1 Smooth Muscle Research Group, Faculty of Medicine, University of Calgary, Calgary, Alberta T2N 4N1; 4 Department of Physiology, Faculty of Medicine, University of Manitoba, Winnipeg, Manitoba, Canada R3T 2N2; 2 Department of Physiology, Faculty of Medicine, University of Iceland, Reykjavik, Iceland IS-101; and 3 Centro de Investigaciones Cardiovasculares, Facultad de Medicina, Universidad Nacional de La Plata, 1900 La Plata, Argentina
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ABSTRACT |
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The properties of delayed
rectifier K+ current
[IK(dr)] of canine airway smooth
muscle cells isolated from small bronchi and its modulation by protein
kinase C (PKC) were studied by whole cell patch clamp.
IK(dr) activated
positive to 40 mV, with half-maximal activation at
16 ± 1.2 mV (n = 15) and average
current density of 31 ± 2.6 pA/pF
(n = 15) at +30 mV. The capacitive
surface area, current density, and voltage dependence of activation of
IK(dr) of
myocytes of ragweed pollen-sensitized dogs were not different from
age-matched control dogs. However, the sensitization reduced the
availability of
IK(dr) between
40 and
20 mV due to a hyperpolarizing shift in the
voltage dependence of steady-state inactivation (
29.9 ± 1.2 in sensitized versus
26.0 ± 0.7 mV in control dogs,
n = 9 and 11, respectively;
P < 0.05). PKC activation
with diacylglycerol analog or phorbol ester depressed
IK(dr) amplitude,
whereas an inactive diacylglycerol analog had no effect. The
hyperpolarizing shift in voltage dependence of inactivation
and/or modulation of
IK(dr) by PKC may
be two mechanisms that contribute to the enhanced reactivity of
bronchial tissues from ragweed pollen-sensitized dogs.
airway smooth muscle; asthma; ragweed pollen sensitization; voltage-gated potassium channel; protein kinase C
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INTRODUCTION |
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THE LEVEL OF CONTRACTILE tone exhibited by smooth muscle cells is dependent on intrinsic (myogenic) and extrinsic (neuronal, epithelial, and hormonal) factors that regulate plasmalemmal ion channel activities, Ca2+ release from the endoplasmic reticulum, intracellular Ca2+ concentration, and the interaction of actin and myosin filaments, i.e., cross-bridge cycling (17). Dysfunctional control of smooth muscle tone contributes to a variety of pathological conditions. For example, asthma has been attributed to alterations in control of airway smooth muscle contractility; anaphylactic bronchospasm is associated with a marked hyperreactivity of airway smooth muscle to a variety of pharmacological and physical stimuli, including histamine (8, 33).
The level of resting membrane potential is important for the control of
airway smooth muscle tone (13, 33). Action potentials are not observed
in bronchial muscle. Rather, contractile tone is postulated to result
from maintained, nonregenerative depolarizations that increase the open
probability of voltage-dependent L-type Ca2+ channels (13, 14, 33).
Sustained Ca2+ influx due to
steady-state activation of L-type
Ca2+ channels occurs over a narrow
range of membrane potentials between 50 and
20 mV and is
known to cause significant elevations in intracellular
Ca2+ levels in isolated airway
myocytes (13, 14). A role for 4-aminopyridine (4-AP)-sensitive delayed
rectifier K+ channels
(Kdr) in contributing to control
of membrane potential, Ca2+
channel activity, and tone is indicated for the airways; exposure of
intact tissues and/or isolated tracheal/bronchial myocytes of
dogs, ferrets, and humans to 4-AP leads to depolarization and contraction (1, 15, 23). The biophysical properties of delayed
rectifier K+ current
[IK(dr)] are
well described for tracheal myocytes; however, little is known
concerning this conductance in smaller-caliber airways. This is
potentially significant since the site of airway dysfunction in
pathological conditions, such as asthma, is either the central (2nd to
6th order) bronchi (immediate asthmatic response) or the peripheral
bronchi (delayed asthmatic response; see Ref. 36).
Previous studies using a canine model of bronchial hyperreactivity demonstrated that tracheal smooth muscle of dogs sensitized with ovalbumin possessed an enhanced contractile response to histamine mediated by H1 receptors, as well as spontaneous phasic contractile activity and an increased myogenic response (4, 5). Similar findings were subsequently obtained for airway smooth muscle from second- to sixth-order bronchi of ragweed-sensitized dogs, including an increased capacity and velocity of shortening of the smooth muscle and increased release of histamine to ragweed pollen antigen (8, 22). The second- to sixth-order bronchi were also shown to be more sensitive to ragweed pollen compared with tracheal smooth muscle by several orders of magnitude (39). The characteristics of bronchial smooth muscle of ragweed-sensitized dogs have been reviewed (43).
