Preferential regulation of rabbit cardiac L-type Ca²⁺ current by glycolytic derived ATP via a direct allosteric pathway

Abstract

1. The activity of Ca²⁺ channels is regulated by a number of mechanisms including direct allosteric modulation by intracellular ATP. Since ATP derived from glycolysis is preferentially used for membrane function, we hypothesized that glycolytic ATP also preferentially regulates cardiac L-type Ca²⁺ channels.

2. To test this hypothesis, peak L-type Ca²⁺ currents (I_Ca) were measured in voltage-clamped rabbit cardiomyocytes during glycolytic inhibition (2-deoxyglucose + pyruvate), oxidative inhibition (cyanide + glucose) or both (full metabolic inhibition; FMI).

3. A 10 min period of FMI resulted in a 40·0 % decrease in peak I_Ca at +10 mV (-5·1 ± 0·6 versus -3·1 ± 0·4 pA pF⁻¹; n= 5, P < 0·01). Similar decreases in peak I_Ca were observed during glycolytic inhibition using 2-deoxyglucose (-6·2 ± 0·2 versus -3·7 ± 0·2 pA pF⁻¹; n= 5, P < 0·01) or iodoacetamide (-6·7 ± 0·3 versus -3·7 ± 0·2 pA pF⁻¹; n= 7, P < 0·01), but not following oxidative inhibition (-6·2 ± 0·4 versus -6·4 ± 0·3 pA pF⁻¹; n= 5, n.s.). The reduction in I_Ca following glycolytic inhibition was not mediated by phosphate sequestration by 2-deoxyglucose or changes in intracellular pH.

4. Reductions in I_Ca were still observed when inorganic phosphate and creatine were included in the pipette, confirming a critical role for glycolysis in I_Ca regulation.

5. With 5 mM MgATP in the pipette during FMI, peak I_Ca decreased by only 18·4 % (-6·8 ± 0·6 versus -5·5 ± 0·3 pA pF⁻¹; n= 4, P < 0·05), while inclusion of 5 mM MgAMP-PCP (β,γ-methyleneadenosine 5′-triphosphate, Mg²⁺ salt) completely prevented the decrease in peak I_Ca (-6·9 ± 0·3 versus -6·5 ± 0·3 pA pF⁻¹; n= 5, n.s.).

6. Together, these results suggest that I_Ca is regulated by intracellular ATP derived from glycolysis and does not require hydrolysis of ATP. This regulation is expected to be energy conserving during periods of metabolic stress and myocardial ischaemia.

In the myocardium, as well as other tissues, ATP synthesis and utilization are subcellularly localized (Lynch & Paul, 1987). Glycolytic enzymes are known to be associated with the cardiac sarcolemma (Pierce & Philipson, 1985; Hazen et al. 1994) and sarcoplasmic reticulum (Pierce & Philipson, 1985). Accordingly, ATP derived from glycolysis has been shown to preferentially support membrane function while ATP generated from oxidative phosphorylation supports contractile function (Weiss & Hiltbrand, 1985). For instance, glycolytic ATP has been shown to preferentially regulate cardiac, as well as skeletal, sarcoplasmic reticular Ca²⁺ transport (Xu et al. 1995). Furthermore, glycolytic ATP regulates the activity of both the sarcolemmal Ca²⁺ pump (Paul et al. 1989; Hardin et al. 1992) and the Na⁺-K⁺ pump (Lynch & Balaban, 1987; Campbell & Paul, 1992) in smooth muscle cells.

Preferential regulation of ATP-sensitive K⁺ channels by glycolysis has been demonstrated in guinea-pig cardiac myocytes (Weiss & Lamp, 1987, 1989). Regulation of the ATP-sensitive K⁺ channel by ATP is the prototype for a putative metabolic feedback of cellular excitability and contractility. As intracellular ATP concentrations decrease, ATP-sensitive K⁺ channels open in a dose-dependent fashion in guinea-pig myocytes (Deutsch & Weiss, 1993) resulting in K⁺ efflux, shortening of the action potential and reduced excitability (Deutsch et al. 1991; Weiss et al. 1992). Ultimately, reduction of excitability reduces contractility preserving intracellular ATP and conferring protection against metabolic stress (Critz et al. 1997).

Like ATP-sensitive K⁺ channels, L-type Ca²⁺ channels are also regulated by ATP. Intracellular injection of ATP in whole-cell patch-clamp studies of guinea-pig myocytes resulted in a 40-60 % prolongation of action potential duration (Taniguchi et al. 1983), which correlated with a corresponding increase in L-type Ca²⁺ current (I_Ca) (Irisawa & Kokubun, 1983). One potential mechanism for this regulation is that intracellular ATP acts as a substrate for phosphorylation of the Ca²⁺ channel by protein kinases (for a review see Hilgemann, 1997). Alternatively, intracellular ATP also regulates Ca²⁺ channel activity directly via an allosteric mechanism not requiring ATP hydrolysis (O'Rourke et al. 1992; Yazawa et al. 1997; Yokoshiki et al. 1997) as originally proposed by Reuter & Scholtz (1977). In contrast to the regulation of ATP-sensitive K⁺ channels by free ATP, direct regulation of I_Ca requires MgATP (Backx et al. 1991; O'Rourke et al. 1992). Direct phosphorylation-independent regulation of I_Ca has also been demonstrated in smooth muscle cells from rat mesenteric artery (Yokoshiki et al. 1997) and this regulation appeared to rely preferentially on ATP derived from glycolysis (Lorenz & Paul, 1997).

