Vol. 275, Issue 5, H1717-H1725, November 1998
Ultrarapid delayed rectifier current inactivation in human
atrial myocytes: properties and consequences
Jianlin
Feng1,2,
Donghui
Xu1,2,
Zhiguo
Wang1,2,3, and
Stanley
Nattel1,2,3,4
1 Department of Medicine and
2 Research Center, Montreal Heart
Institute, Montreal H1T 1C8;
3 Department of Medicine,
University of Montreal, Montreal H3C 3J7; and
4 Department of Pharmacology and
Therapeutics, McGill University, Montreal H3G 1Y6, Quebec, Canada
 |
ABSTRACT |
The ultrarapid delayed rectifier current
(IK,ur) plays a
significant role in human atrial repolarization and is generally believed to show little rate dependence because of slow and partial inactivation. This study was designed to evaluate in detail the properties and consequences of
IK,ur
inactivation in isolated human atrial myocytes.
IK,ur inactivated
with a biexponential time course and a half-inactivation voltage of
7.5 ± 0.6 mV (mean ± SE), with complete inactivation
during 50-s pulses to voltages positive to +10 mV (37°C). Recovery
from inactivation proceeded slowly, with time constants of 0.42 ± 0.06 and 7.9 ± 0.9 s at 80 mV (37°C). Substantial
frequency dependence was observed at 37°C over a clinically
relevant range of frequencies. Inactivation was faster and occurred at
more positive voltages at 37°C compared with room temperature. The
voltage and time dependencies of Kv1.5 inactivation were studied in
Xenopus oocytes to avoid overlapping currents and strongly resembled those of
IK,ur in native
myocytes. We conclude that, while
IK,ur
inactivation is slow, it is extensive, and slow recovery from
inactivation confers important frequency dependence with significant
consequences for understanding the role of
IK,ur in human
atrial repolarization.
potassium channels; action potentials; atrial fibrillation; cardiac
electrophysiology; antiarrhythmic drugs
| |
INTRODUCTION |
A VARIETY OF STUDIES have indicated the presence in
human atrial myocytes of a rapidly activating, highly 4-aminopyridine (4-AP)-sensitive K+ current with
slow and partial inactivation (1, 10, 11, 21). This current has been
variously referred to as
IK,ur, for "ultrarapid delayed rectifier
K+ current" (21), or
Iso, for
"sustained outward current" (1, 11). We refer to the macroscopic
current as IK,ur
in the present manuscript. The voltage- and time-dependent properties
of IK,ur, along
with its sensitivity to small 4-AP concentrations (1, 10, 21), identify
it with currents resulting from the expression of the Kv1.5
K+ channel subunit (7, 19). The
role of Kv1.5 channel subunits in carrying
IK,ur is further
supported by the specific downregulation of
IK,ur in cultured
human atrial myocytes exposed to antisense oligodeoxynucleotides
directed to the NH2-terminal
coding sequence of Kv1.5 (8).
One element of the physiology of
IK,ur that
remains poorly understood is its inactivation properties. Kv1.5
currents show significant, albeit slow, inactivation (17, 19).
IK,ur
inactivation is considered to be slow and partial (1, 10, 11, 21); however, the longest depolarizing pulses used to characterize IK,ur
inactivation have been 4 s in duration, and steady-state inactivation
did not appear to be achieved (1). There is reason to believe that
IK,ur
inactivation may be physiologically relevant, because the end-pulse
current during 80-ms depolarizing pulses (which contains a significant
contribution of
IK,ur; see Refs. 1, 10, 11, 21) shows a statistically significant decrease at rapid
pulsing frequencies (9). We therefore set out to evaluate in detail the
inactivation properties of
IK,ur, with the
use of 50-s depolarizing pulses that allow for a much more accurate
assessment of inactivation than was previously possible with
shorter-duration pulses. In particular, we sought to clarify the
voltage and time dependence of the development of
IK,ur
inactivation at room temperature and body temperature (37°C), the
time dependence of current recovery from inactivation at potentials
comparable to the normal atrial resting potential, 80 mV (23),
and the frequency dependence of
IK,ur at a
holding potential of 80 mV and physiological temperature. IK,ur was
isolated from other currents, particularly the transient outward
current (Ito),
with the use of previously described protocols that rely on the slower
inactivation kinetics and greater 4-AP sensitivity of
IK,ur compared
with Ito (21).
For additional confirmation, we compared the inactivation properties
that we observed for
IK,ur with those
of currents carried by Kv1.5 subunits during 50-s depolarizing pulses
in Xenopus oocytes.
| |
METHODS |
Cell isolation. Specimens of human
right atrial appendage were obtained from the hearts of 30 patients (61 ± 2 yr old, range 42-75 yr) undergoing aortocoronary bypass
surgery. The procedure for obtaining the tissue was approved by the
Ethics Committee of the Montreal Heart Institute. Samples were immersed
in nominally Ca2+-free Tyrode
solution (100% O2, 37°C) of
the following composition (in mM): 136.0 NaCl, 5.4 KCl, 1.0 MgCl2, 0.33 NaH2PO4,
10 dextrose, and 10 HEPES (Sigma), pH adjusted to 7.4 with NaOH. The
myocardial specimens were chopped with scissors into cubic chunks and
placed in a 25-ml flask containing 10 ml of the
Ca2+-free Tyrode solution. The
tissue was gently agitated by continuous bubbling with 100%
O2 and stirring with a magnetic
bar. After an initial 5 min in this solution, the chunks were
reincubated in a similar solution containing 200 U/ml collagenase (CLS
II; Worthington Biochemical) and 4 U/ml protease (type XXIV; Sigma). The first supernatant was removed after 45 min and discarded. The
chunks were then reincubated in a fresh enzyme-containing solution.
