Abstract
Myocardial infarction (MI) causes disturbances in myocardial energy metabolism, ultimately leading to a poor prognosis. Cytosolic glycogen autophagy (glycophagy) and mitochondrial autophagy (mitophagy) are upregulated in MI to optimize energy metabolism but to a limited extent. Asiatic acid (AA), a pentacyclic triterpene derived from the traditional Chinese herb Centella asiatica, displays anti-inflammatory, antioxidant, and antiapoptotic activities. AA has been found to alleviate focal cerebral and liver ischemic injury by reversing mitochondrial dysfunction. In this study, we investigated whether AA exerted cardioprotective effects against MI by activating glycophagy and mitophagy to improve the energy balance. In vitro cardioprotective effects were examined in neonatal mouse cardiomyocytes subjected to oxygen-glucose deprivation for 12 h. Treatment with AA (2–50 μM) significantly increased cell viability and improved the energy metabolism evidenced by increased ATP level and phosphocreatine/ATP ratio. In vivo cardioprotective effects were studied in a mouse model of MI. Administration of AA (5–125 mg·kg−1·d−1, ig) significantly reduced infarct size and ischemic myocardial injury, and improved cardiac function. AA treatment also promoted mitophagy and relieved mitochondrial edema evidenced by increased number of mitophagosomes in ischemic myocardium in vivo and increased mitochondria-light chain 3 (LC3)-II colocalization in ODG-treated cardiomyocytes in vitro. Mitophagy activation was accompanied by activation of the AMPK signaling pathway. Knockdown of AMPK abolished AA-activated mitophagy. Furthermore, we showed that glycophagy was upregulated in OGD cardiomyocytes evidenced by increased starch binding domain protein 1 (STBD1)-GABA type A receptor-associated protein-like 1(GABARAPL1) interaction and extracellular acidification rate, whereas AA treatment further promoted glycophagy accompanied by PI3K/Akt activation. PI3K inhibitor LY294002 or Akt inhibitor GSK690693 blocked the effects of AA on glycophagy and glycolysis. Finally, simultaneous inhibition of glycophagy and mitophagy abolished the cardioprotective effects and energy regulation of AA. These results demonstrate that AA protects ischemic cardiomyocytes by modulating glycophagy- and mitophagy-based energy metabolism through the PI3K/Akt and AMPK pathways.
Keywords: myocardial infarction, asiatic acid, glycophagy, mitophagy, energy metabolism, AMPK signaling
Introduction
Myocardial infarction (MI), the primary component of cardiovascular disease, is one of the leading causes of morbidity and mortality worldwide [1]. Despite treatment advances in recent years, the global medical burden caused by MI remains high [2]. Cardiac mitochondria are responsible for energy production and cardiomyocyte death; their integrity and dynamic nature, which are maintained by their intrinsic proteolytic system, are crucial for cell homeostasis [3]. Under ischemic conditions, ATP and creatine depletion due to mitochondrial injury result in functional and structural left ventricular alterations related to lower myocardial energy reserves [4]. Energy metabolism dysfunction leads to excess mortality from ventricular arrhythmias and progressive heart failure [4]. Therefore, agents that modulate energy homeostasis in MI may be promising therapeutic candidates.
Asiatic acid (AA) is a pentacyclic triterpene derived from Centella asiatica, which has anti-inflammatory, antioxidant, and antiapoptotic effects [5–8]. AA has long been used in herbal therapy for wounds and skin disorders [9]. It is detectable in human serum after oral administration [10], and it causes no significant toxicity in mice after either oral or subcutaneous administration [9]. AA exerts protective effects on focal cerebral ischemia and hepatic ischemia/reperfusion injury by ameliorating oxidative stress and maintaining mitochondrial homeostasis [7, 11], suggesting that it may be a potential therapeutic for mitochondrial and energy metabolism homeostasis.
Mitochondrial autophagy (mitophagy) is a specific autophagic process that selectively degrades damaged mitochondria to decrease reactive oxygen species release and protect against cell apoptosis [3]. Therefore, enhanced mitophagy is important for mitochondrial homeostasis and cardiomyocyte survival during ischemia, hypoxia, and/or other energy shortages [12]. Among the regulators of mitophagy, AMP-activated protein kinase (AMPK) is a classic activator that responds to cell energy stress; it works via various mechanisms, involving inhibition of the mammalian target of rapamycin complex and phosphorylation and activation of Unc-51-like kinase 1 (ULK1) [13]. Phosphorylation of a conserved threonine residue (Thr172) by upstream kinases is essential for AMPK activation [14]. In addition, light chain 3 (LC3) is also considered a modulator of mitophagy. LC3-I is lipidated to LC3-II to induce autophagosome formation, and LC3 lipidation results in an elevated LC3-II/LC3-I ratio or LC3-II expression, which is often used as a marker for mitophagy in numerous tissues, including myocardial tissue [15]. However, it is unclear whether AA maintains mitochondrial and energy homeostasis to protect the ischemic myocardium via mitophagic regulation.