Given that cardiac L-type Ca²⁺ channels are regulated by ATP and that glycolytic ATP has been shown to preferentially influence membrane proteins, we hypothesized that ATP derived from glycolysis would have a more direct influence on cardiac L-type Ca²⁺ channels compared with oxidative phosphorylation. In addition, we hypothesized that this regulation may involve a direct phosphorylation-independent mechanism. We tested these hypotheses in whole-cell voltage-clamped rabbit ventricular myocytes, which were exposed to glycolytic inhibition, oxidative inhibition or both glycolytic and oxidative inhibition (full metabolic inhibition; FMI). Our results establish that peak I_Ca is preferentially regulated by glycolytic ATP via a direct allosteric pathway.

METHODS

Preparation of single cells

Single ventricular cells were obtained from rabbit hearts (New Zealand White, 2-3 kg) using a modification of a previously described technique (Backx & Marban, 1993). All experiments were carried out with the approval of the Animal Care Committee at the Toronto General Hospital in accordance with the guidelines set by the Canadian Council for Animal Care. Briefly, each rabbit was anaesthetized with a mixed solution of pentobarbitone (60 mg kg⁻¹) and heparin (200 i.u. kg⁻¹) given as two intravenous bolus doses separated by 5 min. Before respiratory arrest occurred, the chest was opened through a mid-line sternotomy and the heart was quickly excised, mounted by the aortic root onto a cannula and retrogradely perfused with a modified Hepes-Tyrode solution with 1.0 mM Ca²⁺ and bubbled with 100 % O₂ at 37°C for 2-3 min to wash out all blood residue. The heart was then perfused with a nominally Ca²⁺-free Hepes-Tyrode solution for 10 min and finally with the same Ca²⁺-free solution containing 200 U ml⁻¹ collagenase Type II (Worthington Biochemical Corporation, Lakewood, NJ, USA) for 25-30 min. The enzyme solution was subsequently washed out with a high-K⁺ solution for 5 min. The ventricles were then removed and dissected into small pieces and single cells were isolated by gentle mechanical agitation. Cells were filtered through a nylon mesh, resuspended in high-K⁺ solution with 2 % bovine serum albumin (Sigma-Aldrich) and used within 8 h. Calcium-tolerant, rod-shaped cells with clear, regular cross-striations and no visible blebs were used for electrophysiological recordings.

Solutions and drugs

The modified Hepes-Tyrode solution for cell isolation contained (mM): NaCl, 140; MgCl₂, 1; Hepes, 10; KCl, 4; CaCl₂, 1; D-glucose, 5; pH adjusted to 7.4 with NaOH. The high-K⁺ cell storage solution contained (mM): potassium glutamate, 120; KCl, 20; Hepes, 20; MgCl₂, 1; D-glucose, 10; and K-EGTA, 0.3; pH 7.4. The control bath (external recording) solution contained (mM): NaCl, 145; MgCl₂, 1; CsCl, 5; Hepes, 10; CaCl₂, 1.8; pH adjusted to 7.4 with NaOH. D-Glucose (10 mM) and pyruvic acid (2 mM) were added to the control bath solution as substrate for glycolysis and oxidative phosphorylation, respectively. The control pipette (internal recording) ATP-free solution contained (mM): CsCl, 145; Hepes, 10; MgCl₂, 1; EGTA, 5; pH adjusted to 7.2 with CsOH; the calculated free Mg²⁺ concentration under these conditions was 0.79 mM (CHELATOR Version 1.00; Schoenmakers et al. 1992).

FMI was achieved by the addition of 10 mM 2-deoxyglucose to inhibit glycolysis and 2 mM sodium cyanide to inhibit oxidative phosphorylation, in the absence of substrate (i.e. glucose and pyruvate). Inhibition of glycolysis alone was achieved by inclusion in the bath solution of 10 mM 2-deoxyglucose and 2 mM pyruvic acid (substrate for oxidative phosphorylation). In addition to competitively inhibiting glucose utilization, 2-deoxyglucose also inhibits glycogen phosphorylase (Dringen & Hamprecht, 1993; Oikonomakos et al. 1995) and activates glycogen synthase (Carabaza et al. 1992). To determine the specificity of 2-deoxyglucose we used 100 μM iodoacetamide (IAA), a known inhibitor of glyceraldehyde-3-phosphate dehydrogenase, to inhibit glycolysis. Inhibition of oxidative phosphorylation alone was achieved by inclusion of 2 mM sodium cyanide in the control bath solution containing 10 mM glucose as substrate for glycolysis. In a separate set of experiments 5 mM ATP or 5 mM β,γ-methyleneadenosine 5′-triphosphate (AMP-PCP) complexed as Mg²⁺ salts was included in the control pipette solution. The estimated free Mg²⁺ concentration in the pipette solutions was 1.1 mM for MgATP or MgAMP-PCP (CHELATOR Version 1.00). To investigate the role of the creatine kinase, experiments were conducted with either 1.2 mM NaH₂PO₄ in the bath and 3.0 mM NaH₂PO₄ in the pipette solution, or the same concentrations of NaH₂PO₄ and 20 mM creatine in the pipette solution.

Glibenclamide (1 μM) was added to the bath solution (from a 10 mM stock dissolved in 100 % DMSO) to inhibit ATP-sensitive K⁺ channels during metabolic inhibition. R(+)-methylindazone (IAA94; 10 μM) or 5-nitro-2-(3-phenylpropylamino) benzoic acid (NPPB; 1 μM) were added to the bath solution to block Cl⁻ channels (from a 50 mM stock dissolved in 100 % ethanol). IAA94 and NPPB were purchased from Research Biochemicals International. Glibenclamide, creatine, ATP, AMP-PCP, iodoacetamide, cyanide, 2-deoxyglucose and all other analytical grade chemicals were purchased from Sigma-Aldrich.