Protein kinase C (PKC) of airway and nonairway smooth muscles is activated by a variety of contractile agonists, including acetylcholine and histamine, and enhanced PKC activity has been implicated to contribute to the pathogenesis of asthma (17, 34, 35). We recently obtained evidence that IK(dr) of vascular smooth muscle cells is suppressed by PKC (2, 10, 11). Whether IK(dr) of bronchial myocytes is similarly regulated by PKC is unknown.
Accordingly, in this study, we employed freshly isolated myocytes from bronchi of age-matched, control, and ragweed pollen-sensitized dogs and the standard whole cell patch-clamp technique to 1) characterize the properties of IK(dr) in cells isolated from a small-airway preparation, 2) determine whether the magnitude or properties of 4-AP-sensitive IK(dr) were altered in myocytes from bronchial tissues from ragweed pollen-sensitized dogs, and 3) assess the effect on the magnitude and properties of IK(dr) of activation of PKC.
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METHODS |
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Animal sensitization. Dogs were housed, sensitized, and then killed with a lethal injection of pentobarbital sodium (30 mg/kg body wt iv) according to a research protocol consistent with the Canadian Council on Animal Care and approved by local animal care committees of the Medical Research Council of Canada. Newborn mongrel dogs were immunized within 24 h of birth by intraperitoneal injection of 500 µg of ragweed (Ambrosia elatior) pollen extract as previously described (24). Booster injections were given on a weekly basis for 8 wk and then at a biweekly interval for an additional 8 wk. Age-matched, littermate control dogs received a similar schedule of intraperitoneal injections with adjuvant alone. Sensitization to ragweed pollen extract was confirmed by homologous passive cutaneous anaphylaxis in normal adult dogs. For the experiments reported here, bronchial tissues were obtained from sensitized dogs with a serum immunoglobulin E anti-ragweed antibody titer of equal to or greater than 256 dilutions. We employed tissues from 9 sensitized and 10 control littermate dogs from five different litters. Tissues from three adult mongrel dogs were also employed in some of the preliminary experiments to characterize the outward K+ current components of isolated bronchial myocytes and to define the recording conditions appropriate for the study of IK(dr) of bronchial myocytes with minimal contamination from other conductances.
Cell isolation. Canine lungs were quickly excised and placed in ice-cold Krebs-Henseleit physiological salt solution (see Solutions for composition). The bronchial trees were dissected free from the lung parenchyma and then placed in ice-cold nominally Ca2+-free Krebs-Henseleit solution. Bronchial branches of fourth- to sixth-order airways (which represent the airways involved in the hyperreactive response) were selected according to the method of Shioya et al. (37). The smooth muscle layer was dissected free of adherent epithelial cells and adventitia under a dissection microscope using fine iris scissors, placed in fresh ice-cold nominally Ca2+-free Krebs-Henseleit solution, and stored overnight until tissue digestion the next day. Results of preliminary experiments, performed using myocytes dispersed from tissues on the same day as they were obtained, were not different from those in which the tissues were stored overnight. Freshly dispersed, relaxed single myocytes were prepared from tissue pieces of ~3 mm2 based on a method previously described (30). Briefly, bronchial smooth muscle tissues were placed in a low-Ca2+ physiological salt solution containing collagenase, protease, and elastase (for composition see below) and gently bubbled with 95% O2-5% CO2 at 33-35°C for 20-35 min. After removal from the digestion solution, the pieces were washed several times in low-Ca2+ solution and stored at 4°C until required (within 5 h). Myocytes were liberated from the digested tissue pieces by gentle trituration. Viable myocytes used in the patch-clamp experiments were spindle shaped, optically refractive, and relaxed, as is apparent for the representative myocyte shown in Fig. 1.
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Solutions. The Krebs-Henseleit solution for dissection and overnight storage of intact bronchial tissues contained (in mM) 115 NaCl, 25 NaHCO3, 1.38 KH2PO4, 2.51 KCl, 2.46 MgCl2, 1.91 CaCl2, and 5.56 dextrose (pH 7.4 when gassed with 95% O2-5% CO2) with the exception that the storage solution lacked the added CaCl2. The low-Ca2+ physiological salt solution used in cell isolation contained (in mM) 120 NaCl, 25 NaHCO3, 4.2 KCl, 0.6 KH2PO4, 1.2 MgCl2, and 0.01 CaCl2 (pH 7.4 when bubbled with 95% O2-5% CO2 at room temperature, 20-22°C), supplemented with collagenase (564 U/ml Sigma type 1A; Sigma Chemical), protease (10 U/ml Sigma type XXVII), and elastase (81 U/ml Sigma type IV).