In addition, glycogen autophagy (glycophagy) is a glycogen degradation and utilization process, which triggers liberation of free glucose that can be rapidly utilized by cells with energy requirements [16]. This process is a link between glucose homeostasis and autophagy in neonatal and adult cardiomyocytes, hepatocytes, and other organ cells under conditions of demand for glucose production [17]. Increased glycophagy provides a sufficient supply of glucose to ischemic cardiomyocytes for essential energy metabolism [18]. The potential mechanism of glycophagy involves the binding of starch binding domain protein 1 (STBD1) and the autophagy-related protein 8 (Atg8) family member, GABA type A receptor-associated protein-like 1 (GABARAPL1). STBD1, which consists of a carbohydrate-binding domain and an Atg8-interacting motif, is localized to the glycogen particle and plays an important role in intracellular glycogen trafficking to autophagosome by interacting with GABARAPL1, thereby constituting a specific mechanism of glycophagy [19, 20]. However, it remains unknown whether AA improves the energy supply after MI by regulating glycophagy.
The aims of this study were to determine the energy modulation and cardioprotective effects of AA on oxygen-glucose deprivation (OGD)-treated cardiomyocytes and a mouse MI model, investigate whether mitophagy and glycophagy are responsible for the energy metabolism and protective roles of AA, and elucidate the mechanisms underlying the effects of AA on mitophagy and glycophagy.
Materials and methods
Isolation and culture of neonatal mouse ventricular cardiomyocytes
Neonatal cardiomyocytes were isolated from one-day-old C57BL/6 mice.[21] Briefly, neonatal mice were anesthetized with sodium pentobarbital and sacrificed. Their hearts were removed and rapidly placed on ice. Then, the ventricles were dissected, minced into 1-mm3 pieces, transferred to a sterile tube, and washed in cold phosphate-buffered saline (PBS) to remove blood clots. Minced tissue was digested in PBS supplemented with 1 mg/mL trypsin, 1 mg/mL collagenase type II, and 0.2 mg/mL glucose for 5 min at 37 °C, and incubated with 0.1 mmol/L BrdU to selectively enrich for cardiomyocytes by inhibiting the growth of cardiac fibroblasts. Cardiomyocytes were purified using the differential adhesion method. Isolated cardiomyocytes were plated in a 24-well plate (Corning, NY, New York, USA) at a density of 25,000 cells/cm2 in Dulbecco’s modified Eagle’s medium (DMEM; Thermo Fisher Scientific, Waltham, MA, USA) containing 4.5 g/L glucose and supplemented with 10% fetal bovine serum (FBS; Thermo Fisher Scientific) and 1% penicillin–streptomycin solution (Thermo Fisher Scientific). The cells were cultured at 37 °C in a humidified atmosphere containing 5% CO2.
Cell treatment and transfection
OGD was achieved by culturing cells for 12 h in glucose-free DMEM without FBS in a tri-gas incubator (Heal Force, Shanghai, China) saturated with 1% O2/5% CO2/94% N2 at 37 °C for oxygen deprivation. For cell treatment, AA (Sigma-Aldrich, St. Louis, MS, USA) was dissolved in 5% DMSO and then added to the medium at different concentrations (2, 10, 20, and 50 μmol/L) before OGD. A phosphoinositide 3-kinase (PI3K) inhibitor LY294002 (20 μmol/L; MedChemExpress, Monmouth Junction, NJ, USA) and an Akt inhibitor GSK690693 (10 μmol/L; MedChemExpress) were added to DMEM for 12 h prior to the OGD process. The lentiviral short hairpin RNAs (shRNAs) targeting mice STBD1 and AMPK were purchased from Open Biosystems (Lafayette, CO, USA). To interfere with the expression of STBD1 and AMPK, shSTBD1 and shAMPK were transfected in accordance with the manufacturer’s protocol. After 24 h of incubation, the transfection medium was removed, and the cells were harvested for further analysis.
Cell viability and lactate dehydrogenase release assay
Cardiomyocytes were seeded in 96-well plates at a concentration of 1 × 104 cells/mL. Cell viability was detected using a Cell Counting Kit-8 Kit (CCK-8; Dojindo, Tokyo, Japan). Absorbance at 450 nm was measured using a microplate reader (BioTek Synergy 2; Winooski, VT, USA). The mean optical density (OD) measurements from the indicated groups were used to calculate the percentage of cell viability. Lactate dehydrogenase (LDH) release from cardiomyocytes into the medium was assessed using an LDH Cytotoxicity Assay Kit (Roche, Basel, Switzerland) in accordance with the manufacturer’s instructions.
