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1 Department of Physiology and Cell Biology, University of Nevada School of Medicine, Reno, NV 89557 USA
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(Received 10 December 2004;
accepted after revision 26 January 2005;
first published online 27 January 2005)
Corresponding author S. M. Ward: Department of Physiology and Cell Biology, University of Nevada School of Medicine, Reno, NV 89557 USA. Email: sean{at}physio.unr.edu
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Introduction |
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Neural information, transmitted via ICC-IM, can influence the behaviour of the pacemaker ICC-MY. Electric field stimulation of enteric nerve processes in small muscle bundles of the guinea pig antrum, which contain only smooth muscle cells and ICC-IM, evoked cholinergic neural responses. When the bundles were attached to the ICC-MY network, vagal nerve stimulation induced premature pacemaker potentials (Hirst et al. 2002b), and repetitive stimulation paced the muscles (Beckett et al. 2003). This type of regulation was absent in W/WV mice that lack ICC-IM (Beckett et al. 2003). Although a role for ICC-SEP cannot be ruled out in these responses (these cells are also lost in W/WV mice), the findings suggest that neurally activated depolarization of ICC within the muscle layers can initiate premature slow waves in ICC-MY, and repetitive excitatory nerve stimulation may allow ICC-IM to become the dominant pacemaker.
The physiological evidence suggesting that ICC-IM mediate enteric neural responses and modify pacemaker activity has come primarily from studies of circular muscles of the lower oesophageal sphincter and of the gastric fundus, antrum and pyloric sphincter (Ward & Sanders, 2001; Hirst & Ward, 2003). In the fundus, ICC-IM form a parallel lattice of cells interspersed between nerve terminals and smooth muscle cells in both the circular and longitudinal muscle layers (Burns et al. 1996). In the mouse antrum, a dense population of ICC-IM is found in the circular muscle layer running parallel to the circular muscle cells, but few, if any, ICC-IM are found in the longitudinal layer (Hirst et al. 2002a). Antral ICC-IM have short lateral processes that appear to contact neighbouring ICC-IM cells to form a network (Hirst et al. 2002a). ICC-IM are found in close apposition to enteric nerve terminals and form gap junctions with smooth muscle cells (Horiguchi et al. 2003). As described previously, cholinergic and nitrergic neurotransmission is greatly attenuated in gastric and oesophageal muscles of W/WV mice that lack ICC-IM, suggesting that these cells have a critical role in mediating neurotransmission. These findings have been contested, however, with the suggestion that loss of neural responses in W/WV mice might be due to chronic weakening of the contractile apparatus of the smooth muscles of these animals and unrelated to the role of ICC-IM (Sivarao et al. 2001).
In the present study we have characterized the distribution of ICC-IM in the longitudinal muscle layer of the corpus and antrum of wild-type and W/WV mice to determine if there are regional variations in the density of these cells around the circumference and along the length of the stomach. Studies were also performed to examine the functional relationship between enteric nerve processes and ICC-IM in the longitudinal layer, and to determine whether ICC-IM play a role in the cholinergic and nitrergic control of longitudinal muscle layer excitability. The study provides evidence that, unlike circular muscle layer, there is considerable variation in the density of ICC-IM and the contacts between these cells and enteric nerves in the longitudinal muscle layer of the corpus and antrum. Finding regions of longitudinal muscle that lacked ICC-IM provided the opportunity to test whether ICC-IM are important mediators of cholinergic and nitrergic neural responses in gastric muscles isolated from identical wild-type animals.
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Methods |
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C57Bl6 wild-type and W/WV mutant mice between the ages of 30 and 60 days were used for the described studies. Animals were obtained from the Jackson Laboratory (Bar Harbour, MN, USA). The animals were maintained and the experiments were performed in accordance with the National Institutes of Health Guide for the Care and Use of Laboratory Animals, and the Institutional Animal Use and Care Committee at the University of Nevada approved all procedures used. On days of experiments, mice were anaesthetized with isoflurane (Baxter, Deerfield, IL, USA) prior to cervical dislocation and decapitation.