The standard bath solution employed in the whole cell voltage-clamp
experiments contained the following (in mM): 120 NaCl, 3 NaHCO3, 4.2 KCl, 1.2 KH2PO4,
0.5 MgCl2, 10 glucose, 1.8 CaCl2, and 10 N-2-hydroxyethylpiperazine-N'-2-ethanesulfonic
acid (HEPES; pH 7.4 corrected with NaOH). The standard whole cell
pipette solution contained the following (in mM): 110 potassium
gluconate, 30 KCl, 0.5 MgCl2, 5 HEPES, 5 Na2ATP, and either 0.05 ethylene glycol-bis(-aminoethyl ether)-N,N,N',N'-tetraacetic
acid (EGTA) or 10 1,2-bis(2-aminophenoxy)ethane-N,N,N',N'-tetraacetic acid (BAPTA), as stated in RESULTS, to produce
varied levels of internal Ca2+
buffering (pH adjusted to 7.2 with KOH).
Electrophysiological recordings.
Isolated myocytes were placed in a 300-µl bath and allowed to settle
to the bottom of the chamber before a constant flow of bath solution
(~0.5 ml/min) at room temperature (20-22°C) was initiated. A
standard whole cell patch-clamp recording technique was employed in the
experiments (16). Pipettes were prepared from capillary glass (7052 glass; Richland Glass) using a Sutter P-87 puller (Sutter Instruments) and MF-83 microforge (Narashige Scientific Instrument
Laboratory). Tip resistances of 1-3 M were obtained when filled
with pipette solution. Recordings were performed using either an
Axopatch 1-D or 200A amplifier (Axon Instruments). Pipette potential
and capacitance were nulled and a 5- to 15-G
seal formed with the
membrane. To correct for junction potential, 20 pipettes were nulled in
pipette solution and then transferred to bath solution. A consistent
value of 10 mV for the junction potential was obtained and employed to
correct all voltage-clamp protocols and current recordings. Whole cell
voltage-clamp protocols were applied using pClamp software (Axon
Instruments). Data were filtered at 2 kHz by an on-board 8-pole Bessel
filter before digitization with a Labmaster TL-1-125 or Digidata
1200 analog-to-digital converter (Axon Instruments) and
stored on hard disk in a 486 PC clone. Data were displayed and analyzed
off-line using pClamp (Axon Instruments) and Origin software (MicroCal
Software).
Values for net and tail currents due to
IK(dr) were
normalized for cell capacitance and expressed as picoamperes per
picofarad. Cell capacitance was determined by integration of the
capacity transient evoked by 20-mV command steps. Current-voltage
(I-V) relations for net and tail
currents were determined in the following manner: net current due to
IK(dr), with
minimal contamination by other conductances, was measured at the end of
250-ms pulses and plotted as a function of command step voltage between
80 and +30 mV. Tail currents evoked by repolarization to
40 mV following steps to between
80 and +30 mV and due to
deactivating Kdr exclusively were
calculated as the difference between the peak amplitude of the tail and
sustained current levels at 200 ms. Drug-sensitive difference currents
were determined by digital subtraction of current traces obtained in
the presence of drug from those recorded in its absence.
Drugs. 4-Phorbol 12,13-dibutyrate
(PDBu), Na2ATP, EGTA, BAPTA,
tetraethylammonium chloride
(TEA+), and 4-AP were obtained
from Sigma Chemical; 1,2- and 1,3-isomers of
dioctanoyl-sn-glycerol
(diC8) were obtained from
Serdary Research Laboratories. A stock solution of PDBu (10 mM) was
prepared in dimethyl sulfoxide. Diacylglycerol (DAG) analogs were
dissolved in hexane. All were added directly to the bath solution at
the desired concentration on the day of use. Vehicle controls had no
effect on currents recorded. 4-AP was dissolved in bath solution, and
the pH was readjusted to 7.4.
Statistics. All values in the text and Figs. 1-9 are presented as means ± SE. Data were compared by paired or unpaired Student's t-test for single comparisons. Individual data points in the I-V relations for myocytes exposed to single treatments were compared with a paired Student's t-test, and in experiments where multiple treatments were applied to single cells, the data points were compared by repeated-measures ANOVA followed by Bonferroni's test for multiple comparisons. A level of P < 0.05 was considered to be statistically significant.
