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Am J Physiol Heart Circ Physiol 293: H563-H573, 2007. First published March 30, 2007; doi:10.1152/ajpheart.00469.2006
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Effect of intracellular Ca2+ and action potential duration on L-type Ca2+ channel inactivation and recovery from inactivation in rabbit cardiac myocytes

Julio Altamirano and Donald M. Bers

Department of Physiology, Stritch School of Medicine, Loyola University Chicago, Maywood, Illinois

Submitted 8 May 2006 ; accepted in final form 26 March 2007


   ABSTRACT
TOP
 ABSTRACT
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Ca2+ current (ICa) recovery from inactivation is necessary for normal cardiac excitation-contraction coupling. In normal hearts, increased stimulation frequency increases force, but in heart failure (HF) this force-frequency relationship (FFR) is often flattened or reversed. Although reduced sarcoplasmic reticulum Ca2+-ATPase function may be involved, decreased ICa availability may also contribute. Longer action potential duration (APD), slower intracellular Ca2+ concentration ([Ca2+]i) decline, and higher diastolic [Ca2+]i in HF could all slow ICa recovery from inactivation, thereby decreasing ICa availability. We measured the effect of different diastolic [Ca2+]i on ICa inactivation and recovery from inactivation in rabbit cardiac myocytes. Both ICa and Ba2+ current (IBa) were measured. ICa decay was accelerated only at high diastolic [Ca2+]i (600 nM). IBa inactivation was slower but insensitive to [Ca2+]i. Membrane potential dependence of ICa or IBa availability was not affected by [Ca2+]i <600 nM. Recovery from inactivation was slowed by both depolarization and high [Ca2+]i. We also used perforated patch with action potential (AP)-clamp and normal Ca2+ transients, using various APDs as conditioning pulses for different frequencies (and to simulate HF APD). Recovery of ICa following longer APD was increasingly incomplete, decreasing ICa availability. Trains of long APs caused a larger ICa decrease than short APD at the same frequency. This effect on ICa availability was exacerbated by slowing twitch [Ca2+]i decline by ~50%. We conclude that long APD and slower [Ca2+]i decline lead to cumulative inactivation limiting ICa at high heart rates and might contribute to the negative FFR in HF, independent of altered Ca2+ channel properties.

calcium ion current; excitation-contraction coupling; calcium ion buffers

DURING EXCITATION-CONTRACTION coupling in cardiac ventricular myocytes, Ca2+ current (ICa) through voltage-dependent L-type Ca2+ channels (LCC) triggers sarcoplasmic reticulum (SR) Ca2+ release. Upon sustained depolarization, whole cell ICa reaches a peak and then declines at a rate that is dependent on both membrane potential (Vm) and local Ca2+ concentration ([Ca2+]) at the inner mouth of the LCC (3, 8, 18). However, Ca2+-dependent inactivation occurs faster than the Vm-dependent inactivation and accounts for most of the channel inactivation during the cardiac action potential (AP; see Ref. 3). Ca2+ influx via ICa itself contributes to significant Ca2+-dependent inactivation (e.g., Ba2+ or Na+ currents through LCC inactivate much more slowly; reviewed in Ref. 3), and evidence for this is even seen in unitary current recording (15). However, the large SR Ca2+ release induced by ICa in cardiac myocytes further hastens ICa inactivation (during square pulses or AP; see Refs. 31, 36, and 37) such that Ca2+-dependent (vs. Vm-dependent) inactivation is by far predominant under physiological conditions.

Many molecular details of Ca2+-dependent inactivation have been described (6, 24, 28, 46–48). Calmodulin (CaM) tethered to a CaM-binding domain in the COOH terminal of the LCC {alpha}1C-subunit serves as the local Ca2+ sensor. Because this CaM is strategically located near the conduction pathway, a Ca2+-dependent conformational change allows CaM to bind another effector site(s) (including the IQ domain in the COOH terminal of {alpha}1C), thereby blocking conduction.

Statically elevated intracellular [Ca2+] ([Ca2+]i) can either increase or decrease ICa, probably reflecting the coexistence of Ca2+-dependent inactivation and facilitation of ICa (12, 47). ICa facilitation is seen upon repeated activation of ICa as an increase in the amplitude and time constant of inactivation and involves Ca2+- and CaM-dependent kinase II (CaMKII) phosphorylation of LCC (2, 7, 14, 43, 45). It is not clear whether the same CaM molecule is involved in Ca2+-dependent ICa inactivation and facilitation, but both processes can occur simultaneously. Depending on the [Ca2+]i level and kinetics of change, one or the other may dominate the whole cell ICa.

Although the process of ICa inactivation has been studied in detail, there is little information about how [Ca2+]i influences the kinetics of recovery from inactivation or LCC availability, especially in the context of ventricular myocytes with AP waveforms and Ca2+ transients. These are issues we address in the present study. Indeed, recovery from inactivation has been studied but mainly with respect to the Vm dependence (not [Ca2+]i dependence). The interplay between these may be especially important physiologically with changes in heart rate and inotropic state, that is, the ICa available during a given AP may depend on the prior AP waveform, rate of local [Ca2+]i decline, and diastolic [Ca2+]i and Vm.

This interplay between [Ca2+]i and Vm dependence of ICa may be even more important in pathophysiological states. For instance, the typical positive force-frequency relationship (FFR) in humans (and most mammals) is flattened or reversed in HF (1, 3, 22, 26). This may result from reduced SR Ca2+ uptake, which limits the ability to raise the SR Ca2+ content in heart failure (HF; see Refs. 10, 25, 26, 29, 41). However, the prolongation of AP duration (APD) and [Ca2+]i decline in HF may limit the recovery from inactivation of ICa at higher frequency and hence contribute to the blunted or reversed FFR in HF. Indeed, a frequency-dependent decrease in ICa was found in human HF (19, 27, 39, 40). Sipido et al. (39, 40) suggested that the high diastolic [Ca2+]i typical of HF could slow LCC recovery from inactivation, affecting availability, especially at high stimulation frequency, and exacerbated by the long APD in HF. The APD prolongation in HF is due in large part to the downregulation of outward K+ currents, including transient outward and inward rectifier channels (5, 29, 42).

