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(Circulation Research. 1999;85:e7-e16.)
© 1999 American Heart Association, Inc.

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Ca²⁺ Influx Through Ca²⁺ Channels in Rabbit Ventricular Myocytes During Action Potential Clamp

Influence of Temperature

José L. Puglisi, Weilong Yuan, José W. M. Bassani, Donald M. Bers

From the Department of Physiology (J.L.P., W.Y., D.M.B.), Loyola University Chicago, Maywood, Ill; Departamento de Engenharia Biomédica (J.L.P., J.W.M.B.), Universidade Estadual de Campinas, UNICAMP, Brazil.

Correspondence to Donald M. Bers, PhD, Department of Physiology, Loyola University Medical School, 2160 South First Ave, Maywood, IL 60153. E-mail dbers{at}luc.edu

	Abstract

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Abstract—Ca²⁺ influx via Ca²⁺ current (I_Ca) during the action potential (AP) was determined at 25°C and 35°C in isolated rabbit ventricular myocytes using AP clamp. Contaminating currents through Na⁺ and K⁺ channels were eliminated by using Na⁺- and K⁺-free solutions, respectively. DIDS (0.2 mmol/L) was used to block Ca²⁺-activated chloride current (I_Cl(Ca)). When the sarcoplasmic reticulum (SR) was depleted of Ca²⁺ by preexposure to 10 mmol/L caffeine, total Ca²⁺ entry via I_Ca during the AP was 12 µmol/L cytosol (at both 25°C and 35°C). Similar Ca²⁺ influx at 35°C and 25°C resulted from a combination of higher and faster peak I_Ca, offset by more rapid I_Ca inactivation at 35°C. During repeated AP clamps, the SR gradually fills with Ca²⁺, and consequent SR Ca²⁺ release accelerates I_Ca inactivation during the AP. During APs and contractions in steady state, total Ca²⁺ influx via I_Ca was reduced by 50% but was again unaltered by temperature (5.6±0.2 µmol/L cytosol at 25°C, 6.0±0.2 µmol/L cytosol at 35°C). Thus, SR Ca²⁺ release is responsible for sufficient I_Ca inactivation to cut total Ca²⁺ influx in half. However, because of the kinetic differences in I_Ca, the amount of Ca²⁺ influx during the first 10 ms, which presumably triggers SR Ca²⁺ release, is much greater at 35°C. I_Ca during a first pulse, given just after the SR was emptied with caffeine, was subtracted from I_Ca during each of 9 subsequent pulses, which loaded the SR. These difference currents reflect I_Ca inactivation due to SR Ca²⁺ release and thus indicate the time course of local [Ca²⁺] in the subsarcolemmal space near Ca²⁺ channels produced by SR Ca²⁺ release (eg, maximal at 20 ms after the AP activation at 35°C). Furthermore, the rate of change of this difference current may reflect the rate of SR Ca²⁺ release as sensed by L-type Ca²⁺ channels. These results suggest that peak SR Ca²⁺ release occurs within 2.5 or 5 ms of AP upstroke at 35°C and 25°C, respectively. I_Cl(Ca) might also indicate local [Ca²⁺], and at 35°C in the absence of DIDS (when I_Cl(Ca) is prominent), peak I_Cl(Ca) also occurred at a time comparable to the peak I_Ca difference current. We conclude that SR Ca²⁺ release decreases the Ca²⁺ influx during the AP by 50% (at both 25°C and 35°C) and that changes in I_Ca (and I_Cl(Ca)), which depend on SR Ca²⁺ release, provide information about local subsarcolemmal [Ca²⁺]. The full text of this article is available at http://www.circresaha.org.

Key Words: Ca²⁺ current • cardiac muscle • excitation-contraction coupling • sarcoplasmic reticulum Ca²⁺ release

	Introduction

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In mammalian cardiac muscle, Ca²⁺ influx during the action potential (AP) triggers the release of additional Ca²⁺ from the sarcoplasmic reticulum (SR) via Ca²⁺-induced Ca²⁺ release¹ and results in muscle contraction. To achieve [Ca²⁺]_i balance and a steady-state level of contraction, Ca²⁺ influx must equal Ca²⁺ efflux during successive cardiac cycles. An accurate measurement of those fluxes is needed to understand excitation-contraction coupling. We previously studied the Ca²⁺ removal fluxes and their relative contributions to relaxation in different species (rabbit, rat, and ferret) at different temperatures.^{2 3 4} We also measured Ca²⁺ influx via Ca²⁺ current (I_Ca) during square voltage-clamp pulses and contractions in rat and rabbit at room temperature.⁵ However, the Ca²⁺ entry during an AP will be different than during a square voltage pulse. In the present study, we focus on quantitative measurements of Ca²⁺ influx via Ca²⁺ channels during the rabbit ventricular AP. These values should be useful in the overall quantitative understanding of Ca²⁺ fluxes in rabbit myocytes.

