The involvement of inositol 1,4,5-trisphosphate (IP3) formation in the voltage-dependent regulation of intracellular Ca2+ concentration ([Ca2+]i) was examined in smooth muscle cells of the porcine coronary artery. Slow ramp depolarization from −90 to 0 mV induced progressive [Ca2+]i increase. The slope was reduced or increased in the presence of Cd2+ or (±)-1,4-dihydro-2,6-dimethyl-5-nitro-4-(2-[trifluoromethyl]-phenyl)pyridine-3-carboxlic acid methyl ester (Bay K 8644), respectively. The decrease in [Ca2+]i via the membrane hyperpolarization induced by K+ channel openers (levcromakalim and Evans blue) under current clamp was identical to that under voltage clamp. The step hyperpolarization from −40 to −80 mV reduced [Ca2+]i uniformly over the whole-cell area with a time constant of ∼10 s. The [Ca2+]i at either potential was unaffected by heparin, an inhibitor of IP3 receptors. Alternatively, [Ca2+]i rapidly increased in the peripheral regions by depolarization from −80 to 0 mV and stayed at that level (∼400 nM) during a 60-s pulse. When the pipette solution contained IP3 pathway blockers [heparin, 2-aminoethoxydiphenylborate, xestospongin C, or 1-[6-[((17β)-3-methoxyestra-1,3,5-trien-17-yl)amino]hexyl]-1H-pyrrole-2,5-dione (U73122)], the peak [Ca2+]i was unchanged, but the sustained [Ca2+]i was gradually reduced by ∼250 nM within 30 s. In the presence of Cd2+, a long depolarization period slightly increased the [Ca2+]i, which was lower than that in the presence of heparin alone. In coronary arterial myocytes, the sustained increase in the [Ca2+]i during depolarization was partly caused by the Ca2+ release mediated by the enhanced formation of IP3. The initial [Ca2+]i elevation triggered by the Ca2+ influx though voltage-dependent Ca2+ channels may be predominantly responsible for the activation of phospholipase C for IP3 formation.
Smooth muscle cells (SMCs) in large coronary arteries have relatively low membrane excitability under physiological conditions, because of their low expression level of the voltage-dependent Ca2+ channel (VDCC) (Kuriyama et al., 1998; Yamamura et al., 1999). Membrane hyperpolarization in vascular SMCs (VSMCs), induced by an application of ATP-sensitive K+ (KATP) channel openers, results in a decrease in intracellular Ca2+ concentration ([Ca2+]i), presumably because of the suppression of VDCC activity (Quayle et al., 1997). Another potential mechanism for the decrease in [Ca2+]i induced by KATP channel openers has been suggested to be as follows. The membrane hyperpolarization suppresses the phospholipase C (PLC) activity required for inositol 1,4,5-trisphosphate (IP3) formation in rabbit mesenteric arterial SMCs (Itoh et al., 1992). The elevated PLC activity that occurred during the stimulation of G protein-coupled receptors with an agonist may be voltage-sensitive. It has been shown in guinea pig coronary arterial SMCs (CASMCs) that the membrane hyperpolarization induced by a command pulse under voltage clamp reduces the [Ca2+]i, which is elevated in the presence of an agonist. This phenomenon was also observed in the absence of Ca2+ influx, but only in the presence of agonist, strongly suggesting that the PLC activated by agonist stimulation is voltage-dependent in guinea pig CASMCs (Ganitkevich and Isenberg, 1993).
IP3 is a key second messenger that regulates many important physiological functions. The hydrolysis of phosphatidylinositol 4,5-bisphosphate catalyzed by PLC produces IP3, which results in the release of Ca2+ from the sarcoplasmic reticulum (SR) through the IP3R (Berridge, 1993; Clapham, 1995; Berridge et al., 2003). The phosphoinositide-specific superfamily of PLC enzymes has been classified into four isoforms: β, γ, δ, and ε (Rhee and Bae, 1997; Lymn and Hughes, 2000; Rhee, 2001). Among these isoforms, PLCβ is functionally coupled to heterotrimeric G proteins in response to agonist and is responsible for the generation of IP3 from phosphatidylinositol 4,5-bisphosphate (Berridge, 1993; Somlyo and Somlyo, 1994). The PLCβ protein is expressed in VSMCs (Lymn and Hughes, 2000; LaBelle et al., 2002), including the aorta (Blayney et al., 1996, 1998; Cueille et al., 2003; Ocharan et al., 2005), renal artery (Blayney et al., 1998), tail artery (LaBelle et al., 2002), and coronary artery (Cueille et al., 2003; Ansari et al., 2009).
