Nicotine Promotes Cardiomyocyte Apoptosis via Oxidative Stress and Altered Apoptosis-Related Gene ExpressionZhou X.a · Sheng Y.a · Yang R.a · Kong X.a, b
aDepartment of Cardiology, The First Affiliated Hospital of Nanjing Medical University, Nanjing, and bDepartment of Cardiology, Jiangsu Shengze Hospital, Wujiang, PR China Corresponding Author
Department of Cardiology
The First Affiliated Hospital of Nanjing Medical University
No. 300 Guangzhou Road, Nanjing 210029 (PR China)
Tel./Fax +86 25 8367 2050, E-Mail firstname.lastname@example.org
Objective: To investigate the effect of nicotine on cardiomyocyte apoptosis in vitro and explore the potential mechanisms involved. Methods: The MTT assay was used to detect the viability of cultured cardiomyocytes exposed to different concentrations of nicotine (0.1–100 µM). Laser confocal microscopy, TUNEL assay and flow cytometry were utilized to detect cardiomyocyte apoptosis. Oxidative stress was evaluated by the levels of lactic dehydrogenase, malondialdehyde and superoxide dismutase in the supernatant of culture media. Real-time PCR was conducted to identify mRNA expression changes in apoptosis-related genes between the nicotine and the control group. Results: Nicotine was found to inhibit cardiomyocyte viability in a concentration-dependent manner. Our results demonstrated that nicotine can promote cardiomyocyte apoptosis and the antioxidant glutathione can protect cardiomyocytes from apoptosis via inhibition of nicotine-induced oxidative stress. Real-time PCR indicated that the expression of Bcl-2, Pax3, Bmp4 and Slug was down-regulated in the nicotine group, while the expression of P53, Bax and Msx1 was up-regulated. Conclusion: Nicotine promotes cardiomyocyte apoptosis by inducing oxidative stress and disrupting apoptosis-related gene expression.
© 2010 S. Karger AG, Basel
Smoking is a serious health concern that is associated with cancer and pulmonary and cardiocerebrovascular disease. Nicotine is the addictive component and most harmful ingredient contained within cigarettes. Different groups have investigated the effects of nicotine on apoptosis both in vitro and in vivo, and they have revealed a correlation between nicotine exposure and apoptosis. Several studies have reported that nicotine has a protective effect via inhibiting apoptosis [1,2,3,4,5,6,7], while others have shown the opposite [8,9,10,11,12]. Recent studies have shown that nicotine can increase oxidative stress, which may be associated with apoptosis [13,14,15,16].
Apoptosis, also termed programmed cell death, plays a critical role in the pathogenesis of a variety of cardiovascular diseases. It has been shown that apoptosis occurs in myocardial tissue samples from patients suffering myocardial infarction, dilated cardiomyopathy and end-stage heart failure [17,18,19]. Based on the above facts, we hypothesized that nicotine can induce cardiomyocyte apoptosis, which may be involved in the pathogenesis of nicotine-induced cardiovascular diseases. Induction of oxidative stress and altered expression of apoptosis-related genes such as P53, Bax, Bcl-2, Pax3, Bmp4, Slug and Msx1 may be responsible for cardiomyocyte apoptosis. The present study was conducted to determine the effect of nicotine on cardiomyocyte apoptosis in vitro and explore the potential mechanisms using cultured neonatal rat cardiomyocytes.
The experimental procedures were conducted in accordance with international guidelines and approved by the ethics committee of Nanjing Medical University. Rat hearts were surgically removed from 1- to 3-day-old Sprague-Dawley rats obtained from the Laboratory Animals Center of Nanjing Medical University, washed instantly in cold D-Hanks solution, and then minced into 1–3 mm3 pieces. Thereafter, the minced tissue was subjected to 6–8 cycles of proteolytic dissociation by magnetic stirring (10 min, 37°C) with 0.06% trypsin solution (Gibco, USA). Supernatants from each cycle were pooled and centrifuged. In the end, the cell pellet was resuspended in DMEM supplemented with 20% calf serum (Gibco, USA), 100 U/ml penicillin and 100 mg/ml streptomycin. Selective adhesion was performed after a 1.5 h incubation at 37°C in a humidified atmosphere (5% CO2 and 95% air) in order to obtain a high purity of cardiomyocytes. Subsequently, 0.1 mM bromodeoxyuridine (Sigma, USA) was added to the medium for the first 48 h of culture to inhibit the growth of fibroblasts.
