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Cardiovascular Research 2000 47(2):336-349; doi:10.1016/S0008-6363(00)00107-3
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Copyright © 2000, European Society of Cardiology

Facilitation of L-type calcium currents by diastolic depolarization in cardiac cells: impairment in heart failure

Stéphanie Barrère-Lemaire, Christophe Piot, Florence Leclercq, Joël Nargeot and Sylvain Richard*

Institut de Génétique Humaine, CNRS, UPR 1142, 141 Rue de la Cardonille, 34396 Montpellier Cedex 5, France

* Corresponding author. Tel.: +33-499-61-9939; fax: +33-499-61-9901 srichard{at}igh.cnrs.fr

Received 5 November 1999; accepted 19 April 2000


   Abstract
Top
 Abstract
1 Introduction
 2 Methods
 3 Results
 4 Discussion
 References
 
Objective: Decay kinetics of the voltage-gated L-type Ca2+ current (ICaL) control the magnitude of Ca2+ influx during the cardiac action potential. We investigated the influence of changes in diastolic membrane potential on ICaL decay kinetics in cardiac cells. Methods: Cells were isolated enzymatically from rat ventricles, human right atrial appendages obtained during corrective heart surgery and left ventricles from end-stage failing hearts of transplant recipients. The whole-cell patch-clamp technique was used to evoke ICaL by a 100-ms depolarizing test pulse to –10 mV. Conditioning potentials between –80 and 0 mV were applied for 5 s prior to the test pulse. Results: Depolarizing the cells between –80 and –50 mV prior to the test pulse slowed the early inactivation of ICaL both in rat ventricular and human atrial cells. This slowing resulted in a significant increase of Ca2+ influx. This type of facilitation was not observed when the sarcoplasmic reticulum (SR) Ca2+ content was depleted using ryanodine which reduced the rate of inactivation of ICaL, or when Ba2+ replaced Ca2+ as the permeating ion. Facilitation was favored by intracellular cAMP-promoting agents that, in addition to increasing current peak amplitude, enhanced the fast Ca2+-dependent inactivation of ICaL. Facilitation was impaired in atrial and ventricular human failing hearts. Conclusion: Decay kinetics of ICaL are regulated by the diastolic membrane potential in rat and human cardiomyocytes. This regulation, which associates slowing of ICaL inactivation with reduced SR Ca2+ release and underlies facilitation of Ca2+ channels activity, may have profound physiological relevance for catecholamines enhancement of Ca2+ influx. It is impaired in failing hearts, possibly due to lowered SR Ca2+ release.

KEYWORDS Adrenergic (ant)agonists; Ca-channel; Heart failure; Membrane potential; Myocytes; Serotonin (5HT)


   1 Introduction
Top
 Abstract
 1 Introduction
2 Methods
 3 Results
 4 Discussion
 References
 
The excitation–contraction coupling and contractile force of the myocardium depend upon initial activation of transmembrane L-type Ca2+ channels [1,2]. These channels are closed at negative resting membrane potentials and are gated primarily by membrane depolarization during the action potential. Their activity is also regulated by various transmitters, hormones and intracellular messengers. Best known, β-adrenergic stimulation increases intracellular cAMP which in turn activates protein kinase A and, ultimately, leads to increased open channel probability via a phosphorylation process [3,4]. Ca2+ channel antagonists, which bind to the Ca2+ channel protein and decrease its functional activity, and β-adrenergic blockers, which lower intracellular cAMP, represent two distinct classes of therapeutic agents used to decrease Ca2+ influx in the myocardium [4,5].

In addition to peak current amplitude generated by channel opening, the kinetics of inactivation of the L-type Ca2+ channels are important to determine the duration and, thereby, the amount of Ca2+ entry during membrane depolarization [2–5]. For example, an increase in the frequency of activation of Ca2+ channels induces an increase of Ca2+ entry resulting from a slowing of the decay of ICaL both in animal and human cardiomyocytes [6–15]. This mechanism, referred to as frequency-dependent facilitation, occurs over a wide range of frequencies (0.5–5 Hz) corresponding to heart rates encountered in pathophysiology and could be involved in the force–frequency relation of the healthy myocardium [13,14]. We have recently shown that moderate depolarization in the cell diastolic membrane potential also promotes facilitation of ICaL in rat ventricular cells [9,11]. In the present study, we investigated regulation of this process by β-adrenergic receptors stimulation and intracellular cAMP, a major regulatory pathway of cardiac function. In addition, we bring evidence for the presence of depolarization-induced facilitation in human cardiomyocytes. We also investigated this regulation in cells isolated from end-stage failing human hearts.


   2 Methods
Top
 Abstract
 1 Introduction
 2 Methods
3 Results
 4 Discussion
 References
 
2.1 Rat ventricular myocytes
The investigation conforms with the Guide for the Care and Use of Laboratory Animals published by the US National Institute of Health. Cells were enzymatically isolated from 6- to 10-week old Wistar Kyoto rats as described before [9,11]. Briefly, rats were anesthetized with 30 mg/kg of pentobarbital sodium injected intraperitoneally. The hearts were quickly removed and perfused retrogradely through the aorta (at 37°C) with a Ca2+-free solution containing (in mmol/l): NaCl (112), KCl (6), MgCl2 (2), NaHCO3 (4), KH2PO4 (1.5), HEPES (25), glucose (5.8), penicillin G (60 µg/ml), streptomycin (100 µg/ml) and 0.002% phenol red indicator (pH adjusted at 7.5 with KOH). This step was followed by 20-min perfusion of the same solution supplemented with Ca2+ (10 µmol/l) and 0.8 mg/ml type A collagenase (Boehringer Mannheim). Following perfusion, the ventricles were chopped and incubated in fresh enzymatic solution for 5 min (at 37°C). The tissue samples were rinsed with (in mmol/l): Choline Cl (119), KH2PO4 (1.5), MgCl2 (1.7), HEPES (25), glucose (5.8), succinate (5), pyruvic acid (10), creatine (5), and BSA (1 mg/ml), penicillin (60 µg/ml), and 0.002% phenol red indicator (pH adjusted to 7.4 with TEA-OH). The cells were then mechanically dispersed and stored in this solution at 4°C until electrophysiological recordings.

