Spermidine alleviates cardiac aging by improving mitochondrial biogenesis and function

Exogenous spermidine supplementation restores polyamine metabolism and attenuates cardiac aging

To study polyamine metabolism in the myocardium of 3-month-old (young) and 24-month-old (old) rats, we assessed ODC and SSAT protein expression, as well as polyamine contents. The expression of ODC decreased, while the expression of SSAT increased, in older rats compared with younger ones (Figure 1A). Meanwhile, both spermine (SPM) and spermidine (SPD) contents were significantly reduced in the myocardium of older animals, which showed significant elevations in both polyamines after 6-week treatment with SPD (Figure 1B). To evaluate the potential anti-aging effects of SPD on myocardial tissue, three age-related parameters were assessed: 1) senescence-associated-β-galactosidase (SA-β-gal) activity, 2) accumulation of p21 and p16 proteins, and 3) changes in the ultrastructure of myocardium. We found that SA-β-gal staining and p21/p16 expression was increased in the hearts of older rats, and these effects were attenuated by SPD supplementation (Figure 1C and 1D). Abnormalities in the left ventricular ultrastructure, including sarcomere deformation, lipofuscin deposition, decreased mitochondrial matrix density, mitochondrial cristae disorganization, and inner mitochondrial membrane damage, were also evident in aged rats after electron microscopy evaluation. However, following SPD treatment, sarcomere structure was clear, and the mitochondria were tightly packed between the myofibrils and had intact outer and inner membranes with distinct cristae (Figure 1E). These data showed that in aging hearts SPM and SPD contents are reduced, denoting impaired polyamine metabolism, whereas SPD supplementation markedly protected against heart senescence through replenishment of the cardiac total polyamine pool.

Figure 1. Age-dependent changes in polyamine metabolism and effect of spermidine on cardiac aging in rats. (A) Representative immunoblot bands for ODC and SSAT and quantification of protein levels in the myocardium of 3-month-old (Young) and 24-month-old (Old) rats. GAPDH was used as loading control. (B) Polyamine content, including putrescine (PU), spermidine (SPD), and spermine (SP) in the myocardium of young, old, and SPD-treated old rats (SPD group) evaluated by high-performance liquid chromatography. (C) Representative immunoblot bands for p21 and p16 and quantification of protein levels. GAPDH was used as loading control. (D) Positive area of senescence-associated β-galactosidase (SA-β-gal) staining. Scale bars: 20 μm. (E) Representative transmission electron microscopy images showing ultrastructural changes in the myocardia. Quantification of the area of cells occupied by mitochondria (%); Magnification: ×10,000. n = 6 for each group. ^# P < 0.05 vs. young control, ^## P < 0.01 vs. young control, ^* P < 0.05 vs. old.

Exogenous spermidine supplementation protects against H₂O₂-induced cardiomyocyte aging

To evaluate the effect of SPD on cardiomyocyte aging, we induced senescence in primary neonatal rat cardiomyocytes (NRCMs) and H9C2 myoblasts by treatment with 40 μmol/L H₂O₂ for 4 h (see Supplementary Material for optimization protocol). After H₂O₂ treatment, the expression of ODC decreased, and the expression of SSAT increased significantly compared with control cells (Figure 2A). Next, senescent NRCMs were treated with different doses of SPD (0–100 μmol/L) for 48 h. Results showed that SPD increased cell viability in a dose-dependent manner (Figure 2B). However, a decrease in cell viability was observed with SPD concentrations ≥20 μmol/L. We verified that after treatment with 40 μmol/L H₂O₂, supplementation with 10 μmol/L SPD for 6, 12, 24, 36, and 48 h increased cell viability in a time-dependent manner (Figure 2C). We further observed that the SA-β-gal–positive staining area in both NRCMs and H9C2 cells (Figure 2D), as well as p21 and p16 expression in NRCMs (Figure 2E), were significantly increased after senescence induction with H₂O₂; these changes were in turn prevented by supplementation with 10 μmol/L SPD. These data indicated that an imbalance in polyamine metabolism occurs in cardiomyocytes after H₂O₂-induced aging, and this could be ameliorated by exogenous SPD supplementation.

Figure 2. Expression changes in ODC and SSAT and effect of SPD in H₂O_2- treated cardiomyocytes. (A) Western blot analysis of ODC and SSAT expression in NRCMs treated with or without 40 μM H₂O₂. GAPDH was used as loading control (n = 4). (B, C) Concentration- and time-dependent effect of SPD on the viability of NRCMs treated with 40 μM H₂O₂ (n = 8). (D) SA-β-gal staining in NRCMs (top) and H9C2 cells (bottom). The positive area of SA-β-gal staining is shown in the right-hand graph (n = 8). Scale bars: 20 μm. (E) Representative immunoblot bands for p21 and p16, and quantification of protein levels in NRCMs (n = 4). ^# P < 0.05 vs. control, ^## P < 0.01 vs. control, ^* P < 0.05 vs. H₂O₂, ^** P < 0.01 vs. H₂O₂.

