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MS 8920 Received 2 November 1998; accepted after revision 14 February 1999.
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ABSTRACT |
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INTRODUCTION |
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Many regions of the gastrointestinal tract generate slow waves and associated myogenic contractions; these occur at low frequencies in the absence of nervous activity or applied agonists (Tomita, 1981; Sanders, 1992). Although it was initially thought that this reflected the generation of electrical activity by gastrointestinal smooth muscle cells (Connor et al. 1974; El-Sharkaway & Daniel, 1975; Tomita, 1981), more recently it has been suggested that the generation of slow waves involves an interaction between interstitial cells of Cajal (ICC) and smooth muscle cells (Thuneberg, 1982; Smith et al. 1987; Ward et al. 1994; Sanders, 1996). According to this idea a potential generated in the layer of ICC would be conducted to nearby smooth muscle cells and in some way cause them to contract.
Recently it was shown that, during each slow wave, ICC lying in the myenteric region of guinea-pig antrum repetitively generate a large amplitude, long-lasting, depolarizing membrane potential change which spreads electrotonically to the nearby longitudinal and circular muscle layers. In the circular layer, but not the longitudinal layer, the ensuing depolarization triggers a secondary regenerative response which is abolished by low concentrations of caffeine (Dickens et al. 1999). The secondary responses last for several seconds but the events underlying them are not understood. In this report we describe a series of electrophysiological observations on small segments of the circular smooth muscle layer of the antral region of stomach. Depolarization evoked regenerative potentials which involved the voltage-activated release of Ca2+ from intracellular stores. After release, Ca2+ appears to activate an unknown set of membrane channels and the ensuing depolarization activates L-type Ca2+ (CaL) channels.
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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. Adult guinea-pigs of either sex were stunned, exsanguinated, and the stomach removed. The stomach was immersed in oxygenated physiological saline (composition (mM): NaCl, 120; NaHCO3, 25; NaH2PO4, 0·1; KCl, 5; MgCl2, 2; CaCl2, 2·5; and glucose, 11·1; bubbled with 95 % O2-5 % CO2) and cut along the greater curvature, so allowing the mucosa to be dissected away. In the initial experiments, the serosa was carefully removed under a dissecting microscope and preparations consisting of four to six bundles of circular muscle with the longitudinal layer intact were isolated. The preparations were pinned serosal surface uppermost in a recording chamber whose base consisted of a microscope coverslip coated with Sylgard silicone resin (Dow Corning Corp., Midland, MI, USA). A transducer was attached to one end of the preparation and intracellular recordings were made using a sharp microelectrode (90-150 M
) filled with 0·5 M KCl. In other experiments, single bundles of circular muscle (diameter 150-200 µm, length < 1 mm), with the longitudinal layer attached, were pinned out and the circular layer impaled with one or two independently mounted microelectrodes. These preparations consisted of the longitudinal layer, ICC lying in the myenteric region and the circular muscle layer. In the remaining experiments, single bundles of circular muscle (diameter 150-200 µm, length 300-600 µm), with the longitudinal layer removed, were pinned out and impaled with two independently mounted microelectrodes. These preparations were routinely examined, using an inverted compound microscope, to verify that the longitudinal layer had been removed. Preparations, either with an intact longitudinal layer or with the longitudinal layer removed, were labelled with an antibody to c-Kit and examined with either a fluorescence microscope or a confocal microscope to determine the distribution of ICC (see Burns et al. 1997; Dickens et al. 1999). To examine the excitability of the bundles of circular muscle one electrode was used to pass current and the other to record changes in membrane potential. Signals were amplified with an Axoclamp-2A amplifier (Axon Instruments), low pass filtered (cut-off frequency 1 kHz), digitized and stored on computer for later analysis. Some of the records used to illustrate the results were passed through a three-point moving average filter to reduce recording noise, this did not change the time course or amplitude of the responses in a detectable manner. Preparations were constantly perfused with physiological saline solution warmed to 35°C. The acetoxymethyl ester form of bis-(aminophenoxy) ethane-N,N,N',N'-tetraacetic acid (BAPTA AM), caffeine, cobalt chloride, nifedipine and cyclopiazonic acid (CPA), each obtained from Sigma, were used in some experiments.