During an AP, Ca²⁺ can enter the cell through sarcolemmal Ca²⁺ channels (I_Ca) and via Na⁺/Ca²⁺ exchange (I_Na/Ca), but under normal conditions, it is generally accepted that I_Ca is the major source of Ca²⁺ influx.^{6 7} I_Ca has been intensively studied using voltage clamp, mainly with square pulses that allow characterization of channel kinetics and other intrinsic properties.⁸ Using such kinetic parameters, the behavior of I_Ca during an AP has been modeled.^{9 10} However, this modeling has been limited because the Ca²⁺ influx during an AP is a complicated function of not only time and voltage but also local Ca²⁺ at the inner mouth of the channel. The use of the AP clamp technique has provided some unique insight into the time course of I_Ca under more physiological conditions.^{11 12 13} Nevertheless, it is difficult to assess accurately the integrated Ca²⁺ entry via I_Ca during the normal AP because of contaminating ionic currents (Na⁺, K⁺, Cl^-, and Na⁺/Ca²⁺ exchange) and unknown SR Ca²⁺ loading and release. Yuan et al¹⁴ used AP clamp to study isolated I_Ca during the AP waveform in rabbit and rat ventricular myocytes at room temperature. However, in that study [Ca²⁺]_i was heavily buffered, specifically to allow comparative study of I_Ca properties free of the normal influence of local [Ca²⁺]_i. It is clear that Ca²⁺-dependent inactivation of I_Ca is important in affecting the amount of Ca²⁺ entry.^{15 16 17 18} Indeed, during Ca²⁺ release from the SR, [Ca²⁺]_i near the Ca²⁺ channel mouth may be much higher than the bulk [Ca²⁺]_i measured with fluorescence indicators, consistent with theoretical calculations.^{19 20} This high local [Ca²⁺]_i may contribute greatly to I_Ca inactivation during the AP. In the present study, we allowed normal SR Ca²⁺ release to occur and evaluated its effect on I_Ca during the AP. Moreover, we used the effect of SR Ca²⁺ release on I_Ca inactivation to provide indirect information about local [Ca²⁺]_i near the Ca²⁺ channel.

Having normal Ca²⁺ transients occur during AP clamp creates additional complications in isolating I_Ca from potentially contaminating currents. While K⁺ free, Cs solutions can block most K⁺ currents; it is more difficult to perform experiments in Na⁺-free solutions to prevent Na⁺ currents and Na⁺/Ca²⁺ exchange. This is because Na⁺/Ca²⁺ exchange is so important in the steady state in extruding Ca²⁺ from the cell (which entered via I_Ca). We have overcome this problem and measured I_Ca during AP clamp in Na⁺- and K⁺-free solution with 0.2 mmol/L DIDS to block Ca²⁺-activated Cl^- current (I_Cl(Ca)).^{21 22} AP waveforms were first recorded under more physiological ionic conditions in perforated current clamp mode. These AP waveforms were then applied to cells as the command voltage templates, under conditions where currents other than I_Ca were blocked, but Ca²⁺ transients and contractions were relatively normal. Experiments were done at both 25°C and 35°C using appropriate AP waveforms.

We found that in rabbit myocytes the total amount of Ca²⁺ entry did not change significantly between 25°C and 35°C during AP clamp. However, as the SR Ca²⁺ load and SR Ca²⁺ release increased, the amount of Ca²⁺ entry was reduced to 50% of that seen without SR Ca²⁺ release. The quantity of Ca²⁺ that entered in the first 10 ms (which may trigger SR Ca²⁺ release) was larger at 35°C. Kinetic differences in I_Ca inactivation as SR Ca²⁺ release increases were used to infer changes in local [Ca²⁺] near Ca²⁺ channels and indicate peak SR Ca²⁺ release early in the AP.

	Materials and Methods

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New Zealand White rabbits were obtained from Myrtle's Rabbitry (Thompson Station, Tenn) and cared for according to AAALAC guidelines. Ventricular myocytes were isolated as described,² and cell shortening, APs, and ionic currents were measured at 25°C and 35°C (using 200 µg mL^-1 amphotericin B in perforated patch). Steady-state APs were recorded in current-clamp mode in physiological ionic conditions, triggered by depolarizations (0.5 Hz) at both 25°C and 35°C. The bathing normal Tyrode's (NT) contained the following (mmol/L): NaCl 140, KCl 6, CaCl₂ 2, MgCl₂ 1, glucose 10, and HEPES 5, pH adjusted to 7.4 with NaOH. The pipette solution contained the following (mmol/L): KCl 30, K⁺-aspartate 110, NaCl 8, and HEPES 5, pH adjusted to 7.2 with KOH (pipette liquid junction potential=-13 mV was corrected during recording).

I_Ca was measured in cells depleted of Na⁺ (by 30 minutes in 0 Na⁺/0 Ca²⁺ NT) and in Na⁺-free NT (Na⁺ replaced by TEA and K⁺ by Cs, pH 7.4, with TEA-OH). Perforating patch electrodes contained the following (mmol/L): CsSO₄ 80, CsCl 55, MgCl₂ 10, HEPES 10, and EGTA 0.1. The liquid junction potential (-3 mV) was not corrected. DIDS (0.2 mmol/L) was usually added to block I_Cl(Ca) without reducing I_Ca.^{21 23} Initially, standard I-V curves were assessed with square voltage-clamp pulses (holding potential, E_H=-70 mV, 200-ms steps to -20 to +40 mV). Then, AP waveforms recorded in current clamp were used for trains of AP clamps. These were preceded by application of 10 mmol/L caffeine in NT for 10 seconds to empty the SR. This brief exposure to Na⁺-containing solution allowed extrusion of the SR Ca²⁺ load via Na⁺/Ca²⁺ exchange but without appreciable Na⁺ entry (<0.5 mmol/L). AP clamps were given every 30 seconds to avoid Ca²⁺ overload (with the Na⁺/Ca²⁺ exchanger blocked). After 10 AP clamps, twitch contraction amplitude achieved steady state and was similar to the value at 0.5 Hz in field-stimulated intact cells. Protocols were repeated at 25°C and 35°C, and the P/4 method was used for leak subtraction.