Spermidine attenuates cardiac senescence via inhibiting ROS accumulation and improving mitochondrial function

To confirm the mitochondrial-protective action of SPD, myocardial mitochondrial oxygen consumption and anti-oxidative ability were measured in isolated mitochondria after SPD administration in vivo. Compared with the young heart, the aged myocardium showed decreases in mitochondrial State 3 respiration, respiratory control ratio (RCR), and P/O ratio, and an increase in proton leakage, whereas exogenous SPD supplementation reversed these defects (Figure 3A, a–d). In addition, SPD supplementation reversed the decrease in superoxide dismutase (SOD) and catalase (CAT) expression and activity observed in the aging myocardium (Figure 3B a–d). Mitochondria are responsible for producing ATP, the main cellular energy molecule, in a process that also generates small amounts of reactive oxygen species (ROS). Therefore, ATP and ROS production, and mitochondrial membrane integrity are important indicators of mitochondrial function. We found that ATP levels decreased in H₂O₂-treated NRMCs, and SPD supplementation inhibited such decrease (Figure 3C). In addition, SPD treatment attenuated oxidative stress in H₂O₂-treated H9C2 cells, evidenced by a significant reduction in the fluorescence intensity of the mitochondria-specific superoxide indicator triphenylphosphonium-linked hydroethidium (mitoSOX) (Figure 3D) and the intracellular superoxide indicator dihydroethidium (DHE) (Figure 3E). Mitochondrial transmembrane potential (ΔΨm) was next monitored by the potentiometric dye tetramethylrhodamine ethyl ester (TMRE). In H9C2 cells, ΔΨm was significantly decreased by H₂O₂; however, after addition of SPD, ΔΨm remained intact (Figure 3F). In summary, these findings indicate that SPD alleviates aging-associated mitochondrial damage in vivo and vitro through inhibiting oxidative stress.

Figure 3. Effect of SPD on mitochondrial respiration and ROS accumulation in the aged heart and in H₂O₂-treated cardiomyocytes. (A) Mitochondrial oxidative phosphorylation (OXPHOS) efficiency was evaluated in the rat myocardium. Measurements included mitochondrial oxygen consumption States 3 and 4 (a), respiratory control rate (RCR) (b), P/O ratio (c), and proton leakage (d). Respiration was induced with pyruvate/malate (5 mM each) as energizing substrates and ADP (200 μM) to initiate State 3 respiration (n = 8). (B) Western blot analysis of SOD and CAT expression (a, b), and colorimetric detection SOD and CAT activity (c, d). n = 4 for protein expression and n = 8 for activity assay. ^# P < 0.05 vs. young control, ^## P < 0.01 vs. young control; ^* P < 0.05 vs. old, ^** P < 0.01 vs. old. (C) ATP content of cardiomyocytes measured by luminometry in NRCMs (n = 8). (D) Superoxide production in mitochondria detected by MitoSOX staining in H9C2 cells. (E) ROS production in H9C2 cells detected by DHE in H9C2 cells. (F) Mitochondrial transmembrane potential (ΔΨm) detected by TMRE in H9C2 cells. Quantification of the mean fluorescence intensity of MitoSOX, DHE, and TMRE are displayed on the right side of the graphs (n = 8). ^# P < 0.05 vs. Control, ^## P < 0.01 vs. Control, ^* P < 0.05 vs. H₂O₂ group, ^** P < 0.01 vs. H₂O₂ group.

Spermidine stimulates mitochondrial biogenesis and function via activation of the SIRT1/PGC-1α signaling pathway

To evaluate potential correlations between polyamine metabolism and the SIRT1/PGC-1α signaling pathway, a key regulator of mitochondrial biogenesis, western blot was used to detect the expression of ODC, SSAT, p21, and members of the SIRT1/PGC-1α signaling pathway in cardiac tissue of rats aged 3, 6, 12, and 24 months. We found that the expression of SIRT1, PGC-1α, NRF1, NRF2, and TFAM decreased depending on age (Figure 4B). Age-dependent changes seemed also apparent for ODC, SSAT, and p21 expression, i.e. ODC decreased, while SSAT and p21 increased with age (Figure 4A). A strong negative correlation was observed between SSAT and PGC-1α, NRF1, NRF2, and TFAM expression (Figure 4C). In turn, a significantly positive correlation was seen between ODC and both SIRT1 and NRF1 (Figure 4D). Meanwhile, SPD administration increased cardiac expression of SIRT1 and other mitochondrial biosynthesis-related proteins in the oldest rats (Figure 4E).

