Cilostazol Inhibits Oxidative Stress–Induced Premature Senescence Via Upregulation of Sirt1 in Human Endothelial Cells
Objective— Cilostazol, a selective inhibitor of PDE3, has a protective effect on endothelium after ischemic vascular damage, through production of nitric oxide (NO). The purpose of the present study was to clarify the molecular mechanisms underlying the preventive effect of treatment with cilostazol on oxidative stress–induced premature senescence in human endothelial cells.
Methods and Results— Prematurely senescent human umbilical vein endothelial cells (HUVECs) were induced by treatment with hydrogen peroxide (H2O2) as judged by senescence-associated β-galactosidase assay (SA-βgal), cell morphological appearance, and plasminogen activator inhibitor-1 (PAI-1) expression. Treatment with H2O2 caused 93% of the cells to be SA-βgal positive, whereas 46% of cilostazol (100 μmol/L)-treated cells were positive. HUVECs treated with other cAMP-elevating agents and DETA-NO showed a reduction of SA-βgal–positive cells as well. Cilostazol increased phosphorylation of Akt at Ser473 and of endothelial nitric oxide synthase (eNOS) at Ser1177, with a dose-dependent increase in Sirt1 expression. Moreover, the effect of cilostazol on premature senescence was abrogated through inhibition of Sirt1.
Conclusions— Our results indicated that cilostazol exerted protective effects against endothelial senescence and dysfunction, and enhancement of NO production is a key mediator in upregulation of Sirt1.
The phenomenon of human aging is known to be a critical cardiovascular risk factor. Cellular senescence of endothelial cells has been proposed to be involved in endothelial dysfunction and atherogenesis.1 The lesions of human atherosclerosis have been extensively studied histologically, and these studies have demonstrated that there are vascular cells that exhibit the morphological features of cellular senescence.2
See accompanying article on page 1577
The telomere hypothesis is a widely accepted explanation of the occurrence of cellular senescence.3 Cessation of cell division after extended propagation in culture for a few weeks or months is related to the attrition of telomeres, which is termed replicative senescence. In addition to telomere attrition, some stressors such as oxidative stress elicit similar growth arrest within just a few days, referred to as stress-induced premature senescence (SIPS). Both types of senescence are accompanied by a specific set of changes in cell function, morphology, and gene expression.4 In addition to the above changes, recognized biomarkers of senescent cells include staining for β-galactosidase at pH of 6.0 as opposed to endogenous lysosomal enzyme detected at pH of 4.0 in normal cells.5
According to the free-radical theory, reactive oxygen species (ROS) may be potential candidates responsible for senescence and age-related diseases, and on production of high levels of ROS, the redox balance is disturbed and cells shift into a state of oxidative stress, which subsequently leads to premature senescence with shortening telomeres.6
A PDE3 inhibitor, cilostazol, is used as a vasodilating antiplatelet drug for treating intermittent claudication, and in preclinical studies was shown to have a protective effect on endothelial cells by increasing eNOS activity.7 Cilostazol increases intracellular cAMP content accordingly and activates protein kinase A (PKA) or PI3K/Akt signaling.8 As recently shown, endothelial NO can protect against a state of oxidative stress, and activation of eNOS and subsequent production of NO delay endothelial cellular senescence.9,10
In yeast, Sir2 (silent information regulator-2) has been identified as an NAD+-dependent histone deacetylase.11 Mammalian sirtuin 1 (Sirt1), the closest homolog of Sir2, regulates the cell cycle, senescence, apoptosis, and metabolism, by interacting with a number of molecules, including p53, PML, and PPAR-γ.12–14 A recent study showed that production of NO by caloric restriction increases Sirt1 expression and suggested that eNOS may be involved in regulating the expression of Sirt1 in murine white adipocytes.15 Therefore, we consider that the protective effect of cilostazol against vascular senescence may be attributed to upregulation of Sirt1.
In the present study, cilostazol inhibited oxidative stress–induced premature senescence, and the increased expression of Sirt1 by this drug played a critical role in prevention of endothelial senescence.
