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Am J Physiol Heart Circ Physiol 283: H715-H724, 2002. First published April 11, 2002; doi:10.1152/ajpheart.01000.2001
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Vol. 283, Issue 2, H715-H724, August 2002

Electrophysiological response of rat atrial myocytes to acidosis

Kimiaki Komukai, Fabien Brette, and Clive H. Orchard

School of Biomedical Sciences, University of Leeds, Leeds LS2 9NQ, United Kingdom


    ABSTRACT
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

The effect of acidosis on the electrical activity of isolated rat atrial myocytes was investigated using the patch-clamp technique. Reducing the pH of the bathing solution from 7.4 to 6.5 shortened the action potential. Acidosis had no significant effect on transient outward or inward rectifier currents but increased steady-state outward current. This increase was still present, although reduced, when intracellular Ca2+ was buffered by 1,2-bis(2-aminophenoxy)ethane-N,N,N',N'-tetraacetic acid (BAPTA); BAPTA also inhibited acidosis-induced shortening of the action potential. Ni2+ (5 mM) had no significant effect on the acidosis-induced shortening of the action potential. Acidosis also increased inward current at -80 mV and depolarized the resting membrane potential. Acidosis activated an inwardly rectifying Cl- current that was blocked by 4,4'-diisothiocyanostilbene-2,2'-disulfonic acid (DIDS), which also inhibited the acidosis-induced depolarization of the resting membrane potential. It is concluded that an acidosis-induced increase in steady-state outward K+ current underlies the shortening of the action potential and that an acidosis-induced increase in inwardly rectifying Cl- current underlies the depolarization of the resting membrane potential during acidosis.

action potential; potassium current; chloride current; perforated patch


    INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

CARDIAC MUSCLE becomes acidic in a number of pathological conditions (27). Acidosis decreases the strength of contraction of the heart [for review see Orchard and Kentish (27)] and can induce arrhythmias [for review see Orchard and Cingolani (26)], both of which impair cardiac function. There are many mechanisms whereby acidosis can induce arrhythmias, among which is changing action potential duration: longer action potentials can produce triggered activity, shorter action potentials decrease the refractory period and can lead to premature contraction, and regional effects on the action potential can produce QT dispersion and reentry tachyarrhythmia. Previous studies have shown marked effects of acidosis on the action potential in ventricular cells (3, 19, 21) that could produce arrhythmias by these mechanisms. However, despite the prevalence of atrial arrhythmias, the effect of acidosis on the atrial action potential is unknown.

We have recently shown that acidosis prolongs the action potential in rat ventricular cells by inhibition of the steady-state K+ current (ISS). However, the response in atrial cells may be different. These cells have three distinct Ca2+-independent, depolarization-induced outward K+ currents: a rapidly activating, rapidly inactivating current (fast transient outward current, ITO,f), a rapidly activating slowly inactivating current (slow transient outward current, ITO,s), and a rapidly activating noninactivating current (ISS) (5, 6). Although the kinetics of atrial ISS are similar to those of ventricular ISS, which is inhibited by acidosis, rat atrial ISS is carried by Kv1.5 (1, 2, 4, 25), whereas rat ventricular ISS appears to be carried by Kv1.2 or Kv2.1 (29). Thus the response of the action potential to acidosis may be different in rat atrial and ventricular cells.

Acidosis also depolarizes the resting membrane potential in ventricular cells, but the mechanism is unknown (26). Although a change in resting membrane potential will contribute to the electrophysiological response to acidosis, the response of the resting membrane potential to acidosis in atrial cells is unknown.

In the present study we investigated the effect of acidosis on the action potential and resting membrane potential and the underlying currents in rat atrial cells.


    METHODS
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

Cell Isolation

Male Wistar rats weighing 220~250 g were stunned and then killed by cervical dislocation. The heart was quickly removed and washed with isolation solution (see Solutions and Chemicals) containing 0.5 mM CaCl2. The aorta was cannulated and retrogradely perfused with isolation solution containing 0.5 mM CaCl2 at 10-12 ml/min and at a temperature of 36.5 ± 0.5°C. After good contraction of the heart was confirmed, it was perfused with isolation solution containing 0.1 mM EGTA (Sigma Chemical, St. Louis, MO) for 4 min and then with isolation solution containing 0.8 mg/ml collagenase (Type I, Worthington Biochemical) and 0.08 mg/ml protease (Type XIV, Sigma) for 10 min. At the end of the perfusion, both atrial appendages were dissected from the heart and cut into small pieces, which were incubated with isolation solution containing 1% bovine serum albumin in addition to the above enzymes, for 10 min at 37°C. The tissue was filtered and the filtrate centrifuged at 40 g for 60 s. The supernatant was removed, and the cells were resuspended in isolation solution containing 0.5 mM CaCl2 and stored at room temperature. This process was repeated four to five times with the remaining tissue.

Experimental Setup

An aliquot of cells was transferred to the experimental chamber, which was mounted on the stage of an inverted microscope (Diaphot, Nikon; Tokyo, Japan). In some experiments, the cells were illuminated with long wavelength (>600 nm) light, and the image was detected using a CCD camera (WAT-92A, Watec, Japan) and monitor (model PM-950, Ikegami Tsushinki; Utsunomiya, Japan).