Morphological studies
Immunohistochemical analysis was performed on strips of corpus and antral muscles of wild-type and W/WV mutant mice. Whole mount preparations were prepared after removing the mucosa by sharp dissection. The remaining strips of tunica muscularis were pinned to the base of a dish filled with Sylgard elastomer (Dow Corning Corp., Midland, MI, USA) with the circular muscle layer facing upward. Tissues were fixed in either acetone (4°C; 10 min) or paraformaldehyde (4% w/v in 0.1 M phosphate buffer (PB) for 20 min at 4°C). Following fixation, preparations were washed for 60 min in phosphatebuffered saline (PBS; 0.01 M, pH 7.4). Incubation of tissues in 1% bovine serum albumin for 1 h at room temperature containing 0.3% Triton X-100 was used to reduce nonspecific antibody binding. For examination of ICC, tissues were incubated overnight at 4°C with a rat monoclonal antibody raised against Kit protein (ACK2; 5 µg ml–1 in PBS; Gibco-BRL, Gaithersburg, MD, USA). Immunoreactivity was detected using fluorescein-isothiocyanate (FITC)-conjugated secondary antibody (FITC-anti rat; Vector Laboratories, Burlingame, CA, USA; 1:100 in PBS, 1 h, room temperature). Control tissues were prepared in a similar manner, omitting ACK2 from the incubation solution. Alternatively, tissues were prepared for cryostat sections as previously described (Iino et al. 2004).
For double labelling, tissues were fixed, washed and preincubated in bovine serum albumin for 1 h (1% in PBS) before being incubated sequentially in a combination of primary antibodies. Tissues were incubated overnight at 4°C in primary antibodies. For double-label immunostaining, the first incubation was carried out for 24 h at 4°C. The tissue was subsequently washed in PBS before being incubated in a second antibody for an additional 24 h at 4°C. The combinations of antibodies used were rat/sheep and rat/goat (Table 1). Following incubation in primary antibodies, tissues were incubated separately in secondary antibodies (FITC and Texas Red (TR)). All secondary antibodies were purchased from Vector Laboratories, and diluted to 1:100 in PBS. Secondary incubations were performed for 1 h at room temperature. Control tissues were prepared by either omitting primary or secondary antibodies from the incubation solutions. All the antisera were diluted with 0.3% Triton X-100 in 0.01 M PBS (pH 7.4). Tissues were examined with a Zeiss LSM 510 Meta confocal microscope (Zeiss, Germany) with an excitation wavelength appropriate for FITC (488 nm) and TR (595 nm). Confocal micrographs are digital composites of Z-series scans of 10–15 optical sections through a depth of 3–40 µm.
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After animals were killed, stomachs, including approximately 5 mm of oesophagus and 10 mm of duodenum, were removed and placed in Krebs–Ringer bicarbonate solution (KRB). The gastric corpus and antrum were isolated by a surgical incision across the stomach above a distinct transition line in mucosal structure that separates the fundus from the corpus. A second incision was made across the terminal antrum just above the pylorus. The antrum (12 x 6 mm) was opened along the lesser curvature, and gastric contents were washed away with KRB. The antrum was subsequently pinned to the base of a Sylgard silicone elastomer (Dow Corning Corp., Midland, MI, USA) dish and the mucosa was removed by sharp dissection. For separate studies on the corpus (22 x 10 mm) and antrum, the stomach was divided by an incision across the region of the incisura angularis (see Fig. 1). The separate regions of the stomach were isolated and placed in a recording chamber with the serosal aspect of the muscle facing upward.
