Vol. 277, Issue 1, H119-H127, July 1999
Voltage-dependent K+ current
in capillary endothelial cells isolated from guinea pig
heart
Michael
Dittrich and
Jürgen
Daut
Institut für Normale und Pathologische Physiologie,
Universität Marburg, D-35037 Marburg, Germany
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ABSTRACT |
Capillary fragments were isolated from guinea
pig hearts, and their electrical properties were studied using the
perforated-patch and cell-attached mode of the patch-clamp technique. A
voltage-dependent K+ current was
discovered that was activated at potentials positive to 
20 mV
and showed a sigmoid rising phase. For depolarizing voltage steps from
128 to +52 mV, the time to peak was 71 ± 5 ms
(mean ± SE) and the amplitude of the current was 3.7 ± 0.5 pA/pF in the presence of 5 mM external
K+. The time course of
inactivation was exponential with a time constant of 7.2 ± 0.5 s at
+52 mV. The current was blocked by tetraethylammonium (inhibitory
constant ~3 mM) but was not affected by charybdotoxin (1 µM) or
apamin (1 µM). In the cell-attached mode, depolarization-activated
single-channel currents were found that inactivated completely within
30 s; the single-channel conductance was 12.3 ± 2.4 pS.
The depolarization-activated K+
current described here may play a role in membrane potential oscillations of the endothelium.
coronary circulation; membrane potential oscillations; electrophysiology of endothelium
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INTRODUCTION |
ENDOTHELIAL CELLS show complex changes in membrane
potential on the application of vasoactive agonists (20-22) and
are able to produce membrane potential oscillations (18,
19). Many of the functions of the endothelium, for
example, the release of vasoactive compounds and the regulation of the
hydraulic conductivity of the vascular wall, are influenced by the free
intracellular Ca2+ concentration,
which increases with hyperpolarization and often shows pronounced
oscillations (2, 11, 13, 15, 29). Thus the electrical activity of the
endothelial cells will almost certainly have profound effects on
endothelial function. As yet, the role of different ion channels in
generating the electrical responses of the endothelium is still far
from clear.
Most of our present knowledge of the electrophysiology of vascular
endothelial cells has been derived from studies on
1) cultured macrovascular
endothelial cells, 2) cell lines
derived from endothelium, and 3)
intact endothelium of large vessels (6, 23). There is, however,
relatively little information available on the electrical activity of
capillaries. ATP-sensitive K+
channels (12), Ca2+-activated
K+ channels (30), and
voltage-gated Ca2+ channels (1)
have been found in cultured microvascular endothelial cells. The significance of these findings is uncertain
because it is well known that endothelial cells may undergo profound
changes in metabolism and electrical properties during culture (28). Gögelein and co-workers managed to obtain single-channel
recordings from freshly isolated cerebral capillaries and found
nonselective cation channels (25) and inwardly rectifying
K+ channels (10).
Here we report the first whole cell recordings from fragments of
freshly isolated coronary capillaries. We give a quantitative description of the most prominent current in these cells, a
voltage-dependent K+ current that
activates rapidly on depolarization and inactivates very slowly.
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METHODS |
Isolation of capillaries.
Coronary capillaries were obtained from guinea pig hearts by enzymatic
dispersion. Guinea pigs weighing 250-450 g were decapitated and
their hearts rapidly excised. The isolated heart was perfused at a
constant flow rate of 10 ml/min using a peristaltic pump. The heart was
submerged in a small organ bath warmed to 37°C. The perfusing
solution contained (in mM) 130 NaCl, 15 KCl, 2 CaCl2, 0.8 MgCl2, 1 NaH2PO4,
2 Na-pyruvate, 10 glucose, and 10 HEPES. The pH was 7.4 (adjusted with
NaOH); the temperature was 37°C. The elevated
K+ concentration of the perfusate
caused cardiac arrest. Within 15-20 min, coronary perfusion
pressure increased to a steady level between 60 and 100 mmHg,
indicating recovery of energy metabolism. Subsequently, as a test of
adequate perfusion of the coronary blood vessels, 1 µM adenosine was
applied for 1-2 min, which elicited a reversible reduction of
perfusion pressure by ~50%.