2.2 Human atrial and ventricular cells
Myocytes were isolated as described before [13,16–18]. Briefly, fragments of the right atrial appendage were obtained during open heart surgery (before cardio-pulmonary bypass) from ten patients aged 48 to 79 years in accordance with the institutional guidelines for human subject research of the University Hospital. The clinical diagnosis was aortic or mitral disease (stenosis or insufficiency) or coronary artery disease. Most patients had previously received medication comprising Ca2+ channel antagonists and/or β-adrenergic blockers and/or angiotensin-converting enzyme inhibitors until surgery. During anesthesia, all patients received a benzodiazepine, D-turbocurare, morphine, pentobarbital and antibiotics. Samples of the left ventricle and right atrium were obtained from transplant recipients (aged 44 to 56 years) with either ischemic (two patients) or dilated cardiomyopathy (one patient). These patients had end-stage heart failure (HF; New York Heart Association, functional class IV) with a severe alteration of the left ventricular function and an ejection fraction (EF) less than 20%. Their therapeutic treatment included only ACEI and diuretics. The procedures were approved by the Ethical Committee of the University Hospital. The solutions and enzymes used for transportation and dissociation have been detailed previously [13,16–18]. After the enzymatic procedure, the tissue was stored in the same solution as used for rat cells. The myocytes were dispersed by mechanical agitation in the recording chamber just prior to electrophysiological experiments. Only rod shaped, well relaxed and striated myocytes were subjected to experiments.

2.3 Electrophysiological recordings
ICaL was measured 2 to 10 h after cell dispersion using the whole-cell patch-clamp technique at room temperature (20–22°C). Conditions were optimized to eliminate contaminating voltage-gated inward Na+ and outward K+ currents [9,10,13,16]. Bath solutions contained (mmol/l): TEACl (130), CaCl2 (2), MgCl2 (1.1), 4-AP (4), HEPES (25), dextrose (22), and phenol red (17.7 mg/l); adjusted to pH 7.4 with TEAOH; 290–310 mOsm/l. To measure the waveforms of Ba2+ currents through Ca2+ channels in the same conditions, 2 mM BaCl2 replaced CaCl2. Recording pipettes contained (in mmol/l): CsCl (130), EGTA (10), HEPES (25), Mg-ATP (3), Mg-GTP (0.4); adjusted to pH=7.4 with CsOH; the osmolarity of the buffer was 290–310 mOsm/l. Junction potentials between the intra-pipette solution and the reference electrode were cancelled before obtaining the tight seals. Experiments were performed using large low-resistance pipettes (<3 M{Omega} when filled with the recording solution). After seal formation (resistance ranging between 1 and 20 G{Omega}) and membrane disruption, residual series resistance after electronic compensation were ≤1.2 M{Omega} and could not introduce major errors of voltage clamp (<4 mV) for currents <2 nA. Whole cell membrane capacitances, determined by integrating the capacitive current recorded during a 5 mV voltage step from a HP of –80 mV, ranged between 60 and 200 pF. The voltage-clamp circuit was provided by an Axopatch 200A patch-clamp amplifier. Experimental parameters, such as holding potentials (HPs) and test potentials were controlled with an IBM PC connected through a Tecmar Labmaster analog interface (Axon instruments, Burlingame, CA, USA) to the electrophysiological equipment. Data acquisition and analyses were performed using the PCLAMP software (Axon instruments). Signals were filtered at 3–5 kHz prior to digitization and storage. ICaL was recorded at a test pulse of –10 mV (100 ms duration) delivered from a HP of –80 mV. A voltage-ramp protocol was also used in some cells. No T-type ICa was evident in human atrial and ventricular cells, as well as in rat ventricular myocytes [9,13,16,18]. Although a tetrodotoxin-sensitive low threshold ICa flowing through Na+ channels was recorded sometimes, this current had no significant contribution here because it is observed only from very negative HPs (it is 95% inactivated at –80 mV) and it is small at a test pulse of –10 mV [18].

2.4 Analysis of ICaL
The effects of conditioning membrane potentials on ICaL were examined using the paradigm illustrated in Fig. 1A. Peak ICaL were measured as the difference between the maximal inward current amplitude at –10 and the zero current level. The Ca2+ entry was quantified by integrating ICaL (pA·ms) during the 100-ms duration test pulse, rather than measuring only peak current. The decay of ICaL was also measured. Inactivation was best fitted by the sum of two exponential components using the formula

Formula
where, ICaL(fc) and ICaL(sc) are respectively the current amplitudes of the fast and slow components of ICaL and {tau}(fc) and {tau}(sc) are respectively their related time constants of inactivation. This procedure has been described in detail previously [9,11]. The zero time was set slightly before peak currents to determine ICaL(fc) and ICaL(sc) and, in all cases, the sum of ICaL(fc)+ICaL(sc) accounted for peak ICaL.


Figure 1
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  		Fig. 1 Modulation of ICaL by conditioning potentials in rat ventricular myocytes. (A) Experimental protocol: ICaL were evoked at –10 mV following 5-s conditioning pulses to various potentials. (B) Typical waveforms of ICaL evoked from –80 and –50 mV. Note the change in the waveform of ICaL. In the inset, currents have been scaled for direct comparison of their kinetics. (C) Complete representative relationship between normalized Ca2+ entry (quantified by integrating ICaL (pA·ms) during the test pulse: see Methods) and conditioning voltages in the same cell as in (B). Note the graded increase between –80 and –40 mV.

 
2.5 Statistical analysis
All averaged or normalized data are presented as mean ±S.E.M. The significance between groups of data was assessed using Student's t test (for paired and unpaired samples as appropriate). Results were considered not significant (ns) with P>0.05, significant (*) with 0.01<P<0.05, very significant (**) with 0.001<P<0.01 and extremely significant (***) with P<0.001. Box and whiskers representation was also used as noted (Fig. 7). The box extends from the 25th percentile to the 75th percentile, with a horizontal line at the median (50th percentile). Whiskers extend down to the smallest value and up to the largest (showing the range of data).


Figure 7
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  		Fig. 7 Link of the depolarization-induced facilitation to cAMP and fast decay kinetics of ICaL in human atrial cells. (A) The decay of ICaL was best fitted by two time constants. The box and whiskers representation (see Methods) shows that the fast and slow time constants are separated by an order of magnitude. Analysis was performed on 48 cells (***P<0.001). (B) Illustration of the preferential increase of ICaL(fc) (fc for fast component; sc for slow component) by db-cAMP (100 µmol/l). The time constants {tau}(fc) and {tau}(sc) were respectively 5 and 110 ms (for both traces). (C) Preferential increase of ICaL(fc) by Iso (2 µmol/l). (a) Illustration for a single cell. {tau}(fc) and {tau}(sc) were respectively 5.3 and 70 ms (for both traces). (b) Box and whiskers representation of the effect of Iso on the contribution of ICaL(fc) to ICaL. Analysis was performed on 30 cells (***P<0.001). (D), (a) Depolarization-induced interconversion between ICaL(fc) and ICaL(sc). The experiment was performed using the protocol shown in the inset in the presence of 1 µM Iso. The conditioning potential at –80 mV induced preferential activation of ICaL(fc) whereas the conditioning potential at –50 mV promoted mainly activation of ICaL(sc). The observed voltage-dependent changes in ICaL(fc) and ICaL(fc) were reciprocal. {tau}(fc) and {tau}(sc) were fixed at respectively 8 and 120 ms (for both traces). (b) Box and whiskers representation of the contribution of ICaL(fc) to global ICaL in cells exhibiting facilitation (F) or no facilitation (NF) either in the presence or absence of Iso. Analysis was performed on 20 and 25 cells respectively. Results were extremely significant (***P<0.001).