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RESULTS |
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Morphological studies. Bronchial myocytes isolated from control and sensitized dogs and employed in the whole cell patch-clamp studies were identical to the representative cell shown in Fig. 1. Myocytes from control dogs had an average length of 228 ± 36 µm (range 155-307 µm, n = 59 from three separate cell isolations). There was no difference in the capacitive cell surface area of myocytes from control and sensitized dogs: values of 4,210 ± 290 (n = 16) and 3,710 ± 338 (n = 13) µm2 (P > 0.05) were obtained based on a specific membrane capacitance of 1 µF/cm2 and average cell capacitances of 42.1 ± 2.9 and 37.1 ± 3.4 pF (P > 0.05), respectively. The cells were relaxed on isolation and were capable of repeated contractions to elevated external K+ or histamine (data not shown).
Identification of macroscopic
K+ currents.
The identity and properties of macroscopic
K+ currents of canine bronchial
myocytes have not been characterized in detail. For this reason, we
conducted a series of preliminary experiments to identify conditions
that would permit selective recordings of
IK(dr). These
initial experiments demonstrated the presence of
1) inward L-type
Ca2+ currents that showed rundown
within the first 2-3 min of recording and
2) two dominant components of
macroscopic outward K+ current in
bronchial myocytes of adult and young control or ragweed pollen-sensitized dogs. In the presence of minimal intracellular Ca2+ chelation with 0.05 mM EGTA
in the pipette solution, command steps from a holding potential of
60 mV to between
80 and +30 mV every 10 s evoked net
outward K+ current positive to
40 mV, which displayed voltage- and time-dependent activation
and was "noisy" positive to 0 mV (Fig.
2A).
Outward current evoked during voltage steps increased exponentially
with applied voltage and on average reached a density of 58 ± 4 pA/pF at +30 mV (Fig. 2B).
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Characteristics of
IK(dr) in bronchial
myocytes from control dogs.
The kinetics and voltage dependence of activation, deactivation, and
inactivation of bronchial smooth muscle
IK(dr) were
determined. Activation during a 250-ms test pulse to +20 mV and
subsequent deactivation at 40 mV were both best fit with two
exponentials (Table 1). Slow
inactivation of
IK(dr) was
apparent during command pulses to potentials positive to 0 mV.
Longer-duration, 4- to 10-s steps to +20 mV were applied to bronchial
myocytes to examine the time dependence of inactivation.
IK(dr) decayed
relatively rapidly during the first 4 s of the command pulses and
thereafter exhibited only a very slow decline in amplitude. A
biexponential function was found to best describe the inactivation
kinetics, and, at +20 mV, the average values for the fast and slow time constants (Table 1) were similar to those previously reported for other
smooth muscle cells (e.g., 0.14 and 1.1 s at +5 mV for canine tracheal
myocytes; see Ref. 7). The voltage dependence of
IK(dr) activation
and inactivation was studied. Figure 5
illustrates representative examples of
1) tail currents recorded at
40 mV after steps to between
80 and +30 mV (Fig.
5A) and
2) whole cell currents evoked by a
double-pulse protocol that employed 4-s conditioning pulses to between
110 and +20 mV followed by command steps of 200-ms duration to
+20 mV (Fig. 5B). Figure
5C shows the average values for
normalized peak tail current versus voltage and normalized IK(dr)
availability as a function of command and conditioning step voltages,
respectively. Peak tail current amplitude at
40 mV
after command steps to between
80 and +30 mV was measured (i.e.,
peak minus sustained amplitude after 200 ms), normalized to the maximal
amplitude, and plotted as a function of command step voltage.
Availability after a 4-s conditioning pulse was determined from the
amplitude of current present at the end of 200-ms pulses to +20 mV and
plotted as a function of conditioning pulse voltage. The plots of
normalized current versus voltage of the command step or conditioning
pulse were best fit with single Boltzmann functions (Fig. 5). Values
for half-maximal potential (V0.5) and
slope for activation and inactivation curves from 15 and 9 myocytes,
respectively, were averaged and are given in Table 1. Similar values
for these parameters were also obtained in two myocytes using 10-s
conditioning pulses (not shown).
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Comparison of IK(dr) in bronchial myocytes from control and sensitized dogs. Whether the properties of IK(dr) were altered in myocytes obtained from ragweed pollen-sensitized dogs was determined by applying similar voltage-clamp protocols under identical recording conditions to those described for cells from control dogs. Representative families of IK(dr) of bronchial myocytes of control and ragweed pollen-sensitized dogs and average I-V relations for end-pulse and tail current amplitude of 15 and 13 control and sensitized myocytes, respectively, are shown in Fig. 6. Plots of average normalized current versus voltage for activation and inactivation of IK(dr) in myocytes from sensitized dogs are shown in Fig. 5. Average values for time constants of activation, deactivation, and inactivation, as well as the voltages for half-maximal activation and inactivation, and values for slope factors are given in Table 1. No statistically significant difference was noted for 1) end-pulse and tail current densities (Fig. 6), 2) voltage dependence of activation (Fig. 5), or 3) kinetics of activation, inactivation, or deactivation (Table 1) of IK(dr) between myocytes isolated from the bronchi of sensitized versus control dogs.
