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CARDIOVASCULAR |
1 Laboratory of Experimental Cardiology, University of Leuven, Belgium
2 Department of Cardiology, Academic Hospital Maastricht, The Netherlands
3 Department of Medical Physiology, University Medical Center, Utrecht, The Netherlands
4 Department of Pharmacology and Toxicology, Comenius University, Bratislava, Slovakia
5 Department of Physiology, Charles University, Medical School Plzen, Czech Republic
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Abstract |
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-adrenergic stimulation. We studied the properties of ICaL in myocytes from the dog with chronic atrioventricular block (cAVB) that has cardiac hypertrophy and an increased susceptibility to TdP. Peak ICaL densities at baseline (K+- and Na+-free solutions, 10 mmol l–1
[EGTA]pip) in cAVB were comparable to control, but inactivation was shifted to the right, resulting in a larger window current area in cAVB. Under -adrenergic stimulation, the window current area was increased and shifted to the left, but less so in cAVB (maximum at –27 mV, versus
–32 mV in control). ICaL during a step to –35 mV showed a transient reduction immediately after the peak. Test steps to 0 mV, simultaneous recording of [Ca2+]i and manipulation of sarcoplasmic reticulum (SR) Ca2+ release showed that this resulted from inhibition and fast recovery of ICaL with SR Ca2+ release. The extent of this dynamic modulation was larger in cAVB than in control (23 ± 2% of the initially available current, versus 13 ± 3%; P < 0.05). Early afterdepolarizations (EADs) in cAVB myocytes under -adrenergic stimulation typically occurred in the window current voltage range and after decline of [Ca2+]i. In conclusion, in cAVB, the larger window current, its rightward shift and enhanced dynamic modulation by SR Ca2+ release may contribute to an increased incidence of EADs in cAVB under -adrenergic stimulation.
(Received 3 November 2006;
accepted after revision 24 November 2006;
first published online 30 November 2006)
Corresponding author K. R. Sipido: Laboratory of Experimental Cardiology, KUL, Campus Gasthuisberg O/N 7th floor, Herestraat 49, B-3000 Leuven, Belgium. Email: Karin.Sipido{at}med.kuleuven.ac.be
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Introduction |
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Previous studies on the mechanisms of EADs implicate reactivation of L-type Ca2+ current, ICaL, during prolonged repolarization (January & Riddle, 1989), or of Na+ currents in conditions of abnormal Na+ channel activity (Boutjdir et al. 1994; Studenik et al. 2001). Under -adrenergic stimulation, the shift of gating mode of Ca2+ channels into the high activity mode two facilitates EADs (Tanskanen et al. 2005). Furthermore, Ca2+ release from the sarcoplasmic reticulum is an important modulator of Ca2+ current through inactivation (Puglisi et al. 1999; Sham et al. 1995) and subsequent recovery from release-dependent inactivation (Sipido et al. 1995). Na+–Ca2+ exchange current, associated with Ca2+ release, can also contribute to EADs under adrenergic stimulation (Volders et al. 2000).
To study the TdP associated with cardiac remodelling, and the underlying cellular and molecular mechanisms, animal models have proven very useful. The dog with chronic atrioventricular block (cAVB) and biventricular hypertrophy has a high susceptibility to TdP-type arrhythmias and reiterates many characteristics of the clinical syndromes (Vos et al. 1998; van Opstal et al. 2001). At the cellular level, we observe hypertrophy and functional remodelling, with an increased susceptibility to EADs (Volders et al. 1998; Sipido et al. 2000). We postulated that loss of K+ currents forms the substrate for an increased incidence of EADs, which could be facilitated by an upregulated Na+–Ca2+ exchange current, in combination with increased sarcoplasmic reticulum (SR) Ca2+ release. Recently, Stengl et al. further emphasized how the lack of a significant increase in the slow delayed rectifier K+ current, IKs upon -adrenergic stimulation promotes EADs (Stengl et al. 2006). Although in a previous study (Sipido et al. 2000) we did not see differences in density of the peak inward ICaL in cAVB cells in basal conditions, it is possible that different kinetics or regulation of ICaL could contribute to the EADs under -adrenergic stimulation. Given the increased Ca2+ release in cAVB (Sipido et al. 2000), the Ca2+-dependent modulation of ICaL could also be altered in cAVB. In the current study we therefore investigated the properties of the Ca2+ current in cAVB myocytes, in particular its kinetics and its modulation by Ca2+ release from the SR at baseline and under -adrenergic stimulation.
