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Cardiovascular Research 1999 41(3):629-640; doi:10.1016/S0008-6363(98)00238-7
© 1999 by European Society of Cardiology
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Copyright © 1999, European Society of Cardiology

Anoxia generates rapid and massive opening of KATP channels in ventricular cardiac myocytes

Andreas Knopp, Stephan Thierfelder, Rolf Koopmann, Christoph Biskup, Thomas Böhle and Klaus Benndorf*

Institut für Physiologie, Abt. Herz-Kreislauf-Physiologie, Friedrich-Schiller-Universität Jena, D-07740 Jena, Germany

* Corresponding author. Tel.: +49-3641-934351; Fax: +49-3641-933202.

Received 5 June 1998; accepted 10 June 1998


   Abstract
Top
 Abstract
1 Introduction
 2 Methods
 3 Results
 4 Discussion
 References
 
Objective: The aim was to improve the measurement of both the time course and amplitude of anoxia-induced KATP-channel current (IKATP) in isolated heart cells to specify the role of these channels in the time course of K+ accumulation in the ischemic myocardium. Methods: Ionic currents in isolated ventricular heart cells of the mouse were measured with a patch clamp technique under normoxic conditions (atmospheric pO2), during wash-out of oxygen, and under anoxic conditions (pO2<0.2 mmHg). During the measurement, the actual pO2 in the close proximity of the cell was determined with an optical technique by exciting Pd-meso-tetra(4-carboxyphenyl)porphin with light flashes of 508–570 nm and evaluating the quenching kinetics of the emitted phosphorescence signal at 630–700 nm. These quenching kinetics steeply depend on pO2 and can be evaluated best at pO2 values near 0 mmHg. Results: Out of 28 cells, 23 cells started to develop IKATP at pO2 values between 0 and 0.4 mmHg, i.e. in the range of the level of half maximum activity of the cytochrome oxidase. The remaining five cells developed IKATP between 0.4 and 1.8 mmHg. With respect to the time course, 18 out of 27 cells started to develop IKATP within the first minute after pO2 had decreased to values below 0.2 mmHg. The amplitude of IKATP induced by anoxia and various metabolic inhibitors was large, 29±12 and 48±21 nA (+40 mV), respectively. The anoxia-induced IKATP was significantly smaller than IKATP induced by metabolic inhibitors. During the pulses of 50 ms duration to +40 mV, the amplitude of IKATP decayed and, after clamping back to –80 mV, IKATP generated large tail currents. This suggests a notable change in the concentration gradient of K+ ions in the time range of tens of milliseconds. Conclusions: The results in isolated myocytes indicate that KATP channels open sufficiently rapidly after starting anoxia and generate sufficiently large conductance at maintained anoxia to explain both the time course and magnitude of the ischemic K+ accumulation if an appropriate counter-ion flux is available.

KEYWORDS ATP-sensitive K+ channels; Oxidative metabolism; Oxygen measurement; K+ efflux


   1 Introduction
Top
 Abstract
 1 Introduction
2 Methods
 3 Results
 4 Discussion
 References
 
Extracellular K+ accumulation during acute myocardial ischemia contributes to the generation of ventricular arrhythmias [1], leading in a considerable number of cases to sudden cardiac death. With respect to the time course of this K+ accumulation, it is well established that it starts during the first minute of ischemia and reaches maximum speed in the subsequent 5 min [2, 3]. The pathway along which K+ leaves the cells has been controversial for nearly two decades. At present, two major mechanisms have been put forward as contributing to the accumulation of K+: (1) As a consequence of the inhibition of oxidative metabolism, the cytosolic concentration of lactate and inorganic phosphate increases and these ions subsequently leave the cell. K+ is thought to be coupled to this efflux (via an unknown pathway) to balance the charge movement [4, 5]. (2) A special type of K+ channel opens, thereby generating a sufficiently high conductance. ATP-sensitive K+ channels (KATP channels) are the most likely among the K+ channels. Three experimental lines of evidence exist that KATP channels mediate the ischemic K+ efflux: (a) In inside-out patches, these channels open by a drop of the cytosolic [ATP] to levels below 20–100 µmol/l [6–8]. (b) In both cell-attached patches and whole-cell recordings of isolated cardiac myocytes, opening of KATP channels can be induced by either uncoupling oxidative phosphorylation [9], inhibition of the cytochrome oxidase [10]or removal of oxygen [11, 12]. Under all of these conditions, the current through KATP channels substantially exceeds that of any other K+ current. (c) Sulfonylureas that are known to be potent blockers of KATP channels [13, 14]preferentially reduce the first phase of K+ accumulation [15–17]. However, there are also experimental findings that seem to rule out KATP channels as mediators of K+ accumulation: (1) The [ATP] in the ischemic myocardium was found to remain in the range of several mmol/l [18], which is much too high to induce massive opening of KATP channels. (2) The whole cell current generated by KATP channels (recorded with the patch clamp technique) is too small to explain the K+ efflux. Much larger current, however, has been described by Isenberg et al. [19]using two-microelectrode voltage clamp before the discovery of KATP channels. (3) The time-course of opening of KATP channels is too slow to explain the start of K+ accumulation within the first minute of ischemia.