Direct visualization of IP3 in murine Purkinje cells has shown that membrane depolarization itself facilitates the formation of IP3, and this depends on Ca2+ influx and depolarization (Okubo et al., 2001). In contrast to the results in VSMCs (Ganitkevich and Isenberg, 1993), the preceding activation of PLC by an agonist is not required for the depolarization-induced enhancement of IP3 formation in neurons (Okubo et al., 2001). This striking observation indicates that Ca2+-induced IP3 formation and subsequent Ca2+ release from internal Ca2+ stores is functional in neurons. However, there seems to be a discrepancy between the results in VSMCs and neurons, because the [Ca2+]i was independent of the voltage status when the [Ca2+]i was elevated by caffeine in guinea pig CASMCs (Ganitkevich and Isenberg, 1993). It was reported that, in the absence of exogenous agonists, membrane depolarization directly activates muscarinic M3 receptors, which induces the activation of Gq proteins and PLC and subsequent IP3 generation in mouse tracheal SMCs. This direct activation induces an increase in local Ca2+ release, Ca2+ spark, and contraction in mouse tracheal SMCs (Liu et al., 2009). In addition, the VDCC activation and subsequent metabolic Ca2+ release that involves G protein/PLC activation, IP3 synthesis, and Ca2+ release from the SR participate in the sustained arterial contraction through depolarization-evoked RhoA/Rho-associated kinase activity in the VSMCs. These phenomena are independent of the change in the membrane potential itself, or the mere release of Ca2+ from the SR, and require the simultaneous activation of VDCC and the downstream metabotropic pathway with concomitant Ca2+ release (Fernández-Tenorio et al., 2011).
The present study was undertaken to elucidate whether IP3-induced Ca2+ release takes place during depolarization in the absence of receptor stimulation by an agonist in porcine CASMCs, using two-dimensional confocal Ca2+ imaging analysis under voltage clamp. To inhibit IP3-induced Ca2+ release, heparin, 2-aminoethoxydiphenylborate (2-APB), and xestospongin C, which block IP3 binding to receptors in the SR, and 1-[6-[((17β)-3-methoxyestra-1,3,5-trien-17-yl)amino]hexyl]-1H-pyrrole-2,5-dione (U73122), an inhibitor of PLC, were added to the cytoplasm by using recording pipettes. These results constitute evidence in support of the notion that IP3 production is facilitated during prolonged depolarization in the absence of agonist stimulation, but it requires a preceding increase in [Ca2+]i and corresponding outward currents in CASMCs.
Materials and Methods
All experiments were approved by the Ethics Committee of Nagoya City University and conducted in accordance with the Guide for the Care and Use of Laboratory Animals of the Japanese Pharmacological Society.
Single SMCs from a porcine coronary artery were prepared as described previously (Yamamura et al., 1999). In brief, whole hearts of 6-month-old young pigs were obtained from a local slaughterhouse (Nagoya Meat Market, Nagoya, Japan). Small pieces of the left circumflex coronary artery were dissected, cleaned of blood and surrounding tissues, and immersed for 40 min in Ca2+-free Krebs' solution containing 0.2% collagenase (Amano Enzyme, Nagoya, Japan), 1.0% albumin (bovine, fraction V; Seikagaku Kogyo, Tokyo, Japan), 0.1% papain (Sigma-Aldrich, St. Louis, MO), and 0.2% trypsin inhibitor (Sigma-Aldrich) at 37°C. After this incubation, the solution was replaced with Ca2+- and collagenase-free Krebs' solution. Cells were isolated by gentle agitation with a glass pipette, and a few drops of cell suspension were placed in a recording chamber. After these cells had settled, the recording chamber was continuously perfused with the HEPES-buffered solution at a flow rate of 2 ml/min.
The Ca2+-free Krebs' solution had an ionic composition of 112 mM NaCl, 4.7 mM KCl, 1.2 mM MgCl2, 25 mM NaHCO3, 1.2 mM KH2PO4, and 14 mM glucose. The pH was adjusted to 7.4 by gassing with a mixture of 95% O2 and 5% CO2. The HEPES-buffered solution for electrophysiological recording had an ionic composition of 137 mM NaCl, 5.9 mM KCl, 2.2 mM CaCl2, 1.2 mM MgCl2, 14 mM glucose, and 10 mM HEPES. The pH was adjusted to 7.4 with 10 N NaOH. The pipette solution contained 140 mM KCl, 1 mM MgCl2, 10 mM HEPES, and 2 mM Na2ATP. The pH was adjusted to 7.2 with 1 N KOH. A Ca2+ fluorescence indicator was also added to the pipette solution as described below.
Electrophysiological studies were performed in single SMCs, using whole-cell patch-clamp configuration with a CEZ-2400 amplifier (Nihon Kohden, Tokyo, Japan). The electrophysiological recording procedures and data analysis were performed as described previously (Imaizumi et al., 1989). All of the electrophysiological experiments were carried out at 30 ± 1°C.
Confocal Ca2+ Imaging.