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. Transmembrane potentials were recorded with a standard electrometer (Intra 767; World Precision Instruments (WPI), Sarasota, FL, USA). The longitudinal muscle layer in stomachs of mice is approximately 10–15 µm thick in the corpus and 5–10 µm thick in the antrum. Thus, it was necessary to identify the muscle layer in which cells were impaled. This was accomplished on a routine basis by including propridium iodide (0.1% w/v in 3 M KCl) in the electrode filling solution. After intracellular recordings were performed, tissues were fixed with paraformaldehyde (4% (w/v) in 0.1 M PB) and examined with a confocal microscope with the appropriate excitation for propridium iodide (535 nm). Data were recorded onto a PC running AxoScope 8.0 data acquisition software (Axon Instruments), and hard copies were printed using Clampfit analysis software (Axon Instruments). All experiments were performed in the presence of nifedipine (1 µM) to reduce contractions and facilitate impalements of cells for extended periods. It has been previously demonstrated that slow waves in the gastric antrum are not affected by nifedipine (Suzuki & Hirst, 1999; Beckett et al. 2003). In all experiments, parallel platinum electrodes were placed on either side of the muscle strips, and neural responses were elicited by square wave pulses of electrical field stimulation (EFS; 0.1–0.5 ms pulse durations, one pulse –20 Hz, train duration of 1000 ms, 10–15 V) using a Grass S48 stimulator (Quincy, MA, USA). Isometric force measurements
Separate mechanical experiments were performed on gastric corpus and antrum using standard organ bath techniques. Muscles were dissected as described for electrophysiological experiments, and the mucosa was removed by sharp dissection. Muscle strips (approximately 4 x 2 mm) were isolated and attached to a fixed mount and to a Fort 10 isometric strain gauge (WPI). The muscles were immersed in organ baths maintained at 37 ± 0.5°C with oxygenated KRB. A resting force of 1.0–2.0 mN was applied, which was shown to set the muscles at optimum length (data not shown). This was followed by an equilibration period of 1 h, during which time the bath was continuously perfused with oxygenated KRB. Neural responses were elicited by square-wave pulses of EFS (0.1–0.5 ms pulse durations, one pulse –20 Hz, train duration 10–30 s, 10–15 V) using a Grass S48 stimulator. Signals were recorded using Biopac (Biopac Systems, Santa Barbara, CA, USA), and analysed using Acknowledge software (Biopac Systems, version 3.5.1).
Solutions and drugs
The bath chambers was constantly perfused with oxygenated KRB of the following composition (mM): NaCl 118.5, KCl 4.5, MgCl2 1.2, NaHCO3 23.8, KH2PO4 1.2, dextrose 11.0, CaCl2 2.4. The pH of the KRB was 7.3–7.4 when bubbled with 97% O2–3% CO2 at 37 ± 0.5°C. Muscles were left to equilibrate for at least 1 h before experiments began. For electrophysiological experiments, nifedipine was obtained from Sigma (St Louis, MO, USA) and dissolved in ethanol at a stock concentration of 10 mM before being added to the perfusion solution at a final concentration of 1 µM. Atropine, apamin, N
-nitro-L-arginine (L-NA), tetrodotoxin and propridium iodide were also obtained from Sigma, and dissolved in de-ionized H2O before being diluted in KRB to the final concentration stated in the results section.
Analysis of data
Data are expressed as means ± standard errors of the mean. Student's t test and the Mann–Whitney rank sum test were used where appropriate to evaluate differences in the data. P values of less than 0.05 were taken as a statistically significant difference. The ‘n’ values reported in the text refer to the number of muscle strips used for each experimental protocol. Each muscle strip used in ‘n’ values was taken from a separate animal. Several electrical parameters were analysed: (i) resting membrane potential (RMP), (ii) slow wave amplitude, (iii) duration of slow wave, (iv) inter-slow-wave period, and (v) frequency. Changes in slow-wave duration or inter-slow-wave cycle in response to electrical field stimulation were analysed by averaging the duration of five spontaneous events prior to EFS, and this was compared to the slow wave immediately following EFS. Figures displayed were made from digitized data using Adobe Photoshop 4.0.1 (Adobe Co., Mountain View, CA, USA) and Corel Draw 7.0 (Corel Corp., Ontario, Canada).
