Cardiovascular Research

Cardiovascular Research 1997 35(2):273-282; doi:10.1016/S0008-6363(97)00092-8
© 1997 by European Society of Cardiology

This Article Abstract FREE Full Text (PDF) E-letters: Submit a response Alert me when this article is cited Alert me when E-letters are posted Alert me if a correction is posted Services Email this article to a friend Similar articles in this journal Similar articles in PubMed Alert me to new issues of the journal Add to My Personal Archive Download to citation manager Request Permissions Disclaimer Google Scholar Articles by Shigematsu, S. Articles by Arita, M. Search for Related Content PubMed PubMed Citation Articles by Shigematsu, S. Articles by Arita, M. Social Bookmarking          What's this?

Copyright © 1997, European Society of Cardiology

Anoxia-induced activation of ATP-sensitive K⁺ channels in guinea pig ventricular cells and its modulation by glycolysis

Sakuji Shigematsu^* and Makoto Arita

Department of Physiology, Oita Medical University, Hasama, Oita 879-55, Japan

^* Corresponding author. Tel.: +81 (975) 86-5652; fax: +81 (975) 49-6046.

Received 13 May 1996; accepted 3 March 1997

	Abstract

Top Abstract 1 Introduction 2 Methods 3 Results 4 Discussion References

 
Objective: Exposure to anoxia has been reported to activate ATP-sensitive potassium (K⁺_ATP) channels in isolated ventricular myocytes. We aimed to investigate the mechanisms underlying the anoxia-induced activation of K⁺_ATP channels. Methods: Guinea pig ventricular myocytes were isolated using collagenase digestion. Action potentials and membrane currents were recorded in the whole-cell mode of patch clamp. Exposure to anoxia was performed in a semi-closed airtight chamber, which prevented the diffusion of atmospheric oxygen into anoxic perfusate. Results: Exposure to glucose-free anoxia shortened the action potential duration (APD) to less than 20% of control in 13±3 min. Subsequent reoxygenation rapidly and completely restored the APD. The time-independent large outward current which developed during anoxia was completely suppressed by reoxygenation or by the application of glibenclamide, a K⁺_ATP channel blocker. The presence of extracellular glucose did not prevent APD shortening during anoxia, although it significantly decreased the rate of shortening. Reoxygenation-induced restoration of the APD was inhibited after a long-lasting anoxia. In addition, repeated exposures to anoxia/reoxygenation progressively impaired the recovery of APD during reoxygenation. Conclusions: Activation of K⁺_ATP channels occurs during anoxia. The primary source of ATP that regulates the channel activity seems to be oxidative phosphorylation. ATP derived from anaerobic glycolysis (attained by the increase of extracellular glucose) was observed to partially suppress the channel activity only when oxidative phosphorylation was severely impaired during anoxia.

KEYWORDS Anoxia; Reoxygenation; Potassium channel, ATP-sensitive; Patch clamp; Ventricular myocyte; Glucose; Oxidative phosphorylation; Glycolysis

	1 Introduction

Top Abstract 1 Introduction 2 Methods 3 Results 4 Discussion References

 
ATP-sensitive potassium (K⁺_ATP) channels, which were first discovered in cardiac myocytes, are activated by decreases in the intracellular concentration of ATP [1]. Results of several pharmacological studies with sulfonylurea K⁺_ATP channel blockers suggest that activation of K⁺_ATP channels is important in the shortening of the action potential duration and the increase in cellular K⁺ efflux during cardiac ischemia [2–4]. However, it has not been confirmed yet whether these channels actually open during the early phase of ischemia or not [5, 6]. While the direct measurement of current through K⁺_ATP channels during ischemia is not feasible in whole-heart or tissue preparations, there is experimental evidence that exposure to anoxia [7, 8]or simulated ischemia [9]activates K⁺_ATP channels in isolated ventricular myocytes. However, the precise mechanisms underlying the anoxia-induced activation of the K⁺_ATP channels are unclear.

The lack of sufficient oxygen suppresses oxidative phosphorylation, a major pathway of ATP synthesis. Although activation of K⁺_ATP channels has been reported to be regulated preferentially by ATP derived from glycolysis [10], the inhibition of oxidative phosphorylation by 2,4-dinitrophenol (DNP) [11–13]or cyanide [14, 15]has been shown to activate K⁺_ATP channels in isolated cardiac cells. Conversely, the DNP-induced activation of the K⁺_ATP channels has been partially reversed by enhanced glycolysis [12]. Therefore, it remains controversial whether the ATP derived from either glycolysis or oxidative phosphorylation alone is sufficient to regulate K⁺_ATP channels in myocytes.

The present study investigated the putative regulatory mechanism(s) of K⁺_ATP channel activation in single myocytes under anoxic conditions. Particular attention was paid to the role of the two major ATP synthetic pathways—oxidative phosphorylation and glycolysis.

	2 Methods

Top Abstract 1 Introduction 2 Methods 3 Results 4 Discussion References

 
2.1 Cell preparation and experimental set-up
Single ventricular myocytes from guinea pigs were prepared according to a technique described previously [16]. In brief, the guinea pig underwent cervical dislocation, and the heart was quickly excised and mounted on a Langendorff's apparatus. The heart was rinsed for 5 min with nominally calcium-free Tyrode solution, consisting of (in mM) NaCl 137, KCl 5.4, MgCl₂ 0.5, NaH₂PO₄ 0.16, NaHCO₃ 3.0, glucose 5.5, and HEPES 5 (pH 7.4, adjusted with NaOH). The heart was then enzymatically digested by perfusing it with nominally calcium-free Tyrode solution containing collagenase (0.05 mg/ml, type I, Yakult, Tokyo). After 5–6 min of digestion, the heart was perfused with a recovery solution of the following composition (in mM): KCl 5, glutamic acid 70, taurine 10, oxalic acid 10, KH₂PO₄ 5, HEPES 5, glucose 11, and O,O'-bis(2-aminoethyl)ethyleneglycol-N,N,N',N'-tetraacetic acid (EGTA) 0.5 (pH 7.4, adjusted with KOH) [17]. The temperature of all these perfusates was maintained at 36±0.5°C. The heart was then minced in 50 ml of the recovery solution to disperse the cells. A small amount of the precipitate was transferred to the recording bath (volume, 0.5 ml) located at the bottom of a semi-closed airtight chamber that was mounted on the stage of an inverted microscope (TMD; Nikon, Tokyo) (Fig. 1).

