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GASTROINTESTINAL, HEPATIC, PULMONARY, AND RENAL
Intestinal Disease Research Program, McMaster University, Hamilton, Ontario, Canada
Received for publication
June 20, 2006
Accepted
August 30, 2006.
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Abstract |
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; Koh et al., 1998; Thomsen et al., 1998). Along the length of the intestine, there are multiple intrinsic pacemaking sites that create a stepwise gradient in frequency and, hence, transmit pacemaker-evoked slow waves toward the distal intestine, prompting anal propagation of the slow wave (Diamant and Bortoff, 1969). Without muscle excitation, slow waves do not normally bear action potentials, and little contractile activity is associated with them (Lammers and Slack, 2001). During muscle excitation, neural or otherwise, action potentials are generated by depolarized smooth muscle cells. In the circular and longitudinal muscle layers of the small intestine, the membrane potential is only sufficiently depolarized during the slow-wave plateau phase (Lammers and Slack, 2001). Because of the propagating nature of the slow waves, a peristaltic motor pattern occurs. A critically important component of peristaltic motor activity during digestion is that high-frequency pacemaker activity switches from the most proximal part of the intestine to areas where luminal contents are located (Bercik et al., 2000; Seerden et al., 2005). This results in a complex pattern of peristaltic and "anti"-peristaltic movements that aid in mixing and absorption. Although the mechanism of this erratic creation of dominant pacemaker sites has not been extensively studied, it is likely that local neural excitation plays an important role. Hence, the interaction between neurotransmitters and ion channels displayed by pacemaker cells needs to be elucidated to build a comprehensive understanding of neural regulation of pacemaker activity.
The ICC-AP form extensive networks at both sides of the myenteric plexus region, associated with smooth muscle cells of both the circular and longitudinal muscle layers and surrounding the ganglia of the myenteric plexus (Faussone-Pellegrini, 1985). The close approximation of the myenteric plexus with the ICC-AP network suggests neural regulation of intestinal pacemaking. Indeed, studies recording the electrical slow wave in canine colonic smooth muscle have demonstrated cholinergic regulation, showing that acetylcholine decreases slow-wave frequency, increases slow-wave duration, and enhances intestinal contractility (Huizinga et al., 1984; Hara and Szurszewski, 1986; Sanders and Smith, 1986). However, it is difficult to establish a direct link between neurotransmitter release and control of ICC since the aforementioned studies were performed on preparations including smooth muscle, ICC, and nerves. Recently, two studies have addressed the question of whether or not neurotransmitters directly regulate pacemaker currents in ICC. In isolated ICC of the murine small intestine, noradrenaline modulates pacemaker currents by decreasing pacemaker frequency (Jun et al., 2004). In ICC of the murine stomach, acetylcholine increased pacemaker frequency (Kim et al., 2003).
We have shown in cultured ICC-AP that significant ether-a-go-go-related gene (ERG) K+ current is generated between potentials of –30 and –20 mV (McKay et al., 2006) and that the ERG K+ current is a key regulator of voltage oscillation frequency and plateau potential duration (between –30 and –20 mV) (Zhu et al., 2003). Several groups have investigated the modulatory capability of ERG K+ currents on different cell types via the activation of G-protein-coupled receptors, such as muscarinic receptor, thyrotropin-releasing hormone receptor, 
1A-adrenergic receptor, and 
-adrenergic receptors (Schledermann et al., 2001; Hirdes et al., 2004; Thomas et al., 2004). Second messengers, such as PKC and PKA, cAMP, and PIP2, have been reported to modulate the ERG K+ channel through phosphorylation sites, interaction with a cyclic nucleotide binding domain, and positively charged amino acids, respectively (Barros et al., 1998; Kiehn et al., 1998; Cui et al., 2000; Bian et al., 2001; Thomas et al., 2003).
Epperson et al. (2000) have reported the isolation of mRNA of muscarinic acetylcholine receptors (M2 and M3 subtypes) from cultured and freshly isolated murine ICC-AP. Cholinergic control of pacemaking could be regulated through activation of the expressed M2 receptor (M2R) or M3 receptor (M3R) proteins in ICC-AP. Therefore, we hypothesized that activation of a muscarinic receptor subtype on ICC results in modulation of the ERG K+ current, an important regulator of the pacemaker current. Thus, the objectives for this study were to determine whether the M2R and/or M3R proteins were expressed on ICC, to determine whether the ERG K+ current in ICC is a downstream target of muscarinic receptor activation, and to investigate by what mechanism muscarinic receptor activation modifies ERG K+ channel behavior.