Integrated I_Ca (in pC/pF) was converted to total cellular Ca²⁺ entry by multiplying by the cell surface to volume ratio (4.58 pF/pL cell volume²⁴ ) and dividing by Faraday's constant, the valence of Ca²⁺ and the fraction of nonmitochondrial cell volume (0.65 L cytosol/L cell,¹ ), resulting in Ca²⁺ flux expressed in µmol/L cytosol.

Currents were normalized to cell capacitance (C_m). C_m was measured (with ion currents blocked) by a -5-mV voltage step (E_m) from -80 mV, with currents analyzed¹⁴ using C_m=_c/E_m[I₀/(1-(I/I₀))], where I₀ and _c are the peak and time constant of capacitance current relaxation, fit by a single exponential (extrapolated to the time of E_m), and I is the steady-state current after E_m. Series resistance compensation was not used. Cells used had on-cell access resistance <5 M (3.31±0.4 M), C_m=229±18 pF, and membrane resistance, R_m=463±51 M, giving a membrane charging time constant of 0.71±0.05 ms.

Data are presented as mean±SEM. Student's unpaired or paired t test was used to determine statistical significance. Values of P0.05 were considered as significant.

	Results

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AP Measurements
Figure 1 shows examples of APs recorded in perforated whole-cell current-clamp mode in rabbit ventricular myocytes at 25°C and 35°C. As temperature increased, the resting membrane potential (E_m) was slightly hyperpolarized and AP duration and overshoot were reduced (see Table). In 8 experiments, the action potential duration (APD) measured at the level of 50% repolarization (APD₅₀) decreased from 114±13 ms (mean±SEM) to 74±14 ms (P<0.05). These APD₅₀ values indicate a Q₁₀ of 1.52±0.11. This is similar to the Q₁₀ of 1.46 that can be extrapolated from AP duration recordings in isolated rabbit ventricular muscle,²⁵ whereas Kiyosue et al²⁶ reported an AP duration Q₁₀ of 2.5±0.4 for guinea pig ventricular myocytes. The resting E_m decreased from -72±1.6 to -81±1.7 mV (P<0.05). Despite changes in resting E_m, the AP amplitude was not significantly different (131±1.2 mV at 25°C, 127±2.5 mV at 35°C). Such changes in APD and resting E_m were reversed after returning to the initial temperature. The AP waveforms in Figure 1 were used as the command E_m templates for subsequent AP clamp experiments.

View larger version (10K): [in this window] [in a new window]   Figure 1. Steady-state APs recorded in current clamp with normal extracellular solution at 25°C and 35°C. As temperature increases, APs became shorter and slightly hyperpolarized. APD50=114±13 ms at 25°C, 74±14 ms at 35°C, Em resting=-72±1.6 mV at 25°C, -81±1.7 mV at 35°C. For both cases, P<0.05, n=8. These APs were used as command Em for AP clamp experiments.

View this table: [in this window] [in a new window]   Table 1. Different Parameters Measured at 25°C and 35°C

I_Ca Measurements Using Square Pulses
With all K⁺ replaced by Cs (to block K⁺ currents) and Na⁺-free conditions (to block I_Na, Na⁺/K⁺-ATPase, and Na⁺/Ca²⁺ exchange), all ionic current is blocked by 1 µmol/L nifedipine or 1 mmol/L Cd.¹⁴ Figure 2 shows that the currents recorded under our experimental conditions were almost completely blocked by 300 µmol/L Cd at all voltages (for both AP clamp and traditional square pulses), and similar results were found for 2 µmol/L nifedipine. The 95.1±0.5% block of I_Ca by 300 µmol/L Cd is consistent with a K_i of 15 µmol/L, similar to the values (6 to 10 µmol/L) we have measured in concentration-response experiments (not shown). These results indicate that all of the ionic current in Figures 4 through 9 is carried by Ca²⁺ channels. The lack of other ionic currents under our recording conditions ensures that linear leak subtractions (P/4) used in subsequent experiments should be adequate.

View larger version (17K): [in this window] [in a new window]   Figure 2. Currents in AP clamp and square voltage-clamp pulses are blocked by Cd. A, An AP (as in Figure 1) was used as the command voltage before and after addition of 300 µmol/L CdCl2 to the bath solution. B, Same cell, but using a square depolarizing pulse from -80 to 0 mV for 200 ms. Recordings were under ionic conditions intended to isolate ICa (eg, Na+-free, K+-free solutions containing Cs and DIDS, see Materials and Methods). C, Pooled data from 3 cells (as in panel B) showing almost complete block of current by 300 µmol/L Cd.

View larger version (19K): [in this window] [in a new window]   Figure 4. Ca2+ currents after the blockage of DIDS-sensitive current. A, Superimposed ICa traces at 25°C and 35°C. Before 6 ms, ICa is higher at 35°C, but after 6 ms, this is reversed. B, Superimposed normalized ICa traces emphasize kinetic differences (eg, time to peak ICa is shorter at 35°C). C, Time to peak ICa (5.90±0.45 ms at 25°C, 2.90±0.37 ms at 35°C; P<0.05, n=8). D, Exponential time constants of ICa decline based on double-exponential fits (fast=12.2±1.9 ms at 25°C, 4.20±0.80 ms at 35°C and slow=51.0±5.9 ms at 25°C, 20.1±2.8 at 35°C; P<0.05, n=8).

View larger version (20K): [in this window] [in a new window]   Figure 5. Ca2+ currents measured during AP clamp. AP clamp templates for 25°C and 35°C are shown at top, and corresponding ICa measured at steady-state AP are shown below. Currents shown are after P/4 leak subtraction, using hyperpolarizing pulses. The tenth AP pulse after depletion of SR Ca2+ by caffeine exposure represented an apparent steady state. Both traces are from the same cell.