High-performance liquid chromatography analysis
ATP and phosphocreatine (PCr) levels in OGD cardiomyocytes and ischemic heart tissue were measured using a high-performance liquid chromatography (HPLC) system (SIL-20A; SHIMADZU, Kyoto, Japan) with detection at 205 nm. Briefly, the heart was quickly removed from the anesthetized mouse and the left ventricular wall was collected. Then, the wall was homogenized with pre-cooled 80% methanol, and supernatants were assayed after centrifugation. Similarly, cardiomyocytes were quickly extracted with precooled 80% methanol, and the supernatants were collected after centrifugation. The following chromatographic conditions were used: (1) mobile phase A (10 mmol/L KH2PO4, 2.5% acetonitrile, 5.8 mmol/L tetrabutylammonium hydroxide, pH 6.0), (2) phase B (10 mmol/L KH2PO4, 25% acetonitrile, 5.8 mmol/L tetrabutylammonium hydroxide, pH 5.5), and (3) reversed phase (250 mm × 4.6 mm × 5 µm C18 column). ATP and PCr concentrations were determined by dividing the peak area of the samples by the corresponding standard product.
Mitophagic activity detection
A canonical double transfection-based method was used to monitor the mitophagy in cardiomyocytes. [22, 23] Briefly, neonatal cardiomyocytes were transfected with adeno-associated virus containing GFP-LC3-II (Hanbio Inc., Shanghai, China). For Mito-DsRed delivery, the Mito-DsRed DNA sequence was cloned from pDsRed2-Mito (Clontech, 632421) and inserted into the pLVX-puro plasmid (Clontech, 632164) and then transfected into HEK293T cells using a set of plasmids (Clontech, 631247) for lentivirus packaging. After transfection with GFP-LC3-II and Mito-DsRed, the cells were treated and imaged for GFP and DsRed by fluorescence microscopy.
Measurement of extracellular acidification rate
The extracellular acidification rate (ECAR) was measured using a Seahorse XF96 extracellular flux analyzer (Agilent, North Billerica, MA, USA), which reflects the activity of glycolysis. Briefly, cardiomyocytes were seeded in 96-well assay plates at 8 × 103 cells/well and incubated at 37 °C before analysis. ECAR was recorded immediately after completing OGD. The unit for ECAR was mpH/min.
Animal experiments
Adult C57BL/6 mice (weighing 20–25 g and aged 6–8 weeks) were obtained from Shanghai Model Organisms (Shanghai, China) and cared for in accordance with the Guide for the Care and Use of Laboratory Animals published by the National Institutes of Health (8th Edition, 2011; Bethesda, MD, USA). Animal procedures were approved by the Tongji University Committee on Animal Care (Shanghai, China). All experiments conformed to the international guidelines on the ethical use of animals. AA was dissolved in 5% dimethyl sulfoxide (DMSO) to form a solution of 500 mg/mL, and then diluted to a volume of 0.5 mL with a specific concentration according to different dosage gradient groups (5, 25, 50, and 125 mg·kg−1·d−1). The AA solution was administered orally to the mice using an intragastric tube on a daily basis for 7 consecutive days after the surgery.
MI models were established surgically by ligation of the left anterior descending (LAD) coronary artery in anesthetized mice as previously described [24]. Briefly, the mice were placed in an induction chamber; they received 2% isoflurane, and were then connected to a small rodent ventilator. Afterward, the chest was opened to expose the heart, left auricle, and LAD. The LAD was permanently ligated with an 8-0 nylon suture, and the chest cavity was closed. After reinstallation of spontaneous respiration, the mice were extubated and allowed to recover from anesthesia. Sham-operated mice underwent the same procedure, excluding LAD ligation. The mice received intraperitoneal injections of buprenorphine (0.1 mg/kg) every 12 h for up to three days and intraperitoneal injections of carprofen (5 mg/kg) every 12 h for up to one day after the surgery.
The following studies were conducted. Study 1: The mice were divided into Sham, MI, MI + DMSO, and MI + AA (5, 25, 50, and 125 mg·kg−1·d−1) groups (n = 10). The mice were euthanized seven days after MI, and the hearts were harvested. Study 2: The subgroups were the same as in Study 1 (n = 8–10). After completing the echocardiographic examination, the mice were sacrificed 28 days after MI for further analysis. Study 3: The survival curves of Sham, MI, MI + DMSO, and MI + AA (25 mg·kg−1·d−1) groups (n = 16–25) up to eight weeks after surgery were recorded.
Determination of myocardial infarct size
Myocardial infarct size at seven days after MI was evaluated using 2,3,5-triphenyltetrazolium chloride (TTC) staining. The left ventricle was isolated and cut into 4-mm slices perpendicular to the axis of the LAD, from the base to the apex. Slices were immediately immersed in 1% TTC (pH 7.4) at 37 °C in the dark for 10 min to distinguish the infarcted tissue from viable myocardium. Infarct size (fibrosis area) of the heart at 28 days after MI was determined using the Masson Trichrome Stain Kit (G1340; Solarbio, Beijing, China) in accordance with the manufacturer’s instructions. Infarct size was calculated as a percentage of the left ventricle.