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Results |
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The Kit monoclonal antibody ACK2 was used to examine the regional distribution of ICC in the longitudinal muscle layer of the murine corpus and antrum. Confocal image reconstructions of whole mounts of the gastric corpus revealed ICC at three different locations within the stomach wall. ICC were observed within the circular and longitudinal muscle layers (ICC-IM) running parallel to the long axis of the muscle fibres. ICC were also observed along the intermuscular plane between the circular and longitudinal muscle layers, at the level of the myenteric plexus (ICC-MY; Fig. 1A, C and E). The morphologies of these different populations of ICC were similar to those previously published for the murine stomach (Burns et al. 1996; Hirst et al. 2002a). Examination of the spindle-shaped ICC located within the longitudinal muscle layer revealed a marked heterogeneity in the distribution of these cells along the longitudinal and transverse axes of the corpus (Fig. 1B, D and F). The density of ICC-IM in the longitudinal muscle layer was the greatest close to the border with the fundus (0–2 mm; 5.2 ± 0.2 ICC-IM per random 100 µm transecting line perpendicular to the long axis of the muscle fibres). The density of ICC-IM decreased from the corpus to the antrum (2–4 mm from the border of the fundus, the density was reduced to 2.1 ± 0.4 cells per 100 µm transecting line (P < 0.01 compared with 0–2 mm), and 5–6 mm from the fundus the density approached 0 cells per 100 µm transecting line, and 4 mm oral to the incisura angularis no ICC-IM were observed in the corpus, P < 0.01). Around the stomach, the density of ICC-IM in the corpus was greatest along the greater curvature and decreased towards the lesser curvature of the stomach. Towards the lesser curvature (5 mm from the greater curvature), the density of ICC-IM was 1.8 ± 0.2 cells per 100 µm transecting line compared with 5.2 ± 0.2 ICC-IM along the greater curvature (Fig. 2A–D; P < 0.001). In W/WV mutant animals, ICC-IM were absent from the longitudinal and circular muscle layers of the corpus (Fig. 2G–J).
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Relationship between enteric nerves and ICC-IM
It has been previously shown that excitatory and inhibitory motor nerves form an intimate relationship with ICC-IM in the circular muscle layer of the gastric fundus and antrum (Ward et al. 2000; Beckett et al. 2003). To determine if this relationship exists between enteric motor nerves and ICC-IM within the longitudinal muscle layer of corpus and antrum, double-labelling immunohistochemical experiments using antibodies to either vesicular acetylcholine transporter (vAChT) or nitric oxide synthase (nNOS) were performed with an antibody against the Kit receptor.
In the gastric corpus, vAChT-like (vAChT-LI) immunoreactivity revealed a dense network of varicose nerve fibres within the circular and longitudinal muscle layers and in the region of the myenteric plexus (Fig. 3). At a distance of 0–2 mm from the gastric fundus border, there were 6.1 ± 0.3 vACT-LI nerve fibres per 100 µm cross-section transecting line perpendicular to the axis of the longitudinal muscle. At 5 mm from the fundus border, this number had decreased to 1.3 ± 0.14 fibres per 100 µm cross-section transecting line (P < 0.001 compared with 0–2 mm). There was also a decrease in the number of vAChT nerve fibres within the longitudinal layer from the greater to lesser curvature. The number of fibres at a distance 0–2 mm from the fundus border decreased from 6.1 ± 0.3 along the greater curvature to 1.7 ± 0.3 fibres per 100 µm transecting line at a distance of 6 mm from the greater to lesser curvature (P < 0.001). Double labelling with vAChT and Kit revealed that in the corpus varicose, nerve fibres containing vAChT-LI were closely apposed to ICC-IM (Fig. 3). vAChT nerve fibres were associated with ICC-IM for distances over up to 200 µm. In six animals, 212 ICC-IM were counted; of these, 198 ICC-IM (93.4%) were observed in close association with one vAChT-LI nerve fibre, six ICC-IM (2.8%) were observed with two vAChT nerves, and six ICC-IM (2.8%) did not appear associated with any vAChT-LI immunoreactive nerve fibres, and two ICC-IM (1%) shared one vAChT nerve (Fig. 3).
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Although ICC-IM were absent from the longitudinal layer of the corpus and antrum of W/WV mutants, the number of vAChT-LI and nNOS-LI nerves per 100 µm cross-section transecting line perpendicular to the axis of the longitudinal muscle was similar to that of wild-type animals for all regions (Fig. 6).