View larger version (25K): [in this window] [in a new window]   Figure 6. ICa integrals during steady-state AP clamp. A, Total Ca2+ influx (5.64±0.23 µmol/L cytosol at 25°C, 6.01±0.22 µmol/L cytosol at 35°C) and amount entering during the first 10 ms of AP (0.79±0.13 µmol/L cytosol at 25°C, 1.89±0.20 µmol/L cytosol at 35°C; P<0.05, n=8). B, Temporal evolution of ICa integral during the AP.

View larger version (28K): [in this window] [in a new window]   Figure 7. Effects of SR Ca2+ load after SR Ca2+ depletion by caffeine pulse. A and B, Twitch contraction amplitude progressively increased as the SR Ca2+ load was replenished. C and D, ICa traces recorded simultaneously with contractions during AP clamp pulses (after P/4 leak subtraction as in Figure 5). Pulse numbers 1, 3, 5, 7, and 10 are shown with progressively greater ICa inactivation as the SR refills. E and F, For each pulse number indicated from panels C and D, the current at pulse 1 (with zero SR Ca2+ load) was subtracted to yield the difference currents. These difference currents show how SR Ca2+ release alters ICa for sequential pulses.

View larger version (17K): [in this window] [in a new window]   Figure 8. Putative SR Ca2+ release flux. Assuming that the traces from Figure 7E and 7F reflect the subsarcolemmal [Ca2+]i as sensed locally by the Ca2+ channels, the time derivatives of those traces would reflect the rate of Ca2+ release from the SR. Peak Ca2+ release occurred at 5.4 ms at 25°C and 2.5 ms at 35°C. The units are not explicitly meaningful with respect to Ca2+ flux.

View larger version (18K): [in this window] [in a new window]   Figure 9. Changes in the amount of Ca2+ influx during APs as the steady state is achieved after a caffeine-induced emptying of SR Ca2+ load. The amount of Ca2+ that enters via Ca2+ channels decreased from 12 µmol/L cytosol (without SR Ca2+ release) to 6 µmol/L cytosol (steady state) at either temperature. Pooled data from experiments as in Figure 7.

Some initial experiments with conventional square voltage-clamp pulses were carried out in the absence of DIDS but with Na⁺- and K⁺-free solutions to prevent Na⁺, K⁺, and Na⁺/Ca²⁺ exchange currents. The I_Ca records often showed a shoulder or hump, especially at 35°C.^{21 27} To test whether this might be due to I_Cl(Ca), we used 0.2 mmol/L DIDS, which blocks I_Cl(Ca) without altering I_Ca (at [DIDS] up to 0.5 mmol/L).^{21 23} At 25°C, DIDS had little effect on I_Ca recorded during steps from -70 to 0 mV (Figure 3A), consistent with our previous observations at room temperature. However, at 35°C, the control I_Ca for the same depolarization had a prominent notch, which was abolished by DIDS (Figure 3B). This DIDS-sensitive current resembles the I_Cl(Ca) previously described in rabbit ventricular myocytes.^{21 22} The DIDS-sensitive outward current was largest at potentials where I_Ca and contraction were also large and that are different from the expected Cl^- equilibrium potential of -37 mV (Figure 3C, top curve). We refer to this current as I_Cl(Ca). Peak outward I_Cl(Ca) at 35°C in Figure 3A occurred 17 ms after the start of depolarization.

View larger version (15K): [in this window] [in a new window]   Figure 3. Currents recorded at 25°C and 35°C in control conditions and after the addition of 0.2 mmol/L DIDS to the bathing solution. A, Currents recorded at 25°C with square pulses from a holding potential of -70 to 0 mV (200-ms duration). B, Same conditions as in panel A but at 35°C. The DIDS-sensitive or difference current is more prominent at 35°C. C, I-V relationship for ICa at both temperatures (in the presence of DIDS) and also for DIDS-sensitive current from one cell.

Figure 3C shows the I-V relationship for I_Ca measured with square voltage-clamp pulses at 25°C and 35°C in the presence of DIDS. Maximum I_Ca was at 10 mV at both temperatures. On the other hand, there were large changes in the I_Ca amplitude and time course. The maximal peak I_Ca at 35°C was 77±19% larger than at 25°C (Q₁₀=1.8±0.2, n=8). Increasing temperature can also speed activation and inactivation.²⁸ Figures 4A and 4B show raw and normalized traces of I_Ca at 25°C and 35°C. The time to peak I_Ca decreased significantly from 5.9±0.5 ms at 25°C to 2.9±0.4 ms at 35°C.

Voltage-clamp limitations at very short times hamper precise measurement of the rising phase of I_Ca. For example, the membrane-charging time constant (0.7 ms) indicates that 3 ms will elapse before achieving 90% voltage control. This will distort the initial I_Ca time course measured, compared with ideal voltage clamp. Thus, the activation kinetics of I_Ca cannot readily be analyzed, and even the times to peak should be taken only as relative values or approximations. Warming from 25°C to 35°C should affect capacitance currents much less than ion channel gating. Thus, the relative changes in peak I_Ca and time to peak I_Ca at 25°C versus 35°C are likely to be meaningful (if imprecise in quantitative terms). These voltage-clamp limitations will have much less impact on I_Ca decline or integrals. For example, even without any correction for capacitance, the integrated I_Ca would then be underestimated by <30% (due mainly to outward capacitative current during depolarization). However, using P/4 to compensate for linear leak and capacitance (as we have) reduces the error by much more than a factor of 10, resulting in an error of much less than 3% in I_Ca integrals.

