDGF-BB–stimulated (A) or 15% FCS-stimulated (B) DNA synthesis in BVSMCs. Cells were cultured in serum-deprived DMEM for 48 hours. Thereafter, cells were incubated in DMEM containing PDGF-BB (10 ng/mL, A) or 15% FCS (B) for 21 hours in the presence or absence of the indicated concentrations of Dex followed by a pulse with 1 µCi/mL [3H]thymidine for an additional 3 hours. DNA synthesis was measured as trichloroacetic acid–insoluble radioactivity and data are presented as mean±SEM (n=4). Differences compared with PDGF-stimulated controls were statistically significant (*P<0.05, Fisher's PLSD).


*    Discussion
up arrowTop
up arrowAbstract
up arrowIntroduction
up arrowMethods
up arrowResults
*Discussion
down arrowReferences
 
Because glucocorticoids are well known to be potent stimulators of osteoblastic differentiation,20 21 22 38 we utilized Dex, a potent synthetic glucocorticoid, to induce BVSMCs to acquire osteoblastic characteristics. As shown in this study, Dex enhanced not only in vitro calcification but also several phenotypic markers for osteoblastic differentiation in BVSMCs, such as ALP expression, type I collagen production, and cAMP responsiveness to PTH (Figures 1Up, 2Up, and 3Up). Moreover, we confirmed the potency of Dex on osteoblastic differentiation by utilizing Saos-2 cells as a positive control. Regarding ALP activity and the cAMP response to PTH, the responsiveness of BVSMCs to Dex was less prominent than that of Saos-2 cells (Tables 1Up and 2Up). The less potent effect of Dex on BVSMCs may be ascribed to a heterogeneous population of cells in the BVSMC culture. Interestingly, Dex exerted no effect on PICP production by Saos-2 cells, whereas Dex increased PICP secretion by BVSMCs in the absence of ß-GP (Table 1Up). Early phenotypic markers for osteoblastic differentiation such as PICP may not be affected by Dex in well-differentiated osteoblastic cells. However, the precise mechanism by which such differences of responsiveness to Dex are induced remains to be clarified.

Several key factors in bone mineralization have been demonstrated in calcified lesions of arterial walls, such as matrix vesicles, BMP-2, osteopontin, matrix Gla protein, osteocalcin, and type I collagen.6 7 8 9 10 11 39 We previously demonstrated the significance of ALP, osteopontin, and PTH-related peptide in an in vitro model of vascular calcification by utilizing BVSMCs.12 13 Recently, a key regulatory factor in osteoblastic differentiation, Osf2/Cbfa1, has been identified. BMP-7 induces expression of the osteoblastic isoform, followed by its enhancement of the osteocalcin gene in nonosteoblastic cells.16 Therefore, the Osf2/Cbfa1 gene is thought to be 1 of the "master genes" of as well as a molecular marker for osteoblastic differentiation. In this study, we showed the presence of the Osf2/Cbfa1 gene in cultured BVSMCs as well as in Saos-2 cells (Figure 4AUp and 4BUp). Additionally, we cloned a 5' partial sequence of the bovine osteoblast–specific Osf2/Cbfa1 transcript by reverse transcription–polymerase chain reaction by using total RNA from BVSMCs in preliminary experiments (K.M. et al, unpublished data, 1998). This evidence suggests that cultured VSMCs may be committed to differentiate into osteoblastic cells under certain conditions. However, whether the transcript detected in BVSMCs is the osteoblast-specific isoform remains to be confirmed. Furthermore, Dex enhanced the gene expression of Osf2/Cbfa1 in a time dependent manner in BVSMCs (Figure 4CUp). Therefore, it is likely that Dex may promote osteoblastic differentiation of VSMCs by increasing the expression of the Osf2/Cbfa1 gene.

Linkage of phenotypic gene induction to the downregulation of proliferation is the hallmark of differentiation in numerous cell types.40 Some agents inhibiting the proliferation of osteoblast-lineage cells, such as hydroxyurea, can induce osteoblastic differentiation. It is therefore possible that antiproliferative agents of VSMCs may induce osteoblastic differentiation under certain conditions. Moreover, 17ß-estradiol has been reported to promote osteoblastic differentiation of bovine vascular cells and in vitro calcification without affecting cell growth.41 In this study, we examined the hypothesis that Dex may inhibit the proliferative capacity of BVSMCs, resulting in osteoblastic differentiation. Because Dex did not affect DNA synthesis in the presence of 15% FCS (Figure 5BUp), Dex may directly induce osteoblastic differentiation of BVSMCs without affecting their growth.

The mechanisms of glucocorticoid action on atherogenesis remain to be evaluated. When applied as anti-inflammatory drugs at high doses, glucocorticoids suppress the development of atherosclerosis in experimental animals, despite enhancement of hypertriglyceridemia and hypercholesterolemia.32 33 34 35 Glucocorticoids have also been shown to inhibit the proliferation of cultured VSMCs and the thrombin-induced expression of growth factors.31 42 On the other hand, glucocorticoids are capable of decreasing the expression of hepatic LDL receptors, stimulating the net synthesis of apoB-100 and apoB-48 and decreasing their intracellular degradation.43 These changes are potentially atherogenic, and the strong correlation between an increased serum cortisol level in humans and the extent of coronary artery disease has also been documented.37 In this study, we have shown that Dex increases in vitro calcification by promoting osteoblastic phenotypes in BVSMCs. Taking into consideration that chronic treatment with glucocorticoids induces osteoporosis and that vascular calcification is often associated with osteoporosis, it is suggested that Dex may develop and exacerbate vascular calcification. Further studies are necessary to clarify the long-term effect of glucocorticoid administration on the development of vascular calcification, especially calcified atherosclerotic plaque lesions.


*    Acknowledgments
 
This work was supported in part by grants-in-aid 08671177 (to Y.N.) and 09770793 (to A.S.) for scientific research from the Ministry of Education, Science, and Culture of Japan.

Received April 7, 1998; accepted February 4, 1999.


*    References
up arrowTop
up arrowAbstract
up arrowIntroduction
up arrowMethods
up arrowResults
up arrowDiscussion
*References
 

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