Electrophysiological recording

Cells were transferred into a 1 ml perfusion bath placed on the stage of an inverted microscope and superfused with control bath solution containing glucose and pyruvate at a rate of 1-2 ml min⁻¹. All experiments were performed at room temperature (20-22°C). The gigaseal, whole-cell voltage-clamp configuration of the patch-clamp technique (Hamill et al. 1981) using an Axopatch 200A or 200B amplifier (Axon Instruments) was utilized. A ground silver-silver chloride wire enclosed in an agar bridge (Backx & Marban, 1993) was used to avoid oxidation of the silver upon wash-in of cyanide and subsequent junction potential shifts. Recording pipettes were prepared from thin-walled borosilicate glass (1.5 mm diameter; World Precision Instruments) using a Flaming-Brown micropipette puller (model P-87; Sutter Instruments). The pipette tip was heat polished with a heating filament (model MF-83; Narishige, Japan) and when filled with internal solution, tip resistance was typically 2-3 MΩ. Uncompensated series resistance was typically 2-6 MΩ and compensation was 60-70 %. After membrane rupture, the cell capacitance was estimated automatically by custom-designed computer software and by integrating the area under the capacity transient generated from a 20 mV depolarization from a holding potential of -40 mV. Currents were digitized at 10 kHz, filtered at 2 kHz and stored off-line for analysis.

To isolate I_Ca, internal and external K⁺ was replaced by Cs⁺, Cl⁻ currents were blocked with 10 μM IAA94 or 1 μM NPPB, potent inhibitors of Cl⁻ channels (Pope et al. 1991), and Na⁺ channels were inactivated by a 100 ms depolarizing prepulse to -40 mV. After completion of metabolic inhibition, nifedipine (10 μM) or Cd²⁺ (300 μM) was added to the bath solution to obtain nifedipine- or Cd²⁺-sensitive I_Ca. Peak I_Ca was sampled every 5 s by 200 ms depolarizing pulses to 0 mV from a holding potential of -80 mV, after 100 ms prepulses to -40 mV. Peak I_CaI-V relationships were measured between -40 and +80 mV in 10 mV increments. During the time course of our metabolic inhibition experiments, leak currents were estimated by measuring the current recorded at the holding potential (-80 mV) and the prepulse potential (-40 mV).

Experimental protocols and data analysis

This study consisted of eight groups of cells each representing different metabolic interventions in the absence or presence of ATP or AMP-PCP in the pipette solution and each cell was used as its own control. After making a gigaseal, gaining access into the myocytes and compensating for cell capacitance and series resistance, myocytes were repeatedly depolarized to 0 mV from a holding potential of -80 mV with a 100 ms prepulse to -40 mV. This was done for 5 min while the cells were superfused with external control solution containing glucose and pyruvate in order to allow complete cellular dialysis and endogenous run-down of I_Ca. Following the 5 min equilibration period, a control peak I_CaI-V relationship (depolarizations from -40 to +80 mV) was recorded. This was followed immediately by application of bath solution containing the desired metabolic substrates and inhibitors, while the myocyte was again depolarized every 5 s. After a 10 min exposure to the metabolic inhibitors, a test I-V relationship was recorded and compared with the control I-V relationship. At the end of the protocol a nifedipine (10 μM) or Cd²⁺ (300 μM) current was collected.

The first group of cells was exposed to 2-deoxyglucose and sodium cyanide (FMI) in the absence of the metabolic substrates glucose and pyruvate. In the second group of myocytes, addition of 2-deoxyglucose in the continued presence of pyruvate to maintain oxidative phosphorylation exclusively inhibited glycolysis. The third group of cells was exposed to sodium cyanide to inhibit oxidative phosphorylation in the continued presence of glucose to support glycolysis. In a fourth and fifth group of myocytes, inorganic phosphate alone or inorganic phosphate and creatine, respectively, were utilized under control conditions and during glycolytic inhibition with 2-deoxyglucose. In a sixth group of cells IAA (100 μM) was used to inhibit glycolysis as opposed to 2-deoxyglucose. The above experiments on cells from groups one to six were conducted in the absence of exogenous ATP. The seventh group of cells was exposed to FMI while exogenous ATP was provided via the pipette. The eighth group of cells was also exposed to FMI but with AMP-PCP complexed with Mg²⁺ in the pipette solution.

Curve fitting

Custom-designed and commercially available Microcal Origin (Microcal Software Inc., Northampton, MA, USA) software was used to analyse data. Limiting maximal slope conductances were calculated by fitting the I-V data below +40 mV with the Boltzmann distribution function. The time of half I_Ca inactivation was calculated from the ratio of I_Ca at +10 mV (peak) and I_Ca at the end of the depolarizing step and is referred to as time to half I_Ca inactivation (t_½). All data are presented as means ±s.e.m. and Student's t test for paired data was used for statistical analysis. An experimental α-level of P < 0.05 was considered statistically significant.

RESULTS

Metabolic inhibition in the absence of exogenous ATP

Figure 1A and B summarizes the changes in I_Ca due to endogenous run-down. In these experiments, myocytes were initially superfused with an external Tyrode solution (see Methods) containing 10 mM glucose and 2 mM pyruvate as substrates for glycolysis and oxidative phosphorylation, respectively, with no ATP added to the pipette. After gaining access, myocytes were dialysed for 2-3 min while series resistance was compensated. Next, from a holding potential of -80 mV, I_Ca was then recorded following prepulses to -40 mV followed by depolarizations to 0 mV applied every 5 s for a period of 5 min. This period allowed complete cell dialysis, equilibration and endogenous run-down of I_Ca. Figure 1A indicates that, under our conditions, I_Ca decreased by 10-20 % within the first 3-4 min of recording. After 3-4 min, peak I_Ca remained relatively constant for at least 20 min as shown by the raw traces in Fig. 1A (inset). In fact, no significant difference was observed between mean peak I_Ca recorded at 0 mV 7-8 min after the whole-cell configuration was established (open circle; 6.3 ± 0.4 pA pF⁻¹) and 20 min later (filled circle; 5.8 ± 0.5 pA pF⁻¹) as illustrated in Fig. 1B. In all experiments in this study, we allowed the myocytes to equilibrate for 7-8 min prior to recording a control I-V relationship. Thereafter the effects of metabolic interventions were measured during the subsequent period, which was 10 min in duration.

