Spontaneous transient outward currents and delayed rectifier
K+ current: effects of
hypoxia
C.
Vandier1,
M.
Delpech2, and
P.
Bonnet2
1 Unité Mixte de Recherche (UMR)
Centre National de la Recherche Scientifique 6542, Physiologie des
Cellules Cardiaques et Vasculaires, Faculté des Sciences, 37200 Tours Cedex; and 2 UMR Centre
National de la Recherche Scientifique 6542, Physiologie des
Cellules Cardiaques et Vasculaires, Faculté de Médecine,
37032 Tours Cedex, France
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ABSTRACT |
Single smooth muscle cells of rabbit
intrapulmonary artery were voltage clamped using the perforated-patch
configuration of the patch-clamp technique. We observed spontaneous
transient outward currents (STOCs) and a steady-state outward current.
Because STOCs were tetraethylammonium sensitive and activated by
Ca2+ influx, they were believed to
represent activation of
Ca2+-activated
K+ channels. The steady-state
outward current, which was sensitive to 4-aminopyridine, was the
delayed rectifier K+ current. In
cells voltage clamped at 0 mV, we found that STOCs were not randomly
distributed in amplitude but were composed of multiples of 1.57 ± 0.56 pA/pF. The mean frequency of STOCs was 5.51 ± 3.49 Hz.
Ryanodine (10 µM), caffeine (5 mM), thapsigargin (200 nM), and
hypoxia (PO2 = 10 mmHg) decreased
STOCs. The effect of hypoxia on STOCs was partially reversible only if the experiment was conducted in the presence of thapsigargin. Hypoxia
and thapsigargin decrease steady-state outward current. Thapsigargin
and removal of external Ca2+
abolished the effect of hypoxia, suggesting that hypoxia decreases steady-state outward current by a
Ca2+-dependent mechanism.
pulmonary artery smooth muscle cells; calcium ion regulation; sarcoplasmic reticulum; potassium ion
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INTRODUCTION |
THE MECHANISM BY WHICH hypoxia causes pulmonary
vasoconstriction has not been elucidated. Studies of the hypoxic
response reported that hypoxia directly acts on smooth muscle cells
(18, 25) and on membrane K+
channels. Indeed, hypoxia inhibited delayed rectifier
K+ channels (2, 21) and activated
Ca2+-activated
K+ channels (2, 5). Also, hypoxia
inhibited and activated L-type
Ca2+ currents, respectively, in
proximal and distal pulmonary artery smooth muscle cells (8).
The Ca2+ present in the
sarcoplasmic reticulum (SR) seems to play an important role in the
hypoxic response of pulmonary smooth muscle cells (21, 23, 25). Several
investigators suggested that hypoxic mobilization of
Ca2+ from internal stores was the
initial event induced by hypoxia (21, 23, 25). Salvaterra and Goldman
(23) showed that hypoxia induced a later hypoxic response phase that
consists of an activation of Ca2+
influx, in part, through channels other than L-type
Ca2+ channels. This late hypoxic
phase was completely blocked by thapsigargin (23). Then external
Ca2+ and both the release of
Ca2+ from SR and the depletion of
this Ca2+ appear to be important
in the hypoxic response.
Recently, we showed in rabbit pulmonary artery rings that hypoxia,
which has no effect on resting tone of rabbit intrapulmonary artery
rings, increased the amplitude of the norepinephrine
phasic-induced contraction in the absence of external
Ca2+ (27). This effect was blocked
by ryanodine. This suggested an important role of the
Ca2+ present in the SR for the
hypoxic response of rabbit pulmonary artery cells.
We have examined the effect of hypoxia on spontaneous transient outward
currents (STOCs), which reflect
Ca2+ release of superficial SR
(4), and on outward current activated by a ramp protocol having a low
speed of depolarization. We show here that hypoxia
decreases the delayed rectifier K+
current via a Ca2+-dependent
mechanism and decreases STOCs in intrapulmonary artery smooth muscle
cells. Both effects appear to result from a depletion in
Ca2+ concentration of superficial
SR.
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METHODS |
Cell isolation. All animal experiments
were conducted according to the ethical standards of the
Ministère Français de l'Agriculture. Rabbits of either sex
(2-3 kg) were killed by cervical dislocation. Before the left and
right proximal intrapulmonary arteries (external diameter
2-3
mm) were dissected, the lung was removed and the pulmonary arteries
were perfused with a cold physiological salt solution (PSS) that
contained 10 µM sodium nitroprusside. Vessels were opened along their
longitudinal axis and incubated in
Ca2+-free solution for 10 min.