 
2.6 Solutions
Isoproterenol (Iso), serotonin, and dibutyryl-cAMP (db-cAMP) (Sigma) and ryanodine (Biomol) were prepared as 1 or 10 mmol/l stock solutions in distilled H2O and subsequently diluted at the desired working concentrations in test solutions. The control and test solutions were applied by using a multiple capillary perfusion system (200 µm inner diameter, flow-rate 0.5 ml/min) placed in close proximity of the cell (<0.5 mm). Each capillary is fed by a reservoir 50 cm above the bath. Rapid and complete solution changes were made by switching the opening from one capillary to the next.


   3 Results
Top
 Abstract
 1 Introduction
 2 Methods
 3 Results
4 Discussion
 References
 
3.1 Depolarization-induced facilitation of ICaL in rat ventricular cells
To study the amplitude and the waveforms of ICaL at various resting membrane potentials, rat ventricular cells were voltage clamped at HP –80 mV, and subsequently depolarized to various conditioning potentials between –80 and –10 mV for 5 s prior to voltage steps to –10 mV (which activates maximal current amplitude) for 100 ms in duration (Fig. 1A). The 5-s conditioning pulse duration was selected to achieve steady-state conditions since no further changes in ICaL waveforms were evident during conditioning pulses of longer duration. When this protocol was applied, a facilitation of ICaL was induced by conditioning prepulses of moderate amplitude (Fig. 1B) in 88% of the cells (n=24) leading to a substantial increase of Ca2+ influx as measured by integrating Ca2+ entry (see Methods) during the test depolarization (Fig. 1C). This increase resulted mainly from the slowing of ICaL decay (Fig. 1B). On average, the increase in Ca2+ entry promoted by depolarizing the conditioning potential from –80 to –50 mV was 39±2%. In most cells this depolarization-induced facilitation of Ca2+ channel activity was still marked at –40 mV. Further depolarization to voltages less negative than –40 mV resulted in clear voltage-dependent inactivation of Ca2+ entry due to the decreased availability of the Ca2+ channels for opening [4]. No further change of the decay of ICaL was observed at these voltages (data not shown). However, 12% of the cells lacked the voltage-dependent facilitation of ICaL which could be related to a distinct physiological state of the cells (e.g. phosphorylation level).

3.2 Modulation by Isoproterenol and dibutyryl-cAMP
We investigated whether the depolarization-induced facilitation of ICaL is modulated by β-adrenergic receptor stimulation in rat ventricular cells. This model appeared suitable because facilitation was observed in nearly all cells. Extracellular application of Iso, a β-adrenergic agonist, increased the amplitude of ICaL evoked at –10 mV by 130±40% (n=8) when applied at its maximally effective concentration (2 µmol/l). ICaL peak amplitude slowly increased and reached a plateau within 30 s to 40 s which reflected the time required to achieve steady-state phosphorylation of Ca2+ channels by the activated catalytic subunit of protein kinase A [3,4]. We found that Iso could enhance (when present in control conditions: Fig. 2Aa) or even unmask (when absent: Fig. 2Ab) the depolarization-induced facilitation of ICaL in seven out of eight cells. The increase of Ca2+ entry induced by depolarizing the cells from –80 to –50 mV during the conditioning prepulse was enhanced by a factor 2 (87±9%) after maximal stimulation of the Ca2+ channels by Iso. This enhancement was observed at all test pulses activating ICaL; that is, it did not reflect a shift in the current–voltage relationship as observed using a ramp protocol (Fig. 2B). Fig. 2B also shows that the larger depolarization-induced facilitation of ICaL in the presence of Iso (compare vertical bars) occurs at all voltages of the current–voltage relationship. Since enhancement of the activity of cardiac Ca2+ channels following β-adrenergic receptor stimulation occurs mainly via intracellular cAMP production [2–5], we assessed the direct activation of protein kinase A by cAMP. Addition of 100 µmol/l of the membrane permeable analog db-cAMP to the extracellular perfusion solution could mimic the effect of Iso and enhanced facilitation by a factor 2 (85±16%; n=4). This value was very similar to that obtained for Iso which suggested that cAMP mediates most of the effect of Iso.


Figure 2
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  		Fig. 2 Enhancement of depolarization-induced facilitation of ICaL by Iso in rat ventricular myocytes. (A), (a) The depolarization-induced facilitation of ICaL, present in control conditions (squares), was enhanced during exposure to Iso (2 µmol/l: circles). Ca2+ entry was quantified by integrating ICaL (pA·ms) during the test pulse. (b) Same experiment as in (a) but facilitation was absent in control. It was enhanced by Iso (2 µmol/l). Typical waveforms of ICaL evoked from –80 and –50 mV are shown. (B) Same experiment as in (A) but ICaL was evoked using a voltage-ramp protocol delivered from –80, –60, –40 mV as shown in the left panel. In this myocyte, facilitation was present prior to exposure to the drug. Note the voltage-dependent facilitation at all voltages. Note also the greater facilitation in the presence of Iso and the larger effect of Iso at –40 mV (e.g., compare bars between –80 and –40 mV).

 
3.3 Link with Ca2+-dependent inactivation
Depolarization-dependent facilitation reflects a reduction in the fast decay kinetics of ICaL. In various cardiac cells, including human, there is a joint dependence of the inactivation of ICaL on both voltage and intracellular Ca2+ [2,4,16,19,20]. In this context, ICaL-triggered Ca2+ release from the SR is an important feed-back signal in determining the rate of inactivation of the L-type Ca2+ channel [20–24]. To examine whether the depolarization-induced change in the decay kinetics of ICaL depends on the amount of Ca2+ released from the SR, we used ryanodine to deplete the SR Ca2+ content and tested the effects. The results showed that ryanodine reduces the rate of inactivation of ICaL (Fig. 3A). In these conditions, no facilitation could be induced following conditioning depolarizations (Fig. 3B). The increase of Ca2+ entry that is normally induced by depolarizing the cells from –80 to –50 mV was abolished in all of five cells tested. The slow decay of ICaL was the limiting factor. On average, the potentiation of ICaL observed during facilitation decreased from 42±7% to 8±1% after ryanodine (***P<0.001). Therefore, we concluded that depolarization-dependent facilitation involves SR Ca2+ release-induced inactivation of ICaL.