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RESULTS |
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Slow waves recorded from the antral region of guinea-pig stomach
When intracellular recordings were made from smooth muscle cells in the circular muscle layer, a characteristic sequence of membrane potential changes was associated with each contraction (Fig. 1A). Cells had negative membrane potentials in the range -61 to -67 mV (mean ± S.E.M., -62·4 ± 0·8 mV, n = 5 observations, where each n value represents a measurement from a preparation taken from a separate animal) and discharged slow waves at frequencies of 2·5-3·7 waves min-1. During each slow wave cycle, the membrane potential was stable and then interrupted by a depolarization which triggered a secondary component. Slow waves had peak amplitudes in the range 31·9-41·4 mV (36·3 ± 1·7 mV, n = 5) and half-widths in the range 4·7-6·1 s (5·4 ± 0·3 s, n = 5). Each slow wave was associated with a contraction which had a maximum amplitude in the range 1·6-10·2 mN (5·0 ± 1·9 mN, n = 5), reached after the start of the slow wave (Fig. 1A). These observations are similar to those reported recently (Dickens et al. 1999).
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A, recordings of a slow wave (upper trace) and associated contraction (lower trace) recorded in control solution. B, recordings from the same cell some 15 min after adding nifedipine (1 µM) to the physiological saline. It can be seen that the slow wave was little changed but the amplitude of the associated contraction was much reduced. C, recordings obtained from the same cell, 1 min after adding caffeine (1 mM) to the physiological saline. It can be seen that both the secondary component of the slow wave and the remaining contraction were abolished. The force calibration applies to each contraction record, the voltage calibration applies to each voltage record. The time calibration refers to all recordings. | ||
Nifedipine (1 µM) reduced the amplitudes of the contractions associated with each slow wave (Fig. 1B) with the peak amplitude in nifedipine being 0·5 ± 0·1 mN (n= 5). However, the peak amplitude (35·2 ± 1·9 mV, n = 5) and the half-widths of the slow waves (5·3 ± 0·2 s, n = 5) were unchanged (Fig. 1B). This observation shows that CaL channels are activated during each slow wave and Ca2+ entry via these channels is responsible for the majority of the tension developed. However, CaL channels do not appear to provide a dominant conductance change during the slow wave. Caffeine (1 mM) abolished the secondary component of the slow wave to reveal a passive primary component with an amplitude in the range 6·1-11·2 mV (8·4 ± 1·0 mV, n = 5). At the same time the residual contraction was abolished (Fig. 1C).
These observations are consistent with the suggestion that during each slow wave a depolarization initiated elsewhere triggers a secondary depolarization in the circular smooth muscle layer. Although CaL channels are activated during the secondary phase and allow Ca2+ entry which triggers a contraction, a separate set of channels provides the dominant conductance change. These channels are associated with a small contraction and their activation is in some way inhibited by caffeine (Dickens et al. 1999).
Slow waves recorded from single bundles of circular muscle isolated from guinea-pig antrum
In the next experiments, single bundles of the circular smooth muscle layer were dissected out, pinned in a recording chamber and impaled with two independent microelectrodes. In the initial experiments, the longitudinal muscle layer, along with ICC of the myenteric region, were left attached. Slow waves were recorded from the circular muscle layer which resembled those recorded from preparations containing several bundles of muscle (Fig. 2A). As with the larger preparations (Fig. 1), caffeine (1 mM) abolished the secondary component of the slow waves (Fig. 2B) and revealed a passive primary component with an amplitude of 8·0 ± 0·7 mV (n= 4). Such preparations had resting membrane potentials in the range -61 to -70 mV (-64·1 ± 1·6 mV, n = 6); slow waves occurred at 2-4·8 waves min-1 (3·2 ± 0·4 waves min-1, n = 6) and had peak amplitudes in the range 24·0-38·6 mV (33·2 ± 2·1 mV, n = 6) with half-widths in the range 2·6-8·1 s (5·8 ± 0·8 s, n = 6). Unlike the recordings made from large preparations, those obtained from the smaller preparations invariably displayed membrane noise both between slow waves and most noticeably during the falling phases of the secondary components (Fig. 2A). The membrane noise was reduced in the presence of caffeine. When these preparations were labelled with an antibody to c-Kit, a few ICC were found to be scattered through the circular muscle layer and a network of ICC was found to lie in the myenteric region between the longitudinal and circular muscle layers (see Burns et al. 1997; Dickens et al. 1999).