They were then cut into small pieces and placed in 5 ml of a
Ca2+-free solution containing 191 U/ml collagenase (CLS 2; Worthington Biochemical), 0.22 U/ml pronase E
(Sigma), and 3 mM dithiothreitol at 37°C for 20-23 min on a
tridimensional agitator. The tissue was washed and placed in
Ca2+-free solution without enzymes
and gently agitated for 40 min. The tissue was then strongly agitated
with a polished wide-bore Pasteur pipette to release the cells. Cells
were stored at 4°C and used between 2 and 10 h after isolation. PSS
solution contained (in mM) 138.6 NaCl, 5.4 KCl, 1.8 CaCl2, 1.2 MgCl2, 0.33 NaH2PO4, 10 HEPES, and 11 glucose; pH was adjusted to 7.4 with NaOH. The Ca2+-free solution was a PSS
solution without added Ca2+.
Electrophysiology. For
electrophysiological recording, the cells were placed in a 1-ml volume
bath and continuously superfused by gravity at the rate of 4 ml/min
from reservoirs. The reservoirs could be switched manually to allow
addition and removal of different solutions to the bath as required.
Total solution exchange in the bath was reached ~1.5 min after
switching from control solution. Cell membrane currents were recorded
with a List EPC-7 patch-clamp amplifier (List Electronics, Darmstadt,
Germany). Patch pipettes were pulled from borosilicate glass
capillaries and had resistance of 4-5 M
. The head stage ground
was connected to an Ag-AgCl pellet that was placed in a side bath
filled with the pipette solution, connected to the main bath via an
agar bridge containing 3 M KCl. The junction potentials
between the electrode and the bath were canceled by using the voltage
pipette offset control of the amplifier. The capacitances of the
pipette and the cells were canceled. The series resistance was also
canceled at 50-85%.
We used the perforated-patch technique to record whole cell membrane
currents (22), with amphotericin B included in the patch pipette at 240 µg/ml. Briefly, pipette tips were filled by dipping the tip of the
pipette into the pipette solution, and then the pipette was backfilled
with pipette solution containing amphotericin B. After the gigaseal
between the pipette and the cell was realized, the electrical access to
the cytoplasm was monitored by applying
10-mV pulses for 10 ms
from a holding potential of
60 mV and monitoring the capacitive
transient. This current was filtered at 5 kHz and sampled at 50 kHz.
Typically, access was gained within 10 min and was stable within 30 min. All the experiments started after these 30 min. The pipette
solution contained (in mM) 122 glutamic acid, 25 KCl, 1 MgCl2, 10 HEPES, and 1 EGTA; pH
was adjusted to 7.2 with KOH. In 94 cells, final access resistance was
estimated to be 28 ± 8 M
(range 10-50 M
). Only cells
with series resistance <30 M
were kept.
The voltage-clamp protocol used to evaluate the cell current-voltage
(I-V)
characteristics was a voltage ramp of 0.03 mV/ms from
90 to 0 mV. The holding potential was
60 mV. Data were sampled at 670 Hz
and filtered at 150 Hz.
I-V
relationships were also generated in voltage-clamped cells held at a
membrane potential of
60 mV and then were stepped in 10-mV
increments to command potentials between
90 and 0 mV. The
voltage steps were 400 ms in duration, with 5-s intervals between
steps. The data were sampled at 5 kHz and filtered at 1 kHz. The
voltage-ramp protocol was checked by comparing the
I-V
relationships obtained from the voltage ramp and the current measured
at the end of the 400-ms voltage steps
(n = 11). The results were identical.
To characterize STOCs, the membrane potentials of the cells were held
at a maintained voltage of 0 mV. The STOCs were recorded using a DAT
recorder (DTR-1204 recorder; Biologic) for later analysis. Then STOCs
were sampled at 1 kHz and filtered at 200 Hz for analysis.
Voltage-clamp protocols were generated, and the data were captured with
a PC using a labmaster TL1-125 interface (Scientific Solutions)
and pClamp 5.5.1 software (Axon Instruments). The analysis was realized using pClamp and Origin software (Microcal Software, Northampton, MA).
O2 tension.
The external solution was equilibrated with either air
(PO2 ~150 mmHg) or
N2
(PO2 <10 mmHg) in a reservoir. PO2 in the output of the recording
chamber was monitored with an
O2-sensing electrode (oxymeter,
781 Strahkelvin Instruments). The interval of time between the
switching of bath perfusion to a measured drop in
PO2 was ~3 s, and saturation of the bath at PO2 of ~10 mmHg was
achieved within ~50 s.