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Methods |
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Total atrioventricular block was created in adult mongrel dogs of either sex (n = 9) as previously described (Vos et al. 1998). Animal handling was in accordance with the Dutch Law on Animal Experimentation and the European Directive for the Protection of Vertebrate Animals Used for Experimental and Other Scientific Purposes (86/609/EU). At the time of sacrifice, AVB duration was 48 ± 7 days and weight 24 ± 1 kg; control animals were of matched weight (n = 7, 27 ± 2 kg). Dogs received full anaesthesia. After premedication (1 ml (5 kg)–1: 10 mg oxycodone HCl, 1 mg acepromazine and 0.5 mg atropine, I.M.), sodium pentobarbital (20 mg kg–1 I.V.) was given. Dogs were artificially ventilated with a mixture of oxygen, nitrous oxide (40:60%) and halothane (0.5–1% vapour concentration). Upon thoracotomy, heparin was administered I.V. The hearts were quickly excised and washed in cold cardioplegic solution. Heart weight/body weight was significantly larger in cAVB dogs (12.1 ± 0.4 g kg–1, versus 8.6 ± 0.3 g kg–1 in controls, P < 0.001). Single myocytes were enzymatically isolated from the midmyocardial layer of the left ventricular free wall, as previously described (Volders et al. 1998).
Experimental setup
The experimental setup was built around an inverted microscope (Nikon Diaphot) for simultaneous recording of ionic currents and of intracellular Ca2+ (Sipido et al. 2000). Cells were placed in a perfusion chamber and superfused with external solution at a flow rate of 4 ml min–1. Rapid changing of the extracellular solution was accomplished with a solution switcher, controlled by PClamp 8.0 software (Axon Instruments). The temperature of the bath was controlled and set at 37°C.
Membrane currents were recorded using the whole-cell ruptured patch-clamp technique; patch pipettes had a resistance of 1.5–2.5 M
when filled with internal solution. Membrane currents were recorded with an Axopatch 1D amplifier, filtered at 2 kHz, and sampled and digitized at 4 kHz using a Digidata 1200A analog-to-digital converter and pCLAMP 8.0 software (Axon Instruments). The data acquisition program also controlled the command potential. Membrane currents (pA) were normalized to cell capacitance (pF). The average cell capacity in controls was 169 ± 8 pF (30 cells) versus 182 ± 7 pF in cAVB (37 cells). The trend towards larger capacity was not statistically significant but within the same range as the values previously reported for larger groups, which were statistically significant: Verdonck et al. (2003) found capacitance values of 165 ± 5 pF (91 cells; 5 hearts) in control and 185 ± 4 pF (122 cells; 9 hearts) for cAVB.
In most of the experiments, the bulk changes in [Ca2+]i were buffered by inclusion of 1 mmol l–1 EGTA in the pipette solution. For some experiments, 80 µmol l–1 K5Fluo-3 was added to this pipette solution to report on the presence of Ca2+ release. In other experiments [Ca2+]i was not buffered, and we included 50 µmol l–1 K5Fluo-3 to a pipette solution without EGTA to record [Ca2+]i as previously described (Sipido et al. 2000), or loaded the cell with indo 1, through its ester form indo-1, AM (10 µmol l–1 for 10 min, followed by washing).
Solutions and experimental protocols
Cells were stored in normal Tyrode solution (mmol l–1): 137 NaCl, 5.4 KCl, 0.5 MgCl2, 1.8 CaCl2, 11.8 sodium Hepes and 10 glucose, pH 7.40. Whole-cell recording was established in this solution, but to measure Ca2+ currents we switched to a Na+- and K+-free solution containing (mmol l–1): 120 NMDGCl, 20 TEACl, 11 TEA-Hepes, 0.5 MgCl2, 1.8 CaCl2, 10 glucose, pH 7.4. The internal pipette solution contained (mmol l–1) 130 NMDGCl, 10 NMDG-Hepes, 0.5 MgCl2, 4 MgATP, 10 EGTA, pH 7.20. With these solutions there was a measured offset potential of 8–9 mV, true Vm being more positive than Vpipette. All potentials obtained in these solutions have been corrected in order to be able to compare data obtained in these conditions to data obtained in normal Tyrode solution.