In the present study, we assess the effect of Vm and elevated diastolic [Ca2+]i on ICa inactivation, availability, and recovery from inactivation under conditions where [Ca2+]i is heavily buffered at known levels in ruptured patch clamp. We also extend this to more a physiological setting in perforated patch, showing how slowed [Ca2+]i decline and prolonged APD influence ICa at different frequencies. We conclude that both slowed [Ca2+]i decline and prolonged APD (in the range seen in HF) can reduce ICa, even in normal ventricular myocytes.


   MATERIALS AND METHODS
TOP
 ABSTRACT
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Cardiac myocyte isolation. Myocytes were isolated from adult male New Zealand White rabbits following standard procedures reviewed and approved by the Institutional Animal Care and Use Committee of the Loyola University Chicago. Briefly, animals were anesthetized (pentobarbital sodium; 50 mg/kg), and the heart was quickly removed, rinsed in cold 0 mM Ca2+ Tyrode, placed on a Langendorff apparatus, and perfused through the aorta with a 0 Ca2+ solution for 5 min at 37°C. The heart was then perfused for 10–15 min with a solution containing collagenase B (Boehringer Mannheim). Left ventricle tissue was removed and cut into small pieces. Further incubation of tissue was done for 5–10 min, and digestion was stopped by adding a BSA-containing solution (1 mg/ml). Tissue was gently minced and filtered. Cells were stored in 50 µM Ca2+ at room temperature and used within 8 h. Only quiescent, rod-shaped myocytes with clear striations were included in this study. Before the experiment, cells were placed on laminin-coated cover slips.

Whole cell ruptured-patch and cytosolic Ca2+ buffering. The perfusion chamber was placed on the stage of an inverted microscope (Diaphot-TMD, Nikon), and membrane currents were measured in the whole cell-ruptured patch-clamp configuration (9). Ca2+ or Ba2+ was the charge carrier under extracellular and intracellular conditions designed to suppress all monovalent cationic currents. With the use of the patch pipette (1–2 M{Omega}), [Ca2+]i was buffered using 10 mM EGTA [dissociation constant (Kd) ~150 nM] or a combination of 5 mM 1,2-bis(2-aminophenoxy)ethane-N,N,N',N'-tetraacetic acid (BAPTA)·4Cs (Kd ~190 nM) plus 1 mM dibromo-BAPTA·4K (Kd ~1.80 µM). The later combination was preferred since those are fast Ca2+ chelators; their Kd values cover a broader spectrum compared with EGTA and are less pH sensitive.

Free [Ca2+] tested were (nM) 0, 100, 300, and 600 (free [Ca2+] was calculated with MaxChelator; http://www.stanford.edu/~cpatton/maxc.html; see Ref. 4). The highest practical [Ca2+]i was 600 nM because of progressive cellular contracture at higher levels. In preliminary studies, we attempted to use 2,3-butanedione monixime (BDM) to prevent contraction and allow higher [Ca2+] levels. However, BDM itself modified ICa gating, even at low [Ca2+]i, so this approach was abandoned. In addition, these solutions contained (in mM) 105 CsCl (120 in the EGTA-containing solution), 5 Tris-ATP, 20 HEPES, 0.3 GTP-Na, and 1 free Mg2+, pH 7.2 (CsOH). The gigaohm seal was achieved in normal Tyrode solution containing (in mM): 140 NaCl, 6 KCl, 10 glucose, 10 HEPES, 1 MgCl2, and 2 CaCl2, pH 7.4 (NaOH). Upon attaining the whole cell configuration, cells were bathed with recording solution (0 K+, Cs+ substituted) containing 2 CaCl2 or 1 BaCl2. Experiments were performed at 20–23°C. Voltage control and data acquisition were achieved by using a patch-clamp amplifier (Axopatch 200) and pClamp 8 software (Axon Instruments).

Perforated-patch and AP clamp. We used a perforated patch clamp (amphotericin B; see Ref. 32) to test for the effect of APD on ICa recovery from inactivation in nondialyzed cells at physiological temperature with Ca2+ transients. AP waveforms were used as depolarizing conditioning Vm command pulses. The pipette solution contained (in mM) 80 CsMES, 40 CsCl, 10 HEPES, 1 KCl, 1 CaCl2, and 240 µg/ml amphotericin B (from a stock of 60 mg/ml in DMSO), pH 7.4 (CsOH). CaCl2 was included in the pipette solution to monitor any accidental disruption of the plasma membrane and cytosolic dialysis. The electrodes had a resistance of 1–1.2 M{Omega}. The giga seal was obtained in normal Tyrode at 37°C, and the cell remained in that solution until attaining whole cell configuration (~15–30 min; access resistance ≤11 M{Omega}). Next, the external solution was switched to the ICa recording solution containing (in mM) 140 NaCl, 6 CsCl, 10 glucose, 10 HEPES, 1 MgCl2, 2 CaCl2, 0.02 niflumic acid, and 0.02 TTX, pH 7.4 (CsOH). In this solution, Cs+, niflumic acid, and TTX were used to block contaminating K+, Cl– and Na+ currents, respectively. All junction potentials were corrected. The experiments were performed at 37°C.

Two basic AP waves (see GoGoGoGoGoFig. 6A) were generated with the software LabHeart (30), one simulating a normal steady-state rabbit AP at 1 Hz (normal AP; ~180 ms) and the second was a longer AP simulating a steady-state HF (1 Hz; HF AP; ~330 ms). The duration of each AP was decreased by 10 and 15% to simulate the AP shortening at 2 and 3 Hz, respectively. Specific voltage-clamp protocols are described in RESULTS and in Fig. 6A.