Two-dimensional Ca2+ images were obtained by using a fast scanning confocal fluorescent microscope (RCM-8000; Nikon, Tokyo, Japan) equipped with a Fluor 40×, 1.15 NA objective lens (water immersion; Nikon), and Ratio3 software (Nikon) (Imaizumi et al., 1998; Yamamura et al., 2001, 2002). Freshly isolated myocytes were loaded with 100 μM indo-1 (Dojin Laboratories, Kumamoto, Japan) by diffusion from the recording pipette for a few minutes. Indo-1 acetoxymethyl ester (indo-1/AM; Dojin Laboratories; 5 μM) was loaded for 10 min without electrophysiological recording, and then removed for 10 min at room temperature (24 ± 1°C). The argon ion laser excitation wavelength was 351 nm, and the emission wavelengths were 405 and 485 nm. The microscope resolution was 0.33 × 0.27 μm (one pixel) and 0.5 μm in the z-axis direction. The Ca2+ image was scanned over a full frame (512 × 512 pixels). Confocal Ca2+ images were obtained as the average of eight sequential images for 264 ms (33 ms for each). The global [Ca2+]i was defined as the average [Ca2+]i in the whole-cell area. The peripheral and central [Ca2+]i was measured as the average of five circles of 2-μm diameter, which were located at the cell edge or in the center area, not including the nucleus, respectively. The calibration of the indo-1 signal was performed by a method reported previously (Grynkiewicz et al., 1985). The data analysis was performed as described previously (Imaizumi et al., 1998; Yamamura et al., 2001).
Pharmacological reagents including Evans blue [(6,6′-[(3,3′-dimethyl-[1,1′-biphenyl]-4,4′-diyl)bis(azo)]bis[4-amino-5-hydroxy-1,3-naphthalenedisulphonic acid] tetrasodium salt] and heparin were obtained from Sigma-Aldrich. 2-APB was from Calbiochem (La Jolla, CA), and cadmium chloride and xestospongin C [(1R,4aR,11R,12aS,13S,16aS,23R,24aS)-eicosahydro-5H,17H-1,23:11,13-diethano-2H,14H-[1,11]dioxacycloeicosino[2,3-b:12,13-b′]dipyridine] were from Wako Pure Chemicals (Osaka, Japan). HEPES was from Dojin Laboratories, and iberiotoxin was from Peptide Institute Inc. (Osaka, Japan). Levcromakalim [(3S,4R)-3,4-dihydro-3-hydroxy-2,2-dimethyl-4-(2-oxo-1-pyrrolidinyl)-2H-1-benzopyran-6-carbonitrile] and (±)-1,4-dihydro-2,6-dimethyl-5-nitro-4-(2-[trifluoromethyl]-phenyl)pyridine-3-carboxylic acid methyl ester (Bay K 8644) were provided by Kirin Brewery (Tokyo, Japan) and Kyowa Hakko Kogyo (Tokyo, Japan), respectively. Levcromakalim, glibenclamidue [5-chloro-N-[4-(cyclohexylureidosulfonyl)phenethyl]-2-methoxybenzamide], 2-APB, and U73122 were dissolved in dimethyl sulfoxide (DMSO) at concentrations of 10 to 100 mM as a stock solution. Bay K 8644 and xestospongin C were dissolved in ethanol at concentrations of 20 and 30 μM, respectively, as stock solutions.
Pooled data are shown as the mean ± S.E.M. Statistical significance between two groups and among groups was determined by Student's t test and Scheffé's test after one-way analysis of variance, respectively. Significant difference is expressed as *, p < 0.05 or **, p < 0.01 in the figures.
[Ca2+]i Dependence on the Membrane Potential and Ca2+ Influx.
Simultaneous measurements of the [Ca2+]i and membrane potential were performed under confocal microscopy with 100 μM indo-1 under voltage clamp in single SMCs freshly isolated from coronary artery. The resting [Ca2+]i in the whole-cell area (as the global region) at −90 mV was 104 ± 10 nM (n = 7; Fig. 1, A and B). The [Ca2+]i was progressively increased by a slow ramp depolarization in a voltage-dependent manner and reached 390 ± 21 nM at 0 mV (n = 7). There was no significant difference between the peripheral and central [Ca2+]i levels observed at any of the potentials examined. Voltage-dependent outward currents, which were primarily composed of large-conductance Ca2+-activated K+ (KCa1.1 or BKCa) channel currents and voltage-dependent K+ channel currents, were simultaneously observed during the ramp pulse (Yamamura et al., 1999).