Evaluation of animal cardiac function
The cardiac function in the mice was verified by echocardiography 28 days post-MI. Two-dimensional-guided M-mode echocardiography was used to determine left ventricular chamber volume at systole and diastole and contractile parameters such as left ventricular end-diastolic dimension (LVEDD), left ventricular end-systolic dimension (LVESD), left ventricular end-diastolic volume (LVEDV), and left ventricular end-systolic volume (LVESV). Left ventricular fractional shortening (LVFS) was calculated as follows: FS (%) = (LVEDD − LVESD)/LVEDD × 100%. Left ventricular ejection fraction (LVEF) was derived as follows: EF (%) = (LVEDV − LVESV)/LVEDV × 100%. All measurements were based on the average of at least three cardiac cycles.
Immunofluorescence
Cardiomyocytes or cardiac tissue were fixed for 15 min in 4% paraformaldehyde at room temperature and then permeabilized for 20 min with 0.25% Triton X-100 at room temperature. After blocking in 4% bovine serum albumin for 7 min at room temperature, the sections were incubated with primary antibodies (1:100 cardiac troponin I [cTnI] and 1:100 CD31; Abcam, Cambridge, MA, USA) and subsequently with 1:500 secondary antibody conjugated to Alexa Fluor-488 and Alexa Fluor-549 (Life Technologies, Waltham, MA, USA). The TUNEL assay (Roche) and wheat germ agglutinin (WGA; Sigma) staining were performed according to the manufacturer’s instructions. After final washing, the coverslips were mounted on the slides using 50% glycerin, and the nuclei were further stained with DAPI (Beyotime, Jiangsu, China). The tissues and cells were observed using a confocal laser scanning microscope (Olympus, Tokyo, Japan), and analysis was performed by Image-Pro Plus (Media Cybernetics Inc., Bethesda, MA, USA).
Transmission electron microscopy
Heart tissue samples were fixed in 2.5% glutaraldehyde overnight. Subsequently, they were incubated with 1% osmium tetroxide for 2 h in dark. After washing in distilled water, the samples were incubated with 2% uranyl acetate for 2 h at room temperature and then dehydrated in an ethanol concentration gradient. Finally, the samples were embedded in molds with fresh resin. Ultrathin sections were obtained using the EM UC7 ultramicrotome (Leica, Solms, Germany), stained with lead citrate, and examined using Tecnai G2 T12 (FEI Co., Hillsboro, OR, USA).
Western blot analysis
Heart tissue protein was extracted from the left ventricular myocardium using a lysis buffer (100 mmol/L Tris-HCl, 4% SDS, 20% glycerine, 200 mmol/L DTT, phosphatase, and protease inhibitors, pH 6.8). For whole cell lysates, cells were lysed with the Cell Total Protein Extraction Kit (Sangon, Shanghai, China). Protein concentrations were measured using a bicinchoninic acid (BCA) assay kit (Beyotime) in accordance with the manufacturer’s instructions. Proteins were resolved by 8%–12% sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) and electrotransferred to nitrocellulose membranes (Millipore, Darmstadt, Germany). The membranes were probed overnight at 4 °C with the following primary antibodies: STBD1 (1:1000, 11942-1-AP; Proteintech, Rosemont, IL, USA), GBARAPL1 (1:1000, 26632; Cell Signaling, Beverly, MA, USA), phosphorylated Akt (p-Akt) at Ser473 (1:1000, 4060; Cell Signaling), Akt (1:1000, 9272; Cell Signaling), phosphorylated glycogen synthase kinase 3 beta (p-GSK3β) at Ser9 (1:1000, 9323; Cell Signaling), GSK3β (1:1000, 9832; Cell Signaling), LC3-I/ II (1:1000, 4108; Cell Signaling), p-AMPKα at Thr172 (1:1000, 2535; Cell Signaling), AMPKα (1:1000, 5832; Cell Signaling), or GAPDH (1:5000, 60004-1-lg; Proteintech). After washing, the membranes were incubated with secondary antibody for 2 h at room temperature. Finally, proteins were detected by the Odyssey Infrared Imaging System (Li-Cor, Lincoln, NE, USA) and quantified using ImageJ software (NIH). For all Western blot analyses, unless specifically noted, protein levels were calculated from the ratio of corresponding protein/GAPDH.