View larger version (23K): [in this window] [in a new window] [Download PowerPoint slide]   Fig. 1 Schematic illustration of a semi-closed airtight chamber designed to maintain an anoxic environment. Ultra-pure argon gas (99.9995%) was released from the bottom of the chamber. Air was expelled from the chamber through a small opening in the sliding top-plate. A patch electrode can be inserted through this opening in the top-plate for application to the cell placed in the recording bath. The anoxic perfusate was delivered through a stainless steel tube. Temperature of perfusates and argon gas was maintained at 36±0.5°C.

 
A semi-closed airtight chamber was specially designed to perfuse the cells under anoxic conditions while measuring ionic currents with a patch-clamp pipette. The top of the chamber was covered by a transparent acrylic plate. A patch electrode was inserted into the chamber through a small opening in the top-plate. Since the top-plate is free to move horizontally, a patch electrode can be applied easily to the cells. Ultra-pure argon gas (99.9995%) was released from the bottom of the chamber (2 l/min) and expelled through an opening in the top-plate. Since argon is heavier than air, the atmospheric air close to the surface of the recording bath was expelled and replaced by ultra-pure argon gas. Diffusion of atmospheric oxygen into the anoxic perfusate was completely blocked by the flow of argon.

Under control conditions, the cells were perfused with normoxic Tyrode solution of the following composition (in mM): NaCl 137, KCl 5.4, MgCl₂ 0.5, NaH₂PO₄ 0.16, NaHCO₃ 3.0, CaCl₂ 1.8, glucose 5.5, and HEPES 5 (pH 7.4, adjusted with NaOH). The partial pressure of oxygen (PO₂) in the normoxic Tyrode solution was determined using an O₂ monitor (POG-200BA, Unique Medical, Tokyo) and was approximately 150 mmHg. The anoxic solution was made by bubbling the glucose-free Tyrode solution with pure nitrogen gas (99.99%). The bubbling of nitrogen gas was performed under positive pressure (>30 cmH₂O) for more than 3 h. The PO₂ of the anoxic solution was less than 3.8x10^–8 mmHg, as determined by redox reaction with resazurine [18]. The anoxic solution was delivered to the bath in the chamber through stainless steel tubing. The flow rate of the perfusates was maintained at approximately 20 ml/min. The temperature of the perfusing solutions and argon gas was maintained at 36±0.5°C.

2.2 Recordings of action potentials and membrane currents
The action potentials and membrane currents were recorded using a patch-clamp amplifier (CEZ-2100; Nihon Kohden, Tokyo). Patch electrodes were fabricated from a capillary tube (Drummond Scientific Co., Broomall, PA) using a micropipette puller (P-97; Sutter instrument Co., USA) and a heat polisher (MF-83; Narishige, Tokyo) and filled with solution of the following composition (in mM); KCl 150, HEPES 10, and EGTA 20 (pH 7.2, adjusted with KOH). The resistance of the pipette ranged from 3 to 4 M. A ‘giga-ohm seal’ was established between the pipette and the cell surface by gentle suction. This was followed by application of greater suction to disrupt the membrane patch. Action potentials and transmembrane ionic currents were recorded in the whole-cell patch-clamp mode.

2.3 Experimental protocol
Action potentials were elicited at 1 Hz using a current clamp mode of patch clamp. After a stabilization period with constant action potential configuration in the normoxic Tyrode solution (usually >10 min), the cell perfusate was switched to the glucose-free anoxic solution that was delivered to the bath through a different stainless steel tubing. Reoxygenation was established by changing the anoxic perfusate to a normoxic one with or without 5.5 mM glucose. In some experiments, anoxic solution that contained glibenclamide (1 µM) or glucose (5.5 mM) was applied from another anoxic reservoir. Glibenclamide was a kind gift from Hoechst (Hoechst Japan Co., Tokyo) and stock solution of glibenclamide (1 mM) was made by dissolving the drug in 5% dimethyl sulphoxide (Sigma Chemical Co., St. Louis, MO). Details of the experimental protocol are shown at the top of each figure.

Membrane currents were recorded using a ramp voltage-clamp method. The ramp voltages (+60–140 mV, 10 mv/s) were applied from the holding potential of –40 mV at 30-s intervals. After the constant current–voltage relationship was obtained in the control normoxic tyrode solution, the cell was perfused with the glucose-free anoxic solution. Reoxygenation or application of glibenclamide was made by changing the anoxic perfusate to normoxic perfusate or anoxic solution that contained glibenclamide (1 µM). Details of the experimental protocol for current measurements are also shown in the top of each figure.

2.4 Data acquisition and statistical analysis
The action potential and ionic current signals were stored on magnetic tape using a PCM data recording system (PCM-501ES; SONY, Tokyo), replayed and processed on a personal computer (PC-9801; NEC, Tokyo) equipped with an analog-to-digital converter (ADX-98; Canopus, Kobe). The voltage signals were digitized at a sampling interval of 1 ms. The statistical data are expressed as the mean±SEM. The statistical significance of differences between data was determined using Student's unpaired t-test. A value of P<0.05 was regarded as statistically significant. The investigation conforms with the Guide for the Care and Use of Laboratory Animals (NIH Publication No. 85-23, revised 1985). All procedures met the guidelines stipulated by the Physiological Society of Japan and the Ethical Committee of Oita Medical University for Animal Experiments.