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A-C, slow waves recorded from a single bundle of circular muscle with the longitudinal muscle layer remaining attached. D-F, regenerative potentials triggered in a single bundle of circular muscle which lacked the longitudinal muscle layer. The secondary component of the slow wave (A) was abolished by caffeine (1 mM, B) and returned after washing out the caffeine (C). Similarly the regenerative potentials (D), initiated by passing current (2 nA for 2 s, indicated by bars) through a second intracellular electrode, were abolished by caffeine (1 mM, E) and restored by washing out caffeine (F). The time and voltage calibrations apply to all traces. | ||
In the remaining experiments, recordings were made, again with two independent microelectrodes, from single bundles of circular muscle but with the longitudinal muscle layer removed. When these preparations were labelled with the antibody to c-Kit, a few ICC were found to be scattered through the circular muscle layer but the network of ICC lying in the myenteric region between the longitudinal and circular muscle layers had been removed. Clearly the dissection must also have removed the outer layer of longitudinal muscle: an observation confirmed routinely by viewing the preparations with an inverted microscope. Impalement of these preparations was associated with a negative membrane potential. Immediately after setting up the preparations, these were often more negative than -70 mV and stable. After superfusing the preparation with warmed physiological saline for 90-120 min, the membrane gradually depolarized until the resting potential lay in the range -55 to -64 mV (-60·8 ± 0·4 mV, n = 26), it then remained at these values for several hours. The preparations had input resistances in the range 1-9·5 M (3·3 ± 0·5 M, n = 22). The entire time course of each electrotonic potential was well described by a single exponential; these had time constants in the range 75-360 ms (160 ± 15 ms, n = 22). The membrane potential recordings showed membrane noise. Inspection of the two simultaneous membrane potential recordings indicated that the noise patterns were common to both recording points even if the electrodes were 200-400 µm apart. The membrane noise often summed to generate long-lasting waves of depolarization with peak amplitudes in the range 18-34 mV and half-widths in the range 3·5-5·8 s. These occurred at a frequency of 1-4 waves min-1 and were superimposed on irregular baselines. Occasionally these waves of depolarization triggered the discharge of one or more muscle action potentials. Although the discharge of action potentials was readily abolished by nifedipine (1 µM) the slower waves of depolarization invariably persisted. Caffeine (1 mM) abolished the slow depolarizations (n= 5) but failed to reveal the rhythmical discharge of small amplitude depolarizing potentials generated by ICC in preparations with an intact longitudinal layer (Figs 1C and 2B; see Dickens et al. 1999). As the myenteric layer of ICC had been removed from the bundles of circular muscle, the observations suggest that the bundles of circular smooth muscle are themselves rhythmically active. Whether this originates from the few ICC scattered through each bundle of circular muscle or from some intrinsic property of the smooth muscle cells is not known. The membrane noise persisted in the presence of nifedipine but was abolished by caffeine.
Regenerative potentials evoked in single bundles of antral muscle by depolarizing currents
The previous observations suggest that smooth muscle cells in the circular muscle layer can spontaneously generate long-lasting depolarizing waves. In intact tissues these are triggered by the regular flow of depolarizing current from ICC to the circular muscle layer (Dickens et al. 1999). When depolarizing current was passed through one of the intracellular recording electrodes in a bundle of circular muscle, a long-lasting wave of depolarization was recorded by the second electrode (Fig. 2D). Like the secondary component of a slow wave (Fig. 2A-C), these were abolished by caffeine (1 mM, n = 5, Fig. 2E) and restored by washing out the caffeine (Fig. 2F). The evoked depolarizations will be termed regenerative potentials. They had peak amplitudes in the range 17·6-38·0 mV (25·6 ± 1·7 mV, n = 22) and half-widths in the range 2·1-6·1 s (3·7 ± 0·3 s, n = 22). Occasionally regenerative potentials also triggered a discharge of muscle action potentials. When this occurred the muscle action potentials were brief, with half-widths in the range 26-92 ms (49 ± 8 ms, n = 8). Muscle action potentials, unlike regenerative potentials, were abolished by nifedipine (1 µM).
Regenerative potentials were triggered by a positive movement of the membrane potential rather than by changing the membrane potential to a particular threshold. Thus a regenerative potential was triggered either by a step of depolarizing current or at the offset of a hyperpolarizing current (Fig. 3). In many tissues membrane hyperpolarizations cause rebound excitation by reactivating channels. Such responses often have larger amplitudes than those triggered by depolarizing currents and usually occur shortly after the break of hyperpolarizing current flow. Neither of these patterns was observed with this tissue (Figs 3 and 7). Responses triggered by depolarizing currents had a mean peak amplitude of 26·2 ± 2·6 mV (n= 7). In the same preparations the responses triggered by hyperpolarizing currents had a mean peak amplitude of 25·9 ± 2·7 mV. Furthermore, there was invariably a delay from when the membrane potential returned to its resting value before a regenerative potential was initiated (Figs 3 and 7). In the preparations where the regenerative potentials triggered muscle action potentials, they were triggered by the regenerative potential and never at the end of a hyperpolarizing electrotonic potential.