Figure 1
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  		Fig. 1. Effect of diastolic intracellular Ca2+ concentration ([Ca2+]i) on peak Ca2+ (ICa) or Ba2+ (IBa) current density and inactivation rate. A: ICa measured in a cell dialyzed with a Ca2+-buffered solution containing 100 nM free [Ca2+]i at various test depolarizations [holding potential (VHold) –80 mV]. B: peak ICa (top) and IBa (bottom) density as a function of membrane potential (Vm) measured in cells dialyzed with one of various [Ca2+]i [0 (circle), 100 (square), 300 (triangle), and 600 (diamond) nM]. C: ICa and IBa inactivation time course. Ca: normalized ICa and IBa evoked by a 0-mV step in a cell dialyzed with a solution containing 100 nM free Ca2+. Cb and Cc: time to 50% current decay (T50%) for ICa (b) and IBa (c) plotted as a function of [Ca2+]i. Each data point represents the mean value (±SE) of 4–9 cells.

 

Figure 2
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  		Fig. 2. Diastolic [Ca2+]i and Vm dependence of ICa steady-state inactivation. A: double-pulse protocol (inset) was applied to cells dialyzed with one of various [Ca2+]i [0 (circle), 100 (square), 300 (triangle), and 600 (diamond) nM] and relative ICa amplitude in response to P2 as a function of the voltage of P1. P1 is a 2-s conditioning step to different Vm, which induces activation and some degree of steady-state inactivation. P2 is a constant test pulse (0 mV) used to measure the resultant ICa amplitude and hence the fraction of channels not inactivated (available) by P1. A Boltzmann relation was fitted to the ICa values measured at different [Ca2+]i (solid lines). B: plots of the values of the Boltzmann coefficient (k; left) and Vm at which 50% of the channels are available (V50, right) obtained from the Boltzmann relation as a function of diastolic [Ca2+]i. Each data point represents the mean value ± SE of 4–10 cells.

 

Figure 3
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  		Fig. 3. Diastolic [Ca2+]i and ICa recovery from inactivation. A: double-pulse protocol (inset) was used in cells dialyzed with one of various Ca2+ concentrations ([Ca2+]) [0 (circle), 100 (square), 300 (triangle), and 600 (diamond) nM]. The ratio of the peak amplitude of ICa in response to P2 over that evoked by P1 (P2/P1) is plotted as a function of the period of time between P1 and P2 from a VHold of –85 mV. Solid lines represent single exponential fits. B: plot of the recovery time constant ({tau}) values as a function of [Ca2+]i for ICa (bullet) and IBa ({circ}). Each data point represents the mean value ± SE of 4–9 cells. The {tau} was statistically different at all [Ca2+]i compared with 0 nM, for both ICa and IBa (P < 0.05).

 

Figure 4
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  		Fig. 4. Vm and Ca2+ dependence of ICa recovery from inactivation. A: time constant ({tau}) of ICa recovery from inactivation was both Vm and Ca2+ dependent. The protocol shown in Fig. 3 was applied at 3 different VHold (–85, –70, and –55 mV) and different diastolic [Ca2+]i [0 (circle), 100 (square), 300 (triangle), and 600 (diamond) nM]. Inset emphasizes the increase in {tau} as a function of [Ca2+]i from a VHold = –70 mV (P < 0.05). B: recovery from inactivation of IBa evaluated as described for ICa.

 

Figure 5
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  		Fig. 5. Diastolic [Ca2+]i and frequency-dependent changes on ICa and IBa amplitude. Trains of 10 pulses to 0 mV were applied at a frequency of 1 Hz to cells dialyzed with one of various [Ca2+] [0 (circle), 100 (square), 300 (triangle), and 600 (diamond) nM]. Normalized ICa (A) and IBa (B) amplitudes were plotted as a function of time. The cumulative effect of high [Ca2+]i on slower recovery from inactivation caused a progressive ICa amplitude decline during repeated stimulation at [Ca2+]i ≥100 nM. IBa showed a negative staircase at all [Ca2+]i tested, as expected for its slower recovery from inactivation time course.

 

Figure 6
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  		Fig. 6. Effect of action potential duration (APD) and frequency on ICa recovery from inactivation. Action potential (AP) templates of different duration were used as depolarizing waves under perforated patch-clamp conditions. A: protocol: a conditioning train of 5 AP templates (x5) of similar duration was followed by a square test pulse (P2; from a VHold –86 to +10 mV, 200 ms) at a variable time interval. To normalize peak ICa, a control square pulse (P1), with similar amplitude and duration as P2, was applied before the conditioning AP train. Right: two groups of AP templates used in the conditioning train: Normal (N APs, black traces) of ≤180 ms and long APs (≤330 ms) simulating heart failure (HF) conditions (HF APs, gray traces). Solid lines represent steady-state APs at 1 Hz; dashed and dotted lines on each group simulate a 10 and 15% decrease in APD at 2 and 3 Hz, respectively. Those APs in the conditioning trains were applied at one of three different frequencies. B: representative traces of ICa recovery from inactivation with conditioning trains of AP waves of two different durations at 1 Hz. Top: normal AP (180 ms); bottom: HF AP (330 ms). The relative position for P1, the 5th AP in the conditioning train, and P2 is indicated at top. C: ratio of peak ICa at P2 over that at P1 (P2/P1) plotted as a function of the interval between the beginning of the 5th AP and the beginning of P2. Open symbols, normal APs; filled symbols, HF APs at three different frequencies [1 (circle), 2 (square), and 3 (triangle) Hz]. Inset emphasizes the delay in the onset of recovery when long APs were used. Each data point represents the mean value ± SE of 5–7 cells.

 
Statistics. All data were stored in a personal computer for off-line analysis and are reported as means ± SE. Data were compared by paired Student's t-test or ANOVA followed by all pairwise multiple comparison (Holm-Sidak; SigmaStat 3.0; SPSS). A probability level <0.05 was considered significant.