Materials and Methods
Cilostazol was kindly provided by Otsuka Pharmaceutical Co Ltd, Tokyo, Japan. Forskolin, rolipram, NG-nitro-l-arginine methyl ester hydrochloride (L-NAME) and LY294002 were purchased from Sigma. Myristoylated cell-permeable PKA inhibitor peptide sequence14–22 amide (PKAI) was from Alexis Biochemicals. (Z)-1-[2-(2-aminoethyl)-N-(2-ammonioethyl)amino] diazen-1-IM1,2 diolate (DETA-NO), S-nitrosoacetyl penicillamine (SNAP), 8 Br-cGMP, and 8 Br-cAMP were from Cayman Chemical. N-acetyl-cystein (NAC) was purchased from Calbiochemo.
Human umbilical vein endothelial cells (HUVECs) were purchased from CAMBREX (Walkersville, Md), and maintained in endothelial growth medium (EGM-2, EGM-2 singleQuots, CAMBREX). Population doubling levels (PDL) were calculated as described previously,16 and all experiments were performed at PDL of 8 to 9.
Measurement of cAMP Level
HUVECs were plated in 96-well plates at a density of 5×103 cells per well and cultured overnight. After 15-minute incubation with cilostazol, the medium was aspirated and a lysis buffer was added. cAMP concentration was determined using a cAMP EIA kit (Amersham Biosciences) according to the manufacturer’s instructions.
Inhibition of Sirt1
Proliferating cells were washed 3 times with growth medium and exposed for 24 hours to the indicated concentrations of sirtinol (Calbiochem) or nicotinamide (NAM, Wako Chemical Industries) diluted in medium. After exposure, the dishes were washed 3 times with inhibitor-free medium and cultured. Proliferating cells were transfected with 200 pmol/L siRNA for Sirt1 (GAT GAA GTT GAC CTC CTC A14 and TGA AGT GCC trichloroacetic acid (TCA) GAT ATT A) or control siRNA (Darmacom Co.) using silMPORTER (Upstate Cell Signaling Solutions).
Senescence-Associated β-Galactosidase (SA-βgal) Staining
HUVECs were grown in 100-mm collagen-coated dishes to 80% confluence. HUVECs were pretreated with vehicle (0.05% DMSO), cilostazol (1 to 100 μmol/L), forskolin (0.1 to 1 μmol/L), rolipram (10 to 100 μmol/L), DETA-NO (50, 100 μmol/L), or NAC (3, 5 mmol/L) diluted in EGM-2 medium for 3 days. HUVECs were washed 3 times with EGM-2 and then treated for 1 hour with 100 μmol/L H2O2 diluted in EGM-2. After the treatment, HUVECs were trypsinized, reseeding at the density of 1×105 in 60-mm dishes and cultured with EGM-2 containing these compounds for 10 days. At 10 days after treatment with H2O2, HUVECs were fixed and the proportion of SA-βgal–positive cells was determined as described by Dimri et al.5
NOS Activation Assay
NOS activity was determined using an NOS assay kit (Calbiochem) according to the manufacturer’s instructions.
BrdU Incorporation Assay
BrdU incorporation was analyzed using a commercial kit (Roche).
Cells were lysed on ice for 1 hour in buffer (50 mmol/L Tris-HCl, pH 7.6, 150 mmol/L NaCl, 1%NP-40, 0.1%SDS, 1 mmol/L dithiothreitol, 1 mmol/L sodium vanadate, 1 mmol/L phenylmethylsulfonyl fluoride, 10 μg/mL aprotinin, 10 μg/mL leupeptin, and 10 mmol/L sodium fluoride). Equal amounts of protein were separated by SDS-polyacrylamide gel electrophoresis and transferred to nitrocellulose membranes. After blocking, the filters were incubated with the following antibodies; antiphospho-eNOS (Ser1177), antiphospho-Akt (Ser473), anti-Akt (Cell Signaling Technology), antieNOS (BD Transduction Laboratories), antiacetyl-p53 (Lys373/382), antip53, anti-Sirt1 (Santa Cruz Biotechnology Inc), anti-PAI-1 (Molecular Innovations Inc), and anti-β-actin (Sigma). After washing and incubation with horseradish peroxidase-conjugated antirabbit or antimouse IgG (Amersham) for 1 hour, the antigen-antibody complexes were visualized using an enhanced chemiluminescence system (Amersham).
