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(Circulation Research. 1997;81:512-525.)
© 1997 American Heart Association, Inc.

Articles

Ionic Remodeling Underlying Action Potential Changes in a Canine Model of Atrial Fibrillation

Lixia Yue, Jianlin Feng, Rania Gaspo, Gui-Rong Li, Zhiguo Wang, , Stanley Nattel

From the Department of Medicine, Montreal Heart Institute (J.F., R.G., G.-R.L., Z.W., S.N.) and University of Montreal (R.G., G.-R.L., Z.W., S.N.), and the Department of Pharmacology and Therapeutics, McGill University (L.Y., S.N.), Montreal, Quebec, Canada.

Correspondence to Stanley Nattel, MD, Research Center, Montreal Heart Institute, 5000 Bélanger St, Montreal, Quebec H1T 1C8, Canada.

	Abstract

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Abstract Rapid electrical activation, as occurs during atrial fibrillation (AF), is known to cause reductions in atrial refractoriness and in adaptation to heart rate of the atrial refractory period, which promote the maintenance of AF, but the underlying ionic mechanisms are unknown. In order to determine the cellular and ionic changes caused by chronic atrial tachycardia, we studied right atrial myocytes from dogs subjected to 1, 7, or 42 days of atrial pacing at 400/min and compared them with myocytes from sham-operated dogs (pacemaker inserted but not activated). Rapid pacing led to progressive increases in the duration of AF induced by bursts of 10-Hz stimuli (from 3±2 seconds in sham-operated dogs to 3060±707 seconds in dogs after 42 days of pacing, P<.001) and reduced atrial refractoriness and adaptation to rate of the atrial refractory period. Voltage-clamp studies showed that chronic rapid pacing did not alter inward rectifier K⁺ current, rapid or slow components of the delayed rectifier current, the ultrarapid delayed rectifier current, T-type Ca²⁺ current, or Ca²⁺-dependent Cl^- current. In contrast, the densities of transient outward current (I_to) and L-type Ca²⁺ current (I_Ca) were progressively reduced as the duration of rapid pacing increased, without concomitant changes in kinetics or voltage dependence. In keeping with in vivo changes in refractoriness, action potential duration (APD) and APD adaptation to rate were decreased by rapid pacing. The response of the action potential and ionic currents flowing during the action potential (as exposed by action-potential voltage clamp) to nifedipine in normal canine cells and in cells from rapidly paced dogs suggested that the APD changes in paced dogs were largely due to reductions in I_Ca. We conclude that sustained atrial tachycardia reduces I_to and I_Ca, that the reduced I_Ca decreases APD and APD adaptation to rate, and that these cellular changes likely account for the alterations in atrial refractoriness associated with enhanced ability to maintain AF in the model.

Key Words: Ca²⁺ channel • K⁺ channel • cardiac arrhythmia • cardiac ionic current • action potential

	Introduction

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Atrial fibrillation is the most frequently encountered sustained arrhythmia in clinical practice.^{1 2 3} Since early in the century, the presence of multiple simultaneous reentry circuits has been considered central to the maintenance of AF.⁴ Brief refractory periods favor the occurrence of reentrant arrhythmias,^{5 6} thereby promoting multiple-wavelet reentry in AF.^{7 8 9} There is evidence indicating the important role of reduced atrial refractoriness^{10 11 12} and of the occurrence of multiple-wavelet reentry^{13 14 15} in clinical AF. Although much is known about these functional mechanisms of AF, the nature of cellular abnormalities and, in particular, changes in ionic currents that promote AF has not been determined.

One of the limitations in the study of the cellular mechanisms of AF has been the lack of a reliable animal model of atrial pathology associated with sustained AF. It has recently been shown that chronic rapid atrial activation, whether by the electrical maintenance of AF^{16 17} or by rapid atrial pacing,¹⁸ results in atrial dilation,^{16 18} ultrastructural abnormalities,¹⁸ and markedly increased susceptibility to sustained AF.^{16 17 18} Electrophysiological abnormalities contributing to AF in this model include reductions in the atrial refractory period^{17 18} and a decrease in the adaptation to rate during the refractory period,¹⁸ changes that are qualitatively similar to those in patients with increased vulnerability to AF.¹⁹ An ability to maintain AF develops gradually in the rapid-activation AF model, allowing for systematic studies of underlying cellular mechanisms. Furthermore, the model is likely relevant to electrical remodeling promoting AF maintenance in patients with AF.¹⁷ The purpose of the present study was to evaluate the changes in action potential properties and in ionic currents in atrial myocytes from dogs subjected to rapid atrial pacing and to relate these changes to refractory period abnormalities associated with the ability to sustain AF. In particular, we wished to examine currents that we have previously found to be important in governing canine atrial repolarization: I_K1, I_K, I_Ca, I_Cl.Ca, and a novel ultrarapid delayed rectifier current that we have designated I_Kur.d.^{20 21} Preliminary results of these experiments have been presented in abstract form.²²

	Materials and Methods

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Preparation of the Canine Model
We used a previously described approach to produce dogs with the ability to maintain sustained AF.¹⁸ Fifteen mongrel dogs weighing 27.3±2.4 kg were anesthetized with pentobarbital (30 mg/kg IV). A programmable pacemaker was inserted in a subcutaneous pocket with sterile techniques, and a tined atrial pacing lead was positioned in the right atrial appendage under fluoroscopic guidance. The dogs were then permitted to recover for 24 hours, after which atrial pacing was initiated at 400/min. Pacing was maintained for 1, 7, or 42 days (n=5, 5, and 7, respectively, for each group). These groups will be designated P1, P7, and P42 dogs, respectively, throughout this article. Five additional dogs underwent pacemaker insertion, but the pacemaker was not activated to provide atrial pacing; they served as sham-operated controls and were studied 7 days after pacemaker insertion. An observation period of 7 days was chosen for the sham-operated dogs because the median observation period in rapidly paced dogs was 7 days, and in previous studies we have noted that atrial electrophysiological properties and AF duration are unaltered in sham-operated dogs observed for up to 42 days.²³ The sham-operated dogs will be designated P0 dogs, in contrast to the rapidly paced P1, P7, and P42 dogs.

On the study day, dogs were anesthetized with subcutaneous morphine (2 mg/kg SC) and intravenous -chloralose (120 mg/kg), and a right thoracotomy was performed. The atrial ERP was measured with the extrastimulus technique at basic cycle lengths ranging from 150 to 400 milliseconds, with the use of twice–diastolic-threshold current (2-millisecond square-wave pulses, Digital Cardiovascular Instruments stimulator). At least 2 minutes was allowed at each basic cycle length before measuring ERP. ERP measurements required a total of 15 minutes and were performed in duplicate at all cycle lengths. The average of the two ERP determinations (which were always within 5 milliseconds of each other) was taken to represent ERP at a given cycle length within a given dog. AF was then induced 10 times by atrial burst pacing (10 Hz, 2-millisecond pulses, four times diastolic-threshold current), except when AF was sustained (>30 minutes), in which case, direct-current cardioversion was applied, and AF reinduction was not performed. At least 30 minutes was allowed after cardioversion, and the return of the atrial refractory period to precardioversion values was confirmed before proceeding. When AF was not sustained, the mean duration of AF during the 10 AF inductions was taken to represent AF duration in that animal. The hearts were then excised, and single atrial cells were obtained as described below.

Cell Isolation
Single cells were isolated by previously developed methods.^{20 21} Immediately after cardiac excision, the hearts were immersed in Tyrode's solution (for composition of solutions, see below) at room temperature. All solutions used for dissection and perfusion were equilibrated with 100% O₂. The right coronary artery was cannulated, and the right atrium was dissected free and perfused with Tyrode's solution at 37°C for 5 minutes until the effluent was clear of blood. Any leaking arterial branches were ligated with silk thread to ensure adequate perfusion. The tissue was then perfused at 12 mL/min with nominally Ca²⁺-free Tyrode's solution for 20 minutes, followed by 40-minute perfusion with the same solution containing collagenase (100 U/mL, CLSII, Worthington Biochemical) and 1% bovine serum albumin (Sigma Chemical Co). A small piece of tissue from a well-perfused region was minced, and the cells were harvested. Cells were kept at room temperature in a high-K⁺ storage solution.