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Materials and Methods |
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Immunohistochemistry
Explant Cultures. After 2 to 3 days of incubation, glass coverslips containing explanted cultures were transferred to a separate four-well dish. Explants were washed 2 x 10 min with 0.01 M phosphate-buffered saline (PBS) and fixed in ice-cold acetone for 10 min followed by washes for 3 x 10 min in 0.01 M PBS. For 1 h, explants were blocked with 5% goat serum diluted in background reducing agent (DakoCytomation, Carpinteria, CA). Cells were then incubated overnight at 4°C in either M2R rabbit anti-mouse antibody (1:100; Chemicon International, Inc., Temecula, CA) or M3R rabbit anti-human antibody (1:100; recognizes second extracellular loop of receptor, and there is 95% homology to mouse M3R) (US Biological Inc., Swampscott, MA) and CD117 (c-kit) rat anti-mouse antibody (1:200; Cedarlane Laboratories Ltd.). Explants were washed 3 x 20 min, incubated with the secondary antibodies, Cy5 goat anti-rabbit IgG (1:100; Jackson ImmunoResearch Laboratories, Inc., West Grove, PA) and Cy2 goat anti-rat IgG (1:50; Jackson Immuno-Research Laboratories, Inc.), and washed 3 x 20 min before mounting on a glass slide.
Frozen Murine Jejunal Sections. Adult CD-1 mice were sacrificed in accordance with procedures approved by the Animal Ethics Committee at McMaster University. A midline incision was made, and the jejunum was removed a placed in oxygenated Krebs solution. The tissue was flushed several times with Krebs solution to remove luminal contents. The tissue was transferred to a separate Petri dish containing 0.01 M PBS, where it was then flushed with PBS and sectioned transversally into 1- to 2-cm pieces. Pieces were then embedded in optimal cutting temperature compound (Sakura Finetek, Torrance, CA) and transferred to liquid nitrogen until frozen. Frozen blocks were sectioned on the cryostat at a thickness of 5 to 9 µM and transferred to glass slides. Slides were kept frozen at –70°C until 2 to 3 h before use.
Slides were fixed in ice-cold acetone for 10 min followed by washes for 3 x 10 min in 0.01 M PBS. For 1 h, tissue containing slides were blocked with 5% goat serum (diluted in background reducing agent; DakoCytomation). Slides were then incubated overnight at 4°C in either M2R rabbit anti-mouse antibody (1:100; Chemicon International, Inc.) or M3R rabbit anti-human antibody (1:100; recognizes second extracellular loop of receptor, and there is 95% homology to mouse M3R) (US Biological Inc.) and CD117 (c-kit) rat anti-mouse antibody (1:200; Cedarlane Laboratories Ltd.). Tissue was washed 3 x 20 min, incubated with the secondary antibodies, Cy3 goat anti-rabbit IgG (1:500; Jackson ImmunoResearch Laboratories, Inc.), and Cy2 goat anti-rat IgG (1:50; Jackson ImmunoResearch Laboratories, Inc.) and washed 3 x 20 min before mounting with a glass coverslip.
Electrophysiology
Cell cultures were visualized using a Nikon Diaphot inverted microscope (Nikon, Mississauga, ON, Canada). Whole-cell recordings were obtained using an Axopatch 200B amplifier (Axon Instruments, Union City, CA), and data were acquired using pClamp 9.2 software (Axon Instruments). Pipettes were fabricated to a resistance of 2 to 4 M
. Only cells with series resistance of <15 M were studied, to which series resistance compensation of 
70 to 75% was employed. Cells were constantly perfused in a solution containing 135 mM NaCl, 5 mM KCl, 2 mM CaCl2, 1 mM MgCl2, 5 mM glucose, 5 mM HEPES, and 0.33 mM NaH2PO4, pH 7.35 with NaOH. Pipettes were filled with an internal solution containing 135 mM Cs aspartate, 5 mM KCl, 1 mM MgCl2, 0.1 mM EGTA, and 5 mM HEPES, pH 7.25 with CsOH.