In the presence of 100 μM Cd2+, which completely blocked VDCCs, the increase in the [Ca2+]i evoked by the ramp depolarization was significantly reduced at potentials positive to −40 mV (p < 0.05 versus control). The [Ca2+]i rise at 0 mV was reduced by approximately 80% in the presence of Cd2+ (162 ± 17 nM; n = 4; p < 0.01 versus control of 373 ± 11 nM; Fig. 1C). A Cd2+-insensitive component, however, does exist. To exclude the possibility of an insufficient Cd2+ concentration, higher concentrations of Cd2+ were used in CASMCs. The [Ca2+]i decrease induced by higher concentrations (500 μM and 1 mM; n = 4 for each) tended to be larger than that by a lower concentration (100 μM; Fig. 1C), but the Cd2+-insensitive component remained. The removal of extracellular 2.2 mM Ca2+ resulted in more severe inhibition of the [Ca2+]i increase at 0 mV (by 95 ± 2% reduction; n = 5). On the other hand, in the presence of 1 μM Bay K 8644, which is an activator of VDCCs, the [Ca2+]i was increased significantly at potentials positive to −70 mV. The [Ca2+]i in the absence and presence of Bay K 8644 at 0 mV was 388 ± 5 and 561 ± 55 nM (n = 5; p < 0.01), respectively (Fig. 1D). The [Ca2+]i increase induced by the ramp pulse seemed to be saturated in the presence of Bay K 8644 at potentials positive to −20 mV. The changes in [Ca2+]i elevation caused corresponding alterations in the amplitude of the outward currents in the presence of Cd2+ or Bay K 8644. The relationship in the presence of Bay K 8644 was shifted to the right by a further addition of Cd2+ and became identical to that in the presence of Cd2+ alone (data not shown). These results indicate that most of the [Ca2+]i increase during the slow ramp depolarization was caused by Ca2+ influx through VDCCs, and the effect was enhanced by Bay K 8644. It is, however, notable that the substantial increase in [Ca2+]i occurred even in the presence of Cd2+, which may suggest Ca2+ release from intracellular storage or, alternatively, Ca2+ influx through a Cd2+-insensitive pathway during the slow ramp depolarization.
[Ca2+]i during the Hyperpolarization Induced by K+ Channel Openers.
The relationships between the decrease in [Ca2+]i and membrane hyperpolarization caused by the activation of K+ channels by two different types of openers were determined by using the slow ramp pulse. Evans blue has been reported to activate KCa1.1 channels via an enhancement of the α-subunit (Hollywood et al., 1998; Wu et al., 1999; Yamada et al., 2001). In preliminary experiments, it was confirmed that the application of 100 μM Evans blue induced membrane hyperpolarization by 13.5 ± 1.0 mV (n = 3; p < 0.01 versus a resting membrane potential of −40.8 ± 0.7 mV), which was abolished by the addition of 100 nM iberiotoxin (to −35.5 ± 0.8 mV; n = 3), a specific inhibitor of the KCa1.1 channel. The simultaneous measurement of [Ca2+]i and the membrane potential under current clamp indicated that the average resting membrane potential and [Ca2+]i were −43.4 ± 2.6 mV and 154 ± 9 nM, respectively, under the control conditions (n = 4). The application of 100 μM Evans blue caused the membrane hyperpolarization to −55.0 ± 1.4 mV (n = 4; p < 0.05 versus control) and, correspondingly, a decrease in the [Ca2+]i to 119 ± 11 nM (n = 4; p < 0.05; Fig. 2). The effect of levcromakalim, an activator of the KATP channel, was also examined under the same experimental conditions, except for the removal of 2 mM ATP in the pipette solution to enhance KATP channel activity. The application of 10 μM levcromakalim induced a membrane hyperpolarization of 13.3 ± 1.0 mV (n = 6; p < 0.01 versus resting of −42.3 ± 1.8 mV). Levcromakalim-induced hyperpolarization was abolished by the exposure to 10 μM glibenclamide (to −40.7 ± 2.4 mV; n = 6). In the simultaneous measurement of the membrane potential and [Ca2+]i, the application of levcromakalim induced the hyperpolarization of myocytes from −44.4 ± 0.9 to −57.8 ± 0.6 mV (n = 5; p < 0.01) and decreased the [Ca2+]i from 152 ± 8 to 119 ± 14 nM (n = 5; p < 0.05; Fig. 2C). The relation between the membrane potential and [Ca2+]i in the absence and presence of Evans blue or levcromakalim was in good accord with the relationship determined by the slow ramp under voltage clamp (p > 0.05 versus the fitting line; χ2 test). When the effects of Evans blue and levcromakalim on [Ca2+]i were examined by using the slow ramp in the voltage clamp mode in the same manner as shown in Fig. 1B, the relationship between [Ca2+]i and the membrane potential was identical to that in their absence (n = 4 for each; data not shown). These results confirm that the decrease in [Ca2+]i by membrane hyperpolarization caused by the activation of K+ channels by these openers may be identical to that induced by hyperpolarization in a voltage-clamp configuration.
Moreover, in the presence of Bay K 8644, the effects of Evans blue and levcromakalim on [Ca2+]i were examined by using 5 μM indo-1/AM without the electrophysiological recording. The average [Ca2+]i in the presence of 1 μM Bay K 8644 was 239 ± 10 nM (n = 15; p < 0.01 versus control of 126 ± 3 nM; Fig. 3, A and B). By the addition of 100 μM Evans blue [Ca2+]i was reduced to 165 ± 3 nM (n = 15; p < 0.01), which then was returned to 202 ± 6 nM by the further addition of 100 nM iberiotoxin (n = 15; p < 0.01 versus Evans blue alone). Before the addition of 10 μM levcromakalim the average [Ca2+]i was 242 ± 6 nM in the presence of Bay K 8644 (n = 22; p < 0.01 versus control of 125 ± 2 nM; Fig. 3, C and D). [Ca2+]i was reduced to 165 ± 2 nM (n = 22; p < 0.01) by levcromakalim and returned to 203 ± 7 nM by the further addition of 10 μM glibenclamide (n = 22; p < 0.01 versus levcromakalim alone). It is clear that the effect of K+ channel openers in decreasing the [Ca2+]i was marked when the slope of the relationship between [Ca2+]i and membrane potential was steep in the presence of a VDCC activator.