Cardiomyocytes were randomly divided into 3 groups: cells cultured in normal culture media (the control group); cells treated with nicotine (the nicotine group) (Sigma, USA); cells treated with nicotine and antioxidant (the antioxidant group). Glutathione (250 µM; Sigma, USA) was used as the antioxidant in the present study, blocking the effect of oxidative stress on cultured cardiomyocytes.
Cardiomyocytes were exposed to different concentrations of nicotine (0.1–100 µM) for 48 h and cell viability was detected using the MTT assay according to the manufacturer’s instructions. Briefly, cells were plated at a density of 1 × 105/ml in a 96-well plate and incubated with 20 µl of 5 mg/ml MTT solution (Amresco, USA) for 4 h at 37°C following treatment with various concentrations of nicotine. Thereafter, 150 µl of DMSO (Amresco, USA) was added to each well to dissolve the dye crystal formazan and the plate was shaken for 10 min to make sure that all purple crystals were dissolved. The amount of MTT formazan was quantified by determining the absorbance at 490 nm using a microplate reader.
Cell slices were prepared by innoculating myocardial cells (1 × 106/l) onto a 6-hole board with coverslips, then they were stainedwith Annexin V and propidium iodide (PI) (BD Pharmingen) and observed using laser confocal microscopy. Normal cells could only give off a weak green fluorescence. In the early stages of apoptosis, the cell membranes were stained with Annexin V and gave off a vibrant green fluorescence, while the nucleolus was not stained with PI. The cells in the media and late stages of apoptosis were highly stained by both Annexin V and PI, therefore the cell membranes gave off green fluorescence and the nucleolus gave off a red fluorescence.
The TUNEL Apoptosis Detection Kit (KeyGEN, China) was used to detect apoptosis following the manufacturer’s instructions. Briefly, cells grown on coverslips were fixed with a freshly prepared fixation solution (4% paraformaldehyde in PBS, pH 7.4) for 1 h at RT and then incubated with Blocking solution (3% H2O2 in methanol) for 10 min at room temperature. After washing with PBS, cells were incubated in permeabilization solution containing 0.1% tritonX-100 and 0.1% sodium citrate for 2 min on ice. Thereafter, 50 µl of TUNEL reaction mixture (45 µl equilibration buffer, 1 µl biotin-11-dUTP and 4 µl TdT) was added and the coverslips were incubated for 60 min at 37°C in wet and dark atmospheres. 50 µl of Streptavidin-HRP solution was then added to the samples and the coverslips were incubated for 30 min at 37°C in wet and dark chambers. Finally, DAB substrate was added for coloration (10 min, room temperature) and the coverslips were observed under light microscopy. Apoptotic cells stained brown and normal cells purple-blue.
Oxidative stress was evaluated by detecting levels of lactic dehydrogenase (LDH), malondialdehyde (MDA) and superoxide dismutase (SOD) in the supernatant of culture media according to the instructions of the detection kit (Jian-Cheng, China). The increase in LDH leakage reflected the damage to the myocardial cell membrane by free radicals. The change in MDA content reflected the degree of lipid peroxidation in the cell membrane. The activity of SOD indirectly reflected the capacity to remove oxygen free radicals.
The Annexin V-FITC apoptosis detection kit (BD Pharmingen) was used to detect apoptosis according to the manufacturer’s instructions. Briefly, 1 × 105 cells were collected and washed twice with cold PBS. 5 µl of Annexin V and 5 µl of PI were added to the cells, which were resuspended in 500 µl 1× binding buffer. The cells were gently vortexed and incubated for 15 min at room temperature in the dark, then they were analyzed by flow cytometry within 1 h. Annexin V labeled with a fluorophore could identify cells in the early stage of apoptosis, and PI, a fluorescent nucleic acid binding dye, was responsible for staining cells in the medium and late stages of apoptosis. Analysis was based on gating a subpopulation of cells by forward scatter versus side scatter. The intermediate to large mononuclear cell population was the gated region used to calculate the apoptotic rate. The apoptotic rate was calculated as the percentage of Annexin V-positive and PI-negative cells divided by the total number of cells in the gated region.