Figure 1. Effect of full metabolic inhibition on L-type Ca²⁺ current.

A, once the whole-cell configuration was achieved, I_Ca was sampled at 0.2 Hz and I-V relationships were recorded at the times indicated by the open and filled circles. The typical raw traces (inset) indicate that I_Ca run-down was minor over the 10 min sampling period. B, mean I-V relationships showing that sampling I_Ca at 0.2 Hz for 10 min did not significantly reduce I_Ca under our conditions (n= 4). C, I_Ca was sampled for 5 min before the indicated interruption in order to record a control I-V relationship. This was followed by external wash-in (arrow) of 2-deoxyglucose (10 mM) and sodium cyanide (2 mM) (full metabolic inhibition, FMI). The raw traces (inset) demonstrate that I_Ca prior to FMI (open circle) was larger than I_Ca after FMI (filled circle). D, mean I-V relationships which show that, on average, 10 min of FMI resulted in a 40 % reduction of I_Ca at +10 mV (n= 5, P < 0.01).

Figure 1C shows the peak I_Ca amplitude before and after (indicated by the arrow) application of an external solution containing 10 mM 2-deoxyglucose and 2 mM sodium cyanide (FMI), inhibitors of glycolysis and oxidative phosphorylation, respectively. Note that a 10 min period of FMI (filled circle) resulted in a nearly twofold decrease in I_Ca compared with control (open circle). FMI did not affect current inactivation (i.e. t_½= 33.2 ± 1.3 ms for control versus 30.5 ± 2.6 ms for FMI; n= 5, n.s.) as illustrated by the raw traces in Fig. 1C (inset). Figure 1D shows the mean I-V relationships taken before and 10 min after FMI. On average, FMI reduced I_Ca compared with control from -5.1 ± 0.6 to -3.1 ± 0.4 pA pF⁻¹ (P < 0.01, n= 5) at +10 mV, while mean maximum slope conductance (see Methods) was decreased from 159 pS in control to 108 pS.

In all the experiments shown in Fig. 1 and subsequent experiments, I_Ca is reported as the nifedipine- or Cd²⁺-subtracted current, where nifedipine or Cd²⁺ was added at the end of the experimental protocol. This procedure is a valid measure of I_Ca provided that metabolic inhibition does not activate time-dependent leak currents. We used currents measured at -80 and -40 mV as indicators of the magnitude of leak currents during the time course of our experiments. A summary of leak currents for control, FMI, glycolytic and oxidative inhibition at -80 and -40 mV is shown in Fig. 2A. After a 10 min period of metabolic intervention, current at -80 mV was -314.1 ± 44.5 pA for FMI, -278.7 ± 52.8 pA for glycolytic inhibition and -334.8 ± 35.3 pA for oxidative inhibition, compared with -339.4 ± 22.6 pA for control at the same time point (n= 5, n.s.). Current at -40 mV was -117.2 ± 7.8 pA for FMI, -129.0 ± 35.1 pA for glycolytic inhibition and -146.2 ± 21.8 pA for oxidative phosphorylation compared with -137.6 ± 29.2 pA for control (n= 5, n.s.). Figure 2B illustrates currents recorded in the presence of 10 μM nifedipine from control and after metabolic inhibition. Following a 10 min period with control bath solution the mean current at +10 mV in the presence of nifedipine was -39.1 ± 12.2 pA compared with -35.4 ± 9.8 pA for FMI, -34.2 ± 9.8 pA for glycolytic inhibition and -36.2 ± 2.4 pA for oxidative inhibition (n= 5, n.s.). Therefore, under our experimental conditions metabolic inhibition did not activate time-dependent leak currents.

Figure 2. Holding current in control and after various metabolic interventions.

A, summary of holding current at -80 mV (▪) and -40 mV () for control, full metabolic inhibition (FMI), glycolytic inhibition (GI) and oxidative inhibition (OI). It is clearly evident that the holding current was similar at both -80 and -40 mV regardless of metabolic intervention (n= 5 for all groups). B, mean I-V relationships recorded 10 min after control conditions, FMI, GI and OI in the presence of 10 μM nifedipine were not significantly different (n= 5 for all groups).

Figure 3A illustrates the changes in peak I_Ca before and after (indicated by the arrow) application of the glycolytic inhibitor 2-deoxyglucose (10 mM) in the presence of pyruvate as substrate for oxidative phosphorylation, without ATP added to the pipette solution. The protocols used in these experiments were the same as those described in Fig. 1. For the representative cell shown in Fig. 3A, a 10 min 2-deoxyglucose exposure period reduced I_Ca recorded at 0 mV by almost twofold. The raw traces in Fig. 3A (inset) demonstrate that exposure to 2-deoxyglucose (filled circle) caused a decrease in peak I_Ca compared with control (open circle). Similar to FMI, glycolytic inhibition reduced I_Ca without significantly affecting the time course of inactivation (t_½= 30.4 ± 1.2 ms in control versus 30.4 ± 0.8 ms following glycolytic inhibition; n= 5, n.s.).

Figure 3. Effect of glycolytic and oxidative inhibition on L-type Ca²⁺ current.