The time course of I_Ca decline was fit with a double-exponential decay curve, and both fast and slow time constants were 2.5 to 2.9 times longer at 25°C (_fast=12.2±1.9 at 25°C, 4.20±0.80 ms at 35°C; _slow=51.0±5.9 at 25°C, 20.1±2.8 at 35°C. The fraction of I_Ca decline in the fast component was 0.80±0.03 at 25°C, 0.67±0.03 at 35°C, Figure 4D). The slower inactivation at 25°C resulted in an absolute value of I_Ca that was larger than at 35°C at all times longer than 6 ms in the pulse shown in Figure 4A.

I_Ca Measurements Using AP Clamp
Figure 5 shows I_Ca during steady-state AP clamps at 25°C and 35°C. During the AP clamp, I_Ca activated rapidly and then inactivated, but the time course differs from that observed for square pulses. The peak value of I_Ca and the inactivation kinetics are affected by temperature in a manner qualitatively similar to results obtained with square pulses. At 25°C, peak I_Ca is smaller, but there is a striking sustained component of I_Ca during the AP plateau phase. This sustained component is less prominent at 35°C and may reflect a balance between the gradually increasing driving force for I_Ca as repolarization proceeds and the progression of channel inactivation. The larger peak and more pronounced inactivation at 35°C make the Ca²⁺ influx through the Ca²⁺ channels more concentrated in the early phase of the AP at this temperature.

The I_Ca records were integrated to quantify the amount of Ca²⁺ entering through Ca²⁺ channels. Figure 6A shows the amount that enters during a steady-state twitch over the whole AP and also during the first 10 ms. Temperature did not significantly change the total Ca²⁺ influx via I_Ca during the steady state, temperature-appropriate AP (5.64±0.23 µmol/L cytosol at 25°C, 6.01±0.22 µmol/L cytosol at 35°C). This fact rules out increased total Ca²⁺ influx as an explanation for hypothermic inotropy, in which contractile force increases several fold upon cooling from 35°C to 25°C.²⁵ Cooling did affect channel kinetics, and the smaller peak current at 25°C was balanced by longer duration such that the integrated flux was unaltered. If we consider only the Ca²⁺ that entered during the first 10 ms (which could serve as the trigger for SR Ca²⁺ release), there was a significant increase at higher temperatures (0.79±0.13 µmol/L cytosol at 25°C versus 1.89±0.20 µmol/L cytosol at 35°C; P<0.05, n=8). At 35°C, almost one third of the Ca²⁺ entered the cell during the first 10 ms. Figure 6B shows the temporal evolution of the I_Ca integral at both temperatures. Although the final values were similar, the integral rose faster at 35°C.

SR Ca²⁺ Release Induces I_Ca Inactivation
Successive AP clamp I_Ca records, initiated immediately after depletion of SR Ca²⁺ by a caffeine pulse, allowed us to monitor the effect of SR reloading on I_Ca. Figures 7A and 7B show consecutive contractions during this SR Ca²⁺ refilling process at 25°C and 35°C. As the SR is refilled, the amplitude (and rate of rise) of contractions increased progressively to reach 10% of resting cell length at 25°C. This is consistent with an increasing amount of SR Ca²⁺ release as the SR refilled toward a level that is normal for steady-state twitch conditions. The natural AP waveform may be expected to change as the SR refills. However, we chose to use the same AP clamp waveform at each sequential pulse because this allows us to infer changes in I_Ca from pulse to pulse, which are Ca²⁺ dependent rather than E_m dependent, as steady state is attained.

Figures 7C and 7D illustrate the corresponding simultaneously recorded AP clamp I_Ca records. The peak I_Ca did not change appreciably during successive AP clamps, but I_Ca traces showed progressively more inactivation. Because the voltage waveform was the same for each pulse and the peak I_Ca was not different, the greater I_Ca decline is unlikely to be due to inactivation that is either voltage dependent or dependent on Ca²⁺ entering via I_Ca. On the other hand, the progressive increase in SR Ca²⁺ release may be expected to contribute to progressively stronger Ca²⁺-dependent inactivation of I_Ca (see also References 16 through 18^{16 17 18} ).

Subtraction of the first I_Ca trace (where there is no SR Ca²⁺ release) from the consecutive traces provides a putative SR Ca²⁺ release–sensitive I_Ca (Figure 7E and 7F). The time course of this difference current may also indicate the time course of local [Ca²⁺]_i change due to SR Ca²⁺ release into the region near the L-type Ca²⁺ channels. If we assume that the peak of those traces corresponds to the maximum inactivation induced by the SR Ca²⁺ release, those peaks should reflect the time of maximum local [Ca²⁺]_i due to Ca²⁺ released from the SR. At 35°C, we infer that maximal local [Ca²⁺]_i occurred within the first 20 ms, whereas at 25°C, the peak does not occur until 75 ms. This 20-ms value inferred for peak local [Ca²⁺]_i near the Ca²⁺ channel at 35°C is also consistent with the time to peak of Ca²⁺-activated Cl^- current in Figure 3B.

