Vagal inhibition in the antral region of guinea pig stomach

Emma J. Dickens, F. R. Edwards, and G. D. S. Hirst

Department of Zoology, University of Melbourne, Parkville, Victoria 3052, Australia


    ABSTRACT
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ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
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The effects of vagal stimulation in the presence of a muscarinic antagonist were examined on three distinct rhythmically active cells located in guinea pig antrum. Vagal stimulation inhibited contractions of the circular muscle layer but did not change their rate of occurrence. With the use of intracellular recording techniques, these stimuli were found to initiate inhibitory junction potentials in the circular layer but produced smaller potential changes in driving and follower cells. Inhibition of the circular muscle layer involved two separate components. The dominant component was independent of changes in membrane potential and was abolished by nitro-L-arginine. After abolishing Ca2+ entry into smooth muscle cells with a Ca2+ antagonist, vagal stimulation continued to inhibit the residual contractions associated with each slow wave. When the cyclic changes in intracellular Ca2+ concentration associated with each slow wave were measured, they were found to be unchanged by vagal stimulation. The observations suggest that vagal inhibition of stomach movements does not alter pacemaker activity in the stomach; rather, it results from a change in the sensitivity of smooth muscle contractile proteins to Ca2+.

smooth muscle; slow waves; driving cells; inhibition


    INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

MANY REGIONS of the gastrointestinal tract generate slow waves and contract rhythmically at low frequencies in the absence of stimulation (26, 33). Slow waves appear to result from the interaction between two distinct groups of cells: activity either originates in or is coordinated by interstitial cells of Cajal (ICC), with smooth muscle cells responding by generating complete slow waves (27). Several observations give rise to this view. ICC isolated from mouse gastrointestinal tract generated waves of electrical activity with time courses similar to slow waves recorded in situ (22). Intestinal preparations taken from mutant mice that lack myenteric ICC failed to generate slow waves, although they can generate action potentials (20, 35). Similar findings were obtained from preparations in which the development of ICC was impaired with an antibody to c-kit, a component of ICC (25, 34, 37).

When recordings were made from the guinea pig stomach, three different types of rhythmically active cells were identified (15). One type, called driving cells, were identified as being ICC lying in the myenteric region of guinea pig antrum (ICC-MY). They generated large-amplitude, long-lasting, depolarizing membrane potential changes, called driving potentials. Driving potentials spread passively to the other two cell types, muscle cells lying in either the longitudinal or the circular muscle layers. In the circular layer (slow wave cells), the depolarization triggered a secondary regenerative component that amplifies the potential change (15). Depolarizing isolated bundles of circular muscle (31) triggered similar regenerative responses that lasted for several seconds. They appear to involve the release of internal Ca2+, following the production of a second messenger rather than activation of conventional voltage-dependent ion channels (16). Sufficient Ca2+ is released from intracellular stores during each regenerative component to trigger a small contraction, but the major source of Ca2+ for contraction results from Ca2+ entry via L-type Ca2+ channels that are activated during the secondary component of the slow wave (31). In the longitudinal layer (follower cells), the depolarization fails to trigger a regenerative component (15).

Mechanical activity of the stomach wall, initiated by ICC, is modulated by neural inputs running in the vagus. Vagal inputs to the stomach contain preganglionic axons; some innervate intrinsic excitatory motor neurons lying in the myenteric plexus and others innervate intrinsic inhibitory neurons (28). This report is concerned with the vagal inhibitory projection to the antrum. This pathway is activated by stomach distension, which causes inhibition of muscular activity leading to gastric accommodation (13). Inhibition is mediated by the release of two inhibitory transmitter substances. One transmitter evokes a nonadrenergic, noncholinergic (NANC) inhibitory response (9) that is associated with an inhibitory junction potential (IJP) (2, 23). This transmitter is not nitric oxide (NO) but has been suggested to be either ATP or the neuropeptide vasoactive intestinal polypeptide, which is found in many intrinsic inhibitory nerves (17). The other transmitter is probably NO, and this may be the dominant transmitter in the stomach (14). Recently it has been shown that in several regions of the gastrointestinal tract inhibitory fibers predominantly innervate intramuscular ICC (ICC-IM), where they cause the generation of the second messengers responsible for inhibition of muscular activity (7, 38). Unfortunately, to date it has not been possible to identify ICC-IM with the use of electrophysiological techniques. Presumably they form part of an electrical syncytium with neighboring smooth muscle cells, and although they may have specialized roles they cannot be distinguished on electrical grounds from nearby smooth muscle cells. This study has reexamined the responses to vagal stimulation in the guinea pig stomach, taking into account the three different electrophysiologically identifiable types of rhythmically active cells present in the stomach wall. The experiments suggest that inhibitory responses were largely confined to the circular muscle layer in which NO appeared to reduce the sensitivity of contractile proteins to increases in intracellular Ca2+ concentration ([Ca2+]i).


    METHODS
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INTRODUCTION
METHODS
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The procedures described have been approved by the animal experimentation ethics committee at the University of Melbourne. Guinea pigs of either sex were stunned and exsanguinated, and their stomachs, along with the vagal nerve trunks to the posterior side of the stomach, were removed. The antrum was removed, taking care to maintain the integrity of the vagal trunk, and immersed in oxygenated physiological saline (in mM: 120 NaCl, 25 NaHCO3, 1.0 NaH2PO4, 5 KCl, 2 MgCl2, 2.5 CaCl2, and 11 glucose) bubbled with 95% O2-5% CO2. The mucosa was dissected away, and the preparations were pinned out, serosal surface uppermost, in a recording chamber whose base consisted of a microscope coverslip coated with Sylgard silicone resin (Dow Corning, Midland, MI).