The patch-clamp amplifier (8800 Total Clamp System; Dagan, MN) was controlled by a personal computer through an analog-digital interface (model 1401, Cambridge Electronic Design; Cambridge, UK) by using CED Patch and Voltage Clamp Software (Cambridge Electronic Design), which was also used for data acquisition. Voltage and current were also monitored with the use of an oscilloscope (OS4100, Gould; Hainault, UK).

Measurement of Action Potential and K+ Currents

The perforated patch-clamp technique was used to monitor the action potential and K+ currents (14, 15, 19, 20). Electrodes (2-4 MOmega ) were made from glass capillaries (GC150F-15, Clark Electromedical Instruments; Reading, UK) by using a vertical puller (PP-83, Narishige; Tokyo, Japan). The tip of the electrode was filled with pipette solution (see Solutions and Chemicals) and backfilled with pipette solution containing 200-400 µg/ml amphotericin B. When the electrode tip was in the bath, junction potential was electronically offset to zero. After a seal was made (>1 GOmega ), capacitance was compensated and holding potential was set to -40 mV. Five-millivolt depolarizing steps (20 ms in duration) were applied to monitor pore formation. The pipette solution contained 1 mM CaCl2 to ensure that accidental rupture of the membrane resulted in cell death. Electrical access was usually obtained within 15-30 min. After the capacity transients became constant, series resistance was compensated and measurements were made. Changing between control and acid solutions while the electrode tip was in the bath solution was used to ensure the absence of artifactual changes of potential. Action potentials were measured in current clamp mode. Three- to five-millisecond current pulses, sufficient to trigger an action potential, were injected every 2 s from the amplifier. The duration of the action potential was measured at 25, 50, and 90% repolarization (APD25, APD50, and APD90, respectively). Depolarization-induced K+ currents were recorded in the presence of 0.1 mM CdCl2 (as were the recordings shown in Fig. 3 when K+ was replaced with Cs+) to block Ca2+ current (ICa). 4-Aminopyridine (4-AP) was not used for these experiments because of its nonspecific inhibitory effects on K+ currents (5). To measure ITO,f, holding potential was set to -80 mV. After a 40-ms prepulse to -40 mV [to inactivate Na+ current (INa)], 500-ms test pulses to voltages between -40 and +40 mV were applied; pulses were applied every 2 s. ITO,f was measured as the difference between the peak outward current and the current remaining at the end of pulse. ISS was monitored from a holding potential of -20 mV to avoid contamination by transient outward current (ITO), which is inactivated at this potential (Ref. 6, see also Fig. 3A). A series of 500-ms test pulses to voltages between -40 and +40 mV was applied; pulses were applied every 2 s. ISS was measured at the end of the pulses. Inward rectifier current (IK1) was monitored from a holding potential of -40 mV. A series of 500-ms pulses to voltages between -120 and -30 mV was applied; pulses were applied every 2 s. IK1 was measured at the end of the pulses. All experiments were performed at room temperature.

Measurement of Cl- Current

Conventional whole cell patch clamp was used to measure Cl- currents (ICl) because Cl- diffusion between the pipette and the cell interior is restricted in the perforated patch configuration. A 3 M KCl-agar bridge-AgCl pellet was used as the bath electrode. Holding potential was set to -40 mV. Two-second test pulses to voltages between -120 and +40 mV (in 20-mV increments) followed by 400-ms pulses to +40 mV were applied; pulses were applied every 10 s (10). Current was measured at 2 s. All experiments were performed at room temperature.

Solutions and Chemicals

During isolation and cell storage, the isolation solution used contained (in mM) the following: 130 NaCl, 5.4 KCl, 1.4 MgCl2 · 6H2O, 0.4 NaH2PO4, 10 creatine (Sigma), 20 taurine (Sigma), 5 HEPES (Sigma), and 10 glucose. pH was adjusted to 7.30 with NaOH. When the action potential and K+ currents were measured using the perforated patch-clamp technique, the composition of the extracellular solution was (in mM) the following: 108 NaCl, 5 KCl, 1 Na2HPO4 · 12H2O, 1 MgSO4 · 7H2O, 20 Na acetate, 10 glucose, 1 CaCl2, 10 HEPES, and 5 U/l insulin. pH was first adjusted to 7.40 with NaOH, and then HCl was used to decrease the pH of an aliquot of this solution to 6.50. The composition of the pipette solution was (in mM) the following: 130 KCl, 10 NaCl, 1.4 MgCl2 · 6H2O, 1 CaCl2, and 5 HEPES. pH was adjusted to 7.10 with KOH. When ICl was measured by using conventional whole cell clamp, the composition of the extracellular solution was (in mM) the following: 120 TEACl (Sigma), 10 CsCl (Sigma), 1 MgCl2 · 6H2O, 10 HEPES, 2 BaCl2 (Sigma), and 10 glucose, 1 CaCl2, 2 4-AP (Sigma), and 0.01 nifedipine (Sigma). pH was first adjusted to 6.5 with TEAOH (Sigma), and then an aliquot of the solution was adjusted to pH of 7.4 by further addition of TEAOH. The composition of the pipette solution was (in mM) the following: 110 CsCl, 20 TEACl, 5 MgATP (Sigma), 5 EGTA, and 5 HEPES. pH was adjusted to 7.1 with CsOH.