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The superimposed upper pair of traces show regenerative potentials triggered by passing depolarizing and hyperpolarizing current pulses (lower traces) through the second intracellular electrode. Note the regenerative potentials had similar amplitudes when initiated by a current of either polarity. | ||
When preparations were stimulated with threshold currents, some stimuli failed to trigger a response (Fig. 4A). When they occurred, the regenerative potentials started after variable delays but had constant amplitudes (Fig. 4A). As the stimulus strength was increased, each stimulus initiated a regenerative potential but the latency continued to fluctuate (Fig. 4B). A further increase in the strength of stimulation reduced the latency fluctuations. Generally when the stimulating strength was increased to some 3 times its threshold value, such fluctuations were essentially absent (Fig. 4C). However, even though the latency no longer varied the regenerative potentials continued to be initiated after a finite delay (Fig. 4C).
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Each set of superimposed traces shows successive potentials triggered by passing depolarizing current pulses (indicated by bars) through a second intracellular electrode. The current intensities were 1 (A), 2 (B) and 3 nA (C). Note that the threshold stimulus in A failed to trigger a regenerative potential on one occasion, but, when triggered, the potentials were of similar amplitudes irrespective of current magnitude. Doubling the stimulus strength (B) produced no further increase in amplitude of the responses. Increasing the stimulus strength further (C) dramatically reduced the variability of latency of the responses but did not abolish their long latency completely. The voltage and time calibrations apply to all traces. | ||
The effect of changing the stimulus strength on the latency of the regenerative potential was examined in seven experiments. A single experiment is illustrated in Fig. 5. Sample traces, shown in the upper part of the figure, were chosen to reflect the mean latency with which a particular current initiated a regenerative potential (Fig. 5A). The lower part of the figure shows the relationship between latency of onset of the response and stimulating current (Fig. 5C). It can be seen that as the current intensity was increased, the latency shortened towards a minimum value of about 1 s. A similar pattern of behaviour was observed in each experiment, with the minimum latencies in the range 0·9-1·4 s (1·1 ± 0·1 s, n = 7).
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A, superimposed traces of selected potentials triggered by depolarizing current pulses of increasing intensity. The intensities of the applied currents were +1, +2, +3, +4 and +5 nA (B). The time calibration applies to all traces. C shows the relationship between current strength and mean latency. Each point is the mean of 10 determinations from a single preparation, the error bars represent ± 1 S.E.M. Note that although the latency decreases with increased strength it approaches a minimum value in this preparation of about 1 s. | ||
The previous observations suggest that a depolarization activates some pathway with slow onset kinetics. If this were the case one might expect that a brief depolarization that terminated before the end of the latency period would also trigger a response. This was found to be the case, brief depolarizing pulses which terminated before the minimum latency period readily evoked regenerative potentials, again with latencies from the start of the depolarizing pulse in excess of 1 s, even though the membrane potential had returned to its resting value before the regenerative potential occurred (Fig. 6).
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A, a regenerative potential triggered by a 2 nA depolarizing current pulse lasting for 5 s. B, a regenerative potential triggered by a 4 nA depolarizing current pulse lasting for 1 s. Note that although the membrane potential initially returned to its resting value, the shorter current pulse continued to initiate a regenerative potential. The voltage, current and time calibrations apply to all traces. | ||
Regenerative potentials were also triggered by periods of membrane hyperpolarization (Fig. 3). They also occurred after a considerably latency. An experiment is illustrated in Fig. 7. It can be seen that weak hyperpolarizing currents initiated regenerative potentials with mean latencies of over 5 s. With more intense currents the latency fell to a minimum of about 3 s (Fig. 7C). A similar pattern of behaviour was observed in six other experiments, where the minimum latencies lay in the range 1·2 to 3·4 s (2·5 ± 0·3 s, n = 7). In each experiment the minimum latency of the regenerative potentials triggered by hyperpolarizing currents exceeded that determined with depolarizing currents.
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A, superimposed traces of selected responses triggered by hyperpolarizing current pulses of increasing intensity. The intensities of the applied currents were -1, -2, -3, -4 and -5 nA (B). The time calibration applies to all traces. C shows the relationship between current strength and mean latency. Each point is the mean of 10 determinations from a single preparation, the error bars represent ± 1 S.E.M. Note that the latency decreases with increased strength towards a minimum of about 3 s. | ||
Regenerative potentials had long refractory periods. An experiment is shown in Fig. 8. An absolute refractory period preceded a partial refractory period. If a current pulse was injected 5 s after a conditioning pulse, it failed to trigger a regenerative component (Fig. 8A). When the test pulse and conditioning pulse were separated by some 10-20 s, the test pulses triggered regenerative potentials which had reduced amplitudes and slow rising phases (Fig. 8B-D). When separated by longer intervals the amplitude of the regenerative potential increased until the amplitude of the test response returned to that of the conditioning response (Fig. 8E). An absolute refractory period in the range 5-7 s was detected in five other preparations examined. They had partial refractory periods of 18, 20, 20, 22 and 25 s.