A region of antrum was immobilized with fine pins, and the serosa was carefully removed under a dissecting microscope. A force transducer was attached to the cut end of the circular muscle bundles at the greater curvature. We were unable to determine whether the mechanical activity of the longitudinal layer was changed by vagal stimulation. In this region of the stomach, the longitudinal layer is thin compared with the circular layer, and as a result tension recordings were dominated by mechanical activity from the circular layer. Preparations were viewed with an inverted compound microscope, and intracellular recordings were made using sharp microelectrodes (90-150 MOmega ) filled with 0.5 M KCl. Signals were amplified with an Axoclamp-2A amplifier (Axon Instruments, Foster City, CA), low-pass filtered (cutoff frequency: 1 kHz), digitized, and stored on a computer for later analysis.

Preparations were constantly perfused with physiological saline solution warmed to 35°C containing hyoscine (1 µM), which abolished the effects of vagally activated excitatory motor neurons. In each experiment, the vagus was drawn into a suction electrode for stimulation. The preparation was stimulated with a train of eight stimuli delivered at 10 Hz, and the stimulus strength increased until a maximal inhibitory mechanical response was detected. The stimulus intensity was increased by a further 20% and left unchanged throughout the experiment. It was assumed that this procedure activated all of the vagal preganglionic fibers present.

To examine a range of likely physiological situations, short bursts of high-frequency stimuli (1-8 impulses at 10 Hz) and longer trains of stimuli (train duration 50 s) were applied. In preliminary experiments using long trains of stimuli, it was found that the inhibitory responses were little changed when the stimulation frequency was increased to >2 Hz. Therefore, when long trains of stimuli were applied the responses produced by a stimulation frequency of 1 and 2 Hz were analyzed. Those produced by long trains of higher-frequency stimuli were poorly sustained, and the responses readily fatigued.

In some experiments, changes in [Ca2+]i were determined as described previously (4). In brief, preparations were loaded with fura PE3 and subsequently illuminated with light of two wavelengths (340 and 380 nm), alternating at a frequency of 12 Hz. In initial experiments, this switching rate was found to allow an accurate measure of the time course of the change in [Ca2+]i associated with each slow wave. Photons emitted at a wavelength of 510 nm were counted during each period of illumination to give separate measures of the concentrations of the fura-Ca2+ complex and free fura. The time courses of changes in [Ca2+]i were obtained by taking a ratio of these values; no attempt has been made to correlate the ratio values with absolute changes in [Ca2+]i. Maximal signals were obtained when the microscope was focused on the central plane of the bundles of circular smooth muscle. The area of illumination was restricted to a 100-µm2 window, with the recording electrode placed in the center of this square. As such, [Ca2+]i changes reflected the changes occurring in the same bundle of muscle as that from which the membrane potential recording was being made. Since the circular layer constituted the thickest layer present in the stomach wall, ~90% of the total wall thickness, the signals were dominated by emissions from the circular layer.

All data are expressed as means ± SE. The Student's t-test was used to determine whether data sets differed significantly.

Drugs used in this study were nifedipine, hyoscine, nitro-L-arginine (L-NNA), apamin, tetrodotoxin, and guanethidine (Sigma Chemical, St. Louis, MO). Preparations were loaded using fura PE3-AM (Calbiochem) and pluronic F-127 (Sigma Chemical).


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ABSTRACT
INTRODUCTION
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DISCUSSION
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General observations. When intracellular recordings were made from the antral region of the stomach, three distinct patterns of membrane potential change were associated with each contraction of the circular muscle layer (Fig. 1). The first cells encountered generated small-amplitude rhythmical depolarizations (follower potentials), which usually triggered bursts of action potentials (Fig. 1A). The action potentials were abolished by adding nifedipine (1 µM) to the physiological saline, with the resulting follower potentials having time courses and amplitudes similar to those identified as originating in the longitudinal layer (15). If the electrode was advanced through this layer, cells generating driving potentials (waves of depolarization with peak amplitudes in excess of 40 mV and rapid rising phases like those recorded from ICC-MY) were occasionally detected (Fig. 1B) (15). Further downward movement invariably led to the impalement of a cell that generated a slow wave like those recorded from smooth muscle cells lying in the circular layer (Fig. 1C) (15).


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Fig. 1.   Recordings from the 3 different types of rhythmically active cells present in the wall of the antral region of guinea pig stomach. A: successive follower potentials (top trace) recorded from the outer layer of the guinea pig antrum along with a simultaneous recording of the rhythmical contractions of the circular layer (bottom trace). The resting membrane potential was -62 mV. B: successive driving potentials (top trace) recorded from a driving cell and a simultaneous recording of contractions of the circular layer (bottom trace). The resting membrane potential was -61 mV. C: successive slow waves (top trace) recorded from the circular layer and simultaneous recording of the rhythmical contractions of this layer (bottom trace). The resting membrane potential of the cell was -64 mV. The voltage calibration bar applies to each membrane potential recording; the force calibration bar applies to each contraction record. The time calibration bar refers to all recordings.