Only quiescent rod-shaped cells showing clear cross striations were used. A small aliquot of the solution containing the isolated cells was placed in a 1-mL chamber mounted on the stage of an inverted microscope. Five minutes was allowed for cell adhesion to the bottom of the chamber, and then the cells were superfused at 3 mL/min with Tyrode's solution, modified as described below to record specific currents. Experiments were performed at 35°C for the study of action potentials and all currents except for I_Kur.d. We have previously shown that I_Kur.d amplitude at room temperature is not different from that at 35°C and that I_Kur.d kinetics are easier to resolve at room temperature.²¹ The bath temperature was maintained at 35°C with a Peltier-effect device (N.B. Datyner).

Solutions
Tyrode's solution contained (mmol/L) NaCl 126, CaCl₂ 2 (except as otherwise indicated), KCl 5.4, MgCl₂ 0.8, NaH₂PO₄ 0.33, dextrose 10, and HEPES 10, pH 7.4 (adjusted with NaOH). The high-K⁺ storage solution contained (mmol/L) KCl 20, KH₂PO₄ 10, dextrose 10, glutamic acid 70, ß-hydroxybutyric acid 10, taurine 10, and EGTA 10, along with 1% albumin, pH 7.4 (adjusted with KOH).

The pipette solution for action potential recording contained (mmol/L) GTP 0.1, potassium aspartate 110, KCl 20, MgCl₂ 1, ATP-Mg 5, HEPES 10, Na₂-phosphocreatine 5, and EGTA 0.05, pH 7.4 (adjusted with KOH). The same pipette solution was used to study K⁺ currents, except that the pipette EGTA concentration was increased to 10 mmol/L. The Tyrode's solution was modified for K⁺ current measurement by the addition of Cd²⁺ (200 µmol/L) to block I_Ca and I_Cl.Ca and atropine (200 nmol/L) to eliminate any basal activity of acetylcholine-dependent K⁺ current. For studies of I_K, 2 mmol/L 4AP was added to the superfusate to block I_to and I_Kur.d. E-4031 (5 µmol/L), a highly selective blocker of I_Kr, was used to separate I_Kr from the slower drug-resistant component, I_Ks. I_to was studied in the presence of 10 mmol/L TEA (to inhibit I_Kur.d²¹ ) and 5 µmol/L E-4031 (to suppress I_Kr), with the use of short depolarizing pulses to minimize I_Ks. I_Kur.d was recorded at room temperature (21°C to 23°C). I_K1 was identified on the basis of sensitivity to 0.5 mmol/L Ba²⁺ and typical time and voltage dependence. Contamination by I_Na was prevented by using an HP of -50 mV and/or by isomolar substitution of Tris for extracellular Na⁺.

The extracellular solution for studies of I_Ca and I_Cl.Ca contained (mmol/L) TEA-Cl 126, CsCl 5.4, MgCl₂ 1, CaCl₂ 2, NaH₂PO₄ 0.33, dextrose 10, and HEPES 10, pH 7.4 (adjusted with CsOH). In studies of I_Ca, 2 µmol/L ryanodine was added to suppress I_Cl.Ca, which could otherwise overlap with and contaminate I_Ca.²⁴ The pipette solution used to record I_Ca contained (mmol/L) CsCl 20, cesium aspartate 110, MgCl₂ 1, EGTA 10, GTP 0.1, ATP-Mg 5, HEPES 10, and Na₂-phosphocreatine 5, pH 7.4 (adjusted with CsOH). The same pipette solution was used to record I_Cl.Ca, except that the EGTA concentration was reduced to 0.02 mmol/L in order to allow for the Ca²⁺ transients necessary for free I_Cl.Ca expression.²⁴ In order to record I_Cl.Ca, it is necessary to allow for L-type Ca²⁺ current; thus, all I_Cl.Ca recordings are contaminated by overlapping I_Ca. To isolate I_Cl.Ca and minimize contamination by I_Ca, we used two approaches: (1) currents were recorded before and after 5 to 10 minutes of exposure to 2 µmol/L ryanodine (sufficient time to suppress fully the outward current component), and (2) currents were recorded before and after substitution of methanesulfonate for Cl^- in the bath. The outward current component sensitive to ryanodine or extracellular Cl^- substitution, which is I_Cl.Ca,^{21 24} was then obtained by digital subtraction. T-type Ca²⁺ current was recorded with the same solutions as described above for L-type Ca²⁺ current, but NaH₂PO₄ was eliminated from the bath, and Na₂-phosphocreatine was omitted in the pipette solution. T-type current was separated from L-type current by recording current at holding potentials of -90 and -50 mV and subtracting the current at an HP of -50 mV from that obtained at an HP of -90 mV.

Data Acquisition and Analysis
The whole-cell patch-clamp technique was used to record ionic currents in the voltage-clamp mode, and action potentials were recorded in current-clamp mode. Borosilicate glass electrodes (1-mm outside diameter) were filled with pipette solution and connected to a patch-clamp amplifier (Axopatch 1-D, Axon Instruments). Electrodes with tip resistances of 1 to 2 M were used to record whole-cell currents, and 3- to 5-M electrodes were used to record action potentials.

Cells with resting potentials in the normal physiological range, negative to -70 mV, were selected for action potential recording, after verifying that mean resting potential was not altered in paced dogs. Action potentials were elicited by 2-millisecond twice-threshold pulses. APD stabilized within 15 action potentials at each frequency, and steady state APD was measured to 20% (APD₂₀), 50% (APD₅₀), and 95% (APD₉₅) of full repolarization. Voltage command pulses were generated by a 12-bit digital-to-analog converter controlled by pClamp 6 software (Axon Instruments). Recordings were low pass–filtered at 1 to 5 kHz (half the sampling frequency). Data were sampled at rates varying from 2 to 10 kHz (with sampling at 10 kHz used for the action potential and the rapidly activating currents, I_Ca, I_to, I_Cl.Ca, and I_Kur.d, and sampling at 2 kHz used for the slower currents, I_Kr and I_Ks) and then stored on the hard disk of an IBM-compatible computer. In some experiments, we applied the action potential as a voltage command waveform for voltage clamp ("AP clamp" technique). The action potential was first recorded and then stored as a BPA (binary parameter) file in pCLAMP 6. The stored waveform was then used to voltage-clamp the cell, and net current flow during the action potential was recorded. The action potential was then immediately recorded again, to ensure its stability. In some experiments, AP clamp with the control action potential waveform was performed before and after the rapid application of a blocking drug (changeover within 300 milliseconds via a capillary tube adjacent to the cell).

Tip potentials (2 to 8 mV) were zeroed before formation of the membrane-pipette seal in Tyrode's solution. Junction potentials (10 to 11 mV) after rupture were corrected for action potential recordings only. Mean seal resistance averaged 12.5±0.9 G. Several minutes after seal formation, the membrane was ruptured by gentle suction to establish the whole-cell configuration. R_s was electrically compensated to minimize the duration of the capacitive surge on the current recording and the voltage drop across the clamped cell membrane. R_s along the clamp circuit was estimated by dividing the time constant obtained by fitting the decay of the capacitive transient by the calculated membrane capacitance (the time integral of the capacitive response to a 5-mV hyperpolarizing step from an HP of -60 mV divided by the voltage drop). Before R_s compensation, the decay of the capacitive surge was a single exponential function of time with a time constant of 519±43 microseconds (cell capacitance, 76.8±5.9 pF). Precompensation R_s values averaged 7.2±0.6 M. After compensation, the time constant was reduced to 147±10 microseconds, and R_s was reduced to 2.1±0.2 M. Currents recorded during the present study rarely exceeded 1.5 nA. The mean maximum voltage drop across the R_s was thus in the range of 3 mV. Cells with significant leak currents were rejected, and leakage compensation was not used.