	3 Results

Top Abstract 1 Introduction 2 Methods 3 Results 4 Discussion References

 
3.1 Alterations of action potentials during anoxia and reoxygenation
The first series of experiments evaluated the effect of anoxia on action potentials stimulated at 1 Hz (Fig. 2). Changes in action potential duration (APD) (panel A) and action potential configuration (panel B) were recorded before, during, and after the cell was exposed to anoxic conditions. Following equilibration with normoxic solution, the cell was exposed to glucose-free anoxic solution. In a representative myocyte, the APD began to shorten after 6 min (lag period) of anoxia, resulting in a spike-like action potential in 10 min. Upon reoxygenation (in the continuing absence of glucose), the APD was rapidly restored to the pre-anoxic length. Similar changes were observed in 9 different cells during anoxia–reoxygenation. The APD decreased to <20% of the control value in 13±3 min (n = 10), but increased to pre-anoxic length within 20 s following reoxygenation.

View larger version (18K): [in this window] [in a new window] [Download PowerPoint slide]   Fig. 2 Typical effects of glucose-free anoxia on the action potential of a single ventricular myocyte. Action potentials were elicited at 1 Hz using a patch clamp in the current-clamp mode. (A) Serial changes in the action potential duration at the 90% repolarization level (APD90) before, during, and after anoxic intervention. In the control (normoxic) conditions, glucose (5.5 mM) was present. (B) Action potential configurations at each point indicated in panel A. The superimposed action potentials (right end) represent 18 consecutive action potentials recorded just after the start of reoxygenation.

 
3.2 Alteration of membrane currents during anoxia and reoxygenation
Since the induction of anoxia produced a marked shortening of action potential, we investigated the underlying changes in membrane currents. Fig. 3 illustrates recordings of membrane current elicited using ramp voltage-clamp, according to the protocol illustrated in the inset of panel A. The quasi-steady-state current–voltage relationships obtained from the same experiment are shown in panel B. Under normoxic conditions, the quasi-steady-state current (Fig. 3Ba) was thought to be a potassium current because the reversal potential of this current (–72 mV) was close to the theoretical equilibrium potential of K⁺ (–88.5 mV), and the inward-rectifying property of inward rectifier K⁺ channels was clearly seen. After 5 min of anoxia, the outward current increased at potentials positive to –60 mV (extra-outward current) and inward rectification disappeared (Fig. 3Bc). Following reoxygenation, the extra-outward current rapidly disappeared and the current–voltage relationship returned to that observed during pre-anoxic conditions (Fig. 3Bd,e).

View larger version (27K): [in this window] [in a new window] [Download PowerPoint slide]   Fig. 3 Effects of glucose-free anoxia on the quasi-steady-state membrane current recorded using a ramp voltage-clamp (cf. inset) at 30-s intervals. (A) A consecutive recording of the membrane current. (B) The current–voltage relationship obtained from each point indicated in panel A.

 
To identify the channel responsible for the anoxia-induced extra-outward current, selective blockade of K⁺_ATP channels with glibenclamide was examined (Fig. 4). As in Fig. 3, anoxia provoked an outward current at potentials positive to –60 mV. Glibenclamide (1 µM), applied during anoxia, abolished this extra-outward current within 1 min. The shortening of action potential produced by anoxia was also completely restored by the introduction of glibenclamide (data not shown).

View larger version (31K): [in this window] [in a new window] [Download PowerPoint slide]   Fig. 4 Effects of glibenclamide (1 µM) on the anoxia-induced outward current. (A) A typical recording of the quasi-steady-state membrane current elicited using a ramp voltage-clamp at 30-s intervals (same ramp voltage-clamp protocol as in Fig. 3). (B) The current–voltage relationship obtained from each point indicated in panel A.

 
3.3 Influence of glucose on the anoxia-induced shortening of the action potential duration
As shown above, the exposure of isolated myocytes to anoxic conditions caused activation of K⁺_ATP channels and shortened the APD. Since the anoxic solution did not contain glucose, it is possible that the activation of K⁺_ATP channels was simply provoked by the removal of extracellular glucose, an important substrate for glycolysis as well as for oxidative phosphorylation. To examine this possibility, the influence of extracellular glucose on the APD was investigated (Fig. 5). The removal of external glucose did not significantly shorten the APD, under normoxic conditions. A typical example is shown in Fig. 5A, in which the action potential did not shorten for 120 min. No significant change in the APD was recognized in the 5 cells tested, during a mean observation period of 67±25 min. Since there was no external supply of glucose, it seemed likely that glycogen and/or lipids stored in the cell might have been utilized as a substrate for glycolysis and oxidative phosphorylation during this period [10, 19]. However, anoxia in the presence of glucose (5.5 mM) produced a marked shortening of APD (Fig. 5B). Similar results were observed in 4 other cells tested.

View larger version (23K): [in this window] [in a new window] [Download PowerPoint slide]   Fig. 5 Typical changes in action potential duration at the 90% repolarization level (APD90) in myocytes subjected to various interventions as indicated at the top of each panel. Action potentials were stimulated at 1 Hz. (A) Removal of extracellular glucose alone did not shorten the APD90 under normoxic conditions during 120 min of observation. (B) Exposure to anoxia markedly shortened the APD90 even in the presence of extracellular glucose (5.5 mM) and reoxygenation rapidly restored the APD.

 
The activation of K⁺_ATP channels during anoxia seems to be preferentially mediated by the inhibition of oxidative phosphorylation due to the lack of oxygen. However, anaerobic glycolysis might have some modulatory effects on the activation of K⁺_ATP channels. To examine this possibility, the time course of action potential shortening during anoxia was compared in the presence and absence of extracellular glucose. For this analysis, the time course of action potential shortening was divided into two phases (Fig. 6A). Phase I refers to the period in which the action potential either did not change or changed slowly after the onset of anoxia. Following this phase, the action potential rapidly shortened until it became a spike-like contour. The latter phase was defined as phase II. Quantitatively, the duration of phase I was the time elapsed from the onset of anoxia to the time when the APD reached 80% of the control value. The duration of phase II was defined as the time from the end of phase I to the time when the APD shortened to 20% of the control value (Fig. 6A). The duration of phase II was significantly prolonged in the presence of 5.5 mM glucose (5.2±3.1 min in the absence of glucose, n = 7, vs. 15.5±3.2 min in the presence of glucose, n = 5; P<0.05), whereas the duration of phase I was not significantly different (8.4±4.6 min in the absence of glucose vs. 11.5±4.5 min in the presence of glucose) (Fig. 6B).

