J Zheng and J Fang contributed equally to this work.
Leptin protects cardiomyocytes from serum-deprivation-induced apoptosis by increasing anti-oxidant defence
Article first published online: 14 JUN 2010
© 2010 The Authors. Clinical and Experimental Pharmacology and Physiology © 2010 Blackwell Publishing Asia Pty Ltd
Clinical and Experimental Pharmacology and Physiology
Volume 37, Issue 10, pages 955–962, October 2010
How to Cite
Zheng, J., Fang, J., Yin, Y.-J., Wang, X.-C., Ren, A.-J., Bai, J., Sun, X.-J., Yuan, W.-J. and Lin, L. (2010), Leptin protects cardiomyocytes from serum-deprivation-induced apoptosis by increasing anti-oxidant defence. Clinical and Experimental Pharmacology and Physiology, 37: 955–962. doi: 10.1111/j.1440-1681.2010.05415.x
See editorial commentary on page 953
- Issue published online: 14 JUN 2010
- Article first published online: 14 JUN 2010
- Accepted manuscript online: 14 JUN 2010 12:00AM EST
- Received 15 April 2009; revision 4 June 2010; accepted 7 June 2010.
- superoxide dismutase
1. Leptin, an important adipose-derived hormone, can be associated with cardiac pathophysiology; however, the role of leptin in cardiomyocyte apoptosis is poorly understood. The present study examines serum-deprivation-induced apoptosis in primary cultured cardiomyocytes treated with leptin.
2. Cardiomyocytes were subjected to serum deprivation in the presence or absence of leptin (5 or 50 nmol/L) for 48 h. Apoptosis was determined by Hoechst 33258 and Annexin V-FITC/propidium iodide dual staining. Cell viability, malondialdehyde (MDA) content, caspase 3 activation, and the expression and enzyme activity of superoxide dismutase (SOD) were measured. Small interference RNA (siRNA) targeting SOD1 and SOD2 were used to knockdown their expression and measure apoptosis.
3. Serum deprivation caused nearly 30% of apoptosis in cardiomyocytes, and an approximately 60% decrease in cell viability. The mRNA levels and the activated form of caspase 3 were greatly increased. In the presence of leptin, the apoptotic rate was reduced to approximately 15%, cell viability was increased and the activation of caspase 3 was partially inhibited. Additionally, the augmented lipid peroxidation (MDA formation) was abolished, and the impaired activities of SOD1 and SOD2 were restored by leptin. The mRNA expression of SOD2, but not SOD1, was stimulated by leptin. Transfection with siRNA that cause deficiency of either SOD1 or SOD2 attenuated the anti-apoptotic effects of leptin.
4. The results suggest that leptin inhibits serum-deprivation-induced apoptosis in cardiomyocytes by activating SOD. The present study outlines the direct actions of leptin in cardiac disorders that are related to elevated leptin levels.
Cardiomyocyte apoptosis has been well-documented as having a significant role in cardiomyocyte loss and the development of heart disease.1 Owing to the highly organized and regulated nature of the apoptotic process, apoptosis is believed to be amenable to therapeutic manipulation.2 A growing number of studies focus on understanding the hormones and cytokines associated with apoptosis. Among them are adipokines, the signalling molecules secreted by adipose tissue, such as tumour necrosis factor-α (TNF-α), interleukin-6, adiponectin, leptin, resistin, and so on. With the rapid increase in the prevalence of obesity, adipokines have attracted research attention as an important mechanistic link between obesity and various complications including cardiac dysfunction.3,4 The complex role of adipokines in cardiomyocyte apoptosis is still to be fully characterized.