Data Analysis
In order to ensure representativity of voltage-clamp data, similar numbers of cells from each heart were studied with each protocol (ie, the cells were distributed evenly across dogs). Group data are presented as mean±SEM. Nonlinear curve fitting was performed with the Clampfit routine in pCLAMP 6 (Chebyshev algorithm). Statistical comparisons among groups were performed with ANOVA. If significant effects were indicated by ANOVA, a t test with Bonferroni's correction or Dunnett's test was used to evaluate the significance of differences between individual mean values. A two-tailed P<.05 was taken to indicate statistical significance.

	Results

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Electrophysiological Changes Produced by Sustained Rapid Atrial Pacing
Rapid pacing resulted in an increased duration of induced AF, which averaged 3.4±2.4 seconds in P0 dogs and 24±28 seconds (P=NS), 81±55 seconds (P<.05 versus control), and 3060±707 seconds (P<.001 versus control) in P1, P7, and P42 dogs, respectively. Atrial ERP was reduced in paced dogs, from a mean of 141±5 milliseconds at a cycle length of 400 milliseconds in P0 dogs to 115±6 milliseconds (P<.05 versus P0 dogs), 91±9 milliseconds (P<.05 versus P0 dogs), and 88±9 milliseconds (P<.01 versus P0 dogs) at the same cycle length in P1, P7, and P42 dogs, respectively. In P0 dogs, the ERP decreased consistently when the basic cycle length was reduced from 400 to 150 milliseconds, with a mean change in ERP of -43.6±10.3 milliseconds in all five P0 dogs over this cycle-length range. The tachycardia-dependent reduction in ERP was strongly affected by rapid pacing, with the mean change in ERP at a cycle length of 150 milliseconds relative to that at 400 milliseconds averaging -20.3±4.5 milliseconds (P<.05 versus P0 dogs) in P1 dogs, -5.6±1.0 milliseconds (P<.01 versus P0 dogs) in P7 dogs, and -1.0±3.3 milliseconds (P<.05 versus P0 dogs) in P42 dogs. In summary, rapid atrial pacing caused a progressive increase in the duration of AF, associated with decreases in atrial ERP and in ERP adaptation to increases in atrial rate.

Action Potential Changes Associated With Changes in AF Duration
Resting membrane potential was not altered by rapid pacing, averaging -63.6±0.9 mV (n=65 cells) in P0 dogs compared with -64.1±0.8 mV (n=68), -62.3±0.7 mV (n=66), and -63.9±0.7 mV (n=69) in P1, P7, and P42 dogs, respectively (P=NS). Only cells in which action potentials were stable for at least 20 minutes were used for analysis. Action potential measurements were begun 5 minutes after cell rupture. In cells used for action potential studies, the resting potential averaged -74.1±0.6, -73.5±0.6, -74.1±0.7, and -73.3±0.7 mV in P0, P1, P7, and P42 dogs, respectively (n=25 cells for each). Action potential amplitude at 1 Hz averaged 120±1 mV in P0 cells, 118±1 mV in P1 cells (P=NS versus P0 cells), 109±1 mV in P7 cells (P<.001 versus P0 cells), and 100±1 mV in P42 cells (P<.001 versus P0 cells, n=25 cells per group).

Rapid pacing did not substantially alter APD₂₀ but produced important and qualitatively similar changes in APD₅₀ and APD₉₅. Mean values of APD are shown for all groups of dogs in Table 1. Repolarization occurred progressively and highly significantly earlier as pacing duration increased. In addition to reducing APD, pacing also caused highly significant reductions in rate-dependent APD changes, as illustrated in Fig 1 and summarized in Table 2. In P0 dogs (Fig 1A), APD decreased as stimulation frequency increased. Within 1 day of the onset of rapid pacing, rate-dependent APD abbreviation was attenuated (Fig 1B). After 7 days, there was a marked reduction in the rate dependence of APD (Fig 1C), and after 42 days of rapid pacing, there was virtually no rate dependence of the APD discernible (Fig 1D). Figs 1E and 1F show mean data for APD₂₀, APD₅₀, and APD₉₅ over a wide range of frequencies in 10 cells each from P0 and P42 dogs, along with changes in ERP measured in vivo. The changes in ERP resulting from rapid pacing are very similar to those in APD₉₅ and are consistent with the notion that alterations in repolarization are largely responsible for in vivo changes in refractoriness. Because of their limited rate adaptation, ERP and APD changes in rapidly paced dogs were reduced at higher frequencies; eg, ERP at 6.7 Hz was 87±7 milliseconds in P42 dogs, an 11% reduction compared with the value of 97±6 milliseconds in P0 dogs, and APD₉₅ at 6 Hz was 86±3 milliseconds in P42 dogs, a reduction of 15% from the value of 102±3 milliseconds in P0 dogs.

View this table: [in this window] [in a new window]   Table 1. Changes in APD at Various Degrees of Repolarization Caused by Rapid Atrial Pacing

View larger version (25K): [in this window] [in a new window]   Figure 1. Action potential recordings from representative cells obtained from a sham-operated (P0) dog (A) and dogs subjected to 1 (B), 7 (C), and 42 (D) days of atrial pacing at 400/min. Action potentials were recorded in current-clamp mode at the frequencies indicated. The relationships between APD20, APD50, and APD95 over a wide range of frequencies in vitro (10 cells in each group) and ERP in vivo (five dogs in each group) are shown for P0 dogs (E) and P42 dogs (F) as the mean±SEM (where error bars are not visible, they fall within symbol for the mean).

View this table: [in this window] [in a new window]   Table 2. Changes in APD Caused by Rapid Atrial Pacing: Rate-Dependent Decreases in APD95

I_K1
Cell capacitance was not altered in paced dogs; it averaged 73.7±0.7 pF (n=150) in P0 dogs compared with 75.8±0.3 pF (n=150), 74.2±0.6 pF (n=150), and 76.7±0.8 pF (n=150) in P1, P7, and P42 dogs, respectively. Nonetheless, all current amplitudes are presented as current densities to control for intercell variability in size. I_K1 was measured as the 0.5 mmol/L Ba²⁺-sensitive current upon 300-millisecond pulses from an HP of -40 mV to voltages ranging from -120 to -10 mV (Fig 2A). No clear changes in I_K1 were observed in paced dogs (Fig 2B through 2D). Mean I_K1 density-voltage relations were not changed in paced dogs (Fig 2E), consistent with the lack of any pacing-induced change in the resting potential.

View larger version (28K): [in this window] [in a new window]   Figure 2. A through D, Recordings of IK1 obtained with the voltage protocol shown in the inset at 0.1 Hz in representative cells obtained from a sham-operated dog (A) and dogs subjected to 1 (B), 7 (C), and 42 (D) days of rapid atrial pacing. The recordings shown were obtained by subtracting recordings in the presence of 0.5 mmol/L Ba2+ from recordings before Ba2+ in the same cell (Ba2+-sensitive current). E, Mean±SEM data for IK1 density as a function of test potential (TP) (n=25 cells per group).

I_to
Depolarizing 100-millisecond pulses from -80 mV elicited typical I_to in P0 dogs (Fig 3A). I_to was progressively reduced in paced dogs (Fig 3B through 3D), with no obvious change in the time course of the current. The amplitude of I_to was quantified as the difference between peak and end-pulse steady state current. Mean I_to density decreased progressively at all voltages as pacing duration increased (Fig 3E, n=25 cells per group). Although I_to density decreased in paced dogs, the form of the I-V curve did not change. This is best appreciated by assessing the I-V curves expressed with the currents normalized to maximum current in each cell, as shown in Fig 3F. The normalized I-V curves for all groups are superimposed.

