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Brief periods of ischemia and reperfusion that precede sustained ischemia lead to a reduction in myocardial infarct size. This phenomenon, known as ischemic preconditioning, is mediated by signaling pathway(s) that is complex and yet to be fully defined. AMP-activated kinase (AMPK) is activated in cells under conditions associated with ATP depletion and increased AMP/ATP ratio. In the present study, we have taken advantage of a cardiac phenotype overexpressing a dominant negative form of the α2 subunit of AMPK to analyze the role, if any, that AMPK plays in preconditioning the heart. We have found that myocardial preconditioning activates AMPK in wild type, but not transgenic mice. Cardiac cells from transgenic mice could not be preconditioned, as opposed to cells from the wild type. The cytoprotective effect of AMPK was not related to the effect that preconditioning has on mitochondrial membrane potential as revealed by JC-1, a mitochondrial membrane potential-sensitive dye, and laser confocal microscopy. In contrast, experiments with di-8-ANEPPS, a sarcolemmal-potential sensitive dye, has demonstrated that intact AMPK activity is required for preconditioning-induced shortening of the action membrane potential. The preconditioning-induced activation of sarcolemmal KATP channels was observed in wild type, but not in transgenic mice. HMR 1098, a selective inhibitor of sarcolemmal KATP channels opening, inhibited preconditioning-induced shortening of action membrane potential as well as cardioprotection afforded by AMPK. Immunoprecipitation followed by Western blotting has shown that AMPK is essential for preconditioning-induced recruitment of sarcolemmal KATP channels. Based on the obtained results, we conclude that AMPK mediates preconditioning in cardiac cells by regulating the activity and recruitment of sarcolemmal KATP channels without being a part of signaling pathway that regulates mitochondrial membrane potential.
AMP-activated kinase (AMPK) is activated by stress and in conditions associated with ATP depletion and a consequent increase in the AMP/ATP ratio. Once activated, AMPK phosphorylates several downstream substrates, the overall effect of which is to switch off ATP-consuming pathways (e.g., fatty acid synthesis and cholesterol synthesis) and to switch on ATP-generating pathways (Carling et al., 1989; Hardie, 2003; Carling, 2004). So far, it has been established that AMPK, directly or indirectly, regulate the activity/expression of many intracellular proteins, including acetyl-coenzyme A carboxylase, 3-hydroxy-3-methylglutaryl-coenzyme A reductase, glycerol phosphate acyl transferase, glycogen synthase, elongation factor-2, mammalian target of rapamycin, p70 ribosomal protein S6 kinase, hormone-sensitive lipase, cystic fibrosis transmembrane regulator (CFTR), 6-phosphofructo-2-kinase, endothelial nitric oxide synthase, insulin receptor substrate-1, phosphatidylinositol 3-kinase, glucose transporter 1 and 4, phosphoenolpyruvate carboxykinase, glucose-6-phosphatase, fatty acid synthase, and probably many others that will be identified in the future (for detailed description of AMPK targets, see Hardie, 2003; Hardie et al., 2003). In the heart, AMPK is activated by different types of stress, both of those occurring under physiological and pathophysiological conditions (Kudo et al., 1995; Coven et al., 2003; Sakamoto et al., 2004; Young et al., 2005). AMPK is a heterotrimeric complex comprising of a catalytic α subunit and regulatory β and γ subunits. There are two isoforms of the catalytic subunit termed α1 and α2. α2 subunit is primarily expressed in liver, heart, and skeletal muscle (Hardie, 2003; Carling, 2004). It has been previously shown that mice overexpressing dominant negative α2 subunit in the heart respond to myocardial ischemia by accelerated ATP depletion, early development of myocardial contracture, and decreased glucose uptake (Xing et al., 2003).
Murray et al. (1986) discovered that brief periods of blood vessel occlusion and reperfusion administered prior to a sustained ischemic episode lead to a reduction in infarct size. This cardioprotective phenomenon (now called early preconditioning) enhances the survival of cardiac cells under conditions that induce myocardial infarction. This and subsequent studies have indicated that brief ischemia or hypoxia are likely to switch on intracellular signaling pathways that ultimately result in an increased cellular tolerance to metabolic stress (Yellon and Downey, 2003). As AMPK is activated in the heart by hypoxia/ischemia (for review see Russell, 2003), we have hypothesized that this enzyme, and in particular α2 isoform that is predominant in the heart, might play a role in mediating preconditioning in the heart.