Figure 3
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  		Fig. 3 Effect of ryanodine on voltage-dependent facilitation. (A) Typical waveform of ICaL evoked at –10 mV from a HP of –80 mV in control conditions (control) and after extracellular exposure to 1 µM/l ryanodine. Maximal effect was generally achieved after 10 min. (B) The depolarization induced facilitation of ICaL, present in control conditions (a) was markedly abolished after steady-state effect of ryanodine (b). Typical waveforms of ICaL evoked at –10 from –80 and –50 are shown. (C) Bar graphs show the averaged decrease (mean±S.E.M., n=5) of ICaL facilitation after ryanodine (1 µM/l). The effect was extremely significant (***P<0.001).

 
We next examined the effect of replacing extracellular Ca2+ by Ba2+, which slows the decay of Ca2+ channel current, on facilitation to assess the effect of cAMP independently of Ca2+. In these conditions, Ca2+-dependent inactivation of Ca2+ channel current is completely removed and only purely voltage-dependent inactivation occurs [2,4,9,19]. When the conditioning potential was gradually changed from –80 to –50 mV prior to the test pulse, no increase of current peak amplitude was observed (Fig. 4A, left panel; Fig. 4B). In fact, the peak current decreased gradually and the β-adrenergic receptor stimulation by Iso enhanced this decrease (Fig. 4A, right panel; Fig. 4B) which is related to the leftward shift of the steady-state inactivation curve described before [25]. For example, the percentage of decrease observed for a conditioning prepulse of –50 mV (vs. –80 mV) was enhanced from 14±2 to 23±5% (*P<0.02; n=9). These results contrasted markedly with the large increase obtained using Ca2+ as the charge carrier (Fig. 1). Furthermore, analyzing the variations in integrated Ca2+ and Ba2+ entry during the test pulse confirmed that there is no marked increase of Ba2+ currents between –80 and –50 mV (although a small slowing of the overall current decay may occur) and that Iso induces a large decrease (Fig. 4C). These results suggested therefore that facilitation of Ca2+ channel activity does not occur independently of Ca2+and does not reflect a direct effect of cAMP-dependent phosphorylation on the channels.


Figure 4
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  		Fig. 4 Effect of conditioning potentials on Ba2+ currents in rat ventricular myocytes. (A) Typical waveforms of Ba2+ currents evoked at –10 mV from various conditioning voltages between –80 and –30 mV in the absence (left panel) and presence (right panel) of Iso (1 µmol/l). Note the decrease in current peak amplitude with little change in the decay kinetics. (B) Bar graphs show the averaged decrease (mean ±S.E.M.) of Ba2+ currents peak amplitude for various conditioning prepulses between –80 and –50 mV in the absence and presence of Iso. The significance is quoted as defined in Methods for each test. (C) Comparison of the variations of Ca2+ and Ba2+ entry (quantified by integrating currents (pA·ms) during the test pulse at –10 mV: see Methods) evoked from conditioning depolarizations at –80 and –50 mV in the absence and presence of Iso. Note the variations in opposite directions.

 
3.4 Depolarization-induced facilitation of ICaL in human atrial cells
Another objective of this study was to investigate whether the depolarization-induced facilitation of ICaL could be observed in human cardiomyocytes. In human atrial cells, ICaL started to activate between –40 and –30 and reached maximal peak amplitude between –10 and 0 mV as described before [5,16,17,20]. To study the waveforms of ICaL at various conditioning diastolic membrane potentials, cells were voltage-clamped using the protocol employed for rat ventricular cells with a test pulse at –10 mV (Fig. 1A). Two types of responses were observed. In some cells, Ca2+ influx was increased following moderate pre-depolarizations (Fig. 5A) which resulted mainly from the slowing of current decay. As described in rat cardiomyocytes, the increase was graded with increasing conditioning prepulses between –80 and –40 mV (Fig. 5A). This behavior, observed in 13 cells, could lead to a substantial increase of Ca2+ entry (up to 50%) during the test pulse. In this group, the averaged increase was 22±5% (between –80 and –50 mV). The Ca2+ entry decreased as the conditioning prepulse was further depolarized from –40 to 0 mV, due to voltage-dependent steady-state inactivation of Ca2+ channels (Fig. 5A). On the other hand, no significant facilitation occurred in 18 other cells in which only the voltage-dependent decrease of Ca2+ channels availability for opening was evident between –50 and –10 mV (Fig. 5B). This cell population was characterized by slow inactivation kinetics as illustrated in the inset of Fig. 5B. The superimposition of the two curves suggests that the facilitation phenomenon overlaps with the classical steady-state inactivation of ICaL (Fig. 5B). We found no evident morphological difference between cells that exhibited and cells that did not exhibit facilitation.


Figure 5
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  		Fig. 5 Modulation of ICaL by conditioning potentials in human atrial myocytes. ICaL were evoked as shown in Fig. 1A. (A) Cell illustrating depolarization-induced increase of Ca2+ entry through Ca2+ channels in a human atrial cell. Ca2+ entry was quantified by integrating ICaL (pA·ms) during the test pulse and normalized. Inset: typical waveforms of ICaL evoked to –10 mV from –80 and –50 mV. (B) Same experiment in another cell with no increase of Ca2+ entry. In inset: typical waveforms of ICaL evoked to –10 mV from –80 and –50 mV in this cell population. The superimposed dashed curve was taken from (A).

 
All human atrial myocytes exhibiting the depolarization-induced facilitation of ICaL also exhibited the high-frequency induced facilitation (Fig. 6Aa,b) which we described previously [13,14]. As a corollary, none of the cells lacking the frequency-dependent facilitation exhibited the voltage-dependent facilitation. The two phenomena were not additive; i.e. there was no high frequency-induced facilitation of ICaL when the cells were held at a diastolic membrane potential of –50 mV because, as described for rat ventricular cells before [11], fast inactivation of ICaL was already greatly reduced at a HP of –50 mV (data not shown). Furthermore, as shown above for rat ventricular cells, both types of facilitation were blunted using Ba2+ as the charge carrier through the Ca2+ channels (n=5; data not shown) because the Ca2+-dependent fast decay of the current was dramatically reduced. Finally, as also observed in rat ventricular cells, cAMP-promoting agents enhanced both types of facilitation. This is illustrated (Fig. 6Aa,b) for serotonin (1 µmol/l) which activates the cAMP cascade via the 5HT4-receptor in human atrial myocytes [17]. The membrane permeable analog of cAMP, db-cAMP (100 µmol/l) had similar effects (Fig. 6B). Iso had, on average, more discrete effects (Fig. 6C) owing to some variability among cells (two cells lacked facilitation even after Iso) which contrasted with the results obtained in rat ventricular cells. Such variability may result from pathology or drug treatment and, possibly, reflect a down-regulation of β-adrenergic receptors [26,27].