All chemicals were purchased from BDH Laboratory Supplies (Poole, UK) unless otherwise mentioned. Amphotericin B was purchased from Sigma, and a 50 mg/ml stock solution was made with DMSO (Sigma) immediately before the experiments and diluted into the pipette solution before use. 1,2-bis(2-aminophenoxy)ethane-N,N,N',N'-tetraacetic acid (BAPTA)-AM was purchased from Molecular Probes (Eugene, OR), and a 1 mM stock solution was made with DMSO. A 10 mM stock solution of nifedipine and a 100 mM stock solution of 4,4'-diisothiocyanostilbene-2,2'-disulfonic acid (DIDS; Sigma) were made with DMSO and kept in light-resistant containers. A 10 mM stock solution of strophanthidin (Sigma) was made with ethanol. All stock solutions were diluted into the perfusate immediately before use. Addition of DIDS (final concentration up to 0.1 mM) did not alter pH of perfusate.

Statistical Analysis

Data are expressed as means ± SE for n cells. Paired or unpaired t-tests (two-tailed) were performed as appropriate. When an unpaired t-test was used, the variances were tested by F test. Statistical significance was taken as P < 0.05.


    RESULTS
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

Effect of Acidosis on Action Potential in Atrial Myocytes

Isolated atrial myocytes were current clamped at 0.5 Hz. Figure 1 shows representative action potentials recorded at control pH and during acidosis. Acidosis significantly shortened APD90 from 79 ± 7 to 48 ± 6 ms (P < 0.05, n = 5) but did not alter APD25 (4 ± 0 ms at control pH, 4 ± 0 ms in acidosis) or APD50 (12 ± 1 ms at control pH, 11 ± 1 ms in acidosis). Acidosis also caused depolarization of the resting membrane potential from -79 ± 1 mV at control pH to -76 ± 2 mV during acidosis (P < 0.05, n = 5). This change in resting potential is similar to that reported previously in ventricular cells. However, the abbreviation of the action potential by acidosis is in contrast to the prolongation reported in ventricular cells (see INTRODUCTION).


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  		Fig. 1.   Effect of acidosis on the atrial action potential. Action potentials recorded from a representative atrial cell at control pH and during acidosis. Dashed line indicates 0 mV.

Subsequent experiments were designed to investigate the current(s) responsible for these changes. Currents were normalized to cell capacitance and presented as current density. Mean cell capacitance, calculated from capacitance transients during a 10-mV hyperpolarizing pulse from -80 mV, was 53 ± 3 pF (n = 38).

Effect of Acidosis on ITO in Atrial Myocytes

The ITO in rat atrial myocytes has two components: ITO,f and ITO,s (6), which have time constants of inactivation of ~180 ms and ~3 s, respectively (6). Thus at the end of the 500-ms voltage-clamp pulses used to monitor ITO (see METHODS), ITO,f will have decayed by ~94%, whereas ITO,s will have decreased by only ~15%. In addition, at the stimulation frequency used in the present study ITO,s will be mostly inactivated due to its slow recovery from inactivation (6). Thus the current measured as the difference between peak current and current at the end of the pulse (see METHODS) will reflect predominantly ITO,f.

Figure 2, A and B, shows original traces of total outward current in a representative cell, elicited using the protocol described in the METHODS and shown in the inset in Fig. 2A, at control pH (A) and during acidosis (B). Figure 2C shows the current at +60 mV in control and acidosis. When the currents are superimposed by offsetting one of the currents, it is apparent that the amplitude and time course of ITO,f were identical in control and acidosis (Fig. 2C, inset); because the currents can be superimposed in this way, this suggests that the acidosis-induced increase in outward current (compare Fig. 2, B with A) is a rapidly activating, noninactivating current (see Effect of Acidosis on ISS in Atrial Myocytes), but that ITO,f is unchanged during acidosis. Figure 2D shows mean current density-voltage relationships of ITO,f showing that acidosis did not alter ITO,f: current at +60 mV was 8.0 ± 1.0 pA/pF in control and 6.7 ± 1.5 pA/pF in acidosis [not significant (NS), n = 6].


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  		Fig. 2.   Effect of acidosis on depolarization-induced outward current from a holding potential of -80 mV. A and B: original traces of depolarization-induced outward current in a representative atrial cell in control (A) and acidosis (B). Inset: schematic of protocol: cell was held at -80 mV and depolarized to a series of test voltages between -40 and +60 mV for 500 ms at 0.5 Hz. Arrow indicates zero current level. Capacitance of the cell was 40 pF. C: original traces of the current at +60 mV in control (open circle ) and acidosis (). When the currents are superimposed, it is clear that the amplitude and time course were identical (inset). D: mean ± SE current density-voltage relationship of fast transient outward current (ITO,f) in atrial cells at control pH and during acidosis. Transient outward current was measured as the difference between the peak outward current and the current remaining at the end of pulses (see text for details).