Quantitative PCR
Quantitative PCR (qPCR) was used to determine the mRNA level of AMPKα both in cardiomyocytes and in mouse heart tissue. Total RNA was extracted using TRIzol reagent (Thermo Fisher Scientific). The RNA (1000 nmol) was subjected to reverse transcription using the PrimeScript RT Reagent Kit with gDNA Eraser (TaKaRa, Dalian, China). Then, 20-μL reactions with primers (GENEWIZ, South Plainfield, NJ, USA) were detected by Light Cycler 480II (Roche) using SYBR Green PCR Master Mix (Thermo Fisher Scientific). Relative quantitative analysis of the change in expression levels was obtained using the ΔΔCT method and normalized to GAPDH. The primer sequences were as follows:
5ʹ-GGCCTCACCCTGAGAGAGTA-3ʹ (AMPKα forward),
5ʹ-TCACAGGGTTCTTCCTTCGC-3ʹ (AMPKα reverse),
5ʹ-GGAGCGAGACCCCACTAACAT-3ʹ (GAPDH forward),
5ʹ-ACATACTCAGCACCGGCCTC-3ʹ (GAPDH reverse).
Co-immunoprecipitation
The whole proteins from cardiomyocytes were extracted using lysis buffer for immunoprecipitation. The lysates were centrifuged, and protein concentrations were measured using the BCA assay. A part of the supernatant was used as input control, while the rest was immunoprecipitated overnight at 4 °C by gently rocking with anti-STBD1 antibody. Approximately 4 μL antibody was used (400 μg total protein). Protein A/G agarose beads (Santa Cruz, Dallas, TX, USA) were added to the immunoprecipitates for 2 h with gentle shaking at room temperature. The precipitated proteins were washed three times with lysis buffer and boiled with 5× loading buffer, followed by Western blot analysis as previously described. Rabbit normal IgG (Santa Cruz) served as the negative control.
Statistical analysis
Data are expressed as the mean ± standard error of the mean (SEM). Statistical analyses were performed using SPSS 24.0 (IBM, Armonk, NY, USA). Significant differences between means were analyzed using the unpaired Student’s t-test or one-way analysis of variance followed by Tukey’s post hoc test. Analysis of survival was performed using log-rank (Mantel–Cox and Gehan–Breslow–Wilcoxon) tests applied to Kaplan–Meier survival curves. P < 0.05 was considered statistically significant.
Results
AA attenuates ischemic myocardial injury and promotes energy metabolism
We first determined the effects of AA on OGD cardiomyocytes. As shown in Figs. 1a and 1b, 2–50 μmol/L AA significantly decreased LDH release and upregulated cell viability compared with the OGD + DMSO or OGD groups. Importantly, 10 μmol/L AA showed the best cytoprotective effects on OGD cardiomyocytes. The TUNEL assay further indicated that 10 μmol/L AA had the strongest cellular antiapoptotic effects (Fig. 1c). Next, energy metabolism was investigated in OGD cardiomyocytes to further determine the cardioprotective roles of AA. The PCr/ATP ratio is an important indicator for myocardial energetic reserve, and it is reduced in patients with severe coronary artery disease and acute MI [25]. AA concentration starting from 10 μmol/L significantly increased the ATP level and PCr/ATP ratio, suggesting a higher energy metabolism level and energy reserve (Fig. 1d, e). In addition, DMSO did not show any effect on cell function under normal and OGD conditions (Fig. S1).
Next, dose gradients of AA (5, 25, 50, and 125 mg·kg−1·d−1) were used to determine the effective dose of AA in vivo. As shown in Fig. 2a, oral administration of AA starting from 5 mg·kg−1·d−1 relieved cellular apoptosis compared with the MI (seven days) or the MI + DMSO groups, and 25 mg·kg−1·d−1 showed the best cardioprotective effect. Also, acute cardiac infarct size determined by TTC staining was markedly reduced in the MI + 25 mg·kg−1·d−1 AA group (Fig. 2b). In addition, the long-term protective effects of AA on MI were explored. As shown in Figs. 2c and 2d, AA ≥ 5 mg·kg−1·d−1 improved the LVEF and fractional shortening (FS) compared with the chronic MI (28 days) and MI + DMSO groups. Especially, 25 mg·kg−1·d−1 AA treatment in MI mouse showed the best cardioprotective effects on improving EF and FS. Masson trichrome staining revealed reduced cardiac infarct size (fibrotic area) of chronic MI in 25 mg·kg−1·d−1 AA-treated mouse myocardium (Fig. 2e). In addition, cardiac myocyte hypertrophy after chronic MI was alleviated by AA treatment (25 mg·kg−1·d−1) (Fig. 2f). However, no differences in cardiac angiogenesis were observed between the AA-treated group (25 mg·kg−1·d−1) and control (Fig. 2g). The survival rate was significantly higher in the MI mice that had received 25 mg·kg−1·d−1 AA than in the MI (56 days) and MI + DMSO groups (Fig. 2h). In addition to the in vitro promotion of energy metabolism, AA treatment restored the ATP level and PCr/ATP ratio in the acute (7 days, Fig. 2i, j) or chronic MI myocardium (28 days; Fig. 2k, l). These results demonstrated that AA could modulate the acute and chronic cardioprotective and energy metabolic effects on MI cardiomyocytes both in vitro and in vivo. An AA concentration of 10 μmol/L AA in vitro and a dose of 25 mg·kg−1·d−1 in vivo showed the best protective effects and were used in subsequent studies.