Figure 6
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  		Fig. 6 Modulation of the depolarization-induced facilitation of ICaL by the cAMP pathway in human atrial cells. (A), (a) The depolarization-induced facilitation of ICaL, present in control conditions (squares), was markedly enhanced during exposure to serotonin (1 µmol/l) in a human atrial myocyte. Typical waveforms of ICaL evoked from –80 and –50 mV are shown. In inset (right panel): complete relationship between Ca2+ entry and conditioning voltages in the same cell. (b) Enhancement of the high-frequency induced facilitation of ICaL by serotonin in the same cell. The rate of stimulation was increased from 0.1 to 1 Hz. Steady-state was obtained at the fourth stimulation (i.e., within 3 s). In inset (right panel): bars compare Ca2+ entry at 0.1 Hz (white bars) and 1 Hz (black bars) in the absence and presence of serotonin. Note the enhancement by serotonin. (B) Enhancement of the depolarization-induced facilitation of ICaL by db-cAMP (100 µmol/l). (C) Bar graphs show averaged enhancements (mean ±S.E.M.) of depolarization-induced facilitation of ICaL by Iso (2 µmol/l), db-cAMP (100 µmol/l) and serotonin (1 µmol/l). The number of experiments for each bar is quoted. Significant statistical differences are indicated as *P<0.03.

 
We next investigated the effect of cAMP-promoting agents in further details. Facilitation was clearly linked to the presence of the fast inactivating component of ICaL (Figs. 2Ab and 5AGo). The decay of macroscopic ICaL was indeed best described by the sum of two exponentials with time constants separated by an order of magnitude (Fig. 7A). They varied between 4 and 17 ms (mean 9±1 ms) for the fast one and between 37 and 200 ms (mean 92±7 ms) for the slow one at a test depolarization of –10 mV as determined in a large population of cells (n=48). These time constants underlay two kinetically distinct current components that we will refer to as ICaL(fc) and ICaL(sc). The relative contribution of each component determines the global waveform of ICaL and varies among cells. A double exponential fit to the decay phases of ICaL evoked from each conditioning potential tested provided the amplitudes of ICaL(fc) and ICaL(sc) in each cell. We found that both db-cAMP and Iso, when applied at low rates of stimulation and from HP –80 mV, promoted preferentially ICaL(fc) (Fig. 7B,C). On average, Iso increased the contribution of ICaL(fc) to the global ICaL from 22±4% to 57±4% (n=30; ***P<0.001). When facilitation was observed, it was related to reduced fast inactivation of ICaL rather than to increased peak current amplitude. This change reflected modifications in the amplitudes of ICaL(fc) and ICaL(sc) with no major changes in {tau}(fc) and {tau}(sc). The double exponential fits to the decay phases of ICaL revealed that changing the conditioning voltage from –80 to –50 mV resulted in a reduction in the amplitude of ICaL(fc) and a corresponding increase in the amplitude of ICaL(sc). Since there is only a minor change on global peak ICaL amplitude (Fig. 7Da), the changes in ICaL(fc) and ICaL(sc) amplitudes are, therefore, reciprocal and reflect only a change in the inactivation process of the same population of Ca2+ channels. ICaL(fc) was predominant (75±3% of total ICaL, n=20) only in the fraction of cells exhibiting facilitation (Fig. 7Db). Similar analysis of facilitation in rat ventricular cells led to similar conclusion [9,11]. It should be noted that, in contrast, the inactivation of Ba2+ currents and Ca2+ currents in the presence of ryanodine could be best fitted by a single exponential corresponding to the slower component of ICaL (data not shown).

3.5 Facilitation in failing hearts
We investigated facilitation in myocytes isolated from failing hearts of transplant recipients. These patients had drug treatment limited to ACE inhibitors and to diuretics (no Ca2+ channel antagonist, no β-adrenergic blocker). Fig. 8A shows that facilitation was impaired in most atrial myocytes isolated from end-stage HF patients with a low ejection fraction (<20%) as compared to cells isolated from the non-HF patients with an ejection fraction >40%. On average, facilitation decreased from 10±2% (n=31) to –2±3% (n=11; ***P<0.001). Both the β-adrenergic- and the 5HT4-receptors stimulations were inefficient to induce significant facilitation (ns, P>0.05) though large effects could be observed in a few individual cells. Facilitation was also almost absent in HF ventricular cells (Fig. 8B). Although Iso was able to promote very substantial facilitation (>50%) in three cells, the averaged effect was below significance at the cell population level. These findings contrasted markedly with observations made in rat ventricular cells in which Iso enhanced facilitation. We determined that, the slow component of inactivation ICaL(sc) was the predominant component of ICaL in human HF cells lacking facilitation (data not shown). Therefore, facilitation is not observed in HF mainly because the decay kinetics of ICaL are slower than in normal cells.


Figure 8
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  		Fig. 8 Impairment of the depolarization-induced facilitation of ICaL in human heart failure. (A) Bar graphs show averaged (mean±S.E.M.) facilitation of ICaL induced by a 5-s conditioning prepulse from –80 to –50 mV (as described in Fig. 1A) in human atrial cells. Increase or decrease of ICaL observed in cells isolated from failing hearts (HF) were compared to that observed in non-failing hearts (non-HF). Iso (1 µmol/l) and serotonin (1 µmol/l) did not unmask a significant facilitation of ICaL in HF cells. Significant statistical differences is indicated as ***P<0.001. (B) Bar graphs show averaged (mean±S.E.M.) facilitation of ICaL induced by a 5-s conditioning prepulse from –80 to –50 mV in human ventricular cells from failing hearts. Iso (1 µmol/l) had no significant effect.

 

   4 Discussion
Top
 Abstract
 1 Introduction
 2 Methods
 3 Results
 4 Discussion
References
 
In the present study, we emphasize that variations of the diastolic membrane potential at physiological range regulate the decay kinetics of ICaL and thereby, control transmembrane Ca2+ entry into cardiomyocytes. The new findings are: (i) depolarization-dependent regulation of ICaL decay kinetics occurs not only in rat but also in human cardiomyocytes; (ii) it is the basis of the phenomenon referred to as facilitation; (iii) facilitation reflects removal of the SR Ca2+ release-induced inactivation of ICaL during depolarization and is favored by intracellular cAMP, presumably via enhanced SR Ca2+ load; and, (iv) facilitation is defective in cardiomyocytes isolated from end-stage failing human hearts.

