Abstract The membrane potential (Em) of pulmonary arterial smooth muscle cells (PASMCs) regulates pulmonary arterial tone by controlling voltage-gated Ca2+ channel activity, which is a major contributor to [Ca2+]i. The resting membrane is mainly permeable to K+; thus, the resting Em is controlled by K+ permeability through sarcolemmal K+ channels. At least three K+ currents, voltage-gated K+ (KV) currents, Ca2+-activated K+ (KCa) currents, and ATP-sensitive (KATP) currents, have been identified in PASMCs. In this study, both patch-clamp and quantitative fluorescent microscopy techniques were used to determine which kind(s) of K+ channels (KV, KCa, and/or KATP) is responsible for controlling Em and [Ca2+]i under resting conditions in rat PASMCs. When the bath solution contained 1.8 mmol/L Ca2+ and the pipette solution included 0.1 mmol/L EGTA, depolarizations (−40 to +80 mV) elicited both KCa and KV currents. Removal of extracellular Ca2+ and increase of intracellular EGTA concentration (to 10 mmol/L) eliminated the Ca2+ influx–dependent KCa current. 4-Aminopyridine (4-AP, 5 to 10 mmol/L) but not charybdotoxin (ChTX, 10 to 20 nmol/L) significantly reduced KV current under these conditions. In current-clamp experiments, 4-AP decreased Em (depolarization) and induced Ca2+-dependent action potentials; this depolarization increased [Ca2+]i in intact PASMCs. Neither ChTX nor the specific blocker of KATP channels, glibenclamide (2 to 10 μmol/L), caused membrane depolarization and the increase in [Ca2+]i. However, pretreatment of PASMCs with ChTX enhanced the 4-AP–induced increase in [Ca2+]i. These results suggest that the 4-AP–sensitive KV currents that are active in the resting state are the major contributors to regulation of Em and thus [Ca2+]i in rat PASMCs.
- voltage-gated K+ channels
- Ca2+-activated K+ channels
- ATP-sensitive K+ channels
- membrane potential
- intracellular Ca2+
Elevation of [Ca2+]i plays a primary role in triggering contraction in vascular smooth muscle cells, including PASMCs. Both Ca2+ influx through sarcolemmal Ca2+ channels and Ca2+ release from intracellular Ca2+ stores (eg, sarcoplasmic reticulum) contribute to the rise in [Ca2+]i. By controlling Ca2+ influx via voltage-gated Ca2+ channels, Em in vascular smooth muscle cells is an important regulator of vascular tone.1 2 The Ca2+ current through voltage-gated Ca2+ channels is proportional to the amount of time a Ca2+ channel spends in the open state, which is strongly regulated by Em. It has been proposed that the voltage dependence of Ca2+ channels underlies the Em dependence of vascular tone, with 3-mV depolarization increasing Ca2+ influx as much as twofold.1 2 In vascular smooth muscle cells, Em is controlled by K+ permeability through sarcolemmal K+ channels. Because of high membrane input resistance of 2.6 to 17 GΩ in resting vascular smooth muscle cells,3 4 5 a very small change in outward K+ currents should result in a marked change in Em.
At least four types of K+ channel current have been identified in vascular smooth muscle cells2 6 7 : (1) IK(V), which is composed of transient (A-type)6 8 9 and steady state (delayed rectifier) components,10 11 12 (2) IK(I),13 (3) IK(Ca),14 and (4) IK(ATP).3 15 Em appears to be controlled by an equilibrium between the outward currents provided by the K+ channels and the inward currents provided by Ca2+16 17 and Cl− channels.18 19 However, it has been proposed that under resting physiological conditions, K+ permeability through KV channels is responsible for determining the resting Em16 20 21 22 23 24 25 and depolarization-dependent repolarization,1 2 whereas KATP and KCa channels contribute to the hyperpolarization induced by various exogenous and endogenous vasodilators15 and the repolarization following increased [Ca2+]i,26 respectively. In vascular smooth muscle cells, the KATP and KCa channels, as targets of endogenous vasodilators and negative feedback regulators of vascular tone, link Em to the state of cellular metabolism and [Ca2+]i homeostasis.