Drugs
Bethanechol, phorbol 12-myristate 13-acetate (PMA), and atropine were purchased from Sigma-Aldrich (Oakville, ON, Canada), bisindolylmaleimide was from Calbiochem, EMD Biosciences (San Diego, CA), and E4031 was from Alomone Laboratories (Jerusalem, Israel). Stock concentrations of bethanechol and E4031 were formulated in nanopure water. Dimethyl sulfoxide was used to dissolve PMA, and bisindolylmaleimide and ethanol were used to dissolve atropine. Final concentration of dimethyl sulfoxide and ethanol that cells were exposed to was less than 0.001%.
Analysis
Data were analyzed using Clampfit (Union City, CA) and Origin software (Northampton, MA). A log concentration response curve was formulated for bethanechol and fit using Origin software [Y = A1 + (A2–A1)/(1 + 10log(IC50–x) x nH), where A1 and A2 represent minimal and maximal concentrations, respectively, x represents concentration, IC50 represents the concentration at which 50% of the response in inhibited, and nH represents the Hill Slope. A Boltzmann relationship [I/Imax = 1/{1 + [exp(V0.5)V)/k]}, where I represents current and V represents voltage] was used to determine changes to half-maximal voltage of activation in control and bethanechol-treated cells. The number of cells used was represented by n. Data were statistically analyzed using a Student's t test where data with p < 0.05 were considered significant.
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To investigate whether or not the expression of M2R or M3R was an effect of culture, double labeling studies with CD117 antibody were performed on tissue sections from the murine jejunum. Distinct M2R-L immunoreactivity was found in the circular and longitudinal smooth muscle layers as well as the ganglia of the myenteric plexus (Fig. 1B). M2R-L immunoreactivity was not detected on c-kit-positive cells in the myenteric plexus (Fig. 1B). Similarly, M3R-L immunoreactivity was identified throughout the circular and longitudinal muscle layers on smooth muscle cells, myenteric ganglia, and c-kit-positive cells (Fig. 1D). Not all cells exhibiting c-kit immunoreactivity showed M3R-L immunoreactivity. Interestingly, distinct regions of M3R-L immunoreactivity were found on c-kit-positive cells, which were associated with the myenteric ganglia (Fig. 1D, arrows).
Bethanechol Concentration Response. Using ionic conditions employed in a previous study to evoke ERG K+ currents in ICC-AP (McKay et al., 2006), a hyperpolarizing voltage protocol was applied from a holding potential of 0 mV, stepping in 10-mV increments from –120 to +20 mV. This resulted in the current profile and current-voltage relationship illustrated in Fig. 2, A and B, where maximal peak current occurred at a potential of –100 mV. In our previous study, the ERG K+ channel-specific blocker, E4031, inhibited hyperpolarization-evoked current at –100 mV by 80% (McKay et al., 2006). To determine whether the muscarinic agonist, bethanechol, modulated the ERG K+ current in ICC, a concentration-response relationship was established by adding increasing concentrations of bethanechol (10–8 to 10–4 M) to the preparation (Fig. 2C; n = 3–7) while measuring the current amplitude evoked by stepping from 0 to –100 mV. Using a concentration-response fit, half-maximal inhibition was measured at 3.4 µM. In seven cells, inhibition of 37 ± 4% was achieved using 100 µM bethanechol.
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The time constant of recovery from inactivation proved difficult to measure because of its rapid time course and interference from capacitive transients. Deactivation was best fit using double exponentials. The fast component of deactivation was increased at all measured potentials following the addition of bethanechol (Fig. 4C, n = 6). The slow component of the deactivation time constant was increased by bethanechol at potentials between –50 and –100 mV and unaffected by bethanechol at potentials of –30 and –40 mV (Fig. 4D, n = 6). Compared with control conditions, the process of deactivation was inhibited when cells were hyperpolarized from 0 to –50 and –40 mV, resulting in markedly increased steady-state currents.
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To ensure that modulation of the current was not due to current run down, cells were pulsed with a hyperpolarizing protocol every 2 min for 46 min. Comparisons were made with the resultant current over time to the initial recoding. Under the conditions of this study, the current measured by hyperpolarizing from 0 to –100 mV decreased by 4.2 ± 3.3% at the 46-min time point (n = 6).