Figure 4. SPD prevents age-associated depletion in myocardial polyamines and alterations of SIRT1/PGC-1α signaling pathway proteins. Representative immunoblot bands for ODC, SSAT, and p21 (A), and for SIRT1, PGC-1α, NRF1, NRF2, and TFAM (B) in myocardium from 3-, 6-, 12- and 24-month-old rats. GAPDH was used as loading control. (n = 10). ^* P < 0.05 vs. 3 months, ^** P < 0.01 vs. 3 months. (C) Correlation between ODC and (a) SIRT1, (b) PGC-1α, (c) NRF1, (d) NRF2, and (e) TFAM in cardiac tissue from rats of different ages. (D) Correlation between SSAT and (a) SIRT1, (b) PGC-1α, (c) NRF1, (d) NRF2, and (e) TFAM in cardiac tissue from rats of different ages (n = 10). (E) Representative immunoblot bands for SIRT1, PGC-1α, NRF1, NRF2, and TFAM in the myocardium of young (3 months of age), old (24 months of age), and SPD-treated (6 weeks) old rats. (n = 4). ^# P < 0.05 vs. young, ^## P < 0.01 vs. young, ^* P < 0.05 vs. old, ^** P < 0.01 vs. old. (F, G) Expression of SIRT1, PGC-1α, NRF1, NRF2, and TFAM measured by western blot in NRMCs treated with 0, 2.5, 5, 10, or 20 μM SPD for 24 h (F) or 10 μM SPD for 0, 6, 12, 24, and 48 h (G). Quantification of protein expression is shown on the right-hand side of the graphs (n = 4). ^* P < 0.05 vs. control, ^** P < 0.01 vs. control.

In vitro studies were also performed using NRCMs. We treated NRCMs with various doses of SPD (0-20 μmol/L) for 24 h after senescence induction with H₂O₂. SPD exposure (5 and 10 μmol/L) significantly increased the expression of PGC-1α, NRF1, and NRF2, while 10 μmol/L SPD increased the expression of SIRT1 and TFAM (Figure 4F). Using a fixed dose of 10 μmol/L SPD, we further observed that the expression of these proteins increased in a time-dependent manner (Figure 4G). These data suggest that SPD might delay aging of cardiomyocytes by activating the SIRT1/PGC-1α signaling pathway.

To confirm that SPD-induced mitochondrial biogenesis and functional improvement is mediated by SIRT1 activation, we examined the alterations in SIRT1/PGC-1α signaling resulting from the depletion of the polyamine pool through exposure to a combination of difluoromethylornithine (DFMO) and mitoguazone (MGBG), to induce polyamine synthesis inhibition, or to the SIRT1 inhibitor EX-527. As shown in Figure 5A, SPD enhanced the expression of SIRT1, PGC-1α, NRF1, NRF2, and TFAM in NRCMs pre-treated with H₂O₂. Meanwhile, both DFMO and EX527 abolished the SPD-mediated increase in protein expression. We next measured the expression of mitochondrial oxidative phosphorylation (OXPHOS) complex I, II, and III subunits, and ATP production in NRCMs (Figure 5B and 5C) and analyzed Δψm changes in H9C2 cells (Figure 5D). We noted that SPD treatment reversed the H₂O₂-induced decrease in OXPHOS complex protein expression, ATP levels, and Δψm, whereas DFMO and EX527 partly abrogated these effects.

Figure 5. Inhibition of polyamine biogenesis and SIRT1 activity attenuates SPD-induced mitochondrial biogenesis and functional improvement in aging cardiomyocytes. For in vitro studies, NRMCs and H9C2 cells were cultured as follows: normal culture (Control), H₂O₂ treatment-induced aging (H₂O₂), H₂O₂ plus SPD (H₂O₂ + SPD), H₂O₂ plus SPD and DFMO (DFMO), or H₂O₂ plus SPD and EX527 (EX527). (A) Representative immunoblot bands for SIRT1, PGC-1α, NRF1, NRF2, and TFAM, and quantification of protein expression in NRMCs. GAPDH was used as loading control (n = 4). (B) Representative immunoblot bands for OXPHOS complexes I (NDUFV2), II (SDHB), and III (UQCRC2), and quantification of protein expression in NRMCs (n = 4). (C) ATP content measured by luminometry in NRMCs (n = 8). (D) Mitochondrial transmembrane potential (ΔΨm) detected by JC-1 fluorescence staining in H9C2 cells. Mean fluorescence intensity is displayed on the right of the graphs (n = 6). ^# P < 0.05 vs. Control, ^## P < 0.01 vs. Control, ^* P < 0.05 vs. H₂O₂, ^** P < 0.01 vs. H₂O₂, ^$ P < 0.05 vs. H₂O₂ + SPD, ^$$ P < 0.01 vs. H₂O₂ + SPD. For in vivo studies, the rats were divided into five groups: 1) young (3 months old), 2) old (24 months old), 3) SPD (24-months-old rats treated by SPD for 6 weeks), 4) DFMO (24-month-old rats treated with SPD and DFMO plus MGBG), and 5) EX527 (24-month-old rats treated with SPD and EX527). (E) Cardiac aging evaluated by SA-β-gal staining ex-vivo. (F) SA-β-gal staining quantification. Scale bars: 20 μm (n = 6). (G) Mitochondrial DNA (mtDNA) copy number detected by real-time PCR (n = 8). (H) Mitochondrial oxidative phosphorylation (OXPHOS) efficiency was evaluated based on mitochondrial State 3 and 4 oxygen consumption (a), respiratory control rate (RCR) (b), P/O ratio (c), and proton leakage (d); n = 8. ^# P < 0.05 vs. young, ^## P < 0.01 vs. young, ^* P < 0.05 vs. old, ^** P < 0.01 vs. old, ^$ P < 0.05 vs. SPD, ^$$ P < 0.01 vs. SPD. n = 6 for each group.