Regulation of PA tone and contraction depends largely on resting Em and [Ca2+]i in PASMCs. Membrane ionic channels are major contributors in controlling Em and, subsequently, [Ca2+]i. Thus, the primary goal of the present study was to use the selective blockers of respective K+ channels to identify those K+ channels that are responsible for regulating Em and, more importantly, [Ca2+]i under resting physiological conditions. The data provide evidence that 4-AP–sensitive IK(V) is the major K+ current controlling resting Em and [Ca2+]i in the primary cultures of rat PASMCs.
Materials and Methods
Cell Preparation and Culture
Rat primary cultured (3 to 7 days) PASMCs were used for the present study. The methods used for isolation and culture of PASMCs are described elsewhere.5 Briefly, the right and left branches of the rat main PA, with some intrapulmonary arterial branches, were aseptically removed from Sprague-Dawley rats (125 to 250 g) and placed in 1 mL Hanks’ balanced salt solution (Sigma Chemical Co) containing 1.5 mg collagenase (Worthington Biochemical Co) for 20 minutes at 37°C. The adventitia was then dissected free of the vessel, and the endothelium was carefully rubbed off with forceps. Single PASMCs were obtained by digesting the resultant PA smooth muscle tissue with 1.5 mg/mL collagenase, 0.5 mg/mL elastase (Sigma), and 1 mg/mL bovine serum albumin (Sigma). Cells were incubated in a humidified atmosphere of 5% CO2 in air at 37°C for 3 to 7 days after being plated onto coverslips and fed twice weekly with 10% FBSCM (Irvine Scientific). Twelve to 24 hours before experiments, the bovine serum concentration in FBSCM was decreased to 0.3% in order to stop cell proliferation.
Recording of Membrane Currents and Em in Single PASMCs
The patch-clamp technique was used to measure membrane currents and Em under voltage-clamp or current-clamp (current=0) mode, respectively, in the whole-cell configuration.5 27 Pipettes were pulled from borosilicate glass capillaries (VWR Scientific) with a vertical pipette puller (PP-83, Narishige) and fire polished with a Narishige microforge. The pipettes used for recording whole-cell currents and resting Em had resistances of 2 to 4 MΩ when filled with the regular pipette solution (≈134 mmol/L KCl). Coverslips containing the cells were placed in the recording chamber (volume, ≈0.75 mL) and superfused with external (bath) solution at a rate of 1.3 mL/min. Voltage-clamp command potentials were generated by using an Axopatch-1D patch-clamp amplifier (Axon Instruments) under the control of pclamp software. The interpulse interval in the voltage-clamp experiments was 10 to 25 seconds. Data acquisition was carried out with a digital interface (TL-1 DMA, Axon Instruments) coupled to an IBM-compatible computer. Membrane current and voltage were monitored on a storage oscilloscope (Tektronix). Whole-cell currents were filtered at 2 kHz (−3 dB), digitized at 1 to 2 kHz, and stored on hard disk of the computer for off-line analysis. Series resistance and whole-cell capacitance were compensated (60% to 70%) by adjusting the internal circuitry of the patch-clamp amplifier. The leakage currents were subtracted by using P/4 protocol in pclamp software. All electrophysiological experiments were performed at room temperature (22°C to 24°C).
Measurement of [Ca2+]i in Single PASMCs
The [Ca2+]i in PASMCs was measured by using the fluorescent dye fura 2 and a quantitative fluorescent microscopy system (Photoscan M-series, Photon Technology International).28 The cells grown on 25-mm coverslips were incubated in 3 mL 10% FBSCM containing 5 μmol/L of the acetoxymethyl ester form of fura 2 (fura 2-AM) for 30 minutes at room temperature under an atmosphere of 5% CO2/95% air. The fura 2–loaded cells on coverslips were then superfused (at a rate of 2.2 mL/min) with the standard bath solution (PSS) for 30 minutes at 32°C. This allows sufficient time to wash away any extracellular fura 2-AM and for intracellular esterase to cleave cytosolic fura 2-AM into the active fura 2.