Therefore, we have taken advantage of the cardiac phenotype overexpressing dominant negative form of α2 AMPK subunit to analyze the role, if any, that AMPK play in preconditioning in the heart.
All experiments have been done on male mice overexpressing dominant negative α2 subunit of AMPK (transgenics) and male littermate controls expressing wild type of α2 subunit of AMPK (wild type). Generation, breeding, phenotype characteristics, and genotyping of these mice have previously been described in detail (Xing et al., 2003).
Mice were killed by cervical dislocation (according to UK Home Office procedures), and the heart rapidly removed and placed in ice-cold Tyrode’s solution at 4°C. The aorta was then cannulated and secured using silk suture and the heart attached to a Langendorff perfusion apparatus. Hearts were perfused at a constant flow rate of 5 ml/min at 37°C with oxygenated (95% O2, 5% CO2) Tyrode’s solution (in mM: NaCl 136.5, KCl 5.4, CaCl2 1.8, MgCl2 0.53, glucose 5.5, HEPES-NaOH 5.5, pH 7.4) for a period of 70 min or 30 min for preconditioned hearts. Preconditioning was induced by four cycles of 5 min ischemia (achieved by placing the heart into Tyrode’s solution degassed with 100% Ar at 37°C and switching off perfusion) and 5 min reperfusion. After the stabilization period or preconditioning protocol, the heart was subjected to 5 min of global ischemia. Hearts were snap-frozen in liquid nitrogen and stored at −80°C (Du et al., 2006). Freeze-clamped heart tissues were pulverized to a powder in liquid nitrogen. A 15-fold mass excess of ice-cold Lysis Buffer containing 50 mM Tris/HCl pH 7.5, 1 mM EGTA, 1 mM EDTA, 1% (by mass) Triton-X 100, 1 mM sodium orthovanadate, 50 mM sodium fluoride, 5 mM sodium pyrophosphate, 0.27 M sucrose, 0.1% (by volume) 2-mercaptoethanol, and “complete” proteinase inhibitor cocktail (obtained from Roche, 1 tablet per 50 ml) was added to the powder tissue and homogenized on ice using KINEMATICA POLYTRON (Brinkmann, CT). Homogenates were centrifuged at 13,000g for 10 min at 4°C to remove insoluble material. The supernatant was collected and protein concentration measured by the Bradford Method using bovine serum albumin as the standard. Lysates were snap frozen in aliquots in liquid nitrogen and stored at −80°C. The specific AMPKα1 and AMPKα2 antibodies were kindly donated by Grahame Hardie (University of Dundee). Lysates (50 μg) were used to immunoprecipitate AMPKα1 or AMPKα2. The lysates were incubated at 4°C for 1 h on a shaking platform with 5 μl of protein G-Sepharose coupled to 2 μg of AMPKα1 or AMPKα2 antibody. The immunoprecipitates were washed twice with 1 ml of Lysis Buffer containing 0.5 M NaCl, and twice with 1 ml of Buffer A (50 mM Tris/HCl pH 7.5, 0.1 mM EGTA, and 0.1% (by volume) 2-mercaptoethanol). Phosphotransferase activity towards AMARA peptide were then measured as described previously (Sakamoto et al., 2005, 2006).
Ventricular cardiomyocytes were dissociated from wild type and transgenic mice using an established enzymatic procedure (Mora et al., 2003; Du et al., 2006). In brief, hearts were retrogradely perfused (at 37°C) with medium 199, followed by Ca2+-EGTA-buffered low-Ca2+ medium (pCa = 7), and finally low-Ca2+ medium containing pronase E (8 mg/100 ml), proteinase K (1.7 mg/100 ml), bovine albumin (0.1 g/100 ml, fraction V), and 200 μM CaCl2. Ventricles were cut into fragments in the low-Ca2+ medium enriched with 200 μM CaCl2. Cells were isolated by stirring the tissue (at 37°C) in a low-Ca2+ medium solution containing pronase E and proteinase K supplemented with collagenase (5 mg/10 ml). The first aliquot was removed, filtered through a nylon sieve, centrifuged for 60 sec (at 300-400 rpm), and washed. Remaining tissue fragments were re-exposed to collagenase, and isolation continued for two to three such cycles (the duration of each cycle was 4-5 min).
