Membrane currents in canine bronchial artery and their
regulation by excitatory agonists
Q. J.
Li and
L. J.
Janssen
Asthma Research Group, Father Sean O'Sullivan Research
Center, St. Joseph's Hospital, Department of Medicine, McMaster
University, Hamilton, Ontario, Canada L8N 4A6
 |
ABSTRACT |
The bronchial
vasculature plays an important role in airway physiology and
pathophysiology. We investigated the ion currents in canine bronchial
smooth muscle cells using patch-clamp techniques. Sustained outward
K+ current evoked by step depolarizations was significantly
inhibited by tetraethylamonium (1 and 10 mM) or by charybdotoxin
(10
6 M) but was not significantly affected by
4-aminopyridine (1 or 5 mM), suggesting that it was primarily a
Ca2+-activated K+ current. Consistent with
this, the K+ current was markedly increased by raising
external Ca2+ to 4 mM but was decreased by nifedipine
(10
6 M) or by removing external Ca2+. When
K+ currents were blocked (by Cs+ in the
pipette), step depolarizations evoked transient inward currents with
characteristics of L-type Ca2+ current as follows:
1) activation that was voltage dependent (threshold and
maximal at
50 and
10 mV, respectively); 2) inactivation that was time dependent and voltage dependent (voltage causing 50%
maximal inactivation of
26 ± 22 mV); and 3) blockade
by nifedipine (10
6 M). The thromboxane mimetic U-46619
(10
6 M) caused a marked augmentation of outward
K+ current (as did 10 mM caffeine) lasting only 10-20
s; this was followed by significant suppression of the K+
current lasting several minutes. Phenylephrine (10
4 M)
also suppressed the K+ current to a similar degree but did
not cause the initial transient augmentation. None of these three
agonists elicited inward current of any kind. We conclude that
bronchial arterial smooth muscle expresses Ca2+-dependent
K+ channels and voltage-dependent Ca2+ channels
and that its excitation does not involve activation of Cl
channels.
potassium; calcium; chloride; adrenergic; U-46619
 |
INTRODUCTION |
THE BRONCHIAL
VASCULATURE plays a very important role in maintaining the normal
function of the respiratory system: it nourishes the airway wall,
conditions inspired air, and is involved in defense and clearance of
the airways. Dysfunction of bronchial vasculature may contribute to
asthma, particularly exercise- and cold/dry air-induced
bronchoconstriction (22), and to edema formation in the
lung occurring after acute lung injury with smoke inhalation and acid
aspiration (1, 4, 17); also, adequate bronchial blood flow
is now recognized to be vital to postoperative success after lung
transplantation (12, 16).
Like most systemic vasculature, the bronchial circulation is regulated
primarily via an excitatory adrenergic innervation and inhibitory
nonadrenergic noncholinergic innervation (3, 25) and is
also regulated by blood-borne autacoids, including inflammatory
mediators; the primary constrictor autacoid among the latter is
thromboxane A2 (3, 15).
Constriction and relaxation of vascular smooth muscle is mediated in
part through changes in membrane conductances: in general, excitatory
stimuli cause opening of voltage-dependent Ca2+ channels,
usually via activation of Cl
channels and subsequent
membrane depolarization, whereas vasodilators activate K+
channels, leading to membrane hyperpolarization and decreased Ca2+ influx (13). Although much work has been
done to investigate the ion channels and their regulation in other
systemic arteries, there have been no electrophysiological studies of
the bronchial vasculature. The purpose of this study, then, was to
classify the ion channels present in freshly dissociated canine
bronchial arterial smooth muscle cells and their regulation by
excitatory agonists.
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MATERIALS AND METHODS |
Cell isolation.
Adult mongrel dogs (15-30 kg; either sex) were killed by
intravenous injection of pentobarbital sodium (30 mg/kg), and lobes of
lung were excised. Bronchial vasculature (0.5-1 mm OD) on third- to fifth-order airways was exposed by removing overlying connective tissue and parenchyma. Bronchial veins were generally sparse or completely absent in these intraparenchymal airways (since the bronchial circulation drains into the pulmonary veins) and are easily
distinguished from the bronchial arteries on the basis of diameter and
relative wall thickness; we used only bronchial arteries in these studies.