View larger version (21K): [in this window] [in a new window] [Download PowerPoint slide]   Fig. 6 Effects of extracellular glucose on the time course of action potential shortening during anoxia. (A) Definitions of phases I and II in action potential shortening. (B) Comparison of the effects of extracellular glucose on the duration of phases I and II in action potential shortening (cf. panel A). In the case of glucose-(+) (closed bars; n = 5), glucose (5.5 mM) was present before and during anoxia. In the case of glucose-(–) (open bars; n = 7), extracellular glucose was removed at the time when anoxia was started.

 
3.4 Effects of long-lasting anoxia or repeated exposures to anoxia
In preliminary experiments, we found that restoration of the APD by reoxygenation was successful only when the reoxygenation occurred soon after the action potential assumed a spike-like appearance. If the introduction of reoxygenation was delayed beyond 5 min after the action potential reached this stage, the APD could not be restored. Fig. 7 shows such an example. In the absence of glucose, the cell was exposed to long-lasting anoxic conditions that persisted for 5 min after the action potential shortened to a spike-like appearance. Subsequent reoxygenation did not restore the APD and application of glucose was unable to restore it. However, since the application of glibenclamide completely restored the APD (‘e–f’ in Fig. 7), it is evident that the sustained APD shortening seen after reoxygenation is due to the persistent activation of K⁺_ATP channels.

View larger version (18K): [in this window] [in a new window] [Download PowerPoint slide]   Fig. 7 Effects of long-lasting anoxia on action potentials. Top panel: Typical changes in the action potential duration at the 90% repolarization level (APD90) in a myocyte exposed to long-lasting anoxia, followed by reoxygenation. This was then followed by application of glucose (5.5 mM) and glibenclamide (1 µM) as indicated by bars on the top. Bottom panel: Action potentials recorded at each point indicated in the top panel.

 
Repeated exposure to anoxia progressively impaired the recovery of APD during reoxygenation. A typical example is shown in Fig. 8, in which the myocyte was exposed to anoxia–reoxygenation 4 times. The exposure to anoxia shortened the action potential to a spike-like appearance in each trial; however, the extent of recovery of the APD seen upon each successive reoxygenation progressively decreased as the number of anoxic exposures increased. In addition, the duration of anoxia required for the APD to assume the spike-like appearance was gradually reduced. After a fourth period of anoxia APD remained shortened during reoxygenation and exposure to extracellular glucose (5.5 mM) did not restore the APD. In contrast, application of glibenclamide (1 µM) led to a complete recovery of the APD (‘i–j’ in Fig. 8). Therefore, persistent K⁺_ATP channel activation is thought to underlie the particular type of APD shortening caused by repeated exposures to anoxia–reoxygenation, as was the case for long-lasting anoxia (Fig. 7).

View larger version (24K): [in this window] [in a new window] [Download PowerPoint slide]   Fig. 8 Effects of repetitive exposures to anoxia–reoxygenation on action potentials. Top panel: Typical changes in the action potential duration at the 90% repolarization level (APD90) in a myocyte exposed to 4 cycles of anoxia–reoxygenation. Extracellular glucose (5.5 mM) did not restore the APD. However, the application of glibenclamide (1 µM) led to complete restoration (‘j’). Bottom panel: Superimposed action potential configurations following reoxygenation (a–b, c–d, or e–f) or the application of glibenclamide (i–j). Each tracing represents a consecutive action potential (at 1 Hz), recorded between the points indicated by corresponding letters in the top panel.

 

	4 Discussion

Top Abstract 1 Introduction 2 Methods 3 Results 4 Discussion References

 
Anoxia-induced activation of K⁺_ATP channels in isolated ventricular myocytes has been reported by Benndorf and co-workers [7, 8, 20]. Since anoxia is a central feature of ischemia, the single-cell anoxia model may be a useful tool to investigate electrophysiologic changes in the heart subjected to ischemia, where the patch-clamp techniques are applicable. However, the oxygen demand of single myocytes is so small that the partial pressure of oxygen (PO₂) of the perfusate must be lowered below 0.15 mmHg to induce anoxia-related electrophysiologic changes [21]. The use of a closed (airtight) chamber to maintain the PO₂ at such a low level prevented the use of a patch-clamp electrode. A semi-closed air-excluding chamber was designed for this study by modifying an open-dish chamber that was introduced by Stern et al. [22]and later used with patch clamp by Benndorf et al. [7, 8, 20]. The ‘semi-closed’ chamber system developed for this study offered an advantage over the open-dish chamber methods used by previous investigators [7, 8, 20]because this semi-closed design did not require a delicate or fine control of the laminar flow of argon gas.

4.1 Anoxia activates K⁺_ATP channels and shortens action potential duration
In this study, the effects of anoxia on K⁺_ATP current and APD of single ventricular cells are similar to those reported by Benndorf et al. [7, 20]. Perfusion with glucose-free, anoxic solution shortened the APD of single ventricular myocytes and produced a large time-independent outward current. This current was completely suppressed by reoxygenation or by application of glibenclamide. Therefore, it is reasonable to assume that exposure to anoxia activates K⁺_ATP channels in single ventricular cells. Anoxia is assumed to suppress oxidative phosphorylation, an aerobic pathway for ATP synthesis, and activate K⁺_ATP channels. However, ATP derived from anaerobic glycolysis has been reported to preferentially regulate K⁺_ATP channels [10]. Agents that uncouple (2,4-dinitrophenol [11–13]) or inhibit (cyanide [14, 15]) oxidative phosphorylation also activate K⁺_ATP channels in isolated ventricular myocytes. Thus, it is unclear whether the activation of K⁺_ATP channels in cardiac cells is regulated primarily by ATP derived from glycolysis, oxidative phosphorylation, or from a combination thereof.