The aim of the present study was to further improve our cell-physiological approach with patch clamp measurements under hypoxic conditions [12]in order to uncover the role of KATP channels and to learn more about the cellular mechanisms underlying ischemic K+ accumulation. To this end, we present simultaneous measurement of ionic current with the patch clamp technique and oxygen tension (pO2) in the close proximity of the cell with an optical method. pO2 was obtained from evaluation of the phosphorescence quenching kinetics of Pd-meso-tetra(4-carboxyphenyl)porphin [20]. These kinetics depend steeply on pO2. Our results indicate that KATP channels open sufficiently rapid after starting anoxia and also generate a sufficiently large current to explain both the time-course and magnitude of ischemic K+ accumulation if a respective counter-ion flux is present.


   2 Methods
Top
 Abstract
 1 Introduction
 2 Methods
3 Results
 4 Discussion
 References
 
2.1 Chemicals
Collagenase was obtained from Sigma (type I) or Worthington Biochemical Corporation. Ascorbate oxidase was obtained from Sigma. Pd-meso-tetra(4-carboxyphenyl)porphin (PTP) was purchased from Medical Systems (Greenvale, NY, USA). The PTP powder consisted of 80% bovine serum albumin (BSA), 8% NaCl, 3% Trizma base and 9% Pd-meso-tetra(4-carboxyphenyl)porphin.

2.2 Isolation of single mouse myocytes
The method of isolating mouse ventricular myocytes has been described previously [21]. In brief, after mounting the heart on a Langendorff apparatus, the organ was perfused for 5 min with a nominally Ca2+-free Tyrode solution containing (in mmol/l) 140.0 NaCl, 5.8 KCl, 0.5 KH2PO4, 0.4 Na2HPO4, 0.9 MgSO4, 11.1 glucose and 10.0 HEPES (pH was adjusted to 7.1 with NaOH). Perfusion was continued for 30 min with enzyme solution, which was prepared from Ca2+-free Tyrode solution by adding 200 mg/l collagenase (Sigma Type I or Worthington) and adding defined amounts of Ca2+. The initial Ca2+ concentration was set to 10 µmol/l and increased subsequently in a step-like fashion to a final concentration of 100 µmol/l. Digestion was finished by perfusing the hearts for 3 min with recovery medium. This medium contained (in mmol/l): glutamic acid 50.0, HEPES 20.0, taurine 20.0, glucose 10.0, MgSO4 3.0, EGTA 0.5, KCl 30.0, KH2PO4 30.0, pH 7.3 (KOH). The cells were stored in this solution prior to experimental use. During all perfusion steps, the temperature was kept at 37°C.

2.3 Isolation of single guinea-pig myocytes
Ventricular heart cells of adult guinea pigs of either sex were isolated as described previously [11]. In brief, after extirpation, the heart was mounted on a Langendorff perfusion apparatus where it was first washed with 50 ml of a nominal Ca2+-free solution containing (in mmol/l) NaCl 140.0, KCl 10.0, MgCl2 1.0, glucose 10.0 and HEPES 5.0, pH 7.3 (NaOH). Perfusion was continued for 40 min with collagenase solution, which consisted of Ca2+-free solution to which 1000 mg/l fatty acid-free albumin (Sigma), 300 mg/l collagenase (Worthington) and 2 µmol/l CaCl2 were added. The Ca2+ concentration was raised in increments of 10 µmol/l at 10 min intervals. Finally, the heart was perfused with 50 ml of recovery solution. Before experimental use, the cells were incubated for 1 h in recovery solution. Perfusion was carried out at 37°C.

2.4 Electrophysiology
The patch clamp experiments were performed on the stage of an inverted microscope in an experimental chamber that was designed to insulate the bath against atmospheric oxygen by a layer of ultra pure argon while enabling free access for the patch pipette [22]. The volume of the experimental chamber was 0.1 ml and it was perfused at a rate of 0.3–0.4 ml/min. The control Tyrode solution used in the bath contained (in mmol/l) NaCl 150.0, KCl 5.4, CaCl2 2.5 or 3.6, MgCl2 0.5 and HEPES 10.0, pH 7.4 (NaOH). In part of the recordings, pO2 near the cell was simultaneously measured, as described in detail below. Inhibition of mitochondrial ATP synthesis was induced either by anoxia, by the uncouplers 2,4-dinitrophenol (DNP) or carbonyl cyanide(4-trifluoromethoxy)phenylhydrazone (FCCP), or by rotenone, which blocks the respiratory chain between flavin mononucleotide and ubiquinone. All experiments were carried out at 37°C.

Glass pipettes were pulled from borosilicate glass and their tips were heat polished. The final resistance was between 0.7 and 3 M{Omega} after filling with a solution containing (in mmol/l): 150.0 KCl, 5.0 HEPES and 10 mmol/l EGTA, pH 7.3 (adjusted using KOH).

Ionic current was recorded in the whole-cell configuration with a patch clamp technique using either an Axopatch 200 A or 200 B amplifier (Axon Instruments, Foster City, USA). In the experiments designed to determine the magnitude of IKATP, only pipettes with a resistance of 0.7 to 1.7 M{Omega} were used and the series resistance was compensated carefully, leaving about 20% uncompensated for at the beginning of the measurements. The holding potential was generally set to –80 mV. Voltage-dependent Na+ channels were not blocked.

Currents were filtered with a cut-off frequency of 10 kHz (4-pole Bessel). Recording and analysis of the data were performed on a PC-80486 or a Pentium PC with ISO2 software (MFK, Niedernhausen, Germany). All traces were recorded at a sampling rate of 10 kHz (12-bit resolution).