In in vivo studies, we further observed that depletion of the polyamine pool and inhibition of SIRT1 activity independently increased SA-β-gal-positive staining area (Figure 5E and 5F) and reduced mitochondrial DNA (mtDNA) copy number (Figure 5G) in the myocardium of aged rats treated with SPD. Moreover, SPD-induced improvement in cardiac mitochondrial respiratory function was also remarkably reduced, as indicated by a decrease in mitochondrial State 3 respiration (Figure 5H-a), RCR (Figure 5H-b), and the P/O ratio (Figure 5H-c), and an increase in proton leakage (Figure 5H-d). These results strongly suggest that SPD attenuates cardiac aging by activating SIRT1 and enhancing mitochondrial biogenesis, which leads to improved mitochondrial function in cardiomyocytes.

SIRT1 mediates spermidine-induced mitochondrial biogenesis through deacetylation and nuclear translocation of PGC-1α

To verify that SPD-induced upregulation of SIRT1 results in deacetylation and nuclear translocation of PGC-1α, the SIRT1-selective inhibitor EX527 and the inhibitors of polyamine biogenesis were tested in vivo and vitro. Nuclear expression of SIRT1/PGC-1α signaling proteins was significantly decreased in cardiac tissue from aged rats, and these changes were partially reversed by SPD supplementation. In turn, depletion of the polyamine pool and inhibition of SIRT1 activity hampered the upregulation of SIRT1, PGC-1α, NRF1, NRF2, and TFAM induced by SPD (Figure 6A and 6B). Since SIRT1 deacetylase activity is nicotinamide adenine dinucleotide (NAD+)-dependent, we also measured changes in NAD+ content. Consistent with SIRT1 expression data, significantly decreased NAD+ levels were observed in the aged myocardium, compared with young controls. SPD supplementation increased cardiac NAD+ contents in aged rat hearts, and this effect was abrogated by both polyamine depletion and EX527 (Figure 6C).

Figure 6. SIRT1 is required for SPD-induced PGC-1α activation in aging cardiomyocytes. (A) Western blot detection of SIRT1, PGC-1α, NRF1, NRF2, and TFAM in nuclear fractions isolated from cardiac tissue. (B) Quantification of protein expression based on L-lamin as a nuclear loading control (n = 4). (C) Quantification of NAD+ levels by fluorimetry (n = 8). ^# P < 0.05 vs. young, ^## P < 0.01 vs. young, ^* P < 0.05 vs. old, ^** P < 0.01 vs. old, ^$ P < 0.05 vs. SPD, ^$$ P < 0.01 vs. SPD. (D) Co-localization of SIRT1 (red) and PGC-1α (green) by immunofluorescence in H9C2 cells. Nuclei were stained with DAPI (blue) (n = 8); scale bars: 20 μm. (E, F) Western blot and immunoprecipitation (IP) analysis of PGC-1α acetylation status in NRCMs. Quantification is shown on the right-hand side of the graphs. GAPDH was used as loading control (n = 4). ^# P < 0.05 vs. Control, ^## P < 0.01 vs. Control, ^* P < 0.05 vs. H₂O₂, ^** P < 0.01 vs. H₂O₂, ^$ P < 0.05 vs. H₂O₂ + SPD, ^$$ P < 0.01 vs. H₂O₂ + SPD.

Next, in vitro studies were performed on H9C2 cells and NRCMs. Using immunofluorescence, we determined that co-localization of PGC-1α and SIRT1 was significantly reduced after exposing H9C2 cells to H₂O₂. Exposure to SPD prevented this effect, but failed to do so in the presence of DFMO or EX527 (Figure 6D). Acetylated PGC-1α levels were next measured by western blot and co-IP in NRCMs (Figure 6E and 6F). Compared with control cells, H₂O₂ treatment significantly increased the acetylation status of PGC-1α, while SPD treatment had the opposite effect. By contrast, both DFMO and EX527 exposure increased the acetylation status of PGC-1α. Taken together, these data demonstrate that SPD stimulates mitochondrial biogenesis in senescent cardiomyocytes by preserving cellular NAD+ levels and inducing SIRT1-mediated deacetylation of PGC-1α.