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Inhibition of IK(dr) by
PKC activation.
Activation of PKC is thought to play an important role in the signal
transduction pathway of histamine, which contracts airway smooth
muscle, and alterations in the activity of PKC isoenzymes have been
implicated in the pathogenesis of asthma and bronchospasm of
hyperresponsive airways of sensitized animals (4, 17, 35). Accordingly,
we determined the effects on
IK(dr) of direct PKC activation with an analog of DAG,
1,2-diC8, and the phorbol ester
PDBu. Eleven myocytes from control and sensitized dogs were used (6 and
5 myocytes, respectively), and a similar change in IK(dr) during PKC
activation was observed in all cells tested. Figure
7 shows the effects of the inactive DAG
analog 1,3-diC8 followed by
1,2-diC8 on
IK(dr) amplitude.
To monitor the time course of effects of the DAG analogs on
IK(dr) amplitude,
depolarizing steps to +10 mV followed by a brief step to 40 mV
were applied every 15 s. Figure 7A
shows a representative example of the change in normalized end-pulse
current amplitude under control conditions and during sequential
exposure to the inactive DAG analog
1,3-diC8 (10 µM) followed by
active DAG analog 1,2-diC8 (1 and
10 µM). The representative current recordings shown in Fig. 7,
insets, were obtained at the four time
points indicated by the arrows. No effect on
IK(dr) amplitude
was evident with 1,3-diC8, but
end-pulse amplitude slowly declined in a concentration-dependent
fashion upon exposure to 1,2-diC8.
Figure 7B shows superimposed,
representative difference current traces obtained by digital
subtraction of traces recorded at 30-s intervals during exposure to
1,2-diC8. The onset of inhibition
of IK(dr)
required ~60 s, so the initial difference currents showed no change.
However, the amplitude of the outward 1,2-diC8-sensitive difference
current increased with time, reaching a peak sustained level within
3-5 min. The difference currents at peak inhibition by
1,2-diC8 were very similar to
those determined for 4-AP inhibition (Fig. 4).
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DISCUSSION |
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In this study, the whole cell voltage-clamp technique was used to characterize the properties of IK(dr), as well as to determine the effect of ragweed pollen sensitization and direct activation of PKC with a DAG analog or phorbol ester on this conductance of isolated canine bronchial myocytes. We show for the first time that 1) the voltage dependence of inactivation for IK(dr) in bronchial myocytes is different from that in the trachea, 2) inactivation is shifted to more negative potentials in myocytes from sensitized compared with age-matched control dogs, and 3) PKC activation reduces IK(dr) amplitude. Reduced IK(dr) due to decreased availability and/or inhibition by PKC activation may have implications for altered control of airway smooth muscle tone in asthma.
Two K+ channels,
BKCa and
Kdr, were shown to contribute to
macroscopic currents of canine bronchial myocytes on the basis of pharmacology, Ca2+ sensitivity,
and kinetic behavior. Noisy macroscopic current carried by
BKCa was present positive to
20 mV with minimal intracellular Ca2+ chelation and was selectively
inhibited with 0.1 mM TEA+. In
contrast, macroscopic current due to
Kdr was insensitive to low
TEA+ and high levels of
intracellular Ca2+ chelation, but
it was readily blocked by 4-AP and displayed slow, time-dependent
inactivation. The presence of these two channel types in bronchial
myocytes is consistent with previous studies of airway myocytes of dog,
ferret, pig (7, 26, 29), and humans (1, 18, 41).
IK(dr) of canine
bronchial myocytes was studied using 10 mM BAPTA in the pipette to
provide for substantial chelation of intracellular
Ca2+ and minimal contamination of
whole cell currents by
Ca2+-activated conductances, such
as BKCa,
Ca2+-activated
Cl
, and
Ca2+-dependent nonselective cation
currents. Additionally, depolarizing voltage-clamp steps were limited
to potentials of +30 mV or less to avoid voltage-dependent activation
of BKCa. The ability of 4-AP to
substantially inhibit tail currents produced during deactivation of
Kdr at
40 mV as well as
end-pulse current amplitude is consistent with minimal contamination by
non-Kdr channel activity. The
presence of
IK(dr) in all
canine bronchial myocytes studied here differs from that reported for
human myocytes in which the current was only present in 17% of cells
employed (41). However, Janssen (18) described
IK(dr) in human
bronchial smooth muscle that was sensitive to 4-AP, and Adda et al. (1)
described the presence of mRNA encoding the voltage-gated
K+ channels Kv1.1, Kv1.2, and
Kv1.5 in human airway muscle.