At the end of the voltage-clamp pulse used to elicit ITO,f (Fig. 2), membrane current is due to a small residual ITO,f (see above), ITO,s and ISS (see Effect of Acidosis on ISS in Atrial Myocytes). Thus although ITO,s will be small under the conditions of the present experiments (see Effect of Acidosis on ITO in Atrial Myocytes), it is possible to estimate this current by subtracting ISS at the end of a 500-ms test pulse from a holding potential of -20 mV (see Effect of Acidosis on ISS in Atrial Myocytes) from the current at the end of the 500-ms test pulse to the same potential, but from a holding potential of -80 mV, used to elicit ITO (see also Ref. 6). Although small, ITO,s estimated in this way was not altered by acidosis: the current at +60 mV was 2.0 ± 0.7 pA/pF in control and 2.5 ± 0.6 pA/pF in acidosis (NS, n = 6).

Thus acidosis does not appear to alter ITO,f or ITO,s making it unlikely that changes in ITO underlie the observed changes in the action potential.

Effect of Acidosis on ISS in Atrial Myocytes

ISS was evoked from a holding potential of -20 mV to avoid contamination by ITO,f and ITO,s, which are inactivated at this potential, and was measured at the end of 500-ms test pulses (Fig. 3A, inset, and see METHODS). Figure 3A shows original traces of ISS in a representative cell at control pH. The cell is the same as presented in Fig. 2, A-C; note that ITO is absent at this holding potential (cf. Fig. 2). Figure 3B shows original traces from the same cell during acidosis showing that acidosis increased this current. Figure 3C shows the effect of acidosis on the current density-voltage relationship showing that acidosis significantly increased ISS at all potentials: at +60 mV the current increased from 10.5 ± 1.8 to 14.7 ± 2.2 pA/pF (P < 0.01, n = 6).


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  		Fig. 3.   Effect of acidosis on depolarization-induced outward current from a holding potential of -20 mV. A and B: original traces of depolarization-induced outward current in the same cell as in Fig. 2, A-C, in control (A) and during acidosis (B). Inset: schematic protocol: the cell was held at -20 mV and hyperpolarized or depolarized to a series of test voltages between -40 and +60 mV for 500 ms at 0.5 Hz. Arrow indicates zero current level. C: mean ± SE current density-voltage relationship of steady-state current (ISS) in atrial cells at control pH and during acidosis. *P < 0.05 vs. control. **P < 0.01 vs. control. D: mean ± SE current density-voltage relationship of ISS in atrial cells at control pH and during acidosis when K+ in the pipette solution and perfusate was replaced with Cs+. ISS was measured at the end of pulse as described in the text.

To determine whether this current was carried by K+, the effect of replacing the K+ in the pipette and bathing solutions with Cs+ was investigated. Figure 3D shows that Cs+ almost completely abolished this current at control pH, and that in the presence of Cs+, acidosis did not increase outward current. Thus it appears that ISS is carried by K+ and increased by acidosis.

Effect of Acidosis on ISS in Presence of Intracellular Ca2+ Buffering

In rat ventricular myocytes, acidosis increases intracellular [Ca2+] and hence Ca2+-sensitive outward current (15). It seemed possible, therefore, that the increase in ISS observed during acidosis in the present study was due to the Ca2+-sensitive outward current. The above experiment was therefore repeated in the presence of BAPTA to buffer intracellular Ca2+. After the perforated patch configuration was established, 200-ms depolarizing pulses from -40 to 0 mV were applied every 2 s, and ICa and cell contraction were monitored. The cell was then exposed to 5 µM BAPTA-AM for 10 min; contraction was completely abolished within 5 min, although ICa remained. The perfusate was changed to BAPTA-free Tyrode solution, and measurements were made as described above from a holding potential of -20 mV.

Figure 4A shows the mean current-voltage relationship obtained at control pH and during acidosis in the presence of BAPTA. At control pH, BAPTA did not significantly alter ISS at any test potential (compare Fig. 3C and Fig. 4A, open symbols). In the presence of BAPTA, acidosis still increased ISS: at +60 mV the current increased from 8.0 ± 1.1 to 9.3 ± 1.7 pA/pF (n = 7, P < 0.01) and returned to 7.6 ± 1.1 pA/pF after acidosis (NS vs. precontrol). However, Fig. 4B shows the difference current (acidosis - control; i.e., the increase in current produced by acidosis) in the absence and presence of BAPTA, showing that the increase induced by acidosis was significantly smaller in the presence of BAPTA.