2.5 Deoxygenation of Tyrode solution
In the experiments for measuring the amplitude of IKATP with anoxic solution, oxygen was removed as described previously [12, 23].

In the experiments with concomitant measurement of pO2, anoxic Tyrode solution was prepared by removing the oxygen directly before reaching the experimental chamber using a special deoxygenizer. This deoxygenizer consisted of a gas-tight brass tube (internal diameter, 60 mm; external diameter, 65 mm; height, 170 mm). Within this tube, an oxygen-free gas atmosphere was created with the help of pure nitrogen, which was blown through the tube (flow rate, 200–250 ml/min). The bath solution was deoxygenized by slowly passing it through (flow rate, 0.4 ml/min) a Silastic silicon tube (length, 12 m; internal diameter, 1.47 mm; external diameter, 1.96 mm; Aromando, Düsseldorf, Germany) that was wound within the deoxygenizer. While passing through the deoxygenizer, the oxygen solved in the Tyrode diffused easily through the silicone wall of the tube into the nitrogen atmosphere, and was washed off. The solution leaving the deoxygenizer was completely free of oxygen (measured using the technique described below) and passed to the experimental chamber through high-performance liquid chromatography (HPLC) tubing. Before entering the experimental chamber, the solution was warmed to 37°C.

2.6 Oxygen measurement
The oxygen concentration was determined from the oxygen-dependent quenching kinetics of the phosphorescence signal of PTP. In Tyrode solution, oxygen is the only quencher and the process can be described by the Stern-Vollmer equation

Formula (1)
where {tau}0 is the lifetime in the absence of oxygen and qk is the quenching constant [20, 24]. The lifetime, {tau}, was obtained from the decay of the phosphorescence signal using the Oxyspot® system (GMS; Kiel, Germany). In detail (Fig. 1), the phosphorescence dye PTP was excited at 508–570 nm by short light flashes (decay time constant, 12 µs) that were generated by a xenon flash lamp inside the Oxyspot® system. After filtering, the flashes were directed via a light guide to the microscope where they reached the experimental chamber via a beam splitter and the objective (x40). The phosphorescence signal was collected by the objective and directed to the bottom of the microscope from where it was carried via a light guide to the photomultiplier of the Oxyspot® system. Data acquisition at a frequency of 0.5 MHz started 30 µs after the start of the flash in order to cut off the rising phase of the phosphorescence signal. To improve the signal-to-noise ratio, an average of 40 decay time courses were made, which were elicited in intervals of 100 ms. The averaged phosphorescence decay was fitted off-line with an own routine with a single exponential (Fig. 2A). At low oxygen concentrations, the decay was monoexponential. At higher concentrations of oxygen, the quality of the monoexponential fit became worse. The method is therefore restricted to oxygen concentrations below 70 mmHg (see also [24]).


Figure 1
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  		Fig. 1 Scheme of the experimental setup for the simultaneous measurement of pO2 and ionic currents. Patch clamp was performed in an experimental chamber that allowed one to isolate the atmospheric oxygen from the bath solution by an argon layer. The dye Pd-meso-tetra(4-carboxyphenyl)porphin (PTP), dissolved in the bath solution, was excited through the objective with light flashes of 508–570 nm. This excitation generated a phosphorescence signal (630–700 nm) that decayed with a rate depending on the pO2. The phosphorescence signal was directed to a photomultiplier, digitized and stored on a computer for evaluation. Light in the range of 527–617 nm was used in combination with a TV-camera to observe the cell under study. For further explanation, see text.

 

Figure 2
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  		Fig. 2 Quenching time constant, {tau}, of the phosphorescence signal of Pd-meso-tetra(4-carboxyphenyl)porphin. (A) Quenching time constant at three values of pO2. The decaying time courses could be reasonably fitted with single exponentials, yielding {tau} directly. The lowest pO2 generated the slowest decay, whereas the highest pO2 generated the fastest decay. (B) Determination of {tau}0 and qk according to the Stern-Volmer equation (Eq. 1). Temperature, 37°C. Tyrode solution, containing PTP and ascorbate oxidase (10 units), was filled in a gas-tight glass vessel. Equal amounts of ascorbate were added leading to a successive reduction of pO2 and to an increase in {tau} (see text). At the time when {tau} did not further increase upon addition of ascorbate, the solution was thought to be free of oxygen, yielding {tau}0, which was determined to be 560 µs. The diagram shows 1/{tau} plotted as a function of the amount of {Delta}ascorbate added. The three last additions of ascorbate did not alter 1/{tau} further, i.e. [O2] was zero. Transformation of {Delta}ascorbate into –{Delta}pO2 (upper axis) was done as described in the text. The titration curve was linear at 160 mmHg>–{Delta}pO2>70 mmHg and was therefore fitted using Eq. 1. The slope was used to determine the quenching constant, qk, which was found to be 380 (mmHg s)–1.

 
For visualization of the cell under study, the experimental chamber was also continuously illuminated by light of 572–617 nm, which was produced by a halide lamp plus filter combination that was above the microscope. The wavelengths of this light did not overlap with the wavelengths of the light employed by the oxygen measuring system.