Chemicals. Stock solutions of sodium
nitroprusside (10 mM) and ryanodine (20 mM) were prepared in distilled
water and then diluted in PSS at an appropriate concentration.
4-Aminopyridine (4-AP), caffeine, tetraethylammonium chloride (TEA),
and cadmium (Cd2+) were prepared
daily in PSS. Anthracene-9-carboxylic acid (9-AC; 1 mM) was dissolved
in DMSO, which was present at 0.05% after dilution. Stock solution of
thapsigargin (2 mM) was prepared in DMSO, which was present at 0.01%
after dilution. When we used 9-AC and thapsigargin, the same percentage
of DMSO was added to all of the experimental solutions. All chemicals
were from Sigma (St. Quentin Fallavier, France).
Statistics. Data are expressed as
means ± SD. We used Wilcoxon's test and randomization test for
paired comparisons. Differences were considered significant at
P < 0.05. Statistical analysis was
realized using Stat, a software developed in this laboratory by Dr. J. Y. Le Guennec.
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RESULTS |
Membrane currents recorded in perforated
patch. Membrane currents were recorded in single smooth
muscle cells from the intrapulmonary artery in PSS solution. Ten step
depolarizations between
90 and 0 mV from a holding potential of
60 mV elicited both an outward steady-state current and STOCs
(Fig.
1A).
The steady-state outward current was not always evident when STOCs were
superimposed upon it. TEA (1 mM) in the bath solution abolished STOCs,
and the steady-state outward current was then clearly visible. It was
an outward current that activated near
30 to
40 mV, and
its amplitude increased with depolarization. The addition of 1 mM 4-AP
to the bath solution containing 1 mM TEA almost totally abolished the
steady-state outward current (n = 11).
These two types of outward currents were observed in the same cell in
response to a voltage ramp from
90 to 0 mV (Fig.
1B). With the voltage-ramp protocol,
we clearly see that the STOCs and steady-state outward current
activated at about the same voltage of
30 to
40 mV.

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Fig. 1.
Effects of tetraethylammonium (TEA) and 4-aminopyridine (4-AP) on
steady-state outward current and on spontaneous transient outward
currents (STOCs) elicited by either voltage steps
(A) or voltage ramps
(B). In control, depolarization of
an intrapulmonary artery cell from 90 to 40 mV induced a
steady-state inward current of small amplitude. When the cell was
depolarized further than 40 mV, the steady-state current was
outward, and amplitude varied irregularly with spontaneous peaks in
which the maximal value could reach ~250 pA and 100 ms duration
(STOCs). The same phenomenon was observed when the same cell was
depolarized from 90 mV up to 0 mV by the voltage ramp protocol.
In both protocols, 1 mM TEA in the bath solution abolished STOCs. In
presence of 1 mM TEA and 1 mM 4-AP, the steady-state outward current
was also almost totally suppressed. Arrows in
A and
B and dashed line in
B indicate the 0 current
level.
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The effect of 9-AC, a chloride channel blocker, was tested on four
cells with the same voltage-clamp protocols. Step depolarizations between
90 and 0 mV from a holding potential of
60 mV
elicited the two outward currents. 9-AC (1 mM) had no effect on the
steady-state outward current or on STOCs (Fig.
2A).
Changing the bath solution to one that contained 1 mM 9-AC and 1 mM
4-AP almost totally abolished the steady-state outward current but did
not affect the STOCs. Similar results were obtained from the same cell
in response to the voltage-ramp protocol (Fig.
2B). The steady-state outward current was an outward current activated at
30 to
40 mV,
blocked by 1 mM 4-AP, and not affected by TEA or 9-AC. Characteristics that correspond represented the delayed rectifier
K+ current that has already been
observed in these cells (9, 20, 21, 26).

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Fig. 2.
Effects of anthracene-9-carboxylic acid (9-AC) and 4-AP on steady-state
outward current and STOCs elicited by voltage steps
(A) or voltage ramps
(B) in the same cell. Cell was first
depolarized by steps of 10 mV, which induced steady-state outward
current and STOCs (A: control). 9-AC
(1 mM) had no effect on STOCs. Addition of 1 mM 9-AC and 1 mM 4-AP to
the bath solution had little effect on STOCs, but the steady-state
outward current was totally suppressed. Similar observations were made
when the cell was depolarized by voltage ramps
(B: 90 to 0 mV membrane
potential). Arrows in A and
B and dashed line in
B indicate the 0 current level.