Total RNA was isolated from cultured cardiomyocytes using TRIzol reagent (Invitrogen, Karlsruhe, Germany). Briefly 1.0 × 107 cardiac cells were homogenized in 1 ml Trizol and 200 µl of chloroform was added. After mixing, the samples were centrifuged at 12,000 g for 15 min (4°C), and the upper aqueous phase was transferred to a tube containing an equal volume of isopropanol, mixed thoroughly using a vortex device and centrifuged at 12,000 g for 10 min (4°C). The supernatant was discarded and the precipitated RNA pellet was washed using 1 ml of 75% ethanol and centrifuged at 12,000 g for 5 min (4°C). The final pellet was allowed to air-dry for 5–10 min and was then resuspended in RNAse-free water. The concentration of RNA was quantified by measuring the optical density at 260 nm. The reverse transcription mixture contained 1 µg of total RNA, 0.5 µg of oligo d(T) primer, 4 µl of 5× RT buffer, 0.5 mM deoxynucleotides, 50 U of RNase inhibitor and 200 U of reverse transcriptase (Promega, USA) in a total volume of 20 µl. The reaction was carried out at 42°C for 1 h followed by heat inactivation at 95°C for 5 min. Real-time PCR was performed on the ABI 7300 Real Time PCR System following the manufacturer’s instructions. The reaction mixture consisted of 12.5 µl of SYBR Premix Ex Taq containing TaKaRa Ex Taq HS, dNTP mixture, Mg2+ and SYBR Green I (ABI, USA), 0.3 µM of primer, 1 µl template DNA and ddH2O q.s.’ed to 25 µl. The amplification reactions were carried out according to the following cycling protocol. Initial denaturation for 5 min at 95°C was followed by 40 cycles of 94°C for 30 s, 60°C for 30 s and 72°C for 40 s. The final elongation phase was performed for 10 min at 72°C. The primers used in this study are shown in table 1. The fold change of mRNA was analyzed using the 2–ΔΔCt method .
Data are presented as mean ± SD and differences between groups were analyzed using 1-way ANOVA with the SPSS 15.0 statistical package. Scheffé post hoc test was used if the ANOVA was significant. p < 0.05 was considered statistically significant.
The early morphology of cultured cardiomyocytes was observed using an inverted microscope. The newly separated cells were round, and gradually the cells became spindle-shaped. They gradually stretched and grew pseudopodia, with a presentation that was triangular and polygonal in shape (fig. 1). Almost all the cells adhered to the wall after 12 h, and some single cells presented spontaneous pulses with different frequencies and rhythm. After 72 h, pseudopodia of the cells were interwoven into a net, which gradually turned into cellular clusters that were in a radial array. With synchronized and strong pulses, some functional syncytial cells were formed, and the synchronized pulse frequency was about 60–120 times/min.
There were no significant differences in cell viability between the cardiomyocytes exposed to nicotine (0.1 and 1 µM) and the control; however, the viability of cardiomyocytes exposed at 10 and 100 µM was significantly different from the control (p < 0.05). Our findings suggest that nicotine inhibits cardiomyocyteviability in a concentration-dependent manner (table 2). The lowest effective concentration (10 µM) was chosen as the effective nicotine concentration for the nicotine group.
Cardiomyocytes grown on coverslips were stained with Annexin V/PI and observed using laser confocal microscopy. Normal cells in the control group could only give off weak green fluorescence. Some apoptotic cells could be observed in the nicotine group. In the early stages of apoptosis, the cells gave off a green fluorescence, while in the media and late stages of apoptosis, the cells gave off green and red fluorescence (fig. 2a, b).
Cardiomyocytes grown on coverslips were stained using the TUNEL assay and observed using light microscopy. Some brown apoptotic cells could be observed in the nicotine group, while only purple-blue cells could be observed in the control group (fig. 3a, b).
Oxidative stress was evaluated by detecting LDH, MDA and SOD from the supernatant of culture media of the different groups. Our results showed significant increases in LDH and MDA levels and a decrease in SOD activity in the nicotine group compared with the control group (p < 0.05). However, there was no significant difference between the antioxidant and control groups (table 3).
The results of flow cytometry demonstrated that the apoptotic rate of cardiomyocytes in the nicotine group increased significantly compared with that in the control group (p < 0.05), and the apoptotic rate in the antioxidant group declined remarkably in comparison with that in the nicotine group (p < 0.05; fig. 4a–c).