   RESULTS
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 GRANTS
 REFERENCES
 
ICa dependency on diastolic [Ca2+]i. Figure 1 shows amplitude and inactivation of the membrane current carried either by Ca2+ or Ba2+ (ICa or IBa, respectively) in cells dialyzed with various [Ca2+]i. Cells were depolarized from a holding potential (VHold) of –80 mV in 10-mV increments every 5 s (raw traces are shown in Fig. 1A). Peak ICa and IBa amplitude did not show significant changes (P > 0.05) at the different [Ca2+]i tested. ICa and IBa inactivation rate was evaluated as the time to 50% current decay (T50%) in response to a depolarizing step to 0 mV. As expected, from the low capability of Ba2+ to induce LCC inactivation (or trigger SR Ca2+ release), IBa inactivation was incomplete compared with almost complete ICa inactivation during the 200-ms voltage steps. High diastolic [Ca2+]i did not affect the rate of IBa inactivation at any [Ca2+]i. ICa inactivation was accelerated by elevating [Ca2+]i but only significantly so at 600 nM (P < 0.05 vs. 0, 100, and 300 nM). The fact that IBa inactivation was unaffected by rather high [Ca2+]i (600 nM) indicates that the faster ICa inactivation at 600 nM [Ca2+]i was likely mediated by Ca2+ influx and release rather than the high diastolic Ca2+ per se. This may be most apparent at 600 nM [Ca2+]i because the 4 mM BAPTA is more than half-saturated, such that the local buffering of Ca2+ entry and release by BAPTA and dibromo-BAPTA is somewhat less able to clamp [Ca2+]i near the channel mouth. Indeed, Ca2+-dependent inactivation is still functional (note the difference in T50% for ICa vs. IBa at all [Ca2+]i). These results are consistent with a Kd for Ca2+-dependent inactivation >600 nM (as directly estimated at 4 µM for Ca2+-dependent inactivation; see Ref. 13).

Steady-state ICa availability (Vm dependence) was also measured at different diastolic [Ca2+]i (i.e., the fraction of channels available for activation at any given Vm). A two-square pulse voltage-clamp protocol was applied (Fig. 2A, inset). P1 is a 2-s conditioning step to different Vm, which induces activation and some degree of steady-state inactivation. P2 is a constant test pulse (0 mV) used to measure the resultant ICa amplitude and hence the fraction of channels not inactivated (available) by P1. Availability was fit with a Boltzmann function (normalized ICa = 1/{1 + exp[(V50 – VP1)/k]}, where VP1 is the voltage of P1, V50 is the Vm at which 50% of the channels are available, and k is the Boltzmann coefficient, which indicates voltage sensitivity of the channels).

The symbols in Fig. 2A are means ± SE of 4–10 different cells. Each individual experiment was fitted separately, and the solid lines were generated using the mean values for V50 and k (Fig. 2B). Neither V50 or k were systematically affected by increased [Ca2+]i. V50 was more negative (P < 0.05), and the slope was reduced (P < 0.05) only at 600 nM [Ca2+]i (vs. 0 nM). Similar results were observed in both ICa and IBa (for IBa only a negative shift of V50 was seen at 600 nM; data not shown). Thus there might be a slight reduction in steady-state availability at high diastolic [Ca2+]i, but this effect is negligible at normal diastolic Vm (~80 mV).

Recovery from inactivation at different diastolic [Ca2+]i was assessed as well (Fig. 3). A two-square pulse protocol was applied (Fig. 3A, inset). P1 and P2 are both steps to +10 mV, but P1 is 2 s long to induce maximal inactivation. A variable time (t) separated P1 and P2. The ratio of ICa amplitude evoked by P2 to that evoked by P1 (P2/P1) is a measure of the fraction of channels recovered from inactivation. The time course of normalized ICa (P2/P1) was progressively slower with increasing [Ca2+]i (similar results were obtained with IBa). These results indicate that increased diastolic [Ca2+]i over the physiological range slows ICa and IBa recovery from inactivation. The parallel effects on ICa and IBa indicate that this [Ca2+]i effect is independent of whether inactivation during P1 was Ca2+ or Vm dependent. On the other hand, the longer {tau} values for IBa (including with [Ca2+]i ~0) might indicate that, if anything, recovery from Vm-dependent inactivation is slower than from Ca2+-dependent inactivation.

Because ICa recovery from inactivation is Vm dependent, we tested the effect of increased [Ca2+]i at three different VHold (–85, –70, and –55 mV). The results for ICa and IBa are summarized in Fig. 4 (note that the data at –85 mV are from Fig. 3B). It is clear that the voltage dependency of ICa recovery from inactivation is increased by high [Ca2+]i. The slowest ICa recovery occurred at VHold of –55 mV and [Ca2+]i of 600 nM. Indeed, there seems to be synergy between how elevated [Ca2+]i and depolarized Vm slow recovery from inactivation.

Slow ICa recovery from inactivation at increased [Ca2+]i implies that an increasing fraction of channels would remain inactivated at ≥1 Hz, and channels may accumulate in the inactivated state. We measured the beat-dependent changes in ICa and IBa peak amplitude at different [Ca2+]i (Fig. 5). Trains of 10 steps to 0 mV (VHold = –80 mV; 200 ms) were applied at a frequency of 1 Hz. Figure 5A shows cumulative ICa inactivation for [Ca2+]i ≥100 nM and progressive decrease in ICa (except in Ca2+-free EGTA solution where Ca2+-dependent ICa facilitation is evident). Because IBa had a slower recovery from inactivation time constant (Fig. 4B), the decrease in IBa amplitude was larger and occurred at all [Ca2+]i tested. Neither ICa nor IBa showed a clear dose-dependent effect of [Ca2+]i. Slower stimulation frequency (0.2 Hz) did not cause any decrease in ICa or IBa amplitude (data not shown). This amplitude decrease at 1 Hz could be interpreted as a frequency-dependent decrease in channel availability.

APD and ICa recovery from inactivation in nondialyzed cells. To extend these results under controlled conditions to a more physiological context, we carried out further studies using perforated patch with AP waveforms at 37°C and allowing normal Ca2+ transients to occur. We prolonged both APD and [Ca2+]i decline to simulate HF conditions on normal LCCs.