Microscopic examination of the medium was performed every 15 min to
determine the number and quality of the isolated cells. When the yield
appeared to be maximal, the chunks were suspended in a storage solution
of the following composition (in mM): 20 KCl, 10 KH2PO4,
10 glucose, 70 glutamic acid, 10 
-hydroxybutyric acid, 10 taurine,
10 EGTA, and 0.1% albumin, pH adjusted to 7.4 with KOH, and gently
pipetted. 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 were allowed for cell adhesion to
the bottom of the chamber, and then the cells were superfused at 3 ml/min with a solution containing (in mM) 136.0 NaCl, 5.4 KCl, 0.8 MgCl2, 1.0 CaCl2, 0.33 NaH2PO4, 10 HEPES, and 5.5 glucose, pH adjusted to 7.4 with NaOH. Experiments were conducted at room temperature (23-25°C) or at 37°C
(with the use of a Peltier-effect device). All studies were performed within 12 h of the completion of cell isolation.
Whole cell patch-clamp methods. The
whole cell patch-clamp technique was employed to record ionic currents
in the voltage-clamp mode. Borosilicate glass electrodes (1.0-mm OD)
were used, with tip resistances of 1.5-3 M
when filled with (in
mM) 0.1 GTP, 110 potassium aspartate, 20 KCl, 1.0 MgCl2, 10 HEPES, 5 EGTA, 5 Mg2ATP, and 5 Na2-creatine phosphate (pH
adjusted to 7.4 with KOH) and connected to a patch-clamp amplifier
(Axopatch 200A; Axon Instruments). Command pulses were generated by a
12-bit digital-to-analog converter controlled by pCLAMP software
(Axon). Recordings were low-pass filtered at 5 kHz and stored on the
hard disk of an IBM compatible computer.
Offset voltages generated when the pipette was inserted in Tyrode
solution (2-8 mV) were zeroed before formation of the
membrane-pipette seal. Mean seal resistance averaged 10.9 ± 1.8 G (n = 35). Several minutes after seal formation, the membrane was ruptured by gentle suction to establish the whole cell configuration for voltage clamping.
The series resistance
(Rs) was
estimated by dividing the time constant obtained by fitting the decay
of the capacitive transient by the calculated cell membrane capacitance
(the time integral of the capacitive surge measured in response to 5-mV hyperpolarizing steps from a holding potential of 60 mV, divided by the voltage drop). Before
Rs compensation,
the capacitive time constant was 548 ± 34 µs (cell capacitance:
79 ± 3.8 pF, n = 35). After
Rs compensation,
the time constant was reduced to 145 ± 12 µs. The initial
Rs was calculated
to be 6.8 ± 0.2 M, and
Rs was reduced to
2.0 ± 0.1 M after compensation. Currents recorded during this
study rarely exceeded 2 nA, and the voltage drop across Rs did not exceed
5 mV. Cells with significant leak currents were rejected, and leakage
compensation algorithms were not used.
To minimize possible contamination from delayed rectifier
(IK), inward
rectifier
(IK1), and
acetylcholine-dependent
(IK,ACh) currents, the following chemicals were used in the extracellular solution for
IK,ur recording:
tetraethylammonium chloride (10 mM, to inhibit
IK; Sigma),
atropine (100 nM, to inhibit
IK,ACh; Sigma),
and CdCl2 [200 µM, to
block the Ca2+ current
(ICa);
Sigma]. The sodium current
(INa) was
suppressed by isomolar replacement with choline chloride (Sigma) for
NaCl in the bath solution. In some experiments,
IK1 was
suppressed with the use of 0.5 mM
BaCl2, which does not affect
IK,ur (21).
Functional expression of Kv1.5 in Xenopus
oocytes. The Kv1.5 cDNA was subcloned into pSP64
(Promega) for oocyte expression. cRNA for injection into oocytes was
prepared as previously described (22) with the mMESSAGE mMACHINE kit
(Ambion) using SP6 polymerase after linearization of the plasmid with
EcoR I. The samples were dissolved in
0.1 M KCl, stored at 80°C, and diluted immediately before
injection. Stage V-VI Xenopus oocytes
were injected with 46 nl of cRNA (~300 ng).
Whole cell two-microelectrode voltage-clamp
recording. Whole cell macroscopic currents were
recorded with conventional two-electrode techniques as previously
described (6). Microelectrodes were filled with 3 M KCl and had
resistances in the range of 0.8 M. The bath solution contained (in
mM) 100 NaCl, 5 KCl, 0.3 CaCl2, 2 MgCl2, and 10 HEPES. The pH was
adjusted to 7.4 with Tris base. Signals were acquired with a Geneclamp
amplifier, digitized at 10 kHz, and low-pass filtered at 3 kHz.