Tissues were either used immediately or stored at 4°C for use the
next day. We found no functional differences in tissues that were
studied immediately compared with those used after 24 h of
refrigeration. Tissues were transferred to digestion solution that
contained collagenase and elastase (see composition in
Solutions) and were incubated at 37°C for 1 h, after
which they were gently triturated to liberate individual myocytes.
These were allowed to settle and adhere to the bottom of a recording
chamber (1 ml volume) and were superfused with standard Ringer solution
at a rate of 2-3 ml/min; unless indicated otherwise, experiments
were conducted at room temperature. The dissociated cells were studied within 8 h after dissociation. Electrophysiological responses were
tested in cells that were phase dense and appeared relaxed.
Electrophysiological study.
The majority of recordings were made using the nystatin
perforated-patch configuration of the conventional patch-clamp
recording technique. Pipette tip resistances ranged from 3 to 5 M
,
and access resistances ranged from 9 to 39 M
(70-80%
compensated). Membrane currents were filtered at 5 kHz and sampled at 2 Hz. Acquisition and analysis of data were accomplished using Axopatch 200B and pCLAMP8 software (Axon Instruments, Foster City, CA).
Solutions.
Digestion solution contained collagenase (blend F; 0.9 U/ml; Sigma),
elastase (type IV, 12.5 U/ml), and BSA (1 mg/ml) in Ca2+-
and Mg2+-free Hanks' solution (pH 7.4).
For most of the recordings, we used standard Ringer solution containing
the following (in mM): 130 NaCl, 5 KCl, 1 CaCl2, 1 MgCl2, 20 HEPES, and 10 D-glucose (pH 7.4).
Ca2+-free media was prepared by omitting CaCl2
and adding 1 mM EGTA. The pipette solution generally contained the
following (in mM): 140 KCl, 1 MgCl2, 0.4 CaCl2,
20 HEPES, 1 EGTA, and 150 U/ml nystatin (pH 7.2).
For recordings of Ca2+ currents, on the other hand, we used
Ringer solution with Ca2+ substituted by 5 mM
Ba2+ and a modified pipette solution. When the
perforated-patch configuration was employed, we used a pipette solution
consisting of the following (in mM): 130 CsCl, 10 tetraethylammonium
(TEA), 1 MgCl2, 20 HEPES, 5 EGTA, and 150 U/ml nystatin (pH
7.2). For whole cell recordings of Ca2+ current, this
pipette solution was supplemented with 4 mM ATP (Na+ salt)
and 0.3 mM GTP (Na+ salt).
Chemicals.
All chemicals were obtained from Sigma Chemical. TEA, 4-aminopyridine,
phenylephrine, and caffeine were prepared as aqueous stock solutions.
Nifedipine and U-46619 were dissolved in absolute ethanol and then
diluted with bathing medium; the final concentration of ethanol in the
application pipette was 0.01%. Phenylephrine, caffeine, and U-46619
were applied by pressure ejection (Picospritzer II; General Valve,
Fairfield, NJ) from micropipettes placed close to the cells, whereas
ion channel blockers were applied directly via the bathing medium.
Statistics.
Data are expressed as means ± SD. Statistical significance
(P < 0.05) was determined using a two-tailed
Student's t-test.
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RESULTS |
Outward K+ current.
At resting membrane potentials, the canine bronchial arterial smooth
muscle cells were quiescent, with an input resistance ranging from 1 to
10 G
and no spontaneous current activity whatsoever (Fig.
1A); none of the cells we
studied (~100 cells from >25 animals) exhibited any spontaneous
transient inward currents like those we have described previously in
airway smooth muscle cells (8) or that others have found
in vascular smooth muscle (21, 23). At more depolarized
potentials, these cells generally exhibited spontaneous transient
outward currents (STOCs; Fig. 1, B and C).