The phosphorescence lifetime, {tau}0, and the quenching constant, qk, were determined using the following procedure: A glass vessel with a volume of 14.79 ml was completely filled with PTP containing Tyrode solution and 10 U of ascorbate oxidase. In order to prevent oxygen contamination from the atmosphere, the vessel was sealed using a butylrubber ring. Furthermore, the vessel was provided with an oxygen impermeable septum (Chrompack, Frankfurt, Germany), through which defined amounts of dissolved ascorbate could be injected without any notable oxygen entry. The test solution was warmed to 37°C in a water bath. For excitation of PTP and sampling of phosphorescence, a bifurcated, randomly mixed (salt and pepper) light guide was used whose single peripheral end was positioned at the glass wall of the vessel. For determination of {tau}0 and qk, an average of ten flashes was sufficient. Oxygen was removed in a stepwise fashion from the test solution by injection of 10 µl aliquots of ascorbate (13 mmol/l in Tyrode). The oxygen concentration in the vessel decreased correspondingly by equal amounts [24] according to the equation

Formula (2)
The equilibrium of this reaction is shifted far to the right, i.e. ascorbate is completely oxygenated at least down to pO2=5 mmHg [24]. After 4–5 min, the reaction was complete. Then, a phosphorescence signal was recorded and {tau} was determined. After oxygen was used up, further addition of ascorbate left {tau} unchanged, yielding {tau}0 directly. The quenching constant was obtained by plotting 1/{tau} as a function of the concentration of added ascorbate, {Delta}[ascorbate] (Fig. 2B). {Delta}[Ascorbate] was transformed into {Delta}[O2] according to Eq. 2. Using a solution coefficient of 3.94x10–5 (ml O2)/(ml solutionxmmHg), {Delta}[O2] was transformed into a decrease of oxygen tension (–{Delta}pO2), which was expressed in units of mmHg. At –{Delta}pO2>70 mmHg, the plot was approximately linear (Fig. 2B), with qk being the slope of the regression line. As a result, {tau}0 and qk were determined to be 550 µs and 370 mmHg–1 s–1, respectively (at 37°C). At –{Delta}pO2<70 mmHg the data deviate from linearity. This is most likely caused by the rapid decay time constant, which is in the range of the duration of the excitation flash. If not otherwise mentioned, a concentration of 0.4 g/l PTP powder (corresponding to 49.3 µmol/l PTP) in the extracellular Tyrode solution was used in all experiments.

For oxygen determination with metalloporphins, it is generally recommended to add more than 0.5% BSA to the Tyrode solution in order to prevent self-quenching [25]. We found that {tau}0 was independent of the amount of PTP powder (0.1 to 0.8 g/l) added to the Tyrode solution. Because of this result, experiments were performed without additional BSA so that the solution contained only 0.032% BSA when using 0.4 g PTP powder/l.

2.7 Statement
The investigation conforms with the Guide for the Care and Use of Laboratory animals published by the US National Institutes of Health (NIH Publication No. 85-23, revised 1996).

2.8 Statistics
Statistical data are given as mean±SD. Student’s t-test was used to test for statistical significance (P<0.05).


   3 Results
Top
 Abstract
 1 Introduction
 2 Methods
 3 Results
4 Discussion
 References
 
3.1 Time relationship between onset of anoxia and IKATP
Two experiments in which there was simultaneous measurement of whole cell current and pO2 are illustrated in Fig. 3. At t=0 s, the perfusion was switched from control Tyrode solution to anoxic Tyrode solution containing PTP. During the first tens of seconds, two processes overlap: pO2 decreases and the concentration of PTP increases. The latter means that measurement of pO2 is impossible before PTP increases to a sufficiently high level, which was reached after about 30 s in Fig. 3. This latency until pO2 could be recorded was not limiting because further time elapsed until IKATP appeared. In order to evaluate the latency between onset of anoxia and the appearance of IKATP, we defined the time when pO2 passed 0.2 mm Hg as the beginning of anoxia. At this pO2, the cytochrome oxidase has been reported to reach half maximum activity [26]. We considered the latency until the first indication of IKATP as well as its half maximum amplitude. In the experiment shown in Fig. 3A, the first indication of an extra outward current appeared 160 s after switching to the anoxic solution (arrow). This extra current has been demonstrated previously to be caused by the opening of KATP channels [12, 27]. The corresponding pO2 was determined to be about 0.2 mmHg. Half maximum IKATP was reached 225 s after switching to anoxic solution. Reoxygenation caused an immediate rise in pO2 levels (upper diagram) followed by a decrease in IKATP within a couple of seconds.


Figure 3
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  		Fig. 3 Time course of pO2 (squares) and ionic current (measured as late current at pulses of 50 ms duration to +40 mV; diamonds) when exposing a mouse myocyte to anoxic solution. (A) Cell with a rapid appearance of IKATP. Thirty to forty seconds after switching to anoxic solution, the concentration of PTP was high enough to measure pO2. The first increase in the net current, which may be attributed to IKATP (arrow), appeared at pO2=0.2 mmHg. (B) Cell with a delayed appearance of IKATP. In this cell, the extra current appeared 230 s after exceeding pO2=0.2 mmHg.

 
In the experiment illustrated in Fig. 3B, longer lasting anoxia was needed to induce IKATP. This finding fits with earlier observations where an approximately exponential distribution for the respective latencies was found [12, 27]. Panel B thus indicates that latency is a property of the individual isolated cell and that it cannot be explained by the time needed for the decay of pO2 in the experimental chamber.