Hypoxia of isolated cardiomyocytes has been performed as described previously (Mora et al., 2003; Jovanović et al., 2006a). Cardiomyocytes were placed into Tyrode’s solution (in mM: NaCl 136.5, KCl 5.4, CaCl2 1.8, MgCl2 0.53, glucose 5.5, HEPES-NaOH 5.5, pH 7.4), plated out on glass coverslips, and paced to beat by field stimulation (parameters of the stimulation: 5-20 mV depending on cellular threshold, 5 msec, 1 Hz). Beating cardiomyocytes were perfused with Tyrode solution at a rate of 3 ml/min, and under these conditions, the partial pressure of O2 (PO2) in perfusate was 140 mmHg. To induce preconditioning, we have exposed cardiomyocytes to four episodes of 5 min-long hypoxia/5 min-long reoxygenation followed by hypoxia (termed here as long-lasting hypoxia) that was maintained until cell death (Budas et al., 2004). To induce hypoxia, Tyrode solution was bubbled with 100% argon (PO2 = 20 mmHg). The moment of cell death was defined as the point when cells have become rounded (ratio of diameters <2; normal ratio of diameters is >4, Jovanović et al., 1998; Mora et al., 2003). A time point when 50% of cells survived hypoxia was calculated by linear interpolation of the obtained data for each experiment.
Cell morphology, size parameters, and sarcolemmal/mitochondrial membrane potential were monitored in cells exposed to the preconditioning/hypoxia as previously described (Du et al., 2006). To measure sarcolemmal membrane potential, cells were loaded with di-8-ANEPPS according to the manufacturer’s instruction (Invitrogen, Paisley, UK), and imaged using laser confocal microscopy in line-scan mode (LSM-510, Zeiss, Gottingem, Germany). Cells were scanned under control conditions and then exposed to hypoxia with and without preconditioning. Cells were scanned at 20 min after the beginning of hypoxia. Fluorescence was detected/imaged at 488 nm excitation wavelength and emission was captured at >505 nm. To measure the mitochondrial membrane potential, cells were loaded with JC-1 according to the manufacturer’s instructions (Invitrogen) and continuously monitored with LSM-510. Fluorescence was imaged at 488 nm excitation wavelength and emission was captured at 530 and 590 nm for red and green channels, respectively.
Cells were superfused with Tyrode solution. Pipettes (resistance 3-5 M), were filled with (in mM): KCl 140, MgCl2 1, HEPES-KOH 5, amphotericin B (Sigma, 240 μg/ml) (pH 7.3). The membrane potential was held at −40 mV and the currents were evoked by a 400 msec current step (to +80 mV) recorded directly to hard disk using an Axopatch-200B amplifier, Digidata-1321 interface and pClamp8 software (Axon Instruments, Inc., Foster City, CA) (Budas et al., 2004; Jovanović et al., 2006a).