Electrical activity of the longitudinal muscle of gastric corpus and antrum
Electrical activity was measured in longitudinal muscles of the corpus using standard intracellular recording techniques. Muscle cells were impaled 2 mm from the border with the fundus along the greater curvature. To confirm that longitudinal cells were impaled, cells were filled with propridium iodide (0.1% w/v) while electrical recordings were made (Fig. 7). Longitudinal muscle cells of the corpus had resting membrane potentials of –46 ± 1.0 mV. Cells impaled in 75% of the muscles had small irregular membrane potential oscillations (n = 20; Fig. 7C). Cells in the remaining 25% of muscles had more regular (slow-wave) oscillations with an average amplitude of 3.7 ± 1.0 mV, duration of 2.3 ± 0.1 s, and frequency of 8.9 ± 0.3 cycles min–1 (Fig. 7C).
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Electrical responses to nerve stimulation
Previous studies have shown that ICC-IM are required for postjunctional responses to cholinergic and nitrergic motor neurones in circular muscles of the gastric fundus and antrum (Burns et al. 1996; Ward et al. 2000; Beckett et al. 2002; Suzuki et al. 2003). Here we examined electrical responses to nerve stimulation in longitudinal muscles of the corpus and antrum. EFS (1–20 Hz, 0.1–0.5 ms for 1 s) of corpus muscles evoked biphasic responses. The first phase consisted of a rapid depolarization or excitatory junction potential (EJP) averaging 5.0 ± 0.5, 7.0 ± 0.5, 8.0 ± 1.0 or 9.0 ± 1.5 mV in amplitude, and 0.5 ± 0.02, 0.9 ± 0.1, 0.8 ± 0.1 or 0.7 ± 0.1 s in duration at 1, 5, 10 and 20 Hz, respectively. The EJP was followed by a longer and more pronounced hyperpolarization or inhibitory junction potential (IJP) averaging 5.0 ± 2.0, 8.0 ± 1.0, 9.0 ± 1.0 or 11.0 ± 1.0 mV in amplitude, and 0.8 ± 0.1, 2.0 ± 0.2, 2.5 ± 0.1 or 2.8 ± 0.2 s in duration for 1, 5, 10 and 20 Hz stimuli, respectively (n = 9; Fig. 8). Addition of the nitric oxide synthase inhibitor L-NA (100 µM) abolished IJPs at 5–20 Hz, and prolonged the duration of EJPs produced by trains of impulses. In the continued presence of L-NA, atropine (1 µM) abolished the EJPs and unmasked a fast IJP that averaged 7 ± 1.6 mV in amplitude and 1.45 ± 0.2 s in duration at 20 Hz (n = 6). The fast IJP was blocked by apamin (0.2 µM) or tetrodotoxin (1 µM). These data suggest that corpus longitudinal muscles are innervated by both excitatory cholinergic and nitregic/purinergic inhibitory neurones (Mackenzie & Burnstock, 1980).
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Neural responses to EFS in the corpus and antrum of W/WV mutant animals
Muscles of wild-type mice have reduced ICC-IM in the longitudinal muscle layer, as described previously. But the circular muscle within intact muscles contains an abundance of ICC-IM from corpus through the antrum (Burns et al. 1996). It is possible that a portion of the responses we noted, particularly in the antrum could have been due to neurally mediated responses via circular muscle ICC-IM. Therefore, we also recorded from muscles of W/WV mice, which lack all ICC-IM in the corpus and antrum. Resting membrane potentials of longitudinal muscle cells of corpus muscles of W/WV mice, recorded 2 mm from the fundus border as in wild-type animals, averaged –48.6 ± 1.3 mV (n = 5; Fig. 10), and the recordings lacked the characteristic noisy baseline observed in recordings from wild-type muscles (compare Fig. 7C with Fig. 7E and Beckett et al. 2004). EFS (0.3 ms, 1–20 Hz for 1 s) produced little or no postjunctional responses under control conditions, and responses were not unmasked by addition of L-NA (100 µM), or the combination of L-NA and atropine (1 µM, n = 5; Fig. 10A–C).