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STOCs, which were observed in >75% of the cells recorded with the
perforated-patch technique, were activated around
30 to
40 mV, and their amplitude increased with depolarization (Figs. 1 and 2); they were abolished by 1 mM TEA and insensitive to 1 mM 4-AP
and 1 mM 9-AC, suggesting that STOCs represented a
Ca2+-activated
K+ current rather than a chloride
current.
STOCs were evaluated in 19 cells, with an example represented in Fig.
3. The membrane potential was held at 0 mV,
and the amplitude and frequency of STOCs were measured over 60-s
periods. STOC amplitude was normalized according to cell capacitance.
The smallest STOC that we could distinguish had an amplitude of 20 pA
(corresponding to 0.76 pA/pF); below this value, we could not accurately differentiate STOCs from the membrane noise. These 19 cells
show that, although STOCs were not of uniform size, their amplitudes
were not randomly distributed. All-point amplitude histograms of
membrane current revealed peaks that represented STOC size to be
composed of multiples of 1.57 ± 0.56 pA/pF
(n = 19). The frequency of STOCs was
5.51 ± 3.49 Hz (n = 19).

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Fig. 3.
Example of the nonuniformity of STOC amplitude in a cell voltage
clamped at 0 mV. A: continuous
recording of membrane current. Average frequency of STOCs was 6.41 Hz
in this example. Bottom trace shows
the selected region on an expanded time scale.
B: all-point amplitude histogram of
the record shown in A. Distribution of
the STOC amplitudes as a function of the number of events recorded
distinguished four classes of STOCs shown by four peaks of amplitudes
indicated by asterisks (*). We demonstrated that the amplitude interval
between each peak of each class was constant and of 1.57 ± 0.56 pA/pF (n = 19). Arrow in
A indicates the 0 current level.
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In 30% of cells when the membrane potential was maintained at 0 mV for
>10 min, we observed a decrease of STOC activity with time. In 70%
of cells, STOC activity was stable for 25 min. To evaluate this
rundown, five cells were held at 0 mV for 25 min, and we measured the
frequency of STOCs during 60 s, after 4 min, after 14 min, and after 24 min. At these three different times (4, 14, and 24 min), the
frequencies of STOCs were 1.29 ± 0.58, 0.96 ± 0.53, and 0.55 ± 0.51 Hz, respectively. At these three different
times, the frequencies were not significatively different.
Role of extracellular
Ca2+ in STOC
activity.
To investigate the role of external
Ca2+ in STOCs, we first changed
the bathing solution (PSS) to a
Ca2+-free solution containing 1 mM
EGTA in three cells. In Fig.
4A, the
cell was voltage clamped with the voltage-ramp protocol. This protocol
elicited STOCs and the steady-state outward current. Removal of
extracellular Ca2+ for 10 min
totally abolished STOCs at all ramp voltages. This suggested that
Ca2+ influx was indispensable for
STOC activity. The removal of extracellular Ca2+ was also associated with an
increase in steady-state outward current (Fig.
4A).

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Fig. 4.
Importance of extracellular Ca2+
for STOC activity. A: typical trace of
current elicited by a voltage-ramp protocol in a cell recorded first in
control conditions (control) and then after exposure to 0 Ca2+ solution (0 Ca). Removal of
the external Ca2+ abolished STOCs
and increased the steady-state outward current.
B: in a different experiment, the
cell, whose membrane potential was held at 0 mV, was superfused by
saline solution containing 1 mM
Cd2+.
Cd2+ (1 mM) reversibly decreased
the frequency of STOCs. It also reversibly depressed the steady-state
outward current. Arrows in A and
B and the dashed line in
A indicate the 0 current level.
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In three cells, we tested some properties of the L-type
Ca2+ current to see if this was
one pathway for Ca2+ to enter the
cell and activate STOCs. The L-type
Ca2+ current of rabbit pulmonary
artery is abolished by external
Cd2+ (7). Figure
4B shows a cell voltage clamped at 0 mV for 6 min. During the first minute, the cell was bathed with PSS
solution, and it showed STOCs. When the bath solution was changed to a
solution that contained 1 mM Cd2+,
the frequency of STOCs rapidly decreased and was stable after 2 min in
this solution. The effect of Cd2+
was reversible, and the activity of STOCs recovered in PSS solution after a 2-min wash. Thus, under these conditions of 0-mV voltage clamp
with 1 mM external Cd2+, the STOCs
were not totally abolished, and the mean frequency of STOCs decreased
from 0.51 ± 0.16 Hz in PSS solution to 0.16 ± 0.05 Hz in
solution with Cd2+
(n = 3). In these cells,
Cd2+ also provoked a reversible
decrease of the steady-state outward current.
