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1Department of Pharmacology and Experimental Therapeutics and Cardiovascular Institute, Loyola University Chicago, Maywood; and 2Department of Pharmacology, Northwestern University, Chicago, Illinois
Submitted 16 January 2006 ; accepted in final form 25 October 2006
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ABSTRACT |
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 TOP ABSTRACT MATERIALS AND METHODSRESULTSDISCUSSIONGRANTSREFERENCES |
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-phorbol 12-myristate 13-acetate (10 nM), and the inhibition of IKv by AVP was prevented by the PKC inhibitor calphostin C (250 nM). These results suggest that the stimulation of Ca2+ spiking by physiological concentrations of AVP involves PKC-dependent inhibition of KCNQ5 channels and increased AP firing in A7r5 cells.
potassium channel; signal transduction; membrane potential; calcium; vascular smooth muscle; M current
We have previously demonstrated that concentrations of [Arg8]-vasopressin (AVP) that may be found in the systemic circulation (10–500 pM) modulate the frequency of L-type Ca2+ channel-dependent Ca2+ spikes in A7r5 rat aortic smooth muscle cells. The stimulation of Ca2+-dependent action potentials, which underlie AVP-stimulated Ca2+ spiking, involves activation of a novel signaling pathway (5). Treatment of A7r5 cells with 100 pM AVP leads to increased tyrosine phosphorylation of Kv1.2 delayed rectifier K+ channels (3). We previously speculated (3) that tyrosine phosphorylation of Kv1.2 channels inhibits their function and thereby induces the membrane depolarization that is required for L-type Ca2+ channel activation.
The signal transduction pathway leading to Kv1.2 phosphorylation and Ca2+ spiking requires activation of protein kinase C (PKC) (3, 14). The present study provides the first electrophysiological evidence that activation of PKC at physiological concentrations of AVP leads to inhibition of an outward voltage-sensitive K+ current, depolarizes the membrane, and induces action potential generation in vascular smooth muscle cells. This mechanism may be of fundamental importance to understanding how vasoconstrictor hormones regulate vascular smooth muscle excitability.
The identity of the K+ channels involved in the actions of AVP may be inferred from the electrophysiological and pharmacological characteristics of the Kv current that is inhibited. These characteristics do not match those expected of Kv1.2 channels, but rather fit the known properties of KCNQ channels, a family of Kv channels whose best known roles are as mediators of the "M current," a slowly activating, delayed rectifier K+ current targeted by acetylcholine in the regulation of neuronal excitability. Although KCNQ channels have no previously identified role in vasoconstrictor actions, we have detected expression of KCNQ5 in A7r5 cells and in isolated rat aortic smooth muscle and provide evidence that these channels are targeted in a PKC-dependent manner by physiological vasoconstrictor concentrations of AVP.
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MATERIALS AND METHODS |
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TOPABSTRACTMATERIALS AND METHODSRESULTSDISCUSSIONGRANTSREFERENCES |
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after being filled with internal solutions. Series resistance was not compensated.
The standard bath solution contained (in mM) 5 KCl, 130 NaCl, 10 HEPES, 2 CaCl2, 1.2 MgCl2, and 5 glucose, pH 7.3. Standard internal (pipette) solution contained (in mM) 110 K-gluconate, 30 KCl, 5 HEPES, 1 K2EGTA, and 2 Na2ATP, pH 7.2. Osmolality was adjusted to 268 mosmol/l with D-glucose. Amphotericin B (200 µg/ml) in internal solution was used for membrane patch perforation. Experiments in whole cell perforated-patch configuration were started with series resistance (Rs) below 30 M; cells with an abrupt decrease in Rs were discarded. Stable control currents were recorded for 10–20 min before bath application of pharmacological agents.