Experiments were conducted at room temperature (20-22°C).
Data analysis. Group data are
presented as the means ± SE. Nonlinear curve fitting was performed
with the Clampfit routine in pCLAMP 6 (Chebyshev algorithm). Ionic
current data, which are displayed along with fitted curves in Figs.
1-7, are represented with the use of the data reduction algorithm
in pCLAMP 6, so that the relation between fitted curves and data is
clear; if all data points were shown, fitted curves would be obscured
by data points. Statistical comparisons among groups were performed
with analysis of variance (ANOVA). If significant effects were
indicated by ANOVA, a t-test with
Bonferroni correction or a Dunnett's test was used to evaluate the
significance of differences between individual means. A two-tailed
P < 0.05 was taken to indicate
statistical significance.
| |
RESULTS |
Time-dependent inactivation of
IK,ur.
When a 100-ms pulse was applied to +40 mV, 5 ms after a prepulse to +40
mV to inactivate
Ito, a rapidly
activating and slowly inactivating current was observed at 37°C,
along with a small tail current during an 80-ms repolarization to
20 mV (Fig.
1A). These findings are typical of
IK,ur (21).
Figure 1B shows a recording obtained
with a 50-s pulse to +40 mV after a similar prepulse. During the
prolonged pulse, considerable inactivation is observed. When the data
shown in Fig. 1B were fitted by
nonlinear curve-fitting methods, the best fit was provided by a
biexponential equation. The curve fit to the data in Fig.
1B is shown in Fig. 1C, with time constants of 1.0 and 6.8 s. Note that extensive data reduction was used to provide the data
points in Fig. 1C, otherwise, the
fitting would be totally obscured by overlying data points; however,
curve fitting was performed with the complete set of data shown in Fig.
1B. A similar approach is taken to
illustrate curve fits to inactivation data throughout this paper (e.g.,
see Figs. 3A and
7A).
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Fig. 1.
Ultrarapid delayed rectifier K+
current (IK,ur)
inactivation during depolarizing pulses at 37°C.
A:
IK,ur recorded
with a 100-ms pulse to +40 mV, preceded by a 100-ms prepulse (pp) to
+40 mV to inactivate transient outward current
(Ito).
B:
IK,ur recorded
with the use of a 50-s pulse to +40 mV. A 100-ms prepulse was used to
inactivate Ito.
C: best biexponential curve fit to
data in B. Data reduction was used to
provide the data points shown, to illustrate the fit without obscuring
the fitted curve with overlapping data points. TP, test potential.
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To evaluate further the possible nature of the slowly but extensively
inactivating component, we studied its response to the K+ channel blocker 4-AP. Figure
2A shows
the response to 4-AP of current elicited by a 50-s pulse to +40 mV,
after a 100-ms prepulse to inactivate
Ito. A large
inactivating component was seen before 4-AP. Exposure to 10 µM 4-AP
substantially depressed the inactivating current. At 250 µM, 4-AP
fully suppressed the inactivating current, leaving only a
time-independent component. Note that 4-AP had no effect on current at
the end of the 50-s pulse (i.e., there was no sustained component to
4-AP-sensitive current). Figure 2B
shows the concentration dependence of 4-AP inhibition of the inactivating current. The latter was measured on the basis of the
difference between peak and end-pulse current during a 50-s pulse to
+40 mV, with a 100-ms prepulse used to eliminate
Ito. The
concentration-response curve at 37°C (mean ± SE of 6 cells) provided an EC50 for current
inhibition of 10.2 ± 1.2 µM, and virtually complete inhibition of
current was seen at 100 µM. Results at room temperature are shown in
Fig. 2B (mean ± SE of 6 cells) and
indicate an EC50 of 21.6 ± 2.3 µM, significantly higher (P < 0.01) than at 37°C. The 4-AP sensitivity of the inactivating current is compatible with that previously reported for
IK,ur (1, 13,
21), with reported EC50 values in
the range of 6-50 µM at room temperature and full inhibition at
250-500 µM. The properties of the slowly inactivating current
therefore identify it with
IK,ur. Note that
there was no significant 4-AP-sensitive sustained component at 37°C
after 50 s at +40 mV. Mean current at 37°C inhibited by 500 µM
4-AP at the end of a 50-s pulse averaged 4 ± 1 pA, compared with
819 ± 97 pA for peak 4-AP-sensitive current after a 100-ms prepulse
to inactivate
Ito, and 821 ± 97 pA for inactivating current during a 50-s pulse after a 100-ms
prepulse to remove
Ito
(n = 6). These results show that
IK,ur inactivates completely during 50-s pulses and that the size of the inactivating current during a 50-s pulse (after a prepulse to inactivate
Ito) is an
accurate reflection of
IK,ur amplitude
as measured by 4-AP-sensitive current. Because a 50-s pulse was
sufficient to achieve complete inactivation of
IK,ur at 37°C
and near-complete inactivation at room temperature, 50-s pulses were
used to study
IK,ur
inactivation properties. At least 1 min was allowed at the holding
potential between pulse protocols to ensure full recovery from
inactivation.