Leptin, the protein product of the obese (ob) gene, was originally considered to be an adipose-derived hormone.5 Recent studies, however, have shown that leptin is produced by a variety of tissues including the heart.6 Plasma leptin concentrations are elevated in patients with heart disease, such as chronic heart failure7,8 and ischemic heart disease,9 showing a potential relationship between leptin and cardiac pathophysiology. Leptin exerts its biological effects through a family of membrane-bound receptors termed Ob-R.10 The expression of Ob-R in cardiac tissue has been confirmed,6,11,12 suggesting direct effects of leptin on the heart. In fact, a wide range of direct cardiac effects has been experimentally shown for leptin. Leptin stimulated fatty acid oxidation and lowered triglyceride content, without affecting glucose oxidation, in working perfused rat hearts.13 In isolated ventricular myocytes, leptin produced negative inotropic effects through a nitric-oxide-dependent pathway,14 as well as through JAK/STAT and MAP kinase signalling.15 In addition, leptin was shown to induce hypertrophy in rat neonatal cardiomyocytes,16–18 and to enhance proliferation of human pediatric ventricular myocytes and a murine HL-1 cell line.19
In terms of apoptosis, leptin is known to induce apoptosis in adipose tissue,20 but is also reported to inhibit apoptosis in other tissues, such as pancreatic islets21 and cardiac tissue.22 High levels of cardiac apoptosis were observed in leptin-deficient ob/ob and leptin-resistant db/db obese mice. The increased apoptosis in ob/ob, but not db/db, mice was reduced towards normal levels by leptin repletion, suggesting an anti-apoptotic role for leptin.22 In the rat myoblast cell line, H9c2 a short-term (1 h), but not long-term (24 h), pretreatment with leptin attenuated the subsequent apoptosis induced by H2O2.23 Based on these findings, the present study examined whether pathophysiological concentrations of leptin have anti-apoptotic effects on primary cultured rat neonatal cardiomyocytes under serum-deprived conditions, and investigated the underlying mechanisms.
Cardiomyocytes from 1- to 3-day-old Sprague–Dawley rats were prepared as we described in previous literature.24,25 Briefly, the ventricles were digested with 0.2% trypsin (Amresco, Solon, OH, USA). Cells were filtered through a nylon mesh, purified by Percoll gradient, resuspended in Dulbecco’s Modified Eagle Medium (DMEM; Sigma-Aldrich, St. Louis, MO, USA) containing 10% fetal calf serum and 10% neonatal calf serum, preplated for 60 min, seeded at a concentration of 5 × 105 cells/mL, and cultured in 95% air/5% CO2 in a humidified incubator at 37°C.
The protocol was in accordance with the institutional guidelines of Second Military Medical University, China, and the Guide for Care and Use of Laboratory Animals published by the US NIH (publication No. 96-01).
After 3–4 days of culture, the cardiomyocytes were synchronized in serum-free medium for 24 h, and then continually cultured for 48 h in serum-containing medium (the Serum group), or serum-free medium (the SF group), or serum-free medium containing recombinant rat leptin (Cat. No. 003-17; Phoenix Pharmaceuticals, Burlingame, CA, USA) at a final concentration of 5 or 50 nmol/L (the SF + leptin group). Finally, the cells and media were harvested for measurement.
In the small interference RNA (siRNA) experiments, the cardiomyocytes were synchronized in serum-free medium for 12 h before the addition of siRNA. After 6 h incubation with siRNA, the medium was changed and the cardiomyocytes were incubated with fresh serum-free medium for another 6 h, and then leptin was administered and the cells were incubated for a further 48 h. All siRNA were custom-synthesized by Shanghai GenePharma, Shanghai, China. They were administered at a final concentration of 100 nmol/L with the aid of transfection agents (Cat. No. AM4511; Applied Biosystems, Foster City, CA, USA). Mock-transfection (transfection without siRNA) and negative control siRNA (sense: 5′-UUCUCCGAACGUGUCACGUTT-3′, antisense: 5′-ACGUGACACGUUCGGAGAATT-3′) were used as controls. Optimal sequences of siRNA against SOD1 (sense: 5′-CCAUUAAACUGUAAUCUUATT-3′, antisense: 5′-UAAGAUUACAGUUUAAUGGTT-3′) and SOD2 (sense: 5′-CCAUUAAUUGUGUAUCUCATT-3′, antisense: 5′-UGAGAUACACAAUUAAUGGTG-3′) were screened out during preliminary experiments.
Analysis of cell apoptosis
Cell apoptosis was detected by Hoechst 33258 dye staining (Hoechst 33258 kit; Beyotime, Jiangsu, China) and Annexin V-FITC/propidium iodide dual staining (Annexin V-FITC kit; Bender MedSystems, Burlingame, CA, USA). Both staining procedures were carried out in accordance with the manufacturers’ instructions.