Voltage-clamp command potentials were generated using an Axopatch 200B amplifier under control of pCLAMP8 software. Currents were recorded by application of 1- or 5-s voltage steps from a –74-mV holding potential to test potentials ranging from –94 to +46 mV. Voltage commands were applied every 5 or 10 s; results were then normalized to the membrane capacitance. Whole cell currents were digitized at 10 kHz and filtered at 1 kHz for 1-s voltage step duration and digitized at 2 kHz and filtered at 200 Hz for 5 s voltage step duration. We averaged 1,000 points (corresponding to 100-ms recording time with a 1-s voltage step or 500-ms recording time with a 5-s voltage step) to obtain end-pulse steady-state K+ current. Whole cell capacitance was compensated. Leak subtraction was performed for some experiments (see legends) by extrapolation of the linear portion of the current-voltage (I-V) curve negative to –69 mV as described by Passmore et al. (40). Liquid junction potentials were calculated using Junction Potential Calculator provided by pCLAMP8 software and subtracted off-line. To analyze the voltage dependence of channel activation, we fitted the conductance calculated from the tail currents (measured at –30 mV after steps from –74 mV to voltages between –94 and +36 mV) with a Boltzmann distribution: G/Gmax = 1/[1 + exp(V0.5 – V)/s], where G/Gmax is fractional maximal conductance, V0.5 is the voltage of half-maximal activation, and s is the slope factor.
Current-clamp recording of membrane potential was performed in current-clamp fast mode with I = 0 pA. Data are presented as means ± SE. For comparisons between two groups, Student's t-test, paired or unpaired as appropriate, was used for statistical analysis, with P values <0.05 considered statistically significant.
[Ca2+]i measurements. Essentially as described previously (5, 6), A7r5 cells were grown to confluence on glass coverslips or six-well plates. The cells were washed twice with control medium (135 mM NaCl, 5.9 mM KCl, 1.5 mM CaCl2, 1.2 mM MgCl2, 11.5 mM glucose, and 11.6 mM HEPES, pH 7.3) and then incubated in the same medium with 2 µM fura-2 AM, 0.1% bovine serum albumin, and 0.02% Pluronic F127 detergent for 90–120 min at room temperature (20–23°C) in the dark. The cells were then washed twice and incubated in the dark in control medium for 1–5 h before the start of the experiment. Fura-2 fluorescence (at 510 nm) was measured in cell populations at room temperature with a Perkin-Elmer Life Sciences LS50B fluorescence spectrophotometer or a BioTek fluorescence plate reader. Background fluorescence was recorded before cells were loaded with fura-2 (6-well plates) or determined at the end of the experiment by quenching the fura-2 fluorescence for 10–15 min in the presence of 5 µM ionomycin and 6 mM MnCl2 in Ca2+-free medium (coverslips). After background fluorescence was subtracted, the ratio of fluorescence at 340-nm excitation to that at 380 nm was calculated and calibrated in terms of [Ca2+]i.
Calibration of fura-2 fluorescence in terms of [Ca2+]i was carried out as described previously (4) using solutions of known Ca2+ concentration to construct a standard curve. The Ca2+ concentration of the standard solutions was calculated using software (MaxChelator, version 6.60) that accounts for binding of Ca2+ to each constituent of the solution. For analysis of fluorescence ratios recorded from cells, the equation [Ca2+]i = Kd··[(R – Rmin)/(Rmax – R)] (18) was fit to the standard curve (using SigmaPlot software; Systat Software, Point Richmond, CA) and used to convert ratios (R) into [Ca2+]i. In situ calibration of fura-2 fluorescence by direct determination of minimum and maximum ratios [Rmin and Rmax, respectively (18)] from within cells yields similar calibrated values. Traces shown are representative of at least three similar experiments.