Steady-state inactivation curves were constructed by normalizing the peak current at each test potential to the maximal current, and values were fitted with a Boltzmann equation: I/Imax
= (1 –
A)/{1 + exp[(V
–
V
)/k]}
+
A; where V is the potential of half-maximal inactivation, k is the slope factor, A is the amplitude of the non-inactivating Ca2+ current. Activation curves were derived from current–voltage relations and described by: I/Imax
= 1/{1 + exp[(V
–
V)/k]}. The dose–response curve for isoproterenol was fitted using a logistic equation of the form: I/I0
=
Imax
+ (1 –Imax)/{1 + (EC50/[iso])n}, with EC50 being the concentration of isoproterenol at 50% activation, and n being the Hill coefficient.
For action potential recordings, the pipette solution contained (mmol l–1): 120 potassium aspartate, 20 KCl, 10 potassium Hepes, 10 NaCl, 4 MgATP, 0.05 K5Fluo-3, pH 7.20, and the external solution was normal Tyrode. To measure Ca2+ currents in the absence of Ca2+ buffering, EGTA was omitted from the pipette solution and NMDG+ was replaced by 120 Cs+ and 20 TEA+; in the external solution, NMDG+ was replaced by Na+, and 200 µmol l–1 lidocaine (lignocaine) was added to block the Na+ current.
Lidocaine and Cd2+ were prepared as 200 mmol l–1 stock solutions in water; Ni2+ was made as a 2.5 mol l–1 stock solution. Caffeine (10 mmol l–1) was directly dissolved into the external solution. Niflumic acid (100 mmol l–1) and nifedipine (10 mmol l–1) stock solutions were prepared in DMSO and diluted 1:1000 before use. Isoproterenol was prepared as a 3 mmol l–1 stock in water with 1 mmol l–1 ascorbic acid added; solutions were made freshly for each day of experiments.
Statistics
Data are shown as means ± S.E.M. and they were compared using Student's t test or two-way ANOVA; P < 0.05 was considered significant.
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Results |
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The peak inward currents elicited from a holding voltage of –80 or –40 mV were identical, indicating there was no T-type current present, and values were not different between control (n = 15) and cAVB (n = 16) myocytes (Fig. 1). The current at the end of the 300 ms pulse (indicated by ‘x’) had similar amplitude in cAVB and control. The amplitude of this current was practically the same over the voltage range –20 to + 20 mV, i.e. the range in which the action potential plateau typically occurs.
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–22 ± 2, n
= 9, versus
–27 ± 1, n
= 10, in control; P < 0.05). The combined inactivation and activation curves are shown in Fig. 1C. The voltage range and amplitude of window currents was compared as the overlapping area under the activation and inactivation curves between –50 and 0 mV (inset), and expressed as arbitrary units (a.u.); it was 3.5 ± 0.2 a.u. in cAVB and significantly larger than in control cells, where it was 2.7 ± 0.2 a.u.; the maximum was at –17 mV in control and at –13 mV in cAVB. Faster inactivation of L-type Ca2+ current
Since we previously reported that the amplitude of sarcoplasmic reticulum Ca2+ release is larger in cAVB (Sipido et al. 2000), we measured the time course of inactivation of ICaL by fitting two exponentials to the decline of the current during a step to +10 mV. As illustrated in Fig. 2, the fast time constant was significantly shorter in cAVB. The amplitude of the fast declining current component was not significantly different for cAVB and control (71 ± 5% in cAVB versus 69 ± 3% in control).
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-adrenergic stimulation
We established a dose–response curve for -adrenergic increase of ICaL by washing in increasing concentrations of isoproterenol and measuring peak inward current during a test step to +10 mV. The peak current at each concentration was normalized to the baseline value. Figure 3A shows that the maximal increase at this potential was comparable for cAVB and control, with a Kd of 128 ± 38 nmol l–1 in control and 94 ± 13 nmol l–1 in cAVB (NS).