A single cell of interest was identified and illuminated with light from a 75-W xenon lamp. Filtered emitted light at wavelengths of <550 nm was passed to a rotating chopper disk, allowing light to pass alternately through two 10-nm bandpass filters centered at 340 and 380 nm (Omega Optical). Excitation light was reflected by a dichroic mirror and focused onto the cell being studied with a ×40 Nikon UV-Fluor objective. Emitted light was low pass–filtered (510 nm) and passed through an aperture positioned over the cell. Fura 2 fluorescence (510-nm light emission excited by 340- and 380-nm illumination) from this area of the cell and background fluorescence were measured by using a photomultiplier tube via an Olympus IMT2 microscope equipped for epifluorescence microscopy. The fluorescence signals were collected continuously and stored in a 386 IBM-compatible computer (Everex Computer System) for later analysis.
[Ca2+]i was calculated from the ratio (R) of measured 510-nm fluorescence signals (F) elicited at 340 nm and 380 nm, according to the following equations: F340, cell and F380, cell indicate the measured fluorescence from the cell when illuminated at 340 nm and 380 nm, respectively; F340, bkg and F380, bkg, the background fluorescence signals, respectively; Kd (225 nmol/L), the dissociation constant of the Ca2+:fura 2 complex; Sf2, 340 fluorescence of the Ca2+-free fura 2; Sb2, 380 fluorescence of the Ca2+-saturated fura 2; and Rmin and Rmax, the ratio (F340/F380) of the fluorescence signals of fura 2 that is free of Ca2+ and the ratio of the fluorescence signals of the Ca2+:fura 2 complex at the highest [Ca2+], respectively. Rmax and Rmin were determined by using methods described elsewhere.29 30
Reagents and Solutions
The standard extracellular PSS used for recording IK and Em and for measuring [Ca2+]i contained (mmol/L): NaCl 141, KCl 4.7, CaCl2 1.8, MgCl2 1.2, HEPES 10, and glucose 10, buffered to pH 7.4 with 5 mol/L NaOH. In Ca2+-free PSS, the CaCl2 was replaced by MgCl2, and 0.5 to 1.0 mmol/L EGTA was added. The perfusion speeds are 1.3 and 2.2 mL/min in the patch-clamp setup and the quantitative fluorescent microscopy system. That is, the perfusion speed used for measuring [Ca2+]i is ≈1.69 (2.2/1.3) times faster than that used for measuring Em. The internal (pipette) solution used for recording IK and Em consisted of the following (mmol/L): KCl 125, MgCl2 4, HEPES 10, EGTA 10, and Na2ATP 5, buffered to pH 7.2 with 1 mol/L KOH. Usually, ≈90 μL KOH (1 mol/L) was needed to buffer the 10-mL internal pipette solution (original pH, ≈4.85) to pH 7.2. Thus, actual [K+]i was slightly increased from 125 to 134 mmol/L. This increase in [K+]i would slightly shift the K+ equilibrium potential by ≈2 mV (the calculated K+ equilibrium potential was shifted from −84.0 to −85.8 mV). In some experiments, the EGTA concentration in the Ca2+-free pipette solution was reduced from 10 to 0.1 mmol/L in order to diminish intracellular Ca2+ chelation.
4-AP (Aldrich Chemical Co) was dissolved directly into PSS on the day of use. The solution containing 4-AP was buffered to pH 7.4 by using HCl before each experiment. ChTX (Accurate Chemical & Scientific Co) was dissolved in DMSO (Sigma) and diluted to a final concentration of 10 to 20 nmol/L in the bath solution. Glibenclamide (Sigma) was dissolved in DMSO to make a stock solution of 100 mmol/L; an aliquot of the stock solution was diluted 1:50 000 to 1:10 000 into PSS to make a final concentration of 2 and 10 μmol/L, respectively. Similar dilutions of DMSO alone into PSS were used as controls and had no effect on K+ currents, Em, or [Ca2+]i.
Data are expressed as mean±SEM. Statistical analysis was performed by using paired or unpaired Student’s t test or ANOVA, as appropriate. Differences were considered to be significant at P<.05.