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The traces show regenerative potentials initiated by a pair of stimuli of constant strength (3 nA) and constant duration (5 s) (indicated by bars) applied at varying intervals (A, 5 s; B, 10 s; C, 15 s; D, 20 s; E, 25 s). Note that with a short separation (A), the second stimulus failed to trigger a response. As the separation was increased the regenerative potentials gradually reappeared, albeit with slower rising phases (B-D). With a longer separation responses of equal amplitude were initiated (E). The voltage and time calibrations apply to all traces. | ||
Inhibition of regenerative potentials by CPA or BAPTA but not by Co2+
The previous sections have indicated that positive movements of the membrane potential trigger regenerative potentials in single bundles of circular smooth muscle. These appear not to involve CaL channels. To examine the possibility that other Ca2+ channels were involved the effect of cobalt ions, Co2+, on the regenerative potentials was examined. In each of three preparations, regenerative potentials could be obtained in the presence of Co2+ (0·1, 0·3 and 1 mM). In each preparation Co2+ (0·1 mM) increased the frequency of occurrence of the myogenic depolarizations detected in the absence of stimulation. This lasted for some 20-40 min. After this time the preparations became quiescent but it was still possible to evoke myogenic responses. When the concentration of Co2+ was increased to 0·3 mM regenerative potentials were readily obtained without a change in their threshold (Fig. 9A and B). When the concentration of Co2+ was increased to 1 mM, the input resistances of the preparations fell to about 50 % of their control values but an increase in stimulation strength continued to trigger regenerative potentials.
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A and B, regenerative potentials triggered by 2 nA of depolarizing current (indicated by bars) in control solution and 30 min after adding Co2+ (0·3 mM), respectively. C-E, recorded from another preparation, regenerative potentials triggered by 4 nA current pulses (indicated by bars). The regenerative potential (C) was abolished some 15 min after adding CPA (30 µM) to the physiological saline (D). The potential was restored after washing with drug-free solution for 30 min (E). The voltage and time calibrations apply to all traces. | ||
Regenerative potentials were abolished by cyclopiazonic acid (CPA; 30 µM), an inhibitor of Ca2+-ATPase. In each of the four preparations examined, the amplitudes of the regenerative potentials were slowly reduced over a period of 12-15 min until they were abolished (Fig. 9C and D). Washing out CPA slowly restored the responses until they returned to their control values after 20-45 min (Fig. 9E). In each preparation, when the responses had been abolished by CPA, an increase in stimulation strength failed to restore the response.
Regenerative potentials were inhibited by adding BAPTA to the physiological saline. Superfusing the preparations with physiological saline containing BAPTA (10 µM) for 15 min, reduced the amplitude of regenerative potentials to about half their control value. The mean peak amplitude of control responses was 26·5 ± 3·8 mV; after treatment with BAPTA (10 µM) the responses had peak amplitudes of 13·5 ± 1·2 mV (n= 4). At the same time, the spontaneous generation of myogenic responses was abolished in each of the three preparations examined. Increasing the concentration of BAPTA to 20 µM, again superfused for 15 min, more markedly reduced the amplitude of the regenerative potentials. In a separate group of preparations, the control responses had peak amplitudes of 24·4 ± 2·2 mV; after treatment with BAPTA (20 µM) the responses had peak amplitudes of 3·2 ± 0·6 mV (n= 8). An experiment from the second series is illustrated in Fig. 10. It can be seen that regenerative potentials were initiated in control solution (Fig. 10A) but were much suppressed by treating the preparation with BAPTA (Fig. 10B). A common feature of the residual responses detected in preparations that had been pre-treated with these concentrations of BAPTA was that they appeared to be made up of unitary potential changes. Increasing the concentration of BAPTA AM, for example to 50 µM for 20 min, completely abolished the responses suggesting that they involved only a change in the internal concentration of Ca2+ ([Ca2+]i). The nature of the unitary potentials will be dealt with in a separate report.