Spermidine stimulates mitochondrial biogenesis through PGC-1α

To further assess the involvement of PGC-1α in the stimulation of mitochondrial biogenesis mediated by SPD, siRNA-mediated PGC-1α knockdown was performed in H9C2 cells (Figure 7A). Following transfection with scrambled control (ci) or PGC-1α-targeted siRNAs (si) for 6 h, cells of the two groups were cultured with SPD for 48 h. Western blot analyses showed that, as expected, SPD exposure significantly upregulated the expression of PGC-1α, NRF1, NRF2, and TFAM in ci-transfected cells. PGC-1α siRNA significantly down-regulated the expression of PGC-1α and TFAM. However, the expression of these proteins was not altered, in SPD-treated cells after PGC-1α knockdown (Figure 7B). These results strongly suggest that SPD promotes mitochondrial biogenesis through PGC-1α.

Figure 7. Deletion of PGC-1α abrogates SPD-mediated enhancement of mitochondrial biogenesis. (A) Western blot analysis of PGC-1α expression in H9C2 cells transfected with siRNA against PGC-1α (si) or control (scrambled) siRNA (ci). (B) Western blot and quantification analysis of PGC-1α, NRF1, NRF2, and TFAM expression in transfected H9C2 cells in the absence (ci and si) or presence of SPD (ci + SPD and si + SPD). GAPDH served as loading control (n = 4). ^* P < 0.05 vs. Scramble Control.

Animal experiments

Three-month-old and 24-month-old male Wistar rats were purchased from the experimental animal centre of Harbin Medical University. The animals were acclimated for 4 weeks in a quiet environment, at room temperature (25 °C) with 50% humidity, a 12 h/12 h light/dark cycle, and standard rodent chow and water provided ad libitum. Subsequently, rats were randomly separated into five groups: (1) Control: 3-month-old rats intraperitoneally injected with normal saline for 6 weeks; (2) Old: 24-month-old rats intraperitoneally injected with normal saline for 6 weeks; (3) SPD: 24-month-old rats intraperitoneally injected with SPD (Sigma, St. Louis, MO, USA) dissolved in normal saline (10 mg/kg/d) for 6 weeks before sacrifice; (4) DFMO, 24-month-old rats were given SPD for 6 weeks and simultaneously provide with 2% α-difluoromethylornithine (DFMO; Sigma, St. Louis, MO, USA) in drinking water and intraperitoneally infused with mitoguazone (MGBG, Sigma, St. Louis, MO, USA) dissolved in normal saline (15 mg/kg/d) in the last week; and (5) EX527, 24-month-old rats were administered SPD for 6 weeks and simultaneously provide with EX527 intraperitoneally (a SIRT1 inhibitor; Selleck, Shanghai, China) at 5 mg/kg/d in the last week.

Transmission electron microscopy and mitochondrial area measurement

A portion of the left ventricular apex, approximately 3.0 × 4.0 × 1.0 mm³ in size, was placed in glutaraldehyde phosphate buffer for 30 min and then cut into small pieces of approximately 1~2 mm³ to continue fixation for 24 h at 4 °C. After conventional dehydration, soaking, embedding, and dyeing procedures, 50-70 nm ultrathin slices were obtained. Myocardial ultrastructure was observed using a transmission electron microscope (TEM; S4800 Hitachi, Tokyo, Japan). Mitochondrial area measurement was performed as previously described [62]. Briefly, two left ventricle tissue pieces from each rat were randomly chosen. Images were acquired at a magnification of 10,000× when an area containing longitudinal myofilaments surrounded by a mitochondrial network was observed. Individual mitochondria and myofilaments were delineated, and mitochondrial areas were quantified using Image J software.

Determination of polyamine content

Extraction and benzoylation of polyamines from the left ventricular tissue of rats was based on our previously published methods [37]. Polyamine content was measured using high-performance liquid chromatography (HPLC; Waters Co., Milford, USA). Polyamine derivatives were separated on a Hypersil ODS C18 column (250×4.6 mm, 5 μm; Waters Co.), eluted with methanol and distilled water (v/v, 65/35) at 30 °C, and monitored using an ultraviolet detector at 229 nm. The polyamine concentration was expressed in nanomoles of amines per gram of wet tissue (nmol/g w.w.).