Whole-Cell Currents in PASMCs
When the cells were superfused with 1.8 mmol/L Ca2+–containing PSS and dialyzed with the pipette solutions containing 0.1 mmol/L EGTA, several components of whole-cell currents were recorded by depolarizing cells from a holding potential of −70 mV to a series of test potentials ranging from −40 to +80 mV (Fig 1⇓). There were two components of IK as well as ICa and ICl(Ca). ICa and ICl(Ca) were eliminated by removal of extracellular Ca2+ (Fig 1⇓, center); ICl(Ca) was inhibited by the Cl− channel blocker18 19 niflumic acid (10 μmol/L) (data not shown).
Two components of IK, IK(V) and IK(Ca), could be differentiated by the dependence on extracellular Ca2+ (Fig 1⇑). The Ca2+ influx–dependent IK(Ca) component was evidenced by the hump in an N-shaped I-V relation (I-V curve) between 0 and +60 mV,31 which could only be obtained when 1.8 mmol/L Ca2+ was present in the bath solution (compare Figs 1⇑ and 2⇓). Removal of extracellular Ca2+ and increase of intracellular EGTA concentration (from 0.1 to 10 mmol/L) virtually eliminated the Ca2+-dependent component, IK(Ca),32 of the total IK (Figs 1⇑ and 2⇓). This suggests that the KCa (but not KV) channel activity depends greatly on availability of free [Ca2+]i in the resting PASMCs.
The voltage-gated Ca2+-independent IK, IK(V), could then be isolated by using the Ca2+-free bath solution containing 0.5 to 1 mmol/L EGTA and the Ca2+-free pipette solution containing 10 mmol/L EGTA and 5 mmol/L ATP.32 During a maintained depolarization (eg, +80 mV) under these conditions, IK(V) rose to an early peak and then inactivated when the cell was superfused with Ca2+-free PSS (0.5 mmol/L EGTA present) and dialyzed with 10 mmol/L EGTA Ca2+-free pipette solution (Fig 2⇑). Two exponentials were required to best fit the decay of IK(V) (Fig 2⇑): a fast component with a time constant of 30 milliseconds and a slow component with a time constant of 434 milliseconds at +80 mV in this cell. This suggests that IK(V) is generated by K+ efflux through either of two types of KV channel, eg, A-type6 8 9 and delayed rectifier11 12 32 K+ channels, or one type of KV channel that has multiple states.32
Effects of ChTX and 4-AP on IK(V) in PASMCs
When cells were dialyzed with Ca2+-free pipette solution (plus 10 mmol/L EGTA) and superfused with Ca2+-free PSS containing 0.5 mmol/L EGTA, ChTX (10 nmol/L) negligibly affected IK elicited by test potentials from −40 to +80 mV (Fig 3A⇓ and 3C⇓). The ChTX-insensitive IK, however, was significantly decreased by 10 mmol/L 4-AP (Fig 3B⇓ and 3D⇓); this indeed indicated that this current is the voltage-dependent Ca2+-insensitive IK, termed IK(V).22 32 Averaged data shown in Fig 3C⇓ and 3D⇓ demonstrated that ChTX (10 nmol/L) negligibly affected IK(V), which was elicited by depolarizing cells from −70 to +60 mV, whereas bath application of 10 mmol/L 4-AP significantly diminished the steady state IK(V), which was elicited by voltage steps to +60 mV (by 48%, Fig 3C⇓) and to −40 mV (by 85%; Fig 3D⇓, crosshatched bars). It is noteworthy that 4-AP inhibited the transient IK(V) to a greater extent than the steady state IK(V). These data are consistent with the observations by other investigators.9 10 32
Effects of ChTX, Glibenclamide, and 4-AP on Em in PASMCs
In rat primary cultured PASMCs, the average resting Em values that were measured by using current-clamp technique were −41±1 mV (n=27) and −39±1 mV (n=21), respectively, in the presence and absence of 1.8 mmol/L external Ca2+. Extracellular application of either ChTX (10 nmol/L, Fig 4A⇓ and 4B⇓) or glibenclamide (10 μmol/L, Fig 4C⇓) had no effect on Em. 4-AP (5 to 10 mmol/L), however, significantly and reversibly depolarized PASMCs and elicited extracellular Ca2+-dependent action potentials when the cells were superfused with 1.8 mmol/L Ca2+–containing PSS (Fig 4D⇓ and 4E⇓).