The fittings of the kinetics of inactivation and deactivation are
consistent with other smooth muscle preparations, both being best
described by biexponential functions with two time constants on the
order of 0.2-0.5 and 2-5 s (inactivation) and 10 and 60 ms
(deactivation; see Refs. 2, 3, 7, 26, 28, 40, 46). However, few studies
have quoted values for activation time constants due to the overlap
with activation of transient outward currents. We did not observe
evidence in any cell for the presence of "A"-type outward current
(note the transient current in the presence of
1,2-diC8 resulted from the change
in IK(dr); see
Refs. 2 and 10) indicating its absence or complete inactivation at the
holding potential (60 mV) employed. A dual-exponential fit of
IK(dr) activation
in bronchial myocytes is consistent with that of cloned rabbit vascular
Kv1.5 (O. Clément-Chomienne, K. Ishii, M. Walsh, and W. Cole,
unpublished observation). This study shows that
IK(dr) of canine
bronchial myocytes is activated at voltages positive to approximately
45 mV and that it displays time- and voltage-dependent
inactivation. The overlap of voltage ranges for activation and
inactivation suggests that steady-state Kdr window current may be present
between approximately
45 and 0 mV, with peak outward
IK(dr) at
approximately
25 mV. These properties are similar to those
previously reported for
IK(dr) of ferret
tracheal myocytes; Kdr window
current was observed between approximately
50 and
10 mV
at 37°C (15). This more negative activation voltage of
50 mV
compared with the present study is likely a function of the bath
temperature used to record the ferret tracheal
IK(dr);
activation, but not inactivation, of
IK(dr) of smooth
muscle cells is known to occur over a more negative range at higher
temperatures (e.g., see Ref. 10). It is significant that the voltage
for half-maximal availability of
IK(dr) of
bronchial myocytes was less negative compared with the value reported
for canine and porcine tracheal cells:
26 mV versus
46
and
53 mV for whole cell and for single
Kdr channel currents in the
trachea (7). It would seem unlikely that differences in recording
conditions in the two studies can explain the divergent values for
V0.5 of inactivation (only slightly different pipette solutions were employed, and both were done at room temperature). Moreover, under identical recording conditions (temperature, bath and pipette solutions, protocol) to those employed in this study,
V0.5 for
inactivation of
IK(dr) of rabbit
portal vein myocytes was also more negative (approximately
40
mV) than that observed in the present study, even though the
V0.5 for
activation was similar: approximately
16 and
11 mV in
bronchial and portal vein myocytes, respectively (2, 10). The
physiological implication of the less negative voltage dependence of
inactivation is that
IK(dr) of
bronchial myocytes will contribute steady-state current over a wider
range of membrane potentials and, therefore, its importance to control of membrane potential and contractile tone may be greater than in the
trachea. The role of
IK(dr) was
similarly concluded to be enhanced in small versus large pulmonary
arterial vessels (6). In this case, however, smaller vessels were found
to possess more cells with
IK(dr) (6).
We also found that the availability of
IK(dr)
between 50 and
20 mV was decreased in tissues
from sensitized compared with control dogs because of a negative shift
in voltage dependence of inactivation. Although small in magnitude,
this hyperpolarizing shift in availability of
IK(dr) is of
potential significance to the electrical behavior of the
hyperresponsive airways of ragweed pollen-sensitized dogs. The shift in
the inactivation curve predicts that the amplitude of steady-state
IK(dr) will be
reduced over the critical range of membrane potentials between
50 and
20 mV over which the L-type
Ca2+ window current also occurs
(13, 14). Such small changes in current amplitude will have large
effects on membrane potential in cells with a large input resistance,
as is the case for smooth muscle cells (1-15 G
; see
Ref. 31).
It is possible that the difference in
V0.5 for
inactivation between tracheal and bronchial myocytes, as well as the
negative shift in availability of
Kdr channels in the myocytes of
sensitized dogs, may arise from the expression of different
Kdr channels in the two tissues.