RT-PCR. Total RNA was isolated from either A7r5 cells or endothelium-denuded rat aortas using RNeasy mini kit (Qiagen) plus DNase treatment. cDNA was synthesized with iScript cDNA synthesis kit (Bio-Rad), and then one-tenth of the cDNA product was used for PCR. PCR was carried out using Platinum PCR Supermix (Invitrogen) and 10 pmols of forward and reverse primers at the appropriate annealing temperature (dependent on primer pair). Primers were adapted from Ohya et al. (Ref. 39; KCNQ1–3, KCNQ5) and Beisel et al. (Ref. 1; KCNQ4). A portion of the PCR reaction product was run on 0.8% agarose-TBE (Tris-borate-EDTA) gel against a 100-bp ladder as a molecular weight marker (New England Biolabs). Minus-RT controls using the same reaction conditions with A7r5 RNA were negative for all KCNQ primer pairs (not shown). Expected sizes of reaction products are as follows: KCNQ1, 453 bp; KCNQ2, 372 bp; KCNQ3, 424 bp; KCNQ4, 495 bp; and KCNQ5, 240 bp. Products were excised from the gel, purified using the Qiaquick gel purification kit (Qiagen) and cloned into pCR 2.1 vector using a TA cloning kit (Invitrogen) before DNA was sequenced for confirmation. Rat brain RNA was used as a positive control for each primer set.
RNA interference. A short-hairpin RNA (shRNA) lentivirus targeted to the rat KCNQ5, lv-GFP_KCNQ5_shRNA, was constructed using target sequences derived from the predicted KCNQ5 mRNA coding sequence (GenBank accession no. XM_237012), with the assistance of BLOCK-iT RNAi Designer (Invitrogen). Oligonucleotides encoding the 29-nt hairpin target sequence (5'-TCAAGTTGACAGTGGCGGCTACAGAACAG-3') were obtained commercially (Invitrogen), annealed, and cloned into the pSIH1-H1-Puro shRNA lentivirus vector (System Biosciences). The vector is designed to coexpress copGFP protein, a monomeric green fluorescent protein (GFP) cloned from the copepod, Pontellina plumata. Replication-deficient shRNA lentivirus was prepared using the pPACK-H1 Lentivector packaging system and 293 TN producer cell line according to the manufacturer's protocol. Adherent A7r5 cells in culture were infected with lv-KCNQ5_shRNA by using Polybrene (hexadimethrine bromide; Sigma). Five to seven days later, GFP-positive cells were chosen for electrophysiological recordings. A GFP control lentivirus, lv-GFP, was constructed in parallel with pPACK-H1 reagents and was used to infect A7r5 cells for control recordings.
Immunohistochemical detection of KCNQ5. A7r5 cells were subcultured on 12-mm round glass coverslips and infected with lv-GFP_KCNQ5_shRNA as described above. The cells were fixed with 2% paraformaldehyde in phosphate-buffered saline (PBS; 138 mM NaCl, 2.7 mM KCl, 8.1 mM Na2HPO4, and 1.2 mM KH2PO4, pH 7.4) for 15 min, washed twice with PBS, and permeabilized with 0.5% Triton X-100 (Sigma) in PBS for 15 min. After being washed with 0.1% Triton X-100 in PBS, coverslips were blocked with Image-iT FX signal enhancer (Molecular Probes, Eugene, OR) according to the manufacturer's instructions for 30 min, followed by an additional 2-h blocking step with 10% goat serum. Coverslips were washed three times with 0.1% Triton X-100 in PBS and incubated with rabbit polyclonal anti-KCNQ5 antibodies (Chemicon International, Temecula, CA) at a 1:500 dilution for 2 h at room temperature. After three washing steps with 0.1% Triton X-100 in PBS for 15 min each, coverslips were incubated for 2 h at room temperature in the dark with Alexa Fluor 594 goat anti-rabbit IgG secondary antibody (1:400 dilution in 0.1% Triton X-100 plus 10% goat serum in PBS). Three additional washing steps for 15 min with 0.1% Triton X-100 in PBS were performed before coverslips were mounted on glass slides using the SlowFade light antifade kit (Molecular Probes). Cell images were acquired using C Imaging System (Compix) with an Olympus 1X71 inverted epifluorescence microscope (x10 fluorescent objective) and Simple PCI software (version 5.3.1). Two images of each field were captured, at 490-nm excitation wavelength for GFP fluorescence and at 595-nm excitation wavelength for Alexa Fluor 594 fluorescence, respectively. Regions of interest were defined by outlining cells expressing or not expressing GFP fluorescence (identified using 490-nm excitation). Digital images captured using 595-nm excitation were analyzed for mean pixel intensity of regions of interest. Coverslips incubated without primary antibody had no detectable fluorescence with 595-nm excitation (results not shown).