Figure 6A shows the protocol to assess ICa recovery after an AP. After a control square pulse with ICa fully available (P1), a train of five identical APs was given to 1) attain constant steady-state SR Ca2+ load and diastolic [Ca2+]i and 2) asses ICa recovery from inactivation after the last AP by a square test pulse at a variable time interval (P2). Peak ICa evoked by P2 was normalized to the amplitude of ICa at P1 (where both P1 and P2 were from VHold –86 to +10 mV, for 200 ms). The relative ICa (P2/P1) was plotted as a function of the time interval from the start of the last AP clamp depolarization (Fig 6C). The same protocol was applied with two groups of AP templates of different duration in the conditioning train; short (≤180 ms) and long (≤330 ms) duration, simulating normal and HF conditions, respectively (see Fig. 6A, right). These conditioning trains were applied at one of three different frequencies (1, 2, and 3 Hz, with APD adjusted accordingly). Figure 6B shows representative traces of ICa recovery from inactivation when two AP waves of different duration were used in the conditioning trains at 1 Hz [Fig. 6B, top, 180 ms (normal AP) and bottom, 330 ms (HF AP)].

Figure 6C shows pooled data. There was a clear delay in the onset of ICa recovery with long APD, but the subsequent rate of recovery was also slower (see Fig. 6C, inset). In all cases, ICa recovered ~100% within 2 s. Each point represents the mean value from five to seven different cells, and six different APDs were used in the conditioning trains. The curves in Fig. 6C are most relevant for considering ICa restitution at different heart rates or cycle lengths, as often used in programmed stimulation in intact hearts. For example, one can appreciate from these curves that as heart rate increases from 1 to 2 Hz (1,000- to 500-ms interval) ICa would be incompletely recovered between beats.

In Fig. 7 we extend the analysis of the data from Fig. 6C but now with the time axis shifted such that t = 0 is taken at the end of AP repolarization (≥85%). Thus, in Fig. 7A, recovery time is at constant Vhold = –86 mV, and might be expected to be more Vm and APD independent. With longer APDs, ICa recovery was slower and clustered to the right, whereas with short APDs ICa recovery was faster and tightly grouped at left. Double exponential functions were fit to the time course of each P2/P1 dataset. Figure 7, B and C, shows that the slower recovery phase was not different among any group. However, the fast recovery phase was longer with long APD. The relative amplitude of the two exponential components was ~1:1 (data not shown) and was not affected by APD or frequency. Possibly, the slower terminal repolarization in the long APD group causes the slower fast recovery for that group.


Figure 7
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  		Fig. 7. Long APD slows ICa recovery from inactivation. A: recovery time course of the normalized ICa amplitude (P2/P1) following trains with normal APs (open symbols) and HF APs (filled symbols) at three different frequencies [1 (circle), 2 (square), and 3 (triangle) Hz]. Lines represent double-exponential functions for each data set of normalized ICa (solid lines, conditioning train applied at 1 Hz; dashed line, 2 Hz; and dotted lines, at 3 Hz). Notice that the plot displays only the first stage of the recovery process, and all data sets were shifted to time 0 (see text). Fast (B) and slow (C) time constants ({tau}) for the indicated APs and frequencies in the conditioning trains. The fast time constant was greatly affected by APD, whereas the slow constant did not change significantly.

 
The implication from Figs. 6 and 7 is that, with longer APD, ICa recovery was increasingly incomplete because of both a delay in onset (longer APD) and also slower recovery rate after repolarization. This cumulative ICa inactivation would be reflected as a decrease in ICa availability at frequencies ≥1 Hz. Figure 8 shows ICa availability extrapolated from the datasets shown in Fig. 6C and plotted as a function of frequency. These allow us to predict that, at low frequency (~0.5 Hz), ICa availability is ~100%, but it sharply decreases at higher frequencies. This decrease is steeper when long (HF-like) APDs were used. Clearly cumulative ICa inactivation can be expected at higher frequencies, especially when APD is prolonged. The latter prediction was confirmed when trains of 10 APs with different duration (VHold = –86 mV) were applied at frequencies of 1, 2, and 3 Hz. Figure 8B shows representative ICa traces from the same cell using trains of a normal AP (180 ms) and a HF AP (330 ms), both at 1 Hz. Figure 8, C and D, shows pooled data from five different cells at each frequency. Cumulative ICa inactivation was clearly observed at high frequency (≥1 Hz), but this was more evident when long APD were used at all frequencies tested.


Figure 8
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  		Fig. 8. Long APD decreases ICa availability. A: data from Fig. 6 were used to extrapolate ICa availability (%) as a function of frequency. Normalized ICa (P2/P1) following trains of normal APs (open symbols) and HF APs (closed symbols) plotted as a function of frequency [1 (circle), 2 (square), and 3 (triangle) Hz]. With longer APD, recovery of ICa was increasingly incomplete, and the cumulative inactivation is reflected as a decrease in ICa availability at frequencies ≥1 Hz. B: ICa traces in response to trains of 10 APs with two different durations applied at 1 Hz in the same cell. Insets show amplifications of the peak of the main traces to illustrate the progressive decrease in ICa amplitude when a long APD is used. C: pooled data of normalized ICa amplitudes in response to normal (short) APD at three different frequencies (1, 2, and 3 Hz). D: normalized ICa (as in C) in response to trains of long APD (HF AP). Each data point represents the mean value ± SE of 5 cells.

 
Slowed relaxation and [Ca2+]i decline and smaller Ca2+ transient amplitude are hallmarks of HF, along with prolonged APD. To mimic these Ca2+-handling defects (but with normal Ca2+ channel characteristics), we slowed SR Ca2+ uptake by partially blocking the SR Ca2+-ATPase by controlled exposure to the effectively irreversible sarco(endo)plasmic reticulum (SERCA) inhibitor thapsigargin (TG). This allowed us to test the hypothesis that slowed [Ca2+]i decline slows ICa recovery from inactivation. Myocytes were pretreated with TG (100 nM for 5 min), and, to verify the effect of TG on the twitch Ca2+ transient, parallel experiments were done in fluo 3-loaded cells pretreated with TG. The amplitude of the Ca2+ transient ({Delta}F/F0) decreased by ~40% (from 2.40 ± 0.35 to 1.51 ± 0.12; 6–8 different cells; Fig. 9, A and C), and the relaxation time constant for the Ca2+ transient in TG-treated cells was slowed by approximately twofold (from 256 ± 17 to 592 ± 44 ms; Fig. 9, A and B).


