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(Circulation Research. 1995;76:102-109.)
© 1995 American Heart Association, Inc.

Articles

Inhibition and Rapid Recovery of Ca²⁺ Current During Ca²⁺ Release From Sarcoplasmic Reticulum in Guinea Pig Ventricular Myocytes

Karin R. Sipido, Geert Callewaert, Edward Carmeliet

From the Laboratory of Physiology, University of Leuven (Belgium).

Correspondence to K.R. Sipido, Laboratory of Physiology, KUL, Campus Gasthuisberg, Herestraat 49, B-3000 Leuven, Belgium.

	Abstract

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Abstract We have investigated the modulation of the L-type Ca²⁺ channel by Ca²⁺ released from the sarcoplasmic reticulum (SR) in single guinea pig ventricular myocytes under whole-cell voltage clamp. [Ca²⁺]_i was monitored by fura 2. By use of impermeant monovalent cations in intracellular and extracellular solutions, the current through Na⁺ channels, K⁺ channels, nonspecific cation channels, and the Na⁺-Ca²⁺ exchanger was effectively blocked. By altering the amount of Ca²⁺ loading of the SR, the time course of the Ca²⁺ current (I_Ca) could be studied during various amplitudes of Ca²⁺ release. In the presence of a large Ca²⁺ release, fast inhibition of I_Ca occurred, whereas on relaxation of [Ca²⁺]_i, fast recovery was observed. The time course of this transient inhibition of I_Ca reflected the time course of [Ca²⁺]_i. However, the inhibition seen in the first 50 ms, ie, the time of net Ca²⁺ release from the SR, exceeded the inhibition observed later during the pulse, suggesting the existence of a higher [Ca²⁺] near the channel during this time. Transient inhibition of I_Ca during Ca²⁺ release was observed to a similar degree at all potentials. It could still be observed in the presence of intracellular ATP--S and of cAMP. Therefore, we conclude that the modulation of I_Ca by Ca²⁺ release from the SR is not related to dephosphorylation. It could be related to a reduction in the driving force and to a direct inhibition of the channel by [Ca²⁺]_i. The observation that the degree of inhibition does not depend on membrane potential suggests that the Ca²⁺ binding site for this modulation is located outside the pore. The transient nature of the modulation of I_Ca by Ca²⁺ release will contribute to the recovery of I_Ca during prolonged action potentials.

Key Words: Ca²⁺ channel • heart • Ca²⁺ release • sarcoplasmic reticulum

	Introduction

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It is now well established that Ca²⁺-dependent inactivation is an important determinant of the time course of the Ca²⁺ current (I_Ca) during depolarization.¹ Most of the studies on this Ca²⁺-dependent inactivation have either examined the effects of permeating Ca²⁺ or steady state effects of cytoplasmic Ca²⁺. However, under physiological conditions, I_Ca will trigger Ca²⁺ release from the sarcoplasmic reticulum (SR), resulting in a transient 10-fold increase in [Ca²⁺]_i. This Ca²⁺ release flux is severalfold larger than the Ca²⁺ influx through the channel itself.^{2 3} Because the Ca²⁺ channels are very close to the release channel,^{4 5} one can expect significant modulation of I_Ca by this Ca²⁺ release. This has also been suggested from experiments in which Ca²⁺ release was prevented with ryanodine.^{6 7} In atrial cells dialyzed with citrate, sudden inactivation of I_Ca has been ascribed to Ca²⁺ release.⁸ However, little is known about the quantitative importance of the effect of Ca²⁺ release on I_Ca, its dynamics, or its mechanism. Therefore, in the present study, we will examine this modulation in more detail and attempt to identify the underlying mechanisms.

Recent studies of single Ca²⁺ channel currents suggest that the inhibiting effect of Ca²⁺ on the channel results from a decrease in channel open probability. In cell-attached patches, Ca²⁺ permeation through the channel reduced the probability of subsequent reopening of the channel,^{9 10} and from further analysis, it was suggested that Ca²⁺ shifts the gating mode toward a mode with long-lived closed states.¹¹ In excised patches, with Ba²⁺ as a charge carrier, elevation of cytoplasmic Ca²⁺ in a several micromolar range reversibly reduced the open probability of the Ca²⁺ channels.¹²

Consistent with these single-channel data, Hadley and Lederer¹³ found that a step increase in [Ca²⁺]_i produced by photorelease of caged Ca²⁺ from DM-nitrophen reduced the amplitude of the macroscopic I_Ca. This Ca²⁺-dependent inactivation was no longer seen when phosphorylation of the Ca²⁺ channel was stimulated with isoproterenol; therefore, it was suggested that this inactivation might be related to dephosphorylation.