A histogram of the latencies between the time when pO2 reached 0.2 mmHg and the onset of IKATP is plotted in Fig. 4A. As a result, in 18 out of 27 cells, IKATP appeared within the first minute of anoxia and only the remaining nine cells responded later. One cell did not respond within the recording time of 10 min. A plot of the latency between the time when pO2 reached 0.2 mmHg and half maximum IKATP (Fig. 4B) shows that the majority of cells generated substantial IKATP within the first 3 min, and, apart from one cell, all cells had responded after 7 min.


Figure 4
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  		Fig. 4 Statistics of the latency of IKATP. (A) Latency between anoxia and the first indication of IKATP in 27 mouse myocytes. The beginning of anoxia was set to the time when pO2 had fallen to levels below 0.2 mmHg. Binwidth, 40 s. In the majority of cells, IKATP appeared within the first minute of anoxia or even before reaching pO2=0.2 mmHg. Some of the cells, however, responded significantly later. (B) Plot of the latency between the beginning of anoxia and the half maximum IKATP for the same experiments as shown in (A). (C) Histogram of the first indication of IKATP as a function of the actual pO2. Binwidth, 0.2 mmHg; data were from 28 mouse myocytes. The leftmost bin corresponds to those cells in which pO2≤0.2 mmHg had to be reached to induce IKATP. The other bins indicate that, in some of the cells, oxidative phosphorylation produces an insufficient amount of ATP at pO2 levels above that of half maximum activity of the cytochrome oxidase.

 
A different type of histogram is illustrated in Fig. 4C. Herein, the first indication of IKATP is related to the pO2 measured. The largest, most left bin in the histogram corresponds to cells in which IKATP appeared after pO2 had fallen below 0.2 mmHg. The bins on the right side indicate that some cells already developed IKATP at pO2 that clearly exceeded 0.2 mm Hg. Assuming that all of the cytochrome oxidase molecules have the same pO2 value for half maximum saturation, this finding suggests the existence of oxygen gradients within the cytosol.

Conclusively, simultaneous measurement of whole cell IKATP and pO2 around the cell shows that KATP channels open with kinetics that are sufficiently rapid to explain the time course of the early extracellular K+ accumulation in the ischemic myocardium [2–4].

3.2 Magnitude of IKATP
In an attempt to improve quantification of the large IKATP in whole cells, we optimized the clamp conditions by series resistance compensation, as described in Section 2. To alleviate the experiments, most of them were carried out with metabolic inhibitors (DNP, 1–10 µmol/l; FCCP, 10–100 nmol/l; rotenone, 10 µmol/l) instead of anoxia. Fig. 5 A illustrates two superimposed current traces of a mouse myocyte. Under control conditions, the current at +40 mV is only Ito that in most cells slowly decreases during anoxia [28]. After maximal development of IKATP, a current of 84 nA at the end of a 50-ms pulse (late current) was generated by DNP in this cell. Guinea-pig cells developed IKATP of a similarly large amplitude. Fig. 5B illustrates respective traces of a guinea-pig myocyte in which the total current during anoxia reached 88 nA. The largest peak current observed reached 110 nA. In order to quantify IKATP roughly, the late current of the respective control trace was subtracted. In mouse myocytes we obtained an IKATP of 29±12 nA with anoxia (n=16), 48±22 nA with DNP (n=15), 47±20 nA with FCCP (n=21), and 48±20 nA with rotenone (n=6). Since the values with the metabolic inhibitors were not significantly different, they could be lumped together yielding 48±21 nA. The anoxia-induced IKATP was significantly smaller than IKATP induced by the metabolic inhibitors. A histogram of the maximum amplitudes of IKATP is shown in Fig. 6a. The wide distribution illustrates the great variability of the IKATP amplitude among the cells. In anoxic guinea-pig myocytes an IKATP of 57±23 nA has been obtained (n=14) which was significantly larger than the respective value in mouse cells.


Figure 5
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  		Fig. 5 Magnitude of IKATP. The cells were clamped from the holding potential of –80 mV to the test potential of +40 mV at a pulsing rate of 1 Hz. When using optimized clamp conditions (see Section 2), currents as large as 100 nA could be recorded. (A) Current in a mouse myocyte before the appearance of IKATP with a small transient component (only Ito) and after the development of IKATP. IKATP was induced by 5 µmol/l DNP. (B) Current in a guinea-pig myocyte before and after the appearance of IKATP induced by anoxia.

 

Figure 6
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  		Fig. 6 Magnitude of IKATP (a) Histogram of IKATP magnitude in 58 mouse cells. The late current during a pulse of +40 mV minus the late current amplitude of the control trace is plotted. The holding potential was–80 mV. IKATP was induced either by DNP, FCCP, rotenone (metabolic inhibitors) or anoxia. (b) Plot of the pipette’s resistance versus IKATP. Pipettes with a lower resistance allow one to record larger IKATP. The datapoints were fitted to the equation Rpip=axIKATP+b, yielding a=–0.0069 M{Omega}/nA and b=1.487 M{Omega} (r2=0.2951) as parameters.

 
One has to be aware that such large currents cannot be accurately clamped, even when compensating for most of the series’ resistance. In the case of perfect voltage clamp, one would expect independence between IKATP and the pipette resistance. However, the plot of the resistance of the used pipettes versus IKATP (Fig. 6b) shows that dependence does exist. Linear fit of the scattering data points emphasizes this dependence; the wide pipettes with the low resistance were superior for recording large currents. Conclusively, despite the large amplitude of IKATP described herein, our measurements underestimate the true IKATP substantially.