Role of intracellular
Ca2+ stores in
STOC activity.
To investigate the role of intracellular
Ca2+ stores, we tested the effects
of caffeine, ryanodine, and thapsigargin on STOCs.
To test the effect of caffeine on STOCs, eight cells were voltage
clamped at 0 mV in the presence and in the absence of external caffeine. Figure
5A shows a
cell voltage clamped at 0 mV, which showed STOCs in the PSS solution.
The activity of STOCs was stable during this 2 min in PSS solution.
External application of 5 mM caffeine rapidly induced a large transient
increase in STOCs that summated to give a peak outward current of 19 ± 16 pA/pF (n = 8) ~30 s after
the beginning of the application of caffeine. After this time, the
amplitude of the outward current decreased and was stable ~1 min
after the beginning of the application of caffeine. After this period,
the STOCs were totally abolished. The effect of caffeine was slowly
reversible, and STOCs began to reappear after 2 min in PSS solution and
fully recovered after 4 min.

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Fig. 5.
Role of intracellular Ca2+ stores
on STOCs. A: in a cell held at 0 mV, 5 mM caffeine rapidly elicited an increase in STOCs that summated to form
a transient peak of current. Then STOCs were progressively abolished.
After the washout of caffeine, STOCs reappeared.
B: in a different cell, also held at 0 mV, 10 µM ryanodine slowly decreased STOCs. Arrows in
A and
B indicate the 0 current level.
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To confirm the involvement of ryanodine-sensitive
Ca2+ pools in the activation of
STOCs, 10 µM ryanodine was applied in three cells voltage clamped at
0 mV. Figure 5B shows that the
application of 10 µM external ryanodine decreased STOCs gradually but
not completely in 12 min. The effect of ryanodine was not reversible on
the time scale of these experiments. The slow reduction in STOCs is
consistent with the gradual depletion of the
Ca2+ pool by this agent (12, 17).
To test whether STOCs were linked to the accumulation of
Ca2+ into the SR by
Ca2+-ATPase, we used thapsigargin,
which is known to inhibit this Ca2+-ATPase in pulmonary artery
(10). Seven cells were held at 0 mV to test the effect of thapsigargin.
Figure 6 shows a cell with STOCs and a
steady-state outward current in PSS solution. When the bath solution
was changed to one that contained 200 nM thapsigargin, the amplitude
and the frequency of STOCs gradually decreased. Finally, after 7 min of
superfusion of thapsigargin, STOCs were totally abolished. The
inhibitory effect of thapsigargin was not reversible within the time
scale of these experiments. Together these results suggested that STOCs
require Ca2+ in the SR and that
Ca2+-ATPase, which sequesters
Ca2+ into the SR, also has an
important role.

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Fig. 6.
Role of sarcoplasmic reticulum
Ca2+-ATPase in STOCs. Record
illustrates the application, in the bath solution, of 200 nM
thapsigargin to a cell held at 0 mV. Effect of thapsigargin was similar
but faster than that of ryanodine (compare Figs.
5B and 6). Arrow indicates the 0 current level.
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Effects of hypoxia on steady-state outward
current. The effect of hypoxia was tested on 11 cells.
Ramp depolarization between
90 and 0 mV from a holding potential
of
60 mV elicited the two outward currents. For eight cells, 10 min of hypoxia decreased both STOCs and the amplitude of the
steady-state outward current (Fig.
7A).
Upon reoxygenation, the effect of hypoxia was partially reversible on
the steady-state outward current (data not shown) but was not
reversible on STOCs (see Fig. 9). For three cells (which did not
present STOCs), hypoxia increased the amplitude of the steady-state
outward current or had no effect (data not shown).

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Fig. 7.
Effects of hypoxia on steady-state outward current and on STOCs
elicited by voltage ramps. A: when the
cell was superfused by saline solution containing 1.8 mM
Ca2+, 10 min of hypoxia decreased
STOCs and depressed the amplitude of steady-state outward current.
B: when the cell was bathed in 0 Ca2+ solution, 10 min of hypoxia
increased the amplitude of steady-state outward current. Arrows and the
dashed lines in A and
B indicate the 0 current level.