View larger version (31K): [in this window] [in a new window]   Figure 3. A through D, Recordings of Ito obtained with the voltage protocol shown in the inset at 0.1 Hz in representative cells obtained from a sham-operated dog (A) and dogs subjected to 1 (B), 7 (C), and 42 (D) days of rapid atrial pacing. E, Mean±SEM Ito density as a function of test potential (TP) (n=25 cells per group). *P<.05, **P<.01, and ***P<.001 vs P0 Ito at the same TP. F, Ito at each TP normalized to current at +50 mV for each cell.

In order to assess possible mechanisms of the pacing-induced reduction in I_to, we studied the voltage- and time-dependent properties of I_to. The voltage dependence of inactivation was studied with a double-pulse protocol at an HP of -80 mV (Fig 4A), whereas the voltage dependence of activation was determined from the I-V relation upon step depolarization with changes in driving force corrected by dividing each current by the difference between test potential and the mean reversal potential of I_to tails (-73.6±6.4 mV) as previously described.²⁰ As shown in Fig 4A, mean voltage-dependent activation and inactivation curves (based on 10 cells per group) were not altered by pacing.

View larger version (28K): [in this window] [in a new window]   Figure 4. A, Voltage dependence of inactivation (Inact) and activation (Act) of Ito in various groups of dogs. Inact was assessed with a 1-second prepulse from an HP of -80 mV to the test potential (TP) indicated, followed by a 100-millisecond test pulse to +60 mV. Act voltage dependence was assessed by currents recorded upon depolarization from -80 mV to the voltages indicated, with changes in driving force corrected by dividing current during the TP by the driving force, TP-Erev, where Erev is the reversal potential of Ito tail current determined as previously described.20 The symbols are mean±SEM data for each group of cells (n=10 for each). Curves are best-fit Boltzmann relations to mean data in each group of cells. B, Inact time constants (inact) and time to peak current for Ito elicited by 100-millisecond depolarizations to the TPs shown (mean±SEM, n=10 cells per group). C, Recovery time course of Ito, determined by the ratio of current during a 150-millisecond test pulse (P2) from -80 to +50 mV to that during an identical conditioning pulse (P1) with varying P1-P2 interval. The solid curves are best monoexponential fits to the mean±SEM data shown (n=10 cells per group). D, Frequency dependence of Ito, as determined by the ratio of current during the 15th pulse to current during the first pulse of a train of 100-millisecond depolarizations from -80 to +50 mV at the frequencies shown. Results are mean±SEM of 10 cells per group. All protocols in each panel were delivered at 10-second intervals.

The kinetics of I_to activation were evaluated by measuring time to peak current, and inactivation kinetics were analyzed by fitting a monoexponential relation to the time course of current decay during a depolarizing step. Pacing did not alter the time course of I_to activation or inactivation, as shown by the mean results obtained in 10 cells per group (Fig 4B). The recovery kinetics of I_to were assessed at an HP of -80 mV with a paired pulse protocol (150-millisecond pulses, pulse 1 and pulse 2, from -80 mV to +50 mV with a varying pulse 1–pulse 2 interval) as shown in the inset of Fig 4C. Current during pulse 2 was normalized to that during pulse 1 and showed monoexponential recovery as a function of the pulse 1–pulse 2 interval (Fig 4C). The recovery time constant averaged 24.3±3.1, 25.8±3.2, 22.9±2.9, and 21.8±1.8 milliseconds (n=10 in each group), respectively, in P0, P1, P7, and P42 dogs (P=NS). The frequency dependence of I_to was tested with a train of 15 pulses from -80 mV to +50 mV, with current during the last pulse of the train normalized to first-pulse current. Mean data from 10 cells per group show that pacing did not alter the frequency dependence of I_to (Fig 4D). These results suggest that pacing decreased I_to by reducing its maximum conductance, without changing any other biophysical properties of the current.

I_Cl.Ca
I_Cl.Ca is present in canine atrium and can contribute significantly to repolarization of this tissue.²⁰ Fig 5 shows an analysis of the properties of I_Cl.Ca in control and paced dogs. Panels A through D show ryanodine-sensitive currents obtained by subtracting currents recorded (with the voltage protocol shown in panel A) in the presence of 2 µmol/L ryanodine from currents recorded from the same cells with the same protocol before ryanodine infusion. Currents were not obviously altered in cells from paced dogs. Fig 5E shows mean I_Cl.Ca density-voltage relations based on ryanodine-sensitive currents from 25 cells in each group and indicates that pacing did not alter I_Cl.Ca amplitude. Because of the possibility of ryanodine effects on currents other than I_Cl.Ca, we also studied I_Cl.Ca with the use of current sensitive to extracellular Cl^- substitution in an additional 10 cells in each group. Once again, pacing did not appear to alter I_Cl.Ca, which at +40 mV averaged 451±50 pA in P0 cells, 443±43 pA in P1 cells, 459±51 pA in P7 cells, and 467±48 pA in P42 cells (P=NS for intergroup differ-ences). Fig 5F shows the frequency dependence of I_Cl.Ca as determined with 15-pulse trains of 100-millisecond pulses from -80 to +50 mV (n=10 cells per group). The frequency dependence of I_Cl.Ca was not altered by rapid pacing.

View larger version (29K): [in this window] [in a new window]   Figure 5. A through D, Recordings of ICl.Ca obtained with the voltage protocol shown in the inset at 0.1 Hz in representative cells obtained from a sham-operated dog (A) and dogs subjected to 1 (B), 7 (C), and 42 (D) days of rapid atrial pacing. Results shown were obtained by applying the voltage protocol before and after ryanodine and subtracting currents in the presence of ryanodine from control currents. E, Mean±SEM ICl.Ca density as a function of test potential (TP) (n=25 cells per group). F, Frequency dependence of ICl.Ca, as determined by the ratio of current during the 15th pulse to current during the first pulse of a train of 100-millisecond depolarizations from -80 to +50 mV at the frequencies shown. Results are mean±SEM of 10 cells per group. Trains of 15 pulses at the frequencies shown were separated by 10 seconds at the HP.

Delayed Rectifier K⁺ Currents
Delayed rectifier K⁺ currents, both the classical delayed rectifier I_K and the ultrarapid delayed rectifier I_Kur.d, are important in canine atrial repolarization^{20 21} and could therefore underlie the APD changes caused by rapid atrial pacing. Fig 6 shows an analysis of I_K in control and paced dogs. The overall form of original recordings was not altered by pacing (Fig 6A through 6D). Fig 6E shows mean I_K step and tail current densities (n=25 cells per group) and indicates that total I_K was not altered by pacing. Although overall I_K was not changed, this does not exclude subtle, but potentially significant, changes in the components I_Kr and/or I_Ks. Therefore, we applied the highly selective I_Kr blocker, E-4031 (5 µmol/L), and measured I_K with the protocol shown in Fig 6 before and after the drug in 10 cells in each group, in order to separate the E-4031–sensitive component (I_Kr) from the drug-resistant component (I_Ks). No significant change in the individual components was noted. For example, upon stepping to +30 mV, mean I_Kr step current density was 1.1±0.1, 1.2±0.1, 1.1±0.1, and 1.2±0.1 pA/pF, respectively, in P0, P1, P7, and P42 dogs (P=NS). I_Ks density averaged 3.7±0.4, 4.0±0.4, 3.6±0.4, and 3.8±0.4 pA/pF, respectively, in P0, P1, P7, and P42 dogs (P=NS).

View larger version (28K): [in this window] [in a new window]   Figure 6. A through D, Recordings of IK obtained with the voltage protocol shown in the inset (3-second activating pulse to various test potentials [TPs], followed by 1-second repolarization to -30 mV to observe tail currents) at 0.1 Hz in representative cells obtained from a sham-operated dog (A) and dogs subjected to 1 (B), 7 (C), and 42 (D) days of rapid atrial pacing. E, Mean±SEM density of IK step and tail currents as a function of TP (n=25 cells per group).