A and C, in the whole-cell configuration, I_Ca was initially sampled for 5 min prior to the indicated interruption, in order to record a control I-V relationship. This was immediately followed by external wash-in (arrow) of 2-deoxyglucose (2-DG; 10 mM; A) or sodium cyanide (CN; 2 mM; C). The raw traces in A and C (insets) indicate that I_Ca was larger prior to (open circle) than after (filled circle) glycolytic inhibition and was not altered by oxidative phosphorylation, respectively. B and D, mean I-V relationships indicating that, on average, 10 min of glycolytic inhibition resulted in a 43 % reduction of I_Ca at +10 mV (n= 11, P < 0.01) and was completely reversed by 30 min wash-out of 2-deoxyglucose (n= 6) (B). On the other hand, 10 min of oxidative inhibition (n= 5) increased I_Ca by 16 % at 0 mV (P < 0.05) and did not alter I_Ca at +10 mV and more positive membrane potentials (D).

The mean I-V relationships shown in Fig. 3B establish that 10 min of glycolytic inhibition resulted in a 43.3 % reduction of I_Ca at +10 mV from -6.2 ± 0.2 to -3.7 ± 0.2 pA pF⁻¹ (n= 11, P < 0.01). In addition, Fig. 3B demonstrates that following a 30 min 2-deoxyglucose wash-out period I_Ca returned to control levels (-6.2 ± 0.2 pA pF⁻¹ for control versus -5.8 ± 0.3 pA pF⁻¹ after wash-out of 2-deoxyglucose; n= 6, n.s.). Similar to FMI, glycolytic inhibition resulted in a reduced mean slope conductance from 187 to 103 pS, which was completely reversed by wash-out of 2-deoxyglucose.

On the other hand, after 10 min of oxidative inhibition I_Ca increased modestly at 0 mV as shown in Fig. 3C. Examination of the raw traces in Fig. 3C (inset) showed that, as in the previous experiments, inactivation was not changed by oxidative inhibition (i.e. t_½= 32.2 ± 1.3 ms in control (open circle) versus 30.5 ± 1.6 ms following oxidative inhibition (closed circle); n= 5, n.s.). Figure 3D shows the mean I-V relationships before and after application of cyanide. Clearly, the effect of cyanide was limited to a modest increase in peak I_Ca between -30 and 0 mV with no difference recorded at more positive voltages. At +10 mV, mean peak I_Ca was not altered by cyanide application (-6.2 ± 0.4 pA pF⁻¹ in control versus -6.4 ± 0.3 pA pF⁻¹ following oxidative inhibition; n= 5, n.s.) while the mean maximum slope conductances were virtually unchanged (i.e. 168 pS in control versus 161 pS after cyanide).

Like glucose, 2-deoxyglucose is phosphorylated to 2-deoxyglucose-6-phosphate (Oikonomakos et al. 1995). Due to the inability to metabolize 2-deoxyglucose-6-phosphate, the addition of 2-deoxyglucose could in principle sequester the pool of inorganic phosphate. This phosphate sequestration by 2-deoxyglucose could indirectly inhibit ATP production by the mitochondria and contribute to the reduction of I_Ca. Therefore, we conducted experiments in which we added 1.2 mM inorganic phosphate (NaH₂PO₄) to the bath solution and 3 mM NaH₂PO₄ to the pipette solution. The mean I-V relationships shown in Fig. 4A demonstrate that, following a 10 min exposure to 2-deoxyglucose in the presence of NaH₂PO₄, I_Ca decreased by 25.0 % (-6.4 ± 0.3 pA pF⁻¹ for control versus -4.8 ± 0.1 pA pF⁻¹ after 2-deoxyglucose; n= 8, P < 0.05) at +10 mV. Unlike previous experiments where steady-state current levels were established with 10 min of exposure to 2-deoxyglucose, the inclusion of NaH₂PO₄ slowed the rate of decline of I_Ca (data not shown), so that steady state was achieved after 15 min with I_Ca being reduced by 38.4 % (-6.2 ± 0.1 pA pF⁻¹ for control versus -3.8 ± 0.1 pA pF⁻¹ for 2-deoxyglucose; n= 4, P < 0.01) as shown in Fig. 4B. This degree of I_Ca reduction at steady state was quantitatively similar to that observed following application of 2-deoxyglucose in the absence of NaH₂PO₄ (compare Fig. 4B with Fig. 3B).

Figure 4. Effect of glycolytic inhibition on Ca²⁺ current with 2-deoxyglucose or iodoacetamide.

A, a 10 min episode of glycolytic inhibition with 2-deoxyglucose (2-DG; 10 mM) in the presence of NaH₂PO₄ (P_i; 1.2 mM in bath and 3.0 mM in pipette) reduced I_Ca by 25.0 % (n= 4, P < 0.05). B, however, a 15 min exposure to 2-deoxyglucose in the presence of NaH₂PO₄ (1.2 mM in bath and 3.0 mM in pipette) resulted in a greater reduction of I_Ca (38.4 %, n= 4, P < 0.01). C, a functional creatine kinase shuttle (20 mM creatine (CR) in pipette) did not prevent reduction of I_Ca. D, when myocytes were exposed for 10 min to iodoacetamide (IAA; 100 μM) a 44.8 % reduction of I_Ca was observed (n= 7, P < 0.01), which was similar to the reduction observed following glycolytic inhibition by 2-deoxyglucose (Fig. 3B). E, reduction of I_Ca by IAA was greatly attenuated by the inclusion of 5 mM MgATP in the pipette solution (IAA-MgATP), as was also observed when 2-deoxyglucose was used to inhibit glycolysis.