Hoechst 33258, a membrane-permeable DNA dye, stains the nucleus by chelating DNA and, thus, differentiates a normal nucleus from one that has condensed as a result of apoptosis. In the Hoechst staining procedure, cultured cardiomyocytes were fixed with paraformaldehyde, stained with Hoechst 33258, and examined under an Olympus fluorescence microscope. Apoptotic cells were identified based on nuclear condensation and fragmentation, and the percentage of apoptotic cells compared with total cells was calculated.
Annexin V-FITC/PI dual staining can discriminate intact cells (Annexin V−/PI−), apoptotic/early apoptotic cells (Annexin V+/PI−), and necrotic/late apoptotic cells (Annexin V+/PI+). In this staining method, Annexin V-FITC identifies apoptotic cells based on its high affinity for phosphatidylserine (PS), which is translocated from the inner to the outer leaflet of the plasma membrane in early apoptosis.26,27 PI distinguishes viable from non-viable cells. Viable cells with intact membranes exclude PI, whereas the membranes of dead and damaged cells become increasingly permeable to PI. To carry out this staining procedure, the cultured cardiomyocytes were digested with trypsin, washed, dual-stained with Annexin V-FITC and PI, and then analyzed by flow cytometry.
Analysis of caspase 3 expression and activity
Caspase 3 mRNA levels were measured by two-step reverse transcription polymerase chain reaction (RT-PCR). Total RNA extracted from the cardiomyocytes was used to generate cDNA by reverse transcription and processed by quantitative RT–PCR with SYBR Green as the dsDNA binding dye (Qiagen, Shanghai, China). PCR reactions were carried out in a Rotor-Gene 3000 real-time DNA detection system (Corbett Research, Sydney, Australia). β-actin was used as an internal control, and the standard curve method was used to calculate the relative expression levels of caspase 3 mRNA. The PCR primers were: 5′-CATGGCCCTGAAATACGAAGTC-3′ (forward) and 5′-GCAGGCCTGAATGATGAAGAGTTT-3′ (reverse) for caspase 3 (accession number: NM_012922), and 5′-ATGGTGGGTATGGGTCAGAAG-3′ (forward) and 5′-TGGCTGGGGTGTTGAAGGTC-3′ (reverse) for β-actin (accession number: NM_031144).
The activation of caspase 3 was determined by western blot, using the specific monoclonal antibody against the large fragment (17/19 kDa) of activated caspase 3 resulting from cleavage adjacent to Asp175 (Cat. No. 9664, Cell Signaling Technology, Danvers, MA, USA). Cardiomyocytes were lysed with cell lysis buffer and sonicated on ice. Protein concentration was determined using a bicinchoninic acid kit (Pierce, Rockford, IL, USA). Ten micrograms of total proteins were electrophoresed down a 10% SDS-PAGE gel, transferred onto nitrocellulose membrane and blocked with 5% non-fat dried milk. Then the membrane was probed with the primary antibody (1 : 1000), followed by the peroxidase-conjugated secondary antibody, and the protein signal was visualized by chemiluminescence reagents. The amount of cleaved caspase 3 was quantified by densitometry with a FR-200 analysis system (Fu-Ri Technology, Shanghai, China) and normalized to β-actin, an internal standard. The protein levels of treated cells were expressed as percentages relative to protein levels of control.
Determination of cell viability and malondialdehyde content
Cell viability was evaluated by 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyl tetrazonium bromide (MTT) assay based on the formation of formazan from MTT by metabolically active cells, as described in previous literature.24,25,28 MTT (Amresco) was added to the medium to a final concentration of 0.5 mg/mL for the last 12 h of the experiment. The resulting formazan crystals were dissolved in dimethyl sulfoxide, and the absorbance was measured on a microtitre plate reader (Bio-Rad 550, Hercules, CA, USA). The background absorbance (reagents without cells) was subtracted. Cell viability was presented as percentage absorbance, relative to the control absorbance.
To evaluate oxidative damage, the malondialdehyde (MDA) content in culture supernatant was measured using a commercial kit (Jian-Cheng Biochemical Engineering, Nanjing, China).
Measurement of superoxide dismutase activity and mRNA levels
The superoxide dismutase (SOD) activity in the cell lysate was measured using a commercial kit (Jian-Cheng Biochemical Engineering Co.). SOD1 (Cu/Zn-SOD) and SOD2 (Mn-SOD) activity were identified according to the manufacturer’s instructions.