Removal of extracellular Ca2+ had no effect on the 4-AP–induced steady state depolarization that primarily results from a decreased KV conductance but completely abolished the 4-AP–induced action potentials (Fig 4F⇑) that are presumably due to an increased Ca2+ conductance. Composite data shown in Fig 5⇓ indicate that neither 10 nmol/L ChTX (n=9) nor 10 μmol/L glibenclamide (n=13) affected Em (a change of 1.0±0.7 or 1.2±0.9 mV, respectively), whereas 10 mmol/L 4-AP significantly depolarized the cells by 16±1 mV (n=11) and 13±2 mV (n=12) (steady state depolarization), respectively, in the absence and presence of extracellular Ca2+ (Fig 5⇓).
In the presence of extracellular Ca2+, the 4-AP–induced steady state depolarization, in addition to decreased KV conductance, may also be attributable to a sustained Ca2+ influx33 and Ca2+-activated Cl− efflux.18 19 34 The peak of the 4-AP–induced transient depolarization (action potential) was also averaged in Fig 5⇑ (37±4 mV, n=12). Compared with the 13 mV of 4-AP–elicited steady state depolarization, this transient peak was presumably due to the increased Ca2+ influx through voltage-gated Ca2+ channels that resulted from membrane depolarization via the decreased K+ efflux through KV channels.
In the presence of 15 nmol/L ChTX and absence of extracellular Ca2+, 4-AP (0.3 to 10 mmol/L) depolarized PASMCs in a dose-dependent manner (Fig 6⇓). Under conditions in which PASMCs were dialyzed with solutions containing 10 mmol/L EGTA and superfused with Ca2+-free PSS, bath application of 0.3, 1, 3, and 10 mmol/L 4-AP decreased resting Em (depolarization) by 1.9±0.6, 3.2±0.5, 7.5±1.1, and 15.8±1.0 mV, respectively (Fig 6A⇓ and 6B⇓). Furthermore, 4-AP also inhibited IK(V) in a dose-dependent manner (Fig 6A⇓, inset).
In the cells dialyzed with 10 mmol/L EGTA and 5 mmol/L ATP, inability of ChTX and glibenclamide to depolarize PASMCs can be attributable to deactivated KCa and KATP channels.3 26 32 Actually, this is exactly what the data (Figs 4 through 6⇑⇑⇑) showed. The same reagents (4-AP, ChTX, and glibenclamide) were then used to determine, in intact (nondialyzed) cells, which K+ channels (KV, KCa, and/or KATP channels) contribute to regulating [Ca2+]i in PASMCs (see below).
Effects of ChTX, Glibenclamide, and 4-AP on [Ca2+]i in PASMCs
Consistent with the effects on Em, only 4-AP (10 mmol/L), but neither ChTX (20 nmol/L) nor glibenclamide (10 μmol/L), increased [Ca2+]i under resting conditions in intact (nondialyzed) PASMCs (Fig 7⇓). The 4-AP–induced increase in [Ca2+]i featured an initial relatively rapid rise, which after reaching the maximal level, declined gradually to an elevated [Ca2+]i plateau (Fig 7A⇓ and 7D⇓). Bath application of 5 μmol/L verapamil, a blocker of voltage-gated Ca2+ channels, significantly inhibited the 4-AP–induced increase in [Ca2+]i (Fig 8A⇓ and 8C⇓), whereas removal of extracellular Ca2+ abolished the response of [Ca2+]i to 4-AP (Fig 8B⇓ and 8C⇓). These results indicate that the 4-AP–induced increase in [Ca2+]i is due to Ca2+ influx through voltage-gated Ca2+ channels, which are opened by the membrane depolarization resulting from decreased KV channel activity.