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Fig. 2.
A:
IK,ur during 50-s
pulses at 37°C before and after exposure to 10 and 250 µM
4-aminopyridine (4-AP). B:
concentration-response curve for 4-AP inhibition of
IK,ur at room
temperature (RT) and 37°C (n = 6 cells exposed to all concentrations at each temperature, results are
means ± SE).
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Time and voltage dependence of
IK,ur inactivation.
To analyze the time dependence of
IK,ur
inactivation, we applied 50-s pulses from 50 mV to test
potentials between 0 and +50 mV. Experiments were performed at both
room temperature and 37°C, with both temperatures studied for each
cell, and a 100-ms prepulse to +40 mV was delivered 5 (at 37°C) or
10 (at room temperature) ms before the test pulse to inactivate
Ito. Figure
3A
illustrates IK,ur
inactivation during a 50-s pulse to +40 mV in one cell. Note that
inactivation is much faster at 37°C than at room temperature: inactivation was still proceeding (albeit very slowly) at the end of
the 50-s pulse at room temperature, whereas it was completed within
~20 s at 37°C. Data were fitted by biexponential functions of the form I = Afexp(t/
f) + Asexp(t/s),
as illustrated in Fig. 3A, where
I indicates current,
Af is the
amplitude of the fast component,
As is the
amplitude of the slow component, t is time, f is the time constant of
the fast component, and s is the time constant of the slow component. This analysis provided the
mean ± SE time constants for 12 cells per group shown in Fig. 3B. The fast-phase time constants were
significantly voltage dependent at both temperatures
(P < 0.05, ANOVA), whereas
slow-phase time constants were not voltage dependent. The time
constants were much faster at 37°C; for example, at +40 mV the time
constants averaged 1.0 ± 0.1 and 6.8 ± 0.9 s at 37°C
compared with 2.3 ± 0.4 and 22.2 ± 1.3 s
(P < 0.01 for each) at room
temperature. The relative amplitude
(Af/As)
of fast and slow phases of inactivation were also temperature
dependent, as shown in Fig. 3C, with
faster inactivation comprising a larger proportion of overall
inactivation at higher temperatures. Figure
3D shows the results of experiments designed to assess the development of
IK,ur
inactivation at 37°C and +40 mV with the use of conditioning pulses
of varied duration, followed 5 ms later by a 100-ms prepulse to
inactivate Ito
and then 5 ms later by a 100-ms test pulse to +40 mV to elicit
IK,ur. This
protocol was applied before and after the addition of 500 µM 4-AP,
with IK,ur
quantified on the basis of 4-AP-sensitive current during the test
pulse. The fast and slow time constants averaged 1.3 ± 0.3 and 6.3 ± 0.8 s, and
Af and
As averaged 0.65 ± 0.07 and 0.35 ± 0.05, respectively.
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Fig. 3.
A:
IK,ur
inactivation during a 50-s pulse to +40 mV, preceded by a 100-ms
prepulse to +40 mV to inactivate
Ito (interval
between prepulse and test pulse was 5 ms at 37°C, 10 ms at room
temperature). Best-fit biexponentials to current data are shown; data
points shown for room temperature and 37°C were obtained from the
original data set (to which the curves were fitted) with the use of a
data reduction algorithm so as to avoid obscuring the fitted curves by
overlapping data (approach illustrated in Fig. 1,
B and
C). Results shown were obtained from
the same cell at both temperatures. B:
time constants (mean ± SE, n = 12 cells/observation) of
IK,ur
inactivation at room temperature and 37°C, as obtained with curve
fits of the type shown in A.
*** P < 0.001 vs. value at
37°C. C: ratio of the amplitudes
of fast (Af)
and slow (As)
phases of IK,ur
inactivation at room temperature and 37°C.
D: development of
IK,ur
inactivation as studied with conditioning pulses of varying duration
(CPD), followed 5 ms later by a 100-ms prepulse to inactivate
Ito and then a
100-ms test pulse to +40 mV. Protocol was applied before and after 500 µM 4-AP, and the 4-AP-sensitive current was used to distinguish
IK,ur from
background and leak currents. Results are means ± SE for 6 cells,
and the best-fit biexponential relation is shown. TP, test pulse
potential; s, time constant of
the slow component; f, time
constant of the fast component.
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To evaluate the voltage dependence of
IK,ur
inactivation, we used 50-s prepulses from the holding potential of
80 mV to various voltages, followed by 240-ms test pulses to +40
mV (protocol delivered every 60 s). Because
Ito inactivates
fully within 100 ms at 37°C (21), the current at the end of the
test pulse consists of
IK,ur, nonselective cation current (5), and a small leak component. We
separated IK,ur
from the other currents on the basis of its sensitivity to 4-AP, by
performing experiments in the absence and presence of 500 µM 4-AP and
taking the 4-AP-sensitive end-pulse component as a reflection of
IK,ur. Figure
4A shows
currents recorded during the 240-ms test pulse from a representative
cell at 37°C. Figure 4B shows
results from the same cell with the same protocol, after the addition
of 500 µM 4-AP to the bath solution. The 4-AP-sensitive current from
the same cell is shown in Fig. 4C.
Mean data for six cells studied at 37°C are shown in Fig.