To initiate dissociation of the cells, the heart was perfused for 5 min
with nominally Ca2+-free solution,
which otherwise had the same composition as described above. The heart
was then perfused for 10 min with
Ca2+-free solution to which 30 µM Ca2+ and 1 or 1.5 mg/ml
collagenase blend (type H, Sigma) was added. Subsequently, the heart
was removed from the organ bath and washed briefly in a solution
containing (in mM) 65 K-glutamate, 45 KCl, 30 KH2PO4,
3 MgSO4, 0.5 EGTA, 20 taurine, and
10 glucose (pH adjusted to 7.4 with KOH). The heart was then
disintegrated by being gently shaken with a forceps. Drops of the
suspension were transferred immediately to 35-mm petri dishes (Nunc,
Roskilde, Denmark) containing the same solution. After 30 min,
nonadhering cells were washed away with normal physiological salt
solution containing 5 mM K+ (for
composition, see Electrophysiology, solutions, and
reagents). Intact myocytes, spherical
cells of 10-15 µm in diameter, and capillary fragments remained
attached to the bottom of the petri dishes. In seven experiments,
capillary fragments prepared by the method of Langheinrich and Daut
(14) were used. At least 30 min before the start of
electrophysiological recording, 350-400 units of deoxyribonuclease
(DNase I, type IV, Sigma) were added to each petri dish to clean the
surface of the cells.
Electrophysiology, solutions, and reagents.
Patch-clamp recordings were carried out on the stage of an inverted
microscope (Zeiss, IM 35) at room temperature (23°C) within 10 h
after the suspension was seeded on the petri dishes. After a capillary
fragment adhering to the bottom of a petri dish was selected, a Perspex
frame was mounted over the capillary, as described previously (5). The
frame formed a perfusion chamber of 0.75 mm in height, 1.5 mm in width,
and 18 mm in length. The perfusion rate was 5-10 ml/h. The normal
physiological salt solution contained (in mM) 140 NaCl, 5 KCl, 1 MgCl2, 1 NaH2PO4,
2 CaCl2, 10 glucose, and 10 HEPES
(pH 7.4, adjusted with NaOH). In solutions containing higher
K+ concentrations or
tetraethylammonium (TEA; up to 20 mM),
Na+ was reduced to keep osmolarity
constant. Charybdotoxin (CTX) and apamin were purchased from Alomone
(Jerusalem, Israel). The solutions containing CTX or apamin were
prepared immediately before the experiments from stock solutions
(prepared with physiological salt solution) stored at 20°C
for no longer than 4 wk. All other reagents were obtained either from
Merck (Darmstadt, Germany) or from Sigma (St. Louis, MO).
Voltage-clamp experiments were carried out in both the conventional
whole cell mode of the patch-clamp technique
(n = 28) and the
perforated-patch mode (n = 52). For
perforated-patch measurements, pipettes of 1- to 2-µm tip diameter
were made of thin-walled glass 1.5 mm in diameter without filament
(Clark, Reading, UK). They were coated with Sylgard to reduce
capacitance and heat-polished directly before use. The resistance of
the pipettes was 5-8 M
. The pipette solution contained (in mM)
45 KCl, 100 K-aspartate, 1 MgCl2,
0.5 EGTA, and 10 HEPES (pH 7.2, adjusted with NaOH). With this
solution, Donnan potentials at the perforated patch were assumed to be
negligible. The tip of the patch electrode was first filled with
amphotericin-free pipette solution by aspiration, and then the pipette
was backfilled with the same solution to which 300 µg/ml amphotericin
B had been added from a stock solution. Sonication was applied to
improve solvation of amphotericin B. The stock solution contained 20 mg/ml amphotericin B in DMSO and was prepared freshly every day.
Perforation started shortly after seal formation and reached a
steady-state level within 5-10 min. The pipette solution used for
conventional whole cell recordings contained (in mM) 45 KCl, 100 K-aspartate, 10 EGTA, 1 CaCl2, 3 MgCl2, 2 Na2ATP, 0.1 Na3GTP, and 10 HEPES (pH 7.2, adjusted with NaOH). After a gigaseal was formed, the patch membrane
was ruptured by application of suction.
The seal resistance was determined by slow voltage ramps from
120 to +40 mV and back to 120 mV before the patch was
broken by suction or before amphotericin started to perforate the
patch. The membrane capacitance was calculated from the current offsets observed during the ascending and descending voltage ramps in the whole
cell mode.
Recordings were carried out with an EPC-7 patch-clamp amplifier (List,
Germany) and a modified digital audio tape recorder (DTC-55ES, Sony;
sampling rate 44 kHz). In all whole cell recordings, the capacitance
compensation circuitry of the EPC-7 amplifier was used. For analysis,
the data were filtered with an eight-pole Bessel filter before
sampling. The cutoff frequency was one-half the sampling rate, which
ranged from 100 to 5,000 Hz. Potentials were corrected for liquid
junction potentials (5 to 8 mV). The liquid junction
potentials were determined separately for the recording electrode and
for the reference electrode because changes in extracellular
K+ also affected the potential of
the reference electrode.