Fig 7 shows an analysis of I_Kur.d in sham-operated and paced dogs. Currents were elicited by 140-millisecond depolarizations to voltages ranging from -40 to +60 mV, followed by repolarization for 60 milliseconds to -30 mV to record tail currents. An HP of -50 mV and an 80-millisecond prepulse to +30 mV 10 milliseconds before the test pulse were used to suppress I_to and elicit selectively I_Kur.d as previously described.²¹ Original recordings showed the typical voltage and time dependence of I_Kur.d,²¹ with no change in form caused by pacing (Fig 7A through 7D). Mean step current density as a function of voltage (n=25 cells per group) was not altered by pacing (Fig 7E), nor was the frequency dependence of the current, as established by a series of 15 pulses from -50 to +50 mV (n=10 cells per group). The results of the experiments illustrated in Figs 6 and 7 indicate that changes in delayed rectifier currents do not account for the alterations in APD produced by rapid atrial pacing.

View larger version (29K): [in this window] [in a new window]   Figure 7. A through D, Recordings of IKur.d obtained with the voltage protocol shown in the inset at 0.1 Hz in representative cells obtained from a sham-operated dog (A) and dogs subjected to 1 (B), 7 (C), and 42 (D) days of rapid atrial pacing. E, Mean±SEM IKur.d density as a function of test potential (TP) (n=25 cells per group). F, Frequency dependence of IKur.d, as determined by the ratio of current during the 15th pulse to current during the first pulse of a train of 100-millisecond depolarizations from -80 to +50 mV at the frequencies shown. Results are mean±SEM of 10 cells per group. Trains of 15 pulses at the frequencies shown were separated by 10 seconds at the HP.

I_Ca
I_Ca plays a significant role in maintaining the plateau in canine atrial myocytes²⁰ and has been found to be important in mediating rate-dependent APD changes in human atrium.²⁵ I_Ca is therefore a candidate to underlie the APD changes caused by atrial pacing in dogs. Any contaminating effects of I_Ca rundown were minimized by beginning all studies 5 minutes after membrane rupture, performing protocols in the same sequence with the I_Ca density-voltage relation studied first in all groups of cells, and bracketing protocols by an I_Ca measurement, which was required to vary by <5% over the course of the protocol (otherwise the experiment was rejected). Fig 8A through 8D shows typical I_Ca recordings upon 240-millisecond depolarizing pulses from -50 mV to voltages ranging from -40 mV to +60 mV. Sustained rapid atrial pacing produced a progressive decline in I_Ca amplitude. Peak I_Ca density (mean±SEM, 25 cells per group) is shown as a function of test potential in Fig 8E. I_Ca density was reduced progressively and highly significantly by rapid atrial pacing. For example, at +10 mV, I_Ca density averaged -12.2±0.8 pA/pF in P0 dogs compared with -8.4±0.5 pA/pF (P<.05 versus P0 dogs) in P1 dogs, -5.9±0.4 pA/pF (P<.001 versus P0 dogs) in P7 dogs, and -3.8±0.2 pA/pF (P<.001 versus P0 dogs) in P42 dogs. On the other hand, the form of the I_Ca I-V relation was not changed, as illustrated by Fig 8F, which shows current normalized to the maximum value in each cell; normalized I-V relations are superimposed.

View larger version (25K): [in this window] [in a new window]   Figure 8. Recordings of ICa obtained with the voltage protocol shown in the inset at 0.1 Hz in representative cells obtained from a sham-operated dog (A) and dogs subjected to 1 (B), 7 (C), and 42 (D) days of rapid atrial pacing. E, Mean±SEM ICa density as a function of test potential (n=25 cells per group). *P<.05, **P<.01, and ***P<.001 vs control ICa at the same test potential. F, Mean±SEM ICa obtained by expressing current at each potential relative to peak current at +10 mV in each cell.

To evaluate possible mechanisms involved in the pacing-induced reduction in I_Ca, we studied the voltage- and time-dependent properties of the current as shown in Fig 9. A double-pulse protocol was used to assess the voltage dependence of I_Ca inactivation, as shown in Fig 9A. A 1000-millisecond conditioning pulse to voltages between -90 and +50 mV was applied before a 300-millisecond test pulse to +10 mV (HP, -80 mV; 0.1 Hz). The peak current elicited by the test pulse was normalized to current without a prepulse. Inactivation increased to reach a maximum at 0 mV, decreasing thereafter because of the well-recognized reduction at more positive voltages of Ca²⁺-dependent inactivation.^{26 27} The results between -90 and 0 mV were well-fitted by a modified Boltzmann relation of the form (I_V-I_min)/(I_max-I_min)=1/{1+exp[(V-V_1/2)/s]}, where I_V is I_Ca at prepulse voltage V, I_max and I_min are I_Ca without a prepulse and at the prepulse producing maximum inactivation, respectively, V_1/2 is the voltage for half-maximal inactivation, and s is a slope factor. Pacing did not alter I_Ca inactivation, with V_1/2 averaging -24.6±2.3, -24.3±2.0, -24.5±2.4, and -24.2±3.1 mV in P0, P1, P7, and P42 dogs, respectively (P=NS, n=10 cells per group) and s averaging 6.2±0.7, 7.1±0.8, 6.3±0.64, and 6.2±0.6 mV.

View larger version (20K): [in this window] [in a new window]   Figure 9. A, Voltage dependence of inactivation (Inact) and activation (Act) of ICa in various groups of dogs. Inact was assessed with a 1-second prepulse from an HP of -80 mV to voltages between -90 and +50 mV, followed by a 300-millisecond test pulse to +10 mV. Act voltage dependence was assessed by currents recorded upon depolarization from -80 mV to the test potential (TP) indicated, with changes in driving force corrected by dividing current during the test pulse by the driving force, TP-Erev, where Erev is the reversal potential determined from the voltage intercept of the ascending limb of the ICa I-V relation illustrated in Fig 8E. The symbols are mean±SEM data for each group of cells (n=10 for each). Curves are best-fit Boltzmann relations to mean data in each group of cells. B, Recovery time course of ICa, determined by the ratio of current during a 300-millisecond test pulse (P2) from -80 to +10 mV to that during an identical conditioning pulse (P1) with a varying P1-P2 interval. The solid curves are best monoexponential fits to the mean±SEM data shown (n=10 cells per group). C, Inact time constants (1 and 2) from biexponential curve fits to decay of ICa during 300-millisecond depolarizations to the TPs shown (mean±SEM, n=10 cells per group).

The voltage dependence of I_Ca activation was determined (Fig 9A) by dividing peak current during depolarizing test pulses by the driving force (calculated as the difference between test potential and I_Ca reversal potential from the I-V relation). The results were well-fitted by a Boltzmann relation, with half-maximum activation voltage averaging -12.1±0.9, -10.6±1.1, -11.7±1.1, and -10.3±0.9 in P0, P1, P7, and P42 dogs, respectively (n=10 cells per group, P=NS).

The recovery kinetics of I_Ca were studied with a two-pulse protocol as shown in Fig 9B. The I_Ca recovery time course at -80 mV was not altered by rapid atrial pacing (n=10 cells per group). Recovery was well-fitted by a monoexponential function, with time constants averaging 31.4±4.5, 27.5±3.3, 31.3±2.8, and 29.8±2.9 milliseconds in P0, P1, P7, and P42 dogs, respectively (P=NS). The time course of I_Ca decay was fitted by a biexponential function, as previously reported.^{25 28} As shown in Fig 9C, the time constants of I_Ca inactivation were voltage dependent but not altered in paced dogs compared with control dogs (n=10 cells per group). The results shown in Fig 9 indicate that the reduction in I_Ca density caused by sustained rapid atrial pacing was not due to changes in the voltage dependence or kinetic properties of I_Ca.