4D. Total current amplitude (measured
at the end of the pulse) decreases as prepulse voltage becomes more
positive, with a steady-state value attained at about +10 mV. The
4-AP-resistant component shows no voltage-dependent change, whereas the
4-AP-sensitive component shows voltage-dependent inactivation that is
complete at +10 mV. Results were fitted by Boltzmann functions as
shown. Similar experiments were conducted at room temperature, giving
the mean data from six cells shown in Fig.
4E. The results at room temperature
are qualitatively similar to those at 37°C. Figure
4F shows the normalized inactivation curves for 4-AP-sensitive current at room temperature and 37°C. The
half-inactivation voltage averaged 23.1 ± 2.6 mV at room temperature and 7.5 ± 0.6 mV at 37°C
(P < 0.001).
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Fig. 4.
Determination of the voltage dependence of
IK,ur
inactivation. Pulses (50-s duration) to various conditioning potentials
were followed by a 240-ms test pulse to +40 mV.
IK,ur at the end
of the test pulse was separated from nonselective cation current and
any leak by performing the protocol before and after the addition of
500 µM 4-AP, with
IK,ur being the
4-AP-sensitive component. A: currents
recorded in one cell at 37°C during the 240-ms test pulse after
50-s conditioning pulses to 100 mV (largest current) and
50, 30, 10, +10, and +30 mV (consecutively
decreasing currents), with the use of the protocol shown in
B. B:
currents from the same cell and protocol in
A recorded during the conditioning and
240-ms test pulse after the addition of 500 µM 4-AP.
C: 4-AP-sensitive currents obtained by
subtracting currents in B from those
in A.
D: mean ± SE currents at the end
of the 240-ms test pulse recorded in 6 cells at 37°C with the
approach illustrated in A-C. Results
are shown for total current, current in the presence of 4-AP, and
4-AP-sensitive current obtained by digital subtraction.
E: results obtained in the same
fashion and shown in the same format as in
D but at room temperature.
F: best-fit Boltzmann functions to
4-AP-sensitive current (mean ± SE of 6 cells each) at room
temperature and at 37°C, with data normalized to maximum current
under each condition. 4-AP-res, 4-AP resistant; 4-AP-sens, 4-AP
sensitive.
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Time-dependent recovery from inactivation and
frequency dependence of current. A two-pulse protocol
was used to study time-dependent recovery of
IK,ur at
37°C, as illustrated in Fig.
5A. A pair
of 50-s pulses (first pulse designated
P1 and second pulse designated P2) from 80 to +40 mV
were applied with varying coupling intervals, with the
P2 pulse preceded by a 100-ms
prepulse (ending 5 ms before P2)
to inactivate
Ito. As shown in
Fig. 5A, there was very little time-dependent current at short
P1-P2
intervals, with a progressive recovery in current as the
P1-P2
interval increased. Mean ± SE data from six cells are shown in Fig.
5B, along with the best-fit biexponential function. Recovery was complete within ~20 s and was
always a biexponential function of the
P1-P2
interval. The fast and slow recovery time constants averaged 419 ± 59 ms and 7.2 ± 0.9 s, and the amplitude of the fast and
slow components averaged 0.61 ± 0.05 and 0.43 ± 0.05, respectively.
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Fig. 5.
Time-dependent recovery and frequency dependence of
IK,ur.
A: currents recorded at 37°C
during 50-s test pulses (P2)
from one cell, after a 50-s conditioning pulse
(P1) to +40 mV with
P1-P2
intervals of 50 s (largest current) and 20, 6, 0.6, 0.2, and 0.1 s,
respectively (progressively decreasing currents). Test pulse was
preceded by a 100-ms prepulse to inactivate
Ito.
B: recovery of
IK,ur based on
amplitude of inactivating current during 50-s test pulses recorded as
illustrated in A. Results at each
P1-P2
interval are normalized to current at a
P1-P2
interval of 50 s. Results are means ± SE from 6 cells.
C: currents recorded at 37°C with
the pulse protocol shown, delivered 100 ms after a train of 100 pulses
at the frequencies (F) indicated.
Currents are shown following a 100-ms prepulse to inactivate
Ito, which was
followed 5 ms later by a 140-ms test pulse to record
IK,ur and a 60-ms
repolarization to 20 mV to record
IK,ur tail
currents. The largest current was recorded at 0.1 Hz, and currents
decreased progressively as frequency increased to 4 Hz.
D: mean ± SE tail currents and
4-AP-sensitive step currents elicited with the protocol shown in
C by test pulses after trains of 100- or 200-ms pulses at the frequencies indicated. Results are from 6 cells
at each data point.
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The data provided above indicate that, although
IK,ur inactivates
slowly, it is capable of inactivating fully, and recovery from
inactivation is relatively slow. These observations would suggest the
possibility of significant frequency dependence. We therefore studied
the frequency dependence of
IK,ur with the
use of a train of 100 pulses (from a holding potential of 80 to
+40 mV) of 100- or 200-ms duration, followed by a single 100-ms
prepulse to inactivate
Ito, and then 5 ms later a test pulse (140 ms) to +40 mV, followed by repolarization
for 60 ms to 20 mV to record IK,ur tail
current. The protocol was applied before and after 500 µM 4-AP, and
the frequency dependence of
IK,ur was
determined in the following two ways:
1) based on the tail current, which with the protocol used represents only
IK,ur, and
2) 4-AP-sensitive step current.