3.3 Large IKATP alters the driving force for K+
At large IKATP, pulses of 50 ms duration (+40 mV) caused time-dependent amplitude of the total current. During the pulse, the outward current decayed (time constant, 18.7±4.8 ms; n=29). Interpretation of this decay is complicated because it is a priori not clear to what extent inactivation of Ito contributes. In order to understand the nature of the decay, we looked through the data for experiments in which Ito was inactivated only to a minor extent before IKATP developed. Fig. 5A shows such an experiment in a mouse cell. At the time of development of substantial IKATP, decay of the total current was much larger than that of Ito in the control trace. Fig. 5B shows a typical trace obtained from a ventricular myocyte of the guinea-pig before and during anoxia. Guinea-pig cells do not generate Ito. Nevertheless, after the development of IKATP, a distinct decay is present. It is therefore concluded that, in mouse myocytes as well, the decay is not dominated by inactivation of Ito. The simplest explanation of the decay is that K+ is depleted inside the cell because of the large outwardly directed IKATP.

The large outward current during the pulse at +40 mV should also generate extracellular K+ accumulation in narrow extracellular spaces if the flown out K+ ions do not equilibrate rapidly with the K+ ions of the bulk solution. As a consequence of notable accumulation of K+ ions, clamping back to the holding potential should generate a large inwardly directed tail current [29]. Such tail currents were observed regularly. Furthermore, the amplitude of the tail current should increase proportionally with IKATP at +40 mV. This prediction was tested by considering tail currents during the development of IKATP. Fig. 7 A illustrates that the amplitude of the tail currents in fact corresponds to the amplitude of IKATP at +40 mV. The time course of four characteristic current amplitudes during the whole experiment is shown in Fig. 7B. During both the phase of appearance and disappearance of IKATP, the amplitude of decay (difference between open diamonds and filled squares) and tail current (difference between filled diamonds and open squares) increased concomitantly with the amplitude of the total current at +40 mV.


Figure 7
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  		Fig. 7 Large IKATP alters the driving force for K+ rapidly. (a) Nine consecutive current pulses were recorded in intervals of 5 s during the rising phase of IKATP that was induced by 5 µmol/l DNP. The holding potential was –80 mV and the pulse potential was +40 mV. The amplitude of both the decay during the pulse and the tail current after repolarization increased in proportion to the amplitude of IKATP. The symbols correspond to the times at which current amplitudes for the plot in (B) were measured. (B) Long time course of the same experiment as shown in (A). The steady state current at –80 mV (open squares), the peak current at +40 mV (open diamonds), the late current at +40 mV (filled squares), and the current amplitude after clamping back to –80 mV (measured 1.0 ms after the clamp step) were plotted. The time courses show the concomitant changes in the amplitude of decay and tail current with the amplitude of IKATP.

 
Conclusively, when depolarizing to +40 mV, opening of KATP channels may generate significant changes in the intra- and extracellular K+ concentration in the time range of several tens of milliseconds.


   4 Discussion
Top
 Abstract
 1 Introduction
 2 Methods
 3 Results
 4 Discussion
References
 
4.1 Oxygen measurement
We combined the patch clamp technique with an optical method for the determination of pO2 [20, 25]. This technique provides the following unique advantages: (1) pO2 is measured in the close proximity of the cell, i.e. the measured pO2 most likely represents the actual pO2 that the cell is exposed to. (2) At variance to the alternative technique of oxygen electrodes, the quality of the measurement is best at the lowest levels of pO2 (resolution ±0.1 mmHg) because the quenching kinetics are slowest. This allows one to measure pO2 in the range of interest when studying hypoxic inhibition of metabolism and related processes. (3) Measurements are free of drifts and are independent of the dye concentration, provided that it is high enough to obtain a reasonable signal-to-noise ratio. In summary, the described combination of patch clamp and oxygen measurement might become a powerful technique for studying ionic currents under conditions of very low oxygen tension.

4.2 Oxygen tension and KATP channel opening
In previous reports, we related the appearance of IKATP to the time of switching to anoxic solution [12, 27, 30], because we could not measure pO2. The present results show that this latency was partly caused by the time needed for pO2 reduction in the measuring chamber. Hence, one of the key findings of the present report is that, in the majority of our isolated cells, anoxia (pO2<0.2 mm Hg) opens KATP channels rapidly, namely within the first 2 min.

However, the term ‘anoxia’ should be discussed in more detail. We arbitrarily defined anoxia as a pO2≤0.2 mmHg (half maximum activity of the cytochrome oxidase [26]) and determined the latency as the time period between reaching pO2=0.2 mmHg in the measuring chamber and the appearance of IKATP (Fig. 4C). It would be optimal if the time period between anoxic block of oxidative metabolism and the opening of KATP channels is measured after instantaneously reducing pO2 to zero or if pO2 reduction is at least much faster than the response of the channels. This was not the case in our experiments. It is notable that despite the slow decrease of pO2 in our experimental chamber, the majority of cells developed IKATP at pO2 levels in the range of half maximum activity of the cytochrome oxidase. It is also interesting that five out of 28 cells had already developed IKATP at a pO2 of 0.4–1.8 mm Hg, because, at this pO2, cytochrome oxidase reaches half maximum or even higher activity and, thus, millimolar concentrations of ATP should be maintained. As a consequence, in these cells either the activity of the cytochrome oxidase is down-regulated or the actual pO2 around the mitochondria is reduced with respect to the extracellular pO2. While the first speculation is less likely, experimental and theoretical evidence for substantial oxygen gradients within myocardial cells has been described [31]. At pO2=1 mmHg, these authors reported that the percentage of oxidized cytochrome a+a3 was only 20% (estimated from Fig. 5 in [32]). Further heterogeneities of pO2 might result from the non-uniform distribution of myoglobin, which binds O2 within the cells and contributes to intracellular O2 diffusion [33].