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Fig. 1.
Basal currents. A: at a holding potential
(Vh) of 60 mV and in the absence of any exogenous
agonist, canine bronchial smooth muscle cells exhibited no spontaneous
currents of any kind. In two different cells held at 0 mV,
however, the membrane current was quite "noisy" (B and
C) and often accompanied by spike-like transient outward
currents up to 100 pA in amplitude (C).
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Stepwise depolarizing commands (20-mV increments) from a holding
potential of
70 mV evoked large sustained outward currents often
accompanied by STOCs (Fig. 1; n = 44). The average
amplitude of the current at 70 mV was 49 ± 8 pA/pF
(n = 44). This outward current exhibited little or no
inactivation, decreasing <10% over the course of the depolarizing pulse.
Vascular smooth muscle cells generally exhibit two major types of
K+ currents (Ca2+ activated and delayed
rectifier) that can be distinguished on the basis of sensitivity to
TEA, 4-aminopyridine, and charybdotoxin. We found that TEA
significantly inhibited the sustained and transient outward currents,
leaving only a small outward current devoid of any spike-like outward
currents: currents at +70 mV were reduced to 50 ± 14 and 84 ± 3% of control in the presence of 1 mM TEA (6 cells;
n = 5) or 10 mM TEA (7 cells; n = 6;
Fig. 2A), respectively. These
inhibitory effects of TEA were reversible upon washout of TEA.
Likewise, charybdotoxin (10
6 M) also significantly
inhibited the sustained outward current (67 ± 13% of control at
+70 mV; 5 cells, n = 4) and abolished the large
spike-like outward currents in a reversible fashion (Fig.
2C). 4-Aminopyridine, on the other hand, had no significant effect on the currents at bath concentrations of 1 mM (7 cells; n = 6) nor 5 mM (6 cells; n = 5; Fig.
2B).

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Fig. 2.
Pharmacological sensitivity of outward currents. Step
depolarizations (20-mV increments) from a holding potential of 70 mV
evoked large sustained outward currents upon which spike-like transient
outward currents were often superimposed (see A,
B, and C, left). The sustained and
transient outward currents were markedly suppressed by
tetraethylammonium (TEA; A) or by charybdotoxin
(B) but not by 4-aminopyridine (C).
Left and middle, representative examples of
effects; right, mean effects of TEA (n = 6),
4-aminopyridine (4-AP; n = 5), and charybdotoxin (ChTX;
n = 4).
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These data suggest that a substantial portion of the outward current
evoked by depolarizing pulses is a Ca2+-dependent
K+ current, with little or no contribution from
voltage-dependent delayed-rectifier K+ currents.
Role of Ca2+ influx in
K+ current activation.
Given that the outward currents triggered by membrane-depolarizing
pulses are predominantly Ca2+ dependent, we next
investigated the contribution of external Ca2+ to their activation.
The magnitudes of the outward K+ currents were
significantly reduced, but not abolished, when the external bathing
medium was replaced with a Ca2+-free buffer and were
restored to control levels by reintroduction of external
Ca2+ (Fig. 3A). On
the other hand, their magnitudes were markedly increased when external
Ca2+ concentration was increased to 4 mM (Fig.
3A). On average, the magnitude of the current evoked by a
depolarizing pulse to +70 mV was inhibited 61 ± 8% in
Ca2+-free media (n = 6; P < 0.05) and augmented slightly (but not significantly) to 116 ± 68% of control in 4 mM Ca2+-containing buffer (7 cells
from 6 dogs; Fig. 3C).

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Fig. 3.
Role of external Ca2+ in outward K+
currents. A: K+ currents evoked using step
depolarizations (see legend for Fig. 2) were markedly reduced in size
when external Ca2+ was omitted, markedly increased in size
when Ca2+ concentration ([Ca2+]) was
increased to 4 mM, and then restored to normal amplitudes when
[Ca2+] was returned to 1 mM. B: application of
nifedipine (10 6 M) during step depolarizations also
markedly decreased the amplitudes of the outward currents in a
reversible fashion. Mean amplitudes of K+ currents recorded
under these conditions are given in C (n = 4-6). [Ca2+]ext, external
[Ca2+].