Total RNA was extracted from cardiac ventricular tissue of transgenic and wild type mice using TRIZOL reagent (Invitrogen) according to the manufacturer’s recommendations. Extracted RNA was further purified with RNeasy Mini Kit (Qiagen, Crawley, UK) according to the manufacturer’s instruction. The specific primers for mouse SUR2A and Kir6.2 were designed as follows: For SUR2A: sense, ACTATGGAGTCCGAGAACTA; antisense, AGGTTTGGACCAGTATCACA; for Kir 6.2: sense, ACATGCAGGTGGTGCGCAAG; antisense, AGGGCATCCAGCAGACTGCG. Both set of primers have been previously established to be highly efficient (90.4 and 98.6% of PCR efficiency for SUR2A and Kir 6.2, respectively) and to produce a single product. The reverse transcription (RT) reaction was carried out with ImProm-II Reverse Transcriptase (Promega, Southampton, UK). A final volume of 20 μl of RT reaction containing 4 μl of 5× buffer, 3 mM MgCl2, 20 U of RNasin® Ribonuclease inhibitor, 1 U of ImProm-II reverse transcriptase, 0.5 mM each of dATP, dCTP, dGTP, and dTTP, 0.5 μg of oligo(dT), and 1 μg of RNA was incubated at 42°C for 1 h and then inactivated at 70°C for 15 min. The resulting cDNA was used as a template for real-time PCR. A SYBR Green I system was used for the RT-PCR and the 25 μl reaction mixture contained: 12.5 μl of iQ™ SYBR® Green Supermix (2×), 7.5 nM each primers, 9 μl of ddH2O, and 2 μl of cDNA. The thermal cycling conditions were as follows: an initial denaturation at 95°C for 3 min, followed by 40 cycles of 10 sec of denaturing at 95°C, 15 sec of annealing at 56°C, and 30 sec of extension at 72°C. The real-time PCR was performed in the same wells of a 96-well plate in the iCycler iQ™ Multicolor Real-Time Detection System (Bio-Rad, Hercules, CA). Data was collected following each cycle and displayed graphically (iCycler iQ™ Real-time Detection System Software, version 3.0A, Bio-Rad). Primers were tested for their ability to produce no signal in negative controls by dimer formation and then with regard to the efficiency of the PCR reaction. Efficiency is evaluated by the slope of the regression curve obtained with several dilutions of the cDNA template. Melting curve analysis tested the specificity of primers. Threshold cycle values, PCR efficiency (examined by serially diluting the template cDNA and performing PCR under these conditions), and PCR specificity (by constructing the melting curve) were determined by the same software. Each mouse cDNA sample was measured at three different quantities (and duplicated at each concentration, the corresponding no-RT mRNA sample was included as a negative control, Du et al., 2006; Jovanović et al., 2006b).
For Western blotting experiments, myocardial ischemia/preconditioning have been done as described in “Preconditioning and AMPK activity assay” section. For Western blot analysis, control hearts (hearts perfused with oxygenated Tyrode’s solution for 90 min) or hearts perfused for 30 min plus four cycles of 5 min ischemia/5 min reperfusion plus 20 min in ischemia were analyzed. Rabbit anti-SUR2 and anti-Kir6.2 antibodies were used for immunoprecipitation and Western blotting in this study (Santa Cruz, CA). Ventricular tissue was snap-frozen immediately upon extraction and ground to a powder under liquid nitrogen. The powder was resuspended in 10 ml of tissue buffer (20 mM HEPES, 150 mM NaCl, 1% Triton-X 100, pH 7.5) and homogenized. To obtain the membrane and cytosolic cardiac fraction, mouse ventricular tissue was homogenized in buffer I (TRIS 10 mM, NaH2PO4 20 mM, EDTA 1 mM, PMSF 0.1 mM, pepstatin 10 μg/ml, leupeptin 10 μg/ml, at pH 7.8) and incubated for 20 min (at 4°C). The osmolarity was restored with KCl, NaCl, and sucrose, and the obtained mixture was centrifuged at 500g. The supernatant was diluted in buffer II (imidazole 30 mM, KCl 120 mM, NaCl 30 mM, NaH2PO4 20 mM, sucrose 250 mM, pepstatin 10 μg/ml, leupeptin 10 μg/ml, at pH 6.8) and centrifuged at 7,000g, the pellet removed and supernatant centrifuged at 30,000g. The obtained pellet and supernatant contain the membrane and cytosol fraction, respectively. Protein concentration was determined using Bradford assay. Ten micrograms of the epitope-specific Kir6.2 antibody or 40 μg of the epitope-specific SUR2A antibody was prebound to Protein-G Sepharose beads and used to immunoprecipitate from 50 μg of membrane/cytosol fraction protein extract. The pellets of this precipitation were run on SDS polyacrylamide gels for Western analysis. Western blot probing was performed using 1/1,000 dilution of anti-Kir 6.2 antibody, and detection was achieved using Protein-G HRP and ECL reagents. The band intensities were analyzed using the Quantiscan software (Ranki et al., 2002a; Crawford et al., 2003).