At present, it is not clear why some cells have substantially longer latencies before the appearance of IKATP than others. Since cytosolic ATP predominantly controls the activity of the KATP channels [34], ATP is presumably available for longer in these cells. The reason may be lower ATP consumption and a better ATP buffering capacity by creatine phosphate and other substrates available for ATP production. A depletion of substrate as only reason for the drop in cytosolic ATP is, however, unlikely, because reoxygenation immediately removes IKATP for tens of minutes. This shows that mitochondrial substrate is still available. We therefore favor the idea of a block of glycolysis by an anoxia-induced increase of NADH and that the latency of the onset of this block varies.

4.3 How large is IKATP?
Ionic currents measured with the patch clamp technique with wide pipettes and the facilities for series resistance compensation provided by the amplifier, yielded larger whole cell IKATP than reported previously for both mouse and guinea-pig myocytes [10, 12, 13, 17, 27, 35]. However, at least two arguments may be derived from our experiments that indicate that IKATP is significantly larger than that measured even under our improved conditions. First, we still found a dependence of IKATP amplitude on the pipette’s resistance (Fig. 6b). Second, when clamping repeatedly to the nominal voltage of +40 mV, escape of voltage in the cells with the largest IKATP reached such a high degree that even the Na+ current equivalent disappeared. This suggests that the actual membrane voltage was probably much less depolarized, e.g. –40 to –50 mV, which, in turn, indicates that the true K+ conductance exceeded that derived from the current measurement by several fold. Therefore, alternative voltage clamp techniques are required to accurately measure the full amplitude of IKATP. Among the presently available techniques for voltage clamping isolated heart cells, two techniques are promising: the oil-gap technique introduced by Mitsuiye and Noma [36]and the two-electrode voltage clamp. With respect to the latter technique, it should be emphasized that DNP-induced K+ current of very large amplitude has already been described in guinea-pig cells with two microelectrodes [19]. These authors found a K+ conductance near the reversal potential of about 1.2 µS (estimated from Fig. 6 of this report), which is roughly double that in the present study (400–900 nS; data not shown). Despite the use of two electrodes, Isenberg et al. [19]were also aware that they could not reasonably clamp these large currents. In conclusion, the real amplitude of IKATP is at least twice as large as that measured in the present study and is most likely even larger.

Measurement of larger IKATP in whole cells renews the question of the number of active channels in a heart cell. Taking the maximum measured IKATP at +40 mV to be 50 nA, a single channel current of 2 pA [37–39], and a maximum open probability of 0.7 [12], one calculates, as the estimated lower limit, more than 35,000 channels per cell. Including the enormous voltage escape (see above) and the large (and also not reasonably clamped) current determined by Isenberg et al. [19], an estimate of the lower limit significantly exceeding 100,000 channels per cell seems to be more realistic. This value exceeds even the largest value in the literature, which has been estimated to be 50,000 channels per cell [39]. These authors counted the channels in cell-attached patches of rat cardiac myocytes and extrapolated this number to the total membrane area by assuming that there were ten channels per µm2 of membrane and a total membrane area of 5000 µm2. However, this estimate is certainly too small because neither the exact area of the patch nor the percentage of functioning channels in the patch are known. The latter may be of importance because formation of the patch may significantly reduce the percentage of active channels, which has been shown for voltage-dependent Na+ channels [40].

It is also notable that in guinea-pig myocytes, which produced large IKATP in our studies and also in the work of Isenberg et al. [19], a much lower number (2000–3000) of channels per cell has been estimated from the analysis of both single channel currents [41]and whole-cell currents [10]. While in the small whole-cell currents limitations of the voltage clamp could have introduced significant error, the reason for this difference in the single channel data is not clear.

4.4 Role of KATP channels during ischemia
Both the large amplitude and the short latency of IKATP induced by either anoxia or chemical inhibition of the respiratory chain strongly support the idea that KATP channels are the predominant pathway for K+ efflux in the ischemic myocardium [2, 3]. However, K+ conductance alone cannot explain the K+ efflux during ischemia because, even in cases where it substantially exceeds all other conductances, it would always fix the membrane potential such that the influx of K+ ions balances the efflux. The amount of K+ efflux in the ischemic myocardium therefore requires an equal flux of a counter-ion, either an efflux of anions or an influx of cations, as the generator of the driving force. At present, neither the nature of this current in the ischemic myocardium has been elucidated nor has evidence been presented that the ischemic cell does not develop currents in addition to those observed in single myocytes during anoxia. Assuming that the currents in the anoxic cell are representative for those in the ischemic cell, it may be concluded that any conductance available for the counter-ion is small compared to that provided by KATP channels because the reversal potential in the presence of large IKATP remained in the range of the resting potential (Fig. 7B; cf. [12]).