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). The chosen ionic conditions (with 5 mM K+ inside and out; hence, a K+ equilibrium potential of 0 mV) allowed observation of these currents and produced a significant sustained inward current when cells were depolarized from –60 to –40 and –30 mV. Following the addition of 100 µM bethanechol, the average value of the current amplitude was increased at –50 mV by 26.1 ± 14.7% and significantly increased at –40 mV by 58.7 ± 9.7% (Fig. 5B, n = 6, p < 0.05). Conversely, outward current between 0 and +20 mV was significantly decreased (Fig. 5B). Bethanechol reduced tail currents at –60 mV, evoked following depolarizing pulses from –60 up to 20 mV (Fig. 5C). Tail currents from –20 to –60 mV were reduced by 23.9 ± 5.7% (p < 0.01, n = 6) and from 20 to –60 mV by 30.6 ± 6.5% (p < 0.001, n = 6). All changes in current amplitude evoked by bethanechol treatment were completely prevented by pretreatment with atropine (5 x 10–6 M) (n = 2) and 2 µM E4031 applied following bethanechol treatment consistently inhibited steady-state currents at –50 and –40 mV by 66.8 ± 8.1% and 72.7 ± 12% and tail currents evoked from a prepulse potential of 20 mV by 81 ± 5.7% (Fig. 5, B, inset, and C, inset; n = 3). The time constant of activation was best fitted using single exponentials under control conditions and following treatment with bethanechol. The time constant of activation in the presence of bethanechol exhibited no change from control except consistently at a potential of –40 mV (Fig. 6). At –40 mV, a significant increase from 48.7 ± 7.1 to 80.3 ± 6.8 ms was observed (n = 6, p < 0.05).
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In comparison with treatment with bethanechol alone (Fig. 4C), PKC activation did not affect the fast time constant of deactivation between –120 and –60 mV but did slow the fast time constant of deactivation between –50 and –30 mV (Fig. 9A; n = 4, p < 0.05). The slow time constant of deactivation was increased at voltages between –80 and –50 mV, unchanged at –40 mV, and decreased at –30 mV (Fig. 9B; n = 4, p < 0.05). In four cells, incubation with bisindolylmaleimide prevented the bethanechol-induced modulation of the fast and slow time constant of deactivation (Fig. 9, C and D; n = 4).
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The effect of PKC activation and PKC inhibition was also examined on the depolarization-evoked ERG K+ current. Cells treated with PMA showed no change in the amplitude of inward current between potentials of –60 and –10 mV; however, outward current between 0 and +20 mV was significantly decreased (Fig. 10A; n = 4, p < 0.05). Tail current was also significantly decreased at prepulse potentials between –20 and +20 mV (Fig. 10B; n = 4, p < 0.05). Conversely, inhibition of PKC with bisindolylmaleimide prevented the increase in inward current at –50 and –40 mV evoked by bethanechol (Fig. 10C). Tail current reduction by bethanechol was diminished by PKC inhibition. In cells pretreated with bisindolylmaleimide, tail current evoked from a prepotential of +20 mV decreased 14.6 ± 4.1% with bethanechol (Fig. 10D; n = 4, p < 0.05), compared with cells treated with bethanechol alone, which were reduced by 30.6 ± 6.5% (n = 6, see Fig. 5B).
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Activation of PKC with PMA significantly increased the time constant of activation at –40 mV from 25.9 ± 5.1 to 62.7 ± 4.5 ms (n = 4, p < 0.01), without influencing the time constant of activation at all other voltages studied (Fig. 11A). Inhibition of PKC diminished the effect of bethanechol on the time constant of activation at –40 mV. An increase of 15.7 ± 6.1 ms was achieved after treatment of bethanechol on four cells pretreated with bisindolylmaleimide (Fig. 11B), compared with 38.8 ± 9.8 ms in six cells treated with bethanechol alone (see Fig. 6).
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Discussion |
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). Hence, it is logical to assume that under normal conditions, when the ICC depolarizes in response to inward currents through pacemaker channels (Farrugia, 1999; Huizinga et al., 2002; Koh et al., 2002; Sanders et al., 2006), the height and duration of the slow-wave depolarization is determined in part by depolarization- and hyperpolarization-evoked ERG K+ currents. We have recently documented ERG K+ currents in ICC that were evoked by depolarization from a resting membrane potential of –60 mV (McKay et al., 2006). These studies were carried out using high intracellular Cs+ and equimolar K+. These same conditions were used in the present study to investigate the effects of muscarinic receptor stimulation so that direct comparisons could be made.