The observation that the 4-AP–induced increase in [Ca2+]i was not maintained at the maximal level suggests that a repolarization mechanism (eg, activation of KCa channels) and Ca2+ extrusion and sequestration processes become active after the membrane depolarization and/or the rise in [Ca2+]i. Indeed, administration of 20 nmol/L ChTX in the presence of 4-AP delayed the decline of 4-AP–induced elevation of [Ca2+]i (Fig 9⇓). This suggests that the membrane repolarization process induced by activation of KCa channels due to a rise in [Ca2+]i at least partially contributes to the negative-feedback regulation of depolarization-induced increase in [Ca2+]i in PASMCs.
Time Courses of 4-AP–Induced Membrane Depolarization and Increase in [Ca2+]i
The time courses tmd, tAP, and tCa were determined as follows: in each experiment, t represents the time from the introduction of 4-AP into the tissue chamber to the beginning of the change in the measured parameter. However, in the present study, Em and [Ca2+]i were measured by using two different experimental setups. The perfusion speeds are 1.3 and 2.2 mL/min (see “Materials and Methods”), respectively, in the patch-clamp setup (for measuring Em) and the quantitative fluorescent microscopy system (for measuring [Ca2+]i). Thus, the perfusion rate is ≈1.69 times faster in the fluorescent microscopy system than in the patch-clamp setup. Therefore, all individual tCa values were multiplied by 1.69 to correct for the faster perfusion rate. The rate of 4-AP–induced action potentials (tAP) in the presence of extracellular Ca2+ is 69±9 seconds (n=27), and the time to initiate membrane depolarization (tmd) in the absence of extracellular Ca2+ is 24±2 seconds (n=19). Moreover, the perfusion-corrected time to increase [Ca2+]i (tCa) is 85±5 seconds (n=57); thus, tmd<tAP<tCa. These results suggest that 4-AP–induced membrane depolarization and action potentials precede the 4-AP–induced increase in [Ca2+]i.
PASMCs Have Voltage-Gated, Ca2+-Activated, and ATP-Sensitive K+ Channels
The primary cultured PASMCs used in the present study possess similar physiological, pharmacological, and biochemical properties compared with the freshly dissociated smooth muscle cells and isolated PA rings.35 36 In this cell preparation, a transient (A-type) KV current, a steady state (delayed rectifier) KV current,5 37 a KCa current (Reference 55 and the present study), and an L-type voltage-gated Ca2+ current38 have been described. The electrophysiological and pharmacological properties of these channels are also comparable to those identified in freshly dissociated PASMCs.4 10 11 12 16 25 39 In the isolated PA rings from which PASMCs were dissociated, cromakalim, the opener of KATP channels, significantly blocks 20 mmol/L K+–induced or hypoxia-induced contraction.40 This result indicates that KATP channels, described in many types of vascular smooth muscle cells,2 7 are also present in rat PA smooth muscle.3
ChTX and glibenclamide block KCa and KATP channels with the concentrations for half block (Ki) of 1 to 10 nmol/L1 26 41 and 20 to 100 nmol/L,41 42 respectively. Although each drug has been shown to block some other types of K+ channel as well, ChTX and glibenclamide appear to be fairly selective inhibitors of KCa and KATP channels, respectively, in vascular smooth muscle cells.2 7 41 4-AP, demonstrated to be without effect on KCa11 and KATP2 channels, blocks KV channels with widely differing affinities; Ki ranges from 10 μmol/L to 10 mmol/L.10 11 22 41 43
Voltage-Gated K+ Channels Play a Major Role in the Regulation of Em in PASMCs
The results from the present study indicate that (1) neither the KCa channel blocker ChTX nor the KATP channel blocker glibenclamide affects Em and [Ca2+]i under resting conditions, in spite of their respective blockade effects on IK(Ca) and IK(ATP); (2) the KV channel blocker 4-AP not only depolarizes but also increases [Ca2+]i in PASMCs via its inhibitory effect on IK(V); and (3) ChTX, albeit without effect on resting [Ca2+]i, enhances the 4-AP–induced rise in [Ca2+]i in PASMCs. These data suggest that IK(V) is the major K+ current that is active under resting conditions and contributes to the regulation of resting Em and thus [Ca2+]i in PASMCs, whereas IK(Ca), activated by increased [Ca2+]i and depolarization, is responsible for membrane repolarization as a negative-feedback pathway in regulating Em and Ca2+ influx–induced increase in [Ca2+]i.2 7 16 26 32