SOD1 and SOD2 mRNA levels were measured by two-step RT–PCR as described earlier for caspase 3. The sequences of PCR primers were: 5′-CGGATGAAGAGAGGCATGTTG-3′ (forward) and 5′-CAATGATGGAATGCTCTCCTGAG-3′ (reverse) for SOD1 (accession number: NM_017050), and 5′-TAACGCGCAGATCATGCAG-3′ (forward) and 5′-AACCTGAGTTGTAACATCTCCCTTG-3′ (reverse) for SOD2 (accession number: NM_017051).
All data were expressed as means ± SEM. The data from the siRNA experiments were analyzed with two-way anova to examine the significances of leptin, SOD siRNA and leptin × SOD siRNA interaction. All the other data were analyzed with one-way anova followed by the Student–Newman–Keuls test to determine the differences between groups. SAS 9.0 statistical software (SAS Institute, Cary, NC, USA) was used for data analysis. Two-tailed P < 0.05 was considered statistically significant.
Leptin reduced apoptosis in myocytes deprived of serum
The apoptotic events were measured by observing nuclear changes using a DNA dye, Hoechst 33258, and by detecting PS externalization, a feature of plasma membrane changes in early apoptosis, using the PS-binding protein, Annexin V.
As shown by the nuclear staining with Hoechst 33258 (Fig. 1a), < 10% of the nuclei in serum-containing culture manifested apoptotic characteristics including condensation and fragmentation. In contrast, a greater percentage of apoptotic nuclei was observed in serum-free culture (25.8 ± 1.3%vs 7.2 ± 1.1%, P < 0.01). The presence of a low concentration (5 nmol/L) of leptin slightly affected the apoptosis without statistical significance (21.4 ± 0.9%, P > 0.05 vs SF), whereas a high concentration (50 nmol/L) of leptin significantly decreased the percentage of apoptotic nuclei (13.6 ± 1.9%, P < 0.01 vs SF).
As shown by the Annexin V/PI dual staining (Fig. 1b), a small percentage (5.9 ± 1.6%) of myocytes cultured in serum-containing medium were stained positive for Annexin V-FITC and negative for PI, showing a small number of apoptotic cells under control conditions. In contrast, the percentage of apoptotic cells in serum-free culture greatly increased to 29.5 ± 0.7% (P < 0.01 vs Serum). Leptin at concentrations of 5 and 50 nmol/L significantly reduced the apoptotic cells to 23.0 ± 0.5% (P < 0.01 vs SF) and 17.5 ± 0.5% (P < 0.01 vs SF), respectively. Contrary to this, the proportion of viable cells (Annexin V−/PI−) was increased by leptin. As to the proportion of Annexin V+/PI+ and Annexin V+/PI− myocytes, which were either at the end-stage of apoptosis, were undergoing necrosis or were already dead, there were no differences between the groups.
Leptin inhibited caspase 3 activation in myocytes deprived of serum
As shown in Fig. 2, caspase 3 mRNA levels were increased by approximately 3.5-fold over a 48 h serum-free incubation, and the cleaved fragment of activated caspase 3 was increased to a greater extent (8.9 ± 0.8-fold over control). The treatment of 5 and 50 nmol/L leptin significantly decreased the caspase 3 mRNA level to 2.4 ± 0.3 and 1.6 ± 0.2-fold over control, and decreased the amount of cleaved caspase 3 to 7.2 ± 0.5 and 2.8 ± 0.5-fold over control, respectively. These results showed that leptin partially inhibited caspase 3 expression and activation during serum deprivation.
Leptin increased cell viability and decreased MDA content in myocytes deprived of serum
As determined by the MTT assay (Fig. 3a), 48 h serum-free culture significantly reduced myocyte viability to 36.6 ± 6.9% of control values (P < 0.01). The treatment with a relatively low concentration (5 nmol/L) of leptin had no impact on the viability of serum-deprived myocytes (42.4 ± 5.1% of control, P > 0.05 vs SF). In myocytes receiving the high concentration (50 nmol/L) of leptin, however, the viability after serum deprivation significantly increased to 67.7 ± 4.4% of control (P < 0.01 vs SF), showing the ability of leptin to increase myocyte viability under serum-deprived conditions. The presence of 50 nmol/L leptin in serum-containing culture also resulted in an increase of myocyte viability (data not shown).