Materials.
Cell culture media were obtained from GIBCO-BRL (Gaithersburg, MD) or MediaTech (Herndon, VA). Fura-2 AM, fura-2 pentapotassium salt, and Pluronic F127 were obtained from Molecular Probes. 4-Phorbol 12-myristate 13-acetate (PMA) was obtained from Calbiochem (San Diego, CA). Correolide was generously provided by Merck Research Laboratories (Rahway, NJ).
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RESULTS |
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TOPABSTRACTMATERIALS AND METHODSRESULTSDISCUSSIONGRANTSREFERENCES |
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Whole cell current-clamp recordings revealed that 100 pM AVP induced a gradual depolarization of the membrane, which eventually triggered repetitive action potential firing with a mean latency of 5.7 ± 0.8 min (n = 6, Fig. 2A).
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, measured under voltage-clamp conditions before and 10 min after application of 100 pM AVP, n = 7, P < 0.005; Fig. 2). This effect was associated with a 10-mV depolarization of the membrane (from –55.9 ± 1.6 to –45.2 ± 1.4 mV, n = 8, P < 0.001) measured before action potential firing in the same cells. If AVP stimulates Ca2+ spiking by inhibition of K+ channels, one might predict that K+ channel blockers would mimic the effects of AVP in A7r5 cells. As shown in Fig. 3A, BaCl2, a nonselective K+ channel blocker (49), stimulated Ca2+ spiking in A7r5 cells in a concentration-dependent manner, reminiscent of the effects of AVP (5). Moreover, 100 µM BaCl2 also mimicked AVP in inhibiting outward K+ currents without affecting the amplitude of L-type Ca2+ current (Fig. 3B) and triggered action potential firing in single A7r5 cells (Fig. 3C).
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20%; Fig. 5B), it did not inhibit current over the physiological range of resting membrane potentials and did not activate Ca2+ spiking in A7r5 cells (not shown), suggesting a nonessential contribution of ATP-sensitive channels to the resting K+ current. A relatively selective inhibitor of Kv channels, 4-aminopyridine (4-AP; 1 mM), induced a slight positive shift of activation voltage dependence without significant inhibition of the current (Fig. 5C). A nonspecific K+ channel blocker, Ba2+ (100 µM), significantly inhibited 70% of K+ currents at negative potentials, with this value diminishing to 40% as the membrane potentials were raised to +36 mV (Fig. 5, D and E). A specific Kv1 family inhibitor, correolide (1 µM) (15, 21), significantly reduced Kv currents in A7r5 cells (by 52 ± 7%; Fig. 5F). Block of IKv by correolide was voltage independent, and there was no shift of the voltage activation curve (not shown). Unlike BaCl2 (Fig. 3), correolide treatment did not stimulate Ca2+ spiking or action potential generation (not shown). KCNQ or Kv7 family channels may contribute to Kv currents in vascular smooth muscle cells (39, 60). We tested the effects of the selective KCNQ channel blockers linopirdine (100 µM) and XE991 (10 µM) and found that both agents significantly inhibited IKv in A7r5 cells (Fig. 5, G and H). We also found that IKv was significantly and reversibly enhanced by 10 µM flupirtine, a KCNQ channel activator (Fig. 5I).