Since no attempt was made in this study to histologically identify the three different types of cells present, they are referred to here on the basis of their electrophysiological characteristics, that is, follower cells, driving cells, and slow wave cells (15). Follower cells had peak negative membrane potentials in the range of -59 to -68 mV (-62.3 ± 0.9 mV; n = 12; each n value represents a measurement from a preparation taken from a separate animal). The peak amplitude of follower potentials, including action potentials, was ~25-55 mV. After blocking L-type Ca2+ channels with nifedipine (1 µM), follower potentials had peak amplitudes in the range of 16.0 to 27.6 mV (22.4 ± 1.7 mV; n = 6). Driving cells had peak negative membrane potentials in the range of -59 to -67 mV (-63.5 ± 0.9 mV; n = 8) and discharged driving potentials with amplitudes in the range of 39.9-57.1 mV (44.4 ± 2.0 mV; n = 8). In nifedipine (1 µM), driving potentials had peak amplitudes in the range of 39.0-52.3 mV (43.8 ± 2.4 mV; n = 6). Slow wave cells had negative membrane potentials in the range of -59 to -71 mV (-64.8 ± 1.8 mV; n = 12). They discharged slow waves, which consisted of an initial depolarization and a secondary regenerative component, at frequencies of 2.5-4.5 waves/min (2.8 ± 0.4 waves/min; n = 8) with peak amplitudes in the range of 20.0-42.2 mV (32.3 ± 2.7 mV; n = 12). In a few preparations (5 of 56), slow waves recorded from the circular layer, like those recorded from follower cells, triggered a discharge of action potentials (see, for example, Fig. 8A). These action potentials were also abolished by nifedipine (1 µM) to reveal conventional slow waves (see Fig. 8B). In all preparations, nifedipine (1 µM) had little effect on the amplitudes of slow waves, which did not trigger discharges of action potentials but dramatically reduced the amplitudes of the contractions associated with each slow wave (31). These observations confirm that three distinct patterns of electrical activity can be identified in the wall of the antrum (15).

Effect of vagal stimulation on electrical activity recorded from follower cells of guinea pig antrum. In all experiments, the muscarinic receptor antagonist hyoscine (1 µM) was added to the physiological saline to block the effects of stimulating vagal excitatory pathways. To ensure that the vagus was being adequately stimulated (see METHODS), simultaneous measurements of the contractile activity of the circular layer were made. Vagal stimulation produced only small hyperpolarizing membrane potential changes in follower cells (Fig. 2). When brief trains of high-frequency stimuli (8 stimuli at 10 Hz) were delivered in the interval between follower potentials, they triggered hyperpolarizations with peak amplitudes of 0.3-1.0 mV (0.8 ± 0.1 mV; n = 5; Fig. 2, A and C). When delivered during the discharge of action potentials at the peak of a follower potential, such trains of stimuli failed to change their rate of discharge and a potential change could not be detected (Fig. 2, B and D). The discharge of action potentials was abolished by adding nifedipine (1 µM) without a detectable change in peak negative potential. The same trains of vagal stimuli (8 at 10 Hz) produced hyperpolarizations of 0.9-2.7 mV (1.8 ± 0.3 mV; n = 5) when delivered during the interval between follower potentials; the values before and after nifedipine were significantly different (P < 0.05). We have no explanation for this difference. When delivered during the plateau of the follower potentials, they initiated similar hyperpolarizations to those detected during the quiescent period, with peak amplitudes of 0.1-2.5 mV (1.1 ± 0.5 mV; n = 5). Changes in membrane potential were not detected in either control solutions or in solutions containing nifedipine (1 µM) when long trains of vagal stimuli (1 or 2 Hz for 50 s) were delivered. Although these trains of stimuli produced marked changes in the amplitudes of the contractions produced by the circular muscle layer both in control (Fig. 2E) and in nifedipine-containing solutions, the rate of occurrence of follower potentials was not consistently changed.


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Fig. 2.   Effect of vagal stimulation on follower potentials recorded from guinea pig antrum. A and B: recordings of follower potentials and associated contractions of the circular layer. Trains of vagal stimuli (; 8 at 10 Hz) were applied before (A) and during (B) a follower potential. The trains of stimuli inhibited the force of circular muscle contraction associated with each follower potential (A, bottom trace) or reduced the basal tension (B, bottom trace). C and D: membrane potential changes are displayed at faster scan speeds. When the stimulus was presented between follower potentials, a small hyperpolarization was detected (C). When presented during the peak of a follower potential, no change in the discharge of muscle action potentials could be detected (D). E: effects of applying a long train of stimuli (2 Hz for 50 s) to the vagus. The discharge of follower potentials was unchanged, but the contractions of the circular layer were inhibited. All recordings were made from the same cell, with a peak negative potential of -63 mV.

Effect of vagal stimulation on driving potentials recorded from guinea pig antrum. The inhibitory effects of vagal stimulation on the driving cells were recorded in solutions with or without added nifedipine (1 µM). Again, simultaneous measurements of the contractile activity of the circular layer were recorded. Brief trains of stimuli (1-8 stimuli at 10 Hz) produced only small-amplitude hyperpolarizations in driving cells (Fig. 3). A train of stimuli (8 at 10 Hz) delivered in the period between driving potentials caused hyperpolarizations of 1.5-2.5 mV (2.0 ± 0.2 mV; n = 4; Fig. 3, A and C). Identical trains of stimuli, delivered during the plateau phases of the driving potentials, initiated hyperpolarizations of 0-2.3 mV (1.2 ± 0.5 mV; n = 4; Fig. 3, B and D). Changes in the time courses and amplitudes of driving potentials were not detected if long trains of vagal stimuli (1 or 2 Hz for 50 s) were delivered. These trains of stimuli had no consistent effect on their rate of occurrence. In some preparations the rate of occurrence increased slightly; in others the rate fell. Again, these stimuli produced marked changes in the amplitudes of the contractions produced by the circular muscle layer (Fig. 3E).