We can take this analysis one step further. To the extent that the difference currents in Figure 7E and 7F reflect the SR-dependent change in local [Ca²⁺]_i near the L-type Ca²⁺ channel mouths, their rates of change may reflect the rate of SR Ca²⁺ release as sensed locally by the Ca²⁺ channel. Figure 8 shows the derivatives of the traces from Figure 7E and 7F. We focus on early times where the change is likely to be dominated by SR Ca²⁺ release rather than reuptake or other processes. Peak Ca²⁺ release appears to occur at 5.4 ms at 25°C and 2.5 ms at 35°C. This coincides with the peak of I_Ca during the AP at the two temperatures (6.1±0.38 ms at 25°C, 2.8±0.4 ms at 35°C; P<0.05, n=8). This may indicate that there is little delay between I_Ca activation and SR Ca²⁺ release. It is also notable that the peak of the putative release flux occurred at the same time for pulses with different SR Ca²⁺ loads and released quantities.

Obviously, features of I_Ca inactivation relate inferentially to the rate of SR Ca²⁺ release, unlike quantifiable measurements with optical indicators. The latter phases of the difference current (or its derivative) may be harder to interpret because of possible cumulative effects of inactivation on subsequent I_Ca during a given AP. On the other hand, I_Ca may respond much more rapidly to highly localized changes in [Ca²⁺]_i in a manner that fluorescent indicators cannot. Even with confocal microscopy, Ca²⁺-dependent signals represent averages over diameters on the order of 500 nm, whereas the space between the release channel and L-type Ca²⁺ channel may be 20 times smaller. Thus, these AP clamp experiments may provide unique insight into the process of excitation-contraction coupling.

SR Ca²⁺ Release Reduces Ca²⁺ Influx During AP by 50%
Figure 9 shows the beat-to-beat change in integrated Ca²⁺ influx during the AP as the SR refills after the caffeine-induced depletion. Regardless of the temperature, the SR Ca²⁺ release progressively inhibited Ca²⁺ influx by 50%. At 25°C, the integrated Ca²⁺ influx at the first pulse (without SR Ca²⁺ loading) was 12.5±0.7 µmol/L cytosol. The value at 35°C for pulse one was 12.3±0.7 µmol/L cytosol. These values are not very different from those reported by Yuan et al¹⁴ for rabbit ventricular myocytes under AP clamp at room temperature with EGTA in the pipette. Their reported value was 13.4 µmol/L cell, which corresponds to 20 µmol/L cytosol after accounting for mitochondrial volume, as we have done in the present study.

	Discussion

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Isolation of I_Ca in the Physiological Context
A novel aspect of our study is that we have isolated I_Ca (by using Na⁺- and K⁺-free solutions and DIDS) while simultaneously having the physiological AP waveform and normal Ca²⁺ transients at both 25°C and 35°C. We used very brief exposures to Na⁺-containing solution with caffeine for periodic unloading of the SR. This prevents progressive Ca²⁺ loading and Ca²⁺ overload in Na⁺-free experiments in contracting cardiac myocytes. Of course, blocking Na⁺/Ca²⁺ exchange and Na⁺ current could alter the local [Ca²⁺]_i at the inner mouth of the Ca²⁺ channel (due to either altered induction of SR Ca²⁺ release or local action of Na⁺/Ca²⁺ exchange).^{29 30} This is an intrinsic limitation of our approach. However, it is not possible to simultaneously isolate I_Ca while having normal Na⁺/Ca²⁺ exchange or Na⁺ current. Because the contractions were comparable to normal twitches without voltage clamp, blocking Na⁺/Ca²⁺ exchange may make only a very minor alteration in the local Ca²⁺ transient and I_Ca time course. Indeed, Bassani et al² found that Na⁺-free conditions did not inhibit Ca²⁺ transient amplitude (if SR Ca²⁺ load was constant) and only modestly slowed [Ca²⁺]_i decline in rabbit and rat ventricular myocytes. Thus, although we have isolated I_Ca effectively and used physiological AP waveforms, there might be minor differences in local [Ca²⁺] that we cause by preventing Na⁺/Ca²⁺ exchange, and those might alter I_Ca slightly. Caffeine can have complicating effects such as phosphodiesterase inhibition and increased myofilament Ca²⁺ sensitivity. The myofilament effects are rapidly reversed on caffeine washout, and our brief (<10 seconds) exposures to caffeine does not result in altered I_Ca amplitude (which would be expected if phosphodiesterase inhibition had a basal or lasting effect).

I_Cl(Ca) has been extensively studied.^{21 22 23 31 32} I_Cl(Ca) was much more prominent at 35°C, although it can be increased at room temperature by isoproterenol.²¹ It is not clear why I_Cl(Ca) was so much more pronounced at 35°C, but it could be due to locally higher [Ca²⁺] (due to increased peak I_Ca), higher Ca²⁺ sensitivity at 35°C, or higher intrinsic activity at 35°C versus 25°C. Although our main concern was to block I_Cl(Ca), this current may normally facilitate rapid repolarization when SR Ca²⁺ load and release are high. This could shorten AP duration, limit Ca²⁺ entry via I_Ca, and thus provide negative feedback on cell Ca²⁺ load.