Besides these observations of inactivation of the Ca²⁺ channel by cytoplasmic Ca²⁺, a number of studies have demonstrated a potentiation of I_Ca by [Ca²⁺]_i, if the elevation is small. Such a potentiating effect of cytoplasmic Ca²⁺ on the channel was originally proposed by Marban and Tsien.¹⁴ Photorelease of caged Ca²⁺ from nitr-5 has been reported to potentiate I_Ca,^{15 16} although a transient inactivation was also observed.¹⁶ Hirano and Hiraoka¹⁷ also observed an increase in channel activity during small increases in [Ca²⁺]_i. Facilitation of I_Ca during repeated depolarizations after a period of rest has also been attributed to an increase in [Ca²⁺]_i.^{18 19 20 21 22} This facilitation was most often noted during experiments in which the cells were perfused with 10 mmol/L EGTA, suggesting that the required increase in [Ca²⁺]_i is small and/or confined to the subsarcolemma. To observe facilitation, the membrane potential between the depolarizing pulses had to be more negative than -50 mV.^{21 22}

In the present study, we investigate the modulation of the macroscopic I_Ca by Ca²⁺ release from the SR. We will examine whether this modulation can be related to changes in phosphorylation and whether it is dependent on membrane potential.

	Materials and Methods

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Cell Isolation
Single ventricular myocytes were isolated enzymatically by a variant of the method described by Mitra and Morad.²³ Guinea pigs were killed by a blow on the neck, the heart was quickly excised, the aorta was cannulated, and the heart mounted on a Langendorff setup for retrograde perfusion. The remaining blood was first washed out with normal Tyrode's solution, followed by 10 minutes of perfusion with a nominally Ca²⁺-free solution. The heart was then perfused with a solution containing 0.7 mg/mL collagenase (Boehringer Mannheim) and 0.1 mg/mL protease (type XIV, Sigma Chemical Co) for 20 minutes. The enzyme solution was then washed out with a low Ca²⁺ solution (0.2 mmol/L). The ventricles were cut off, and the cells were dispersed, resuspended in normal Tyrode's solution, and stored at room temperature.

Voltage Clamp and [Ca²⁺]_i Measurements
The experimental setup was built around an inverted microscope (Nikon Diaphot). The whole-cell variant of the patch-clamp technique was used.²⁴ Pipettes were pulled from borosilicate glass (Jencons Scientific) and had resistances between 1.8 and 2.4 M when filled with 130 mmol/L CsCl. Membrane currents were measured with an Axopatch ID amplifier (Axon Instruments), filtered at 5 kHz, and read into a PC for later analysis. The analog-to-digital converter sampled at 2 kHz, and the data acquisition program also controlled the command potential and various components of the [Ca²⁺]_i measurement system (FASTLAB software, Indec Systems).

The pipette solution contained 50 µmol/L K₅–fura 2 (Molecular Probes). The excitation wavelengths, 360 nm and 380 nm, were selected by narrow bandpass filters mounted on a fast rotor wheel in front of a xenon arc lamp (Cairn Research). This allowed the wavelengths to be alternated at 1 kHz. The excitation light was reflected onto the cell by a dichroic mirror (410 nm, long pass) placed under the objective while the emitted fluorescence passed through the dichroic mirror and was collected by a photomultiplier connected to the side port. A bandpass filter centered at 510 nm selected the maximal emission wavelength. The raw fluorescence signal was filtered at 1600 Hz. The signal of each excitation wavelength was reconstructed by a sample-and-hold circuit (Cairn Research) and read into the PC for later analysis. The background-corrected ratio was calibrated to obtain [Ca²⁺]_i.²⁵ Calibration parameters were obtained from in vivo calibration.²⁶ For the analysis of the time course of the inhibition of I_Ca versus the time course of the [Ca²⁺]_i transient, [Ca²⁺]_i was calculated according to the rate equation to obtain a kinetically corrected [Ca²⁺]_i transient.²

Solutions and Pulse Protocols
The control external solution contained (mmol/L) NaCl 130, CsCl 10, HEPES 10, glucose 10, MgCl₂ 1, and CaCl₂ 1.8, pH 7.40, with NaOH. To isolate I_Ca from other membrane currents, all monovalent cations were substituted with impermeant ones. The pipette solution contained (mmol/L) N-methyl-D-glucamine (NMDG) chloride 110, tetraethylammonium (TEA) chloride 20, HEPES 10, MgCl₂ 0.5, MgATP 4, and K₅–fura 2 0.05, pH 7.20 with TEAOH. The external solution contained (mmol/L) TEA chloride 130 (or NMDG chloride 130), HEPES 10, glucose 10, MgCl₂ 1, and CaCl₂ 1.8 (or 5.4), pH 7.40 with TEAOH. With these external and internal solutions, monovalent cation currents (ie, K⁺, Na⁺, and the nonspecific cation current) and the Na⁺-Ca²⁺ exchange current were effectively blocked. Therefore, the total membrane current almost exclusively represented I_Ca. External solution changes were performed with a fast perfusion system,²⁷ and the removal of Na⁺ was complete in <5 s. The absence of Na⁺ and of Na⁺-Ca²⁺ exchange was confirmed by the following observations: disappearance of Na⁺ current, absence of voltage-dependent relaxation of [Ca²⁺]_i, and absence of voltage-dependent increase of [Ca²⁺]_i at positive potentials.^{2 28}