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  		Fig. 4.   Effect of acidosis on depolarization-induced outward current in the presence of intracellular Ca2+ buffering. A: mean ± SE current density-voltage relationship of ISS in atrial cells in the presence of BAPTA at control pH and during acidosis. After treatment with 5 µM BAPTA/AM, the same protocol was used as in Fig. 3 (see text for details). *P < 0.05 vs. control. **P < 0.01 vs. control. B: means ± SE difference currents (acidosis-control) obtained in the absence and presence of BAPTA. *P < 0.05 vs. in the presence of BAPTA. C: action potentials recorded from a representative atrial cell after treatment with BAPTA at control pH and during acidosis. Dashed horizontal line indicates 0 mV.

Because BAPTA reduced the acidosis-induced increase of ISS, its effect on the response of the action potential to acidosis was investigated to test whether the increase in ISS might underlie the acidosis-induced abbreviation of the action potential. Figure 4C shows that BAPTA altered the configuration of the action potential at control pH (compare Fig 4C, control with Fig. 1, control), presumably by buffering the Ca2+ transient and hence inhibiting inward Na+/Ca2+ exchange current (INa/Ca) during the action potential (inhibition of the exchange using Ni2+ had a similar effect on action potential configuration, see Fig. 5). In the presence of BAPTA, acidosis caused a smaller decrease of action potential duration (Fig. 4C; APD90 decreased from 53 ± 10 to 41 ± 9 ms, n = 6, P < 0.01, i.e., by 11 ± 3 ms, compared with a decrease of 31 ± 10 ms in the absence of BAPTA).


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  		Fig. 5.   Effect of acidosis on action potentials in the presence of Ni2+. Action potentials recorded from a representative atrial cell in the presence of 5 mM Ni2+ at control pH and during acidosis. Dashed horizontal line indicates 0 mV.

These data are compatible with the idea that an acidosis-induced increase in BAPTA-sensitive and BAPTA-insensitive ISS underlie the abbreviation of the action potential observed during acidosis (see DISCUSSION).

Effect of Acidosis on Action Potential in Presence of Ni2+

It seems possible that changes in INa/Ca during acidosis might also contribute to the abbreviation of the action potential (in which case inhibition of INa/Ca by BAPTA would be expected to inhibit action potential shortening during acidosis). This seems feasible because Na+/Ca2+ exchange is inhibited by acidosis (14, 20, 28, 34) and modulated by intracellular Ca2+. To test this idea, we investigated the effect of acidosis on the action potential in the presence of 5 mM NiCl2 to block INa/Ca (8). In the presence of Ni2+, acidosis still shortened the action potential (APD90 decreased from 72 ± 17 to 53 ± 17 ms, n = 3, P < 0.05) (Fig. 5). This decrease was not significantly different from that observed in the absence of Ni2+, making it unlikely that changes in INa/Ca contribute significantly to the abbreviation of the action potential observed during acidosis (although the decrease observed in the presence of Ni2+ was slightly smaller than that observed in control, suggesting that INa/Ca may play a small role).

Effect of Acidosis on IK1 in Atrial Myocytes

IK1 is responsible for late repolarization and maintenance of the resting membrane potential. Thus IK1 was measured at control pH and during acidosis to investigate whether it could contribute to the observed changes in the action potential or resting potential. However, acidosis had no significant effect on IK1 (not shown): at -120 mV the current measured at the end of the 500-ms test pulses used was -10.5 ± 1.1 pA/pF at control pH, -10.4 ± 1.2 pA/pF in acidosis (NS, n = 5). It thus appears unlikely that IK1 plays a role in the observed changes in the action potential and resting potential.

Mechanism of the Resting Membrane Potential Depolarization During Acidosis

To investigate the mechanism responsible for the depolarization of the resting membrane potential during acidosis, the inward shift of holding current at -80 mV at control pH and during acidosis was monitored. Figure 6C shows that acidosis significantly increased the inward current at -80 mV, and that this shift was not significantly different in the presence of BAPTA, when K+ in the pipette and bathing solutions was replaced with Cs+ or when pipette and bathing K+ was replaced with Cs+, and 5 mM Ni2+, 0.1 mM Ba2+, and 20 µM strophanthidin were present in the bathing solution (extracellular MgSO4 was replaced with MgCl2, and Na2HPO4 was omitted to avoid precipitation with Ba2+). Thus it appears unlikely that changes in Ca2+-sensitive currents (including INa/Ca), ICa, K+ currents, or Na pump current (INa/K), underlie the acidosis-induced depolarization of resting membrane potential.


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  		Fig. 6.   Effect of acidosis on resting membrane potential. A: action potentials recorded from representative atrial cell in the presence of 50 µM 4,4'-diisothiocyanostilbene-2,2'-disulfonic acid (DIDS) at control pH and during acidosis. B: mean ± SE acidosis-induced depolarization of resting membrane potential (from top to bottom): in control; +5 mM Ni2+; + intracellular BAPTA; +50 µM DIDS. *P < 0.05 vs. control. C: mean ± SE acidosis-induced inward shift of holding current at -80 mV (from top to bottom) in control; + intracellular BAPTA; with replacement of K+ in the pipette solution and perfusate with Cs+; with K+ in the pipette solution and perfusate replaced with Cs+, and 5 mM Ni2+, 0.1 mM Ba2+, 20 µM strophanthidin present in the perfusate. Numbers of cells are shown in parentheses. All changes in resting membrane potential and holding current were significantly different from baseline (P < 0.05) except that in the presence of DIDS (NS, not significant).