Local Control of the Decrease in [Ca2+]i by Membrane Hyperpolarization.
Because the decrease in the [Ca2+]i induced by membrane hyperpolarization may be locally and not uniformly controlled in CASMCs, the images of [Ca2+]i decrease by step hyperpolarization were analyzed (Fig. 4). The global [Ca2+]i at −40 mV was 187 ± 1 nM (n = 5). The membrane hyperpolarization from the holding potential of −40 to −80 mV caused a global [Ca2+]i decrease to 108 ± 6 nM with a decay time constant of 10.0 ± 0.7 s (n = 5). We analyzed quantitatively whether the [Ca2+]i decrease during hyperpolarization occurred uniformly in CASMCs (Fig. 4, A–C). At the holding potential of −40 mV the [Ca2+]i in the central and peripheral regions was 184 ± 5 and 190 ± 2 nM, respectively (n = 5; p > 0.05; Fig. 4D). With hyperpolarization of the membrane from −40 to −80 mV for 30 s, the [Ca2+]i in the central and peripheral regions was reduced to 105 ± 11 and 104 ± 8 nM (n = 5; p > 0.05) with a decay time constant of 10.6 ± 0.8 and 8.8 ± 0.5 s, respectively (n = 5; p > 0.05; Fig. 4E).
Ca2+ images of CASMCs during hyperpolarization from 0 to −80 mV were also analyzed in the same manner as shown in Fig. 4 (Fig. 5). When the membrane potential was held at 0 mV for 60 s, the global [Ca2+]i was 390 ± 17 nM (n = 5). The hyperpolarization to −80 mV reduced the global [Ca2+]i to 113 ± 7 nM with a decay time constant of 8.0 ± 0.3 s (n = 5). The global [Ca2+]i at 0 mV in the peripheral region was not significantly different from that in the central region (p > 0.05; Fig. 5D). The decay time constant in the peripheral regions (6.7 ± 0.3 s; n = 5) was, however, significantly smaller than those of the global regions (n = 5; p < 0.05) and central regions (8.6 ± 0.5 s; n = 5; p < 0.05) (Fig. 5E). It is clear from the time constants and line-scan Ca2+ image in Fig. 5C that the [Ca2+]i decrease in the peripheral region was faster than in the central region during membrane hyperpolarization.
[Ca2+]i and IP3 Formation during Membrane Hyperpolarization.
The possibility that the decrease in [Ca2+]i during membrane hyperpolarization under voltage clamp is partly caused by reduced IP3 formation was examined in the absence of an agonist. The change in [Ca2+]i induced by hyperpolarization from −40 to −80 mV was measured in the presence of 1 mg/ml heparin in the pipette solution, a blocker of IP3 receptors (IP3Rs) (Fig. 6A). The [Ca2+]i at both −40 and −80 mV was unaffected by heparin (184 ± 6 and 100 ± 2 nM, respectively; n = 4; p > 0.05 versus control of 177 ± 9 and 106 ± 11 nM; Fig. 6B). The decrease in [Ca2+]i was fitted by means of a single exponential function with the time constants of 10.7 ± 1.1 s (n = 4) and 10.6 ± 1.4 s (n = 4; p > 0.05) in the absence and presence of heparin, respectively (Fig. 6C). In addition, there was no significant difference in the inward current amplitude between the control (152 ± 14 pA; n = 4) and heparin-treated cells (141 ± 3 pA; n = 4, p > 0.05) (Fig. 6D). The addition of other blockers of IP3Rs, 50 μM 2-APB or 10 μM xestospongin C, in the pipette solution, did not affect the [Ca2+]i, decay time constant or current amplitude (n = 4 for each; p > 0.05 versus vehicle; Fig. 6, B-D). In these experiments, the pipette solution contained 0.05% DMSO as the solvent even in the control. Similar results were also obtained by the internal addition of 10 μM U73122 (n = 5), a PLC inhibitor, which contained 0.05% ethanol as the solvent. These results suggest that the difference in Ca2+ release by the production of IP3 at −40 and −80 mV may be negligible under these experimental conditions, and the decrease in the [Ca2+]i induced by the step hyperpolarization may be simply caused by the decrease in Ca2+ influx.
Local Control of the [Ca2+]i Increase Induced by Membrane Depolarization.