Figure 9
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  		Fig. 9. Effect of thapsigargin (TG) on the Ca2+ transient amplitude and relaxation time constant in field-stimulated myocytes. Ca2+ transients were recorded in field-stimulated cells (0.2 Hz at room temperature) loaded with fluo 3. A group of cells was pretreated with 100 nM TG during 5 min to partially block the SR Ca2+-ATPase. A: Ca2+ transients (F/F0) for control (Ctl; no TG) and TG-treated cells (+TG). Inset: normalized Ca2+ transients (%) to show the slower relaxation time course on SR Ca2+-ATPase block. The relaxation time course was evaluated by a single exponential fit. B: mean time constant for both groups. The [Ca2+]i transient relaxation was slowed by ~50% after exposure to TG. C: peak Ca2+ transient amplitude ({Delta}F/F0) for Ctl and +TG. Each bar represents the mean value ± SE of 6–8 cells.

 
Figure 10 summarizes data from experiments like those in Figs. 6–8 done with or without the above TG pretreatment. Slowing [Ca2+]i transient decline with TG with control APD had little impact on the fast time constant of ICa recovery (Fig. 10A), but this results in a slight reduction in ICa availability for APs at 1–3 Hz (Fig. 10B). With the HF AP, the effect of TG on fast recovery of ICa was more noticeable, especially for HF AP at 3 Hz (Fig. 10B). Moreover, it is instructive to compare the control AP without TG with the HF AP + TG (i.e., with slowed [Ca2+]i decline and Ca2+ transient amplitude). This shows that the reduced ICa availability becomes increasingly prominent in the HF phenotype as frequency is increased and that slower [Ca2+]i decline significantly slows ICa recovery despite the smaller SR Ca2+ release. This is consistent with the notion that reduced ICa availability may contribute to reduced ICa (and hence Ca2+ transient amplitude) in HF at high heart rates and that both longer APD and altered Ca2+ transient kinetics contribute to this effect.


Figure 10
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  		Fig. 10. Slower [Ca2+]i transient decline slows ICa recovery from inactivation. Because TG decreases peak [Ca2+]i transient by ~40% and slowed its relaxation by ~50%, the role of increased diastolic Ca2+ in ICa recovery from inactivation was evaluated in TG-treated cells. A: fast {tau} of ICa recovery from inactivation in control and TG-treated cells for normal AP and HF AP in the conditioning trains applied at 1 Hz. Inset shows the two different AP waves used. TG slightly slowed ICa recovery (P > 0.05). B: extrapolated ICa availability is smaller in TG-treated cells (black symbols) compared with control cells (white and gray symbols for normal and HF AP, respectively) at high frequency of stimulation (3 Hz; P < 0.05 only for HF AP. Each data point represents the mean value ± SE of 6 and 3 cells for Ctl and +TG, respectively.

 

   DISCUSSION
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 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
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In the present study, we assessed the influence of [Ca2+]i on ICa in both static relatively controlled conditions and physiological conditions with AP and dynamic Ca2+ transients. We find that static elevation of [Ca2+]i in ventricular myocytes slows recovery from inactivation of both ICa and IBa and also that at 600 nM [Ca2+]i the steady-state availability of both ICa and IBa is shifted to more negative Vm. However, [Ca2+]i up to 600 nM had little influence on either the amplitude of fully available ICa or IBa or the kinetics of inactivation. We also show how these effects of [Ca2+]i on ICa recovery in physiological conditions work together with the Vm dependence of ICa to limit ICa at increased heart rate, especially under pathophysiological conditions where APD is prolonged and [Ca2+]i decline is slowed. This limitation of ICa recovery may contribute to the flat or negative FFR seen in HF.

Ca2+- and Vm-dependent ICa inactivation. ICa inactivation in ventricular myocytes is both Ca2+ and Vm dependent (8, 18) and typically exhibits a U-shaped availability curve (3, 17, 23). The minimal ICa availability occurs at Vm where ICa is maximal and is thus a hallmark of Ca2+-dependent inactivation. Ca2+-dependent inactivation is much faster than Vm-dependent inactivation, such that for long conditioning pulses the availability curve becomes sigmoid, reflecting greater Vm-dependent inactivation, especially at positive Vm where ICa is small (as in Fig. 2). Ca2+-dependent inactivation is now known to be mediated by Ca2+ binding to CaM tethered to a CaM-binding domain in the COOH terminal of the LCC {alpha}1C-subunit (6, 24, 28, 46–48). During a normal AP, Ca2+-dependent inactivation is by far dominant over Vm-dependent inactivation, even considering Ca2+ entry via ICa alone. The large Ca2+-induced Ca2+ release of Ca2+ from the SR further accelerates Ca2+-dependent inactivation (36, 38) such that only half as much integrated Ca2+ influx occurs when there is SR Ca2+ release during the AP (31). Indeed, the fast time constant for Ca2+-dependent inactivation of ICa with normal SR Ca2+ release is typically ~6 ms vs. several hundred milliseconds for purely Vm-dependent inactivation. Thus dynamic [Ca2+]i changes during ICa and SR Ca2+ release can clearly influence ICa inactivation and recovery there from.

We found only minor effects of static elevation of diastolic [Ca2+]i on either current inactivation or the Vm dependence of LCC steady-state inactivation. A slight acceleration of ICa inactivation at 600 nM [Ca2+]i (Fig. 1C) was not seen for IBa, so we do not attribute this to the diastolic [Ca2+]i per se but rather to the Ca2+ entry (with greater SR Ca2+ release) superimposed on a high diastolic [Ca2+]i with less capacity of the BAPTA/dibromo-BAPTA to buffer local [Ca2+]i. We did see a shift in ICa and IBa availability at 600 nM [Ca2+]i to more negative Vm. Although this could be a true effect of this [Ca2+]i on channel availability because of binding to an inactivating site, it might also be because of effects of high [Ca2+]i on membrane surface charges potential (i.e., shifting the Vm sensed by the channel). However, because activation curves were not similarly shifted (P > 0.05; data not shown), this might reflect a Vm-dependent Ca2+ dissociation step in channel availability. Overall, our results are consistent with a Kd for Ca2+-dependent inactivation much higher than 600 nM, such as the value of 4 µM estimated in single Ca2+ channel current recording (13). This is also consistent with results in excised "inside out" membrane patches from cardiac and smooth muscle myocytes, where a decrease in the ensemble average IBa and increased inactivation rate were observed with a Kd ~2 µM [Ca2+]i (33, 35).