Real-Time Quantitative Reverse Transcription
Expression of Sirt1 in HUVECs was measured by quantitative RT-polymerase chain reaction (PCR). Total RNA in HUVECs was isolated with ISOGEN (Nippon gene Inc). After treatment with Rnase-free Dnase for 30 minutes, total RNA (50 ng/μL) was reverse transcribed with random hexamers and oligo d(T) primers. The expression level of Sirt1 relative to GAPDH was determined by means of staining with SYBR green dye and a LineGene fluorescent quantitative detection system (Bioflux Co), as recommended by the manufacturer. Primer quality was verified by dissociation curve analysis, the slopes of standard curves, and reactions without RT. The following primers were used: Sirt1 (forward (F) 5′-CCTGACTTCAGATCAAGAGACGGT-3′; reverse (R) 5′-CTGATTAAAAATGTCTCCACGAACAG-3′, GAPDH F 5′-ACCACAGTCCATGCCATCAC-3′; R 5′-TCCACCACCCTGTTGCTGTA-3′).
The animal experiments were approved by our institutional review board. Ten-week-old SPF male wild-type BALB/c mice (n=40, weighing approximately 25 g) were supplied by Charles River Laboratories Inc. Animals were housed under a 12-hour light/dark cycle and fed a normal diet. These mice were administered 25 mg/kg paraquat (1,1-dimethyl-4,4-bipyridinium) (Wako Chemical) by intraperitoneal injection. Then mice were randomly assigned to 2 treatment groups (control group, n=20; cilostazol group, n=20). The each group received gavage administration of vehicle alone or cilostazol 60 mg/kg/d for their lifetime. We made diabetic mice (n=40) by a single intraperitoneal injection of streptozotocin (STZ; 60 mg/kg, Sigma). Tail blood glucose was assayed 3 days after injection using glucose test strips (Roche). The mice were killed by cervical dislocation. The aorta was removed after systemic perfusion with phosphate-buffered saline (PBS) for histological examination. The proportion of SA-βgal–positive cells was analyzed by NIH image software. The primary antibody was purified rat antimouse CD31 (platelet endothelial cell adhesion molecule; PECAM-1) monoclonal antibody from pharmingen. ROS were measured with 2′, 7′-Dichlorodihydrofluorescein, diacetate (DCF) (Sigma). As previously described by Shi et al,17 the aorta was rapidly removed and placed in oxygenated (12% O2, 5% CO2) physiological salt solution (PSS) of the following composition (NaCl 130 mmol/L, KCl 4.7 mmol/L, CaCl2 1.6 mmol/L, MgSO4 1.17 mmol/L, NaHCO3 14.9 mmol/L, KH2PO4 1.18 mmol/L, EDTA 0.026 mmol/L, glucose 1.0 mmol/L). The living aorta was carefully isolated, cannulated (24G, Terumo Co Ltd) at both ends, pressured, and loaded with DCF solution (10 μmol/L) for 10 minutes. Then the aorta was washed by PSS 3 times, embedded in OCT medium, and cryosectioned. TOTO-3 for nuclear staining, secondary antibodies (Alexa Fluor 488 donkey antirat IgG and Alexa Fluor 594 donkey antirat IgG), and antifade reagent were from Molecular Probe (Invitrogen). Fluorescent images were taken and analyzed using a confocal laser microscope (LSM510, Carl Zeiss MicroImaging Co Ltd). Urinary 8-Hydroxydeoxyguanosine (8-OhdG) and creatinine were measured using a DNA damage ELISA kit (Stressgen) and creatinine assay kit (Cayman chemical), respectively.
Telomerase activity was measured with 2 μg protein using a telomerase PCR-ELISA kit according to the manufactures instructions (Chemicon, Temecula).