4.1 Depolarization-induced facilitation of ICaL
A well-known effect of conditioning depolarizations between –50 and 0 mV is to decrease peak ICaL amplitude due to a voltage-dependent decrease in the availability for opening of the Ca2+ channels [4]. However, conditioning depolarizations between –80 and –50 mV do not decrease but instead increase gradually Ca2+ influx during the test depolarization. This type of facilitation evidenced initially in rat ventricular cells [9,11,28] is shown here for the first time in human cardiomyocytes. As described before for the high-frequency induced facilitation [6–15], it is related mainly to a marked slowing of ICaL decay kinetics. The slight increase of ICaL peak amplitude may result only from slower inactivation rather than from an increase in the number of Ca2+ channels available for opening [11]. It has been widely reported that facilitation reflects a genuine effect on Ca2+ channels [6–15,29]. Facilitation is associated with an inward current both at negative and positive voltages [29]. It seems thus unlikely that fast Ca2+-activated chloride currents have a contribution because these currents should be in opposite directions (inward at negative voltages and outward at positive potentials) due to the use of symmetrical chloride concentrations [30].

4.2 Links with intracellular cAMP and Ca2+
The early fast decay of ICaL and its related component ICaL(fc) are determined by a Ca2+ release-induced inactivation in a microdomain inaccessible to EGTA and not by its own influx [2,20–24]. This local Ca2+ signalling can contribute to 65–75% of inactivation [22,23] and has been demonstrated using ryanodine or thapsigargin which both abolish Ca2+ release from the SR or, yet, phospholamban-deficient mice [15,20,22–24]. Our result showed that when the SR-Ca2+ release and thereby the fast Ca2+-dependent inactivation of ICaL are reduced by ryanodine, no depolarization-induced facilitation occurs. Therefore, the slow inactivation of ICaL which underlies the depolarization-induced facilitation, involves reduced Ca2+ release-induced inactivation as demonstrated for frequency-dependent facilitation [7,8,15].

The primary effect of cAMP is to increase peak ICaL due to both an increase in probability of Ca2+ channel opening as well as an increase in the number of functional Ca2+ channels [2–4]. We show here that another effect of cAMP is to enhance the depolarization-induced facilitation of ICaL. Our detailed analysis reveals that facilitation occurs because the fraction of current exhibiting fast Ca2+-dependent inactivation (ICaL(fc)) is enhanced by cAMP. Therefore, there is a greater proportion of channels that can switch from fast to slow inactivation, with ICaL(sc) becoming the major current component during facilitation. Consistently, when the Ca2+-dependent inactivation is abolished using Ba2+ as the permeating ion, β-adrenergic receptors stimulation is unable to promote facilitation. Thus, we conclude that the effect of cAMP-dependent phosphorylation on facilitation is not direct and is related to enhanced Ca2+-dependent inactivation of ICaL resulting from stimulated Ca2+ load of the SR.

In contrast with the high-frequency induced facilitation, the depolarization-induced facilitation occurs in a range of conditioning voltages considered too negative (<–50 mV) to activate significant macroscopic ICaL. Two mechanisms could account for this intriguing phenomenon. Voltage-driven Ca2+ release from the SR [31] or alternatively, Ca2+ entry via a very small number of Ca2+ channels with a high driving force for Ca2+ can generate large localized Ca2+ concentrations and, hence, SR-Ca2+ release. Interestingly, cAMP-dependent phosphorylation induces a leftward shift of the voltage threshold required for channel activation and may favor Ca2+ entry at potentials as negative as –60 mV [25,32]. The precise mechanism(s) determining the coupling between the conditioning depolarization and Ca2+ release-induced inactivation of ICaL will be worth studying in the future.

4.3 Possible physiological relevance
Another form of depolarization-induced facilitation of ICaL has been described in cardiac cells. In this case, strong conditioning depolarizations (>40 mV) increase Ca2+ channels activity when Ba2+ is the permeating cation [25,33]. In contrast, facilitation here only requires moderate depolarization but Ca2+ is mandatory. The slowing of ICaL inactivation associated with reduced SR Ca2+ release, may supply additional Ca2+ ions for contraction and loading of the SR [34,35]. It may be an adaptive mechanism to support the increase in cardiac contraction during exercise and stress in response to catecholamines release [36]. For example, β-adrenergic receptors stimulation has been reported to induce cell depolarization in guinea-pig papillary muscles and sheep Purkinje fibers [37–39]. Activation of chloride currents and ‘pacemaker’ current If is also expected to induce cell depolarization [40,41], thus favouring ICaL facilitation. However, any pathological situation generating ectopic beats and cell depolarization may contribute to the arrhythmogenicity of cAMP promoting agents, and result in deleterious Ca2+ overload. For example, this latter process may participate in the generation of ventricular arrhythmias during acute myocardial infarction and ischemia [42,43] that cause both catecholamines release and cell depolarization.

4.4 Impairment in HF
We found that the depolarization-induced facilitation is variable in human atrial cells which may be related to the underlying pathology. Studies in human cardiomyocytes are often complicated because access to normal tissue is rare. We did not analyze the influence of various diseases (aortic or mitral stenosis or insufficiency, coronary artery disease, atrial fibrillation) and treatments because a valuable statistical analysis would require a very large number of patients and of cells per patient. However, we found that the depolarization-induced facilitation is blunted in most atrial and ventricular myocytes isolated from end-stage HF patients. A similar conclusion was reached before for frequency-dependent facilitation [5,13,14]. Several mechanisms may contribute to impair facilitation of ICaL in HF. The Ca2+ channels may be changed in terms of structure (e.g., nature or stoichiometry of associated subunits) and, therefore, in their gating properties. An alternative possibility is impaired regulation of normal channels. In particular, HF is characterized by an alteration of [Ca2+]i handling [44,45]. The systolic [Ca2+]i transients are reduced owing to crucial defective Ca2+ re-uptake in the SR which results into a decrease in the force of contraction [46,47]. An increased phospholamban inhibition of Ca2+ uptake by the SR-Ca2+-ATPase activity is likely to reduce the negative feed-back of the Ca2+ released from a poorly loaded SR on Ca2+ channels [24]. This failure is likely to explain the slow decay kinetics of ICaL that we observed here in HF myocytes. This may be consistent with the recent single channel data showing that L-type Ca2+ channels from failing hearts remain available for a longer time during depolarization [48]. Lower intracellular cAMP, resulting from the down-regulation of β-adrenergic receptors [22,23] or from an increase in G-protein inhibition [49], may also contribute to lessening of current inactivation and lack of facilitation due to inefficiency to stimulate the SR-Ca2+-ATPase activity [24].