4.2 Source of ATP for control of K⁺_ATP channel activity
Action potential shortening subsequent to the activation of K⁺_ATP channels was not observed immediately after the cells were exposed to anoxic conditions. A lag period of several minutes was required for the observation of anoxia-induced changes. The lag period was likely due to the time required for the [ATP]_i to decline to the threshold level for activation of the K⁺_ATP channels. It is assumed that activation of the K⁺_ATP channels during anoxia is caused primarily by a decrease in [ATP]_i. However, other metabolites also contribute to the activation. Increases in the level of intracellular adenosine diphosphate (ADP) [23, 24], proton [25], and lactate [26, 27]have been shown to decrease the sensitivity of K⁺_ATP channels to [ATP]_i and to increase the open probability of the channels. In the present study, increases in intracellular ADP and/or lactate may have facilitated the activation of K⁺_ATP channels during anoxia. Involvement of intracellular acidification seems unlikely since the intracellular pH was buffered with 10 mM HEPES present in the pipette.

It has been reported that K⁺_ATP channels are regulated preferentially by the ATP generated via glycolysis in permeabilized cells [10]. However, the present results appear to be consistent with the hypothesis that the ATP produced by oxidative phosphorylation regulates the K⁺_ATP channels. The reasons are as follows: (1) Depletion of glucose from the perfusate did not affect the APD (Fig. 5A) and the presence of extracellular glucose exerted only a limited effect on the APD shortening induced by anoxia (Fig. 6). (2) The action potential shortening and activation of K⁺_ATP channels produced by anoxia were rapidly and completely restored by the replenishment of oxygen alone (Figs. 2 and 3). In the latter respect, we could not exclude the possibility that reoxygenation-induced restoration of APD and inhibition of K⁺_ATP channels might have been secondary to the enhanced synthesis of ATP via ‘aerobic’ glycolysis. The amount of ATP synthesized via oxidative phosphorylation by far exceeds ATP produced via aerobic glycolysis. Therefore it is difficult to differentiate precisely the fractional contribution of ATP produced by glycolysis from that of oxidative phosphorylation responsible for the closure of K⁺_ATP channels.

4.3 Role of ATP derived from anaerobic glycolysis
Although ATP produced by oxidative phosphorylation appeared to play a key role in the control of K⁺_ATP channel activity, ATP produced via anaerobic glycolysis also had a limited, but significant regulatory effect on K⁺_ATP channels during anoxia. The rate of APD shortening during anoxia was significantly slower in the presence of extracellular glucose. More precisely, the presence of glucose significantly lengthened the duration of phase II without affecting the length of phase I (Fig. 6B). Phase I seems to reflect the time needed for [ATP]_i to decrease to a threshold level for the activation of K⁺_ATP channels, after the aerobic ATP synthesis was inhibited. Phase I does not appear to reflect the dissipation of intracellular substrates for ATP production (i.e., glycogen or lipid store), because the APD was not substantially altered (for more than 60 min) even in the absence of extracellular glucose, under normoxic conditions (cf. Fig. 5A). In contrast, phase II may reflect the time course of a decline of [ATP]_i after it has reached the threshold concentration for K⁺_ATP channel activation. The presence of extracellular glucose retarded the rate of APD shortening at phase II, perhaps by supplying ATP produced via anaerobic glycolysis. Increased extracellular levels of glucose have been shown to enhance glycolysis [28]. Since the length of phase I was unaffected by the presence of glucose, the contribution of anaerobic glycolysis appeared to become manifest only after the [ATP]_i had been severely decreased after inhibition of oxidative phosphorylation.

Even during phase I, glucose-induced enhancement of anaerobic glycolysis might have increased [ATP]_i and tended to inhibit the APD from shortening. However, the concomitant increase in lactate might have offset such inhibitory effects of ATP on K⁺_ATP channels, because increases in lactate decreased the sensitivity of K⁺_ATP channels to [ATP]_i, and promoted the activation of K⁺_ATP channels [27]. Thus, the length of phase I might have been unaltered even in the presence of extracellular glucose. However, this principle could not be extended to the duration of phase II where severe reduction of [ATP]_i is anticipated to occur. In phase II, the presence of glucose decelerated the shortening of APD, perhaps because the inhibitory effect of ATP (produced by enhanced glycolysis) prevailed over the effect of lactate on K⁺_ATP channels.

Activation of K⁺_ATP channels in isolated myocytes occurs when oxidative phosphorylation is inhibited by metabolic inhibitors, including DNP [11–13]or cyanide [14, 15]. We found that the application of DNP activated K⁺_ATP channels even in the presence of extracellular glucose at a concentration of 5.5 or 11 mM [12]. However, this DNP-induced K⁺_ATP channel activation was reversed when the concentration of extracellular glucose was increased up to 22 mM [12]. These findings support the hypothesis that, under the conditions employed in our present study, the activation of K⁺_ATP channels in myocytes subjected to anoxia is preferentially controlled by ATP derived from oxidative phosphorylation. ATP produced via glycolysis also exerts a limited but significant inhibitory effect on the K⁺_ATP channels when oxidative phosphorylation is severely impaired.