Ca2+ images of CASMCs during depolarization from −80 to 0 mV were analyzed in the same manner as shown in Fig. 4 (Fig. 7). When the membrane potential was held at −80 mV for 60 s, the global [Ca2+]i was 109 ± 8 nM (n = 6). The depolarization to 0 mV elevated the global [Ca2+]i to 387 ± 16 nM, with a rise time constant of 2.8 ± 0.4 s (n = 6). The global [Ca2+]i at 0 mV in the peripheral region was not significantly different from that in the central region (p > 0.05; Fig. 7D). It is noteworthy that the rise time constant in the peripheral regions (1.3 ± 0.2 s; n = 6) was significantly smaller than in the global areas (n = 6; p < 0.05) and central areas (2.9 ± 0.4 s; n = 6; p < 0.05) (Fig. 7E). It is clear from the time constants and Ca2+ line-scan image in Fig. 7C that the [Ca2+]i increase in the peripheral region was faster than that in the central region during membrane depolarization. In Fig. 7C, the contraction of myocytes was clearly observed, approximately 3 s after a depolarizing pulse.
Modulation of [Ca2+]i via IP3 Formation by Membrane Depolarization.
In the next series of experiments, the possibility that membrane depolarization facilitates IP3 formation and subsequent Ca2+ release from storage sites was examined by the use of agents suppressing the IP3-mediated increase in [Ca2+]i. The long depolarization induced a rapid increase in the [Ca2+]i from 107 ± 4 nM to a peak of 371 ± 13 nM in control myocytes (n = 8) and from 103 ± 3 to 392 ± 16 nM in 1 mg/ml heparin-treated myocytes (n = 4; p > 0.05), respectively (Fig. 8, A–C). The [Ca2+]i reached a peak within 5 s of the start of depolarization in both control and heparin-treated cells. The global [Ca2+]i was maintained at a high level in the control cells (343 ± 12 nM at 60 s; 90 ± 2% of the peak; n = 8; p > 0.05 versus the peak). In contrast, the global [Ca2+]i declined gradually to a steady level of 224 ± 9 nM within 60 s in the heparin-treated cells (n = 4; p < 0.01 versus the corresponding peak and also versus the steady level in the control). Heparin reduced the sustained increase in [Ca2+]i at 60 s from the start of depolarization to 42 ± 3% of the peak (n = 4). Likewise, the decay of the outward currents in the heparin-treated cells (from 1061 ± 49 pA at the peak to 507 ± 39 pA at 60 s, a 49 ± 5% decrease; n = 8) was significantly faster and larger than that in the control cells (from 1103 ± 40 pA at the peak to 354 ± 37 pA at 60 s, a 32 ± 3% decrease; n = 4; p < 0.05) (Fig. 8D).
The effects of 2-APB, U73122, and xestospongin C on [Ca2+]i during depolarization were also examined in the same manner. Figure 9, A and B clearly demonstrates that the [Ca2+]i and current amplitude during prolonged depolarization were decreased by 50 μM 2-APB and 10 μM U73122 in the pipette solution, respectively. The [Ca2+]i peaks of 392 ± 39, 417 ± 21, and 405 ± 41 nM were reduced to the steady-state level of 360 ± 24, 242 ± 7 nM (p < 0.01 versus the steady state of the vehicle, 0.05% DMSO), and 254 ± 21 nM (p < 0.05) in the absence and presence of 2-APB or U73122, respectively (n = 4 for each) (Fig. 9C). The maximum amplitude of the outward currents evoked by depolarization was unaffected by 2-APB and U73122 (n = 4; p > 0.05; Fig. 9D). On the other hand, a significant decrease in current amplitude during prolonged depolarization was observed with the internal addition of 2-APB (324 ± 23 pA; n = 4; p < 0.01 versus vehicle at 529 ± 13 pA) and U73122 (324 ± 34 pA; n = 4; p < 0.05). A similar time course for the [Ca2+]i reduction and current decrease during the prolonged depolarization was observed in the case of an internal application of 10 μM xestospongin C (n = 4) containing 0.05% ethanol as the solvent.
The [Ca2+]i increase in the step depolarization from −80 to 0 mV was markedly reduced by the addition of 100 μM Cd2+ to the bathing solution (n = 7; p < 0.01; Fig. 10), and the increase was statistically significant. In the presence of heparin in the pipette solution, the addition of Cd2+ significantly reduced the [Ca2+]i at both the peak and steady state (n = 7; p < 0.01). In the presence of Cd2+, there was no significant difference in [Ca2+]i either at the peak or the steady state between the absence and presence of heparin.
The present study clearly shows, by simultaneous measurement of the Ca2+ image and electrophysiological signal, that the [Ca2+]i is regulated by Ca2+ influx through VDCCs, an effect that is modulated directly by the membrane potential, so that the change in VDCC activity is exerted on [Ca2+]i proportionally. The decrease in the [Ca2+]i induced by K+ channel openers is mainly caused by a reduction of Ca2+ influx resulting from the decrease in VDCC activity mediated by the membrane hyperpolarization in VSMCs. In addition, the sustained increase in [Ca2+]i during depolarization is partly caused by the Ca2+ release mediated by the activation of PLC followed by enhanced IP3 formation in VSMCs (Fig. 11).