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When we changed the bath solution to
Ca2+-free solution for 10 min, we
first observed an increase in the steady-state outward current as
already described in Fig. 4A, and then
hypoxia never decreased the steady-state outward current (Fig.
7B), but, in contrast, hypoxia
increased in amplitude. This effect was observed in all six cells
tested and was clearly seen for membrane potential positive to
20 mV. For two cells, hypoxia also decreased the amplitude of
the steady-state outward current for membrane potential between
40 and
20 mV.
Seven cells were voltage clamped with the voltage-ramp protocol in the
presence and in the absence of external thapsigargin. Figure
8 shows that the amplitude of the outward
current was decreased in the presence of external solution containing
thapsigargin (200 nM) during ~10 min. This effect was observed in all
of the cells tested. When the bath solution was further changed to a
hypoxic solution containing thapsigargin (200 nM), 10 min of hypoxia
increased the amplitude of steady-state outward current in four of the
seven cells (Fig. 8C). This effect,
which was partially reversible, was similar to the effect of hypoxia
observed in Ca2+-free solution
(Fig. 7B). In three cells, hypoxia
had no effect on the amplitude of the steady-state outward current.

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Fig. 8.
Effects of thapsigargin on steady-state outward currents elicited by
voltage ramp in normoxia and in hypoxia. Traces of currents elicited by
a voltage-ramp protocol in a cell recorded in control conditions
(trace A,
PO2 = 145 mmHg) and after exposure to
200 nM thapsigargin (trace B,
PO2 = 145 mmHg). In the same
experiment and in the presence of thapsigargin, 10 min of hypoxia
increased the amplitude of steady-state outward current
(trace C,
PO2 = 10 mmHg). Arrow and the dashed
line indicate the 0 current level.
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Effects of hypoxia on STOCs. To test
the effect of hypoxia on STOCs, cells were superfused with a solution
equilibrated with 100% N2. Figure
9 shows STOCs in a cell held at 0 mV before
and after a period of hypoxia. When the cell was in normoxia, a
PO2 of ~150 mmHg, the activity of
STOCs was stable for the 5 min of recording. Changing the bath solution
to a hypoxic PSS solution decreased the
PO2 to ~10 mmHg, and then we
observed a decrease in the frequency of STOCs. After 10 min of hypoxia,
STOCs had almost totally disappeared. Next, the solution was
reoxygenated by changing the bath solution to normoxic PSS solution for
10 min. Upon reoxygenation, STOCs did not recover within the time scale
of these experiments. To evaluate the effect of hypoxia and
reoxygenation on STOCs, we measured the frequency of STOCs in eight
cells for 60-s periods after 4 min in control conditions, after 9 min
of hypoxia, and after 9 min of reoxygenation. The frequency decreased
from 0.37 ± 0.46 Hz in control to 0.12 ± 0.22 Hz in
hypoxia and to 0.07± 0.19 Hz upon reoxygenation. Compared with
normoxia, the frequencies were significantly lower during hypoxia and
after reoxygenation. This effect was not due to a run-down phenomenon
since, in control experiments, we did not observe a significant
decrease in STOC frequency over a similar time scale (see above).

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Fig. 9.
Effect of hypoxia-reoxygenation on STOCs.
A: membrane currents recorded from a
cell voltage clamped at 0 mV. B:
variation of PO2 in the experimental
chamber during the superfusion of normoxic physiological salt solution
(PSS) and PSS equilibrated with N2
(see METHODS). When the
PO2 in the chamber decreased from
~150 mmHg (normoxia) to 10 mmHg (hypoxia), STOC frequency decreased
progressively. Even after return to normoxia
(PO2 150 mmHg), this effect was not
reversible. Arrow in A indicates the 0 current level.
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Seven cells were voltage clamped at 0 mV during hypoxia and
reoxygenation in the presence of thapsigargin. Figure
10 shows that, during 5 min in a normoxic
PSS solution, STOC activity was stable. The application of 200 nM
thapsigargin to the bath solution in normoxia decreased the amplitude
and frequency of STOCs before totally abolishing them after 8 min. The
PO2 in the bath solution was then
decreased from 150 mmHg to ~10 mmHg in the continued presence of
thapsigargin. This had no visible effect on STOCs, and during 10 min of
hypoxia no STOCs were observed. Upon reoxygenation in the presence of
thapsigargin, STOCs rapidly reappeared but with smaller amplitudes and
lower frequencies than those seen in the initial control condition. In
five of the seven cells held at 0 mV and exposed to hypoxia in the
presence of thapsigargin, we observed STOCs upon reoxygenation.