To study the effect of Ca²⁺ release on the time course of I_Ca, we compared I_Ca for pulses with different amounts of Ca²⁺ release. All other parameters were assumed to be constant, ie, a similar degree of voltage-dependent inactivation and of inactivation by permeating Ca²⁺. This means that the sizes of the depolarizing step and of the peak I_Ca must be comparable. This goal can be achieved during a train of depolarizing pulses at low frequency (0.1 Hz) in Na⁺-free solutions. In the absence of Na⁺-Ca²⁺ exchange, the net efflux of Ca²⁺ from the cell is negligible,^{2 29} and during repetitive depolarizing pulses that activate Ca²⁺ entry through I_Ca, the cell gets loaded with Ca²⁺. Resting [Ca²⁺]_i increases only slightly, suggesting that most of the Ca²⁺ that has entered is sequestered rapidly in the SR. This results in incremental loading of the SR. This approach is illustrated in Fig 1. Before switching to the Na⁺-free solution, cells were held for 3 minutes at -70 mV in a normal Na⁺-containing Tyrode's solution to deplete the SR.²⁸ We then switched to an Na⁺-free solution and set the holding potential at -45 mV to inactivate T-type I_Ca. This step from -70 to -45 mV may have triggered a Ca²⁺ release³⁰ but was not recorded. We then repeatedly depolarized the cell to 0 mV, from a holding potential of -45 mV, with a 10-s interval. The first depolarizing pulse elicited only a very small Ca²⁺ transient (tracing a). Consecutive depolarizing pulses were accompanied by increasing amounts of Ca²⁺ release (tracings b through f). Since these depolarizing pulses were given at low frequency, ie, 0.1 Hz, the peak I_Ca was not affected by rate-dependent modulation.^{31 32} Because of the short duration of the protocol, typically 2 to 3 minutes in Na⁺-free solution, a slow increase of peak I_Ca, during long exposure to Na⁺-free solutions as described by Balke and Wier,³³ also did not interfere with our observations.

View larger version (21K): [in this window] [in a new window]   Figure 1. Pulse protocol to obtain variable amounts of Ca2+ release from the sarcoplasmic reticulum. Cells were held at -70 mV in a normal Na+-containing solution for 3 minutes. The solution was then switched to an Na+-free one, and the holding potential was set at -45 mV. Every 10 s, a depolarizing step to 0 mV (300 ms) was applied. Tracings a through f represent the membrane currents (I, upper tracings) and [Ca2+]i transients (lower tracings) of six consecutive pulses in this Na+-free solution. Every next pulse is accompanied by a larger release as the cell gets loaded with Ca2+ (see text). V indicates membrane voltage.

	Results

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Changes in I_Ca During Ca²⁺ Release From the SR
In Fig 2, a depolarizing pulse with a small amount of Ca²⁺ release (tracings a and a') is compared with a pulse accompanied by a large Ca²⁺ release (tracings b and b'), with either 1.8 mmol/L external CaCl₂ (left panels) or 5.4 mmol/L CaCl₂ (right panels). Each pair of depolarizing steps (a and b, and a' and b') is to the same membrane potential (-20 mV), and for each pair, peak I_Ca values are comparable. Therefore, we assume that the voltage-dependent inactivation and the inactivation by permeating Ca²⁺ are comparable for each pair and that differences in the time course of I_Ca reflect the effect of the different amounts of Ca²⁺ release.

View larger version (24K): [in this window] [in a new window]   Figure 2. Changes in Ca2+ current during Ca2+ release from the sarcoplasmic reticulum. Tracings are shown of membrane currents (I, upper tracings) and [Ca2+]i transients (middle tracings) from the first depolarizing step from -45 to -20 mV after switching to Na+-free solution (labeled a and a') and of a similar step after the cell had been loaded with Ca2+ by 5 to 10 previous steps (labeled b and b'). On the left side of the figure, external CaCl2 was 1.8 mmol/L, and the pulse duration was 2.4 s. On the right, external CaCl2 was 5.4 mmol/L, and the pulse duration was 4 s. The lower tracings are the difference tracings, obtained by subtracting current a from b (tracings with noise of low amplitude, marked by arrow) and subtracting [Ca2+]i transient a from b (higher amplitude noise). These difference tracings were filtered by a five-point averaging method. The dotted line indicates the baseline before the depolarizing step. Tracings from the left and right panels were obtained from two different cells.