Vascular smooth muscle tone is determined by the [Ca2+]i level, which is directly regulated by the membrane potential (Carl et al., 1996). In the range of −90 to 0 mV, the L-type Ca2+ current amplitude was enhanced after the depolarization of the cell membrane and reached a peak at 0 mV (Yamamura et al., 1999). When the VDCC activity was enhanced by the agonist, the depolarization-evoked [Ca2+]i was increased compared with the absence of the agonist. On the other hand, the blockade of VDCCs depressed the extent of the [Ca2+]i increase (>80% decrease). These results indicate that the extent of the Ca2+ influx depends on VDCC activity (or the open probability) and the major pathway of the [Ca2+]i increment by the membrane depolarization is through the VDCCs.
It is still unclear whether the intracellular Ca2+ mobilization during membrane hyperpolarization is induced by the potentiation of K+ channel activities in vascular smooth muscle. It has been suggested that the major mechanism underlying the vasodilating effect of the enhancement of KATP channel activity may be caused by the decrease in Ca2+ influx through VDCCs by the membrane hyperpolarization in VSMCs, resulting in the decrease in vascular tone (Nelson and Quayle, 1995). The Ca2+ mobilization during the membrane hyperpolarization induced by a KATP channel opener was demonstrated by the simultaneous measurement of [Ca2+]i and the membrane potential. The degree of [Ca2+]i decrease induced by membrane hyperpolarization was consistent with that of [Ca2+]i change induced by the membrane potential. Further experiments in voltage-clamped cells showed the application of levcromakalim did not affect the relationship between the [Ca2+]i distribution and the membrane potential. The results obtained from recordings under both the current- and voltage-clamp modes indicate that the membrane hyperpolarization via the opening of the KATP channel causes a decrease in Ca2+ influx through VDCCs, which results in the decrease of the smooth muscle tonus to achieve vascular relaxation.
The commonly used KCa1.1 channel opener 1,3-dihydro-1-[2-hydroxy-5-(trifluoromethyl)phenyl]-5-(trifluoromethyl)-2H-benzimidazol-2-one (NS-1619) (Sellers and Ashford, 1994; Holland et al., 1996) has various side effects such as Ca2+ channel inhibition (Edwards et al., 1994; Holland et al., 1996) and Ca2+ release from internal Ca2+ storage sites (Yamamura et al., 2001). Therefore, Evans blue instead of NS-1619 was used, because it is reported to be a KCa1.1 channel opener that does not affect other types of ion channels (Hollywood et al., 1998). The relationship between [Ca2+]i decrease and membrane hyperpolarization induced by Evans blue is in agreement with that by levcromakalim, which indicates that the change in [Ca2+]i and the membrane potential by the potentiation of KCa1.1 channel activity is similar to that resulting from the activation of the KATP channel. Even though a different K+ channel was activated, the extent of the reduction in [Ca2+]i was equal. It is possible that drugs that activate the K+ channel without Ca2+ channel inhibition may have a relaxant effect similar to KATP channel openers. KATP channel openers are used in the treatment of angina, hypertension, bronchial asthma, hypersensitive urinary bladder, and other diseases (Edwards and Weston, 1995). On the other hand, KCa1.1 channel openers have not come into use as therapeutic drugs to date, although they are proposed as potential targets in hypertension, urinary incontinence, and erectile dysfunction (Lawson and McKay, 2006; Ledoux et al., 2006). The blockade of the KCa1.1 channels, which are highly expressed in VSMCs (Kuriyama et al., 1998), results in a membrane depolarization of several millivolts and an increase in muscle tone, so these channels are thought to regulate the resting membrane potential and muscle tone (Nelson and Quayle, 1995). In addition, it is suggested that the potentiation of KCa1.1 channel activity, such as spontaneous transient outward currents and hyperpolarizations, is triggered by local Ca2+ release from the ryanodine receptor (RyR) on the SR, termed a “Ca2+ spark” (Nelson et al., 1995) that contributes to the negative feedback mechanism for the regulation of smooth muscle tone (Bolton and Imaizumi, 1996; Imaizumi et al., 1999; Jaggar et al., 2000; Ohi et al., 2001; Cheng and Lederer, 2008). The evidence suggests that KCa1.1 channel openers as well as KATP channel openers exert a relaxant effect.
One possible mechanism of Ca2+ sensitization is the activation of the protein kinase C, which is mediated by the formation of diacylglycerol after the activation of PLC by agonists. It is conceivable that the membrane hyperpolarization may inhibit phosphatidylinositol turnover, resulting in a reduction in IP3, or the membrane hyperpolarization produced by K+ channel openers may affect intracellular Ca2+ stores and suppress only the activity of IP3-induced Ca2+ release channels. It is suggested that the membrane potential regulates the membrane-associated enzymatic activity of PLC (Itoh et al., 1992; Ganitkevich and Isenberg, 1993). Whether IP3 production depends on membrane potential and the regulation of [Ca2+]i was examined in VSMCs by using heparin, which is reportedly a blocker of the IP3Rs on the SR. Under voltage-clamp conditions, the internal addition of heparin did not affect the relationship between [Ca2+]i and membrane potential (n = 4; p > 0.05 versus absence of heparin), which indicates that IP3 production, that is, PLC activity may not have contributed to the regulation of [Ca2+]i in this case. In addition, the [Ca2+]i during hyperpolarization was not affected by IP3 production.