In conclusion, we show that facilitation of L-type Ca2+ channels activity can be induced by moderate depolarization of the diastolic membrane potential in cardiomyocytes. This process may have profound physiological relevance for catecholamines enhancement of Ca2+ influx. Facilitation is impaired in human HF, possibly due to both low intracellular cAMP (with low opening probability of the Ca2+ channel) and altered re-uptake of intracellular Ca2+ by the SR which decreases the negative feed-back by the Ca2+ released from the SR on Ca2+-dependent inactivation of Ca2+ channels.

Time for primary review 24 days.


   Acknowledgements
 
We thank Matteo Elia Mangoni, Steve Dubel and Govindan Dayanithi (Montpellier, F) and Jamie Vandenberg (Cambridge, UK) for improving the manuscript. This work was supported by grants from the ‘Association Recherche et Partage’ (to SR), the ‘Association Française contre les Myopathies’ and laboratoires Servier (SBL).


   References
Top
 Abstract
 1 Introduction
 2 Methods
 3 Results
 4 Discussion
 References
 

  1. Fabiato A. Stimulated calcium current can both cause calcium loading in and trigger calcium release from the sarcoplasmic reticulum of a skinned cardiac Purkinje cell. J. Gen. Physiol. (1985) 85:291–320.[Abstract/Free Full Text]
  2. Bers D.M., Perez-Reyes E.W. Ca channels in cardiac myocytes: structure and function in Ca influx and intracellular Ca release. Cardiovasc. Res. (1999) 42:339–360.[Abstract/Free Full Text]
  3. Tsien R.W., Bean B.P., Hess P., Lansman J.B., Nilius B., Nowycky M.C. Mechanisms of calcium channel modulation by β-adrenergic agents and dihydropyridine calcium agonists. J. Mol. Cell Cardiol. (1986) 18:691–710.[ISI][Medline]
  4. McDonald T.F., Pelzer S., Trautwein W., Pelzer D. Regulation and modulation of calcium channels in cardiac, skeletal, and smooth muscle cells. Physiol. Rev. (1994) 74(2):365–507.[Free Full Text]
  5. Richard S., Leclercq F., Lemaire S., Piot C., Nargeot J. Ca2+ currents in compensated hypertrophy and heart failure. Cardiovasc. Res. (1988) 37(2):300–311.[CrossRef]
  6. Lee K.S. Potentiation of the calcium-channel currents of internally perfused mammalian heart cells by repetitive depolarization. PNAS (1987) 84:3941–3945.[Abstract/Free Full Text]
  7. Tseng G.N. Calcium current restitution in mammalian ventricular myocytes is modulated by intracellular calcium. Circ. Res. (1988) 63:468–482.[Abstract/Free Full Text]
  8. Fedida D., Noble D., Spindler A.J. Use-dependent reduction and facilitation of Ca2+ current in guinea-pig myocytes. J. Physiol. (1988) 405:439–460.[Abstract/Free Full Text]
  9. Richard S., Tiaho F., Charnet P., Nargeot J., Nerbonne J.M. Two pathways for Ca2+ channel gating differentially modulated by physiological stimuli. Am. J. Physiol. (1990) 258:H1872–1881.[ISI][Medline]
  10. Zygmundt A.C., Maylie J. Stimulation-dependent facilitation of the high threshold calcium current in guinea pig ventricular myocytes. J. Physiol. (1990) 428:653–671.[Abstract/Free Full Text]
  11. Richard S., Charnet P., Nerbonne J.M. Voltage- and time-dependent interconversion between distinct gating pathways of the high threshold cardiac calcium channel. J. Physiol. (1993) 462:197–228.[Abstract/Free Full Text]
  12. Yuan W., Bers D.M. Ca-dependent facilitation of cardiac Ca current is due to Ca-calmodulin-dependent protein kinase. Am. J. Physiol. (1994) 267:H982–H993.[ISI][Medline]
  13. Piot C., Lemaire S., Albat B., Séguin J., Nargeot J., Richard S. High frequency-induced upregulation of human cardiac calcium currents. Circulation (1996) 93:120–128.[Abstract/Free Full Text]
  14. Lemaire S., Piot C., Leclercq F., Leuranguer V., Nargeot J., Richard S. Heart rate as a determinant of L-type Ca2+ channel activity: mechanisms and implication in force–frequency relation. Basic Res. Cardiol. (1998) 93(suppl. 1):51–59.[Medline]
  15. Delgado C., Artiles A., Gomez A.M., Vassort G. Frequency-dependent increase in cardiac Ca2+ current is due to reduced Ca2+ release by the sarcoplasmic reticulum. J. Mol. Cell. Cardiol. (1999) 31(10):1783–1793.[CrossRef][ISI][Medline]
  16. Ouadid H., Séguin J., Richard S., Chaptal P.A., Nargeot J. Properties and modulation of Ca channels in adult human atrial cells. J. Mol. Cell Cardiol. (1991) 23:41–54.[ISI][Medline]
  17. Ouadid H., Séguin J., Dumuis A., Bockaert J., Nargeot J. Serotonin increases calcium current in human atrial myocytes via the newly described 5-hydroxytryptamine 4 receptor. Mol. Pharmacol. (1992) 41:346–350.[Abstract]
  18. Lemaire S., Piot C., Séguin J., Nargeot J., Richard S. Tetrodotoxin-sensitive Ca2+ and Ba2+ currents in human atrial cells. Receptors Channels (1995) 3:71–81.[ISI][Medline]
  19. Lee K.S., Marban E., Tsien R.W. Inactivation of calcium channels in mammalian heart cells: joint dependence on membrane potential and intracellular calcium. J. Physiol. (1985) 364:395–411.[Abstract/Free Full Text]
  20. Sun H., Leblanc N., Nattel S. Mechanisms of inactivation of L-type calcium channels in human atrial myocytes. Am. J. Physiol. (1997) 272:H1625–H1635.[ISI][Medline]
  21. Sipido K., Calleweart G., Carmeliet E. Inhibition and rapid recovery of calcium release from sarcoplasmic reticulum in guinea pig ventricular myocytes. Circ. Res. (1995) 76:102–109.[Abstract/Free Full Text]
  22. Sham J.S. Ca2+ release-induced inactivation of Ca2+ current in rat ventricular myocytes: evidence for local Ca2+ signalling. J. Physiol. (1997) 500(2):285–295.[Abstract/Free Full Text]