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These data clearly indicate that external Ca2+ can markedly
influence the magnitude of the K+ currents evoked by
depolarizing pulses. We therefore investigated the effect of
nifedipine, a selective blocker of voltage-dependent Ca2+
channels, on these outward K+ currents. Nifedipine
(10
6 M) also markedly reduced K+ currents in
a reversible fashion (Fig. 3B); on average, K+
currents evoked by step depolarizations to +70 mV were inhibited by
72 ± 4% (n = 4; P < 0.05; Fig.
3D).
Voltage-dependent Ca2+ current.
The data presented above provide indirect evidence for
voltage-dependent Ca2+ channels in these cells. We examined
the Ca2+ currents directly by replacing K+ in
the pipette solution with Cs+ to block K+
currents (TEA was also present in the pipette solution, for the same
purpose) and by replacing Ca2+ in the bathing medium with
Ba2+ (5 mM) to augment the inward current.
Depolarizing step commands (10-mV increments) from a holding potential
of
70 mV evoked transient inward currents (Fig.
4). Activation of these currents was time
dependent (peak activation occurring within 24 ± 13 ms;
n = 13) and voltage dependent (threshold approximately
equal to
30 mV, half-maximal at
3 ± 7 mV, and maximal at +20
mV; Fig. 4). The currents also inactivated in a fashion that was time
dependent; mean
for inactivation at
10 mV was 206 ± 53 ms.
Repolarization to the holding potential did not evoke slowly
decaying tail currents reminiscent of Ca2+-dependent
Cl
current (10) nor of T-type
Ca2+ currents (6).

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Fig. 4.
Voltage-dependent Ca2+ currents. A: in a
cell dialyzed with Cs+-containing pipette solution and held
at 70 mV, depolarizing steps evoked inward currents that were
reversibly abolished by nifedipine (10 6 M). B:
mean peak amplitudes of these inward currents plotted against membrane
voltage. C: same cell represented in A was held
under voltage clamp for 5 s at conditioning potentials ranging
from 100 to +30 mV, after which inward Ca2+ currents were
evoked by a step to +10 mV. D: Boltzmann analysis of voltage
dependence of activation ( ) and inactivation
( ) in one cell reveals incomplete voltage-dependent
inactivation and substantial "window current" at voltages more
positive than 30 mV. I/Imax, ratio
of current to maximal current.
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We used a different voltage protocol to examine the voltage dependence
of inactivation as follows: cells were held under voltage clamp at
potentials ranging from
100 to +30 mV (10-mV increments) for 4.9 s ("conditioning pulses"), followed by return to the holding potential (of
70 mV) for 54 ms and then a test pulse to +10 mV (1 s
duration). A typical plot of the magnitude of the test pulses against
voltage of the conditioning pulses is shown in Fig. 4D. Voltage-dependent inactivation was incomplete, and Boltzmann analysis of the data yielded a mean voltage causing 50% maximal inactivation of
26 ± 2 mV (n = 5).
Effects of caffeine on membrane currents.
In addition to Ca2+ influx pathways, intracellular
Ca2+ concentration ([Ca2+]i) can
also be elevated by release of internally sequestered Ca2+
(2, 20). Millimolar concentrations of caffeine trigger
release of internal Ca2+ by enhancing the opening of
ryanodine receptors on the sarcoplasmic reticulum. This release is
transient, however, so it is generally not possible to characterize the
effects of caffeine on membrane currents evoked by a series of
depolarizing step commands. Instead, we examined the effects of
caffeine while holding the membrane potential constant at various
levels. When the cells were held under voltage clamp at 0 mV, caffeine
(10 mM) evoked a substantial outward current that was transient in
nature, rising to a mean peak value of 688 ± 248 pA and then
decaying to baseline over the course of 2-3 s even though caffeine
continued to be applied (Fig.