The results obtained with the conventional whole cell mode and with the
perforated-patch mode were very similar. Therefore, the results
obtained with both approaches were combined. The data are given as
means ± SE, and n denotes the
number of capillaries from which the data were obtained.
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RESULTS |
Electrical properties of coronary capillary fragments.
The capillary fragments could be easily identified by their morphology.
They appeared as thin translucent tubes of 
300 µm in length and
contained a regularly spaced string of nuclei. As measured between the
nuclei, the diameter of the vessel was 3-4 µm. Occasionally the
capillaries had side branches of similar diameter. The capillary
fragments most suitable for whole cell recording were nonbranched,
contained only three to five nuclei, and had an overall length of
30-80 µm (Fig. 1). The endothelial nature of the cells in the capillary fragments was ascertained in
control experiments by multicell RT-PCR (26) with the use of a
hydraulic "cell picker" and specific primers for endothelin-1.
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Fig. 1.
Capillary fragment and cardiomyocyte. Phase contrast photomicrograph of
a capillary fragment containing 4 nuclei. Adjacent cardiomyocyte is
shown for size comparison. Diameter of capillary in region between
nuclei was ~3 µm; lumen can just be discriminated as a dark line.
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The tip of the patch pipette was sealed in most cases to the
perinuclear region near the middle of the capillary fragment; the mean
seal resistance was 76 ± 6 G (n = 20). The capacitance of the capillary fragments was 42 ± 5 pF
(n = 31). The mean membrane potential
in physiological salt solution containing 5 mM
K+ was 34 ± 2 mV
(n = 32). In the
high-K+ solution (145 mM
K+), the membrane potential was
3.5 ± 0.8 mV (n = 25). The
mean steady-state input resistance of the capillary fragments (at
voltages near 0 mV) was 1.8 G (range 0.7-40 G). However, in
capillary fragments with a calculated input resistance >10 G, the
true input resistance could not be determined precisely, because it was
on the same order of magnitude as the seal resistance.
Single freshly isolated endothelial cells (in the same preparation) had
an average membrane capacitance of 9.2 ± 0.8 pF
(n = 30) (N. von Beckerath, M. Dittrich, and J. Daut, unpublished observations), i.e., ~22% of the
capacitance of our small capillary fragments consisting of three to
five cells. This is consistent with the assumption of electrical cell
coupling in the capillary fragments. In view of the high input
resistance of single endothelial cells (6, 32), we assume that a
spatially uniform voltage clamp of the entire capillary fragment could
be achieved.
Depolarization-activated
K+ current.
The most prominent current found in the capillary fragments was a
voltage-activated outward current. A typical recording is shown in Fig.
2A.
Depolarizing voltage steps were applied from a holding potential of
98 mV to test potentials between 38 and +52 mV. The
initial phase of the current is shown on a faster time scale in Fig.
2B. At +52 mV, the time to peak was 71 ± 5 ms (n = 23) with 5 mM external
K+ and 49 ± 4 ms
(n = 16) with 145 mM external
K+. The activation was followed by
a very slow inactivation. The time course of inactivation could be
fitted with a single exponential. At +52 mV, the mean time constant of
inactivation (determined by voltage steps >30 s) was 7.2 ± 0.5 s
in the presence of 5 mM external
K+
(n = 26). With 145 mM external
K+, the time constant of
inactivation (at +52 mV) was 6.4 ± 0.7 s
(n = 9), which is not significantly
different from the values obtained with 5 mM external
K+.
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Fig. 2.
K+ outward current activated by
depolarization. Perforated-patch recording was made with 5 mM external
K+. Same set of current records is
shown on a slow (A) and fast
(B) time scale.
A: voltage protocol
(top). Depolarizing voltage steps of
17-s duration were applied from a holding potential of 98 mV; holding
potential was maintained for 6.8 s before application of test pulse. In
this experiment, current declined to a steady level within 17 s
(bottom).
B: time course of initial phase of
activation. Dotted lines show that at different voltages (from +7 to
+52 mV) the initial current, after decay of capacitive transient, was
similar to steady-state current observed after inactivation of outward
current.