Total Ca²⁺ Influx During AP
Some investigators have tried to assess Ca²⁺ influx via Ca²⁺ channels during the AP by measuring the nifedipine-sensitive current.⁷ Although nifedipine can block I_Ca, by doing so it also prevents SR Ca²⁺ release and the consequent activation of Na⁺/Ca²⁺ exchange current and I_Cl(Ca). Thus, the nifedipine-sensitive current may include currents that are Ca²⁺ activated in addition to I_Ca. Terracciano and MacLeod³³ used a clever approach in guinea pig and rat ventricular myocytes by integrating I_Ca and Na⁺/Ca²⁺ exchange over a full contraction-relaxation cycle, knowing that in the steady state, Ca²⁺ entry must equal Ca²⁺ efflux. In this way, whatever Ca²⁺ entered and left via Na⁺/Ca²⁺ exchange during a cardiac cycle would not show on the integral, whereas each Ca²⁺ ion that entered via I_Ca and left via Na⁺/Ca²⁺ exchange would produce inward movement of 3 charges (2 coming in as I_Ca and 1 entering per Ca²⁺ extruded in exchange for 3 Na⁺). Thus, two thirds of the inward current could be attributed to I_Ca. They found steady-state Ca²⁺ entry via I_Ca of 4 and 14 µmol/L cytosol in rat and guinea pig AP, respectively, at 0.5 Hz and 22°C. However, these authors could not interpret the time course of I_Ca during the AP, because I_Ca and I_Na/Ca would be overlapping in time.

Yuan et al¹⁴ isolated I_Ca using AP clamp in rabbit and rat ventricular myocytes but under conditions where [Ca²⁺]_i transients were inhibited by dialysis with 10 mmol/L EGTA. They found integrated Ca²⁺ entry of 20 and 13 µmol/L cytosol during rabbit and rat AP, respectively, and under similar conditions, Grantham and Cannell⁷ reported 34 µmol/L cytosol during the guinea pig AP (after adjusting their values to the units used here). These values for rabbit and guinea pig are somewhat higher than observed in the present study, but this may be due to lower Ca²⁺-dependent inactivation with EGTA or BAPTA in the cell.

Ca²⁺ can also enter the cell during the cardiac AP via Na⁺/Ca²⁺ exchange, and this would be thermodynamically favored at the very early phase of the AP.⁶ However, no convincing direct measurements of this outward current during an AP have been reported. Under normal physiological conditions, Ca²⁺ entry via this route is likely to be quantitatively small for several reasons. After SR Ca²⁺ release is prevented, Ca²⁺ influx during the AP can still activate a substantial contraction in rabbit ventricle, but if Ca²⁺ channels are also blocked, contractions are abolished.^{6 34} However, if [Na⁺]_i is elevated by blocking the Na⁺/K⁺-ATPase, Ca²⁺ entry via Na⁺/Ca²⁺ exchange during the AP can increase sufficiently to activate large contractions. What probably limits Ca²⁺ entry via Na⁺/Ca²⁺ exchange during the normal AP is the rise in local Ca²⁺ due to both Ca²⁺ entry via Ca²⁺ channels and release from the SR. Thus, such Ca²⁺ entry via Na⁺/Ca²⁺ exchange is probably limited to the first few milliseconds of the AP. This would constrain the integrated Ca²⁺ influx to be probably much less than 1 µmol/L cytosol, and this is consistent with model calculations.³⁵ Grantham and Cannell⁷ reported an upper limit of Ca²⁺ entry via Na⁺/Ca²⁺ exchange to be 30% of the total Ca²⁺ influx during the first 10 ms of the AP, but they overestimated the outward Na⁺/Ca²⁺ exchange current, because I_Ca and Ca²⁺ transients (and early Na⁺/Ca²⁺ exchange current reversal) were prevented by nifedipine. Thus, it is likely that the integrated I_Ca reported in the present study is by far the major Ca²⁺ influx during the AP.

Temperature Alters I_Ca Kinetics but not Total I_Ca Flux
It is well-known that warming accelerates Ca²⁺ channel activation and inactivation and increases peak I_Ca for square voltage-clamp pulses.²⁸ Our results are consistent with these classic observations, even during the more complex AP waveform. However, the higher peak of I_Ca along with the faster inactivation at 35°C resulted in a crossover such that there was more Ca²⁺ influx at 25°C for times longer than 6 ms (Figure 4A). This makes it less obvious how the current integral will change with temperature, especially with changes in AP duration and Ca²⁺ transient. Surprisingly, we found that these changes almost counterbalanced each other, such that the total integrated I_Ca flux during the AP was almost unchanged between 25°C and 35°C. This was true for both steady-state twitches and those when the SR was Ca²⁺ depleted.

During steady-state contraction, Ca²⁺ influx and efflux must be matched. Otherwise, the cell will gradually gain or lose Ca²⁺. If steady-state Ca²⁺ influx during the AP does not change between 25°C and 35°C, the same should be true for Ca²⁺ efflux. Puglisi et al⁴ performed quantitative analysis of Ca²⁺ transport during relaxation at 25°C and 35°C in rabbit ventricular myocytes. Although transport rates and relaxation were much faster at 35°C, the integrated Ca²⁺ extrusion via Na⁺/Ca²⁺ exchange and balance of removal fluxes by Na⁺/Ca²⁺ exchange and SR Ca²⁺ pump were almost the same. Thus, our expectation of flux balance is fulfilled.