5A). Caffeine also evoked
contractions (data not shown). When the same cells were held at
60
mV, however, caffeine had no discernible effect on membrane current
(Fig. 5B).

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Fig. 5.
Membrane currents evoked by caffeine. A: in a cell held
under voltage clamp at 0 mV, caffeine (10 mM; 1 s application from
puffer pipette) evoked large transient outward current that peaked
within 2 s and decayed completely back to baseline. B:
at a holding potential of 60 mV, however, this same cell showed no
response to caffeine. C: in another cell, ramp
depolarizations (from 70 to +50 mV) were used to explore the entire
current-voltage relationship of the caffeine responses; although
caffeine (10 mM) did cause cell contraction (data not shown) and
substantial augmentation of outward current, it did not evoke any
inward current whatsoever.
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We also investigated the effects of caffeine using ramp depolarizing
commands from a holding potential of
70 up to +50 mV at a rate of 120 mV/s. In this way, it is possible to capture the complete
current-voltage relationship of the caffeine-evoked currents, although
it must be understood that both [Ca2+]i (and
thus the degree of current enhancement) and membrane potential are
changing at the same time but at different rates and/or in different
directions. Before application of caffeine, ramp depolarizations evoked
only outward current with a similar current-voltage relationship as the
K+ currents described above (compare Fig. 1 with Fig.
5C); no inward currents were seen in any cell held at
negative membrane potentials (17 cells; n = 9). Upon
application of caffeine (10 mM), the outward currents were increased
499 ± 40% (8 cells, n = 5; P < 0.01), as shown in Fig. 5C. However, there was still no
indication whatsoever of any inward (i.e., Cl
) current in
any of the cells tested (Fig. 5C).
Effect of phenylephrine and thromboxane mimetic on membrane
currents.
In general, the adrenergic innervation, through its actions on
-adrenoceptors, represents the primary excitatory neural input for
the bronchial vasculature (25). Thromboxane A2
is also a potent spasmogen in many vascular beds (15). We
therefore tested the effects of the
-adrenoceptor agonist
phenylephrine and the thromboxane mimetic U-46619 on membrane currents
using the same protocol described above for caffeine-evoked responses.
Although phenylephrine (10
4 M) did cause the cells to
contract (data not shown), it did not evoke any inward current during maintained voltage clamp at
60 mV or when a range of voltages was
tested using ramp depolarizations or incrementing step commands (n = 6; data not shown). Also, although adrenergic
agonists cause release of internally sequestered Ca2+ in
these cells (7), we did not observe a transient
augmentation of K+ currents upon application of
phenylephrine during step commands (Fig.
6A) or ramp depolarizations.
To the contrary, we noted a delayed but prolonged suppression of
K+ currents: for example, Fig. 6A shows the
magnitude of K+ currents evoked in one cell using
depolarizing step commands to +10 mV before and after 2 min of exposure
to phenylephrine (10
4 M). On average, K+
currents were suppressed to 65 ± 13% of control by phenylephrine when bath temperature was 37°C (n = 4); at room
temperature, however, phenylephrine had very little effect on the
currents, reducing them to only 94 ± 8% (n = 4).

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Fig. 6.
Effects of phenylephrine (PE) on membrane currents. Step
depolarizations to +10 mV (from a holding potential of 70 mV)
delivered to a bronchial arterial smooth muscle cell at 10-s intervals
evoked outward K+ currents. A: representative
tracings. B: mean amplitude of currents in B.
Application of PE (10 4 M in puffer pipette; 1 s
application) did not evoke any inward current but did cause a slowly
developing, long-lasting, and reversible suppression of the outward
K+ currents (A and B).
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U-46619 (10
6 M) also did not evoke any inward current
whatsoever in any cells tested (e.g., Fig.
7A) but did cause a marked enhancement of K+ currents in many but not all of the cells
(5 of 11 cells studied, lasting 10-20 s), followed by a marked
suppression of the same (e.g., Fig. 7, A and B).