During the large [Ca²⁺]_i transient, fast inhibition of I_Ca is seen. However, on relaxation of [Ca²⁺]_i, a rapid recovery is observed. This recovery follows the rapid decline of [Ca²⁺]_i, which occurs mostly within 1 s. This is further illustrated in the lower tracings. The membrane current and [Ca²⁺]_i transient of tracing a were subtracted from those of tracing b, and these difference tracings were scaled. It is clear that the inhibition of I_Ca is transient and that its time course reflects the time course of [Ca²⁺]_i. However, it was not possible to completely superimpose the tracings. We chose to align the tracings during the decline of the Ca²⁺ transient, because gradients are less likely to occur during this phase than during the first 50 ms after depolarization, the period during which Ca²⁺ release from the SR occurs.^{2 3} The inhibition of I_Ca during these first 50 to 100 ms apparently exceeded the inhibition seen later on for similar [Ca²⁺]_i levels, suggesting that the actual [Ca²⁺] at the channel during this period is different from what is measured with fura 2. The peak inhibition of I_Ca occurred at 42±13 ms (mean±SD of five cells, steps to -20 mV). This coincided with the end of the rapid upstroke of the [Ca²⁺]_i transient.

The transient inhibition of I_Ca could be clearly observed at the potentials commonly seen during the action potential plateau, between -20 and +20 mV (Fig 3). At potentials positive to +40 mV, I_Ca became small, and the effect of Ca²⁺ release was more difficult to distinguish.

View larger version (12K): [in this window] [in a new window]   Figure 3. Inhibition and recovery of Ca2+ current during Ca2+ release observed at plateau potentials. Membrane currents (I, upper tracings) and [Ca2+]i transients (lower tracings) during pulses to the indicated potentials are shown. Each pulse was preceded by five loading pulses (external CaCl2, 5.4 mmol/L). All tracings were obtained in the same cell.

When looking at the tracings of Figs 2 and 3, one could interpret the data differently, by saying that the larger [Ca²⁺]_i transient induces a transient outward current shift. Therefore, we have to carefully examine the hypothesis that the transient current shift reflects a [Ca²⁺]_i-activated membrane current not related to I_Ca. Because of our solutions, this current could not be carried by monovalent cations, but it could be due to a [Ca²⁺]_i-activated Cl^- current. Such a current is as yet unknown in the guinea pig heart but has been described in rabbit cardiac cells.^{34 35 36} However, its relation to the [Ca²⁺]_i transient is very different from what is observed here, because the [Ca²⁺]_i-activated Cl^- current declines before the peak of [Ca²⁺]_i.³⁶ Also, in the voltage range from -20 to +20 mV, the transient current shift was always outward, although the Cl^- reversal potential was -2 mV. Fig 4A further illustrates that if external Cl^- is reduced from 142 to 32 mmol/L, thereby shifting the Cl^- reversal potential from -2 to +38 mV, the apparent current shift during Ca²⁺ release is still outward. These observations virtually exclude the possibility that this current shift represents a Cl^- current. Another independent argument was provided by experiments in which we reduced the external [Ca²⁺] after loading the SR in the presence of high external Ca²⁺. This procedure resulted in a substantial decrease of the amplitude of I_Ca, which was nevertheless capable of evoking a Ca²⁺ release of comparable amplitude because of the previous loading (Fig 4B). If an outward current, independent of I_Ca, is generated by [Ca²⁺]_i, the net membrane current should shift outwardly, since the inward I_Ca is drastically reduced. Despite a comparable [Ca²⁺]_i transient in tracings a and b, the current of pulse b is not more outwardly shifted than the one of pulse a, confirming that the outward current shift is indeed part of I_Ca.

View larger version (16K): [in this window] [in a new window]   Figure 4. The observed change in membrane current reflects a change in Ca2+ current (ICa). A, Membrane current (I, upper tracing) and [Ca2+]i transient (lower tracing) of a pulse from -45 to 0 mV, with Cl- reversal potential set at +38 mV (external [Cl-], 32 instead of 142 mmol/L; replaced with aspartate). During Ca2+ release, an apparent transient outward current shift is still present. B, Reduction of the inward ICa by decreasing external CaCl2 from 5.4 mmol/L (tracings marked b) to 0.2 mmol/L (tracings marked a). Before each tracing was recorded, the cell was loaded with Ca2+ by 12 pulses to +10 mV in the presence of 5.4 mmol/L CaCl2; therefore, the smaller ICa still elicits a sizable [Ca2+]i release. Despite the reduction in inward current, the current is not shifted more outwardly during Ca2+ release.

Two other types of control experiments were designed to exclude the possibility that the observed change in I_Ca was not related to factors other than the change in Ca²⁺ release from the SR. In the first set of experiments, we determined that the same pulse protocol did not affect the time course of I_Ca when performed in normal Tyrode's solution, thus excluding frequency-dependent effects (n=4, results not shown). As expected at this low frequency, the [Ca²⁺]_i transients did not change, and neither did I_Ca. In a second set of experiments, we again performed the same protocol but in Na⁺-free solution and with 1 mmol/L EGTA in the pipette. Three to 4 minutes after establishing the whole-cell patch, [Ca²⁺]_i transients could no longer be evoked, and the pulse protocol in Na⁺-free solution no longer affected the time course of I_Ca (n=3, results not shown).