Values are shown as mean±SEM in the text and figures. Differences between the groups were analyzed using 1-way analysis of variance, followed by Bonferroni test. Probability values less than 0.05 were considered significant.
Cilostazol Inhibits Oxidative Stress–Induced Premature Senescence in Human Endothelial Cells
To investigate the effect of cilostazol on the senescent phenotype in HUVECs, we induced premature endothelial senescence by addition of H2O2 100 μmol/L for 1 hour. We found that treatment with cilostazol inhibited the senescent phenotype as judged by SA-βgal assay and enlarged and flattened cell morphological appearance at 10 days. Under treatment with H2O2, 93% of cells were SA-βgal positive, versus only 46% of cilostazol (100 μmol/L)-treated cells under the same oxidative conditions (Figure 1A). We found that HUVECs treated with other cAMP-elevating agents showed a reduction of SA-βgal–positive cells as well (forskolin 1 μmol/L; 51%, rolipram 100 μmol/L; 53%). Treatment with cilostazol decreased the specific senescent morphological changes (Figure 1A). Expression of PAI-1 was decreased by treatment with cilostazol (Figure 1A). Treatment with cilostazol restored the rate of BrdU incorporation in prematurely senescent HUVECs (supplemental Figure I, available online at http://atvb.ahajournals.org). In parallel with this, telomerase activity was increased by treatment with cilostazol (supplemental Figure I). Moreover, we examined the effect on cell growth for 12 days after treatment with vehicle, H2O2 and cilostazol. Addition of H2O2 decreased cell number of HUVECs and treatment with cilostazol recovered it (Figure 1B). p53 plays a pivotal role in cellular senescence. Therefore, we examined the expression and acetylation of p53 at Lys373/382, one of the critical targets of Sirt1. As shown in Figure 1B, we observed that H2O2 increased the expression and acetylation of p53, and treatment with cilostazol decreased the acetylation of p53.
Enhancement of cAMP Production and eNOS Activity Induced by Cilostazol
When HUVECs were treated with cilostazol, the cAMP level significantly increased in a concentration-dependent manner at cilostazol concentrations of 1 and 100 μmol/L (data not shown). In the presence of H2O2, cilostazol increased eNOS activity (Figure 2A), expression of eNOS, and the phosphorylation of eNOS at Ser1177 in parallel with the phosphrylation of Akt at Ser473 (Figure 2C). Although exposure to H2O2 affected the total amount of eNOS and Akt, treatment with cilostazol reverted their expression to nearly normal levels (Figure 2C). To investigate the effect of NO on the senescent phenotype in HUVECs, we treated these cells with an NO donor, DETA-NO (100 μmol/L). DETA-NO–treated HUVECs showed decreased SA-βgal–positive cells (Figure 2B), and an increased rate of BrdU incorporation and telomerase activity (supplemental Figure I). These results suggest that the protective effect against a senescent phenotype may be attributed to an increased of NO via eNOS activation by cilostazol.
Treatment With Cilostazol Increased Sirt1 Expression
To explore the mechanism by which cilostazol prevents from premature endothelial senescence, we considered that an increase in NO production could promote the longevity gene, Sirt1. We found that cilostazol significantly increased Sirt1 mRNA and protein in a concentration-dependent manner for 10 days after treatment with H2O2 (Figure 3A). In contrast, Sirt1 mRNA and protein were not altered in the absence of H2O2 treatment (data not shown). To determine whether the expression of Sirt1 was regulated by the increase in NO production, we exposed prematurely senescent HUVECs to either an NO donor (such as DETA-NO or SNAP), a cAMP analog (8 Br-cAMP), or a cGMP analog (8 Br-cGMP). After these treatments, the expression of Sirt1 protein was markedly higher than that in untreated cells (Figure 3B). Furthermore, treatment with an NOS inhibitor, L-NAME, decreased Sirt1 expression (Figure 3C). To clarify the molecular mechanisms by which cilostazol induces SIRT1 expression, we examined the effect of protein kinase inhibitors on the cilostazol-induced phosphrylation of eNOS, Akt and expression of Sirt1 (supplemental Figure II). In the absence of H2O2 treatment, PKAI and LY294002 inhibited the cilostazol-induced phosphorylation of eNOS at Ser1177. The cilostazol-induced phosphorylation of Akt at Ser473 was inhibited by LY294002, however the inhibition by PKAI was not significant. Sirt1 expression was not altered by treatment with PKAI or LY294002. In the presence of H2O2 treatment, PKAI and LY294002 showed the similar effect on the cilostazol-induced phosphorylation of eNOS at Ser1177 and Akt at Ser473, but Sirt1 expression was significantly decreased.