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As can be seen by comparing Fig. 2, A
and B, the amplitude of the initial
current (after decay of the capacitive transient) was similar to the
amplitude of the steady-state current, which suggests that the current
inactivated completely. The residual time-independent current (Fig. 2)
showed a variable degree of outward rectification, which was most
likely due to a Cl
conductance (23). No attempt was made to eliminate this current, because it did not interfere with the time-dependent current
investigated here (see DISCUSSION).
The amplitude of the inactivating outward current was determined by
subtracting the steady-state current observed after inactivation
(measured 30 s after the depolarizing voltage step) from the peak
current. At +52 mV, the amplitude of the depolarization-activated
current was 3.7 ± 0.5 pA/pF with 5 mM external
K+
(n = 31) and 3.3 ± 1.0 pA/pF with
145 mM external K+
(n = 9). If we assume a reversal
potential of 83 mV in 5 mM external K+ (0 mV in 145 mM
K+), we obtain a conductance of
27 pS/pF in 5 mM external K+ and
63 pS/pF in 145 mM external K+.
To investigate the ionic nature of the current, the external
K+ concentration was varied
between 5 and 145 mM. The outward current was activated by short
depolarizing voltage steps to +52 mV, and the reversal potential of the
deactivation tails was determined. In the experiment shown in Fig.
3A, the
tail currents reversed at 49 mV in the presence of 20 mM
external K+. Figure
3B shows that the dependence of the
reversal potential on external K+
was in close agreement with the prediction of the Nernst equation for a
K+-selective current.
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Fig. 3.
K+ selectivity of outward current
analyzed by tail currents. A: whole
cell recording of tail currents in presence of 20 mM external
K+ concentration
([K+]o;
average of 9 sweeps). Outward current was activated by a voltage step
from 88 to +52 mV (duration 200 ms). Deactivation was induced by
hyperpolarizing voltage steps to voltages around expected
K+ equilibrium potential of
50 mV; voltage protocol is shown at
right. Deactivation tails were
recorded at voltages around expected
K+ equilibrium potential of
50 mV. Amplitude of tail currents was obtained by subtracting
current immediately following capacitive transient (remaining after
capacitance compensation) from current recorded 140 ms after
hyperpolarizing voltage jump. Reversal potential determined by
interpolation was 49 mV. B:
dependence of reversal potential of deactivation tails on
[K+]o.
Straight line represents K+
equilibrium potentials according to Nernst equation. Data were taken
from 8 capillary fragments investigated with at least 2 different
[K+]o
values. Time course of deactivation became slower when potentials
became more positive in presence of elevated external
K+. In 7 recordings, we
additionally tested effects of 50 µM
Ba2+ on deactivation tails to
exclude a possible contamination by inward rectifier
K+ currents. Because no difference
in tail currents was observed, data obtained in presence and absence of
Ba2+ were combined.
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Voltage dependence of activation.
To determine the voltage dependence of activation, the potential was
clamped to 98 mV for 2.7 s and then the outward current was
activated by depolarizing voltage steps to various test potentials, as
shown in Fig. 4. The currents
activated by depolarizing voltage steps showed a sigmoid rising phase.
The time required to reach the peak current decreased with increasing
depolarization. The degree of sigmoidicity of the rising phase depends
on the number of sequential conformational changes required to allow
opening of the channel (35, 36). The best fits of the time course were
obtained with an exponential function raised to the fourth power;
exponents of 1, 2, and 3, respectively, gave inferior fits (see legend
to Fig. 4). As shown in Fig. 4B, the
time constants (
) of the fitted curves decreased from 40 ms at 18 mV
to 11 ms at +12 mV (n = 4).
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Fig. 4.
Time course of activation of outward current.
A: outward current was activated by
depolarizing voltage steps from 98 mV using conventional whole
cell mode with 5 mM external K+.
Holding potential of 98 mV was maintained for 2.7 s. Time course
of activation was sigmoid, and time to peak decreased with increasing
depolarization. Smooth lines superimposed on current traces are
least-square fits to I = A [1 exp
(t/)]4 + c, where
I is outward current, A is maximum
amplitude of activated current, t is
time, is time constant of activation, and
c is instantaneous current. Exponents
of 1, 2 and 3, respectively, gave inferior least-square fits; they
described time course less accurately, as could be easily judged by
eye. With an exponent of 4, was 25.4 ms, 13.5 ms, and 10.3 ms at
8, +2, and +12 mV, respectively. Duration of depolarizing pulses
was adjusted to terminate at peak of current. Deactivation tails were
recorded at 34 mV. B:
dependence of time constant of activation on voltage
(n = 4). Amplitude of depolarizing
voltage steps was varied as described in
A.
