Department of Zoology, University of Melbourne, Parkville, Victoria 3052, Australia
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ABSTRACT |
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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
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INTRODUCTION |
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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).
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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 M) 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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RESULTS |
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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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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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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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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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DISCUSSION |
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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).
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ACKNOWLEDGEMENTS |
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We are grateful to Dr. Narelle Bramich for her helpful comments on the manuscript.
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FOOTNOTES |
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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.
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