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When we examined the full range of the current–voltage relation from the two holding potentials at maximal stimulation by 3 µmol l–1 isoproterenol, we also found that the peak inward current was similar for the two holding voltages (Fig. 4A). A comparison between the I–V curves of cAVB (n = 12) and control (n = 10) yielded no statistical differences (ANOVA). However, the increase at the lower potentials was slightly less in cAVB (t test on data at –30 mV, P < 0.05). The I–V curves illustrate the leftward shift of activation. Steady-state inactivation curves were also shifted to the left and inactivation was incomplete even at 0 mV. Figure 4B shows the pooled data for steady-state inactivation and activation. The window area was much larger than in baseline conditions (Fig. 1C). The calculated surface area was 6.8 ± 0.8 (n = 7) in cAVB, not significantly different from the value in control cells (6.3 ± 0.4, n = 8). However the position of both activation and inactivation curves in cAVB was significantly more to the right resulting in a more positive maximum (–27 mV in cAVB, and –32 mV in control).
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Modulation of window current by SR Ca2+ release
To further study and quantify this postulated inactivation and recovery process, we used the voltage protocol illustrated in Fig. 6A. A step to 0 mV was introduced at varying time intervals after the step to –35 mV, to examine the availability of L-type Ca2+ channels. The evolution of the peak of the current at 0 mV then reflects the time course of inactivation and recovery. This time course tracked closely the ‘notch’ on the current trace at –35 mV. A quantitative analysis of the correlation between the peak current at 0 mV, normalized to its maximal value, and the corresponding current at –35 mV, also normalized to its maximal value, yielded a correlation coefficient of 0.94 ± 0.01, with P < 0.001 (illustrated in Fig. 6C). These data are consistent with our hypothesis that the ‘notch’ results from an inactivation and recovery process.
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We further examined the role of SR Ca2+ release by analysing the time course of the Ca2+ window current after suppressing SR Ca2+ release by continuous superfusion of the cells with 10 mmol l–1 caffeine (Fig. 6B). In the presence of caffeine there was no ‘notch’ on the current trace at –35 mV, and the test pulses to 0 mV showed a monotonous inactivation (n = 5). The inactivation at the end of the pulse was actually more pronounced than in the absence of caffeine, probably because of larger cytosolic Ca2+ accumulation during Ca2+ influx as there is no effective SR Ca2+ uptake.
Using this protocol on cells from cAVB (n = 10) and control (n = 9), we saw the typical inactivation and recovery in both groups. To quantify the extent of this modulation, we measured the ratio between the fully available current at 0 mV, Imax, and the minimal current at 25 ms, i.e. the extent of inactivation, and the ratio between Imax and the subsequent maximally recovered current. The difference between these two measurements then indicates the extent of dynamic modulation. As illustrated in Fig. 7, this was significantly larger in cAVB than in control myocytes (23 ± 2% of the initially available current, versus 13 ± 3% in control; P < 0.05).
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Our data on the current modulation so far were obtained in the presence of EGTA and indicated that it is the subsarcolemmal Ca2+ that drives this process. In a number of cells we confirmed that similar processes were present in the absence of Ca2+ buffering. Such experiments are more difficult to interpret as we can not work in Na+-free conditions and Na+ and Na+–Ca2+ exchange currents superimpose on Ca2+currents. To suppress the Na+ current we used a holding voltage of –65 mV and added 200 µM lidocaine. Figure 8A illustrates the inactivation/recovery time course of the current at –35 mV and its relation to the global [Ca2+]i transient in a cAVB myocyte. The current at –35 mV (trace indicated by arrow and labelled ‘a’) now did not have a ‘notch’ but had a transient inward current, presumably Na+/Ca2+ exchange current. The test pulses to 0 mV however, had a time course very similar as in the presence of EGTA with a clear inhibition at 20 ms, with subsequent partial recovery; the inhibition occurred at the same time as the inward current on the trace at –35 mV. The [Ca2+]i transient corresponding to the pulse to –35 mV (black trace, ‘a’) attained a peak value at 40 ms after the onset of the depolarization, slightly lagging behind the maximal inhibition of the current; the decline of the global [Ca2+]i transient was also slower than the recovery of the current as derived from the evolution of the peak current with the test pulse to 0 mV. This protocol was applied to four cAVB cells and five control cells, and the dynamic modulation of the Ca2+ current was analysed. As shown in Fig. 8B, without Ca2+ buffering, the extent of inactivation tended to be larger in cAVB with a significantly larger dynamic modulation than in control (25 ± 7% of the initially available current, versus 8 ± 2% in control; P < 0.05).