Data are presented as mean±SEM, with n representing the number of experiments. The difference between means were assessed using Student’s t-test (non-paired and paired where appropriate), Fisher exact test, Mann-Whitney rank sum test, or by ANOVA on ranks using SigmaStat program (Jandel Scientific, Chicago, IL). P<0.05 was considered statistically significant.
In wild type, preconditioning induced increase in both α1 and α2 AMPK activity (α1 AMPK activity was 14.0±1.3 mU/mg before and 39.3±6.7 mU/mg after preconditioning, n=3, P=0.02; α2 AMPK activity was 10.4±0.2 mU/mg before and 17.7±2.2 mU/mg after preconditioning, n=3, P=0.03, Fig. 1). In contrast, in transgenic mice, preconditioning failed to activate AMPK (α1 AMPK activity was 23.3±7.0 mU/mg before and 25.0±1.8 mU/mg after preconditioning, n=3 P=0.82; α2 AMPK activity was 7.4±2.2 mU/mg before and 9.8±0.1 mU/mg after preconditioning, n=3, P=0.33, Fig. 1). To test whether preconditioning-induced activation of AMPK is associated with cardioprotection, we have used an experimental model of hypoxia and preconditioning that utilize field-stimulated, adult cardiomyocytes, a pure myocardial preparation free of neuronal, vascular and humoral influences (Mora et al., 2003; Budas et al., 2004). Preconditioning significantly increased cellular resistance to hypoxia in wild type, that is, 50% of preconditioned cells were able to survive for 61.7± 6.0 min while the same percentage of non-preconditioned cells survived only 19.3±10.0 min (Fig. 1; n=3, P=0.02). In contrast, preconditioning was inefficient in cells from transgenic mice (Fig. 1; 50% of cells survived hypoxia for 28.3± 10.9 min without and 31.0±7.0 min with preconditioning, n=3 for each, P=0.85).
It has been proposed that preconditioning may activate putative mitochondrial KATP channels and/or inhibit the mitochondrial permeability transition pore (mPTP) (Gross and Peart, 2003; Halestrap et al., 2004). In both cases, the mitochondrial membrane potential would change. Therefore, we have monitored mitochondrial membrane potential on-line in beating cardiomyocytes from wild type and transgenic mice using the fluorescent dye JC-1, a ratiometric dye which is established as a reliable indicator of mitochondrial membrane potential (Reers et al., 1995). During hypoxia alone, the JC-1 ratio steadily rose in both phenotypes and the membrane depolarization reached statistical significance after 15 min of hypoxia (Fig. 2). In both cell types, four cycles of 5 min hypoxia/reoxygenation induced mitochondrial membrane depolarization (Fig. 3A). When preconditioned cells were exposed to sustained hypoxia, JC-1 ratio has risen further and has achieved statistical significance after only 5 min of hypoxia (Fig. 3B). In both cell types, regardless whether they were preconditioned or not, the moment of cell death was associated with the abrupt increase in JC-1 ratio (Fig. 3C).
It has been shown that preconditioning induces activation of sarcolemmal KATP channels, which would shorten action membrane potential duration and decrease intracellular Ca2+ loading, leading to cardioprotection (Suzuki et al., 2002; Budas et al., 2004). Preconditioning activates sarcolemmal KATP channels in hypoxia in single beating cardiomyocytes (Budas et al., 2004). Therefore, we have measured membrane potential under control conditions and in hypoxia with and without preconditioning. In non-preconditioned cardiomyocytes from the wild type, 20-min-long hypoxia induced no significant changes in duration of action membrane potential (the action potential duration (apd) was 0.38±0.02 sec under control conditions and 0.38±0.06 sec in hypoxia, n=4), P=0.99, Fig. 4). Preconditioning shortened duration of action membrane potential in wild type (control apd was 0.38±0.06 msec and apd in hypoxia after preconditioning was 0.29±0.04 sec, n=4, P=0.009, Fig. 4). Non-preconditioned cells from transgenic mice responded to hypoxia similarly to non-preconditioned cells from the wild type (apd was 0.36±0.06 sec under control conditions and 0.41±0.04 sec in hypoxia, n=4, P=0.49, Fig. 5). However, in contrast to the wild type, preconditioning had no effect on action membrane potential duration in cardiac cells from transgenic mice (control apd was 0.33±0.04 msec and apd in hypoxia after preconditioning was 0.43±0.07 sec, n=4, P=0.38, Fig. 5).