Measuring the time course of inhibition of IKv by 10 µM linopirdine revealed that within 15 min of addition of linopirdine, the AVP-sensitive current was fully inhibited such that addition of 100 pM AVP had no further effect (Fig. 6, A and B). The functional effects of KCNQ channel inhibition on A7r5 cell excitability were also examined using current clamp in single A7r5 cells. Treatment of A7r5 cells with 10 µM linopirdine resulted in repetitive action potential firing (Fig. 6C). In fura-2-loaded A7r5 cell monolayers, linopirdine (5 µM) stimulated Ca2+ spiking (Fig. 6D) and flupirtine (10 µM) transiently inhibited AVP-stimulated Ca2+ spiking (Fig. 6E).
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20–30% of the cell population exhibiting GFP fluorescence 5–7 days after infection. Immunofluorescence staining revealed a significant reduction in KCNQ5 immunoreactivity in GFP-expressing cells compared with non-GFP-expressing cells in the same culture (Fig. 8, A and B). Whole cell patch clamp of the lv-GFP_KCNQ5-shRNA-infected fluorescent cells revealed that IKv was significantly smaller than that recorded from fluorescent cells infected with a GFP control virus (lv-GFP; Fig. 8C).
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DISCUSSION |
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TOPABSTRACTMATERIALS AND METHODSRESULTSDISCUSSIONGRANTSREFERENCES |
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The ability of vasoconstrictor hormones to stimulate arterial vasomotion (rhythmic constrictions of resistance arteries) has been recognized for several decades. These constrictions are associated with action potential firing in the vascular smooth muscle cells of the artery wall (17). The presumed mechanism underlying the stimulation of action potential firing is a depolarization of the smooth muscle cell plasma membrane (sarcolemma) to a threshold potential at which the regenerative opening of voltage-sensitive Ca2+ channels would produce the steeply rising phase of the action potential. The entry of Ca2+ into the cytosol would then result in activation of the cell's contractile apparatus.
Membrane depolarization generally involves alterations in transmembrane ion fluxes. Vasoconstrictor hormones may positively or negatively affect a variety of ion channels, including voltage-sensitive Ca2+ channels, nonselective cation channels, K+ channels, and Cl– channels. The physiological ionic conditions in the cytosol and extracellular environment of vascular smooth muscle cells dictate that depolarization of the membrane from its resting potential would most readily occur by either an increase in Ca2+, Na+, or Cl– permeability or a decrease in K+ permeability of the sarcolemma.
Vasoconstrictor-induced membrane depolarization is often attributed to increased inward current. Inward Cl– currents (due to Cl– efflux) are enhanced by 
-adrenergic stimulation in rabbit portal vein myocytes (13), by endothelin-1 or angiotensin II in rat pulmonary artery myocytes (20, 45, 46), and by endothelin-1 or vasopressin in A7r5 cells (56). Furthermore, the stimulation of Cl– efflux has been implicated in the vasoconstrictor effects of endothelin-1 in rabbit basilar artery (10) and norepinephrine in rat resistance arteries (33) and aorta (32). There also is abundant evidence that vasoconstrictor hormones, including AVP, may enhance inward currents via stimulation of nonselective cation channel activity (Ca2+ and/or Na+ influx) in vascular smooth muscle cells (for reviews, see Refs. 34, 47, 54), including A7r5 cells (26, 29, 31, 37, 58). In most cases these effects have been evaluated using agonist concentrations that are orders of magnitude higher than would be found in the systemic circulation under physiological conditions.
Depolarization due to activation of inward currents (e.g., via Cl– or nonselective cation channel activation) would be associated with a decrease in membrane resistance. In contrast, inhibition of K+ currents would favor an increase in membrane resistance. It is noteworthy that we have observed a net increase in membrane resistance in the response to 100 pM AVP in the present study. This finding is consistent with our hypothesis that inhibition of Kv currents is the primary mechanism for membrane depolarization and stimulation of action potential firing by physiological concentrations of AVP in A7r5 cells.