On average, K+ currents were reduced to 63 ± 11%
(n = 4); this electrophysiological response was
accompanied by contraction of the cells (data not shown). We did not
compare the effect of temperature on these responses to U-46619.

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Fig. 7.
Effects of U-46619 on membrane currents. A:
representative tracing showing repeated ramp depolarizations (from 70
to +50 mV) and outwardly rectifying K+ currents accompanied
by spontaneous transient outward currents that they evoked.
B: from data shown in A, certain consecutive
membrane current responses were averaged (indicated by open boxes in
A labeled a-c), decimated (i.e.,
only every 10th sample taken), and then superimposed. U-46619
(10 6 M in application pipette; 1-s applications), applied
two times at different distances from the cell, caused these currents
to be first augmented (b) and then suppressed (c)
compared with control (a).
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DISCUSSION |
The bronchial circulation plays an important role in many aspects
of airway physiology and pathophysiology. Thus a thorough understanding
of the mechanisms underlying excitation-contraction coupling in this
vascular bed would be highly clinically valuable. In this study, we
provide the first electrophysiological description of bronchial
arterial smooth muscle cells.
K+ currents.
We found the K+ current evoked by depolarizing
voltage commands in the bronchial artery to be predominantly
Ca2+ dependent in nature, since it was 1)
markedly and reversibly inhibited by TEA or charybdotoxin but not by
4-aminopyridine; 2) reduced substantially by interfering
with Ca2+ influx using nifedipine or by removing external
Ca2+ but increased by raising external Ca2+;
and 3) enhanced by releasing internally sequestered
Ca2+ using caffeine. Voltage-dependent K+
current, on the other hand, appears to make little or no contribution, since 4-aminopyridine had very little effect on the
depolarization-evoked currents. In general, Ca2+-dependent
K+ currents play an important role in the regulation of
many vascular smooth muscles, as discussed in more detail below
(13, 18, 19).
Ca2+ currents.
The Ca2+ currents that we recorded in these cells exhibited
many characteristics of "L-type" current, including: 1)
threshold and peak activation at approximately
30 and +20 mV,
respectively; 2) half-maximal inactivation at approximately
25 mV; 3)
for inactivation of ~200 ms; 4)
absence of slowly decaying tail currents upon repolarization; and
5) sensitivity to dihydropyridines. Inactivation of these
currents was incomplete, leaving substantial "window current" at
all voltages more positive than
30 mV. This has tremendous physiological importance, since it allows for sustained influx of
Ca2+ during agonist stimulation, which leads to membrane
depolarization. Activation of these Ca2+ currents is
sufficient to evoke contraction, as indicated by the substantial
contractions that we have previously described in intact tissues
exposed to KCl (7).
Cl
currents.
Although many vascular smooth muscle cells exhibit
Ca2+-dependent Cl
currents spontaneously
and/or when internal [Ca2+]i is elevated by
various stimuli (14, 18, 23), some appear not to
(18). Interestingly, we found no evidence for inward current of any kind in these bronchial arterial smooth muscle cells,
neither spontaneously nor in response to voltage-dependent Ca2+ influx (Fig. 4) or release of internally sequestered
Ca2+ induced by caffeine (Fig. 5), phenylephrine (Fig. 6),
or U-46619 (Fig. 7), even though these stimuli are sufficient to cause
cell shortening and substantial augmentation of K+ current
(except for phenylephrine). Thus we conclude that these tissues do not
express functional Cl
channels.
Spasmogen-evoked changes in membrane currents.
In most arterial preparations, agonist-evoked contractions
involve activation of inward Cl
(14) and/or
nonselective cation currents (11), leading to membrane
depolarization and subsequent opening of voltage-dependent Ca2+ channels such as the ones described in this tissue. It
is for this reason that Ca2+ channel blockers are so
effective in the treatment of hypertension.