Recovery of ICa from inactivation. Although both [Ca2+]i and Vm are expected to influence ICa recovery from inactivation, most prior work has focused on the Vm dependence of recovery, where recovery is faster at more negative Vm (3, 8, 16, 19). Imredy and Yue (15) argued that the long-lived Ca2+-dependent inactivated state could become unstable at hyperpolarized Vm, favoring the transition to a Ca2+-independent closed state (recovery from Ca2+-dependent inactivation) and availability for activation. This could explain the negative effect that depolarized Vm has on ICa recovery from inactivation. It is unclear how the cause of inactivation (Ca2+ or Vm) influences the recovery kinetics (although early data did not distinguish a difference; see Ref. 8). Here we found that IBa recovery was slower than ICa recovery at all values of VHold during the recovery phase (Fig. 3). Because more of the ICa (vs. IBa) inactivation is the result of Ca2+-dependent inactivation, this is consistent with recovery from Vm-dependent inactivation being slower. Sipido et al. (38) also showed that, during long depolarizations (4 s) with SR Ca2+ release, ICa could inactivate and recover from Ca2+-dependent inactivation and reactivate ICa (all without repolarization). This emphasizes that the dynamic interplay of [Ca2+]i and Vm during the AP and Ca2+ transient must be considered to understand the physiological situation for ICa recovery from inactivation (see below).

We show that ICa (and IBa) recovery from inactivation is slowed by both elevated [Ca2+]i and depolarized VHold. As expected, the time constant of recovery ({tau}) increased at depolarized VHold, but this increase was much steeper at higher [Ca2+]i (Fig. 4). For example, the ICa recovery {tau} at VHold = –55 mV and 600 nM [Ca2+]i was approximately threefold larger than that at VHold = –55 mV and 0 nM and approximately sixfold larger than that at VHold = –85 mV and 600 nM. This steep increase was clearly dependent on diastolic [Ca2+]i, since IBa displayed qualitatively the same response. This indicates synergistic regulation of recovery from inactivation by [Ca2+]i and Vm.

Effects of steady-state [Ca2+]i on ICa. Steady-state elevation of [Ca2+]i can either increase or decrease ICa in cardiac myocytes, probably reflecting the coexistence of Ca2+-dependent inactivation and CaMKII-dependent ICa facilitation (12, 47). ICa facilitation is seen upon repeated activation of ICa (and not IBa) as an increase in both the current peak and the time constant of inactivation, involving LCC CaMKII-dependent phosphorylation (2, 7, 14, 43, 45). It is unclear if the same CaM molecule is involved in Ca2+-dependent ICa inactivation and facilitation, but both processes can occur simultaneously. Depending on the [Ca2+]i level and kinetics of change, one or the other may dominate the whole cell ICa.

Yamaoka and Seyama (44) found in frog ventricular myocytes that increasing [Ca2+]i (from ~67 nM to 1.94 µM) via Na2+/Ca2+ exchange, facilitated LCC activity and enhanced inactivation rate. However, in many cases, a dual effect has been described. For example, Hirano and Hiraoka (12) found that elevating [Ca2+]i within 200–400 nM by high K+ depolarizations increased LCC open probability and availability (without affecting ICa) and suggested the involvement of a phosphorylation-dependent process (which retrospectively could have been ICa facilitation). However, further [Ca2+]i increase inhibited LCC activity (perhaps reflecting inactivation).

We found no significant effect of resting [Ca2+]i (0–600 nM) on peak ICa and IBa (Fig. 1); however, half-activation of IBa was shifted ~10 mV to negative Vm (vs. ICa). This shift is expected because Ba2+ is not as effective as Ca2+ in screening external membrane surface charge, decreasing the threshold for activation and inactivation (3). Interestingly, there was no shift in Vm dependence of either ICa or IBa at higher [Ca2+]i (a negative shift might be expected because of increased charge screening in the internal surface of the membrane). This suggests that these changes in [Ca2+] do not greatly alter internal surface potential, which is probably more influenced by the higher concentrations of free Mg2+ (1 mM) and K+ (>100 mM). This agrees with a lack of shift in the ICa vs. Vm relation upon an increase in [Ca2+]i in cardiac and smooth muscle myocytes (17, 23).

Although elevating [Ca2+]i from 100 to 600 nM would decrease the electrochemical driving force for Ca2+ (at 0 mV) by 18% (from 131 to 107 mV), no systematic decrease in ICa amplitude was seen (similar to previous results; see Ref. 38). This is presumably because these low [Ca2+]i values do not drive appreciable outward ICa, and the Hodgkin-Katz current equation (dominated by Ca2+ influx for physiological [Ca2+]i) is a better predictor of net ICa than Ohm's law (using the reversal potential for Ca2+).

Frequency-dependent inhibition of ICa in dialyzed cells. We tested the hypothesis that elevated diastolic [Ca2+]i would limit ICa recovery from inactivation at increasing frequency of depolarization. In cells where [Ca2+]i was relatively controlled, ICa recovered ~100% within 2 s, but the rate of recovery depended on [Ca2+]i (Fig. 3). As [Ca2+]i increased (to 100, 300, and 600 nM), the recovery time constant was 28, 59, and 113% slower, respectively (for VHold –85 mV, with greater slowing at more depolarized VHold). Because the conditioning pulses lasted 2 s, conditioning ICa and SR Ca2+ release did not affect ICa recovery (see above). Similar effects were obtained with IBa, but recovery was ~20–50% slower (at a VHold –85 mV). Even for 200-ms pulses at 1 Hz, there was a limitation of ICa that accumulated from pulse to pulse (Fig. 5). However, the extent of ICa diminution at 1 Hz was not strictly correlated with diastolic [Ca2+]i.