AA promotes mitophagy in OGD cardiomyocytes
As AA is involved in the regulation of cellular energy metabolism, we next evaluated the effects of AA on mitophagy, an important autophagic process that maintains energy homeostasis in the MI heart [18]. Transmission electron microscopy (TEM) revealed that the mitochondria in myocytes of the MI group were swollen dramatically compared with the Sham group, revealing serious myocardial injury, while relieved mitochondrial edema and increased number of mitophagosomes (red arrows) were observed in the MI + AA group (Fig. 3a). Western blotting demonstrated that the autophagic indicator, LC3-II expression, was upregulated in AA-treated OGD cardiomyocytes compared with the OGD and OGD + DMSO groups (Fig. 3b). Congruously, the fluorescence results also revealed a significant increase in LC3-II positive puncta in AA-treated OGD cardiomyocytes (Fig. 3c, d). Additionally, the co-localization of LC3-II and mitochondria significantly increased in OGD cardiomyocytes after AA treatment, confirming the increased mitophagic activity in this in vitro model (Fig. 3c, e). These results revealed that AA may be responsible for the promoted mitophagy in ischemic myocardium and cardiomyocytes.
AMPK pathway is involved in AA-induced mitophagy
AMPK is a vital energy and mitophagic modulator of most tissue cells, especially under hypoxic environments [26]. Therefore, we investigated AMPK activation in OGD cardiomyocytes and MI cardiac tissue. We found upregulated expression of p-AMPKα at threonine 172 in OGD cardiomyocytes, and AA treatment further improved the activation of AMPK (Fig. 4a). In addition, there was no significant difference in AMPKα mRNA expression in cardiomyocytes between the AA-treated group and controls (Fig. 4b). Similar results were also observed in the myocardial tissue (Fig. 4c, d). To determine whether AMPK is associated with AA roles in mitophagy, shRNA was used to inhibit AMPK expression (Fig. 4e, f). As expected, AMPK deletion completely reversed the increased LC3-II expression in AA-treated OGD cardiomyocytes (Fig. 4g) and inhibited the increased mitophagic activity induced by AA (Fig. 4h–j). These results suggest that AMPK-regulated mitophagy may be one of the mechanisms by which AA protects the ischemic heart.
AA treatment upregulates glycophagy in OGD cardiomyocytes
The regulation of glycogen degradation and utilization (glycophagy) in AA-treated ischemic myocardium and cardiomyocytes was also investigated. As shown in Fig. 5a, glycophagosomes sequestering the glycogen particles were observed in the myocardium of the MI + AA group (orange arrows). Co-immunoprecipitation was performed to explore the effects of the combination of STBD1 and GABARAPL1, which is critical for the delivery of glycogen to lysosomes in the autophagic pathway [20]. The expression of both STBD1 and GABARAPL1 was elevated in the AA-treated OGD group compared with the OGD and OGD + DMSO group (cell lysates) (Fig. 5b). Similar results were observed by pulling down STBD1. The binding and interaction of STBD1 and GABARAPL1 were significantly enhanced in the OGD + AA group, indicating promoted glycophagy. To evaluate the utilization of glucose produced by glycophagy, ECAR was analyzed using the Seahorse XF96 extracellular flux analyzer to determine the glycolytic activity in cardiomyocytes (Fig. 5c). Glycolysis was significantly promoted in AA-treated OGD cardiomyocytes compared with OGD and OGD + DMSO groups. Furthermore, to evaluate the relationship between glycophagy and the protective effects of AA, shRNA inhibiting STBD1 expression was constructed to decrease STBD1 expression and STBD1-GABARAPL1 binding in OGD cardiomyocytes (Fig. 5d); we found that STBD1 depletion significantly abolished the promoted ECAR in AA-treated cardiomyocytes (Fig. 5c).
PI3K/Akt pathway is responsible for AA-induced glycophagy
The expression and interaction of STBD1 and GABARAPL1 are associated with the activation of PI3K/Akt axis [27–29]. Thus, the downstream mechanisms of glycophagy induced by AA in OGD cardiomyocytes were determined. The protein expression of phosphorylated Akt at serine 473 and phosphorylated GSK3β at serine 9 were markedly elevated compared with the OGD and OGD + DMSO groups (Fig. 6a, b). Importantly, the specific PI3K inhibitor LY294002 and Akt activation inhibitor GSK690693 abolished the effects of AA on upregulation of STBD1 and GABARAPL1 expression (Fig. 6c). In addition, the interaction between STBD1 and GABARAPL1 was also inhibited. Inhibition of PI3K/Akt signaling also abolished the increased glycolysis achieved by AA (Fig. 6d).