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Fig. 3.   Effect of vagal stimulation on driving potentials recorded from the guinea pig antrum. A and B: recordings of driving potentials and associated contractions of the circular layer. Trains of vagal stimuli (; 8 at 10 Hz) were applied before (A) and during the plateau (B) of a driving potential. The trains of stimuli inhibited the force of contraction associated with each driving potential (bottom traces). C and D: membrane potential changes, displayed at faster scan speeds. When presented between driving potentials, the stimuli initiated a small hyperpolarization (C); when presented during the plateau, a hyperpolarization could not be detected (D). E: long trains of vagal stimuli (2 Hz for 50 s) failed to alter the discharge of driving potentials but inhibited the contractions produced by the circular layer. Recordings shown in A, B, C, and D were obtained from the same cell; the peak negative potential was -60 mV. The peak negative potential of the cell shown in E was -65 mV.

Similar experiments were carried out on a further four preparations in the presence of nifedipine. Nifedipine (1 µM) had no effect on the amplitudes or rates of rise of individual driving potentials [control amplitude 44.3 ± 3.0 mV, maximal rate of voltage change (dV/dtmax) 0.48 ± 0.05 V/s; amplitude in nifedipine 43.4 ± 2.0 mV, dV/dtmax 0.47 ± 0.1 V/s; n = 6]. In physiological saline containing nifedipine, a train of vagal stimuli (8 at 10 Hz) applied during the interval between driving potentials initiated hyperpolarizations with a mean peak amplitude of 1.1 ± 0.3 mV (n = 4). When delivered during the plateau phases of driving potentials, such trains of stimuli initiated hyperpolarizations of 1.5 ± 0.4 mV (n = 4). Again, longer trains of stimuli failed to change the properties of driving potentials. These observations suggest that vagal inhibitory inputs are unable to modify the discharge of driving potentials.

Effect of vagal stimulation on slow waves, changes in [Ca2+]i, and contractile activity recorded from the circular layer of guinea pig antrum. Experiments were carried out either in preparations in which changes in membrane potential and associated contractions were measured simultaneously or in preparations in which changes in membrane potential, changes in [Ca2+]i, and contractile activity were all measured together. To monitor changes in [Ca2+]i, preparations were loaded with fura PE3. Fura-loaded preparations generated slow waves identical to those detected in unloaded preparations. Their slow waves occurred at 2-5/min. They had peak amplitudes in the range of 29.7-41.5 mV (32.9 ± 1.4 mV; n = 8) superimposed on negative potentials in the range of -61 to -66 mV (-63.0 ± 0.6 mV; n = 8). Each slow wave was associated with a long-lasting increase in [Ca2+]i that preceded the associated contraction (Fig. 4).


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Fig. 4.   Effect of vagal stimulation on slow waves and associated changes in intracellular Ca2+ concentration ([Ca2+]i) and force. A: successive slow waves (a), associated changes in [Ca2+]i (b), and contractions (c) recorded simultaneously from the circular layer of guinea pig antrum. When the vagus was stimulated with a long train of stimuli (1 Hz for 50 s) during the period shown by a horizontal bar, slow waves (a) and associated changes in [Ca2+]i (b) were largely unchanged, but the baseline tension and the amplitude of each associated contraction fell (c). The train of stimuli caused an increase in peak negative potential of ~2-3 mV, and the membrane potential change of each slow wave was interrupted by successive inhibitory junction potentials (IJPs; a). B: recordings from the same cell and the responses produced when the vagus was stimulated at a higher frequency (2 Hz for 50 s). Although this train of stimuli produced a more marked hyperpolarization (a) and a more profound inhibition of mechanical activity (c), no changes in either the amplitudes of successive increases in [Ca2+]i or baseline [Ca2+]i were detected (b). The peak negative potential of the smooth muscle cell, in the absence of stimulation, was -62 mV. The voltage calibration, emission ratio, force, and time calibration bars apply to both sets of recordings.

Trains of low-frequency vagal stimulation reduced the amplitude of contraction associated with each slow wave and caused a fall in basal tension (Fig. 4). These inhibitory responses were not associated with detectable changes in either the peak increase in [Ca2+]i associated with each slow wave or the level of [Ca2+]i between each slow wave (Fig. 4). Invariably, the inhibition of mechanical activity was more marked when higher frequencies of stimulation were applied (Fig. 4). When stimulation was delivered at 1 Hz, the baseline tension fell by 0.9 ± 0.2 mN (n = 13); at 2 Hz, the baseline tension fell by 1.4 ± 0.3 mN (n = 13); these values were significantly different (P < 0.01). At 1 Hz, the peak amplitude of individual contractions fell from 4.0 ± 0.6 mN to 2.2 ± 0.5 mN (n = 13; Fig. 4A); at 2 Hz, the peak amplitude of individual contractions fell to 0.9 ± 0.4 mN (n = 13; Fig. 4B). The inhibition produced by the two rates of stimulation was significantly different (P < 0.01). Vagal stimulation produced only moderate but sustained increases in the peak negative potential detected between slow waves. Fifty stimuli delivered at 1 Hz caused a hyperpolarization of 1.7 ± 0.2 mV (n = 13, 8 loaded and 5 unloaded preparations); 100 stimuli delivered at 2 Hz caused a hyperpolarization of 2.9 ± 0.2 mV (n = 13). These values were significantly different (P < 0.01). At the start of each train of stimuli, the peak amplitude of the first slow wave was depressed by ~3-7 mV. This was occasionally associated with a slight increase in interval between slow waves (Figs. 4 and 6). The inhibitory mechanical responses and the periods of sustained membrane hyperpolarization persisted in the presence of guanethidine (10 µM) but were abolished by tetrodotoxin (0.1 µM). Since all experiments were carried out in the presence of hyoscine (1 µM), the inhibitory responses produced by vagal stimulation clearly resulted ultimately from activation of intrinsic NANC neurons (9). The alternative possibility, that the responses resulted from antidromic activation of sensory fibers running the vagus, seems unlikely. Such fibers could be activated, but higher stimulating strengths were required; when trains of such stimuli were applied, they invariably produced long-lasting membrane depolarizations.