Is the Modulation of I_Ca by Ca²⁺ Release Related to Dephosphorylation?
To examine the possibility that the rapid inhibition and recovery of I_Ca is related to dephosphorylation and rephosphorylation of the channel, we substituted the ATP of the pipette solution with ATP--S. This stable ATP analogue is capable of donating its thiophosphate group for phosphorylation, but the phosphorylated substrate becomes resistant to dephosphorylation.^{37 38} We also added 100 µmol/L cAMP to the pipette solution to promote phosphorylation (Fig 5).

View larger version (17K): [in this window] [in a new window]   Figure 5. Inhibition and recovery of Ca2+ current during Ca2+ release when dephosphorylation is prevented by substitution of ATP in the pipette solution with ATP--S (4 mmol/L). cAMP (0.1 mmol/L) was also included in the pipette solution to promote phosphorylation. External CaCl2 was 1.8 mmol/L. Membrane currents (I, upper tracings) and [Ca2+]i transients (lower tracings) of the first pulse after switching to the Na+-free solution are shown (depolarization, from -45 mV to the indicated potential).

This substitution of ATP with ATP--S did not inhibit the Ca²⁺ pump of the SR, and contractions could be observed throughout the experiment. Several well-known effects of phosphorylation were observed: an increase of the peak amplitude of I_Ca and the appearance of a time-independent background current compatible with the cAMP-dependent Cl^- current. With this pipette solution, it proved impossible to deplete the SR by resting the preparation, and even the first pulse in Na⁺-free solution was already accompanied by a sizable [Ca²⁺]_i transient. Further loading of the cell often resulted in spontaneous oscillations, making an analysis with current and [Ca²⁺]_i subtractions, as in Fig 2, very difficult. However, even without this subtraction, it is quite clear from the tracings in Fig 5 that a transient inhibition of I_Ca during Ca²⁺ release from the SR also occurred in the presence of ATP--S and cAMP. Similar results were obtained in six other cells. Therefore, it seems very unlikely that the rapid modulation of I_Ca during a depolarizing pulse is related to (de)phosphorylation.

The above conclusion relies on the validity of the assumption that in our experimental conditions phosphorylation was maximal and irreversible. These assumptions were tested in separate experiments. The extent of phosphorylation was examined by applying 200 nmol/L of isoproterenol, with a pipette solution similar to that used in the above experiments, ie, with ATP--S and cAMP. Both I_Ca and the Cl^- current remained stable during this application, indicating that phosphorylation was maximal (n=5). In another set of experiments, we examined the reversibility of the phosphorylation. For this purpose, we used a pipette solution with ATP--S but without cAMP. Application of isoproterenol now induced a large increase in the amplitude of I_Ca and in the amplitude of the holding current at -50 mV, compatible with the induction of a Cl^- current. On washout of isoproterenol, the increase of I_Ca proved irreversible, compatible with the assumption that the Ca²⁺ channel had been irreversibly phosphorylated (n=8). These results are in accordance with the previously published study by Hescheler et al,³⁸ who also observed irreversible phosphorylation with ATP--S in the pipette during whole-cell recording. Similar to the maintained increase in I_Ca, the Cl^- current did not disappear on washout of isoproterenol. Although in the presence of ATP--S the changes in I_Ca and in the Cl^- current induced by isoproterenol were irreversible, such changes were completely reversible with a normal pipette solution, ie, one with normal ATP (n=5). These last experiments further support the assumption that with ATP--S in the pipette, the Ca²⁺ channel was indeed irreversibly phosphorylated.

Is the Modulation of I_Ca by Ca²⁺ Release Voltage Dependent?
The fast modulation of I_Ca could be related to a rapid block and unblock of the channel by cytoplasmic Ca²⁺. Such a mechanism may make the modulation voltage dependent if Ca²⁺ has to move to a blocking site within the pore. Blocking of a channel by a cation from the cytoplasmic site is expected to be more pronounced at the more positive potentials.³⁹ On the other hand, if the rapid modulation is related to binding to a blocking site outside the pore, voltage dependence is not expected. Therefore, we examined whether the degree of inhibition and recovery was different for different membrane potentials.

For this purpose, we recorded 8 to 12 consecutive pulses to the same test potential after switching to Na⁺-free solution, and this protocol was repeated for various membrane potentials, with a 3-minute rest interval in normal Tyrode's solution at -70 mV (Fig 6A). At each potential, we thus obtained I_Ca for 8 to 12 different amplitudes of Ca²⁺ release (Fig 6B, left; Fig 1 also illustrates the first six steps of such a protocol). We then quantified the inhibition of I_Ca by [Ca²⁺]_i as the fraction of current remaining at 150 ms after the depolarizing step and plotted this fraction as a function of the amplitude of the [Ca²⁺]_i transient at the same time (Fig 6B, right). We preferred this procedure over a fitting of the decline of I_Ca with two (or more) exponentials, because it allowed a comparison of I_Ca and [Ca²⁺]_i at the same point in time. The time of 150 ms was chosen because it is well beyond the time of peak [Ca²⁺]_i, and gradients are likely to be less important at this time.