Cilostazol Dose Not Have a Function of Direct Scavenger of Hydrogen Peroxide
It is possible that cilostazol may function as an antioxidant drug. Therefore, we examined the effect of NAC, another antioxidant, on Sirt1 expression, phosphorylation of Akt, and NOS activity as well as senescence markers. As shown in Figure 4A, treatment with NAC (0, 3, 5mmol/L) significantly decreased SA-βgal activity. Phosphorylation of Akt and NOS activity was increased by treatment with NAC (Figure 4B and 4C). However, Sirt1 expression was not altered (Figure 4C). To clarify the effect of cilostazol or NAC on H2O2, we examined whether cilostazol or NAC could scavenge H2O2 radicals. We performed a cell-free, horseradish peroxidase–coupled oxidation analysis.18 We observed that NAC scavenged H2O2 significantly, but cilostazol not (supplemental Figure III). These results indicate that inhibiting H2O2-induced senescence by cilostazol may not be attributable to its direct antioxidative effect such as NAC.
Inhibition of Sirt1 Abrogates the Protective Effect of Cilostazol Against Premature Senescence
To determine the role of endogenous Sirt1 in premature senescence, HUVECs were treated with a Sirt1 chemical inhibitor, sirtinol, a physiological Sirt1 inhibitor, NAM or Sirt1 siRNA. Knockdown of Sirt1 with siRNA was confirmed by Western blotting. As shown in Figure 5A and 5B, Sirt1 inhibition abrogates the effect of cilostazol on SA-βgal activity and the senescent specific morphological changes. Likewise, we found that Sirt1 inhibition had a similar effect to DETA-NO treatment (data not shown). Increased phosphorylation of eNOS at Ser1177 and decreased expression of PAI-1 by cilostazol were no longer observed when Sirt1 was inhibited (Figure 5C). These results indicate that Sirt1 could play an important role in the protective effect of cilostazol against a senescent phenotype.
Administration of Cilostazol Inhibits Vascular Endothelial Senescence Induced by Oxidative Stress in BALB/c Mice
To investigate whether cilostazol has a protective effect on vascular endothelial senescence induced by oxidative stress in vivo, we administrated paraquat, a herbicide that generates superoxide, to BALB/c mice. We performed resection of the thoracic artery of these mice and compared the senescent phenotype in the presence and absence of cilostazol. The number of SA-βgal–staining cells was significantly increased in untreated thoracic arteries, but was decreased in cilostazol-treated thoracic arteries (Figure 6A and 6B). Cross-sections of arteries stained with SA-βgal showed that positive cells were mostly located on the luminal surface and stained for CD-31, indicating that blue staining originated from vascular endothelial cells and not from the extracellular matrix (supplemental Figure IV). To estimate the degree of DNA damage caused by paraquat, we measured urinary 8-OhdG, a marker of DNA damage from oxidative stress. Urinary 8-OHdG level was decreased after cilostazol treatment (supplemental Figure IV). Immunostaining of the sections for Sirt1 showed that Sirt1 expression was increased in aortic endothelial cells by treatment with cilostazol (Figure 6C). To estimate the antioxidative effect of cilostazol on vasculature, we used DCF, cell-permeable fluorogenic probe, to measure ROS within cells by detection of enzymatically formed H2O2. The intensity of green fluorescence indicating DCF-positive cells was markedly increased in untreated thoracic arteries, which was decreased in cilostazol-treated thoracic arteries (supplemental Figure IV). The number of DCF-positive endothelial cells was decreased in cilostazol-treated thoracic arteries (supplemental Figure IV). Next, we used STZ diabetic mice in which the endothelial senescence documented.19 The treatment with cilostazol decreased SA-βgal–positive endothelial cells (supplemental Figure V).