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Fig. 10.
Effect of hypoxia-reoxygenation on STOCs in the presence of
thapsigargin. A: membrane currents
recorded from a cell voltage clamped at 0 mV.
B:
PO2 in the experimental chamber
during the superfusion of normoxic PSS
(PO2 value of 150 mmHg) and PSS
equilibrated with N2
(PO2 <20 mmHg). Thapsigargin (200 nM) abolished STOC activity, and hypoxia had no further effect. Upon
reoxygenation (PO2 recovers to 150 mmHg), we observed a resumption of STOC activity. Arrow in A
indicates the 0 current level.
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DISCUSSION |
Membrane currents recorded in perforated
patch. The steady-state outward current was an outward
current activated at
30 to
40 mV, blocked by 1 mM 4-AP,
and not affected by TEA or 9-AC. These characteristics correspond to
the delayed rectifier K+ current
[IK(dr)]
that has already been observed in these cells (9, 20, 21, 26). Removal
of extracellular Ca2+ was
associated with an increase of the amplitude of
IK(dr). This could be explained by a decrease in a
Ca2+ inward current or by a
screening of surface charges (11).
STOCs, which were observed in >75% of the cells recorded with the
perforated-patch technique, were activated around
30 to
40 mV. Their amplitude increased with depolarization, and they were abolished by 1 mM TEA and were activated by
Ca2+ influx, suggesting that STOCs
represented a Ca2+-activated
K+ current. Furthermore, 1 mM 9-AC
had no effect, suggesting that STOCs were not chloride currents (13).
Importance of external
Ca2+ in STOC
activity.
In Ca2+-free solution, STOCs
totally disappeared, suggesting that a
Ca2+ influx was necessary. Similar
results were found in rabbit cerebral artery and in jejunal smooth
muscle cells (3, 15). However, removal of external
Ca2+ did not suppress STOCs in ear
artery smooth muscle cells (3). This could be explained by a difference
in the arrangement of the Ca2+
stores (3).
The fact that at 0 mV and with
Cd2+ we still observed STOCs
suggests that the L-type Ca2+
current was not indispensable. The decrease in STOC activity in the
presence of Cd2+ may instead
result from an inhibition of Ca2+
leak current, a residual Ca2+
window current of L-type Ca2+
current,
Na+-Ca2+
exchange current (16), or a screening of charge surface (11). Bychkov
et al. (6) showed that Ca2+ entry
through a reverse-mode
Na+-Ca2+
exchanger determines Ca2+ store
refilling and then regulates STOC activity. Because the Na+-Ca2+
exchanger current is blocked by 1 mM
Cd2+ (16), this could explain our
results.
Importance of internal stores of
Ca2+ in STOC
activity.
Caffeine at concentrations >1 mM decreases the threshold of
Ca2+-induced
Ca2+ release (14) before depleting
Ca2+ stores. The initial increase
in STOC activity induced by 5 mM caffeine could be due to the increase
in internal Ca2+ from SR resulting
in Ca2+-induced
Ca2+ release; then, after the SR
was emptied, the STOCs would be abolished. Ryanodine at 10 µM, a
concentration used to deplete the internal store (12, 17), gradually
decreased STOCs. The fact that ryanodine did not completely abolish
STOCs suggested that only a part of the
Ca2+ from the SR responsible for
STOCs was released through the ryanodine receptor, and we cannot
exclude that Ca2+ was also
released through inositol 1,4,5-trisphosphate-induced Ca release.
The internal Ca2+ store dependence
of STOCs was confirmed by the inhibiting action of thapsigargin.
Thapsigargin (200 nM) totally abolished STOCs in rabbit pulmonary
smooth muscle cells. At this concentration, thapsigargin was also known
to abolish STOCs in rat cerebral arterial smooth muscle cells (19).
Thapsigargin is known to block
Ca2+-ATPase of SR in pulmonary
artery (10) and then to deplete
Ca2+ stores (24).
Are STOCs activated by graded release of
Ca2+ from the
SR?