To determine the extent of early K+ accumulation in the ischemic myocardium, the actual driving force of the K+ ions may be estimated as follows: Assuming a K+ accumulation rate of 1 mmol/(l min) [2, 3], an average cellular volume of 67 pl (determined from guinea-pig cells; unpublished) and an extracellular space that was one third the size of the intracellular space [42], one calculates a mean K+ current per cell of 35 pA. Taking into account a conductance of 500 nS per cell, a deviation of only 0.07 mV from the actual resting potential can be calculated. Furthermore, assuming a mean duration of KATP channel activity of 1 min (Fig. 4, 7B) and that the variable latency observed during anoxia (Fig. 4A) is also present in ischemic tissue, the IKATP of an individual cell is most likely manifold larger than 35 pA.

The K+ influx generated by the Na+–K+ pump should be included in this consideration. There is experimental evidence that the pump’s activity is still maintained at the time when KATP channels open [34]. Assuming a pump current of 30 pA at –80 mV at the physiological [Na+]i of 10 mmol/l [43], a rate of decrease of [K+]e of less than 2 mmol/(l min) is calculated. This pump rate is necessary to balance any K+ efflux in order to maintain the Na+ and K+ gradient under physiological conditions. The above estimations of K+ efflux may therefore be considered as efflux in addition to that already compensated for by the pump.

Furthermore, the time course of the counter-ion flux should be considered. In the case where the counter-ions start to flow before the development of IKATP, one would have expected (under our experimental conditions) that notable extra inward current would have appeared at –80 mV because the equilibrium potential for the counter-ion flux is most likely positive at this voltage. Since such a current component did not appear (Fig. 7B), it is speculated that, during ischemia, conductance for the counter-ions develops together with IKATP. Taken together with the finding that the time course of IKATP fits well with the ischemic accumulation of K+ and including the fact that the ischemic K+ efflux can only be as large as the flux of the counter-ion, it seems to be an attractive idea to assume that the flux of counter-ions is coupled to oxidative metabolism, in a similar way to the opening of KATP channels.

Possible candidates for a counter-ion flux are (1) an efflux of phosphate, (2) an efflux of chloride, (3) an efflux of lactate, an influx of Na+ ions through either non-specific channels (4) or voltage-dependent Na+ channels (5). Whereas no convincing experimental evidence exists at present for an efflux of chloride (1) and phosphate (2), lactate, which is produced by anaerobic glycolysis, has been reported to be liberated from the ischemic myocardium to a great extent [44]and it is likely that the major part of the lactate efflux is mediated by a monocarboxylate transporter [45, 46]. Recently, this idea has been confirmed by Northern blot analysis, which showed that one of these transporters (MCT1) is also expressed in the heart [47]. In order to balance the K+ efflux through KATP channels, lactate would have to permeate the membrane as an anion, i.e. the action of the transporter would have to be electrogenic. Support for this hypothesis comes from the results of Gwilt et al. [44]who reported functional interdependence of the efflux of K+ through KATP channels and of lactate by the monocarboxylate transporter inasmuch as they showed that the efflux of both K+ and lactate is inhibited to a similar extent by either glibenclamide, a blocker of KATP channels, or {alpha}-cyano-4-hydroxycinnamic acid, a blocker of the monocarboxylate transporter. Coupling of lactate efflux to the transmembrane K+ gradient has also been described by Trosper and Philipson [48]. In the case of such a functional coupling between the efflux of K+ and lactate, the protons left within the cell should also be considered. Protons are certainly buffered within the cell or may be exchanged for Na+ ions by the Na+–H+ exchanger with the consequence of an intracellular Na+ load. In the ischemic myocardium, the latter is well established [49, 50]. Another experimental result, however, conflicts with the hypothesis of a functional coupling between K+ and lactate efflux: Shieh et al. [51]reported that the monocarboxylate transporter is electroneutral, which means that protons are cotransported with the charged lactate, which indicates that lactate efflux via the monocarboxylate transporter cannot balance K+ efflux though KATP channels. Further experiments will be necessary to solve this puzzle.

The fourth candidate for a counter-ion flux, an influx of cations (4), would require that Ca2+ ions are excluded because, at the required flux rates, the cells would immediately die from Ca2+ overload, which is not the case. Therefore, nonspecific cation channels carrying monovalents might be involved. It has been shown that such channels in the myocardium open at an elevated intracellular [Ca2+] [52]. However, at present, there is no experimental evidence that these channels open to a sufficient extent during ischemia. At least under conditions of anoxia and chemical inhibition of the respiratory chain, opening of these channels was not observed [12].

Na+ influx through voltage-dependent Na+ channels (5) usually lasts for only several tens of microseconds at physiological temperature [53]. From time to time, however, Na+ channels may switch to a non-inactivating gating mode [54]and it has been shown that long openings are favored at increased levels of lysophosphatidylcholine [55], which is considered to be one of the mediators of ischemic injury of the heart [56]. Since the number of Na+ channels available in the membrane is large, switching of only a very small fraction of these channels to the non-inactivating mode would be sufficient to mediate sufficient Na+ influx to balance the measured K+ efflux during ischemia.

In conclusion, the results of this report further strengthen the idea that KATP channels form a major pathway for K+ efflux in the ischemic myocardium by ruling out two important objections against this hypothesis: The conductance generated by KATP channels would be too small to explain the degree of K+ accumulation and too late to explain its initiation during the first minute after starting ischemia.

Time for primary review 18 days.


   Acknowledgements
 
This work was supported by the grant Be1250/10-1 of the Deutsche Forschungsgemeinschaft.


   References
Top
 Abstract
 1 Introduction
 2 Methods
 3 Results
 4 Discussion
 References
 

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