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The extent of activation of the K+
current at different test potentials was determined by analyzing the
deactivation tails, with the assumption that the tail current amplitude
was proportional to the conductance at the (preceding) test potential.
At the peak of the current, the potential was stepped back to 34
mV to induce deactivation. This potential, close to the
Cl equilibrium potential
(ECl), was
chosen to minimize any contamination by a time-dependent
Cl current (see
DISCUSSION).
In Fig. 5 the normalized conductance
derived from the tail currents at 34 mV is plotted against the
potential at which the current was activated. The voltage dependence of
the conductance change was sigmoid and could be described by a
Boltzmann function raised to the fourth power [which is
consistent with an activation mechanism involving four independent and
identical conformational changes (35, 36)]. The potential for
half-maximum activation (V1/2) was 7 mV
(n = 7). A simple Boltzmann function
gave a slightly inferior fit (not shown). We have also analyzed the voltage dependence of activation of the
K+ current in the presence of 145 mM external K+. Under these
conditions, V1/2
was 5 mV (n = 6) and the shape of the
activation curve was very similar to that shown in Fig. 5 for 5 mM
external K+.
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Fig. 5.
Voltage dependence of activation and inactivation. Activation and
inactivation curves were recorded in 5 mM external
K+. Data are means ± SE of 7 activation experiments ( ) and 21 inactivation experiments ( ). For
construction of activation curve, recordings were performed in
conventional whole cell mode using protocol described in Fig. 4.
Amplitude of tail current was taken as a measure of conductance at end
of activating voltage pulse
(gK). Amplitude of tail
current, normalized with respect to maximum current, is plotted against
potential at which current was activated. Averaged activation curve was
fitted by a Boltzmann function raised to the 4th power:
gK= {1 + exp [(V V')/k']}4,
where V' gives position on
voltage scale and k' is slope
factor indicating e-fold change in
conductance per mV. V' was 32 mV, and k' was 15 mV.
Potential for half-maximal activation
(V1/2; which
differs from V' due to 4th
power), was 7 mV. For construction of inactivation curve, recordings
were performed with perforated-patch technique or conventional whole
cell mode using protocol described in Fig. 6. Peak currents elicited by
voltage steps to +52 mV from different holding potentials were
normalized with respect to maximum current and plotted versus holding
potential. Mean inactivation curve was fitted by a Boltzmann function:
gK = {1 + exp [(V V1/2)/k]}1,
with a V1/2
(voltage for half-maximal inactivation) of 50 mV and a
k of 13 mV.
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Voltage dependence of inactivation.
The voltage dependence of steady-state inactivation was determined by
applying depolarizing voltage steps from holding potentials between
128 and +37 mV (Fig. 6). Holding
potentials were maintained for at least 6 s before the application of
depolarizing steps (of 
30 s duration) to +52 mV, which maximally
activated the outward current. The voltage dependence of steady-state
inactivation could be described by a simple Boltzmann function (Fig.
5). With 5 mM external K+, the
half-inactivation potential
(V1/2) was
50 mV and the slope factor
(k), indicating the
e-fold change in conductance per
millivolt (see legend to Fig. 5), was 13 mV
(n = 21). With 145 mM external
K+,
V1/2 was
53 mV and k was 13 mV
(n = 8). Thus the
inactivation curve, like the activation curve, was almost
unaffected by changes in external
K+.
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Fig. 6.
Steady-state inactivation. Whole cell recording was made with 5 mM
external K+. Current was activated
by depolarizing pulses to +52 mV from holding potentials between 128
and +37 mV. Holding potentials were maintained for 6 s.
Upper traces show voltage commands.
Amplitude of activated current, i.e., difference of peak current and
steady-state current after 32 s, was taken as a measure of steady-state
inactivation at holding potential. This capillary showed steady-state
outward rectification, possibly due to a
Cl conductance.
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Recovery from inactivation was studied in eight experiments using the
protocol shown in Fig.
7A. First,
the membrane potential was clamped to +52 mV for at least 28 s to
induce complete inactivation. The potential was then stepped back to
88 mV for various periods of time. With increasing duration of
the hyperpolarization, the peak outward current recorded at +52 mV
became successively larger. The time course of recovery from
inactivation could be described by a single exponential (Fig.
7B); the time constant was 551 ± 94 ms (n = 8).
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Fig. 7.
Recovery from inactivation. A:
complete inactivation of K+
outward current was achieved by depolarizing capillary to +52 mV for 28 s in 5 mM external K+. Removal of
inactivation was initiated by voltage pulses to 88 mV;
subsequently, voltage was again stepped to +52 mV (see
inset). Duration of hyperpolarizing
pulses was progressively prolonged. Peak current measured at +52 mV
increased with increasing duration of hyperpolarization.