The mitochondrial and cytosolic (i.e. myofibrils) creatine kinase catalyses the transfer of high energy phosphate between ATP, ADP, creatine and creatine phosphate, shuttling ATP from the mitochondria to the cytosol. Therefore, it is possible that during glycolytic inhibition under our conditions (i.e. in the absence of creatine) the mitochondria were able to synthesize ATP but this ATP was not able to reach the Ca²⁺ channels. To test for this possibility, we conducted experiments in the presence of 20 mM creatine and 3 mM NaH₂PO₄ in the pipette solution and 1.2 mM NaH₂PO₄ in the bath solution. The mean I-V relationships shown in Fig. 4C indicate that I_Ca at +10 mV decreased by 41.8 % from -6.7 ± 0.3 pA pF⁻¹ for control to -3.9 ± 0.6 pA pF⁻¹ (n= 5, P < 0.01) following 10 min of glycolytic inhibition with creatine and inorganic phosphate. This result was similar to that observed in the absence of creatine and inorganic phosphate, establishing that the mitochondria are unable to sustain I_Ca when glycolysis is inhibited.

Given the significant reduction of I_Ca following application of 2-deoxyglucose, we sought to use another inhibitor of glycolysis to confirm the role of glycolytic derived ATP in modulating I_Ca. The mean I-V relationships in Fig. 4D demonstrate that the application of 100 μM IAA for 10 min caused I_Ca to decrease by 44.8 % (-6.7 ± 0.3 pA pF⁻¹ for control versus -3.7 ± 0.2 pA pF⁻¹ after application of IAA; n= 7, P < 0.01) at +10 mV. The similarity in the degree of reduction of I_Ca with 2-deoxyglucose and IAA suggests that the reduction of I_Ca is indeed the result of glycolytic inhibition (compare Fig. 3B with 4D). The effects of IAA on I_Ca were not due to direct effects of IAA on Ca²⁺ channels since, as illustrated in Fig. 4E, the inclusion of 5 mM MgATP in the pipette solution largely prevented the reduction of I_Ca by IAA (12.5 % reduction of I_Ca from -6.4 ± 0.1 pA pF⁻¹ for control to -5.6 ± 0.2 pA pF⁻¹ at +10 mV for IAA-MgATP; n= 6, P < 0.05).

Previous studies have shown that I_Ca is sensitive to pH (Chen et al. 1996; Schuhmann et al. 1997) and that changes in intracellular pH can occur following the application of 2-deoxyglucose and cyanide (Allue et al. 1996). Therefore, to determine whether the reduction of I_Ca observed following metabolic inhibition was due to changes in pH, we conducted FMI experiments with increased proton buffering conditions. Figure 5A shows that, after a 10 min FMI period in the presence of 50 mM Hepes in the bath and pipette solutions, I_Ca decreased by almost twofold, similar to the reduction observed after FMI with only 10 mM Hepes (see Fig. 1C and D). The mean I-V relationships shown in Fig. 5B demonstrate that following a 10 min period of FMI in the presence of 50 mM Hepes I_Ca at +10 mV decreased by 47.6 % (-6.2 ± 0.1 pA pF⁻¹ for control versus -3.2 ± 0.2 pA pF⁻¹ after FMI; n= 6, P < 0.01). Since HCO₃⁻ is a more physiological buffer than Hepes and since Allue et al. (1996) successfully clamped pH during metabolic inhibition in isolated myocytes with HCO₃⁻, we conducted a set of experiments with 50 mM Hepes plus 25 mM NaHCO₃ in the bath solution. The NaHCO₃ solution was continuously bubbled with a mixture of 95 % O₂ and 5 % CO₂ (pH 7.4). The representative myocyte shown in Fig. 5C illustrates that 10 min of FMI with high Hepes and HCO₃⁻ still decreased I_Ca. The mean I-V relationships in Fig. 5D show that 10 min of FMI reduced I_Ca at +10 mV on average by 38.6 %, from -6.4 ± 0.1 pA pF⁻¹ for control to -3.9 ± 0.2 pA pF⁻¹ after FMI in the presence of high Hepes and HCO₃⁻ (n= 4, P < 0.01). This result was also similar to the reduction of I_Ca observed during FMI with 10 mM Hepes (Fig. 1D).

Figure 5. Effect of full metabolic inhibition on Ca²⁺ current in the presence of high Hepes and/or NaHCO₃.

To determine whether the reduction of I_Ca after FMI is related to alterations in pH_i, proton buffering conditions were improved; 50 mM Hepes was present in the bath and pipette solutions in A-D, plus 25 mM NaHCO₃ in the bath solution in C and D. A and C, in the whole-cell configuration, I_Ca was initially sampled for 5 min before the indicated interruption to record a control I-V relationship. This was followed with external wash-in (arrow) of 2-deoxyglucose (10 mM) and sodium cyanide (2 mM) (FMI). In A, it is evident that 50 mM Hepes in the bath and pipette solution did not prevent the reduction of I_Ca upon wash-in of metabolic inhibitors. C shows that 50 mM Hepes in the bath and pipette and 25 mM NaHCO₃ in the bath did not prevent the reduction of I_Ca upon wash-in of metabolic inhibitors. B and D, mean I-V relationships showing that, on average, metabolic inhibition with 50 mM Hepes only resulted in a 47.6 % reduction of I_Ca (B; n= 6, P < 0.01) compared with a 38.6 % reduction with 50 mM Hepes and 25 mM NaHCO₃ (D; n= 4, P < 0.01).