Our present notion seems to be in contrast to the previous report using saponin-permeabilized ventricular cells of the same species and cell-attached patch-clamp techniques to measure the activity of single K⁺_ATP channels [10]. The application of substrates (fructose-1,6,-diphosphate, phosphoenol pyruvate, NAD, K₂HPO₄, and ADP in the presence of FCCP) for anaerobic glycolytic ATP production inhibited the K⁺_ATP channel activity more efficiently than did the application of substrates (pyruvate, glutamate, creatine, K₂HPO₄, and ADP) necessary for mitochondrial (oxidative phosphorylation-mediated) ATP production. To simulate in situ conditions of the heart, these experiments were conducted in the presence of 2-deoxyglucose (10 mM) and hexokinase (10 IU/ml) (i.e., an exogenous ATP-consuming system). In contrast, the effects of anoxia and removal of glucose, in our present report, were studied using quiescent myocytes. The contraction of myocytes was inhibited by the presence of intracellular EGTA. Thus, the ATP consumption in the present study may have been greatly reduced compared with that seen in the case of permeabilized cells. In intact cells, most ATP generated by mitochondria that are located close to myofilaments is thought to be used for the contractile machinery before it diffuses towards the K⁺_ATP channels in the cell membrane. In contrast, glycolytic ATP is generated by enzymes located at the site of K⁺_ATP channels. Therefore, an ATP concentration gradient exists between mitochondrial ATP production sites and K⁺_ATP channel sites in the cytoplasm. The magnitude of the gradient would be decreased after the rate of ATP utilization in the cytoplasm was increased, a condition similar to the presence of 2-deoxyglucose and hexokinase [10]. Therefore, in permeabilized cell experiments activation of K⁺_ATP channels could have been more sensitive to ATP produced via glycolysis than that produced in mitochondria. In cells in which contraction was inhibited by the intracellular introduction of EGTA (present study), an ATP concentration gradient in the cytoplasm may be altered because ATP consumption of the contractile machinery is minimal. Under these conditions, gating of K⁺_ATP channels would be primarily determined by ATP produced in mitochondria because the rate of production of ATP in mitochondria by far exceeds that via glycolysis. Indeed, in permeabilized myocytes, mitochondrial substrates were as effective as glycolytic substrates in blocking K⁺_ATP channels when the intrinsic level of ATP utilization was lowered [10].

Although extracellular glucose retarded the rate of APD shortening, perhaps due to the enhancement of ATP production via anaerobic glycolysis, the presence of glucose did not prevent the ultimate APD shortening caused by anoxia. We attribute the latter APD shortening to the inhibition of anaerobic glycolysis. Indeed, anaerobic glycolysis is reportedly inhibited in the late phase of ischemia, although it may be enhanced in earlier phases [29, 30]. In this context, it is noteworthy that the restoration of APD by reoxygenation was markedly inhibited after prolonged or repeated exposures to anoxia (Figs. 7 and 8). Such reoxygenation-resistant APD shortening could not be restored by the application of glucose but was completely restored by the application of glibenclamide. We therefore speculate that glycolysis itself may have been inhibited in these specific situations. Suppression of glycolysis eventually leads to the depletion of pyruvate, an essential substrate for oxidative phosphorylation, and results in no production of ATP via oxidative phosphorylation.

4.4 Limitations and implications of present study
K⁺_ATP channels are activated as a result of a decrease in [ATP]_i [1]. The activation of K⁺_ATP channels is known to be facilitated by increases of intracellular ADP, proton, and lactate. All of these changes in the intracellular milieu occur in the setting of ischemia; thus, cardiac K⁺_ATP channels might be activated in the early phase of ischemia, during which the average [ATP]_i still remains high. Several investigators have shown that the application of glibenclamide decreased the efflux of potassium in the initial fast phase but did not affect it in the secondary increase phase [6]. In isolated hearts, application of glibenclamide attenuated the action potential shortening even in the initial few minutes of no-flow ischemia [31, 32], thereby suggesting that K⁺_ATP channels are significantly activated at a very early phase of ischemia. However, glibenclamide used at relatively high concentration (>10 µM) cannot be considered a specific blocker for K⁺_ATP channels. Inhibition of catecholamine-induced chloride channels [33]and other potassium channels [34]have been described. Furthermore, glibenclamide is known to affect calcium release from the sarcoplasmic reticulum [35, 36]and to inhibit lactate production [4, 37]. Thus, it has not been defined yet whether cardiac K⁺_ATP channels actually open and contribute to the shortening of action potential duration and accumulation of extracellular potassium ions during the early phase of ischemia in vivo.

Our present findings are consistent with the hypothesis that the activation of K⁺_ATP channels is mediated primarily by the reduction of [ATP]_i derived from depressed oxidative phosphorylation, provided that the activation of K⁺_ATP channels has occurred within the first few minutes of ischemia. If we assume that the activation of K⁺_ATP channels is primarily regulated by ATP derived from anaerobic glycolysis (and not from oxidative phosphorylation), these channels are anticipated not to open during the early phase of ischemia, because it was reported that anaerobic glycolysis is markedly enhanced during this early phase of ischemia [29, 30].

Time for primary review 29 days.

	Acknowledgements

 
We thank K. Moriyama for secretarial assistance, and M. Ohara for comments. This study was supported by a Grant-in-aid for Scientific Research from the Ministry of Education, Science, Sports and Culture of Japan (#06670064) to M.A.

	References

Top Abstract 1 Introduction 2 Methods 3 Results 4 Discussion References

 