It has been recently shown that the activation and recruitment of these channels mediates preconditioning (Budas et al., 2004). As the preconditioning was not associated with the shortening of action membrane potential in transgenic mice, we have hypothesized that the channel activity and the channel recruitment may be impaired in mice overexpressing dominant negative α2 AMPK. Indeed, perforated patch clamp electrophysiology has revealed that whole cell K+ current is increased by preconditioning in cells from the wild type, but not in cells from transgenic mice (Fig. 6A). In addition, HMR 1098 (30 μM), a selective inhibitor of the opening of sarcolemmal KATP channel (Gogelein et al., 2001), has prevented preconditioning-induced shortening in action membrane potential in cardiomyocytes with intact AMPK (Fig. 6B). At the same time, HMR 1098 (30 μM) did not affect properties of action membrane potential in transgenic cells during preconditioning (Fig. 6B). A blockade of the channel opening with HMR1098 (30 μM) has inhibited the preconditioning-induced cardioprotection (Fig. 7; 50% of preconditioned cells survived hypoxia for 61.7±6.0 min in the absence of HMR 1098 and only 36.0±3.9 min in the presence of HMR 1098, n=3-4, P=0.01 ). HMR 1098 (30 μM) did not decrease survival of preconditioned cardiomyocytes from transgenic mice (Fig. 7; 50% of cells survived hypoxia for 31.0±7.0 min in the absence and 42.5±4.4 min in the presence of HMR 1098, n=3-4, P=0.20).
In addition to the channel activation, preconditioning-induced recruitment of sarcolemmal KATP channels is crucial for cardiac resistance to hypoxia (Budas et al., 2004). Therefore, we have examined whether AMPK is involved in preconditioning-induced stimulation of sarcolemmal KATP channels trafficking. Under control conditions, hearts from wild type and transgenic mice had similar mRNA levels of Kir6.2, the KATP channel pore-forming subunit, and SUR2A, the KATP channel regulatory subunit (Fig. 8; Kir6.2: average cycle threshold was 19.2±0.2 and 19.2±0.2 for wild type and transgenic mice, respectively, n=3, P>0.50; SUR2A: 22.8±0.3 and 22.7±0.1 for wild type and transgenic mice, respectively, n=3, P>0.50). We have probed anti-SUR2 immunoprecipitate of cardiac membrane and cytosol fraction with anti-Kir6.2 antibody. Using this strategy, we have measured only those Kir6.2 and SUR2A subunits that physically assemble in sarcolemma to form a channel (Ranki et al., 2001, 2002b). Western blotting has demonstrated that the levels of Kir6.2 in sarcolemma were not different between wild type and transgenic mice (Fig. 8; the band intensity was 21.0±4.0 and 23.0±5.0 in wild type and transgenic mice, respectively (n=3 for each, P>0.50). The sarcolemmal/cytosolic Kir6.2 ratio under control conditions was also not different between experimental groups (Fig. 8; the ratio was 0.63±0.02 and 0.68±0.09 for wild type and transgenics, n=3 for each, P=0.39). However, in wild type, preconditioning significantly increased sarcolemmal/cytosolic Kir6.2 ratio (Fig. 9, from 0.63±0.02 under control conditions to 1.03± 0.14, n=3 for each, P=0.04). In contrast, in transgenic mice, preconditioning did not significantly affect sarcolemmal/cytosolic Kir 6.2 ratio (Fig. 9, from 0.68±0.09 under control conditions to 0.76±0.14, n=3 for each, P=0.48).
It has been previously shown that overexpression of the dominant negative α2 subunit of AMPK blocks the activation of AMPK, reduces ischemia-stimulated glucose uptake in the heart and leads to an accelerated depletion of ATP/exacerbated diastolic dysfunction, possibly because of increased energy consumption (Xing et al., 2003). As shown in the present study, these effects of suppressed α2 AMPK activity did not result in increase in cardiac sensitivity to ischemia and hypoxia.