The inhibition of contractile activity produced by low-frequency vagal stimulation appeared to result entirely from the release of NO. In the four fura-loaded preparations examined, the vagal inhibition of mechanical activity was abolished ~15-20 min after L-NNA (0.1 mM) had been added to the physiological saline (Fig. 5). L-NNA itself had no effect on [Ca2+]i determined between or during each slow wave; nor did it affect either the amplitude of each slow wave or the amplitude of each associated contraction. Similar observations were made on a further four preparations in which only slow waves and associated contractions were measured.


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Fig. 5.   The effect of nitro-L-arginine (L-NNA) on vagoinhibitory responses of the circular muscle layer. A: small changes in membrane potential produced by vagal stimulation (2 Hz for 50 s; a), lack of changes in [Ca2+]i associated with each slow wave (b), and inhibition of contraction (c) recorded from the circular layer of guinea pig antrum are shown. B: effect of an identical train of stimuli, recorded 20 min after adding L-NNA (0.1 mM) to the physiological saline. It can be seen that, although vagal stimulation continues to produce small changes in membrane potential (a), the inhibition of mechanical activity was blocked (c). All recordings were obtained from the same cell; the peak negative potential, in the absence of stimulation, was -60 mV. The voltage calibration, emission ratio, force and time calibration bars apply to both sets of recordings.

It has previously been shown that vagal inhibitory responses in the stomach consist of a nitrergic and an apamin-sensitive component (14). Our experiments suggest that the apamin-sensitive component of the response to vagal stimulation plays little part in the vagal inhibition of mechanical activity of the stomach. This point was examined in four preparations that were bathed in apamin-containing (0.1 µM) solutions. The small membrane potential changes triggered by vagal stimulation were considerably reduced by apamin, but the inhibition of contractile activity was barely affected (Fig. 6, A and B). In control solution, the peak negative potential increased by 3.2 ± 0.6 mV (n = 4) during a 50-s train of impulses delivered at 2 Hz. In the presence of apamin, similar trains of stimuli increased the peak negative potential by 1.7 ± 0.4 mV (n = 4). The two values were significantly different (P < 0.05). In each experiment, both the vagal inhibition of mechanical activity and the apamin-resistant hyperpolarization were abolished by the subsequent application of L-NNA (0.1 mM; Fig. 6C).


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Fig. 6.   Effect of apamin and L-NNA on responses to vagal stimulation recorded from a smooth muscle cell lying in the circular layer of guinea pig antrum. A: simultaneous recordings of slow waves (top) and associated contractions (bottom) before, during, and after applying a train of stimuli (2 Hz for 50 s) to the vagus. Shortly after starting the train of stimuli, the amplitude of the first slow wave was reduced and there was a sustained inhibition in the amplitude of associated contractions. B: after adding apamin (0.1 µM) to the physiological saline, vagal stimulation failed to change the shape of slow waves, whereas the inhibition of contractile activity was largely maintained. C: further addition of L-NNA (0.1 mM) abolished the remaining inhibitory effects of vagal stimulation. The time, force, and voltage scale bars apply to all traces.

Brief trains of stimuli (1-8 impulses delivered at 10 Hz) initiated IJPs during the intervals between slow waves and during their secondary phases (Fig. 7). The amplitudes of IJPs increased with the number of stimuli presented, usually to a maximum when ~6-8 stimuli were presented. IJPs initiated between slow waves by a train of eight stimuli delivered at 10 Hz had peak amplitudes of 1.4-5.6 mV (4.1 ± 0.4 mV; n = 13). These stimuli consistently reduced the amplitude of the contraction associated with the following slow wave. The amplitudes of IJPs increased when the stimuli were presented during a slow wave (compare Fig. 7, A and G with Fig. 7, B and H). Thus trains of eight stimuli delivered at 10 Hz during the peak of a slow wave initiated IJPs with amplitudes of 9.6-20.0 mV (14.8 ± 1.0 mV; n = 13). These IJPs also had longer durations than did those initiated between slow waves (Fig. 7, G and H). If the train of stimuli was presented during the later part of the slow wave, the regenerative component was truncated and the associated contraction was much reduced (see Fig. 9). IJPs initiated in the period between slow waves were greatly reduced in amplitude by apamin (control amplitude: 4.6 ± 0.7 mV; amplitude in apamin: 0.5 ± 0.1 mV, n = 4; Fig. 7G; P < 0.01), the inhibitory effect of vagal stimulation on contractile activity was little changed (Fig. 7, A and C). The subsequent addition of L-NNA (0.1 mM) abolished the remaining responses produced by vagal stimulation (Fig. 7E). When IJPs were initiated during a slow wave, apamin reduced their amplitudes but was less effective than on IJPs recorded between slow waves. Their amplitudes fell from 16.0 ± 1.8 mV to 12.3 ± 1.9 mV (n = 4; Fig. 7H; P < 0.05). Again, apamin had little ability to block the inhibitory effects of vagal stimulation on contraction. These were abolished by L-NNA (0.1 mM), and at the same time any remaining apamin-resistant components of the IJP were abolished (Fig. 7H).