Voltage-gated K+ channels consist
of pore-forming - and modulatory
-subunits. The voltage
dependence of inactivation of voltage-gated
K+ channels is known to be
affected by association with some
-subunits. For example, Kv1.5,
which is the dominant Kv present in human bronchi (1), displays a more
negative voltage dependence of inactivation but no change in kinetics
when expressed in the presence of Kv
2.1 (45). Alternatively, the
formation of heterotetramers between different
-subunits of the Kv1
family, which possess distinct inactivation properties, can give rise
to channels with novel, intermediate properties, including differences
in voltage dependence of inactivation (32). Therefore, differences in
the expression of a
-subunit and/or increased formation of
heteromultimers of Kv1.5 and Kv1.1, which are expressed in human
airways (1), could be the cellular basis for the difference in
inactivation of
IK(dr) in
myocytes from trachea and bronchi, as well as the negative shift in
availability of
IK(dr) in
myocytes of sensitized compared with control dogs. Further studies will
be required to address these issues.
In this study, we employed two structurally different analogs of DAG
and a phorbol ester to determine the effect of activation of PKC on
IK(dr) of
bronchial smooth muscle. The active DAG analog 1,2-diC8 and the phorbol ester
PDBu were found to cause a decrease in the amplitude of
IK(dr), but the
inactive analog 1,3-diC8 was without effect. This indicates that nonspecific effects of
1,2-diC8, for example, changes in
membrane fluidity or direct channel block, were not involved. The
activity of BKCa, L-type
Ca2+, ATP-sensitive
K+, and nonselective cation
channels in smooth muscle were previously reported to be affected by
PKC activation (see Ref. 27 for a review). A change in these
conductances also cannot account for the present data for the following
reasons: 1) the effects of PKC
activation on Kdr were determined
under conditions that minimized the contribution of
BKCa and
Ca2+-activated
Cl current to macroscopic
current (high intracellular Ca2+
chelation with 10 mM BAPTA and command potentials negative to +30 mV);
2)
1,2-diC8 had no effect on
time-independent current under the recording conditions employed,
suggesting that changes in ATP-sensitive
K+ and nonselective cation
currents were not involved; and 3)
activation of PKC was associated with a decline in
Kdr tail current amplitude at
40 mV where little contamination of macroscopic current by Ca2+ current would be expected.
The ability of PKC activation to suppress
IK(dr) of
bronchial myocytes is consistent with our previous observations on
rabbit portal vein (2, 10). A family of at least 11 related PKC isoforms possessing different properties has been characterized, and at
least five isoforms are known to be expressed in smooth muscle tissues
(9, 17). The effects of 1,2-diC8
and PDBu on
IK(dr) of
bronchial myocytes can be attributed to PKC,
, or
for the
following reasons. PKC
I,
II,
,
,
, and
were identified in immunoblots using isoform-specific antibodies from canine
bronchial preparations (12). The low intracellular
Ca2+ level under our recording
conditions (due to dialysis with pipette solution containing 10 mM
BAPTA) likely precludes a role for the Ca2+-dependent isoforms in the
suppression of
IK(dr) in our
experiments, and the
Ca2+-independent isoform PKC
is
insensitive to DAG and phorbol esters (9). In rabbit portal vein, the
effect of PKC on
IK(dr) was attributed to PKC
(10). Inhibition of
IK(dr) due to
activation of PKC may contribute to dysfunctional control of membrane
potential in asthma. This view is based on the following.
1) Histamine release is elevated in
bronchi of sensitized animals (8).
2) Histamine and other contractile
agonists are known to stimulate DAG production, leading to PKC
activation in airway smooth muscle (35).
3) Activation of PKC causes a tonic
increase in tone in human airway tissue; 100 nM PDBu produces a
sustained contraction that is ~40% of the maximal response to
acetylcholine (34). A role for changes in membrane potential and
activation of voltage-gated Ca2+
channels is suggested by the fact that phorbol ester-induced contractions are dependent on extracellular
Ca2+ influx and inhibited by
treatment with dihydropyridines (34). 4) PKC activation has been
implicated in the pathogenesis of airway smooth muscle hyperreactivity
(33, 35). In light of the ability of DAG analog and phorbol ester to
depress IK(dr),
the lack of a difference in current density in bronchial myocytes of
sensitized versus control dogs argues against an intrinsic difference
in basal PKC activity in the two groups. However, as noted above, the
bronchi of the sensitized dog are known to release more histamine in
response to antigen challenge compared with control animals (8). This
difference in histamine release may be expected to produce
1) an extrinsic change in PKC
activity and 2) an enhanced depression of
IK(dr) in situ
that would not be observed in the isolated myocytes in the absence of
antigen, mast cells, and/or histamine.