There are reports that other vasoconstrictor agonists inhibit K+ currents in vascular smooth muscle cells. For example, several earlier studies suggested that endothelin-1 inhibits Kv channels in arterial myocytes (2, 36, 46, 51, 52). The contribution of this effect to endothelin-1-induced membrane depolarization is not clear, however. In rabbit pial arteriolar myocytes exposed to varying concentrations of endothelin-1, membrane depolarization was associated with a decrease in membrane resistance (EC50 
100 pM for both depolarization and decreasing membrane resistance; Ref. 19), suggesting that inhibition of K+ channels was not the predominant depolarizing mechanism. Angiotensin II (100 nM) has been reported to decrease Kv currents in rabbit portal vein myocytes (8) and rat mesenteric arterial myocytes (24) by a PKC-dependent mechanism, although the contributions of this mechanism to membrane depolarization or Ca2+ signaling at physiological concentrations of angiotensin II have not been examined.
Inhibition of outward K+ currents would suffice to depolarize the membrane, but in addition it also may serve to sensitize the cells to depolarization by small increases in inward current. According to Ohm's law, voltage is proportional to the product of current and resistance. Therefore, an increase in resistance would enable a small current to produce a larger change in voltage. In other words, inhibiting K+ currents may render the cells more excitable such that a small injection of inward current (e.g., by activation of inward Cl– or cation currents) would more effectively depolarize the cells to activate L-type voltage-sensitive Ca2+ channels. Activation of L-type Ca2+ channels also would be more likely to produce a regenerative depolarization (action potential), because each channel opening would provide a larger depolarization when membrane resistance is increased.
Our studies have implicated Kv channels in the AVP signal transduction cascade. However, other classes of K+ channels also are expressed in vascular smooth muscle cells, including inward rectifier (Kir), Ca2+-activated (KCa), and ATP-sensitive K+ (KATP) channels [reviewed by Nelson and Quayle (38)]. Although there is evidence from numerous studies that vasoconstrictor agonists also may regulate these other types of K+ channels, they are unlikely to be responsible for the effects of AVP observed in the present study. KCa channels are unlikely to be active in unstimulated cells, where resting [Ca2+]i is typically <100 nM (5). It is unlikely therefore that inhibition of KCa currents would depolarize the membrane under these conditions. Furthermore, we have found that treatment of A7r5 cells with 100 nM iberiotoxin, a specific inhibitor of BKCa channels, did not inhibit the AVP-sensitive K+ current (Fig. 5A) and did not induce depolarization of membrane potential or trigger Ca2+ spiking. On the other hand, KCa channels may be activated by Ca2+ and/or voltage during the course of the action potential and therefore may play a role in the repolarization or afterhyperpolarization phases of the action potential. KATP channels are inhibited by normal cytosolic ATP concentrations and therefore also are expected to be minimally active in healthy unstimulated cells (38). A study by Dumont and Lamontagne (12) found no effect of glibenclamide, a KATP channel blocker, on AVP-induced contraction, and glibenclamide by itself did not induce contraction of unstimulated aortic rings. We also found no effect of glibenclamide on Ca2+ spiking in A7r5 cells but observed a small inhibition of Kv currents at positive membrane potentials (Fig. 5B). Finally, the current-voltage profile of Kir is not consistent with the currents that we observed to be inhibited by AVP, which are outwardly rectifying.
Although often used as a selective Kv channel blocker, 1 mM 4-AP was not an effective inhibitor of Kv currents in A7r5 cells (Fig. 5, C and D). The effects of 4-AP on native vascular smooth muscle Kv currents are highly variable, with aortic myocytes being particularly insensitive (9, 55). Although higher concentrations of 4-AP have been tested in several vascular myocyte preparations (9), 5 mM 4-AP has been reported to have nonspecific effects attributed to changes in cytosolic pH when perforated-patch recording techniques are used (41).