Figure 6B shows that acidosis significantly depolarized the resting membrane potential in control, in the presence of Ni2+ and BAPTA, as expected from the results above, but that 50 µM DIDS inhibited the acidosis-induced depolarization of the resting membrane potential (Fig. 6, A and B), suggesting that the depolarization is due to acidosis-induced activation of a DIDS-sensitive ICl. To test this idea further, the effect of acidosis on ICl was investigated.

Figure 7, A and B, shows original traces of ICl recorded from a representative cell at control pH (Fig. 7A) and during acidosis (Fig. 7B) in the absence of Na+ and K+ and the presence of Cs+, TEA+, Ba2+, nifedipine, 4-AP, and internal EGTA as described in METHODS. Under these conditions (pipette [Cl-] = 130 mM), an inwardly rectifying current was recorded with a reversal potential of -10 mV (Fig. 7C, open circles); reducing pipette [Cl-] to 20 mM (replaced with aspartate-) markedly decreased the measured current and shifted the reversal potential to -46 mV (Fig. 7C, open triangles). The similarity of the measured reversal potentials to the calculated Cl- reversal potentials (-2 mV and -49 mV, respectively) suggests that the measured current is carried predominantly by Cl-; the difference between the calculated and measured reversal potentials may represent imperfect equilibration between pipette and intracellular solutions. Acidosis increased inward current (Fig. 7C, filled symbols): when pipette [Cl-] was 130 mM, current at -120 mV increased from -4.0 ± 2.1 to -6.5 ± 2.5 pA/pF (n = 5, P < 0.05). Figure 7D shows that the current induced by acidosis (i.e., current in acidosis minus that at control pH) shows strong inward rectification and is decreased when pipette [Cl-] is decreased: the current induced by acidosis at -120 mV was reduced from -2.5 ± 0.6 pA/pF (n = 5) to -0.4 ± 0.2 pA/pF (n = 3; P < 0.05) when pipette [Cl-] was reduced from 130 to 20 mM. The acidosis-induced increase in current was completely inhibited by DIDS (Fig. 7E), although DIDS had no significant effect on membrane current at control pH under these conditions or on resting membrane potential under control conditions (not shown). These data suggest, therefore, that the acidosis-induced depolarization of the resting membrane potential is due to activation of a DIDS-sensitive, inwardly rectifying ICl (ICl,ir).


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  		Fig. 7.   Effect of acidosis on Cl- current. A and B: original traces of Cl- current monitored using ruptured patch in a representative atrial cell at control pH (A) and during acidosis (B); pipette [Cl-] 130 mM. Cell was held at -40 mV and hyperpolarized to a series of test voltages between -120 and +40 mV in 20-mV increments for 2 s followed by 400-ms pulses to +40 mV at 0.1 Hz. Arrow indicates zero current level. C: means ± SE current density-voltage relationships of Cl- current at control pH (open symbols) and during acidosis (filled symbols) when pipette [Cl-] was 130 mM (circles) or 20 mM (triangles). *P < 0.05 vs. control pH. D: means ± SE difference current (acidosis - control) when pipette [Cl-] was 130 mM () or 20 mM (black-triangle). *P < 0.05 vs. 130 mM pipette [Cl-]. E: means ± SE difference current in the absence () and presence () of 0.1 mM DIDS; pipette [Cl-] 130 mM. *P < 0.05 vs. in the absence of DIDS.

Relationship Between Resting Membrane Potential and Action Potential Duration

To test whether the observed change in action potential duration might be secondary to the acidosis-induced change in resting membrane potential per se, current injection was used to depolarize the resting membrane potential by 5 mV. However, this maneuver tended to prolong action potential duration (not shown), making this unlikely.


    DISCUSSION
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

In the present study, acidosis was produced by decreasing the extracellular pH (pHo) from 7.4 to 6.5. Measurement of intracellular pH (pHi) using the fluorescent dye 2',7'-bis(2-carboxyethyl)-5(6)-carboxyfluorescein has shown that pHi decreases by ~0.5 pH units in response to this change in pHo (20). These changes are, therefore, within the range observed pathophysiologically (27). pHi reaches a new steady state within 3 min; measurements in the present study were therefore carried out after 5 min exposure to the acid solution (15). In ventricular cells, the presence of a perforated patch electrode containing 5 mM HEPES does not affect the change of pHi that occurs on exposure to the acid solution (not shown); it therefore seems unlikely that it would do so in atrial cells. The perforated patch-clamp technique was used because acidosis alters the intracellular environment, increasing intracellular [Ca2+] (12), intracellular [Na+] (12), calcium-calmodulin-dependent kinase (CaMKII) activity via the increase in intracellular [Ca2+] (13) and inhibiting protein phosphatase activity (23). We therefore used the perforated patch-clamp technique where possible to minimize disruption to these normal responses, to investigate the physiological response to acidosis.