B: time course of recovery from
inactivation (data from A).
Difference between peak current and steady-state current (after 40 s at
+52 mV) is plotted against duration of hyperpolarization. Envelope
curve could be fitted by a single exponential with a time constant of
468 ms.
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K+ channel
blockers.
As a first step toward a pharmacological characterization of the
outward current, we tested the K+
channel blockers TEA, CTX, and apamin. TEA reversibly inhibited the
voltage-activated current. In the presence of 10 mM TEA, the current
amplitude was reduced to 23 ± 6% of control during depolarizing voltage steps from 98 to +52 mV
(n = 4). Figure
8 shows a dose-response curve in which the
(interpolated) concentration required for half-maximal inhibition
(IC50) was 3.5 mM. In a second
complete dose-response curve IC50
was 2.5 mM. These results suggest that the current described here was
TEA-sensitive with an IC50 of ~3
mM.
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Fig. 8.
Effect of external TEA concentration
([TEA+]o)
on voltage-activated K+ current.
A: depolarizing voltage pulses to +52
mV were applied from a holding potential of 88 mV (voltage
protocol shown at top) were applied
in presence of 0, 0.08, 0.4, 2, 10, and 20 mM external TEA
(conventional whole cell recording). Increasing TEA concentrations
caused increasing block of voltage-activated current.
B: concentration dependence of TEA
block. Difference between peak current at +52 mV and steady-state
current after 40 s, normalized to current under control conditions (0 mM TEA), was plotted against TEA concentration on a semilogarithmic
scale. Solid line was drawn by eye. TEA concentration required for
half-maximal inhibition was estimated by interpolation.
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Some members of the Kv family of
K+ channels are sensitive to CTX
(3, 24). When we tested the effects of CTX on the voltage-activated K+ current, we found that 1 µM
CTX had no effect. Endothelial
Ca2+ activated
K+ channels are sensitive to
either apamin or CTX (23). Application of 1 µM apamin did not have
any effect on the outward current transient or the steady-state current
after complete inactivation. These findings suggest that
Ca2+-activated
K+ currents and
charybdotoxin-sensitive Kv channels (3) did not play a role under our
experimental conditions.
In 4 of 80 experiments, steady-state inward rectification was observed
at potentials negative to the resting potential. Thus it seemed
appropriate to exclude a possible contribution of inward rectifier
channels to the kinetics of the depolarization-activated K+ current (see legend to Fig. 3).
Single-channel recordings.
In 3 of 80 measurements in capillary fragments, voltage-dependent
single-channel currents could be resolved in the cell-attached mode of
the patch-clamp technique. An example obtained with 5 mM
K+ in the external solution is
shown in Fig.
9A. The
top trace shows the transmembrane potential of the patch (inside outside) calculated from the applied extracellular patch
potential and the membrane potential of the capillary fragment
(recorded later in the whole cell mode). When the patch membrane was
depolarized from 138 to +62 mV, single-channel outward currents were
observed. In the first 3 s, two channels opened simultaneously. The
channel activity subsided within 30 s after the depolarizing voltage
step, i.e., the open-state probability of the channels had a time
course similar to that of the whole cell currents. The average
single-channel conductance calculated from recordings at different
potentials in the three capillary fragments was 12.3 ± 2.4 pS
(symmetrical K+). The
extrapolated reversal potentials of the single-channel currents
measured in the cell-attached mode were equal to the membrane
potentials of the capillaries, which is consistent with a
K+-selective channel.
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Fig. 9.
Voltage-activated single-channel currents.
A: cell-attached recording from a
capillary fragment. Upper trace shows
transmembrane potential of patch
(Vm Vp) calculated
from applied pipette potential
(Vp) and
resting potential
(Vm) of
56 mV recorded later in perforated-patch mode. Openings of 2 channels were induced by depolarization from 138 to +62 mV.
Capacitive transient was subtracted. Channel inactivated completely
within 30 s. Small fluctuations of baseline were caused by beginning of
amphotericin perforation. B:
current-voltage relation of single-channel current was linear and had
an extrapolated reversal potential close to 0 mV
(Vm Vp), as
expected in symmetrical K+. Mean
current amplitudes were calculated from single-channel currents of
5-8 openings at each voltage. Conductance determined between +20
and +80 mV was 9 pS.
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Current-clamp experiments.