1. Noma A. ATP-regulated K⁺ channels in cardiac muscle. Nature (1983) 305:147–148.[CrossRef][Medline]
2. Gasser R.N.A., Vaughan-Jones R.D. Mechanism of potassium efflux and action potential shortening during ischaemia in isolated mammalian cardiac muscle. J Physiol (Lond) (1990) 431:713–741.[Abstract/Free Full Text]
3. Kantor P.E., Coetzee W.A., Carmeliet E.E., Dennis S.C., Opie L.H. Reduction of ischaemic K⁺ loss and arrhythmias in rat hearts. Effect of glibenclamide, a sulfonylurea. Circ Res (1990) 66:479–485.
4. Wilde A.A.M., Escande D., Schumacher C.A., et al. Potassium accumulation in the globally ischemic mammalian heart: a role for the ATP-sensitive K⁺ channel. Circ Res (1990) 67:835–843.[Abstract/Free Full Text]
5. Yan G.X., Yamada K.A., Kléber A.G., McHowat J., Corr P.B. Dissociation between cellular K⁺ loss, reduction in repolarization time, and tissue ATP levels during myocardial hypoxia and ischemia. Circ Res (1993) 72:560–570.[Abstract/Free Full Text]
6. Wilde A.A.M., Aksnes G. Myocardial potassium loss and cell depolarisation in ischaemia and hypoxia. Cardiovasc Res (1995) 29:1–15.[Free Full Text]
7. Friedrich M., Benndorf K., Schwalb M., Hirche H. Effects of anoxia on K and Ca currents in isolated guinea pig cardiocytes. Pflügers Arch (1990) 416:207–209.[CrossRef][Web of Science][Medline]
8. Benndorf K., Friedrich M., Hirche H. Anoxia opens ATP regulated K channels in isolated heart cells of the guinea pig. Pflügers Arch (1991) 419:108–110.[CrossRef][Web of Science][Medline]
9. Henry P., Popescu A., Pucéat M., Hinescu M.E., Escande D. Acute simulated ischaemia produces both inhibition and activation of K⁺ currents in isolated ventricular myocytes. Cardiovasc Res (1996) 32:930–939.[Abstract/Free Full Text]
10. Weiss J.N., Lamp S.T. Cardiac ATP-sensitive K⁺ channels. Evidence for preferential regulation by glycolysis. J Gen Physiol (1989) 94:911–935.[Abstract/Free Full Text]
11. Trube G., Hescheler J. Inward-rectifying channels in isolated patches of the heart cell membrane: ATP-dependence and comparison with cell-attached patches. Pflügers Arch (1984) 401:178–184.[CrossRef][Web of Science][Medline]
12. Nakamura S., Kiyosue T., Arita M. Glucose reverses 2,4-dinitrophenol induced changes in action potentials and membrane currents of guinea pig ventricular cells via enhanced glycolysis. Cardiovasc Res (1989) 23:286–294.[Web of Science][Medline]
13. Wu B., Sato T., Kiyosue T., Arita M. Blockade of 2,4-dinitrophenol induced ATP sensitive potassium current in guinea pig ventricular myocytes by class I antiarrhythmic drugs. Cardiovasc Res (1992) 26:1095–1101.[Abstract/Free Full Text]
14. Van der Heyden G., Vereecke J., Carmeliet E. The effect of cyanide on the K-current in guinea-pig ventricular myocytes. Basic Res Cardiol (1984) 80:93–96.
15. Noma A., Shibasaki T. Membrane current through adenosine-triphosphate-regulated potassium channels in guinea-pig ventricular cells. J Physiol (Lond) (1985) 363:463–480.[Abstract/Free Full Text]
16. Taniguchi J., Kokubun S., Noma A., Irisawa H. Spontaneously active cells isolated from the sino-atrial and atrio-ventricular nodes of the rabbit heart. Jpn J Physiol (1981) 31:547–558.[Web of Science][Medline]
17. Isenberg G., Klockner U. Calcium tolerant ventricular myocytes prepared by preincubation in a ‘KB medium’. Pflügers Arch (1982) 395:6–18.[CrossRef][Web of Science][Medline]
18. Allshire A., Piper H.M., Cuthbertson K.S.R., Cobbold P.H. Cytosolic free Ca²⁺ in single rat heart cells during anoxia and reoxygenation. Biochem J (1987) 244:381–385.[Web of Science][Medline]
19. Wisneski J.A., Gertz E.W., Neese R.A., Mayr M. Myocardial metabolism of free fatty acids. Studies with ¹⁴C-labeled substrates in human. J Clin Invest (1987) 79:359–366.[Web of Science][Medline]
20. Benndorf K., Bollmann G., Friedrich M., Hirche H. Anoxia induces time-independent K⁺ current through K_ATP channels in isolated heart cells of the guinea-pig. J Physiol (Lond) (1992) 454:339–357.[Abstract/Free Full Text]
21. Wittenberg B.A., Wittenberg J.B. Oxygen pressure gradients in isolated cardiac myocytes. J Biol Chem (1985) 260:6548–6554.[Abstract/Free Full Text]
22. Stern M.D., Silverman H.S., Houser S.R., et al. Anoxic contractile failure in rat heart myocytes is caused by failure of intracellular calcium release due to alteration of the action potential. Proc Natl Acad Sci USA (1988) 58:6954–6958.
23. Lederer W.J., Nichols C.G. Nucleotide modulation of the activity of rat heart ATP-sensitive K⁺ channels in isolated membrane patches. J Physiol (Lond) (1989) 419:193–211.[Abstract/Free Full Text]
24. Shen W.K., Tung R.T., Machulda M.M., Kurachi Y. Essential role of nucleotide diphosphates in nicorandil-mediated activation of cardiac ATP-sensitive K⁺ channel. A comparison with pinacidil and lemakalim. Circ Res (1991) 69:1152–1158.[Abstract/Free Full Text]
25. Koyano T., Kakei M., Nakashima H., Yoshinaga M., Matsuoka T., Tanaka H. ATP-regulated K⁺ channels are modulated by intracellular H⁺ in guinea-pig ventricular cells. J Physiol (Lond) (1993) 463:747–766.[Abstract/Free Full Text]
26. Keung E.C., Qian L. Lactate activates ATP-sensitive potassium channels in guinea pig ventricular myocytes. J Clin Invest (1991) 88:1772–1777.[Web of Science][Medline]
27. Han J., So I., Kim E.Y., Earm Y.E. ATP-sensitive potassium channels are modulated by intracellular lactate in rabbit ventricular myocytes. Pflügers Arch (1993) 425:546–548.[CrossRef][Web of Science][Medline]
28. MacLeod D.P., Prasad K. Influence of glucose on the transmembrane action potential of papillary muscle. Effects of concentration, phlorizin and insulin, non-metabolizable sugars, and stimulators of glycolysis. J Gen Physiol (1969) 53:792–815.[Abstract/Free Full Text]
29. Neely JR, Liedtke AJ, Whitmer JT, Rovetto MJ. Relationship between coronary flow and adenosine triphosphate production from glycolysis and oxidative metabolism. In: Roy PR, Harris P, editors. Recent advances in studies on cardiac structure and metabolism, vol 8. The sarcoplasm. Baltimore: University Park Press, 1975:301–321.
30. Rovetto M.J., Whitmer J.T., Neely J.R. Comparison of the effects of anoxia and whole heart ischemia on carbohydrate utilization in isolated working rat hearts. Circ Res (1973) 32:699–711.[Abstract/Free Full Text]
31. Cole W.C., McPherson C.D., Sontag D. ATP-regulated K⁺ channels protect the myocardium against ischemia/reperfusion damage. Circ Res (1991) 69:571–581.[Abstract/Free Full Text]
32. Shigematsu S., Sato T., Abe T., et al. Pharmacological evidence for the persistent activation of ATP-sensitive K⁺ channels in early phase of reperfusion and its protective role against myocardial stunning. Circulation (1995) 92:2266–2275.[Abstract/Free Full Text]
33. Tominaga M., Horie M., Sasayama S., Okada Y. Glibenclamide, an ATP-sensitive K channel blocker, inhibits cardiac cAMP-activated Cl conductance. Circ Res (1995) 77:417–423.[Abstract/Free Full Text]
34. Reeve H.L., Vaughan P.F.T., Peers C. Glibenclamide inhibits an voltage-gated K⁺ current in the human neuroblastoma cell line SH-SY5Y. Neurosci Lett (1992) 135:37–40.[CrossRef][Web of Science][Medline]
35. Chopra L.C., Twort C.H.C., Ward J.P.T. Direct action of BRL 38227 and glibenclamide on intracellular calcium stores in cultured airway smooth muscle of rabbit. Br J Pharmacol (1992) 105:259–260.[Web of Science][Medline]
36. Bian, K. Hermsmeyer, K. Enhancement of rat aorta Ca²⁺ channel current by glyburide (Abstract). FASEB J 1992;6:A1544.
37. Venkatesh N., Lamp S.T., Weiss J.N. Sulphonylureas, ATP-sensitive K⁺ channels, and cellular K⁺ loss during hypoxia, ischemia, and metabolic inhibition in mammalian ventricle. Circ Res (1991) 69:623–637.[Abstract/Free Full Text]