Determination of senescence-associated β-galactosidase activity

Senescence-associated β-galactosidase (SA-β-gal) staining was performed according to the instructions of the SA-β-galactosidase Staining Kit (Beyotime Inc., Haimen, China C0602). In brief, cryostat sections were fixed in 3% formaldehyde for 15 min, followed by three washes in phosphate-buffered saline (PBS) at room temperature. Slides were immersed in freshly prepared SA-β-gal staining solution and then incubated at 37 °C overnight. Stained sections were washed twice with PBS and counterstained for 1 min with hematoxylin. Samples were analyzed under an optical microscope by two independent investigators in a blind fashion. All samples were stained in triplicate. The activity of the senescence marker was scored as the percentage of dark blue areas. SA-β-gal staining was also performed, following protocol instructions, on neonatal rat cardiomyocytes and H9C2 cells.

Measurement of antioxidant enzyme activity

Superoxide dismutase (SOD) and catalase (CAT) activities in cardiac tissue were measured using commercial kits (SOD: A001-3-1; CAT: A007-2-1; Jiancheng Bio. Institute, Nanjing, China) with a spectrophotometer (Perkin- Elmer, Norwalk, CT, USA). Protein concentration was measured using the bicinchoninic acid method (Pierce, Rockford, USA) with bovine serum albumin (BSA) as a standard.

Mitochondrial isolation

Mitochondria were extracted from cardiac tissue using a Tissue Mitochondria Isolation Kit (Beyotime Biotechnology, Shanghai, China C3606). Briefly, after treatment, rats’ ventricular muscle tissues were cut and weighed. Samples were cut into smaller pieces and digested with trypsin by adding the appropriate volume of mitochondrial separation reagent. Each sample was ground 20-30 times in a Dounce homogenizer and centrifuged at 600 g for 5 min at 4 °C. Thereafter, the supernatant was transferred and centrifuged at 11,000 g for 10 min at 4 °C. Pelleted mitochondria were divided and one fraction preserved in Mitochondria Storage Buffer (Beyotime Biotechnology, Shanghai, China C3609) for immediate detection of respiratory rate, while the other was stored at −80 °C for western blot analysis.

Determination of mitochondrial respiratory function

State 3 and State 4 oxygen consumption rates, respiratory control rate (RCR), and ADP consumed/oxygen consumed (P/O ratio) with pyruvate and malic acid as substrates were measured in isolated myocardial mitochondria using a Clark oxygen electrode (Hansatech Instruments, Norfolk, UK) at 30 °C in mitochondrial respiration buffer as previously described [36]. When the oxygen electrode reaction chamber temperature was 25 °C, 500 μg of mitochondrial protein was added to the buffer solution. After equilibration for 1 min, 5 mmol/L pyruvate and 5 mmol/L malate were added to the buffer to initiate the respiration test. State 3 respiration begun after addition of 0.2 μmol/L of ADP; once ADP was exhausted, State 4 respiration ensued. RCR, an important index to evaluate mitochondrial respiratory function, reflects mitochondrial oxidative phosphorylation (OXPHOS) coupling efficiency and is expressed by the ratio of the State 3/State 4 oxygen consumption rates, representing ADP availability and depletion, respectively. P/O in turn is the ratio of the number of moles of ADP added to the reaction system to the number of moles of oxygen consumed, which represents the efficiency of mitochondrial OXPHOS.

Mitochondrial DNA copy number estimation

Cardiac tissue mtDNA was extracted using a whole genome DNA extraction kit (Zoman Biotechnology, Beijing, China), and its concentration measured on a Nanodrop 2000 instrument. Real-time PCR was conducted with a 2x SYBR Green qPCR Master Mix Kit in a LightCycler 96 real-time PCR cycler. The total PCR volume was 20 μl, with 1 μL each of 10 μmol/L upstream and downstream primers, 1 μl of cDNA, 7 μL of SYBR Green qPCR Master Mix, and 10 μL of deionized water. Two-step amplification involved activation of the Hot-Start DNA Polymerase for 10 min at 95 °C. PCR conditions involved 40 cycles, with each cycle consisting of denaturing at 95 °C for 15 s, extension at 60 °C for 60 s and melting at 95 °C for 15 s, 60 °C for 60 s, and 95 °C for 15 s. The primers were as follows: Cytochrome B: Forward: 5′-GCAGCTTAACATTCCGCCCAATCA-3′; Reverse: 5'-TGTTCTACTGGTTGGCCTCCGATT-3′; Actin: Forward, 5'-TCGTGCGTGACATTAAAGAG-3′; Reverse: 5'-ATTGCCGATAGTGATGACCT-3′; Calculation formula: ^ΔCT=(β-actin CT)-(Cyt B CT), mtDNA copy number=2×2^ΔCT.

Nuclear isolation

Nuclear and cytosolic protein fractions were extracted from cardiac tissue using the Nuclear and Cytoplasmic Protein Extraction Kit (P0027; Beyotime Biotechnology, Shanghai, China). Briefly, tissue samples were cut into pieces and homogenized in the presence of phenylmethylsulfonyl fluoride (PMSF). Each mixture was then placed in an ice bath for 15 min and then centrifuged at 1500 g for 15 min at 4 ºC to separate cytoplasmic proteins (supernatant) and cell nuclei (precipitate). Corresponding extraction reagents were next added to each fraction following the manufacturer’s protocol. The samples were stored at −80 °C for subsequent western blot analysis.