To get further information on the function of the current, we carried
out current-clamp experiments. Depolarizing current pulses were
injected into capillary fragments in the conventional whole cell
current-clamp mode. Figure 10 shows a
typical experiment with a capillary that had a resting potential of
~20 mV. At this potential the voltage-dependent
K+ current was completely
inactivated. The current pulses caused voltage displacements that
showed no obvious time dependence (the passive membrane time constant
was too fast to be resolved on the time scale of the record). In
contrast, when the membrane potential was changed to about 40 mV
by the application of hyperpolarizing current, a qualitatively
different pattern of voltage changes was observed. Figure
10B shows that, in the presence of a
holding current of 23 pA, superimposition of the same
depolarizing current pulses gave rise to a slow, time-dependent
depolarization if the threshold of activation of the
K+ current (20 mV) was
crossed. When the applied current was switched off, the membrane
hyperpolarized with no obvious delay.
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Fig. 10.
Effect of depolarizing current pulses on membrane potential. Typical
experiment shows effect of depolarizing current pulses (duration 15.4 s) in a capillary fragment (5 mM external
K+; whole cell mode).
Upper traces indicate applied
currents; intervals between current pulses were 46 s.
A: current injection at resting
membrane potential of about 20 mV. At this potential,
voltage-activated K+ current was
almost completely inactivated. Depolarization appears instantaneous
because membrane time constant was fast compared with time scale of
recording. B: membrane potential was
adjusted to 40 to 45 mV by application of a holding
current of 23 pA. In this potential range, voltage-activated
K+ current is only partially
inactivated (see Fig. 5). Depolarizing current steps induced a very
slow depolarization that could be fitted with a single exponential
(continuous line; time constant 6.7 s). Time course of slow
depolarization was probably related to inactivation of
voltage-activated K+ current. When
current was switched back to 23 pA, resulting hyperpolarization to
about 43 mV showed no delay.
|
|
The most likely interpretation of the slow voltage changes shown in
Fig. 10B is that the activation of the
voltage-dependent K+ current
initially impeded the depolarization and that, subsequently, the cells
depolarized slowly as the K+
current inactivated. In four current-clamp experiments similar to that
shown in Fig. 10, the time constant of the change in membrane potential
between 0 and +20 mV was 8.1 ± 2.3 s. This is similar to the mean
time constant of inactivation of the voltage-clamp current at +6 mV,
which was 6.6 ± 1.2 s (n = 8). It
should be noted that the time course of the membrane potential change
on current injection and the time course of inactivation during a voltage step to a fixed potential are not necessarily identical. Nevertheless, the good agreement of the time constants measured in
voltage-clamp and current-clamp experiments (in the same potential range) is consistent with the hypothesis that both effects may be
attributable to the activation and subsequent inactivation of a
voltage-activated K+ current.
Thus, when the membrane potential of capillaries is more negative than
40 mV, depolarizing currents may be antagonized by activation of
an outward current as soon as the threshold of 20 mV is crossed.
| |
DISCUSSION |
Patch-clamp recording from freshly isolated capillary fragments.
Because microvascular endothelial cells in situ are not directly
accessible with patch electrodes, the electrophysiology of microvascular endothelium has so far been investigated mostly in
cultured cells. However, gene expression of channels, receptors, and
metabolic pathways may change rapidly under cell culture conditions (9,
28). On the other hand, in freshly isolated endothelial cells, cell
identification can pose a problem, because single endothelial cells
round up and may not be readily distinguished from rounded up
fibroblasts or smooth muscle cells. In the present study we have used
freshly isolated capillary fragments that can be easily identified by
their morphology. In many vascular beds, endothelial cells were found
to be electrically coupled by gap junctions (7, 8, 17), although in
capillaries it has been difficult to identify gap junctions by electron
microscopy (34). Our finding that the membrane capacitance of single
endothelial cells was about five times smaller than the average
capacitance of the capillary fragments is consistent with the idea that
the endothelial cells in the capillary fragments were electrically coupled and that a spatially homogeneous voltage clamp could be achieved. The electrical length constant of capillaries is probably ~1 mm (6), whereas the length of the capillary fragments was maximally 300 µm.
The steady-state current-voltage relations between 60 and +52 mV
showed a variable degree of outward rectification, similar to the
results summarized by Nilius et al. (23). This finding and the mean
resting potential of 34 mV, which is close to
ECl, suggest that
Cl channels were also
present. Recently it has been shown that the volume-activated
Cl current may also show
some inactivation at potentials greater than +40 mV with time constants
of maximally 200 ms (31). However, our current exhibits much longer
time constants of inactivation, and most of our experiments were
carried out at potentials of up to +52 mV, where the extent of
inactivation of the volume-activated Cl current would be
negligibly small. Thus we consider it unlikely that
Cl currents interfered with
our analysis of the voltage-activated K+ current.