Small inward currents compatible with T-type Ca²⁺ current were noted in control cells, as illustrated in Fig 10A. Ca²⁺ current recorded upon depolarization from -90 to -20 mV is shown by the circle; current recorded upon depolarization from -50 mV is indicated by the diamond. The current inactivated by reducing the holding potential, as determined by digital subtraction, is shown at the bottom of the panel. Corresponding results in a cell from a P42 dog are shown in Fig 10B. Mean±SEM current-voltage relations for T-type current in 10 P0 cells and 10 P42 cells are shown in Fig 10C. The threshold for current activation was -40 mV, and peak current occurred at -20 mV. Rapid pacing did not alter the amplitude or density of T-type current at any voltage. Fig 10D shows the voltage dependence of T-type Ca²⁺ current inactivation, as determined with 1-second prepulses to the voltages indicated, followed by a 240-millisecond test pulse to -20 mV. Inactivation voltage dependence was well-fitted by a Boltzmann relation, with a V_1/2 of -65±8 mV and slope factor of 5.9±0.5 mV in five P0 cells compared with a V_1/2 of -67±8 mV and slope factor of 5.1±0.5 mV in five P42 cells.

View larger version (23K): [in this window] [in a new window]   Figure 10. Properties of T-type Ca2+ current in cells from sham-operated and paced dogs. A, Recordings of Ca2+ current with HPs of -90 and -50 mV (pulse to -20 mV for 240 milliseconds, 0.1 Hz) in a P0 cell (top) and subtracted current (bottom). B, Recordings in same format as for panel A, obtained in a cell from a P42 dog. C, Mean±SEM current density-voltage relation for T-type current in 10 cells each from P0 and P42 dogs. D, Inactivation curve for T-type Ca2+ current, obtained with 1-second prepulses to the conditioning potential (CP) indicated, followed by a test pulse (240 milliseconds, 0.1 Hz) to -20 mV. Results are mean±SEM in five cells for each group.

Role of Ionic Changes in Action Potential Abnormalities Associated With Susceptibility to AF
The ionic studies described above indicate that rapid atrial pacing in dogs causes progressive decreases in I_Ca and I_to, without altering I_K1, I_Kr, I_Ks, I_Kur.d, T-type Ca²⁺ current, or I_Cl.Ca. In order to assess the potential contributions of changes in I_Ca and I_to to action potential alterations in paced dogs, we performed pharmacological studies with the use of AP clamp. Fig 11A through 11C shows the effects of increasing concentrations of nifedipine on the action potential waveform and corresponding currents from a representative P0 cell. The top panels are action potentials recorded at 0.1 Hz in current-clamp mode before and after nifedipine superfusion. Nifedipine caused a concentration-dependent decrease in APD. Mean values for APD under control conditions and at each nifedipine concentration are shown in Table 3. The bottom panels of Fig 11 show nifedipine-sensitive currents obtained by digital subtraction of AP-clamp recordings obtained with the control AP waveform as the voltage-command pulse in the presence of drug from those before drug exposure. The drug-sensitive current showed an initial rapidly decaying inward component, followed by a more sustained inward current whose decrease paralleled repolarization of the imposed (control) waveform. Nifedipine-sensitive current increased to reach a maximum at 10 µmol/L (mean values of peak drug-sensitive current are shown in the inset of Fig 11H). Maximal APD reduction was seen with a nifedipine concentration (10 µmol/L) that in standard voltage-clamp studies of five separate P0 cells inhibited 91±3% of I_Ca elicited by 300-millisecond step depolarizations from -70 to +10 mV at 1 Hz. Fig 11D shows the effects of 10 µmol/L nifedipine in an atrial cell from a dog paced for 42 days. In contrast to the effect in P0 cells, the drug produced only slight action potential abbreviation in the cell from the paced dog (mean APD₉₅ reduction by 10 µmol/L nifedipine was 7.9±1.4% in 10 cells from P42 dogs, P<.001 versus the 57±5% reduction produced by the same nifedipine concentration in control dogs), and only a small nifedipine-sensitive current flowed during the action potential (Fig 11H, bottom). AP clamp results similar to those shown in Fig 11G and 11H were obtained for five cells from P0 dogs and five cells from P42 dogs: the total amplitude of 10 µmol/L nifedipine-sensitive current averaged 809±86 pA in control cells and 118±26 pA in P42 cells, respectively (P<.001). These findings suggest that I_Ca depression is responsible for much of the action potential abbreviation in paced dogs.

View larger version (13K): [in this window] [in a new window]   Figure 11. Action potential recordings at 0.1 Hz (top) and corresponding drug-sensitive AP clamp recordings (bottom) from P0 dog cells exposed to 1 (A), 5 (B), and 10 (C) µmol/L nifedipine (Nif) and from a cell from a dog subjected to 42 days of rapid pacing exposed to 10 µmol/L Nif (D). Action potential recordings are shown before () and after () exposure to Nif. The control action potential waveform was used as a voltage-command pulse to record current before and after exposure to Nif at the concentrations indicated, and Nif-sensitive current (panels E through H) was obtained by digital subtraction of current in the presence of the drug from current in its absence in the same cell. Similar results were obtained in five P0 dog cells and five cells from P42 dogs. ***P<.001 vs N10. Mean±SEM Nif-sensitive currents are shown in the inset of panel H (N1, N5, and N10 indicate current in P0 cells sensitive to 1, 5, and 10 µmol/L Nif; P42,N10, 10 µmol/L–sensitive current in P42 cells).

View this table: [in this window] [in a new window]   Table 3. Effects of Nifedipine on APD in 10 Cells From P0 Dogs Studied at 0.1 Hz

We then sought to evaluate the possible role of I_Ca reductions caused by chronic rapid atrial pacing in the observed changes in the rate dependence of APD. Fig 12A shows the effect of increasing the rate from 0.1 to 2 Hz on the P0 canine atrial action potential. There was a striking acceleration in repolarization, reducing APD₉₅ by a mean of 84.2±6.4 milliseconds in 10 cells. Six weeks of rapid pacing reduced total APD and greatly attenuated rate-dependent APD₉₅ reduction (Fig 12B) to a mean of 1.4±0.1 milliseconds in 10 cells. Cells from P0 dogs exposed to nifedipine showed a substantial reduction in APD and a reduction in APD₉₅ abbreviation (Fig 12C) quantitatively similar (mean of 1.9±0.1 milliseconds in 10 cells) to those produced by rapid pacing. These results suggest that reductions in I_Ca can account for the attenuation in rate-dependent APD changes caused by chronic rapid atrial activation.

View larger version (17K): [in this window] [in a new window]   Figure 12. A through C, Examples of action potentials recorded at 0.1 Hz () and 2 Hz () in cells from a sham-operated dog (A), from a dog subjected to 42 days of rapid atrial pacing (B), and from a sham-operated dog after exposure of the cell to 10 µmol/L nifedipine (Nif) (C). Results similar to those illustrated in each panel were obtained in 10 cells for each group. D, Action potentials from a P42 cell at 1 Hz before (P42) and after (Bay K) exposure to 1 µmol/L Bay K 8644. E and F, Action potential recordings at 0.1 Hz before (control [Ctl], ) and after exposure to 50 µmol/L 4AP () and 2 mmol/L 4AP () in a P0 cell without the addition of Nif (E) and a P0 cell studied in the continuous presence of 10 µmol/L Nif (F).

If reductions in I_Ca are the cause of the loss of the plateau and the abbreviation of the action potential in paced dogs, increasing I_Ca in such cells should be able to restore the plateau. We evaluated this possibility in seven cells isolated from P42 dogs, from which we recorded action potentials at 1 Hz before and after exposure to 1 µmol/L Bay K 8644. Bay K 8644 restored a plateau in all cases and, in one instance, returned APD to the normal range (Fig 12D). Overall, Bay K 8644 increased APD₉₅ from 87±2 to 120±6 milliseconds (P<.001). Bay-K 8644 did not fully normalize APD (APD₉₅ in P0 dog cells at 1 Hz averaged 190 milliseconds; Table 1), consistent with its limited ability to increase I_Ca in P42 cells. The increase in I_Ca by Bay K 8644 was investigated with voltage-clamp steps to +10 mV in five P42 cells, in which the drug (1 µmol/L) increased peak I_Ca from 329±31 to 591±78 pA (P<.05), still substantially less than I_Ca in P0 myocytes, which averaged 810±89 pA. I_Ca in P42 cells exposed to Bay K 8644 was of the same order as I_Ca in P1 cells, in which APD₉₅ at 1 Hz averaged 134±8 milliseconds, similar to the value of 120±6 milliseconds in P42 cells treated with Bay K 8644.