MDA levels were measured to estimate the degree of lipid peroxidation. Compared with serum-containing culture, the supernatant MDA content in serum-free culture was approximately increased by twofold (4.2 ± 0.3 vs 1.9 ± 0.2 μmol/L, P < 0.01). The presence of 5 nmol/L leptin partially inhibited MDA production (3.4 ± 0.2 μmol/L, P < 0.05 vs SF), and 50 nmol/L leptin significantly reduced MDA content to control levels (2.1 ± 0.1 μmol/L, P > 0.05 vs Serum, P < 0.01 vs SF; Fig. 3b).
Leptin increased SOD activity in myocytes deprived of serum
The activity of SOD, a major anti-oxidant enzyme, was determined to estimate the resistance of myocytes to oxidative stress. A significant decline in total SOD activity was observed for the cell lysate of serum-free culture (14.0 ± 0.6 vs 22.2 ± 1.9 U/mg pro, P < 0.01; Fig. 4a). SOD1 and SOD2 activities were decreased in serum-free culture, as compared with the serum-containing control (SOD1: 10.2 ± 0.9 vs 15.9 ± 1.2 U/mg pro, P < 0.01; SOD2: 3.8 ± 0.5 vs 6.3 ± 0.9 U/mg pro, P < 0.01; Fig. 4b). The low concentration (5 nmol/L) of leptin did not affect either SOD1 (11.8 ± 0.6 U/mg pro, P > 0.05 vs SF) or SOD2 activity (5.1 ± 0.5 U/mg pro, P > 0.05 vs SF) under serum-free conditions. The high concentration (50 nmol/L) of leptin significantly elevated both the activity of SOD1 (12.3 ± 0.7 U/mg pro, P < 0.05 vs SF) and SOD2 (7.3 ± 0.6 U/mg pro, P < 0.01 vs SF). In particular, SOD2 and total SOD activity of serum-deprived cells are comparable to those of the serum-containing control treated with 50 nmol/L leptin (Fig. 4a,b).
In contrast to impaired SOD activity, the mRNA level of either SOD isoform did not change under serum-free conditions. The treatment of leptin significantly increased the mRNA expression of SOD2, but not SOD1. In the presence of 5 nmol/L and 50 nmol/L leptin, SOD2 mRNA levels significantly increased to 1.7 ± 0.2 and 2.3 ± 0.2-fold over control, respectively, over a 48 h incubation (Fig. 4c). The time-course analysis for 50 nmol/L leptin showed that SOD2 mRNA expression significantly increased after a 3 h incubation, peaked at 6–12 h, and remained high until 48 h. Also, the total SOD activity increased in a similar pattern. SOD1 mRNA, however, did not change under the treatment of 50 nmol/L leptin (Fig. 4d).
SOD deficiency attenuated the anti-apoptotic effects of leptin
To inhibit specifically the gene expression of SOD, cardiomyocytes were transfected with SOD siRNA for 6 h and recovered in ‘blank’ medium for 6 h before the addition of leptin. All siRNA were given at a final concentration of 100 nmol/L. As shown in Fig. 5a (n = 5 for each group), neither mock-transfection nor transfection with negative control siRNA (NC siRNA) affected the mRNA expression of SOD1 or SOD2. In contrast, SOD siRNA resulted in SOD deficiency. SOD1 and SOD2 siRNA significantly decreased SOD1 and SOD2 mRNA levels to 0.66 ± 0.05 and 0.49 ± 0.06 of the blank medium control group, respectively (P < 0.01). Correspondingly, SOD1 and SOD2 activities were decreased by approximately 25% and approximately 35%, respectively (data not shown). The siRNA-mediated inhibition of SOD expression was highly specific, based on the fact that the siRNA against one SOD isoform did not change the mRNA level of the other SOD isoform (Fig. 5a).
To determine the role of oxidative stress in our model of apoptosis, cells were treated with a cell-permeable SOD mimic Mn-TBAP (manganese (III) tetrakis-(4-benzoic acid) porphyrin, 0.1 mmol/L). The apoptotic rate significantly decreased to 14.3 ± 1.3% (P < 0.01 vs SF, Fig. 5b), showing the protective effect of increased SOD activity in this model.