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A, regenerative potential triggered by a 3 nA hyperpolarizing current pulse. B, the preparation was superfused with a solution containing BAPTA AM (20 µM) for 15 min and then washed with drug-free solution for a further 10 min. Hyperpolarizing current pulses triggered regenerative potentials of reduced amplitude. Washing the preparation with drug-free solution for several hours failed to restore the potentials. The voltage and time calibrations apply to both traces. | ||
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DISCUSSION |
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These observations reconfirm that slow waves persist in the presence of an organic calcium antagonist (Fig. 1; Sanders, 1992; Malysz et al. 1995). However, in the guinea-pig antrum, the contraction associated with each slow wave was much reduced in amplitude indicating that, in this tissue, nifedipine blocked most CaL channels. Thus although CaL channels are activated during each slow wave (Sanders, 1992), they appear to make a negligible contribution to the waveform of the slow wave. Recently it has been suggested that slow waves recorded from the circular smooth muscle layer of the antrum consist of two components, a passive component which results from current flow in nearby interstitial cells and a regenerative component which is triggered in the circular smooth muscle cells by this depolarization (Dickens et al. 1999). All-or-nothing regenerative potentials were initiated in isolated bundles from the circular muscle layer when their membrane potential was moved in a positive direction. These responses had similar time courses and amplitudes to the secondary component of the slow waves; both were abolished by caffeine (Fig. 2). These observations support the view that slow waves of the circular muscle layer consist of two separate components, one of which reflects the response of the circular muscle layer to membrane depolarization (Dickens et al. 1999).
However, all of the observations suggest that the regenerative potentials are unlikely to result from the activation of conventional voltage-activated ion selective channels. Thus regenerative potentials were triggered by a positive movement of the membrane potential, either from rest or from a hyperpolarized value (Fig. 3). Conventional voltage-dependent channels activated by depolarization open when the membrane potential exceeds a particular threshold. Regenerative potentials had distinct thresholds at which they were initiated after a variable latency. However, increasing the stimulus strength failed to reduce their latency to less than 1 s (Figs 4, 5 and 7). One explanation for the delay in onset of the regenerative potentials could be that preparations were not isopotential and the responses were triggered at some point electrically distant from the recording point. This seems unlikely on several grounds. In some experiments, the electrodes were separated by less than 50 µm, in others the electrodes were separated by over 300 µm. Whatever the separation, a minimum common delay was observed whichever electrode was used for current injection. As the preparations were never longer than 600 µm, less than half an electrical length constant (Abe & Tomita, 1968; Cousins et al. 1993), it is difficult to see how an excitable point could consistently have the same electrical separation from both electrodes. Secondly although preparations had low input resistances, suggesting that they contained several hundred smooth cells coupled together into a single syncytium, the entire time course of each electrotonic potential was well described by a single exponential which had a value similar to that determined for other guinea-pig intestinal muscles (Abe & Tomita, 1968; Cousins et al. 1993). These observations suggest that the preparations were electrically short and that electrically remote points are unlikely to exist. Furthermore, when membrane potential recordings obtained with the two electrodes were compared, even with wide electrode separations, the membrane noise detected by each electrode was very similar. Finally a latency of about 1 s represents about six electrical time constants. With a one-dimensional cable, an electrotonic delay of this magnitude implies that the point of initiation would have to be several length constants away, as such a distant point would be barely depolarized by current flow at a distant source. This effect is even more marked with multidimensional cables (Jack et al. 1975). Clearly the delay in the onset of the regenerative potentials is unlikely to result from them being initiated at an electrically remote site. The depolarization-activated voltage-dependent channels present in cells isolated from guinea-pig antrum fail to display the range of kinetic behaviours that would explain our observations (Noack et al. 1992). Similarly, the regenerative potentials are unlikely to result from the activation of hyperpolarization-activated channels. Although some of these channels display slow opening characteristics, which show marked voltage sensitivity (DiFrancesco, 1993), none are readily activated by depolarization.
A probable explanation for the origin of regenerative potentials is that they involve a pathway which increases [Ca2+]i by releasing Ca2+ from an intracellular store. After loading the tissues with BAPTA, which buffers [Ca2+]i to low levels, the amplitudes of regenerative potentials were much reduced (Fig. 10). Regenerative potentials were abolished by either caffeine (Fig. 2) or CPA (Fig. 9), which disrupts the internal storage of Ca2+ in smooth muscle (Uyama et al. 1992), but both persisted in the presence of either nifedipine or Co2+. Presumably the increase in [Ca2+]i triggers the nifedipine-resistant component of contraction (Fig. 1) and activates a set of Ca2+-sensitive channels which produce a depolarization. The identity of the Ca2+-sensitive channels responsible for the regenerative potentials is not known; presumably they are selective for either chloride (Large & Wang, 1996) or cations (Pacaud & Bolton, 1991). In the absence of nifedipine, the depolarization produced by the Ca2+-sensitive channels activates CaL channels to allow Ca2+ entry which triggers a substantial contraction (Fig. 1). As nifedipine readily reduces the contraction without modifying the slow wave, entering Ca2+ must have little or no access to the Ca2+-sensitive channels present in this tissue (see Bramich & Hirst, 1999, for related discussion). An alternative possibility is that a set of Ca2+ channels remains unblocked and effective in the presence of either nifedipine or Co2+ and that these trigger the regenerative release of Ca2+ from an internal store (Fabiato & Fabiato, 1975). However, such channels have not been described in this tissue (Noack et al. 1992). Moreover, if this were the case, the response should occur after a short latency; intestinal Ca2+ channels activate rapidly (Smirov et al. 1992) and the regenerative release of Ca2+ from an internal store occurs abruptly (Schiefer et al. 1995; Imaizumi et al. 1996).