The final set of experiments was performed to assess the potential impact on the action potential of I_to reduction caused by rapid pacing. A widely used tool to study I_to is 4AP, which strongly inhibits I_to at concentrations that have relatively little effect on a variety of other K⁺ currents.²⁹ However, I_Kur.d is very sensitive to 4AP.²¹ Therefore, we first exposed P0 cells to 50 µmol/L 4AP, which fully inhibits I_Kur.d with minimal effects on I_to.²¹ We then superfused the cells with 2 mmol/L 4AP, which causes no additional change in I_Kur.d but fully inhibits I_to.²¹ A representative example is shown in Fig 12E. The lower 4AP concentration raised the plateau and prolonged APD, whereas the higher concentration further raised the plateau and accelerated phase-3 repolarization, causing a net decrease in APD. The effect of blocking I_to on the action potential is indicated by comparing results in the presence of 50 µmol/L 4AP (block of I_Kur.d alone) with those in the presence of 2 mmol/L 4AP (block of I_Kur.d and I_to). As shown by the mean data in Table 4, I_to inhibition reduces APD in P0 cells; however, this may not reflect the result of I_to inhibition in paced dogs, in whom the action potential is importantly altered by a reduced I_Ca. In order to obtain an indication of the potential effect of reduced I_to in the presence of reduced I_Ca, we exposed cells from P0 dogs to 10 µmol/L nifedipine, followed by 50 µmol/L 4AP and then 2 mmol/L 4AP. Typical results (Fig 12F) show small concentration-related increases in APD. The mean data shown in Table 4 indicate that in the presence of 10 µmol/L nifedipine, inhibition of I_Kur.d with 50 µmol/L 4AP increases APD₉₅ to a modest extent (28±2%). Block of I_to (comparing results with 2 mmol/L 4AP, representing block of I_to and I_Kur.d, with those at 50 µmol/L 4AP, blocking I_Kur.d alone) caused a small (10-millisecond) nonsignificant further increase in APD₉₅. Thus, the reduction in I_to occurring in the presence of strongly reduced I_Ca in paced dogs is unlikely to have contributed importantly to changes in the action potential.

View this table: [in this window] [in a new window]   Table 4. Effects of 4AP on APD at 0.1 Hz

	Discussion

Top Abstract Introduction Materials and Methods Results Discussion References

 
We have shown that rapid atrial pacing decreases both APD and APD adaptation to rate. The APD changes are qualitatively similar to, and likely account for, alterations in atrial refractoriness and are accompanied by decreases in I_Ca and I_to without alterations in other ion channel currents flowing during the action potential plateau. The evidence for an important role of reduced I_Ca in the action potential abnormalities in paced dogs includes (1) the similar effects of nifedipine and rapid pacing on action potential morphology, (2) the lack of effect of nifedipine on action potential morphology in cells from dogs exposed to 42 days of rapid pacing, (3) the similar changes in APD adaptation to rate in cells from rapidly paced dogs and cells from sham-operated dogs exposed to nifedipine, and (4) the ability of the I_Ca agonist Bay K 8644 to restore the plateau in cells from rapidly paced dogs.

Comparison With Previous Findings in Models of AF
We are not aware of previous studies of cellular and ionic mechanisms in a chronic animal AF model. Our findings of reduced atrial ERP are consistent with those of Morillo et al¹⁸ and Wijffels et al¹⁷ in animals prone to AF because of chronic atrial tachycardias. In addition, Wijffels et al noted a progressively decreased ERP adaptation to rate in their goat model of tachycardia-induced AF. Preliminary analyses of epicardial mapping data in dogs subjected to chronic rapid atrial pacing point to a role for alterations in atrial ERP, including reduced ERP abbreviation at rapid rates, in the occurrence of multiple-circuit reentry underlying sustained AF.³⁰ Our observations provide likely explanations for ERP abnormalities detected in these earlier studies in terms of both cellular (changes in APD) and ionic (alterations in I_Ca) mechanisms.

Patients with increased vulnerability to AF have also been found to have reduced atrial ERP and decreased ERP adaptation to rate.¹⁹ Underlying ionic mechanisms have not been assessed in AF patients, but some interesting data are available from patients with atrial dilation, a population known to be predisposed to AF. Both I_to^{31 32} and I_Ca^{32 33} are decreased in atrial cells of patients with atrial dilation. Moreover, both absolute APD and APD accommodation to changes in rate are reduced in cells from dilated atria.³² These findings are qualitatively similar to changes that we observed in dogs with increased susceptibility to AF. Koumi et al³⁴ have reported that I_K1 is reduced in myocytes of dilated atria from patients with congestive heart failure. Sakakibara et al³⁵ have indicated (without showing data) that I_Na properties are not altered in patients with presumed atrial disease compared with those without such a presumption.

Comparison With Previous Studies of Ionic Changes in Animal Models of Heart Disease
Although no data are available in the literature regarding ionic changes in animals with atrial disease, a number of studies have been performed on ventricular myocytes of animals with experimentally induced ventricular pathology. In contrast to the reduced ERP and APD seen with atrial disease, APD is increased in ventricular myocytes from failing^{36 37} and hypertrophied^{38 39} hearts. I_to is decreased in dogs with pacing-induced heart failure³⁶ and cardiomyopathic Syrian hamsters.³⁷ Conflicting results have been obtained in studies of I_Ca in ventricular cells of dogs with pacing-induced ventricular failure, with either no change³⁶ or a decrease⁴⁰ having been reported. Similarly, both no change⁴¹ and a decrease³⁸ in I_Ca density have been reported in rats several weeks after acute myocardial infarction, whereas decreased I_Ca has been reported in dogs with 5-day-old infarctions.⁴² In summary, decreases in both I_to and I_Ca have been reported in ventricular myocytes from animals with chronic ventricular disease, but discrepancies remain, particularly for I_Ca.

One methodological aspect of our work that differs from previous studies of ionic changes in diseased cardiac tissue is the use of pharmacological tools and the AP-clamp method to relate critically ionic changes to alterations in the action potential. Many previous studies failed to record action potentials. Only in the work of Kääb et al³⁶ was an effort made to relate ionic changes to action potential alterations, with the use of 4AP and hyperpolarizing current to evaluate the potential importance of I_to suppression in the APD prolongation caused by pacing-induced left ventricular failure. The results shown in Fig 11 of the present study indicate that I_Ca suppression underlies APD abbreviation in paced dogs, whereas those in Fig 12A through 12C point to a prominent role of I_Ca reduction in altered APD adaptation to rate. The observations in Fig 12 suggest that changes in I_to are not likely to contribute importantly to action potential changes in dogs subjected to chronic rapid atrial activation.

Comparison With Previous Observations of Cellular Abnormalities in AF
There is relatively little published information about cellular abnormalities in chronic animal models of AF. Boyden and Hoffman⁴³ showed that action potentials from right atria of dogs with tricuspid insufficiency and atrial tachyarrhythmias (predominantly atrial flutter) were not significantly different from control right atria. Similarly, no significant differences were noted between control atrial action potentials and those of dogs with chronic mitral regurgitation and AF,⁴⁴ and the same was true for right atrial preparations of cats with spontaneous cardiomyopathy and atrial arrhythmias.⁴⁵ Left atrial preparations from moderately or severely dilated left atria of cardiomyopathic cats showed reduced resting potential, action potential amplitude, and phase-0 upstroke velocity, along with increased APD.⁴⁵ Preliminary findings have been published suggesting that atrial monophasic APD is markedly reduced in goats with chronic electrically maintained AF and that APD abbreviation in response to tachycardia is absent and may even be reversed.⁴⁶ We were unable to find published reports of changes in ionic currents in animal models of atrial tachyarrhythmia.