It has been proposed that (1) increasing intracellular ATP concentration from 0 to 1 mmol/L significantly depolarizes rabbit PASMCs,3 (2) hypoxia-induced pulmonary vasoconstriction is inhibited by the KATP channel opener cromakalim,40 44 and (3) anoxia-induced pulmonary vasodilation is inhibited by the KATP channel blocker glibenclamide.44 The physiological role of IK(ATP) in regulating Em and [Ca2+]i under resting conditions in PASMCs is not certain because of the very low open probability of KATP channels at physiological ATP concentrations.42 Hyperpolarization caused by activation of KATP channels, however, has been demonstrated to be the mechanism responsible for vascular relaxation induced by a number of endogenous vasodilators.2 3 7 42
Role of Ca2+-Activated and ATP-Sensitive K+ Channels in Regulating Em and [Ca2+]i
The activity of KCa channels increases steeply with both depolarization and increasing [Ca2+], and the [Ca2+] needed to produce half-maximal activation of IK(Ca) (at 0 mV) is 0.5 to 2 μmol/L2 6 7 14 45 46 in various smooth muscle cells, whereas the KCa channels in rat PASMCs show channel openings only with >300 nmol/L [Ca2+] present on the cytoplasmic side.47 Thus, KCa channels in PASMCs are virtually inactive under resting conditions where Em and [Ca2+]i are −40 mV and ≈0.1 μmol/L, respectively. When the cells are depolarized and [Ca2+]i is increased, however, activation of KCa channels tends to repolarize the cells, close voltage-gated Ca2+ channels, and reduce vascular tone.2 7 16 17 26
The concentration of ATP for half-maximal inhibition of IK(ATP) is 20 to 140 μmol/L, whereas 1 to 3 mmol/L ATP completely blocks KATP channels.3 15 42 48 Internal ATP concentration is usually in the millimolar range42 ; thus, KATP channels in muscle cells would normally be held closed by ATP in the resting state. However, because of the high membrane input resistance in vascular smooth muscle cells in the range of 2.6 to 17 GΩ,1 2 3 4 5 the opening of even a few KATP channels by various endogenous vasodilators and metabolic inhibitors would have substantial effects on Em and vascular tone.2 7 42
Resting Em measured in the primary cultures of PASMCs used in the present study (Em, −40 mV) is somewhat more depolarized than that (Em, −55 mV) reported in freshly dissociated PASMCs.3 4 This discrepancy may result from the higher concentration of ATP (5 mmol/L)3 and Mg2+ (4 mmol/L)25 in the pipette solutions used in the present study. In rabbit PASMCs, Clapp and Gurney3 reported that increasing intracellular ATP concentrations from 0 to 3 mmol/L significantly shifted Em from −70 to −53 mV and virtually blocked the glibenclamide-induced depolarization (from 15 to 1 mV). Furthermore, in canine renal arterial smooth muscle cells, increasing intracellular Mg2+ from 0 to 10 mmol/L caused an ≈20-mV change of resting Em (from −52 to ≈−33 mV) and significantly reduced the 4-AP–induced depolarizations (from 21 to 6 mV).25 These observations imply that ATP- and Mg2+-induced inhibitions of IK(ATP) and IK(V), respectively, may contribute to the relatively more depolarized Em measured in the present study.
The 4-AP–sensitive transient (measured at 5 to 50 milliseconds) and steady state IK(V) (measured at 285 to 295 milliseconds of the 300-millisecond test pulse), at a test potential of −40 mV, were 26±11 and 14±4 pA (Fig 3D⇑), respectively. This indicates that KV channels are active at resting Em in PASMCs. 4-AP–induced blockade of KV channels not only depends on voltage and time but also depends on the state of channels per se.49 The resting state blockade of 4-AP on KV channels occurs at negative voltage, when the channels are mostly at their resting state.49 Blockade of the 4-AP–sensitive KV channels at such a resting state would, therefore, cause membrane depolarization and subsequently open voltage-gated Ca2+ channels, promote Ca2+ influx, and ultimately increase [Ca2+]i. This is indeed what was observed in the present study.