View larger version (25K): [in this window] [in a new window]   Figure 6. Voltage dependence of inhibition of Ca2+ current (ICa) during Ca2+ release. A, Pulse protocol. V indicates membrane voltage. The cell was held at -70 mV in normal Tyrode's solution. Then it was switched to Na+-free solution (5.4 mmol/L CaCl2), and the holding potential was set at -45 mV. Twelve pulses to the test potential were given, and the cell was returned to the normal Tyrode's solution for 3 minutes. The protocol was then repeated for a different test potential. B, Example of four pulses to +40 mV with different amplitudes of Ca2+ release. I indicates membrane current. The current at 150 ms after the depolarizing step (I150ms) is measured as the fraction of the peak current (Ipeak), and the amplitude of the [Ca2+]i transient is measured at that same time. C, Plot of the inhibition of ICa as a function of the amplitude of the [Ca2+]i transient at that time for steps to -20 and to +10 mV. The relative inhibition (slope of the relation) is the same at the two potentials. D, Similar plot as in panel C obtained in a different cell for steps to 0 and to +40 mV (original tracings in panel B). Again, the slope of this relation is the same at the two different potentials.

This protocol was applied to 10 cells, in a total of 25 runs. In all runs, a small increase in resting [Ca²⁺]_i was noted. On average, resting [Ca²⁺]_i was 69.6±14.1 nmol/L (mean±SD) at the start and 132.8±42.8 nmol/L at the end of the run. In most runs, the peak of I_Ca remained stable, or some rundown occurred. In 10 runs, a moderate increase in peak I_Ca was observed, from 13% to 45% (range). If the increase exceeded 20%, the runs were not included in the analysis, and conclusions are based on a total of 18 runs in 7 cells.

Panels C and D of Fig 6 are plots of the inhibition of I_Ca by [Ca²⁺]_i for different membrane potentials, as obtained by the protocol outlined above, in two different cells. If a voltage-dependent mechanism is present, one expects the slope of the plot of the more positive potentials to be steeper; ie, for the same change in [Ca²⁺]_i, the relative decrease in I_Ca will be larger at the more positive potential. In the two cells illustrated in panels C and D, the slope of the plots obtained at the more positive potentials is comparable to the one of the plots obtained at more negative potentials, despite a voltage difference of 30 and 40 mV, respectively. In other cells, the slopes could differ slightly, but no consistent differences were found; ie, sometimes positive voltages were slightly steeper, and sometimes negative voltages were slightly steeper.

This quantitative analysis makes it unlikely that the transient inhibition of I_Ca during Ca²⁺ release results from a strongly voltage-dependent mechanism. Therefore, it is also unlikely that the recovery of I_Ca on relaxation of [Ca²⁺]_i would be voltage dependent. This was investigated by applying the analysis presented in Fig 2 at different potentials. For each potential, currents and [Ca²⁺]_i tracings with a large and a small Ca²⁺ release were obtained, and the difference current and difference [Ca²⁺]_i were calculated (see Fig 2). Fig 7 shows superimposed tracings of these subtracted-current and [Ca²⁺]_i tracings. The recovery from inhibition can be observed at all potentials. In a total of five cells, two to five membrane potentials were compared in this way in each cell. No differences were found, supporting the hypothesis that the recovery from Ca²⁺-dependent inhibition is not voltage dependent.

View larger version (10K): [in this window] [in a new window]   Figure 7. Recovery of Ca2+ current at all po- tentials. A depolarizing step with a minimal Ca2+ release and one with a large Ca2+ release were obtained at each of the indicated potentials. The tracings shown are the current differences (tracings marked by arrows, low-amplitude noise) and [Ca2+]i differences (high-amplitude noise) obtained by subtracting tracings from the pulse with minimal Ca2+ release from the pulse with a large Ca2+ release, as illustrated in Fig 2. Depolarizing steps to the indicated potentials are shown. All tracings are from the same cell.

Reduction in Driving Force due to the Increase in [Ca²⁺]_i
As the increase in cytoplasmic [Ca²⁺] during Ca²⁺ release will shift the reversal potential to more negative values, the resulting decrease in the driving force for Ca²⁺ influx is expected to decrease the amplitude of I_Ca. Can we explain the observed inhibition of I_Ca by this mechanism? We have calculated that an increase in [Ca²⁺]_i from 100 to 500 nmol/L decreases the reversal potential from +140 to +120 mV (with 5.4 mmol/L external CaCl₂ in our experimental conditions, Ca²⁺ is the only permeant cation). The effect of a decrease in driving force will be different at different membrane potentials, although such differences are small; ie, at 0 mV, the current will decrease by 14%; at +40 mV, by 20%. We have measured the decrease in current amplitude for an increase in [Ca²⁺]_i of 500 nmol/L from experiments illustrated in Fig 6. We measured the current amplitude at 150 ms after the depolarizing step, as in Fig 6B, for a [Ca²⁺]_i transient with an amplitude of 100 nmol/L and for a [Ca²⁺]_i transient with an amplitude of 500 nmol/L. For steps to 0 mV, we found that the current amplitude had decreased by 50% to 55% (range, n=3); for steps to +40 mV, by 41% to 73% (range, n=3).