Voltage-dependent
K+ current in
coronary capillaries.
We have found a voltage-activated
K+ current in coronary capillaries
that has not been previously described in endothelial cells. This
current activates and inactivates much more slowly than the A-type
current found by Takeda et al. (27) in cultured aortic endothelial
cells (the only other endothelial voltage-dependent K+ current reported so far).
Because most previous patch-clamp studies were carried out on cultured
macrovascular endothelial cells, it appears possible that the current
reported here is restricted to capillary endothelium. Alternatively,
its expression may be lost during cell culture.
The capillary endothelial K+
current was activated at potentials positive to 20 mV (Fig. 5) and
was characterized by a sigmoid onset. The time to peak was on the order
of 50-80 ms. The rising phase of the current could be described by
an exponential function raised to the fourth power, and steady-state
activation could be fitted by a Boltzmann function raised to the fourth
power. This gives a minimum for the number of sequential conformational changes required to open the channel (4, 35, 36). The time course of
inactivation could be fitted by a single exponential with time
constants between 5 and 10 s. The time constant of recovery from
inactivation was ~0.5 s. The current showed a relatively high
sensitivity to TEA but was not affected by CTX. The kinetics of the
current showed only minor changes when external
K+ was increased from 5 to 145 mM,
whereas the conductance increased more than twofold. The single-channel
conductance was ~12 pS. The properties of the endothelial
K+ current described here differ
substantially from eag or HERG currents and from the slowly activating delayed rectifier in cardiac muscle, but show some similarities with
Shaker-type voltage-activated K+ currents (3, 24).
Possible function of voltage-activated
K+ current.
The activation and inactivation curves (Fig. 5) showed some overlap in
the voltage range from 0 to 10 mV. Multiplication of
relative activation and relative inactivation suggests that ~2% of
the maximal current may flow in the window around 20 mV. Theoretically, this might contribute to the setting of the resting potential in depolarized cells. However, we have not seen a
corresponding "hump" in the steady-state current-voltage
relation. More importantly, however, the voltage-activated
K+ current may have a pronounced
influence on the dynamic behavior of the membrane during membrane
potential oscillations. Depolarization of the cell membrane from a
resting potential of 40 to 45 mV, where 30-40% of
the current is available for activation, to 0 mV may activate ~20%
of the maximum current. The rapid activation of the current would
counteract any further depolarization. Thus the effect of depolarizing
current pulses may be delayed for several seconds until the outward
current is inactivated (see Fig. 7). This may play a role during
activation of the cells with vasoactive agonists, which often elicit
pronounced membrane potential oscillations (11, 13, 15, 33). The
mechanisms underlying the membrane potential oscillations in
endothelial cells have been widely studied (11, 16, 30, 34). The
available evidence suggests that oscillatory changes in intracellular
Ca2+ are accompanied by
synchronized changes in membrane potential, which are partially, but
not entirely, attributable to the opening of
Ca2+-activated
K+ channels. On the other hand,
changes in membrane potential may also influence transmembrane
Ca2+ movements (2). An outward
current that activates rapidly and inactivates slowly would be expected
to modulate the shape and frequency of membrane potential oscillations
and thus, via the effect of membrane potential on
Ca2+ influx, influence
oscillations in intracellular
Ca2+.
| |
ACKNOWLEDGEMENTS |
We thank B. Burk, R. Luzius, R. Graf, A. Mazzola, and E. Hoffmann
for technical and secretarial help and Dr. C. Walther for useful
comments on the manuscript.
| |
FOOTNOTES |
This work was supported by the Deutsche Forschungsgemeinschaft (Da
177/4-4 and Da 177/7-1).
Present address of M. Dittrich: Institut für Neurophysiologie,
Universität zu Köln, Robert-Koch-Strasse 39, D-50931
Köln, Germany (E-mail: michael.dittrich{at}uni-koeln.de).
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 and other correspondence: J. Daut,
Institut für Normale und Pathologische Physiologie der
Universität Marburg, Deutschhausstrasse 2, D-35037 Marburg,
Germany (E-mail: daut{at}mailer.uni-marburg.de).
Received 25 September 1998; accepted in final form 10 March 1999.
| |
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ABSTRACT
INTRODUCTION