The apoptotic rates of serum-deprived cardiomyocytes in the presence or absence of leptin and SOD siRNA are shown in Fig. 5b and Table 1. Two-way anova on apoptotic rates showed the significant main effects of leptin, SOD1/2 siRNA, and leptin × SOD1/2 siRNA interaction. On one hand, SOD1 and SOD2 siRNA both independently increased apoptosis. On the other hand, the significance of leptin × SOD1/2 siRNA interaction suggested an interference of SOD1/2 siRNA with leptin action. As shown in Table 1, the individual presence of either SOD1 or SOD2 siRNA caused more apoptosis in leptin-treated myocytes, suggesting that the deficiency of either SOD isoform attenuated the anti-apoptotic effects of leptin. It is indicated that the anti-apoptotic activity of leptin is dependent on SOD1 and SOD2.
|Group||Apoptotic cells (%)|
|SF (n = 10)||29.5 ± 0.7|
|SF + leptin 5 nmol/L (n = 10)||23.0 ± 0.5|
|SF + leptin 50 nmol/L (n = 10)||17.5 ± 0.5|
|SF + negative control siRNA (n = 6)||31.6 ± 0.7|
|SF + negative control siRNA + leptin 50 nmol/L (n = 6)||17.0 ± 0.8|
|SF + SOD1 siRNA (n = 5)||38.3 ± 0.8|
|SF + SOD1 siRNA + leptin 5 nmol/L (n = 9)||35.5 ± 0.3|
|SF + SOD1 siRNA + leptin 50 nmol/L (n = 11)||28.6 ± 0.4|
|SF + SOD2 siRNA (n = 5)||39.6 ± 0.9|
|SF + SOD2 siRNA + leptin 5 nmol/L (n = 9)||35.2 ± 0.4|
|SF + SOD2 siRNA + leptin 50 nmol/L (n = 11)||30.5 ± 0.5|
|Results of two-way ANOVA|
|Leptin||(2, 59) F = 312.4*|
|SOD1 siRNA||(2, 59) F = 301.5*|
|Leptin × SOD1 siRNA||(3, 59) F = 6.6*|
|Leptin||(2, 59) F = 270.4*|
|SOD2 siRNA||(2, 59) F = 333.8*|
|Leptin × SOD2 siRNA||(3, 59) F = 5.7*|
The present study showed that leptin protected cultured neonatal rat cardiomyocytes against serum-deprivation-induced apoptosis, which is associated with enhanced SOD activity and blunted caspase 3 activation. To our knowledge, we first showed that leptin increased SOD1 and SOD2 activity, and stimulated the expression of SOD2, but not SOD1, in serum-deprived myocytes. We also showed that the deficiency of either SOD1 or SOD2 attenuated the inhibition of apoptosis caused by leptin.
Serum deprivation, a component of ischemia in vivo, has been shown to induce apoptosis in cultured cardiomyocytes.29–31 We also reported apoptosis in serum-deprived cardiomyocytes in earlier work,24 and we found that the treatment of a cell-permeable SOD mimics Mn-TBAP reduced apoptosis in this model by approximately 52% (Fig. 5b), suggesting that oxidative stress contributes to serum-deprivation-induced apoptosis. In the present study, both Hoechst 33258 and Annexin V/PI staining showed the anti-apoptotic effects of leptin. Furthermore, enhanced viability, reduced MDA formation and enhanced SOD activity were observed in leptin-treated myocytes, supporting the anti-apoptotic effect of leptin by increasing antioxidant defence.