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Properties of EADs in cAVB
The experiments above do not examine directly the current underlying EADs in cAVB. However, the data support the potential and likelihood for Ca2+ current being involved in EADs under -adrenergic stimulation, as recovery from inactivation by SR Ca2+ release of the Ca2+ current could contribute a net depolarization and EAD. This hypothesis predicts that EADs are most likely to be observed in the voltage range of the window currents and at a time when the Ca2+ transient has declined substantially. We examined these predictions in two series of experiments. In myocytes studied with microelectrodes, EADs were observed in 4/13 cells in the presence of 300 nmol l–1 isoproterenol and the EAD take-off potential was –35 ± 4 mV, within the range of the maximal window current. In another group of cAVB myocytes we recorded action potentials and [Ca2+]i transients under whole-cell voltage clamp, in the presence of 1 µmol l–1 isoproterenol and stimulated at 0.5 Hz; EADs were observed in 4/10 cells. As illustrated in Fig. 9, in these conditions the [Ca2+]i transient had declined to near baseline values, even at plateau potentials, before the appearance of EADs (at 800–1200 ms after the upstroke of the action potential, [Ca2+]i had declined to 5–10% of its peak value; take-off potential was between –40 and –10 mV). In the example, the EAD was accompanied by a modest increase in [Ca2+]i, whereas the subsequent DAD was accompanied by a large spontaneous increase of [Ca2+]i. These observations are consistent with the concept that under -adrenergic stimulation window currents could contribute to EADs following recovery of ICaL from Ca2+-release-dependent inactivation.
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Discussion |
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Baseline amplitude of Ca2+ currents is unchanged in cAVB but window area is increased
We first established that there was no re-expression of T-type Ca2+ current as described in other models of cardiac remodelling in the dog (Dun et al. 2004).
We next focused on the properties of the L-type Ca2+ current and found that the amplitude of peak currents at baseline was unchanged in cAVB, but that the inactivation curve was shifted to the right and the area of window currents increased. A comparison to the literature shows that the whole-cell L-type Ca2+ current is unchanged in many models of hypertrophy (Tomaselli & Marban, 1999), but that in some models the current is increased, at least transiently (Balke & Shorofsky, 1998; Perrier et al. 2004), sometimes with larger window currents (Ryder et al. 1993). Such data do not necessarily imply that the expression and function of the channels is unchanged. In human heart failure an interesting story has unfolded, as single-channel recordings have indicated that the activity of the individual channels is increased (Schroder et al. 1998). Cheng et al. showed that PKA activation resulted in smaller maximal currents in failing myocytes, suggesting a higher phosphorylation level at baseline (Chen et al. 2002), which could compensate for either a lower number of channels, or a reduced open probability due to altered subunit expression (Hullin et al. 2003). In the present study, a higher level of baseline phosphorylation could help to explain why the whole-cell current is normal despite a downregulation of transcription of the 
subunit in the LV free wall (Ramakers et al. 2005). The somewhat smaller response of peak ICaL to isoproterenol in the negative voltage range and the reduced leftward shift of the activation and inactivation curves could be consistent with this hypothesis. This remains, however, speculative at present and needs to be further investigated.
This hypothesis also does not address why we see a rightward shift of the inactivation curve at baseline. We could not detect this in an earlier series of experiments, which examined a different population and a smaller sample (Sipido et al. 2000). Auxilliary subunits are known to affect the rate of inactivation, as well as levels of functional expression (Takahashi et al. 2003). At present little is known about changes in subunits with remodelling, but we plan to further investigate this potential mechanism.
Modulation of L-type Ca2+ current by Ca2+ release from the SR
Earlier studies have underscored the importance of the inactivation of L-type Ca2+ current by the high local concentration of Ca2+ during Ca2+ release from the SR (Sham et al. 1995; Sipido et al. 1995; Adachi Akahane et al. 1996) and the role of release-dependent inactivation of ICaL on the AP plateau and duration (Fauconnier et al. 2003; Takamatsu et al. 2003). The emphasis in the present study is on the recovery following inactivation and the consequent dynamic modulation of currents in the window voltage, an aspect that has received less attention before. In an earlier study we demonstrated the recovery by subtraction analysis of two pulses with different amplitudes of Ca2+ release (Sipido et al. 1995). Using the novel protocol of the present study we could follow the time course throughout a single pulse. A limitation of the current protocol is that we can apply it only for smaller depolarizing steps, but it is well suited to study the window current voltage range. A major advantage is that it can also be applied to examine inactivation and recovery with Na+ present in the external solution. The protocol clearly demonstrates that the time course of the net current in the presence of Na+ does not reveal the extent of inactivation and recovery of the Ca2+ channels due to superimposition of the Na+–Ca2+ exchange current.