Electromechanical coupling is also sufficient to produce contractions
in the bronchial artery, as indicated by the robust contractions evoked
by high-millimolar concentrations of KCl (7). However, in
the present study, we were unable to identify any spontaneous or
agonist-stimulated inward currents in these cells, although they are
present in other vascular beds (11, 14) and we have been
able to demonstrate them in airway smooth muscle cells using identical
experimental techniques (5, 8, 9). Another mechanism by
which excitatory autacoids can evoke membrane depolarization in
vascular smooth muscle (other than activation of an inward current) is
suppression of outward current (19, 24). The resting
membrane potential in the cells that we studied appears to be as low as
50 mV, as indicated by the current-voltage relationships in Fig. 2.
It is clear, then, that there is a physiologically relevant
K+ conductance active at rest, since only this type of
conductance can hyperpolarize the membrane to this degree; the
equilibrium potentials for all other ions are much more positive than
this (those for Cl
and Mg2+ are both
0 mV,
whereas those for Na+ and Ca2+ are both very
positive). As such, any decrease in the membrane permeability to
K+ will lead to depolarization; in preliminary experiments,
we have found that TEA (5 mM) does evoke substantial contraction in
intact tissues (data not shown). In fact, we did observe substantial suppression of outward K+ current after application of
either phenylephrine or U-46619 (although the latter often briefly
enhanced the K+ current). This suppression lasted several
minutes, long after application of the agonists had ended. Moreover,
adrenergic suppression was essentially abolished at room temperature,
whereas the effect of U-46619 was not; this parallels our previous
finding that adrenergic contractions were essentially abolished by
cooling to room temperature, whereas those evoked by U-46619 were
hardly affected (7). The mechanism underlying this
suppression was not investigated in the present study. However, the
inability of caffeine to suppress K+ currents suggests that
this is not related to changes in Ca2+ concentration (since
caffeine, phenylephrine, and U-46619 all release internal
Ca2+). Instead, it likely involves G proteins (which are
not normally activated by caffeine); these may act directly on the
channels or stimulate downstream events such as activation of
phospholipases and various kinases (20). The effects of
caffeine on membrane current and mechanical activity are not secondary
to inhibition of phosphodiesterase activity, since they were very
transient, resolving to baseline within 20 s after application of
caffeine; this time course mirrors that of caffeine-induced release of
internally sequestered Ca2+.
Thus it appears that excitation-contraction coupling in bronchial
smooth muscle involves membrane depolarization (via suppression of
outward current) with voltage-dependent Ca2+-influx and
nonelectromechanical coupling mechanisms (15). The latter
mechanism may be more important than the former,
particularly at lower temperatures (which are also
physiologically relevant given the cooling of the airway during
accelerated ventilation), since we have previously shown that
excitatory mechanical responses in this tissue involve increased
Ca2+ sensitivity of the contractile apparatus much more
than elevation of [Ca2+]i per se
(7).
In conclusion, canine bronchial artery smooth muscle cells exhibit
Ca2+-dependent K+- and voltage-dependent
(L-type) Ca2+ currents, with little or no evidence of
voltage-dependent "delayed-rectifier" K+ currents or
Cl
currents. The thromboxane mimetic U-46619 caused a
transient increase of outward K+ currents, followed by
marked suppression of the same that lasted several minutes. Adrenergic
stimulation only produced the sustained suppression of K+
currents (not their enhancement), and only at a physiological temperature. Neither agonist activates inward (i.e., Cl
)
current at all in this tissue. These electrophysiological data explain
the mechanical effects produced by these agonists.
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ACKNOWLEDGEMENTS |
These studies were supported by operating funds from the Canadian
Institutes of Health Research and a Scientist Award from the Medical
Research Council of Canada.
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FOOTNOTES |
Address for reprint requests and other correspondence:
L. J. Janssen, Dept. of Medicine, McMaster Univ., 50 Charlton Ave. E., Hamilton, Ontario, Canada L8N 4A6 (E-mail:
janssenl{at}mcmaster.ca).
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. Section 1734 solely to indicate this fact.
First published January 11, 2002;10.1152/ajplung.00421.2001
Received 30 October 2001; accepted in final form 10 January 2002.
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