When the PASMCs were bathed in PSS containing 4-AP, the membrane depolarization (Fig 4D⇑ and 4E⇑) was prolonged by (1) inhibition of KV channels by 4-AP10 11 25 ; (2) Ca2+ influx through L-type voltage-gated Ca2+ channels, which are inactivated incompletely at a voltage range between −40 and −20 mV33 ; (3) inhibition of KCa channels by chelating [Ca2+]i with 10 mmol/L EGTA, as decreasing intracellular EGTA from 10 mmol/L to 0.1 mmol/L significantly shortens the depolarization duration24 ; (4) inhibition of KATP channels by the 5 mmol/L ATP and 1.2 mmol/L MgCl2 present in the pipette solution,1 2 3 7 ; and (5) Cl− efflux through Ca2+-activated Cl− channels.18 19 34 Thus, the inhibition of repolarization that results from activation of KCa, KV, and KATP channels, as well as the inward currents due to Ca2+ influx and Cl− efflux, may account for the prolongation of Ca2+-dependent spikes induced by 4-AP (Figs 4D⇑, 4E⇑, and 5⇑).
In conclusion, sarcolemmal KV channels in PASMCs are active under resting conditions and are responsible for maintaining resting Em4 16 20 21 22 23 25 32 and thereby helping to regulate Ca2+ conductance and [Ca2+]i.1 2 33 Accordingly, KV channel activity plays a fundamental role in regulating PA tone under physiological conditions. Nevertheless, KCa and KATP channels in PASMC membranes respond to changes of intracellular Ca2+ and ATP, respectively.2 Consequently, KCa channel activity appears to play an important role in controlling Em and, subsequently, [Ca2+]i11 26 by controlling repolarization as a negative-feedback regulator in PASMCs, whereas KATP channel activity regulates Em and [Ca2+]i in response to alteration of the cellular metabolic state.2 Furthermore, KCa and KATP channels may also be targets of various exogenous and endogenous vasoactive substances. Development and utilization of pharmacological activators of K+ (KV, KCa, and KATP) channels, especially KV channels, in PASMCs may greatly benefit the treatment of pulmonary hypertension, which is a major contributor to many cardiopulmonary diseases.
Selected Abbreviations and Acronyms
|FBSCM||=||fetal bovine serum culture medium|
|ICa||=||inward Ca2+ current|
|ICl(Ca)||=||Ca2+-dependent Cl− current|
|IK||=||whole-cell K+ current|
|IK(ATP)||=||ATP-sensitive K+ current|
|IK(Ca)||=||Ca2+-activated K+ current|
|IK(I)||=||inwardly rectifying K+ current|
|IK(V)||=||voltage-gated K+ current|
|KATP channel||=||ATP-sensitive K+ channel|
|KCa channel||=||Ca2+-activated K+ channel|
|Kir channel||=||inwardly rectifying K+ channel|
|KV channel||=||voltage-gated K+ channel|
|PASMCs||=||PA smooth muscle cells|
|PSS||=||physiological salt solution|
|tAP||=||time course of the action potential|
|tCa||=||time course of the increase in [Ca2+]i|
|tmd||=||time course of 4-AP–induced membrane depolarization|
This study was supported by a Grant-in-Aid from the American Heart Association, Maryland Affiliate, Inc; a Special Research Initiative Support grant from the University of Maryland School of Medicine; and National Institutes of Health grant HL-54043. Dr Yuan is a Parker B. Francis Fellow in Pulmonary Research and a recipient of the Giles F. Filley Memorial Award from the American Physiological Society. Dr Yuan thanks A. Aldinger, R.T. Bright, and E.M. Santiago for their technical assistance and is also grateful to Drs M.P. Blaustein, M.L. Tod, L.J. Rubin, and M.T. Nelson for their review of the manuscript.
- Received November 17, 1994.
- Accepted April 25, 1995.
- © 1995 American Heart Association, Inc.
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