Glycophagy and mitophagy are key factors for AA to protect ischemic cardiomyocytes
Finally, we determined whether AMPK-regulated mitophagy and PI3K/Akt-regulated glycophagy are involved in AA-mediated cardioprotection and energy metabolism under OGD. Inhibition of the PI3K/Akt pathway (LY294002 and GSK690693) significantly elevated LDH release and decreased cell viability compared with the AA group (Fig. 7a, b). Consistently, depletion of AMPK expression (shAMPK) also abolished the protective roles of AA. Importantly, simultaneous suppression of PI3K/Akt and AMPK (LY294002 + shAMPK or GSK690693 + shAMPK) further aggravated the AA-treated cardiomyocyte injury compared with single PI3K/Akt inhibition or AMPK inhibition. Similar results were also observed in the TUNEL assay and in energy metabolic analysis in vitro (Fig. 7c–e). No significant effects of single-use of shAMPK and PI3K/Akt inhibitors on cell function were observed (Fig. 7 and Fig. S1). These results reveal that mitophagy regulated by AMPK and glycophagy modulated by PI3K/Akt signaling are both involved in AA-induced myocardial protection and energy metabolism.
Discussion
Myocardial infarction is the leading cause of human death worldwide. Altered mitochondrial function and energy metabolism in cardiomyocytes contribute to the pathophysiology of heart injury caused by MI [3]. However, at present, drugs for myocardial infarction, based on ameliorating energy imbalance, are scarce. In this study, we demonstrated that 25 mg·kg−1·d−1 AA in vivo and 10 μmol/L AA in vitro significantly promoted cardiac function and energy metabolism, and reduced infarct size and cell injury; that the AA-induced cardioprotective effects on ischemic myocardium and cardiomyocytes were related to the enhanced mitophagy and glycophagy flux; that AMPKα activation was involved in AA-mediated mitophagy in MI; and that STBD1/GABARAPL1 binding and PI3K/Akt axis were responsible for glycophagy regulated by AA in the ischemic myocardium and cardiomyocytes (Fig. 8). These findings extend previous knowledge and provide insights into the mechanisms underlying the cardioprotection conferred by AA, supporting the fact that AA may be a drug candidate for the treatment of ischemic heart disease.
In the infarcted myocardium, mitochondrial damage results in decreased ATP and PCr level and lower myocardial energy reserve [4]. Ameliorating the energy imbalance may increase the survival rate of cardiomyocytes in the infarct zone, allowing more cells to recover after the blood supply is restored. Various molecular compounds and proteins have ameliorated the energy metabolism disorder in infarcted myocardium, which provides new insights into MI treatment [30–32]. Our research revealed that AA, a pentacyclic triterpene, improved mitophagy- and glycophagy-related energy metabolism of ischemic cardiomyocytes and reduced the infarct size, thereby expanding the possibilities of AA application in the field of MI. In addition, our study also suggests that the optimal concentration and dosage of AA to protect mouse ischemic cardiomyocytes in vitro and in vivo are 10 μmol/L and 25 mg·kg−1·d−1, respectively. Compared with the low-dose group, an AA concentration higher than 10 μmol/L or 25 mg·kg−1·d−1 showed diminished protective effects on cell apoptosis and energy metabolism, but no toxic effects on cardiomyocytes were observed at high doses. Previous reports revealed that AA reduces ischemic myocardial injury by modulating inflammatory response and glycometabolism [33, 34]. In this study, we elucidated a new mechanism underlying the AA-mediated cardioprotection; specifically, AA improves energy metabolism by regulating mitophagy to maintain mitochondrial function and by modulating glycophagy to increase glycogen utilization.
Autophagy is a lysosomal degradation pathway that plays an important role in the maintenance of cellular homeostasis and the recycling of energy and material [35]. Although autophagy has been considered a nonselective process, accumulating evidence suggests the existence of selective autophagy through damaged organelles, including mitochondria, endoplasmic reticulum, ribosomes, and peroxisomes [36, 37]. Among them, mitochondria-specific autophagy, termed mitophagy, has been most extensively studied, and is most closely related to energy homeostasis in ischemic cardiomyocytes [3]. Mitochondrial quality is an important determinant of cell viability, and mitophagy plays a crucial role in this control mechanism. Genetic or pharmacological inhibition of autophagy has been shown to exacerbate myocardial ischemic injury and chronic cardiac remodeling in MI mouse models [38, 39]. Conversely, activation of autophagy limits myocardial damage in response to ischemia and reduces chronic ischemic remodeling and heart failure [39, 40]. However, the excessive activation of mitophagy may weaken its cardioprotective effects due to the loss of mitochondrial quantity and function. Therefore, it is necessary to prevent the overactivation of pharmacological or genetic mitophagy. Our results demonstrated that 10 μmol/L or 25 mg·kg−1·d−1 AA can moderately activate mitophagy in ischemic cardiomyocytes, thereby reducing the excessive consumption of intracellular energy and promoting cardiomyocyte survival.