At 0 mV, the amplitudes of STOCs were not uniform, and the step
increment of 1.57 ± 0.56 pA/pF suggested that
Ca2+, which activated STOCs in
these cells, is uniformly released from the SR and particularly from
superficial SR (4). In cells clamped at 0 mV with a 4-137 mM
K+ gradient, the unitary
conductance for Ca2+-activated
K+ channel would be 86 pS, and
with 1 µM intracellular Ca2+,
the open probability would be 0.25 (1). If we assume that the rise in
Ca2+ close to the channel was 1 µM, then 20 channels would be required to open and to induce a
uniform interval of 1.57 ± 0.56 pA/pF. The maximum amplitude of
STOCs would then represent the activation of 160 channels. Nelson et
al. (19) showed that STOCs are activated by spontaneous release of
Ca2+
(Ca2+ sparks) from the SR close to
the sarcolemma. They suggested that one spark activates 13 Ca2+-activated
K+ channels, which is close to the
value that we estimate.
Effect of hypoxia on
IK(dr).
In the presence of external calcium, hypoxia decreased the amplitude of
IK(dr). This
effect was abolished by removal of Ca2+ in the external
solution. This suggested that hypoxia decreases IK(dr) via a
Ca2+-dependent mechanism. Because
10 min in Ca2+-free solution also
depletes Ca2+ from superficial SR,
this source of Ca2+ was also
important in the effect of hypoxia on
IK(dr). This was confirmed by the inhibitory action of thapsigargin (which was applied
10 min before) on the effect of hypoxia. Indeed, thapsigargin is known
to deplete SR Ca2+ stores (24).
These results are consistent with those of Salvaterra and Goldman (23),
who showed that thapsigargin blocked the elevation of internal
Ca2+ concentration induced by
hypoxia. Also, Post et al. (21) showed that hypoxia could release
Ca2+ of SR, which induced a
decrease in the amplitude of
IK(dr). However, this release of Ca2+ may induce an
increase in STOCs in our cells, but we always observed a decrease in
these currents. This could be explained by the use of the
perforated-patch technique instead of the whole cell configuration of
the patch-clamp technique. The mechanism to explain this difference needs to be demonstrated. Indeed, with the perforated-patch technique, Ca2+ homeostasis is less modified
by intrapipette dialysis (22). Also, Post et al. (21) didn't observe
STOCs in their cells.
Next, we suggested that the decrease in
IK(dr) was
blocked by a depletion in Ca2+
concentration of SR.
Effect of hypoxia on STOCs. Ten
minutes of hypoxia decreased the activity of STOCs, and this effect was
not reversible. The effect of hypoxia may be due to an inhibitory
action of hypoxia on the
Ca2+-activated
K+ channel, on the
Ca2+-release channels of the SR,
on Ca2+ influx, or on
Ca2+-ATPase of the SR. An
inhibitory action of hypoxia on
Ca2+-activated
K+ channel and on L-type
Ca2+ current was not likely
because hypoxia had no effect or activated the
Ca2+-activated
K+ channel (2, 5, 21), and the
L-type Ca2+ current was mostly
inactivated at 0 mV (7). It was shown that Ca2+ release by SR was activated
and not inhibited by hypoxia (23, 25). Then the more likely action by
which hypoxia decreased STOCs was an inhibitory action on the
Ca2+-ATPase of the SR, leading to
a decrease in Ca2+ concentration
of superficial SR. Indeed, thapsigargin is known to block
Ca2+-ATPase of the SR in pulmonary
artery (10) and then to deplete Ca2+ stores (24).
However, because the presence of thapsigargin allows the STOCs to
return after hypoxia, this might implicate a different mechanism of
action. More experiments are needed to discover such new mechanisms.
In conclusion, we showed that hypoxia decreases both STOCs and
IK(dr) by a
Ca2+-dependent mechanism.
Thapsigargin and removal of external
Ca2+ abolished the effect of
hypoxia on
IK(dr),
suggesting that hypoxia decreases
IK(dr) by a
Ca2+-dependent mechanism that
depends on the Ca2+ concentration
of SR.
 |
ACKNOWLEDGEMENTS |
We thank Dr. Ian Findlay for helpful criticisms on the manuscript.
We thank Dr. Dominique Thuringer for useful discussions and for helping
in isolating cells, Dr. Claire Malécot for critical reading of
the paper, Maryse Pingaud for technical assistance, Gilles Pinal for
building some electronic devices, and Chantal Boisseau for secretarial
assistance.
 |
FOOTNOTES |
This work was supported by le Ministère de l'Enseignement
Supérieur et de la Recherche and la Fondation pour la Recherche Médicale.
Address for reprint requests: P. Bonnet, UMR CNRS 6542, Physiologie des
Cellules Cardiaques et Vasculaires, Faculté de Médecine, 2 bis, Boulevard Tonnelé, B.P. 3223, 37032 Tours Cedex, France.
Received 27 May 1997; accepted in final form 20 March 1998.
 |
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