Similar results were obtained with both analyses. Figure
5C shows the 4-AP-sensitive current
from a typical experiment. Increases in frequency caused progressive
reductions in
IK,ur amplitude.
Mean results from six experiments with 200-ms pulses and six
experiments with 100-ms pulses are shown in Fig.
5D and indicate that
IK,ur
demonstrates highly significant frequency dependence
(P < 0.001 for each pulse duration)
over a clinically relevant range of frequencies (0.5-4 Hz).
The final set of experiments in native myocytes was performed to
evaluate possible changes in the reversal potential of
IK,ur during
depolarization to assess the possibility that apparent current
inactivation may be due to accumulation of
K+ in a restricted extracellular
space and a positive shift in the reversal potential. A double-pulse
protocol was used to measure the reversal potential of current at
37°C after a 100-ms (Fig. 6A) and
a 5-s (Fig. 6B) pulse to +40 mV. In
five cells, the reversal potential averaged 75.1 ± 4.1 mV
for the 100-ms pulse and 70.8 ± 6.2 mV for the 5-s pulse
(P = not significant). This small and statistically nonsignificant change in reversal potential can account
for only a 5% decrease in
IK,ur, in
contrast to the 84% decrease after 5 s at +40 mV seen at 37°C in
the experiments illustrated in Fig.
3A.
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Fig. 6.
Reversal of tail currents recorded from one cell at 37°C with the
use of a 100-ms (A) and a 5-s
(B) activating pulse. There was no
significant change in the reversal potential, indicating that the
substantial IK,ur
decrease that occurred during the 5-s pulse could not be attributed to
extracellular K+ accumulation.
Similar results were obtained in a total of 5 cells.
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Inactivation properties of Kv1.5 current expressed in
Xenopus oocytes. We applied a variety of approaches
(including the use of prepulse protocols and the pharmacological probe
4-AP) to isolate IK,ur; however,
it is impossible in native systems that express a variety of currents
to be sure that the currents studied are free from contamination by
other currents and are undistorted by the manipulations used to
suppress other currents. We therefore expressed Kv1.5 channels in
Xenopus oocytes and assessed the
voltage and time dependence of Kv1.5 inactivation at room temperature during 50-s depolarizing pulses as applied to study
IK,ur.
Figure 7A
shows the time course of Kv1.5 current inactivation during a 50-s pulse
from 80 to +40 mV. The data were best fit by a biexponential relation,
with time constants of 1.32 and 16.77 s in the example shown. Figure
7B shows mean time constants for Kv1.5
inactivation as measured at room temperature in 12 oocytes, which are
of the same order as the
IK,ur
inactivation time constants at room temperature shown in Fig.
3B. Figure
7C shows recordings from a
representative experiment to study the voltage dependence of Kv1.5
inactivation. A 300-ms test pulse to +40 mV was preceded by 50-s
conditioning pulses to voltages between 100 and +20 mV. In Fig.
7, only the currents for conditioning pulse voltages of 100,
80, and 60 to 0 mV are shown for the sake of clarity. Figure 7D shows mean data from five
oocytes. The data were well fitted by a Boltzmann relation, as shown.
The best-fit Boltzmann relation to data from each oocyte had a
half-inactivation voltage that averaged 26.1 ± 3.1 mV, of the same
order as values obtained for
IK,ur at room
temperature, as shown in Fig. 4F,
which was 23.1 ± 2.6 mV.
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Fig. 7.
Time and voltage dependence of inactivation of Kv1.5 channels expressed
in Xenopus oocytes.
A: time course of current inactivation
during a 50-s pulse to +40 mV. Reduced original data set (see Fig. 1
legend for explanation) and best-fit biexponential relation are shown.
B: time constants of Kv1.5
inactivation (room temperature) as determined with the types of
experiments illustrated in A. Results
are means ± SE of 12 experiments.
C: voltage-dependent inactivation of
Kv1.5 currents, determined at room temperature with the use of 50-s
conditioning pulses to the voltages indicated in
inset, followed by 300-ms test pulses
to +40 mV. Current evoked by the test pulse decreased progressively as
prepulse voltage became more positive, with no current detectable after
a prepulse to 0 mV. D: mean ± SE
data obtained in 5 experiments with the protocol illustrated in
C. Currents at each conditioning pulse
potential are normalized to current at a conditioning pulse potential
of 100 mV in each experiment. Best-fit Boltzmann relation to
mean data is shown.
|
|
| |
DISCUSSION |
In the present study, we have shown that
IK,ur undergoes
complete, albeit slow, voltage-dependent inactivation at positive potentials. The voltage and time dependencies of
IK,ur
inactivation are quite similar to those of Kv1.5 channels expressed in
Xenopus oocytes. Recovery of
IK,ur from
inactivation at diastolic potentials is also very slow, so that
IK,ur density is
quite sensitive to alterations in frequency over the physiological
heart rate range in humans.
Comparison with previous studies of
IK,ur and Kv1.5
inactivation.