Reduction of temperature from 35°C to 25°C normally produces a large increase in developed force (500%), referred to as hypothermic inotropy.²⁵ The present results indicate that increased Ca²⁺ influx during the AP is unlikely to contribute to this effect. This inotropy may be due more to the slowing of Ca²⁺ removal from the cytosol, a prolonged active state, and increased SR Ca²⁺ content.^{4 25}

I_Ca Inactivation Under Physiological Conditions
Our results clearly indicate that the overall integrated Ca²⁺ influx during the AP via I_Ca is reduced by 50% when normal SR Ca²⁺ release occurs. Similar conclusions were drawn by Trafford et al³⁶ based on experiments in ferret ventricular myocytes with square pulses after caffeine-induced SR Ca²⁺ depletion. They found that integrated Ca²⁺ influx declined from 14.8 to 6.7 µmol/L cytosol (after correction for 30% mitochondrial volume). Terracciano and MacLeod³³ also found that steady-state Ca²⁺ influx during AP clamp in guinea pig ventricular myocytes was increased by 39% after blocking SR Ca²⁺ function with thapsigargin. Sipido et al¹⁶ also demonstrated inactivation of Ca²⁺ channels by SR Ca²⁺ release during long square pulses and further showed that the Ca²⁺ channels could recover as [Ca²⁺]_i declined. Sham et al³⁷ and Adachi-Akahane et al¹⁷ also showed that blocking SR Ca²⁺ release slowed I_Ca inactivation (eg, by 67%), and they also emphasized the apparent local nature of this effect at the dyadic junction. Thus, regardless of species, temperature, or depolarization waveform, it appears that SR Ca²⁺ release inhibits 50% of Ca²⁺ entry, which would otherwise occur via I_Ca.

Inactivation of L-type Ca²⁺ channels depends on both voltage and Ca²⁺.¹⁵ In the absence of divalent cations, when Na⁺ is used as the charge carrier for the Ca²⁺ channel, the half-time for current decline can be >500 ms (at voltages that would correspond to the AP plateau).³⁵ This purely voltage-dependent inactivation is so much slower than I_Ca inactivation under physiological conditions that one may infer that almost all physiological I_Ca inactivation is Ca²⁺ dependent. This also suggests that altered driving forces due to local Ca²⁺ accumulation or depletion¹⁹ are unlikely to contribute significantly to the decline of I_Ca during the pulse.¹⁶ In our experiments, at the first postcaffeine AP clamp, when the SR is depleted of Ca²⁺ (Figures 7 and 9), all Ca²⁺-dependent inactivation must be due to Ca²⁺ entry via the channel. We did not measure how much inactivation occurred during this AP. However, by comparing normalized flux for square pulses with Ca²⁺ versus Na⁺ as charge carrier,³⁵ it can be inferred that Ca²⁺ entry is responsible for reducing total influx by 60% for a 160-ms pulse. Given that the steady-state APs here reduced integrated Ca²⁺ influx by 50%, we infer that Ca²⁺ influx and SR Ca²⁺ release contribute about equally to I_Ca inactivation during the AP.

Thus, it is clear that SR Ca²⁺ release can limit Ca²⁺ influx by feedback on the L-type Ca²⁺ channel. This would serve to limit Ca²⁺ entry when the SR is already full. Conversely, it would allow for greater SR Ca²⁺ replenishment when SR Ca²⁺ release is small (eg, Figure 9).

Local [Ca²⁺]_i Sensed by Channels
From our sequential I_Ca traces (Figure 7), it is clear that inactivation induced by SR Ca²⁺ release starts very rapidly. Indeed, the L-type Ca²⁺ channel may sense released Ca²⁺ much before a fluorescent Ca²⁺ indicator that is distributed throughout the bulk cytoplasm. This is an intrinsic advantage of this electrophysiological signal from molecules perfectly positioned to sense the local [Ca²⁺] of interest. Furthermore, the present measurements are done without adding exogenous Ca²⁺ indicator buffers, which could perturb local Ca²⁺ transients. The amount of Ca²⁺-dependent inactivation was used as an indicator of local [Ca²⁺]_i near the L-type Ca²⁺ channel, which may be located very close to the SR Ca²⁺ release channel. The rate of change of this local signal (Figure 8) may then be a local indicator of the rate of SR Ca²⁺ release sensed very near the release channel. Although it is not practical to calibrate these signals with respect to rates of Ca²⁺ flux, they may provide unique insight into the timing of SR Ca²⁺ release. During the rabbit ventricular AP, peak SR Ca²⁺ release appeared to occur at 2.5 and 5 ms after the start of depolarization at 35°C and 25°C, respectively (which coincided with the time of peak I_Ca). This indicates very tight functional coupling between the Ca²⁺ influx and release channels.

Using currents to indicate local [Ca²⁺] changes may also help to further characterize the relative locations of SR Ca²⁺ release channels, L-type Ca²⁺ channels, Na⁺/Ca²⁺ exchangers, and Ca²⁺-activated Cl^- channels. Indeed, [Ca²⁺]_i buffering that is sufficient to prevent released Ca²⁺ from activating Na⁺/Ca²⁺ exchange does not prevent released Ca²⁺ from profoundly affecting I_Ca inactivation.¹⁷ This suggests that the Na⁺/Ca²⁺ exchanger is not localized discretely as close to the SR Ca²⁺ release channel as is the L-type Ca²⁺ channel. Although our main objective with respect to I_Cl(Ca) in the present study was to block it, we did observe that it also sensed high local [Ca²⁺]_i quickly at 35°C. I_Cl(Ca) reached a peak in 20 ms, similar to the I_Ca inactivation. However, we have insufficient data to determine whether this channel senses the same local [Ca²⁺]_i as the Ca²⁺ channel. Differential [Ca²⁺] sensitivities and delays for impact on the currents may also complicate more detailed comparisons of this nature. Nevertheless, it is clear that changes in I_Ca inactivation can be a valuable indicator of local [Ca²⁺]_i at the location that might be most critical for understanding excitation-contraction coupling.

	Acknowledgments

 
This work was supported by a grant from the United States Public Health Service (HL-30077). The authors are indebted to Christina Hovance and Steven Scaglione for technical assistance and to Drs Eileen McCall and Kenneth S. Ginsburg for critical comments on the manuscript.

Received December 16, 1998; accepted August 12, 1999.

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