In mammalian cells, several types of autophagy, including mitophagy, are regulated by the AMPK pathway [26]. AMPK can directly phosphorylate ULK1 on Ser467, Ser555, Thr574, and Ser637 [13, 41], and it plays a crucial regulatory role in mitophagy under various conditions [42–44]. In our study, the activation of AMPK increased under ischemic conditions in cardiomyocytes, and p-AMPKα expression was further elevated, indicating enhanced activation of the AMPK signaling pathway in the AA-treated MI heart. Inhibition of AMPK expression abolished all of the effects of AA on mitophagy, indicating that AMPK may be a central regulator connecting AA and mitophagy. LC3, one of the mammalian Atg8 orthologs, is typically used as an indicator of autophagic or mitophagic flux [45, 46]. AMPK is necessary and sufficient for LC3 lipidation (LC3-I to LC3-II) to induce autophagosome formation [15], which is consistent with our findings. However, further studies are needed to clarify the specific mechanism downstream of AMPK in AA-treated ischemic cardiomyocytes.
There have been few studies on glycophagy in myocardial infarction; nevertheless, glycophagy may play an important role in maintaining the energy supply of cardiomyocytes after MI. Cytosolic glycogen is localized in the myocyte subcellular regions, including intramyofibrillar, intermyofibrillar, and subsarcolemmal compartments [47]. Glycophagy is a glycogen degradation and utilization process, which triggers the liberation of free glucose that can be rapidly utilized by cells with energy requirements [16]. In the setting of a local high demand for glycogen, glycophagy may support the cytosolic machinery and metabolism in the myocardium under ischemic stress. In addition to increased mitophagic levels, an increased number of glycophagosomes was observed, indicating promoted glycophagy. The increased glycophagic level elevates the content of intracellular free glycogen, thereby providing the necessary energy supply for ischemic cardiomyocytes and maintaining their survival. Here, we verified the role of AA in glycophagy through the PI3K/Akt axis. Mechanistically, the interaction of STBD1 and GABARAPL1 and activation of the PI3K/Akt pathway in OGD cardiomyocytes were also enhanced by AA treatment. Furthermore, loss-of-function analysis suggested that the binding of STBD1 and GABRAPL1 was weakened by a PI3K or an Akt inhibitor. These results demonstrated that the PI3K/Akt axis and STBD1-GABARAPL1 interaction are responsible for the effects of AA on glycophagy in OGD cardiomyocytes. In addition, it is intriguing to observe that LY294002 and GSK690693 inhibit the phosphorylation of Akt at the same level, as well as GSK3β, suggesting that Akt and GSK3β may be comparably activated and that the PI3K/Akt/GSK3β axis is responsible for the AA-mediated glycophagy in OGD cardiomyocytes. However, further studies are needed to unravel how GSK3β affects STBD1. Inactivation of PI3K/Akt and/or AMPK also deteriorated cell injury, indicating that both glycophagy and mitophagy are critical for the cardioprotective roles of AA. Therefore, AA may be a multieffect cardioprotective regulator that increases glycogen utilization by promoting glycophagy and glycolysis and also enhances cardiomyocyte resistance to ischemia, and reduces excessive energy consumption by promoting mitophagy. The PI3K/Akt and AMPK pathways have overlapping effects on myocardial protection [48, 49]; thus, the upstream pathways of the two after AA treatment are worth further investigation.
In conclusion, our results demonstrated that AA improved recovery from myocyte injury, myocardial dysfunction, and energy metabolism disorders after MI by regulating mitophagy to maintain mitochondrial homeostasis and by modulating glycophagy and glycolysis to increase glycogen utilization.
Supplementary information
Acknowledgements
This work was supported by the National Key Research and Development Program of China (2017YFA0105600, 2020YFA0112600), the National Natural Sciences Foundation of China (81870208, 82070260), the Shanghai Pujiang Program (19PJ1408900), the Shanghai Animal Research Program (21140901800), the Science and Technology Development Fund of Shanghai Pudong New Area (PKJ2020-Y23), Shanghai Engineering Research Center of Artificial Heart and Heart Failure Medicine (19DZ2251000), the Major Program of Development Fund for Shanghai Zhangjiang National Innovation Demonstration Zone < Stem Cell Strategic Biobank and Stem Cell Clinical Technology Transformation Platform > (ZJ2018-ZD-004), and the Peak Disciplines (Type IV) of Institutions of Higher Learning in Shanghai.
Author contributions
FQ analyzed the data, designed the study, and wrote the manuscript. FQ, YY, WL, and YSG performed the experiments. ZMZ, ZML, and LG provided technical support and substantial contributions to the conception and design of the study. ZML and LG directed the study, managed the project, interpreted the data, and co-wrote the manuscript.
Competing interests
The authors declare no competing interests.
Contributor Information
Zhong-min Liu, Email: liu.zhongmin@tongji.edu.cn.
Ling Gao, Email: gaoleng5@126.com.
Supplementary information
The online version contains supplementary material available at 10.1038/s41401-021-00763-9.
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