	Discussion

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We have demonstrated that the transient increase in cytoplasmic Ca²⁺ due to Ca²⁺ release from the SR is an important determinant of the time course of I_Ca during depolarization. The inhibition of I_Ca was predicted from previous studies that had demonstrated that elevation of cytoplasmic Ca²⁺ decreases L-type current^{1 13} and reduces the probability of opening of single L-type Ca²⁺ channels.^{9 10 12} However, the rapid recovery was not predicted. Our report is the first that has documented the modulation of I_Ca by Ca²⁺ release from the SR in detail, in the absence of other Ca²⁺-activated currents. In the following paragraphs, we will discuss some of the particular features of this modulation revealed in the present study.

Early Inhibition of I_Ca Apparently Exceeds [Ca²⁺]_i Measured With Fura 2
Although the time course and amplitude of the transient inhibition of I_Ca were clearly related to the amplitude and time course of the [Ca²⁺]_i signal, during the first 50 to 100 ms the inhibition of I_Ca exhibited an "overshoot" (Fig 2, lower tracings). Such an observation could be explained by the existence of a higher [Ca²⁺] near the channel than the cytosolic concentration measured with fura 2. The existence of concentration gradients within the cell is strongly supported by theoretical calculations⁴⁰ and should be particularly important during this early period of the Ca²⁺ transient, when most of the Ca²⁺ release occurs.^{2 3 41} The particular spatial organization of the channels involved, with the Ca²⁺ release channel in proximity to the L-type Ca²⁺ channel, would certainly favor the occurrence of gradients in this subsarcolemmal area.^{4 5} Many previous experimental observations can only be explained by assuming the existence of such gradients,^{30 42 43} and it is very tempting to ascribe our present observation to a similar phenomenon. Membrane currents may indeed be exquisite "sensors" of subsarcolemmal ion concentrations and have been used as such in other cell types.^{44 45} However, such an interpretation remains tentative and at present must be viewed with caution. At least two other causes of the discrepancy must be considered. [Ca²⁺]_i during this period may be falsely underestimated because of the properties of the indicator. Because of its low K_d (200 nmol/L), measurements by fura 2 of [Ca²⁺] >1 µmol/L become uncertain. In skeletal muscle, a comparison of simultaneous measurements of [Ca²⁺]_i by fura 2 and a low-affinity indicator^{46 47} has demonstrated that in this preparation fura 2 underestimates the true [Ca²⁺]_i. The kinetics of the dye may also be too slow to follow the rapid changes in [Ca²⁺]_i, a possible source of error that we tried to compensate for by using the rate equation to calculate [Ca²⁺]_i rather than the equilibrium equation.² A second possible reason for the observed discrepancy is that the relation between inhibition of I_Ca and [Ca²⁺]_i is not necessarily linear. A recent study of Ba²⁺ currents through L-type Ca²⁺ channels in excised patches indeed showed a nonlinear relation.¹²

Therefore, at this point we cannot unequivocally conclude that the discrepancy between the time course of the inhibition of I_Ca and of the [Ca²⁺]_i transient during the period of Ca²⁺ release results from the existence of subsarcolemmal gradients. However, if the exact quantitative relation between the degree of inhibition of the Ca²⁺ channel and [Ca²⁺] at its cytoplasmic side could be established in independent experiments, then analysis of Ca²⁺ channels in situ during Ca²⁺ release could provide an exciting means to explore the actual [Ca²⁺] near the channel during this time.

Mechanism of the Modulation of I_Ca by Ca²⁺ Release From the SR
An unexpected finding was that the observed inhibition is rapidly reversible, because it follows very closely the changes in [Ca²⁺]_i. The mechanism of this rapid modulation was not related to a phosphorylation cycle, since it could still be observed after phosphorylation with ATP--S, and it was not voltage dependent. Therefore, it must be related to Ca²⁺ binding and unbinding to a regulatory cytoplasmic site outside the pore and/or to a reduction in driving force.