As previously reported,20 the concentration of cilostazol (60 mg/kg/d) we administrated in this study was within clinical relevance. In vitro experiments, we used cilostazol at 0 to 100 μmol/L and confirmed a concentration-dependent trend. Given the average plasma concentration of cilostazol orally administered to humans (100 mg/body/d) is about 2 to 10 μmol/L and may be partially higher in our body, our used concentration of cilostazol is comparable to clinical.
Previous studies have shown that overexpression of Sirt1 antagonizes cellular senescence through acetylation of p53 with localization of the PML body.13 Recently, we reported that Sirt1 overexpression prevented the development of oxidative stress–induced premature senescence in human endothelial cells.21 Senescence of endothelial cells leads to endothelial dysfunction and may result in advanced atherosclerotic lesions.2 In fact, it has been reported that endothelial cells in samples of human aortia with atherosclerosis exhibited a senescence-like phenotype, increased expression of PAI-1,22 and decreased production of NO.1 NO production and eNOS expression are severely limited in senescent endothelial cells.23 Although NO is known to be involved in reducing oxidative stress and the progression of atheroscrerosis, the present study suggested that the NO-mediated prevention of premature senescence was attributable to Sirt1 function. These findings implicate the NO-Sirt1 axis as one of the fundamental determinants of endothelial senescence, and the role of Sirt1 as a driver of cellular stress resistance and longevity is noteworthy in the context of its expression profile.
The free-radical theory of aging proposes that degenerative senescence is largely the result of the cumulative effect of ROS. In this study, we used paraquat mice as an oxidative stress model. Moreover, we studied the effect of cilostazol on endothelial senescence used by STZ-diabetic mice as more suitable for clinical settings. In addition, Takase et al recently reported that cilostazol had exhibited an antiatherosclerotic effect on vasculature in ApoE-deficient mice.24 Therefore, we suggest that cilostazol has a beneficial effect on vasculature in clinical settings.
Cilostazol-induced NO production by eNOS activation via a cAMP/PKA- and PI3K/Akt-dependent mechanism was previously confirmed in the porcine thoracic aorta by an ESR25 technique and in clinical practice by endothelial-dependent vasodilation.26 Our results showed that cilostazol phosphorylated Akt via a PKA-independent mechanism (supplemental Figure II). It is suggested that both cAMP/PKA and PI3K/Akt signaling pathways are involved in cilostazol-induced phosphorylation of eNOS, however the contribution of these signaling to the upregulation of Sirt1 in the presence of H2O2 is more critical than that of in the absence of H2O2. Therefore, we suggest that upregulation of Sirt1 by cilostazol is modulated via cAMP/PKA, PI3K/Akt, and eNOS-dependent mechanism under oxidative conditions, but further investigation is needed to elucidate why there is discrepancy of Sirt1 expression.
A recent study showed DETA-NO, an NO-donor, and eNOS transfection activated hTERT and delayed endothelial senescence, indicating that eNOS has an antiatherosclerotic effect even in cases of advanced atherosclerosis. It is therefore suggested that incrased NO bioavailability by other pharmaceutical products such as 3-hydroxy-3-methylglutaryl (HMG)-coenzyme A (CoA) reductase inhibitors or agents with phytoestrogenic properties such as resveratrol may exert a protective effect against endothelial senescence, and this possibility deserves further investigation.
In summary, we showed that cilostazol inhibited oxidative stress–induced premature senescence, and subsequently enhancement of Sirt1 expression played a critical role in inhibition of a senescent phenotype in human endothelial cells. Our results suggest that NO production by cilostazol has a protective effect against endothelial senescence and dysfunction.
Sources of Funding
This work was supported by a Grant-in-Aid for Scientific Research from the Ministry of Education, Science, Culture and Sports of Japan (18590801).
Original received February 4, 2008; final version accepted May 28, 2008.
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