The present study shows that the ERG K+ channel gating in ICC-AP is modulated by activation of a muscarinic G-protein-coupled receptor (probably the M3R), through a PKC-dependent mechanism. The major finding was that steady-state currents between the potentials of –55 and –35 mV, evoked either by depolarization or hyperpolarization, were increased by muscarinic stimulation. It appears that in the presence of bethanechol, the processes of inactivation (upon depolarization) and deactivation (following hyperpolarization) were inhibited within this range of voltages leading to increased steady-state currents. A contributing factor is probably the leftward shift in half-maximal voltage of activation and, therefore, a greater likelihood of channels being open around –40 mV. These findings are not consistent with data from expressed HERG channels, suggesting that they may be unique to the native ICC channel.
Hyperpolarization following depolarization of ICC evokes rapid recovery from inactivation of ERG channels, resulting in large peak inward currents under our experimental conditions followed by partial fast deactivation leading to sustained currents. The peak currents were inhibited by bethanechol, and the time constants of both the fast and slow components of deactivation increased. This decrease in hyperpolarization-evoked peak current is consistent with previous reports in the literature regarding the effect of G-protein-coupled receptor stimulation on ERG K+ currents (Kiehn et al., 1998; Cui et al., 2000; Schledermann et al., 2001; Thomas et al., 2003; Hirdes et al., 2004).
Depolarization-evoked ERG K+ currents at potentials positive to 0 mV were decreased in the presence of bethanechol. Kiehn et al. (1998) attributed a decrease in depolarization-evoked HERG K+ current to a rightward shift in the activation voltage; hence, a lower likelihood of the channel being opened. We observed a marked decrease in channel availability at depolarized potentials.
The muscarinic receptor, M3R was identified on ICC-AP in both neonatal murine explant cultures and adult murine jejunum. In tissue, M3R immunoreactivity was detected on ICC-AP that were in close association with the myenteric plexus. It is possible but entirely speculative that the receptor is only expressed on ICC that are in close approximation to cholinergic nerves and therefore responding to cholinergic stimulation by expression of receptors. In cultures, M3R immunoreactivity was detected in some but not all c-kit-positive cells. The c-kit immunoreactive cells surrounding the explant were M3R immunoreactive, and these were the cells recorded from. We did not find M2R immunoreactivity on ICC-AP of the murine jejunum. This finding is consistent with a recent study by Iino and Nojyo (2006), which did not detect M2R immunoreactivity in the ICC-AP of the guinea pig small intestine (Iino and Nojyo, 2006).
The unique features of muscarinic modulation of ERG K+ currents in ICC may be due to the fact that we recorded from native cells, whereas most if not all reported studies on muscarinic regulation in the literature have been conducted using coexpression of both G-protein-coupled receptors and HERG/ERG K+ channels in expression systems (e.g., Xenopus oocytes, rat GH3/B6, Chinese hamster ovary, human embryonic kidney 293, human tsA-201) (Barros et al., 1998; Kiehn et al., 1998; Cui et al., 2000; Bian et al., 2001; Schledermann et al., 2001; Thomas et al., 2003, 2004; Hirdes et al., 2004). Another possible explanation is that in ICC, the ERG K+ channel may be coexpressed with -subunits (minK or MiRP1), which have been found to coexpress with HERG K+ channels in native tissue and have been shown to influence HERG K+ channel properties (McDonald et al., 1997; Abbott et al., 1999; Weerapura et al., 2002). A study by Cui et al. (2000) looked at the modulatory effect of cAMP on HERG channels coexpressed with either MiRP1 or minK. Compared with cAMP modulation of HERG channels expressed alone that produced a rightward shift in voltage-dependent activation and decreased current amplitude, cAMP produced a leftward, hyperpolarizing shift in voltage-dependent activation and both increased or did not change current amplitude in cells coexpressing HERG and MiRP1 or minK (Cui et al., 2000). It is possible that ERG K+ channels in ICC are coupled with subunits and therefore show similarities in channel modulation to the expression systems coexpressing both HERG and a subunit. Potentially, ERG1b and ERG3 could form the heteromultimeric channel. Interestingly, the inactivation for the expressed ERG3 subunit is slower than ERG1 and ERG2 and is also slower than the activation of ERG3, therefore creating substantial noninactivating steady-state current (Shi et al., 1997). In addition, Hirdes et al. (2005) and Wimmers et al. (2002) have shown that the kinetics of heteromultimeric ERG channels differ from homomultimeric ERG channels. It will be important to determine subunit identities for the ERG channel in ICC, which is the current focus of our laboratory.