Correolide, a selective Kv1 family inhibitor (15, 21), significantly inhibited Kv currents in the present study but was ineffective in stimulation of Ca2+ spiking in A7r5 cells. The reasons for this apparent discrepancy are not immediately clear. Inhibition of IKv by 1 µM correolide was very gradual. About 50% inhibition of IKv was achieved only after 20–30 min, as observed for native rabbit pial arteriolar Kv currents (7). The relatively slow inhibition of IKv by correolide in A7r5 cells may have prevented the Ca2+ spiking response from developing, because of inactivation of L-type Ca2+ channels or other time-dependent adaptive responses of the cells. It also is possible that correolide has nonspecific effects that interfere with the Ca2+ spiking response.
Despite its partial inhibition by correolide, other pharmacological characteristics of the AVP-sensitive Kv current in A7r5 cells (sensitivity to 100 µM Ba2+, insensitivity to 4-AP) do not fit the expected properties of Kv1 family channels. Ba2+-sensitive, 4-AP-insensitive K+ channels have been implicated in the control of membrane potential and vasoconstriction in rabbit renal arcuate arteries (42). In primary rat aortic smooth muscle cells in culture (23) and guinea pig mesenteric artery myocytes in situ (22), Ba2+ (0.5–1.0 mM) induced significant membrane depolarization, increased membrane resistance, and induced repetitive action potential generation. These effects have been attributed to inhibition of K+ conductance, but the identities of the targeted channels have remained elusive.
The Ba2+ sensitivity, along with the electrophysiological characteristics (kinetics and voltage-dependence of activation, absence of time-dependent inactivation) of the A7r5 Kv current are similar to what has been described for the Kv7 family (KCNQ family) of voltage-gated K+ channels (43). These channels are among the most recently identified mammalian K+ channels and are thought to function in excitable cells to maintain a negative resting membrane potential. These channels have not historically been considered among the cohort of vascular ion channels, but recently, KCNQ channels were found to be expressed in murine portal vein (KCNQ1; Ref. 39), and functional vasoconstrictor effects of KCNQ channel blockers were demonstrated in rat and murine pulmonary arteries (28) and murine portal vein (60). No previous studies have demonstrated regulation of these channels by vasoconstrictor hormones.
Inhibition of Kv currents and vasoconstrictor actions of the selective KCNQ channel blockers linopirdine and XE991 have been the basis for postulating a functional role for KCNQ channels in rat and murine pulmonary arteries and portal vein (28, 39, 60). We have found that these same agents inhibit the AVP-sensitive Kv current in A7r5 cells at concentrations that selectively block KCNQ currents in other cells and that linopirdine mimics AVP in the stimulation of action potential generation and Ca2+ spiking. We also found that the KCNQ channel activator flupirtine has the opposite effect (increasing Kv current and suppressing AVP-stimulated Ca2+ spiking). KCNQ5 expression in A7r5 cells (and freshly isolated rat aortic smooth muscle cells) was confirmed by RT-PCR (whereas KCNQ1–4 were undetectable), and RNA interference targeted to KCNQ5 significantly decreased IKv. On the basis of these observations, we conclude that KCNQ5 channel inhibition mediates the stimulation of Ca2+ spiking via physiological concentrations of AVP in A7r5 cells.
KCNQ5 expression has been demonstrated in a number of brain regions as well as in sympathetic ganglia and skeletal muscle (35, 48), but this is the first demonstration of its expression or function in vascular smooth muscle cells. KCNQ5 channel function has been measured using expression systems in which KCNQ5 overexpression yields currents with characteristics reminiscent of neuronal M currents and regulation by M1 muscarinic receptor activation (48). These properties are similar to the electrophysiological characteristics of Kv currents in A7r5 cells and their regulation by activation of V1a vasopressin receptors. The signal transduction pathways involved in regulation of neuronal KCNQ5 currents by G protein-coupled receptor activation are not fully understood, but both hydrolysis of phosphatidylinositol 4,5-bisphosphate and activation of PKC have been implicated (11).