Voltage-sensitive release of Ca2+ from intracellular stores has not been demonstrated in smooth muscle but has been in cardiac muscle (Howlett & Ferrier, 1997). However, again the increase in [Ca2+]i occurs on membrane depolarization after a short latency (Howlett & Ferrier, 1997). An alternative explanation for our findings is that rather than directly triggering Ca2+ release from intracellular stores, a positive change in membrane potential activates a metabolic pathway. Such pathways, when activated by transmitters, lead to the formation of second messengers which in some way trigger membrane potential changes with latencies which approach 1 s (Large, 1982; Cousins et al. 1993). If a positive movement of the membrane potential triggered the formation of a second messenger in this tissue, even a short lasting membrane depolarization would trigger a regenerative potential (Fig. 6). As an example, inositol trisphosphate (IP3) production has been shown to be enhanced by membrane depolarization in some smooth muscles (Itoh et al. 1992; Ganitkevich & Isenberg, 1993). Our observations could be explained if IP3, or some other messenger which released Ca2+ from intracellular stores, was produced at rest and its rate of production increased by a positive membrane potential change. Ca2+ would be released from an intracellular store, the increase in [Ca2+]i may be further amplified by calcium-induced calcium release (Iino, 1989) and an all-or-none regenerative potential would be triggered. Along these lines, it is of interest to note that some second messenger pathways, activated by transmitters, have long partial refractory periods like those detected in this study. The partial refractory period of cholinergic excitatory junction potentials recorded from guinea-pig intestine is some 20-30 s (H. M. Cousins & G. D. S. Hirst, unpublished observations).
In summary, our observations show that bundles of circular smooth muscle of the antrum generate regenerative potential changes which do not involve conventional voltage-sensitive channels. Rather they appear to involve the release of Ca2+ from intracellular stores and the subsequent activation of a population of unknown Ca2+-sensitive channels.
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REFERENCES |
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| Abe, Y. & Tomita, T. (1968). Cable properties of smooth muscle. The Journal of Physiology 196, 87-100 | [Medline] |
| Bramich, N. J. & Hirst, G. D. S. (1999). Sympathetic neuroeffector transmission in the rat anococcygeus muscle. The Journal of Physiology 516, 101-115 | [Abstract/Full Text] |
| Burns, A. J., Herbert, T. M., Ward, S. M. & Sanders, K. M. (1997). Interstitial cells of Cajal in the guinea-pig gastrointestinal tract as revealed by c-Kit immunohistochemistry. Cell and Tissue Research 290, 11-20 | [Medline] |
| Connor, J. A., Prosser, C. L. & Weems, W. A. (1974). A study of pace-maker activity in intestinal smooth muscle. The Journal of Physiology 240, 671-701 | [Medline] |
| Cousins, H. M., Edwards, F. R., Hirst, G. D. S. & Wendt, I. R. (1993). Cholinergic neuromuscular transmission in the longitudinal muscle of the guinea-pig ileum. The Journal of Physiology 471, 61-86 | [Abstract] |
| Dickens, E. J., Hirst, G. D. S. & Tomita, T. (1999). Identification of rhythmically active cells in guinea-pig stomach. The Journal of Physiology 514, 515-531 | [Abstract/Full Text] |
| DiFrancesco, D. (1993). Pacemaker mechanisms in cardiac muscle. Annual Review of Physiology 55, 455-472 | [Medline] |
| El-Sharkaway, T. Y. & Daniel, E. E. (1975). Ionic mechanisms of the control of intestinal electrical activity. American Journal of Physiology 229, 1287-1298 | [Medline] |
| Fabiato, A. & Fabiato, F. (1975). Contractions induced by a calcium-triggered release of calcium from the sarcoplasmic reticulum of single skinned cardiac cells. The Journal of Physiology 249, 469-495 | [Abstract] |
| Ganitkevich, V. Ya. & Isenberg, G. (1993). Membrane potential modulates inositol 1,4,5-trisphophate-mediated Ca2+ transients in guinea-pig coronary myocytes. The Journal of Physiology 470, 35-44 | [Abstract] |
| Howlett, S. E. & Ferrier, G. R. (1997). The 1996 Merck Frosst Award. The voltage sensitive release mechanism: a new trigger for cardiac contraction. Canadian The Journal of Physiology and Pharmacology 75, 1044-1057. | [Medline] |