Effect of Acidosis on Atrial Action Potential

Role of ICa and INa/Ca. We have previously shown that acidosis has little effect on ICa in rat ventricular cells when the perforated patch-clamp technique is used (14, 19, 20). Because the channel that carries this current in atrial cells appears to be the same as in ventricular cells, it seems likely that acidosis has little effect on ICa in atrial cells. Although inhibition of ICa by Ni2+ may contribute to the change in action potential configuration observed in the presence of Ni2+, the observation that acidosis-induced shortening of the action potential still occurred in these conditions suggests that changes in ICa are not necessary to account for this abbreviation of the action potential.

Inhibition of Na+/Ca2+ exchange shortens action potential duration (30). Acidosis-induced inhibition of Na+/Ca2+ exchange (14, 20, 28, 34) could, therefore, explain the observed abbreviation of the action potential during acidosis. However, the present study shows that acidosis shortened the action potential even after the exchange had been inhibited by 5 mM Ni2+ (Fig. 5), making it unlikely that inhibition of Na+/Ca2+ exchange plays a major role in the acidosis-induced shortening of the action potential, although it is possible that it plays a small role (see RESULTS).

Role of ITO,f and ITO,s. In the present study, acidosis did not alter ITO,f (Fig. 2) consistent with the observation that acidosis did not alter early repolarization (Fig. 1). Although atrial ITO,f is distinct from ventricular ITO with respect to its time course of activation and inactivation, the same channels (Kv4.2/4.3) are thought to underlie both currents. The present result is therefore consistent with the observation that acidosis does not alter ITO in rat ventricular cells when holding potential is negative to -60 mV (15). These data make it unlikely that the effect of acidosis on the action potential is mediated by a change in ITO,f.

It is also unlikely that ITO,s is involved in the acidosis-induced shortening of the action potential because ITO,s is mostly inactivated in the conditions used in the present study and appears to be unaffected by acidosis (see RESULTS), consistent with previous reports that Kv1.2, which appears to carry ITO,s, is unaffected by acidosis (32).

Role of ISS. Kv1.5 is the most likely candidate for rat atrial ISS (25), because the time- and voltage-dependent properties of heterologously expressed Kv1.5 channels are similar to those of rat atrial ISS, and anti-Kv1.5 antibody shows high levels of binding in rat atrial cells (2) and exposure of rat atrial cells to antisense oligodeoxynucleotides for Kv1.5 significantly inhibits ISS (4).

The present data show that ISS is inhibited by replacing K+ with Cs+, consistent with a K+ current. ISS also appeared to have a Ca2+-insensitive component and a Ca2+-sensitive component that was only observed during acidosis (BAPTA decreased ISS during acidosis but had no effect on ISS at control pH; Figs. 3 and 4). We are unaware of any previous work showing that ISS is Ca2+ dependent or has two components. However, it is not clear whether these are two different components of ISS or whether the apparently Ca2+-insensitive component is due to incomplete buffering of Ca2+ by BAPTA. In support of the latter suggestion, both components showed the same kinetics (rapidly activating, noninactivating; not shown), and acidosis increased both components of ISS (Fig. 4B).

Previous work has shown that acidosis decreases ISS in rat ventricular cells (see INTRODUCTION), although this may be because in rat ventricle ISS is carried by Kv1.2 or Kv2.1 (29). However, acidosis (pH 7.3-6.3) also inhibits Kv1.5 current expressed in Xenopus oocytes (32), although this difference may be due to differences in the channel's environment and regulation in the two cell types (e.g., 9). The mechanism of action of acidosis on ISS is unclear but is likely to be an effect on channel conductance rather than inactivation; the observation that the increase in ISS produced by acidosis is voltage independent (Fig. 3C) also suggests that it is unlikely that the effect of protons is within the channel pore.

The observation that BAPTA inhibited the acidosis-induced decrease in action potential duration (Fig. 4C) is compatible with the idea that an acidosis-induced increase in the BAPTA-sensitive component of ISS is responsible for much of the action potential shortening observed during acidosis.

Role of IK1. In the present study, acidosis did not alter IK1, making it unlikely that changes in IK1 are responsible for the changes in the action potential observed during acidosis.

IK1 in the rat ventricle appears to be carried by Kir2.1 (24). Although the identity of rat atrial IK1 has not been established, Kir2.1 is pH insensitive (37), consistent with the present data.

Mechanism of acidosis-induced shortening of the atrial action potential. In the present study, acidosis shortened the action potential of rat atrial cells. This is in contrast to previous work, which showed that acidosis prolongs the action potential of rat ventricular cells under identical experimental conditions. The abbreviation observed in the present study is unlikely to be due to the effects of acidosis on INa/Ca, ICa, ITO,f, ITO,s IK1, or ICl,ir, or secondary to the depolarization of the resting membrane potential (see above), but could be due predominantly to the effect of acidosis on ISS, which is increased in atrial cells, but inhibited in ventricular cells and which could, therefore, account for the different response in the two cell types. Unfortunately, there is no specific inhibitor of ISS with which to test this hypothesis, although the observation that BAPTA inhibited the increase in ISS and the abbreviation of the action potential supports the idea.