Is there any relationship between regulation of KCNQ5 and Kv1.2 channels by AVP? A role for Kv1.2 channels in A7r5 cells was suggested by our previous finding that AVP-induced Ca2+ spiking was dependent on PYK2 activation and the associated tyrosine phosphorylation of the Kv1.2 channel protein (3). M1 muscarinic acetylcholine receptor activation can also induce PYK2-dependent tyrosine phosphorylation of Kv1.2 channels and suppression of their activity (16, 25). These findings suggest that M1 receptor activation can initiate the same signaling sequence that we previously implicated for AVP-stimulated Ca2+ spiking in A7r5 cells. As noted above, KCNQ5-mediated M currents also were suppressed in response to M1 receptor activation (48). Both KCNQ channels and Kv1.2 channels are widely distributed in neural tissues (11, 50). If these channels coexist in neurons, their regulation by M1 receptor activation may represent a coordinated response in the stimulation of neuronal excitability. Although the relationship between KCNQ5 and Kv1.2 regulation in A7r5 cells is not completely clear, we may speculate that they are both involved in the regulation of excitability in vascular smooth muscle cells.
In summary, at physiological vasoconstrictor concentrations, AVP stimulates action potential generation and Ca2+ spiking in A7r5 cells via PKC-dependent inhibition of Kv currents. The latter have pharmacological and electrophysiological attributes of KCNQ5 channel currents, and the current amplitudes are decreased by knocking down KCNQ5 expression. Future studies are needed to examine the hypothesis that KCNQ5 channels are mediators in the physiological vasoconstrictor effects of vasopressin and potential targets for therapeutic intervention in cardiovascular diseases.
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ACKNOWLEDGMENTS |
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Present address of C. J. Moran: William Beaumont Hospital, Royal Oak, MI 48073.
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FOOTNOTES |
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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.
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1) subunits in terminal arterioles of rabbit. J Physiol 534: 691–700, 2001.-adrenoceptor-mediated vasoconstriction in the anesthetized rat. Eur J Pharmacol 401: 403–412, 2000.[CrossRef][ISI][Medline]
) and peak inward current recorded at the beginning of the pulse in control (
) followed by application of 10 µM verapamil plus 100 nM iberiotoxin for 10 min (
). Verapamil and iberiotoxin were then replaced with 100 µM GdCl3 (
) for an additional 10 min (n = 4). Peak inward currents recorded in the presence of verapamil plus iberiotoxin and GdCl3 were omitted for clarity. B: representative traces of current recorded in the presence of 100 µM GdCl3 in external solution to isolate Kv component (top). With a holding potential of –74 mV, a voltage step protocol was applied to test potentials from –94 to +36 mV, followed by steps to –30 mV for tail-current recording. Dotted line indicates 0 current level. C: mean current recorded at the end of the pulse (1,000 points recorded over the last 500 ms were averaged) from the traces presented in B after leak subtraction and normalization to the cell capacitance (C = 284 pF). D: voltage dependence of steady-state activation fitted by the single Boltzmann function (solid line, see MATERIALS AND METHODS). Conductance was calculated from the tail currents measured at –30 mV based on a K+ reversal potential of –84 mV. Voltage of half-maximal activation (V0.5) = –38.0 ± 1.6 mV; slope factor (s) = 8.3 ± 0.4 mV (n = 21); G/Gmax, fractional maximal conductance. E: time course of Kv current inhibition by 100 pM AVP recorded by applying a 5-s step to a membrane potential of 0 mV from a holding potential of –74 mV at 15-s intervals. Currents were normalized to mean control current (I/Ic) recorded for 10 min before adding AVP (n = 7). F: I-V curves of mean outward current measured in the presence of 100 µM Gd3+ in control and after exposure to 100 pM AVP for 15 min. After leak subtraction, currents were normalized to the maximal control current (I/Imax; 