With this approach we thus find a small but significant increase in the amplitude of the modulation of the Ca2+ current by SR Ca2+ release in cAVB. This distinguishes the remodelling in the dog with cAVB and compensated hypertrophy from the changes in the dog with heart failure where Ca2+ release from the SR is reduced and Ca2+ current inactivation is slower (O'Rourke et al. 1999; Winslow et al. 1999). The data are also consistent with a preserved Ca2+ removal process in cAVB, necessary to observe recovery of ICaL, again different from the reduced Ca2+ uptake into the SR observed in heart failure (reviewed in Hasenfuss & Pieske, 2002). The extent of inactivation and recovery is only slightly less in the presence of EGTA compared to without Ca2+ buffering, indicating that it results from a local process which is unaffected by buffering of bulk changes in [Ca2+]i and consistent with data from others (Sham et al. 1995; Adachi Akahane et al. 1996).
We did not formally study the relative contribution of Na+–Ca2+ exchange inward current compared to inward current resulting from recovery from inactivation of ICaL. Data in Fig. 8A, and published data on the time course of Na+–Ca2+ exchange during caffeine-induced Ca2+ release, support the hypothesis that these are consecutive events. On activation of Ca2+ release Na+–Ca2+ exchange inward current is large and ICaL is small because of inactivation; on Ca2+ reuptake into the SR, Na+–Ca2+ exchange current wanes and ICaL recovers. Inward Na+–Ca2+ exchange current and ICaL can thus both contribute to EADs, the former by a ‘conditioning phase’ of depolarization during Ca2+ release (Volders et al. 2000), followed by reactivation of ICaL.
Mechanisms of EADs and adrenergic response of Ca2+ current in cAVB
The small rightward shift of the inactivation curve in cAVB does not affect availability of the current at baseline, but can enhance availability during the plateau. It notably leads to a larger window for current re-activation, and this finding could be relevant for the EADs that occur in the presence of IKr block with dofetilide or almokalant (Volders et al. 1998). Another property of ICaL in cAVB that will facilitate EADs at plateau voltages is the small rightward shift of the maximal window current observed in the presence of -adrenergic stimulation.
The response of ICaL to -adrenergic stimulation is largely preserved, even if the peak amplitude at more negative potentials is somewhat smaller. On a background of reduced repolarizing current (Volders et al. 1999), in particular IKs which should shorten the action potential under -adrenergic stimulation (Varro et al. 2000; Stengl et al. 2003; Jost et al. 2005), the preserved ICaL is an important factor in tipping the balance towards action potential prolongation and EAD formation.
As recently demonstrated in an elegant modelling study (Tanskanen et al. 2005), the mode 2 gating of Ca2+ channels under -adrenergic stimulation and the stochastic nature of the channel behaviour can explain the occurrence of EADs within a typical pattern of variable duration of the action potential, as we also observed (Fig. 9). Our data add direct evidence for the recovery of the Ca2+ current during maintained depolarization. The extent of recovery we measure is actually larger than proposed in modelling; this can be due to local [Ca2+]i changes being larger than provided for in the modelling or because the model implements only a limited number of release units.
Though most of the work on the recovery in the present study was done at –35 mV, equally large sustained currents are present at more positive potentials (Fig. 4). We have previously shown that the modulation of ICaL by SR Ca2+ release is not voltage dependent (Sipido et al. 1995), consistent with the location of the Ca2+-binding site in the cytosolic carboxytail of the channel (Peterson et al. 2000). This implies that a similar extent of recovery can also occur at the more positive potentials.
In conclusion, in cAVB, the L-type Ca2+ current has a larger window; with -adrenergic stimulation it occurs at more positive voltages than in the control; the Ca2+ current has an enhanced dynamic modulation related to the large amplitude of SR Ca2+ release. These properties may contribute to the availability of Ca2+ channels and the increased incidence of EADs in cAVB under -adrenergic stimulation.
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