The decrease in current amplitude we have observed for an increase in [Ca²⁺]_i of 500 nmol/L was much larger than predicted by a reduction in driving force alone. As discussed above, [Ca²⁺]_i measured by fura 2 may underestimate the actual [Ca²⁺], but this is less likely to be important for values of 500 nmol/L. We have calculated that to account for the decrease in current amplitude by a reduction of driving force, the actual [Ca²⁺]_i has to be between 20 and 100 µmol/L. Until now, no experimental evidence could support these values. Even with aequorin, a lower affinity Ca²⁺ indicator, peak [Ca²⁺]_i in heart muscle rarely exceeds 1 to 2 µmol/L. Therefore, we conclude that a reduction in driving force due to the increase in cytoplasmic [Ca²⁺] may contribute to but cannot fully explain the observed decrease in I_Ca. What about the local subsarcolemmal [Ca²⁺]? There is no doubt that during Ca²⁺ flux through the L-type Ca²⁺ channel and through the Ca²⁺-release channel, high concentrations will occur close to the channel mouth.⁴⁰ On the other hand, such gradients will disappear very rapidly (in the order of a millisecond) on closure of the channel by diffusion, buffering, and Ca²⁺ removal.⁴⁸ The largest gradients are therefore expected early during the pulse (first 50 to 60 ms) during Ca²⁺ release from the SR. Our calculations were performed for current amplitudes and [Ca²⁺]_i values measured at 150 ms after the depolarizing step, and gradients are more likely to be small, but their existence cannot be dismissed. It has been suggested before that local subsarcolemmal increases in [Na⁺] can induce shifts in reversal potential when the transsarcolemmal Na⁺ currents are measured.⁴⁹ However, it remains difficult at present to predict the effect of local ion accumulation on the driving force. [Ca²⁺] gradients may also induce potential gradients, if they are not accompanied by equal gradients of negative charges. Local accumulation of Ca²⁺ may also affect surface charges. To evaluate a change in reversal potential due to local accumulation of Ca²⁺, ideally one should measure the reversal potential directly across the channel, with reference in this subsarcolemmal space.⁵⁰

Studies of the effect of an increase in cytoplasmic Ca²⁺ on single Ca²⁺ channel current recordings predominantly indicate a reduction in the number and duration of the channel openings rather than a reduction in the single-channel current amplitude.^{9 12} These findings point toward a regulatory effect of Ca²⁺ on channel gating rather than on driving force.

In conclusion, the mechanism underlying the rapid inhibition and recovery of I_Ca observed during Ca²⁺ release from the SR can at present not be identified with absolute certainty. Our data indicate that it does not involve a (de)phosphorylation process. The effect of the increase in the bulk cytoplasmic [Ca²⁺]_i on the reversal potential is too small to account for the observed dramatic effect, although local accumulation of Ca²⁺ may possibly have a more pronounced effect on the driving force. Rapid binding and unbinding of Ca²⁺ to a regulatory site can be involved. Because transmembrane voltage has no effect on this [Ca²⁺]_i-dependent modulation, this regulatory site must be located outside the pore.

Modulation of I_Ca by Ca²⁺ Release: Physiological Implications
In the theory of local control of excitation-contraction coupling, one L-type Ca²⁺ channel controls a cluster of SR release channels.^{3 41 48} Activity of the release channel is a function of the open probability of the sarcolemmal channel and of the amplitude of the current through the sarcolemmal channel. Rapid inhibition of the sarcolemmal Ca²⁺ channel by the Ca²⁺ released from the SR could act as a negative feedback on Ca²⁺ release. To evaluate its role in this context, calculations incorporating the Ca²⁺-dependent inhibition of the sarcolemmal Ca²⁺ channel are needed. Moreover, the model would have to incorporate the fact that apparently this inhibition is readily reversible.

The observation that during maintained depolarization, recovery of I_Ca occurs as [Ca²⁺]_i declines may provide additional insight into the mechanisms responsible for the generation of early afterdepolarizations. These early afterdepolarizations have been ascribed to reactivation of the L-type I_Ca during prolonged depolarization.^{51 52} This reactivation occurs because steady state inactivation and activation curves overlap, creating a "window" of potentials during which Ca²⁺ current can be maintained and recovery from inactivation can occur. Such window currents were demonstrated in single cardiac Purkinje cells.⁵³ Our present observations suggest the existence of an additional mechanism for reactivation of I_Ca, namely, recovery from Ca²⁺-dependent inhibition.

The effects of the Ca²⁺ release–dependent modulation of I_Ca during normal action potentials are more difficult to predict. From Fig 2, it is clear that the amplitude of the current changes is sizable and by itself would have a major effect on the time course of the action potential. However, in normal conditions, Ca²⁺ release will also evoke an inwardly directed Na⁺-Ca²⁺ exchange current, thus offsetting the change in total membrane current. This simultaneous occurrence of the inward Na⁺-Ca²⁺ current with the decrease in I_Ca may explain why the dramatic effects of Ca²⁺ release on I_Ca have not been noticed in previous experiments.

In conclusion, Ca²⁺ release from the SR is an important modulator of the time course of I_Ca during depolarization and will act in concert with previously described voltage-dependent modulation and with modulation by the permeating Ca²⁺ ions. Any change in the amplitude of the Ca²⁺ release will have profound effects on the time course of I_Ca.

	Acknowledgments

 
The authors wish to thank Diane Hermans and Luce Heremans for technical assistance. Dr Sipido is a Postdoctoral Researcher of the National Foundation for Scientific Research, Belgium.

	Footnotes

 
Previously published as a preliminary report in abstract form (Circulation. 1993;88[suppl I]:I-278).

Received February 24, 1994; accepted September 14, 1994.

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