CiteULike    Connotea    Del.icio.us    What's this?

This article has been cited by other articles:

				J. Shimokawa, H. Yokoshiki, and H. Tsutsui Impaired activation of ATP-sensitive K+ channels in endocardial myocytes from left ventricular hypertrophy Am J Physiol Heart Circ Physiol, December 1, 2007; 293(6): H3643 - H3649. [Abstract] [Full Text] [PDF]

				P. Dhar-Chowdhury, M. D. Harrell, S. Y. Han, D. Jankowska, L. Parachuru, A. Morrissey, S. Srivastava, W. Liu, B. Malester, H. Yoshida, et al. The Glycolytic Enzymes, Glyceraldehyde-3-phosphate Dehydrogenase, Triose-phosphate Isomerase, and Pyruvate Kinase Are Components of the KATP Channel Macromolecular Complex and Regulate Its Function J. Biol. Chem., November 18, 2005; 280(46): 38464 - 38470. [Abstract] [Full Text] [PDF]

				J. Chen, J. X. Zhu, I. Wilson, and J. S. Cameron Cardioprotective effects of KATP channel activation during hypoxia in goldfish Carassius auratus J. Exp. Biol., July 15, 2005; 208(14): 2765 - 2772. [Abstract] [Full Text] [PDF]

				J. S. Cameron, K. E. Hoffmann, C. Zia, H. M. Hemmett, A. Kronsteiner, and C. M. Lee A role for nitric oxide in hypoxia-induced activation of cardiac KATP channels in goldfish (Carassius auratus) J. Exp. Biol., November 15, 2003; 206(22): 4057 - 4065. [Abstract] [Full Text] [PDF]

				T. Lu, T. Hoshi, N. L Weintraub, A. A Spector, and H.-C. Lee Activation of ATP-sensitive K+ channels by epoxyeicosatrienoic acids in rat cardiac ventricular myocytes J. Physiol., December 15, 2001; 537(3): 811 - 827. [Abstract] [Full Text] [PDF]

				A. Knopp, S. Thierfelder, B. Doepner, and K. Benndorf Mitochondria are the main ATP source for a cytosolic pool controlling the activity of ATP-sensitive K+ channels in mouse cardiac myocytes Cardiovasc Res, November 1, 2001; 52(2): 236 - 245. [Abstract] [Full Text] [PDF]

				K.-i. Ito, T. Sato, and M. Arita Protein kinase C isoform-dependent modulation of ATP-sensitive K+ channels during reoxygenation in guinea-pig ventricular myocytes J. Physiol., April 1, 2001; 532(1): 165 - 174. [Abstract] [Full Text] [PDF]

				Y. Shimoni, P. E. Light, and R. J. French Altered ATP sensitivity of ATP-dependent K+ channels in diabetic rat hearts Am J Physiol Endocrinol Metab, October 1, 1998; 275(4): E568 - E576. [Abstract] [Full Text] [PDF]

This Article Abstract FREE Full Text (PDF) E-letters: Submit a response Alert me when this article is cited Alert me when E-letters are posted Alert me if a correction is posted Services Email this article to a friend Similar articles in this journal Similar articles in PubMed Alert me to new issues of the journal Add to My Personal Archive Download to citation manager Request Permissions Disclaimer Google Scholar Articles by Shigematsu, S. Articles by Arita, M. Search for Related Content PubMed PubMed Citation Articles by Shigematsu, S. Articles by Arita, M. Social Bookmarking          What's this?

Online ISSN 1755-3245 - Print ISSN 0008-6363

Copyright © 2009 European Society of Cardiology

Oxford Journals Oxford University Press

• Site Map
• Privacy Policy
• Frequently Asked Questions

Other Oxford University Press sites: Oxford University Press Oxford Journals China Oxford Journals Japan American National Biography Booksellers' Information Service Children's Fiction and Poetry Children's Reference Corporate & Special Sales Dictionaries Dictionary of National Biography Digital Reference English Language Teaching Higher Education Textbooks Humanities International Education Unit Law Medicine Music Online Products Oxford English Dictionary Reference Rights and Permissions Science School Books Social Sciences Very Short Introductions World's Classics