The role of AMPK in early preconditioning in the heart has never been studied before. In the present study, we have shown that preconditioning activates both AMPKα1 and AMPKα2 isoforms. We found that a lack of AMPK activation in transgenic mice was associated with a loss of preconditioning suggesting that the activation of AMPK is crucial for preconditioning-induced cardioprotection.
The underlying mechanism of preconditioning is complex and still a matter of vigorous research. The most recent studies have suggested that the preconditioning-induced cardioprotection might be mediated by inhibition of the opening of mPTP (Juhaszova et al., 2004), a non-specific megachannel in the inner mitochondrial membrane. The opening of mPTP results in collapse of the inner membrane potential, uncoupling of respiratory chain, efflux of cytochrome-c and other molecules which, all together, lead to cell death (Weiss et al., 2003; Halestrap et al., 2004). In accord with such view, we have shown that cell death was readily associated with dramatic mitochondrial membrane depolarization. Under any examined conditions, the response of mitochondria to hypoxia was indistinguishable between the experimental groups. In both wild type and transgenic mice, preconditioning has fostered mitochondrial membrane depolarization. It has been shown in Girardi cells and C2C12 miotubes that ischemic preconditioning might be associated with mitochondrial membrane depolarization, which was proposed to be due to activation of putative mitochondrial KATP channels (Minners et al., 2001). These channels were originally described in 1991 (Inoue et al., 1991) and it was suggested that their opening might mediate cardioprotection (Sato et al., 2000). At present, however, the identity/existence of this ion channel is a matter of vigorous discussion (Hanley and Daut, 2005). More recently, it has been suggested that other, non-KATP channel, K+ conductance(s) in mitochondria could be involved in mediating cardioprotection (O’Rourke et al., 2005). Our findings that preconditioning fosters mitochondrial membrane depolarization would be compatible with the idea that the activation of a K+ conductance could mediate preconditioning in the heart. However, as no difference was found between experimental groups in mitochondrial response to preconditioning, it seems that AMPK is not involved in the regulation of mitochondrial membrane potential during preconditioning.
It has been recently shown that preconditioning induces activation and trafficking of sarcolemmal KATP channels (Budas et al., 2004), which, in turn, decreases the duration of action membrane potential and Ca2+ influx, thus promoting cell survival during ischemia/hypoxia (Jovanović and Jovanović, 2001; Kane et al., 2005). In the present study, hypoxia alone did not significantly change the action potential duration in first 20 min in both types of mice and these results would be expected based on previous findings (Verkerk et al., 1996; Budas et al., 2004). Preconditioning has activated sarcolemmal KATP channels resulting in shortening of the action membrane potential, which is also in accord with previous studies (Budas et al., 2004). In contrast, preconditioning did not affect K+ current and action potential duration in cardiomyocytes isolated from the transgenic mice, suggesting that preconditioning-induced activation of sarcolemmal KATP channels require intact AMPK catalytic activity. That the activation of these channels is essential for preconditioning-induced cardioprotection was demonstrated by the findings that HMR 1098, a selective blocker of sarcolemmal KATP channels (Gogelein et al., 2001), inhibited cellular survival afforded by preconditioning in wild type, without having any effects on preconditioned transgenic cardiomyocytes.
It has been shown that a crucial event in mediating preconditioning is recruitment of sarcolemmal KATP channels. The signaling cascade responsible for regulation of KATP channels trafficking is still to be fully understood, but it has been suggested that protein kinase C inhibits recruitment of sarcolemmal KATP channels (Hu et al., 2003). Until now, no enzyme has been associated with the stimulation of recruitment of these channels. KATP channels are composed, in vivo, of Kir6.2 and SUR2A, a pore-forming, and regulatory subunits, respectively, and at least, four accessory proteins (Inagaki et al., 1996; Carrasco et al., 2001; Crawford et al., 2002a,b; Jovanović and Jovanović, 2005; Jovanović et al., 2005). It has been shown that the level of Kir6.2 protein in anti-SUR2 immunoprecipitate accurately reflects the number of KATP channels (Ranki et al., 2001, 2002a,b). In wild type, preconditioning has increased sarcolemmal/cytosolic Kir6.2 ratio confirming previous findings that preconditioning stimulates recruitment of sarcolemmal KATP channels (Budas et al., 2004). However, in transgenic mice no channel recruitment was observed in preconditioning suggesting that α2 AMPK catalytic activity is essential for preconditioning-induced stimulation of KATP channels recruitment to the sarcolemma.