Full metabolic inhibition in the presence of exogenous ATP or ATP analogue

In order to test whether the effects of full metabolic or glycolytic inhibition on I_Ca resulted from depletion of intracellular ATP, as opposed to other effects of metabolic inhibition, we repeated the full metabolic and glycolytic inhibition experiments but included either ATP or AMP-PCP (a non-hydrolysable analogue of ATP) in the pipette solution. Figure 6A shows typical results when the pipette solution contained MgATP (i.e. 5 mM ATP together with 6.2 mM total MgCl₂). After superfusion with a bath solution containing 2-deoxyglucose and sodium cyanide (arrow) for 10 min, I_Ca was reduced by 16 % (see also Fig. 6A, inset) without change in the time course of current inactivation (i.e. t_½= 33.0 ± 1.3 ms in control versus 31.4 ± 1.4 ms following FMI; n= 4, n.s.). The mean I-V relationships in Fig. 6B show that I_Ca decreased by 18.4 % (-6.8 ± 0.6 versus -5.5 ± 0.3 pA pF⁻¹, n= 5, P < 0.05) at +10 mV, compared with a 40 % reduction in the absence of ATP in the pipette (Fig. 1D). Associated with this modest reduction of I_Ca with ATP, the mean maximum slope conductance was also reduced by 15 % from 163 to 142 pS.

Figure 6. Effect of full metabolic inhibition on L-type calcium current in the presence of MgATP or MgAMP-PCP.

MgATP (5 mM) or MgAMP-PCP (5 mM) was included in the pipette solution in A and B, and C and D, respectively. A and C, in the whole-cell configuration, I_Ca was initially sampled for 5 min before the indicated interruption in order to record a control I-V relationship. This was immediately followed by external wash-in (arrow) of 2-deoxyglucose (10 mM) and sodium cyanide (2 mM). The raw traces (insets) demonstrate that in the presence of MgATP (A) I_Ca prior to (open circle) was larger than I_Ca after (filled circle) FMI. However, when MgAMP-PCP was included in the pipette (C) FMI did not alter I_Ca. B and D, mean I-V relationships which show that, on average, 10 min of FMI resulted in a moderate but significant reduction of I_Ca at +10 mV with MgATP (B; n= 5, P < 0.05) and was unaltered with MgAMP-PCP (D; n= 5, n.s.).

Intracellular ATP regulates Ca²⁺ channel activity by phosphorylation-dependent (Hilgemann, 1997) and phosphorylation-independent (O'Rourke et al. 1992; Yazawa et al. 1997; Yokoshiki et al. 1997) pathways. In order to distinguish between these regulatory pathways, experiments were also performed with the non-hydrolysable ATP analogue, AMP-PCP, in the pipette solution. Figure 6C shows that 5 mM MgAMP-PCP (i.e. 5 mM AMP-PCP plus 6.2 mM total MgCl₂) in the patch pipette completely prevented the reduction of I_Ca associated with FMI. With MgAMP-PCP in the pipette, the mean I-V relationships illustrated in Fig. 6D were unaffected by FMI The peak I_Ca at +10 mV was not significantly altered by FMI (-6.9 ± 0.3 pA pF⁻¹ in control versus -6.5 ± 0.3 pA pF⁻¹ following FMI; n= 5, n.s.). Similar to MgATP, inclusion of MgAMP-PCP in the pipette had little effect on the maximum slope conductance (194 pS in control versus 191 pS following FMI; n= 5) and rate of current inactivation (t_½= 43.7 ± 4.5 ms in control versus 41.4 ± 4.1 ms following FMI; n= 5, n.s.).

DISCUSSION

The present study shows that a 10 min period of FMI with 2-deoxyglucose and sodium cyanide reduced peak I_Ca at +10 mV by 40 %, while inhibition of glycolysis alone resulted in a comparable 43 % decrease in peak I_Ca. On the other hand, inhibition of oxidative phosphorylation alone using cyanide did not have a significant effect on peak I_Ca, suggesting that glycolytic derived ATP preferentially regulates I_Ca. This reduction of I_Ca by 2-deoxyglucose did not result from non-specific sequestration of inorganic phosphate since inclusion of NaH₂PO₄ could not prevent the inhibitory effects of this non-hydrolysable glucose analogue. This result is consistent with a previous finding in isolated rat hearts where 2-deoxyglucose reduced ATP levels without affecting inorganic phosphate concentration (Kupriyanov et al. 1991). The rate of I_Ca decline was altered, however, when NaH₂PO₄ was included suggesting that the levels of phosphate may influence the kinetics of I_Ca inhibition.

To further establish the role of glycolytic derived ATP in the regulation of I_Ca, we also blocked glycolysis with 100 μM IAA, which at this low concentration is a relatively specific inhibitor of glyceraldehyde-3-phosphate dehydrogenase (Sabri & Ochs, 1971). However, unlike 2-deoxyglucose, IAA should not create a sink for inorganic phosphate. Nevertheless, IAA significantly reduced I_Ca at +10 mV, similar to 2-deoxyglucose. Taken together these data suggest that substrate metabolism by the mitochondria is unable to maintain I_Ca. However, the mitochondrial creatine kinase is required to shuttle energy from the mitochondria to the cytosol (Aliev & Saks, 1997). Therefore, it is possible that, when creatine kinase is functional, the mitochondria could prevent I_Ca reduction during glycolytic inhibition. However, even in the presence of creatine and phosphate the application of 2-deoxyglucose still caused a significant decrease in I_Ca. These results establish that glycolysis and not oxidative phosphorylation is involved in the ATP-dependent reduction of I_Ca. The above results are consistent with preferential regulation of cardiac I_Ca by glycolysis as previously observed in rat portal vein smooth muscle cells (Lorenz & Paul, 1997). Our results are also consistent with the lack of effect of 1 mM cyanide on I_Ca measured in feline cardiomyocytes (Furukawa et al. 1994). Preferential regulation of cardiac L-type Ca²⁺ channels by glycolytic ATP is also expected from the sensitivity of other cardiac membrane proteins to glycolytic inhibition (Weiss & Lamp, 1987, 1989; Xu et al. 1995).