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Fig. 7.   Effect of apamin and L-NNA on responses produced by brief trains of vagal stimuli delivered before or during a slow wave. Traces at left show responses to trains of vagal stimuli (; 8 at 10 Hz) presented in the intervals between slow waves and recorded in control solution (A), in a solution containing apamin (0.1 µM; C), and in a solution containing both L-NNA (0.1 mM) and apamin (0.1 µM; E). The time courses of the evoked membrane potential changes are overlaid in G. It can be seen that apamin reduced the amplitude of the IJP but that the inhibitory mechanical responses were little changed (C and G, b). L-NNA abolished the inhibition of mechanical responses (E). At right, responses to trains of vagal stimuli (8 at 10 Hz) presented at the peak of the secondary component of a slow wave are shown in control solution (B), in a solution containing apamin (0.1 µM; D), and in a solution containing both L-NNA (0.1 mM) and apamin (0.1 µM; F). The time courses of the evoked membrane potential changes are overlaid in H. It can be seen that apamin reduced the amplitude of the IJP (D and H, b) but that the inhibition of mechanical activity persisted (D). In the presence of L-NNA (F and H, c), the residual IJP and inhibition of mechanical activity were both abolished (F). The upper set of time, force and voltage calibration bars applies to traces A-F. The lower set of time and voltage calibration bars applies to traces G and H.

When these experiments were repeated with L-NNA (0.1 mM) applied before apamin (0.1 µM), L-NNA abolished the inhibitory effects of vagal stimulation on mechanical activity. L-NNA had little effect on the amplitudes of IJPs evoked in the interval between slow waves but reduced the amplitudes of IJPs initiated during a slow wave. L-NNA-resistant hyperpolarizations were abolished by the subsequent application of apamin.

A series of experiments was carried out in preparations in which L-type Ca2+ channels were blocked by the Ca2+ antagonist nifedipine. Nifedipine (1 µM) did not alter the basal level of [Ca2+]i but reduced the amplitude of both the increase in [Ca2+]i and the contraction associated with each slow wave (Fig. 8, A and B). On the occasions in which slow waves were associated with discharges of muscle action potentials (Fig. 8A), the action potentials were abolished by nifedipine; however, the slow waves themselves were little affected (31). In control solutions, the mean change in [Ca2+]i associated with each slow wave was 0.4 ± 0.05 F340/380 (n = 4) and the mean increase in tension associated with each slow wave was 3.5 ± 1.0 mN (n = 4). After adding nifedipine (1 µM) to the physiological saline, each slow wave was associated with a peak increase in [Ca2+]i of 0.2 ± 0.05 F340/380 and a peak increase in tension of 0.8 ± 0.4 mN. After blocking Ca2+ entry with nifedipine, vagal stimulation continued to inhibit the contraction associated with each slow wave (Fig. 8B). The period of mechanical inhibition was not associated with a change in the pattern of [Ca2+]i linked to each slow wave or with the level of [Ca2+]i detected between slow waves (Fig. 8B). Again, the inhibitory responses to vagal stimulation were abolished by L-NNA (0.1 mM; Fig. 8C). These experiments suggest that vagal stimulation was able to inhibit contractions triggered by the release of Ca2+ from intracellular stores.


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Fig. 8.   Effect of nifedipine and L-NNA on responses to vagal stimulation recorded from the circular muscle layer of guinea pig antrum. A: vagal stimulation (1 Hz for 50 s) had little effect on slow waves (a) and associated changes in [Ca2+]i (b) but inhibited contractions of the circular layer (c). Note that in this preparation, slow waves intermittently generated bursts of muscle action potentials like those recorded from smooth muscle cells in the longitudinal layer. B: addition of nifedipine (1 µM) abolished the discharge of action potentials (a) and reduced the amplitude of associated increases in [Ca2+]i (b) and contractions (c). However, vagal stimulation continued to inhibit the amplitude of the contraction associated with each slow wave (c). C: inhibitory response to vagal stimulation was abolished by L-NNA (0.1 mM). All recordings were obtained from the same cell; the peak negative potential, in the absence of stimulation, remained -66 mV throughout. The voltage calibration, calcium ratio, force, and time calibration bars apply to the three sets of recordings.