From the data in Fig. 4, we can expect most Ca2+ channels to recover from inactivation between pulses at 0.5 Hz. However, at 1 Hz, 7–10% of Ca2+ channels would not be available (for 0–600 nM [Ca2+]i), and this fraction would dramatically increase at 2 Hz to ~15–30% (for 0–600 nM [Ca2+]i). This prediction is confirmed by Fig. 5, and the effects can be cumulative. In addition, ICa at 0 nM [Ca2+]i showed a positive staircase (not seen for IBa). We attribute the increase in ICa amplitude in 0 [Ca2+]i to ICa facilitation, which becomes more apparent because of less cumulative Ca2+-dependent inactivation (with no SR Ca2+ release expected in these conditions). This is consistent with the IBa results where ICa facilitation is not expected. These results set up the studies done under more physiological conditions where we used APs (rather than square pulses), normal Ca2+ transients (vs. buffered [Ca2+]i), and perforated patch-clamp at 37°C (to minimize perturbing normal cellular Ca2+ homeostasis).

APD and ICa recovery from inactivation in nondialyzed cells. Sipido et al. (39, 40) reported a frequency-dependent decrease in ICa amplitude in human HF myocytes and suggested that increased diastolic [Ca2+]i might slow ICa recovery from inactivation and underlie this frequency-dependent ICa decrease. Although ICa properties may also change in HF (11, 27, 34), here we assessed the intrinsic effects of prolonged APD and slowed [Ca2+]i decline at different frequencies on normal Ca2+ channels. Because ICa time course is quite different during the AP than during square pulses (19–21, 31, 45), we used AP templates to drive conditioning pulses, with subsequent tests of ICa availability.

With long AP during the conditioning train to simulate the HF AP (Figs. 6 and 7), we saw the expected delay in ICa recovery (due to later repolarization) but also a substantial decrease in the rate of subsequent ICa recovery (Fig. 7, A and B) at all frequencies tested (1–3 Hz). There was no significant difference in the rate of ICa recovery within the long or short AP waveform groups at 1–3 Hz, despite significant differences between long vs. short AP groups. It is possible that slower ICa recovery (prolonged fast {tau}) after the longer APD templates is related to a greater extent of Vm-dependent inactivation (vs. the shorter APD) and a slower intrinsic recovery from Vm-dependent inactivation (of IBa vs. ICa above). Taken together, the delayed onset of ICa recovery and slower recovery rate would cause a progressive increase in the fraction of inactivated channels at frequencies ≥1 Hz. ICa recovered ~96–99% at ~2 s, regardless of APD in the conditioning train, such that little impairment of ICa availability is expected at 0.5 Hz. However, even at 1 Hz, 6–11% of the channels have not recovered from inactivation, and this is exacerbated at higher frequency (2–3 Hz), especially with the longer APD profile. These predictions were confirmed with trains of APs with variable duration applied at three different frequencies (1, 2, and 3 Hz; Fig. 8, B–D). Cumulative ICa inactivation was clearly exacerbated when long APD were used at all frequencies tested. At the 10th pulse, long APD caused cumulative inactivation of ~14% at 1 Hz, as opposed to only ~3% with short APD at the same frequency. This difference dramatically increased at 3 Hz, where long APD caused a decrease in peak ICa of ~55% compared with the 32% decrease caused by short APD. Taken together, these results indicate clear Vm-dependent effects on ICa availability that could contribute to the altered FFR in HF, independent of possible HF-induced Ca2+ channel modification.

[Ca2+]i transients with smaller peak amplitude and slower relaxation time course are also typical in HF, in addition to prolonged APD. To simulate this slowed [Ca2+]i decline in normal myocytes, we used timed treatment with the SERCA inhibitor thapsigargin, which slowed [Ca2+]i decline by ~50% and decreased peak amplitude of the [Ca2+]i transient by ~40%. Slowing [Ca2+]i decline also slowed ICa recovery (despite smaller Ca2+ transient amplitude), especially when long APs were used (Fig. 10A), and this would be reflected as a decrease in ICa availability at high frequencies (Fig. 10B). This indicates that both the kinetics of [Ca2+]i decline and repolarization work independently (or perhaps synergistically) to influence ICa availability in normal cardiac myocytes. The relative effects of these factors in limiting ICa availability in a specific pathophysiological condition (such as HF) will of course depend on 1) the detailed alteration in AP waveform (at different frequencies); 2) the kinetics and amplitude of the Ca2+ transient (at different frequencies), including diastolic [Ca2+]i; and 3) any alterations in ICa characteristics under that condition. This means that the effects of Vm and [Ca2+]i described here are important under pathophysiological conditions but that the specific situation must be directly assessed.

In conclusion, we demonstrated that increased steady-state levels of Ca2+, within the physiological range, affect ICa recovery from inactivation without major effects on other ICa properties. In addition, under more physiological conditions (perforated patch clamp at 37°C), long APs and elevated diastolic [Ca2+]i slowed ICa recovery. We believe that this is an important mechanism that might help to explain, in part, the negative frequency dependence of ICa amplitude in HF (27, 39, 40), where elevated diastolic [Ca2+]i and long APD are common.


   GRANTS
TOP
 ABSTRACT
 MATERIALS AND METHODS
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 GRANTS
REFERENCES
 
This work was supported by National Heart, Lung, and Blood Institute Grants HL-30077 and HL-64724.


   ACKNOWLEDGMENTS
 
We thank J. Acevedo and B. French for technical assistance.


   FOOTNOTES
 

Address for reprint requests and other correspondence: D. M. Bers, Dept. of Physiology, Loyola Univ. Chicago, 2160 South First Ave., Maywood, IL 60153 (e-mail dbers{at}lumc.edu)

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


   REFERENCES
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 ABSTRACT
 MATERIALS AND METHODS
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 DISCUSSION
 GRANTS
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