Previously, the anti-apoptotic effects of leptin were reported in the hearts of ob/ob and db/db obese mice,22 and in a rat cardiac cell line H9c2.23 Also, the cardioprotective effects of leptin were observed in perfused mice hearts, where leptin (10 nmol/L) given during reperfusion reduced infarct size.32 The study in obese mice, as aforementioned, showed increases of cardiac apoptosis resulting from leptin deficiency or non-functional leptin receptors, which was abolished by leptin treatment.22 The study in H9c2 cells showed that H2O2-induced apoptosis was attenuated by pre-incubating cells with 6 nmol/L leptin for 1 h, but not 24 h, before H2O2 exposure.23 Another study addressing hypoxic damage in terms of lactate dehydrogenase and creatine kinase release from cultured cardiomyocytes showed that pretreatment with a very high dose of leptin (3000 ng/mL (187 nmol/L)) for 5 h, but not 1 or 20 h, provided significant protection against the following hypoxia.33 In the aforementioned studies in cells, leptin was exclusively given before the harmful stimuli and its protection was strictly time-limited. The present study did not attempt to pretreat myocytes with leptin for different durations. Instead, the myocytes were continuously exposed to leptin during the period of serum deprivation. The present results show the direct anti-apoptotic activity of leptin, suggesting that leptin is a tropic factor for the survival of cardiomyocytes under conditions of hyperleptinemia, which occurs in patients with obesity34 and certain heart disease.7–9
To understand the involvement of SOD in leptin-mediated inhibition of apoptosis, we measured the mRNA expression and enzyme activity of SOD1 and SOD2, the major SOD isoforms in cytoplasm and mitochondria, respectively. The results showed that leptin enhanced SOD1 and SOD2 activity, and stimulated the mRNA expression of SOD2, but not SOD1. SOD activity does not seem to be strictly correlated to mRNA level, as shown by enhanced activity with unchanged mRNA level for SOD1, as well as moderately enhanced activity accompanied by a much higher mRNA level for SOD2. Undefined post-transcriptional and post-translational regulation are probably both involved in the response of SOD activity to leptin. Despite the discrepancy between SOD mRNA levels and activity, it is suggested that the ability of serum-deprived myocytes to scavenge oxidative radicals is enhanced by leptin in the present study. Furthermore, the role of SOD in leptin-mediated myocyte survival is supported by the inhibition of SOD siRNA on the anti-apoptotic effects of leptin. We used siRNA to knockdown specifically the expression of SOD1/2 and found that myocytes deficient in SOD were susceptible to apoptosis on serum deprivation. The anti-apoptotic activity of leptin was alleviated by the lack of either SOD1 or SOD2, suggesting that SOD1 and SOD2 contribute to leptin-induced apoptosis resistance. The effects of leptin are likely to be direct and specific for SOD. The time-course analysis showed early and persistent induction of SOD2 mRNA and total SOD activity by leptin (Fig. 4d), suggesting a direct effect of leptin on SOD. Besides, leptin failed to change the mRNA level of catalase (data not shown), supporting its specificity for SOD.
The present study showed predominant anti-oxidative activity of leptin, which is not in accordance with a recent study in human monocytes.35 The latter study incubated monocytes with comparable doses of leptin (1 and 10 nmol/L) for no more than 2 h, and found that total SOD activity was increased, but the production of oxidative species was overwhelming.35 This discrepancy with our data might be caused by differences in cell types and incubation time for leptin. In another study, primary cardiomyocytes were serum-deprived for 72 h, a much longer duration compared with the present study, and then incubated in serum-free medium containing similar doses of leptin (1–1000 ng/mL or 0.0625–62.5 nmol/L) for 4 or 72 h.17 Increases of reactive oxygen species (ROS) production were observed in that setting, whereas SOD activity was unknown. Despite all that, the myocytes receiving leptin experienced hypertrophy instead of apoptosis or necrosis, supporting the fact that leptin acted as a tropic factor. The present study did not measure ROS production. There is a possibility that SOD activation in response to leptin might be accompanied by ROS generation. Nevertheless, our results showed that leptin improves the balance between anti-oxidant and oxidative stress.
In summary, our results showed that the obesity-associated hormone, leptin, exerted direct anti-apoptotic effects in serum-deprived cardiomyocytes by relieving oxidative stress and inactivating the intrinsic apoptotic pathway, supporting a cardiac protective role of leptin against stressful stimuli. This study offers insights into the role of leptin in obesity-associated heart disease.
This study was supported by grants from the National Basic Research Program of China (No. 2006CB503807, 2009CB521902), the National Natural Science Foundation of China (No. 30870906, 30800375), the Pujiang Project of Shanghai, China (No. 08PJ14001) and the Project Sponsored by the Scientific Research Foundation for the Returned Overseas Chinese Scholars, State Education Ministry, China (No. (2008) 891).
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