The changes in
IK(dr) of
bronchial myocytes associated with ragweed pollen sensitization or PKC
activation identified in this study may be two contributing factors to
the mechanism(s) responsible for dysfunctional airway smooth muscle
contractility in asthma. This view is based on compelling evidence that
Kdr contributes to the control of
membrane potential and the level of depolarization of smooth muscle
during exposure to contractile or relaxant stimuli (1, 10, 13, 25, 28,
31, 38). Previous studies on intact airways from ferrets and humans
employing 4-AP and charybdotoxin to inhibit
Kdr and
BKCa, respectively, indicated that
IK(dr)
contributes to resting K+
conductance but that BKCa is not
as important under basal conditions (1, 15). Treatment with 4-AP (in
the presence of tetrodotoxin and atropine) caused depolarization of
tracheal smooth muscle membrane potential by at least 5 mV and produced
tonic contraction, the latter being sensitive to inhibition of L-type
Ca2+ channels with dihydropyrines
(1, 15, 23). Tonic contractions due to steady-state
Ca2+ influx in airway smooth
muscle are possible because of the presence of a small L-type
Ca2+ window current, over the same
voltage range as that of Kdr
window current, which is capable of producing marked increases in
intracellular Ca2+ concentration
(13). IK(dr)
activation has been postulated to function as a voltage-dependent,
feedback mechanism for control of
Ca2+ influx in smooth muscle (11,
31); depolarization due to activation of inward currents in bronchial
myocytes carried by T- or L-type Ca2+,
Cl, and/or
nonselective cation channels (13, 19, 20, 21) is offset by the
activation of outward K+ current.
In cerebral resistance arteries, inhibition of
IK(dr) with 4-AP
causes enhanced myogenic reactivity to increased intraluminal pressure
(25). Kdr may function in parallel
with a Ca2+-dependent feedback
mechanism involving BKCa
activation in response to changes in intracellular
Ca2+ concentration (31). Precise
changes in the amplitude of outward current via these voltage- and
Ca2+-dependent feedback mechanisms
would prevent regenerative Ca2+
influx and permit the graded depolarizations and tone development observed in bronchi. Further studies are required to determine whether
alterations in BKCa channels also
contribute to airway hyperresponsiveness.
This study identifies two different mechanisms by which the
contribution of Kdr to control of
membrane potential may be reduced in airway smooth muscle in asthma:
1) a negative shift in voltage dependence of inactivation resulting in reduced steady-state
IK(dr) between
40 and
20 mV and 2)
inhibition by PKC activation. Because the level of membrane potential
is determined by a balance between inward and outward currents in
smooth muscle (31), a decline in steady-state
IK(dr) due to a
change in availability or suppression by PKC activation would result in
depolarization and may contribute to the reported shift in membrane
potential to more positive potentials in sensitized canine airway
tissues (42). A reduction in steady-state IK(dr) may also
contribute to the development of airway hyperreactivity that is
observed in asthma; contractile agonists are known to activate inward
conductances, for example, L-type
Ca2+ (44),
Cl
, and nonselective cation
currents (20, 21), and depolarize tracheal myocytes. A decline in the
voltage-dependent feedback control of membrane potential may therefore
result in enhanced depolarization, greater L-type
Ca2+ channel activation, and an
increased rise in intracellular
Ca2+ concentration during exposure
to contractile agonists. The net result would be greater development of
tone and increased peak force, which are typical of hyperreactive
airway tissues (4, 8, 33, 35). Further experiments on bronchial smooth
muscle are required to assess 1) the
role of IK(dr) in
depolarization of intact bronchial tissues to specific contractile
agonists and 2) whether activation
of phospholipase C-coupled receptors can produce a PKC-dependent
decline in Kdr channel activity in
bronchial myocytes.
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ACKNOWLEDGEMENTS |
---|
We thank Fabrice Chomienne for expert assistance in the isolation of single myocytes.
![]() |
FOOTNOTES |
---|
This study was supported by grants from the Medical Research Council of Canada (MT-13506 to W. C. Cole), Respiratory Health National Centres of Excellence (N. L. Stephens), and Icelandic University and Science Foundations (S. B. Sigurdsson). G. J. Waldron is a Fellow of the Canadian Hypertension Society and Medical Research Council of Canada. S. B. Sigurdsson and E. A. Aiello were Alberta Heritage Foundation for Medical Research Visiting Scientists. W. C. Cole is a Senior Scholar of the Alberta Heritage Foundation for Medical Research.
Address for reprint requests: W. C. Cole, Smooth Muscle Research Group, Univ. of Calgary, 3330 Hospital Dr., NW, Calgary, Alberta, Canada T2N 4N1.
Received 23 December 1997; accepted in final form 7 April 1998.
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