Patients with enhanced atrial vulnerability have decreased atrial ERP adaptation to rate,¹⁹ and patients with chronic AF appear to have reduced ERP and monophasic APD in the right atrial appendage.⁴⁷ Microelectrode studies in right atrial appendages of patients with chronic AF show a significant reduction in APD and in APD adaptation to rate compared with non-AF controls and have a morphology similar to those of our rapidly paced dog cells, with a loss of the action potential plateau.^{48 49} Van Wagoner et al⁵⁰ reported a decrease in I_to and sustained outward current, but not I_K1, density in patients with AF, consistent with our findings.

Potential Mechanisms of Ionic Changes
The mechanisms of ionic changes caused by heart disease are presently unknown. In the present study, whereas I_to and I_Ca density were reduced by sustained rapid atrial pacing, other biophysical properties of the currents, including voltage and time dependence, were unaltered. This finding suggests a decrease in the number and/or conductance of I_to and I_Ca channels, without a change in their fundamental nature, and is consistent with previous work suggesting that I_to reductions in pacing-induced heart failure are due to a decrease in the number of I_to channels without other changes in their properties.³⁶ The molecular basis for decreases in currents associated with cardiac pathology remains unknown, as does the mechanism(s) that stimulates changes in channel expression. The latter could include regional myocardial ischemia, altered ionic fluxes, changes in wall tension, and alterations in autonomic nervous system function.

Limitations of the Model
There is evidence that clinical AF alters atrial properties to promote further AF ("AF begets AF"),¹⁷ and many of the electrophysiological changes in the canine tachycardia model^{16 18 30} resemble those in goats with electrically maintained AF.¹⁷ However, it cannot be assumed that the rapid-pacing dog model of AF is directly analogous to clinical AF, any more than the tachycardia-induced ventricular failure model can be directly extrapolated to all forms of clinical congestive heart failure. On the other hand, the rapid atrial pacing animal model of AF has a variety of electrophysiological features in common with patients predisposed to the arrhythmia.^{10 11 12 19 32} Furthermore, the ionic changes noted in our dogs resemble those reported in patients with atrial dilation,^{31 32 33} and the action potential changes we observed are similar to those previously reported in patients with AF.^{48 49} The decrease in I_to and lack of change in I_K1 that we noted are similar to observations in patients with AF by Van Wagoner et al.⁵⁰ Further work clearly needs to be performed to assess directly the ionic abnormalities in atrial myocytes of patients with AF, in order to determine more clearly the relevance of our observations to the ionic mechanisms underlying AF in various clinical populations.

In the present study, we have focused on action potential abnormalities underlying the ERP changes in the model, along with underlying mechanisms. ERP changes of the type we observed have been noted in both dogs^{16 18 30} and goats¹⁷ with a substrate for AF, as well as in patients with enhanced atrial vulnerability.¹⁹ In all of these models, ERP changes were felt to be important in promoting the maintenance of AF.^{17 18 19} On the other hand, it is possible (even likely) that other electrophysiological abnormalities, such as changes in conduction^{23 30} and in heterogeneity of refractoriness,⁵¹ also play a role. It should be noted that at short cycle lengths closest to those of AF, both APD and ERP are little changed in rapidly paced dogs (eg, see Fig 1E and 1F), suggesting a potentially important role for other factors in AF maintenance. The evaluation of such additional electrophysiological changes, and of potential underlying ionic mechanisms, is beyond the scope of the present study.

Our observation that I_Cl.Ca is not altered in rapidly paced dogs is puzzling, in view of the decrease in I_Ca. The magnitude of I_Cl.Ca is controlled in a complex way by a variety of factors, including the I_Ca trigger for Ca²⁺ release, the density of I_Cl.Ca channels, the Ca²⁺ load in the sarcoplasmic reticulum, sarcoplasmic reticulum functioning, and the physical proximity between elements of the system (Ca²⁺ channels, sarcoplasmic reticulum Ca²⁺ release channels, and Ca²⁺-dependent Cl^- channels). Further work would be necessary to analyze all elements of the system in order to determine why I_Cl.Ca remains constant in paced dogs despite decreased I_Ca.

We worked with a canine model, and canine atrial cells share many ionic properties with human atrial myocytes. These include the presence, order of magnitude, voltage dependence, and kinetic properties of I_to, I_Kr, and I_Ks.^{20 52 53 54} The role of I_Ca in rate-dependent changes in human atrial APD²⁵ is very similar to that noted for canine atrial myocytes in the present study. Like canine atrium, human atrial cells have a significant ultrarapid delayed rectifier current,⁵² although there are differences in rectification properties, sensitivity to tetraethylammonium, and probable molecular basis. On the other hand, human atrium does not appear to possess I_Cl.Ca⁵⁵ or T-type Ca²⁺ currents.²⁵ These ionic differences, as well as the possibility that there are others not yet defined, must be considered when trying to relate our findings to humans.

Potential Significance of Our Findings
Decreased atrial contractility upon conversion to sinus rhythm is a common feature in patients with chronic atrial tachyarrhythmias, a phenomenon sometimes referred to as "atrial stunning."^{56 57} The resulting atrial stasis may be important in promoting thromboembolic phenomena⁵⁸ and may delay improvements in physical capacity after electrical rhythm reversion.⁵⁶ The mechanisms underlying contractile dysfunction following cardioversion of AF are currently unknown. Our results point toward the possibility that the rapid atrial activation rates during AF substantially decrease I_Ca, which could account for an important decrease in contractility.

The present study is the first of which we are aware to establish the ionic and cellular mechanisms associated with susceptibility to sustained AF in a well-defined animal model. Clinical approaches to AF remain limited because of inadequate efficacy and/or adverse consequences of available therapeutic avenues.⁵⁹ The development of improved pharmacological approaches will require a better understanding of underlying ionic mechanisms. Although much has been learned over the past few years about the ionic determinants of normal human atrial repolarization,⁶⁰ relatively little is known about how these properties are altered in patients with AF. The latter may have an important impact on the response to drugs designed to inhibit specific channels whose expression may be altered in AF. Furthermore, a better understanding of the ionic mechanisms leading to AF and underlying molecular triggers may allow for novel therapies to be devised that prevent or reverse the development of the substrate underlying AF. The model we used is potentially helpful in studying the development of the ionic substrate of AF, by virtue of its gradual and predictable progression, and our findings are valuable in showing the ionic changes that lead to AF susceptibility in a well-defined animal model. In addition, rapid atrial activation is a feature of clinical AF, so it is likely that the ionic changes observed in our canine model are relevant to electrical remodeling in patients with AF.

	Selected Abbreviations and Acronyms

 

AF = atrial fibrillation 4AP = 4-aminopyridine APD = action potential duration APD20, APD50, APD95 = APD at 20%, 50%, and 95% repolarization ERP = effective refractory period HP = holding potential ICa = L-type Ca2+ current ICl.Ca = Ca2+-dependent Cl- current IK = classical delayed rectifier current IK1 = inward rectifier K+ current IKr, IKs = rapid and slow components of IK IKur.d = ultrarapid delayed rectifier current (dog) INa = Na+ current Ito = transient outward current I-V = current-voltage P0, P1, P7, P42 = dogs subjected to sham operation and 1, 7, and 42 days of atrial pacing Rs = series resistance TEA = tetraethylammonium V1/2 = half-maximal inactivation voltage

	Acknowledgments

 
This study was supported by the Medical Research Council of Canada, the Quebec Heart Foundation, and the Fonds de Recherche de l'Institut de Cardiologie de Montreal (FRICM). Lixia Yue is the recipient of a studentship award from the Canadian Heart Foundation, and Rania Gaspo is the recipient of a Medical Research Council of Canada/Pharmaceutical Manufacturer's Association of Canada postdoctoral fellowship award. The authors wish to thank Diane Campeau for excellent secretarial assistance.

	Footnotes

 
Previously published in abstract form (Circulation. 1996;94[suppl I]:I-592).

Received November 25, 1997; accepted July 8, 1997.

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