Nicotinamide adenine dinucleotide (NAD+) measurement

NAD+ from cardiac tissue and cultured cells was estimated colorimetrically with a NAD/NADH assay kit (ab65348; Abcam, Cambridge, MA, USA) according to the manufacturer’s instructions.

Cell culture

Primary neonatal rat cardiomyocytes (NRCMs) were obtained from Wistar rats aged 1-3 days as previously described [63]. All experiments were approved by the Animal Care Committee for the Use of Experimental Animals at Harbin Medical University (Heilongjiang, China). Briefly, the hearts were cut into pieces, digested with trypsin for 8 minutes and terminated digestion with DMEM culture solution containing 10% fetal calf serum. After 7 times of digestion, the cells were collected by centrifugation at 600 g for 10 min at 4 °C. Two hours after incubation at 37 °C/5% CO₂, the attached cells were discarded, and unattached cardiomyocytes were re-cultured in collagen-coated 35 mm Petri dishes or 8-, 24- or 96-well plates in DMEM containing 10% fetal bovine serum (FBS) and 1% penicillin or streptomycin, in a humidified incubator at 37 °C/5% CO₂. The media was changed twice a week. The H9C2 cell line (purchased from the Chinese Academy of Sciences in Shanghai, China) was cultured in DMEM containing 10% FBS and 1% penicillin or streptomycin in an incubator at 37 °C with 5% CO₂.

Cell viability assays

For cell viability assessment, NRCMs were cultured for 48 h in 96-well plates and then exposed to 40 μmol/l H₂O₂ over 4 h (see "Supplementary Materials"). The medium was then replaced with normal medium, and various concentrations of SPD (0-100 μmol/L) were added for 48 h. Alternatively, following exposure to H₂O₂, 10 μmol/L SPD was added to fresh culture medium and NRCMs incubated for different time periods up to 48 h. Cell viability was determined using the Cell Counting Kit-8 (Solarbio, Beijing, China CA1210) according to manufacturer's instructions. Absorbance (450 nm) was measured, and results expressed as percentages relative to control (100%).

Cell experiments

NRCMs and H9C2 cells were randomly divided into five experimental groups: (1) Control: high glucose-DMEM with 10% FBS; (2) H₂O₂: 40 μmol/l H₂O₂ for 4 h, and incubation in normal medium for additional 48 h; (3) H₂O₂-SPD: 40 μmol/l H₂O₂ for 4 h, and incubation in normal medium with 10 μM SPD for additional 48 h; (4) H₂O₂+SPD+DFMO (DFMO): 2 mmol/L DFMO plus SPD for 48 h followed by H₂O₂ for 4 h; (5) H₂O₂+SPD+EX527 (EX527): 40 μM EX527 plus SPD for 48 h followed by H₂O₂ for 4h. Aminoguanidine (1 mM) was added to the medium 5 min before SPD treatment to inhibit the activity of PAO in FBS.

Cellular ATP measurement

ATP concentration in primary cultured NRCMs was assayed using an ATP Assay Kit (S0027; Beyotime Biotechnology, Shanghai, China) according to the manufacturer’s instructions and measured on a Luminoskan™ Ascent luminometer (Thermo Fisher Scientific Inc., USA). An ATP calibration curve was used for data normalization.

Mitochondrial transmembrane potential and ROS production assays

To evaluate mitochondrial transmembrane potential (Δψm), the cells were stained with tetramethylrhodamine ethyl ester (TMRE, AAT Bioquest Inc. Sunnyvale, CA, USA) for 20 min at 37 °C, or with JC-1 (Mitochondrial Membrane Potential Assay Kit; C2006, Beyotime Inc., Shanghai, China). ROS production in living cells was assessed by dihydroethidium (DHE) staining (S0063, Beyotime Inc., Shanghai, China). To detect the production of mitochondrial superoxide, the cells were stained with MitoSOX (M36008, ThermoFisher Scientific, Waltham, MA, USA). Images were taken with an Olympus FluoView FV1000 fluorescence microscope (Olympus Optical Co., Ltd., Takachiho, Japan). Results were compiled from 20–45 cells from three or more independent experiments, and the same microscope settings were used for all treatments.