Our results indicated that mRNA expression of P53 and Bax was increased and Bcl-2 was decreased in the nicotine group (p < 0.05), which was in accordance with previous studies [21,22]. In addition, the expression of Pax3, Bmp4 and Slug were down-regulated in the nicotine group (p < 0.05) and the expression of Msx1 was up-regulated (p < 0.05; fig. 5).
The present study was designed to investigate the effect of nicotine on cardiomyocyte apoptosis in vitro and explore the potential mechanisms involved. The results of MTT demonstrated that nicotine inhibits cardiomyocyte viability in a concentration-dependent manner. In the present experiment, the lowest effective concentration (10 µM) detected by the MTT assay was chosen to be the concentration of nicotine used in our study, which is above what Benowitz et al.  have reported as the nicotine plasma levels in smokers. The findings from confocal microscopy, TUNEL assay and flow cytometry suggested that nicotine can promote cardiomyocyte apoptosis, which led us to question the molecular mechanisms of nicotine-induced apoptosis. We hereby confirmed that nicotine-induced oxidative stress and altered expression of apoptosis-related genes are responsible for cardiomyocyte apoptosis.
To evaluate oxidative stress, we measured LDH, MDA and SOD in the supernatant of culture media from all of the different groups. The change in leakage of LDH reflects damage to the myocardial cell membrane, and the increase in its content is usually regarded as the symbol of irreversible injury to cardiomyocytes . MDA is a product of lipid peroxidation, and its content can reflect the degree of lipid peroxidation in the cell membrane and the severity of damage to cells by free radicals . The activity of SOD, which is an important antioxidant enzyme, can indirectly reflect the capacity to remove oxygen free radicals . Our findings suggest that nicotine can induce oxidative stress and that the antioxidant glutathione can block this effect.
Flow cytometry was carried out to detect the apoptotic rate of cardiomyocytes in all of the different groups. The apoptotic rate in the nicotine group was increased significantly compared with that in the control group, while the apoptotic rate in the antioxidant group declined remarkably in comparison with that in the nicotine group. Taken together, nicotine can promote apoptosis and antioxidants can protect cardiomyocytes from apoptosis via inhibiting oxidative stress. These findings support our hypothesis that nicotine-induced oxidative stress may be responsible for cardiomyocyte apoptosis.
The results from the present study demonstrate that the expression of Bcl-2, Pax3, Bmp4 and Slug were down-regulated in the nicotine group and the expression of P53, Bax and Msx1 were up-regulated. The altered expression of these apoptosis-related genes may be responsible for cardiomyocyte apoptosis.
P53, a key tumor suppressor gene, plays a central role in cellular stress response pathways, regulating the expression of multiple genes involved in controlling transcription of apoptosis [27,28]. Bax and Bcl-2, members of the Bcl-2 family, are critical regulatory factors in response to apoptosis. Bax is the pro-apoptotic member and Bcl-2 is the anti-apoptotic member [29,30,31]. Pax3 is required for migration of the cardiac neural crest in the developing heart . A recent study has revealed that Pax3 is required for cardiac outflow tract septation since it blocks p53-dependent processes in the migration of cardiac neural crests . Bmp4, one of the members of the TGFβ superfamily, is a key signaling molecule involved in endothelial-mesenchymal transformation (EMT) during endocardial cushion morphogenesis [34,35]. BMP2/4 signaling in neural crest derivatives is essential for outflow tract development and may regulate a crucial proliferation signal for the ventricular myocardium . It has been suggested that BMP4 mediates apoptosis in both endocardial cushions and ventricular myocardium . Slug, a Snail family member, is involved in the process of EMT which is required for the migration of neural crest cells and the formation of cardiac valves and septa . Msx1, which is required for EMT of the atrioventricular cushions and patterning of the atrioventricular myocardium, has been indicated to regulate survival of secondary heart field precursors and post-migratory proliferation of cardiac neural crest in the outflow tract [39,40,41]. A recent study has reported that Slug and Msx1 lie upstream of the apoptotic factors Bax and Bcl-2 and that a balance between the anti-apoptotic activity of Slug and the apoptotic activity of Msx1 is required for the proper development of the neural crest .
In conclusion, nicotine can promote cardiomyocyte apoptosis by inducing oxidative stress and disrupting the expression of apoptosis-related genes.
This research was supported by the Department of Pathogenic Biology of Nanjing Medical University.
Department of Cardiology
The First Affiliated Hospital of Nanjing Medical University
No. 300 Guangzhou Road, Nanjing 210029 (PR China)
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