In the first studies of
IK,ur, the
current was found to have slow and partial inactivation (20%
inactivation over 2 s at room temperature; see Ref. 21). Firek and
Giles (10) noted a contribution of a slowly inactivating component to
the pedestal current in human atrial cells, with a time constant of 1.7 s at 33°C. The pedestal component showed voltage-dependent
inactivation when the latter was studied with the use of prepulses,
with inactivation increasing as prepulse duration was increased from
400 to 2,500 ms, to a maximum of 50% (10). Amos et al. (1) studied
outward current inactivation upon depolarization of human atrial
myocytes at room temperature and noted a small slowly inactivating
component with a time constant of 1.4 s during 2-s pulses at room
temperature. This slowly inactivating component had many of the
features of IK,ur
(voltage dependence, 4-AP sensitivity, etc.) and had a
half-inactivation voltage of 9 ± 1 mV under their
experimental conditions.
Studies of Kv1.5 inactivation have also been somewhat limited.
Philipson et al. (17) obtained an inactivation time constant for Kv1.5
channels expressed in Xenopus oocytes
in the range of 2 s with the use of 1-s pulses at room temperature. The
half-inactivation voltage was 26 mV. Snyders et al. (19)
expressed Kv1.5 channels in mouse-derived L cells and studied their
biophysical properties at room temperature in greater detail.
Inactivation was found to be slow and partial, averaging 20 ± 2%
after 250 ms at +60 mV and 69 ± 3% during a 5-s pulse to +50 mV.
In three cells subjected to 30-s depolarizing pulses, inactivation
averaged 86 ± 1%. During 5-s pulses, inactivation developed as a
biexponential function with time constants of the order of 200-300
ms and 2-3 s and weak voltage dependence. The recovery from
inactivation after 5-s pulses was monoexponential with a time constant
of 1.65 ± 0.11 s, and the half-inactivation voltage was 25 ± 4 mV.
The present study is consistent with previous observations in the
literature but expands upon them substantially by studying in detail
the time course of development and removal of inactivation in native
myocytes with the use of pulses sufficiently long to achieve steady
state at 37°C. There are no systematic studies in the literature of
IK,ur or Kv1.5
current inactivation with the use of depolarizing pulses of >5 s in
duration. With slow-phase inactivation time constants in the range of
22 s at room temperature (based on our observations), a 5-s pulse is
clearly insufficient to characterize adequately the properties of
inactivation. Contrary to most previous observations using shorter
pulses, we found that IK,ur and Kv1.5
currents inactivate extensively; in fact, inactivation is complete if
depolarization to positive voltages is maintained for sufficient
periods of time. Because of relatively slow recovery from inactivation
at diastolic potentials,
IK,ur shows
significant frequency dependence at physiological temperatures.
Novel aspects and potential
importance. Atrial fibrillation (AF) is the most common
sustained arrhythmia in clinical practice, and the therapeutic options
presently available are suboptimal (16). The frequency-dependent
behavior of human atrial repolarization is an important determinant of
the occurrence of atrial reentrant arrhythmias and, in particular, AF
(2, 3) and may be a target for antiarrhythmic drug action (23). It is
therefore very important that a better understanding be obtained of the
basic ionic mechanisms underlying the response of the human atrial
action potential to changes in heart rate. Because of its slow and
partial inactivation, IK,ur has been
considered a current that is relatively frequency independent under
physiological conditions. The present report shows that this is far
from the case. Although
IK,ur
inactivation develops slowly, it is potentially quite extensive, and
recovery is slow at diastolic potentials. Consequently, the degree of
inactivation at the onset of an action potential will depend on the
relative times the cell spends at voltages positive to 50 mV (at
which inactivation begins at 37°C) versus voltages negative to
50 mV. During rapid arrhythmias, such as AF, atrial myocytes may
remain at voltages positive to 50 mV for most of the cycle, and
substantial IK,ur
inactivation might then be expected. Even with pulse durations as short
as 100 ms, we found that
IK,ur showed
substantial rate-dependent behavior at 37°C over the frequency
range between 0.1 and 4 Hz (Fig.
5D). Consideration of the kinetics
and magnitude of
IK,ur inactivation may thus be important in understanding the rate dependence of human atrial repolarization.
As activation rate increases, action potential duration tends to
decrease, largely because of
ICa inactivation
(4, 15, 24). This rate-dependent acceleration in repolarization
decreases the refractory period and therefore the minimum path length
over which reentry can occur (the wavelength) and thereby promotes the
occurrence of reentrant arrhythmias (18). The rate-dependent decrease
in IK,ur may tend
to offset the decrease in
ICa resulting from tachycardia and thereby result in a longer action potential during
tachycardia than if
IK,ur were rate
independent. Thus the rate-dependent properties of
IK,ur may serve a
protective role against reentrant arrhythmia. Teleologically, a similar
role may be suggested for
IK,ur
downregulation, which has been demonstrated in patients with AF (20).