Western blot analysis

Frozen left ventricular cardiac tissue and isolated nuclear samples were homogenized in ice-cold RIPA lysis buffer (P0013B; Beyotime Inc., Shanghai, China) with 50 μg/ml PMSF and incubated on ice for 40 min. Homogenates were next centrifuged at 10,000 g for 15 min at 4 °C to remove cellular debris and isolate total protein. Cardiomyocytes were harvested and lysed in the same RIPA buffer containing PMSF. Protein concentration was determined using a Bradford assay kit (P0006C; Beyotime Inc., Shanghai, China). Equal amounts of protein from different experimental groups were loaded and separated on 10% SDS-PAGE gels, and electrophoretically transferred onto 0.2 μm PVDF membranes. The membranes were blocked in TBS-T containing 5% (w/v) skim milk at 37 °C for 1 h, and incubated overnight at 4 °C with primary rabbit antibodies specific to GAPDH (10494-1-AP), p21 (10355-1-AP), p16 (10883-1-AP), SOD (24127-1-AP), CAT (21260-1-AP), GLB1 (15518-1-AP), SIRT1 (13161-AP), TFAM (19998-1-AP), NRF1 (12482-1-AP), NRF2 (16396-1-AP), SDHB (10620-1-AP), NDUFV2 (15301-AP), or UQCRC (14742-1-AP). All polyclonal antibodies were from Proteintech (Wuhan Sanying Biotechnology, China). Antibodies against ODC (sc-33539) and SSAT (sc-67159) were from Santa Cruz Biotechnology (Dallas, TX, USA). Goat anti-PGC-1α (ab106814) was from Abcam (Cambridge, MA, USA), and mouse anti-lamin B1 (66095-1-Ig) was from Proteintech (China). Horseradish peroxidase (HRP)-conjugated secondary antibodies (ZB-2301, ZB-2305, ZB-2306; Zhongshan Golden Bridge Biotechnology, Beijing, China) were used to label primary antibodies. Immunoreactive proteins were then developed using ultrasensitive ECL luminescent solution (Proteintech), quantified using a FluorChem Chemiluminescence Imaging System (Protein Simple, San Jose, CA) via densitometry, and normalized to that of GAPDH. The final results were expressed as relative protein levels by normalizing the data to control values.

Immunofluorescence

Following treatment with SPD, H9C2 cells were seeded onto glass cover slips in 24-well plates. For detection of PGC-1α and SIRT1, cells were fixed in 4% paraformaldehyde for 15 min, permeabilized with PBS/0.1% Triton X-100, blocked with 3% BSA in PBS, and incubated with PGC-1α (sc-518038) and SIRT1 (sc-15404) antibodies (Santa Cruz Biotechnology, Dallas, TX, USA) overnight at 4 °C. Fluorescent secondary antibodies (ab150113 and ab150075; Abcam, Cambridge, MA, USA) were used for detection according to the manufacturer’s instructions. Immunofluorescence was assessed with a confocal microscope (Carl Zeiss LSM510, Tokyo, Japan). Confocal laser scanning microscopy analysis of PGC-1α and SIRT1 colocalization was performed for H9C2 cells.

Detection of protein acetylation through co-immunoprecipitation

To detect PGC-1α acetylation in cells, cultured cardiomyocytes were collected and lysed in lysis buffer (P0013C; Beyotime, Shanghai, China) for 30 min at 4 °C. After centrifugation at 14,000 g for 20 min at 4 °C, the supernatants were incubated with 1 μg of anti-PGC-1α antibody overnight at 4 °C. Rabbit IgG (Santa Cruz Biotechnology) was used as control. Each sample was incubated with protein A/G magnetic bead slurry (Selleckchem, Houston, TX, USA) for 2 h at 4 °C to avoid nonspecific binding. A total of 500 μg of protein from each sample was incubated with 2 μg of specific antibody to acetylated lysine (#9441, Cell Signaling Technology, Danvers, MA, USA) overnight at 4 °C according to the instructions of the Protein A/G Magnetic Beads for IP. The blot was probed with rabbit anti-PGC-1α (Abcam) and anti-acetylated lysine antibodies and then treated with appropriate secondary antibodies conjugated to HRP (Santa Cruz Biotechnology, Dallas, TX, USA). Then, western blotting was performed as described above.

siRNA-mediated PGC-1α knockdown

For short interfering RNA (siRNA)-mediated knockdown studies, H9C2 cells were seeded into six-well plates 48 h prior to transfection on serum-free DMEM. Cells were transfected with scrambled, nonspecific siRNA or siRNA targeting the mouse PGC-1α gene (GenePharma, Shanghai, China) using Lipofectamine 2000 Reagent (Invitrogen, Carlsbad, CA, USA) for 6 h according to the manufacturer's instructions. Next, H9C2 cells were treated with SPD for 48 h for subsequent assays.

Statistical analyses

All statistical analyses were performed using SPSS 17.0 (SPSS. Chicago, IL, USA). Values are expressed as mean ± SD. The t-test was used for comparison between two groups. Comparisons between the mean of three or more groups were done by one-way ANOVA. Correlation analysis was performed by linear regression. Differences were considered significant at p < 0.05. Each experiment consisted of at least three replicates per condition.