One problem with this hypothesis is that the effect of acidosis on ISS is greater at more positive potentials (Fig. 3), whereas the effect on the action potential is greater at more negative potentials (Fig. 1). Because many of the repolarizing currents are voltage and time dependent, the comparison of the action potential and the currents monitored under voltage clamp is not straightforward. However, a similar relationship between the configuration of the action potential and a change in ISS to that observed in the present study has been reported in response to phenylephrine (11), which produced a prolongation of late depolarization that was ascribed to a decrease in ISS. It seems possible that the effect of the acidosis-induced increase in ISS on the action potential is masked during the early phase (APD25, APD50) of the action potential by the presence of other large currents (e.g., INa, ITO,f) and thus that an increase in ISS, possibly with a small contribution by altered INa/Ca (see RESULTS), could underlie the abbreviation of the atrial action potential during acidosis.

Effect of Acidosis on Resting Potential

Acidosis-induced depolarization of the resting membrane potential has been observed by many investigators [for review see Orchard and Cingolani (26)], although the mechanism has remained obscure. The present study shows that the acidosis-induced increase in inward current, and hence resting membrane potential, is unlikely to be due to changes in Ca2+-sensitive currents (including INa/Ca), ICa, K+ currents, or INa/K (Fig. 6, B and C).

However, acidosis markedly increased ICl,ir, monitored using whole cell clamp (Fig. 7). This current and the acidosis-induced increase in current were reduced when pipette [Cl-] was decreased (Fig. 7, C and D): when pipette [Cl-] was 20 mM, within the physiological range (10-20 mM; Ref. 16), the acidosis-induced increase in current at -80 mV was 0.19 ± 0.12 pA/pF, similar to the increase in inward current observed during acidosis using perforated patch-clamp recording under more physiological conditions (~0.5 pA/pF; Fig. 6). The slightly higher current observed during perforated patch recording might be due to the high [Cl-] in the pipette and a small permeability of the perforated patch to Cl- causing a small increase in intracellular [Cl-]. The observation that the increase in inward current induced by acidosis was inhibited by decreasing pipette [Cl-] and by DIDS (Fig. 7, D and E), which also abolished the depolarization of the resting membrane potential (Fig. 6), suggests that the depolarization of the resting membrane potential is due to acidosis-induced activation of a DIDS-sensitive ICl,ir. Assuming a membrane resistance of 118 MOmega (range in rat ventricular myocytes 20-200 MOmega , Ref. 36), the small (28 pA) inward shift of current at -80 mV could account for the 3.3-mV depolarization observed during acidosis.

ICl,ir does not appear to be due to protein kinase A-dependent ICl, Ca2+-activated Cl- current, or swelling-induced ICl, because these do not show inward rectification (18, 31, 16). However, this current shows similarities to that carried by volume-regulated Cl (ClC-2) channels, which have recently been found in atrial and ventricular cells (7, 10); the ICl carried by these channels shows inward rectification (10, 17), is of comparable magnitude to that recorded in the present study (10), and is increased by acidosis in Xenopus oocytes (17). The presence of Cd2+, which blocks ClC-2, during measurement of depolarization-induced K+ currents (see METHODS) could then explain why this current was not observed during these K+ current measurements, although the amplitude of ICl,ir at physiological intracellular [Cl-] is sufficiently small (Figs. 6 and 7) that it would be difficult to detect during measurement of relatively large K+ currents, even in the absence of Cd2+. One problem with this hypothesis is that these channels have been reported to be insensitive to stilbene derivatives such as DIDS, which agrees with the observation in the present study that DIDS did not alter ICl,ir at control pH. It is possible, therefore, that acidosis alters the response to DIDS, either by affecting DIDS itself, or DIDS may block the stimulatory action of H+ on ClC-2. Alternatively acidosis may increase ICl,ir, not by acting on the channel, but by causing a DIDS-sensitive increase of intracellular [Cl-], although the possible mechanism is unclear: the Cl-/OH- exchange (or H+-Cl- coinflux) mechanism described by Sun et al. (33) is DIDS insensitive, whereas the DIDS-sensitive Na+-HCO<UP><SUB>3</SUB><SUP>−</SUP></UP> cotransporter and Cl-/HCO3- exchanger (22) would be inhibited in the HEPES-buffered (HCO<UP><SUB>3</SUB><SUP>−</SUP></UP> free) solutions used in the present study (33). In either case, it appears likely that ClC-2 channels carry the acidosis-induced ICl,ir that underlies the depolarization of the resting membrane potential during acidosis.


    ACKNOWLEDGEMENTS

This work was supported by British Heart Foundation and Wellcome Trust.


    FOOTNOTES

Address for reprint requests and other correspondence: C. H. Orchard, School of Biomedical Sciences, Univ. of Leeds, Leeds LS2 9NQ, UK (E-mail: c.h.orchard{at}leeds.ac.uk).

The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

April 11, 2002;10.1152/ajpheart.01000.2001

Received 14 November 2001; accepted in final form 8 April 2002.


    REFERENCES
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ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
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