Since biphasic changes in intracellular pH occur following metabolic inhibition with cyanide and 2-deoxyglucose and since protons modulate I_Ca, we tested whether increasing intracellular pH buffering capacity with the addition of 50 mM Hepes and 25 mM NaHCO₃ would affect the changes in I_Ca observed with metabolic inhibition. Increasing the pH buffering capacity with high levels of HCO₃⁻, previously shown to prevent changes in intracellular pH (Allue et al. 1996), had no effect on the magnitude or rate of I_Ca reduction following metabolic inhibition. Thus, the changes in I_Ca are unlikely to be mediated by changes in intracellular pH.

Prevention of I_Ca reduction following metabolic inhibition by MgATP or MgAMP-PCP in the pipette solution (Fig. 6B and D; cf. Fig. 1D) suggests that metabolic regulation by glycolysis involves direct allosteric phosphorylation-independent effects of ATP on L-type Ca²⁺ channels. These observations are consistent with previous studies in guinea-pig myocytes (O'Rourke et al. 1992; Yazawa et al. 1997) and vascular smooth muscle cells (Yokoshiki et al. 1997) where it was shown that Mg²⁺ must be complexed with ATP or its analogues (AMP-PCP or AMP-PNP (adenylyl imidophosphate)) to modulate I_Ca. We found that MgAMP-PCP completely prevented the reduction of I_Ca associated with FMI.

The more potent effect of MgAMP-PCP compared with MgATP (compare Fig. 6B with Fig. 6D) in preventing I_Ca reduction during FMI might be related to local ATP concentrations. With ATP in the pipette, local ATP concentrations could be reduced by cellular ATPases, which utilize ATP at a rate comparable to the rate of ATP diffusion into the cell from the pipette. This local consumption would not be expected for non-hydrolysable analogues of ATP, like AMP-PCP. In addition, local cellular ATPases may enhance the intracellular ADP concentration locally and thereby compete with ATP for the allosteric binding site, as has been reported for the ATP-sensitive K⁺ channel (Terzic et al. 1995) and CFTR (cystic fibrosis transmembrane conductance regulator) Cl⁻ channel (Anderson & Welsh, 1992; Gadsby et al. 1995).

In this study the effect of FMI on intracellular ATP concentration was not measured. However, depletion of the global intracellular ATP concentration to below 20 μM is associated with rigor cross-bridge formation and square-shaped cell morphology (Fabiato & Fabiato, 1975; Allue et al. 1996). In our studies, when ATP was not included in the pipette, FMI and inhibition of oxidative phosphorylation consistently caused cells to shorten by 50-60 % and become square shaped within 3-5 min of exposure to the inhibitors, suggesting that the intracellular ATP concentration was reduced to less than 20 μM (Allue et al. 1996). However, inhibition of glycolysis and inclusion of MgATP or MgAMP-PCP during FMI was not associated with cell shortening, suggesting that a global reduction of ATP to less than 20 μM did not occur and was not required for I_Ca reduction. These results are expected since contractile proteins have been previously shown to be preferentially supported by oxidative phosphorylation (Weiss & Lamp, 1987, 1989).

It is conceivable that the reduction of I_Ca during FMI is related to the substantial changes in myocyte shape that occur as a result of rigor cross-bridge development. For instance, cell shrinkage as a result of hyperosmotic perfusion has been shown to decrease, while cell swelling has been shown to increase, peak Ba²⁺ currents through Ca²⁺ channels in smooth muscle (Xu et al. 1996). Similarly, cell shrinkage reduces the amplitude of I_Ca and slows its rate of inactivation in guinea-pig ventricular myocytes, secondary to rapid elevation of intracellular Ca²⁺ (Ogura et al. 1997). On the other hand, it has also been demonstrated in guinea-pig cardiomyocytes that I_Ca is not altered by 20 % stretch or hyposmotic stress (Sasaki et al. 1992) in rat cardiomyocytes (Hongo et al. 1996). In our study it is unlikely that the reduction of I_Ca is due to cell shortening since the reductions of I_Ca were observed after glycolytic inhibition (with 2-deoxyglucose or IAA), where cell shortening was not observed. Furthermore, allosteric regulation by MgATP has been shown to occur in the presence of 5 mM intracellular ATP, where rigor and cell shape changes are prevented (O'Rourke et al. 1992; Yokoshiki et al. 1997).

In summary, we have shown that cardiac L-type I_Ca is preferentially regulated by MgATP derived from glycolysis as opposed to oxidative phosphorylation and that regulation by ATP does not require hydrolysis. This preferential utilization of glycolytic ATP by L-type Ca²⁺ channels could reflect the observation that glycolytic enzymes are colocalized with skeletal Ca²⁺ channels (Brandt et al. 1990), similar to A-kinase anchoring proteins (Goa et al. 1997). This regulatory mechanism may be extremely important under both physiological and pathophysiological conditions. Under physiological conditions this phosphorylation-independent mechanism may operate separate from the phosphorylation-dependent mechanisms. It has been demonstrated that the K_d for 50 % potentiation of L-type Ca²⁺ channel activity by internal perfusion of 5 mM ATP is 0.53 mM (Yokoshiki et al. 1997), which is much higher than the K_d (i.e. 15 μM) for phosphorylation (Francis & Corbin, 1994; Hilgemann, 1997). Therefore, this phosphorylation-independent pathway represents an important regulatory control mechanism for I_Ca activity operating at relatively high levels of intracellular ATP. This regulatory mechanism of I_Ca would be expected to work in concert with a shortening of the action potential duration to reduce excitability and contractility, thereby conserving energy during metabolic stress.