The present study demonstrates that ERG K+ channel modulation in ICC-AP through MR activation is PKC-dependent. Barros et al. (1998) and Thomas et al. (2003, 2004) have reported HERG K+ current modulation via a PKC-dependent mechanism based on experiments using PKC activation and inhibition in Xenopus oocytes expressing HERG K+ channel protein. Although it was hypothesized that PKC modulated HERG K+ channels by phosphorylation of PKC-dependent phosphorylation sites, deletion of these sites by mutagenesis did not prevent the response to G-protein receptor agonists or PKC activators (Thomas et al., 2003, 2004). To date, the phosphorylation-independent mechanism responsible for PKC dependent modulation remains unknown.
PKC activation mimicked and PKC inhibition diminished or abolished the modulatory action of bethanechol. However, inhibition of PKC before bethanechol addition did not completely inhibit the effect of bethanechol. This may be due to incomplete inhibition of PKC or modulatory actions of bethanechol that are not sensitive to PKC inhibition such as other second messengers involved in M3R activation (i.e., PIP2). Although the effect of PIP2 on channel modulation was not investigated in the present study, Bian et al. (2001) and Wang et al. (2004) have reported a leftward shift of activation when PIP2 was applied to the pipette, which was prevented when a PIP2 antibody was used.
To extrapolate effects of MR-induced ERG K+ current modulation observed under the present experimental conditions to in vivo small intestinal pacemaking is arduous due the multitude of effects that downstream signaling can have on other ion channels in the ICC, on the biochemical intracellular pacemaking mechanism itself, as well as on smooth muscle cells. We postulate that upon the release of acetylcholine from the enteric cholinergic neurons and stimulation of the M3R on ICC, ERG K+ current activation is increased due to increased steady-state currents and the leftward shift in half-maximal activation voltage. The increased ERG K+ current generated when ICC depolarize through slow-wave activity from –60 to –40 mV would cause an increase in potassium leaving the cell (due to its electrochemical gradient) and result in hyperpolarization of the ICC. Hence, muscarinic stimulation would decrease ICC excitability if the ERG channels were the only target on ICC. The overall effect of cholinergic stimulation of the intestinal smooth musculature appears to be excitatory, resulting in prolonged slow waves when measured in smooth muscle cells (Huizinga et al., 1984; Sanders and Smith, 1986; Seerden et al., 2005). Consistently, in barium follow-through studies in rats, those receiving galantamine (a compound that inhibits acetylcholinesterase) had decreased intestinal barium transit time and increased smooth muscle contractility (Turiiski et al., 2004). Further studies on muscarinic regulation of other ion channels in ICC and smooth muscle cells are needed to resolve the complex actions of cholinergic receptor stimulation.
In summary, the present study shows that the ERG K+ channel in ICC is affected by stimulation of muscarinic receptors, probably the M3 receptor, via a PKC-dependent mechanism. Modulation of the ERG K+ current in ICC-AP will affect the kinetics of pacemaking in the intestinal musculature.
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Acknowledgements |
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Footnotes |
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Article, publication date, and citation information can be found at http://jpet.aspetjournals.org.
ABBREVIATIONS: ICC-AP, -MP, or -MY, interstitial cells of Cajal-Auerbach's plexus or myenteric plexus; ERG, ether-a-go-go-related gene; PK, protein kinase; PIP2, phosphatidylinositol bisphosphate; M2R, M2 receptor; M3R, M3 receptor; PBS, phosphate-buffered saline; PMA, phorbol 12-myristate 13-acetate; M2R-L, M2 receptor-like; M3R-L, M3 receptor-like; HERG, human ether-a-go-go-related gene; E4031, 1-[2-(6-methyl-2-pyridyl)-ethyl-4-(methylsulfonylaminobenzoyl)piperidine.
Address correspondence to: Dr. Jan D. Huizinga, McMaster University, HSC-3N5C, 1200 Main Street West, Hamilton, ON L8N 3Z5, Canada. E-mail: huizinga{at}mcmaster.ca
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) and after 100 µM bethanechol (
). Inset, effect of 2 µM E4031 (
) on bethanechol (
) under control conditions (