There is considerable evidence that
IK,ur is present
in human atrium but not human ventricle (1, 8, 12, 14). This observation makes
IK,ur a
potentially attractive target for the development of ion
channel-selective antiarrhythmic drugs with reduced ventricular
proarrhythmic potential. The rate-dependent behavior of
IK,ur is
important to consider for the therapeutic application of such
compounds. If
IK,ur is
decreased substantially at rapid activation rates (e.g., during AF),
IK,ur blockers
may be much more effective in preventing AF, by virtue of the large amplitude of
IK,ur during
slower sinus rates, than in terminating the arrhythmia.
Several groups have identified a kinetically slower component of
Ito in human
atrial myocytes and have suggested that this component is related to
Kv1.5 channels (1, 10, 11). Our observations provide direct support for
this conjecture. The slowly inactivating component in human atrium has
been analyzed as a monoexponential function (1, 10, 11). Our results
suggest that
IK,ur
inactivation is, in fact, biexponential, as noted for Kv1.5 currents in
our study and previously published findings (19). The observation of
slow, monoexponential decay of outward current in human atrial cells is
probably due to the relatively short pulse durations used [<4 s
at room temperature (1), <2.6 s at 33°C (10)], which would
be insufficient to detect with any precision the slow phase of
IK,ur
inactivation with time constants ranging from 22 s (at room
temperature) to 7 s (at 37°C). The substantial temperature
sensitivity of both the relative magnitude and the rate of fast and
slow IK,ur
inactivation that we observed is important to consider when comparing
various studies in the literature and when analyzing the potential
physiological consequences of
IK,ur inactivation.
Potential limitations.
K+ accumulation in extracellular
clefts may reduce K+ currents
during prolonged depolarizations, giving the impression of inactivation
in the absence of true voltage-dependent inactivation. To address this
possibility, we measured the reversal potentials of tail currents after
short (100-ms) and long (5-s) depolarizing pulses (Fig. 6). If
K+ accumulation is contributing
importantly to current changes during the depolarizing pulse,
significant decreases in reversal potential should be seen during the
longer pulse. We found no statistically significant change at 37°C
in the reversal potential with a 5-s pulse compared with a 100-ms
pulse. On the basis of the actual reversal potentials measured,
K+ accumulation would account for
a 5% decrease in
IK,ur over the course of a 5-s pulse, rather than the 84% decrease observed experimentally.
The potential role of contaminating currents is always a concern in
studies of native cells expressing multiple channel types. Experimental
conditions were designed to minimize potential contamination by
INa,
ICa,
IK, and
IK1. The most
important potential overlapping current remaining was
Ito. We isolated
IK,ur with the
use of prepulses to inactivate
Ito and with the
use of 4-AP-sensitive current, as previously described (21). Even with
these methods, however, potential questions remain with respect to the
completeness of the suppression of overlapping currents and the degree
to which measures used to isolate
IK,ur may have
distorted it. We therefore studied the inactivation properties of Kv1.5
channels expressed in Xenopus oocytes
and compared them with properties noted for IK,ur. The close
similarity in the extent, kinetics, and voltage dependence of
inactivation between Kv1.5 and
IK,ur provides
strong support for the physiological validity of our observations
regarding IK,ur.
We have demonstrated that
IK,ur undergoes
extensive, albeit slow, inactivation and that slow recovery from
inactivation confers substantial rate-dependent properties on
IK,ur over the
physiologically relevant frequency range. The kinetics of
IK,ur
inactivation were determined in detail at both room temperature and
37°C, and a close similarity was noted between the inactivation
properties of
IK,ur and Kv1.5
current, supporting the notion that Kv1.5 channels carry a slowly
inactivating outward K+ current
during depolarization of human atrial myocytes. Our findings point to a
potentially important role of the rate-dependent properties of
IK,ur in
contributing to rate-dependent behaviors of the human atrium, which are
known to be associated with vulnerability to atrial reentrant
arrhythmias. Furthermore, our observations need to be considered in the
development of novel antiarrhythmic agents that target
IK,ur for the
treatment of AF.
| |
ACKNOWLEDGEMENTS |
The hKv1.5 clone that we used was a kind gift of Dr. Arthur M. Brown, Vice President Research, MetroHealth Center, Cleveland, OH. We
thank Nathalie Talbot for technical assistance and Caroll Boyer and
Luce Bégin for secretarial help with the manuscript. We express
deep gratitude to Drs. Michel Carrier, Raymond Cartier, Yves Leclerc,
Conrad Pelletier, Louis Perrault, Michel Pellerin, and Pierre
Pagé for providing human atrial tissue samples.
| |
FOOTNOTES |
This work was supported by grants from the Medical Research Council of
Canada, the Quebec Heart Foundation, and the Fonds de Recherche de
l'Institut de Cardiologie de Montreal. Z. Wang is a Canadian Heart
Foundation Research Scholar.
The costs of publication of this
article were defrayed in part by the
payment of page charges. The article
must therefore be hereby marked
"advertisement"
in accordance with 18 U.S.C. §1734 solely to indicate this fact.
Address for reprint requests: S. Nattel, Research Center, Montreal
Heart Institute, 5000 Bélanger St. East, Montreal, Quebec, Canada
H1T 1C8.
Received 9 April 1998; accepted in final form 20 July 1998.
| |
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Am J Physiol Heart Circ Physiol 275(5):H1717-H1725
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Abstract
Introduction