Each of the previous observations has suggested that vagal inhibition does not change the pattern of [Ca2+]i associated with each slow wave. This seems surprising given that the responses to vagal stimulation are characterized by the occurrence of IJPs and that in control solutions slow waves activate L-type Ca2+ channels to allow Ca2+ entry. This was examined further by presenting brief trains of high-frequency stimuli (8 stimuli at 10 Hz) at various parts of the slow wave cycle while measuring [Ca2+]i. When stimuli were presented in the interval between slow waves, no change in [Ca2+]i occurred during the IJP or during the increase in [Ca2+]i associated with the following slow wave; nevertheless, the contractile response was reduced. If presented at the peak of the regenerative component of each slow wave, the slow wave was transiently interrupted. Although there was a dramatic reduction in the contractile response, the overall change in [Ca2+]i was little affected. If, however, the volley of stimuli was presented later, so that the IJP occurred during the later part of the regenerative response, the IJP terminated the regenerative response and shortened the duration of the associated increase in [Ca2+]i (Fig. 9). At the same time, the contractile response associated with that slow wave was severely attenuated (Fig. 9). Thus a change in the [Ca2+]i pattern associated with each slow wave could only be accomplished if the IJP shortened the secondary component of the slow wave.


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Fig. 9.   Effect of brief trains of vagal stimuli on changes in [Ca2+]i delivered during the regenerative component of slow waves. Each set of traces shows the effect of a brief train of vagal stimuli (; 8 at 10 Hz), on the time courses of slow waves (A and B, a), associated changes in [Ca2+]i (A and B, b), and contractions of the circular muscle layer (A and B, c). Since the volley of impulses was delivered progressively earlier during the regenerative component of the slow wave, both the slow wave and associated increase in [Ca2+]i were progressively shortened. Both recordings were obtained from the same cell; the peak negative potential was -65 mV. The voltage calibration, emission ratio, force and time calibration bars apply to each set of recordings.


    DISCUSSION
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

Our experiments have examined whether vagal inhibitory fibers functionally innervate the three distinct types of rhythmically active cells present in the antrum. We have shown that pacemaker activity is unaffected by vagal stimulation and that the dominant effect is inhibition of force production by smooth muscle cells of the circular muscle layer. In this layer, vagal fibers activate intrinsic neurons that release NO, which in turn appears to reduce the responses of contractile proteins to the increase in [Ca2+]i associated with each slow wave. Only when higher-frequency bursts of stimuli were presented at critical periods of the slow wave could changes in [Ca2+]i be detected.

When recording from the outer layer of smooth muscle cells, follower potentials were detected. Follower potentials occurred synchronously with contractions of the circular muscle. This observation agrees with the finding that slow waves in the circular muscle layer and follower potentials occur at the same time (15). In the longitudinal layer, follower potentials failed to trigger secondary regenerative components (15); rather, they triggered bursts of action potentials (Fig. 1A). The action potentials, but not the follower potentials, resulted from the activation of L-type Ca2+ channels since they, but not the follower potentials, were blocked by the Ca2+ antagonist nifedipine. Vagal stimulation failed to produce substantial hyperpolarizations, even during the peak of a follower potential, and failed to prevent the discharge of muscle action potentials (Fig. 2). In contrast, the same trains of stimuli evoked substantial hyperpolarizations in the circular layer, particularly when the membrane potential was depolarized during a slow wave (Fig. 7). This might suggest that much of the hyperpolarization detected in the longitudinal layer resulted from the passive spread of current from the circular layer, since in this tissue the two layers appear to be electrically connected via ICC-MY (15). If the IJP was initiated in the longitudinal layer, it would be expected to increase in amplitude during a depolarization. Conversely, the increased amplitude of the corresponding IJP triggered in the circular layer may not be transmitted to the longitudinal layer because of the intense conductance change occurring during the plateau phase of the driving potential. Whatever the case, our observations indicate that vagal inhibitory nerve fibers have little ability to alter the electrical properties of the longitudinal muscle layer. Together, these observations suggest that the longitudinal muscle layer of the stomach, as with other regions of the intestine (19, 30), receives a sparse inhibitory innervation. This view is supported by the observation that only a few fibers containing NO synthase innervate the longitudinal layer of many regions of the guinea pig gastrointestinal tract (18).

Brief high-frequency trains of vagal stimuli also failed to evoke substantial hyperpolarizations in driving cells. Again, during the plateau phase of a driving potential an IJP was not detected. Either the conductance change during the plateau phase was sufficiently intense to short-circuit an inhibitory conductance change or driving cells received few inhibitory inputs (32). Moreover, long trains of low-frequency vagal stimulation did not alter the shape or consistently change the rate of generation of driving potentials (Fig. 3). This observation is at first sight surprising. In the heart, even moderate trains of stimuli produce dramatic changes in the discharge of pacemaker action potentials (3, 8, 10, 11). Clearly, the present observations indicate that this is not the case for driving cells. All of the observations presented in this report suggest that pacemaker activity is not under vagal inhibitory control in the stomach, with the frequency of occurrence of slow waves (Fig. 4) being similarly unaffected (23).

In the circular muscle layer, vagal stimulation initiated IJPs that involved apamin-sensitive channels (Fig. 7) and apamin-insensitive but L-NNA-sensitive channels. Two component IJPs have been detected in a number of different intestinal preparations (5, 29), with such IJPs often being followed by a third rebound excitatory phase (35). The ability of apamin to inhibit IJPs differentially, on the basis of where they were initiated during the slow wave cycle, may reflect differing ionic conductances of the two components. If the L-NNA-sensitive component had a reversal potential near the peak negative potential of circular muscle cells, it would fail to generate a net outward current; the resultant IJP would be entirely sensitive to apamin. Conversely, the L-NNA-sensitive component would produce a large-current flow when initiated during a slow wave and the IJP would be less effected by apamin. Alternatively, the blockade produced by apamin may be voltage sensitive. In many preparations, the hyperpolarization associated with apamin-sensitive channels is largely responsible for inhibition of mechanical activity, although an apamin-insensitive component invariably persists (12). In canine gastric fundus, inhibitory nerve stimulation evokes a hyperpolarization, a relaxation, and a concomitant fall in [Ca2+]i. There it was shown that, although a hyperpolarization could suppress Ca2+ entry via L-type Ca2+ channels if these had been activated, the hyperpolarizations themselves made little contribution to the inhibition of mechanical activity. Rather, it was suggested that NO enhanced the uptake of Ca2+ by the sarcoplasmic reticulum or reduced the Ca2+ sensitivity of the contractile apparatus (1).