  23. Adachi-Akahane S., Cleemann L., Morad M. Cross-signaling between L-type Ca2+ channels and ryanodine receptors in rat ventricular myocytes. J. Gen. Physiol. (1996) 108:435–454.[Abstract/Free Full Text]
  24. Masaki H., Sato Y., Luo W., Kranias A., Yatani A. Phospholamban deficiency alters inactivation kinetics of L-type Ca2+ channels in mouse ventricular myocytes. Am. J. Physiol. (1997) 272(41):H606–H612.[ISI][Medline]
  25. Tiaho F., Nargeot J., Richard S. Voltage-dependent regulation of L-type Ca channels by isoproterenol. Pflügers Arch. (1991) 419:596–602.[CrossRef][ISI][Medline]
  26. Bristow M.R., Ginsburg R., Minobe W., et al. Decreased catecholamine sensitivity and beta-adrenergic-receptor density in failing human hearts. N. Engl. J. Med. (1982) 307:205–211.[Abstract]
  27. Muntz K.H., Zhao M., Miller J.C. Downregulation of myocardial β-adrenergic receptors: receptor subtype selectivity. Circ. Res. (1994) 74:369–375.[Free Full Text]
  28. Xu X., Best P.M. Increase in T-type calcium current in atrial myocytes from adult rats with growth hormone-secreting tumors. Proc. Natl. Acad. Sci. USA (1990) 87:4655–4659.[Abstract/Free Full Text]
  29. Tiaho F., Piot C., Nargeot J., Richard S. Regulation of the frequency-dependent facilitation of L-type calcium currents in rat ventricular cells. J. Physiol. (1994) 477:237–252.[Abstract/Free Full Text]
  30. Sorota S. Insights into the structure, distribution and function of the cardiac chloride channels. Cardiovasc. Res. (1999) 42:361–376.[Abstract/Free Full Text]
  31. Howlett S.E., Zhu J.Q., Ferrier G.R. Contribution of voltage-sensitive calcium release mechanism to contraction in cardiac ventricular myocytes. Am. J. Physiol. (1998) 274(2):H155–H170.[ISI][Medline]
  32. Sipido K., Volders P.G.A., Antoons G., Vos M. L-type Ca2+ currents at negative potentials during β-adrenergic stimulation. Biophysical J (2000) 78:371A. Abstract.
  33. Pietrobon D., Hess P. Novel mechanism of voltage-dependent gating in L-type calcium channels. Nature (1990) 346:651–655.[CrossRef][Medline]
  34. Sipido K.R., Stankovicova T., Flameng W., Vanhaecke J. ha, Verdonck F. Frequency dependence of Ca2+ release from the sarcoplasmic reticulum in human ventricular myocytes from end-stage heart failure. Cardiovasc. Res. (1998) 37:478–488.[Abstract/Free Full Text]
  35. Linz K.L., Rainer M. The late component of L-type calcium current during guinea-pig cardiac action potentials and its contribution to contraction. Pflügers Arch. (1998) 436:679–688.[CrossRef][ISI][Medline]
  36. Ross J. Jr., Miura T., Kambayashi M., Eising G.P., Ryu K. Adrenergic control of the force frequency relation. Circulation (1995) 92:2327–2332.[Abstract/Free Full Text]
  37. Nakaya H., Hattori Y., Tohse N., Shida S., Kanno M. β-adrenoceptor-mediated depolarization of the resting membrane in guinea-pig papillary muscles: changes in intracellular Na+, K+ and Cl– activities. Pflügers Arch. (1990) 417:185–193.[CrossRef][ISI][Medline]
  38. Glitsch H.G., Rasch R. An effect of noradrenaline on resting potential and Na activity in sheep cardiac Purkinje fibers. Pflügers Arch. (1986) 406:144–150.[CrossRef][ISI][Medline]
  39. Terris S., Wasserstrom A., Fozzard H. Depolarizing effects of catecholamines in quiescent sheep cardiac Purkinje fibers. Am. J. Physiol. (1986) 25:H1056–H1061.
  40. Harvey R.D., Clarck C.D., Hume J.R. Chloride current in mammalian cardiac myocytes. Novel mechanism for autonomic regulation of action potential and resting membrane potential. J. Gen. Physiol. (1990) 95:1077–1702.[Abstract/Free Full Text]
  41. Pino R., Cerbaï E., Calamaï G., et al. Effect of 5-HT4 receptor stimulation on the pacemaker current If in human isolated atrial myocytes. Cardiovasc. Res. (1998) 40:516–522.[Abstract/Free Full Text]
  42. Ryden L., Ariniego M.D., Arnman K., et al. A double-blind trial of metoprolol in acute myocardial infarction. Effects on ventricular tachyarrhythmias. N. Engl. J. Med. (1983) 308:614–618.[Abstract]
  43. Kimura S., Cameron J., Kozlovskis P.L., Basset A.L., Myerburg R.J. Delayed after depolarization and triggered activity induced in feline Purkinje fibers by alpha adrenergic stimulation in the presence of elevated calcium level. Circulation (1984) 70:1074.[Abstract/Free Full Text]
  44. Gwathmey J.K., Copelas L., MacKinnon R., et al. Abnormal intracellular calcium handling in myocardium from patients with end-stage heart failure. Circ. Res. (1987) 61:70–76.[Abstract/Free Full Text]
  45. Beuckelmann D.J., Näbauer M., Erdmann E. Intracellular calcium handling in ventricular myocytes from patients with terminal heart failure. Circulation (1992) 85:1046–1055.[Abstract/Free Full Text]
  46. Pieske B., Kretschmann B., Meyer M., et al. Alterations in intracellular calcium handling associated with the inverse force–frequency relation in human dilated cardiomyopathy. Circulation (1995) 92:1169–1178.[Abstract/Free Full Text]
  47. Hasenfuss G., Mulieri L.A., Leavitt B.J., Allen P.D., Haeberle J.R., Alpert N.R. Alteration of contractile function and excitation–contraction coupling in dilated cardiomyopathy. Circ. Res. (1992) 70:1225–1232.[Abstract/Free Full Text]
  48. Schröder F., Handrock R., Beuckelmann D.J., et al. Increased availability and open probability of single L-type calcium channels from failing compared with nonfailing human ventricle. Circulation (1998) 98:969–976.[Abstract/Free Full Text]
  49. Eschenhagen T., Mende U., Nose M., et al. Increased messenger RNA level of the inhibitory G-protein {alpha}-subunit Gi{alpha}2 in human end-stage heart failure. Circ. Res. (1992) 70:688–696.[Abstract/Free Full Text]

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