| Iino, M. (1989). Calcium-induced calcium release mechanism in guinea pig taenia caeci. Journal of General Physiology 94, 363-383 | [Abstract] |
| Imaizumi, Y., Henmi, S., Nagano, N., Muraki, K. & Watanabe, M. (1996). Regulation of Ca-dependent K current and action potential shape by intracellular Ca storage sites in some types of smooth muscle. In Smooth Muscle Excitation, ed. Bolton, T. B. & Tomita, T., pp. 337-354. Academic Press, London. | |
| Itoh, M., Seki, N., Suzuki, H., Ito, S., Kajikura, J. & Kuriyama, H. (1992). Membrane hyperpolarization inhibits agonist-induced synthesis of inositol 1,4,5-trisphosphate in rabbit mesenteric artery. The Journal of Physiology 451, 307-328 | [Abstract] |
| Jack, J. J. B., Noble, D. & Tsien, R. W. (1975). Electric Current Flow in Excitable Cells. Oxford University Press. | |
| Large, W. A. (1982). Membrane potential responses of the mouse anococcygeus muscle to ionophoretically applied noradrenaline. The Journal of Physiology 326, 385-400 | [Medline] |
| Large, W. A. & Wang, Q. (1996). Characteristics and physiological role of the Ca2+-activated Cl- conductance in smooth muscle. American Journal of Physiology 217, C435-454. | |
| Malysz, J., Richardson, D., Farraway, L., Christen, M.-O. & Huizinga, J. D. (1995). Generation of slow wave type action potentials in the mouse small intestine involves a non-L-type calcium channel. Canadian The Journal of Physiology and Pharmacology 73, 1502-1511. | [Medline] |
| Noack, T., Deitmer, P. & Lammel, E. (1992). Characterization of membrane currents in single smooth muscle cells from the guinea-pig antrum. The Journal of Physiology 451, 387-417 | [Abstract] |
| Pacaud, P. & Bolton, T. B. (1991). Relation between muscarinic receptor cationic current and intestinal calcium in guinea-pig jejunal smooth muscle cells. The Journal of Physiology 441, 477-499 | [Abstract] |
| Sanders, K. M. (1992). Ionic mechanisms of electrical rhythmicity in gastrointestinal smooth muscles. Annual Review of Physiology 54, 439-453 | [Medline] |
| Sanders, K. M. (1996). A case for interstitial cells of Cajal as pacemakers and mediators of neurotransmission in the gastrointestinal tract. Gastroenterology 111, 492-515 | [Abstract] |
| Schiefer, A., Meissner, G. & Isenberg, G. (1995). Ca2+ activation and Ca2+ inactivation of canine reconstituted cardiac sarcoplasmic reticulum Ca2+-release channels. The Journal of Physiology 489, 337-348 | [Abstract] |
| Smirov, S. V., Zholos, A. V. & Shuba, M. F. (1992). Potential-dependent inward currents in single isolated smooth muscle cells of the rat ileum. The Journal of Physiology 454, 549-571 | [Abstract] |
| Smith, T. K., Reed, J. B. & Sanders, K. M. (1987). Origin and propagation of electrical slow waves in the circular layer of canine colon. American Journal of Physiology 252, C215-224 | [Medline] |
| Thuneberg, L. (1982). Interstitial cells of Cajal: intestinal pacemaker cells? Advances in Anatomy and Embryological Cell Biology 71, 1-130. | |
| Tomita, T. (1981). Electrical activity (spikes and slow waves) in gastrointestinal smooth muscles. In Smooth Muscle: An Assessment of Current Knowledge, ed. Bülbring, E., Brading, A. F., Jones, A. W. & Tomita, T., pp. 127-156. Edward Arnold, London. | |
| Uyama, Y., Imaizumi, Y. & Watanabe, M. (1992). Effects of cyclopiazonic acid, a novel Ca2+-ATPase inhibitor, on contractile responses in skinned ileal smooth muscle. British Journal of Pharmacology 106, 208-214 | [Medline] |
| Ward, S. M., Burns, A. J., Torihashi, S. & Sanders, K. M. (1994). Mutation of the proto-oncogene c-kit blocks development of interstitial cells and electrical rhythmicity in murine intestine. The Journal of Physiology 480, 91-97 | [Abstract] |
We wish to thank Dr Narelle Bramich, Ms Emma Dickens and Dr Frank Edwards for their helpful comments on the manuscript. In addition Ms Emma Dickens carried out the histological experiments reported here; we wish to express our gratitude. This project was supported by a grant from the Australian NH&MRC.
Corresponding author
G. D. S. Hirst: Department of Zoology, University of Melbourne, Parkville, Victoria 3052, Australia.
Email: georgedh{at}clyde.its.unimelb.edu.au
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