In the guinea pig antrum, blockade of apamin-sensitive channels had little or no effect on the ability of low-frequency vagal stimulation to reduce the force of contraction associated with each slow wave. However, inhibiting the formation of NO with L-NNA abolished the vagal responses, an observation that supports the view that NO is the dominant transmitter released by the vagus in the stomach (14). Vagal stimulation inhibited contractions when they resulted either from both Ca2+ entry and internal Ca2+ release or from Ca2+ release alone (Figs. 4 and 8), suggesting that the inhibitory mechanism affected contractile responses from whichever source of Ca2+ they were triggered. Unlike the dog fundus, vagal inhibitory responses were not associated with a change in [Ca2+ ]i. Thus neither resting [Ca2+]i nor peak increase in [Ca2+]i associated with each slow wave was changed (Fig. 4). Clearly, since we were measuring syncytial [Ca2+]i, the properties of a small Ca2+ compartment could have been changed. Nevertheless, the simplest explanation for our observations is that during long periods of low-frequency stimulation neurally released NO produces inhibition solely by reducing the Ca2+ sensitivity of the contractile proteins. This mechanism has not been demonstrated in other tissues. However, cGMP, the second messenger produced in many tissues by NO (21), has been shown to directly reduce the sensitivity of arterial contractile proteins to Ca2+ (24).

Higher-frequency trains of vagal stimulation were able to modify the increase in [Ca2+]i associated with each slow wave but only if they were presented during the secondary component of the slow wave (Fig. 9). If presented before a slow wave, the vagal volley reduced the amplitude of the associated contraction but did not change the [Ca2+]i transient. If presented during the regenerative component, the duration of the [Ca2+]i transient was shortened; at the same time, the secondary component was truncated (Fig. 9). As has been pointed out, the release of Ca2+ from intracellular stores appears to activate sets of Ca2+-sensitive channels in the membranes of circular muscle cells. This provides the dominant conductance change, which moves the membrane potential through a region in which L-type Ca2+ channels are activated (16, 31). An IJP triggered during a slow wave would be expected to reduce Ca2+ entry via L-type Ca2+ channels. Although this was the case, the reduction in the peak increase in [Ca2+]i was slight; the most obvious change was a shortening in the duration of the Ca2+ transient (Fig. 9). This implies that the hyperpolarization produced during a slow wave does not lead to the closure of many L-type Ca2+ channels. Presumably, most L-type Ca2+ channels are activated at potentials very close to the potential at which the regenerative component of the slow wave is initiated, that is, about -40 mV, and the hyperpolarizations produced by vagal stimulation at potentials positive to this barely affect L-type Ca2+ channel opening. IJPs triggered during a slow wave had longer durations than did those triggered in the interval between slow waves (Fig. 8). This might suggest that IJPs are able to transiently inhibit the secondary component of the slow wave (Fig. 7). When initiated later in the second component, IJPs were able to terminate the secondary regenerative component (Fig. 9). It is not clear whether this results from a direct inactivation of the secondary component by a membrane hyperpolarization or whether a second messenger produced by the inhibitory transmitter inactivates the secondary component.

In summary, our observations indicate that inhibitory transmitters released by vagal inputs do not modify the generation of myogenic activity by driving cells. However, the mechanical activity of the circular layer is dramatically changed by vagal activity; the force produced by the circular layer in response to each slow wave is greatly attenuated. This is the case whether contractions are triggered mainly by Ca2+ entering through L-type Ca2+ channels or by internally released Ca2+. Inhibition depends largely on the release of NO and is mediated without any great change in the profiles of [Ca2+]i associated with each slow wave produced by the circular muscle layer. Together, these observations suggest that the dominant effect of neurally released NO in this tissue is to reduce the sensitivity of contractile proteins to an increase in [Ca2+]i. Our experiments have not examined whether inhibition results from a direct action of NO on smooth muscle or whether it must first act on an intermediate set of cells such as ICC-IM. It is clear that the circular layer of guinea pig antrum contains numerous ICC-IM (6, 15). It may well be that, as in other regions of the gastrointestinal tract, ICC-IM are the targets for intrinsic inhibitory fibers and produce the second messenger that ultimately causes inhibition (7, 38).


    ACKNOWLEDGEMENTS

We are grateful to Dr. Narelle Bramich for her helpful comments on the manuscript.


    FOOTNOTES

This project was supported by a grant from the National Health and Medical Research Council of Australia.

Address for reprint requests and other correspondence: G. D. S. Hirst, Dept. of Zoology, Univ. of Melbourne, Victoria 3010, Australia (E-mail: d.hirst{at}zoology.unimelb.edu.au).

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. §1734 solely to indicate this fact.

Received 29 November 1999; accepted in final form 7 March 2000.


    REFERENCES
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ABSTRACT
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RESULTS
DISCUSSION
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