It is generally accepted that AMPK is activated in response to metabolic stress usually associated with ATP depletion and consequent increase in the AMP/ATP ratio (Hardie, 2003; Carling, 2004). Being gated by intracellular ATP, sarcolemmal KATP channels are also activated by decrease in intracellular ATP and increase in intracellular AMP (Noma, 1983; Elvir-Mairena et al., 1996). Once activated, AMPK phosphorylates several downstream substrates, the overall effect of which is to switch off ATP-consuming pathways and to switch on ATP-generating pathways which would counteract energy depletion and thereby, act in a cardioprotective manner (Hardie, 2003; Carling, 2004). More recently, it has been suggested that AMPK may couple membrane transport with the cellular metabolism by inhibiting ion-transport proteins such as the cystic fibrosis transmembrane conductance regulator Cl- channel and the epithelial Na+ channel, thereby limiting the dissipation of transmembrane ion gradients and promoting cell survival (Hallows, 2005). The present finding that AMPK stimulates KATP channels activity/recruitment is an addition to the list of AMPK-governed events that result in the cytoprotective outcome. The activation of sarcolemmal KATP channels would decrease the duration of sarcolemmal action potential which, in turn, would decrease Ca2+ influx, thus preventing Ca2+ overload and cell death. In addition, as sarcolemmal KATP channel subunits physically associate with energy-producing enzymes (Carrasco et al., 2001; Carwford et al., 2002a,b; Jovanović et al., 2005), an increase in these channels also means an increase in sarcolemmal presence of crucial enzymes for cellular metabolism and energy production, which may promote cellular survival under conditions of metabolic stress. In support of this idea are reports suggesting that sarcolemmal KATP channels are likely to have functions additional to their channel property, such as glucose transport (Miki et al., 2002), which could contribute to cardioprotection evoked by preconditioning. In this respect, regulation of not only channel activity but also channel complex levels seem to be an important part of cardioprotective signaling in preconditioning.
We have recently found that preconditioning activates the phosphoinositide-dependent kinase-1 (PDK1)-protein kinase B (PKB)-glycogen synthase kinase-3β (GSK-3β) signaling cascade. This cascade regulates mitochondrial membrane potential and is involved in cardioprotection without having any effects on sarcolemmal membrane potential (Budas et al., 2006). As the inhibition of AMPK abolishes preconditioning-induced sarcolemmal membrane depolarization and cardioprotection without affecting preconditioning-induced changes in mitochondrial membrane potential, it seems that preconditioning activates in parallel at least two signaling pathways, PDK1-PKB-GSK-3β on one and AMPK-sarcolemmal KATP channels on the other side, which could act independently of each other. It is apparent that both of these pathways must be functional in order to confer preconditioning-induced cardioprotection. This idea is quite compatible with the recent report showing that PKB-GSK-3β are not involved in regulation of expression/recruitment of sarcolemmal KATP channels (Brown et al., 2005).
We thank Grahame Hardie for providing AMPK antibodies. We also thank Aventis Pharma (Frankfurt, Germany) for providing HMR 1098. This research was supported by grants from British Heart Foundation, BBSRC, MRC, TENOVUS-Scotland, Wellcome Trust and Anonymous Trust to A.J., and Grants from the National Institute of Health (HL67970) and Established Investigator Award from the American Heart Association to R.T.
Contract grant sponsor: British Heart Foundation; Contract grant sponsor: BBSRC; Contract grant sponsor: MRC; Contract grant sponsor: TENOVUS-Scotland; Contract grant sponsor: Wellcome Trust; Contract grant sponsor: Anonymous Trust; Contract grant sponsor: National Institute of Health; Contract grant number: HL67970; Contract grant sponsor: American Heart Association.