Membrane depolarization activates VDCCs and thus induces Ca2+ release via RyRs, a process that is obligatory for excitation-contraction coupling and other physiological responses in skeletal and cardiac muscles. However, depolarization-induced Ca2+ release and its functional significance, as well as the underlying signaling mechanisms of the G protein/PLC/IP3-mediated Ca2+ release cascade in VSMCs, are largely unknown. It has been reported that membrane depolarization causes a direct activation of muscarinic M3 receptors in the absence of exogenous agonists, which causes the activation of G proteins and PLC and subsequent IP3 generation in mouse tracheal SMCs (Liu et al., 2009) as well as in rat basilar arterial SMCs (del Valle-Rodríguez et al., 2003). In addition, VDCC activation by high K+ stimulation and the subsequent metabolic Ca2+ release pathway that involves G protein/PLC activation, IP3 synthesis, and Ca2+ release from the SR, participates in the sustained arterial contraction in basilar and aortic SMCs (Fernández-Tenorio et al., 2011). These results indicate that L-type Ca2+ channel activation constitutes the key event that triggers depolarization-dependent metabotropic Ca2+ release from the SR and smooth muscle sensitization through the activation of the RhoA/Rho-associated kinase pathway. These phenomena are independent of the change in the membrane potential itself, or at least the release of Ca2+ from the SR, but they do require the simultaneous activation of VDCCs and the downstream metabotropic pathway with concomitant Ca2+ release (Fernández-Tenorio et al., 2011).
In this study, we demonstrated that the sustained phase of [Ca2+]i increase induced by long depolarization is regulated by IP3 production via PLC activation, which is different from membrane hyperpolarization. In the presence of IP3R blockers, the [Ca2+]i level was rapidly increased by depolarization to the same level as in control cells (∼400 nM), but the sustained [Ca2+]i was gradually reduced by ∼250 nM within 30 s. Similar results were obtained from myocytes treated with an inhibitor of PLC. These studies disclose that membrane depolarization increases IP3 production without the need of any stimulation by an agonist and the generated IP3 is required for the sustained increase in [Ca2+]i in VSMCs. In the presence of Cd2+, long depolarization slightly increased the [Ca2+]i, which was apparently lower than that in the presence of heparin alone. These results indicate that the sustained increase in [Ca2+]i during long depolarization is mainly caused by the Ca2+ influx resulting from VDCC activation and subsequently evokes Ca2+ release, an effect that is caused by the enhanced formation of IP3, in VSMCs.
In conclusion, the [Ca2+]i is regulated by the activity of the VDCC, which is linked directly to the membrane potential and [Ca2+]i reduction by K+ channel openers. It is mainly caused by the decrease of VDCC activity by membrane hyperpolarization via the activation of the K+ channel in VSMCs. The initial [Ca2+]i elevation triggered by Ca2+ influx though VDCCs and subsequent Ca2+-induced Ca2+ increase from RyRs on the SR, termed a “Ca2+ hot spot” (Imaizumi et al., 1998; Morimura et al., 2006), may be primarily responsible for the activation of PLC that is required for IP3 formation and to maintain the sustained phase of [Ca2+]i increase in VSMCs.
Participated in research design: Yamamura and Imaizumi.
Conducted experiments: Yamamura.
Performed data analysis: Yamamura, Ohya, Muraki, and Imaizumi.
Wrote or contributed to the writing of the manuscript: Yamamura and Imaizumi.
We thank Dr. Wayne Giles (University of Calgary, Calgary, Canada) for providing data acquisition and analysis programs.
This work was supported by a Grant-in-Aid for Scientific Research on Priority Areas from the Ministry of Education, Culture, Sports, Science, and Technology [Grant 20056027] (to Y.I.); a Grant-in-Aid for Scientific Research (B) from the Japan Society for the Promotion of Science [Grant 23390020] (to Y.I.); a Grant-in-Aid for Young Scientists (B) from the Japan Society for the Promotion of Science [Grant 23790092] (to H.Y.); and a Grant-in-Aid from the Takeda Science Foundation (to H.Y.).
Article, publication date, and citation information can be found at http://jpet.aspetjournals.org.
- smooth muscle cell
- coronary arterial SMC
- vascular SMC
- intracellular Ca2+ concentration
- dimethyl sulfoxide
- indo-1/acetoxymethyl ester
- inositol 1,4,5-trisphosphate
- IP3 receptor
- KATP channel
- ATP-sensitive K+ channel
- phospholipase C
- ryanodine receptor
- sarcoplasmic reticulum
- voltage-dependent Ca2+ channel
- Bay K 8644
- (±)-1,4-dihydro-2,6-dimethyl-5-nitro-4-(2-[trifluoromethyl]-phenyl)pyridine-3-carboxylic acid methyl ester
- KCa1.1/BKCa channel
- large-conductance Ca2+-activated K+ channel.
- Received March 12, 2012.
- Accepted May 14, 2012.
- Copyright © 2012 by The American Society for Pharmacology and Experimental Therapeutics