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Contractile measurements
We also compared mechanical responses of longitudinal muscles of the corpus and antrum using 2 mm x 4 mm muscle strips prepared from the area along the greater curvature. Spontaneous contractile activity was observed in muscles of both regions. Phasic contractions occurred at 7.5 ± 0.4 min–1 in the corpus (n = 11; Fig. 11) and at 3.4 ± 0.24 min–1 in the antrum (n = 15; Fig. 12). It was difficult to determine the period between contractions in the corpus due to the sinusoidal nature of the contractions. The period between contractions in the antrum was 12.8 ± 2.62 s. Spontaneous contractions in the corpus were weaker than in the antrum (i.e. 0.05 ± 0.01 mN for corpus versus 0.45 ± 0.03 mN mg–1 for antrum; P < 0.01). EFS (1–20 Hz, 0.1–0.5 ms duration for 10 s, n = 7) caused an initial relaxation of corpus muscles that was followed by contraction. The two phases of the response were frequency dependent (Fig. 11A). L-NA (100 µM) blocked the relaxation phase and potentiated the contractile phase (Fig. 11B). The contractile event was inhibited by atropine (1 µM; Fig. 11C). After L-NA and atropine, high-frequency stimulation (i.e. 20 Hz) yielded small contractions that were blocked by TTX (1 µM; Fig. 11D). EFS also evoked frequency-dependent contractions in longitudinal corpus muscles of W/WV mice. These averaged 0.11 ± 0.01 and 0.16 ± 0.03 mN at 5 and 10 Hz, respectively. L-NA (100 µM) and atropine (1 µM) had little or no effect on contractile responses to EFS. Any responses that were observed were blocked by TTX (1 µM) (Fig. 11E–H).
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These data suggest that a noncholinergic excitatory transmitter is released in sufficient concentrations to excite longitudinal muscles at frequencies of 5 Hz and above. In a further series of contractile experiments, we explored the hypothesis that the noncholinergic neurotransmitter could be a neurokinin. In corpus muscles of wild-type and W/WV mice, the NK2 antagonist SR-48968 (1 µM) partially inhibited EFS-evoked contractile responses observed after L-NA and atropine. SR-48968 also partially inhibited the phase advancement in slow waves observed in antrum under the same conditions. The NK1 antagonist GR-82334 (1 µM), added in the continued presence of the NK2 antagonist, completely inhibited responses to EFS in the corpus and antrum. These data suggest that higher frequencies of stimulation can release neurokinnins that can directly affect smooth muscle cells or reach the ICC-MY in the antrum (not shown).
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Discussion |
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The general concept that ICC-IM are required for neurotransmission has been questioned by some investigators who suggested an alternative hypothesis that developmental defects in smooth muscle cells may explain the loss of neural responses in gastrointestinal muscles of W/WV animals (Sivarao et al. 2001). Using intraluminal manometric measurements to examine the lower oesophageal sphincter (LES) pressures, these authors found that the LES of in W/WV mice was hypotensive, but the sphincter still relaxed in accordance with the swallow reflex or in response to vagal stimulation. It was concluded that the hypotensive nature of the LES was likely to be due to defects in the smooth muscle cells. These authors appear not to have considered an earlier report that responses to exogenous ACh are identical in wild-type and W/WV muscles (Ward et al. 2000), which suggested that smooth muscle function is normal in mutant animals with compromised Kit signalling. This has been confirmed more recently in fundus muscles from Sl/Sld animals that also lack ICC-IM (Beckett et al. 2002). In the present study we have shown that regions lacking ICC-IM, which occur naturally in wild-type mice, also have reduced nitrergic and cholinergic responses. Our results are consistent with the hypothesis that ICC-IM are critical intermediaries in the mediation of postjunctional responses to the primary excitatory and inhibitory neurotransmitters in gastrointestinal muscles.
We examined the distribution of ICC-IM in the in the longitudinal layers of the corpus and antrum, and found a pronounced gradient in the distribution of these cells in which the numbers of ICC-IM rapidly declined in the circumferential axis toward the lesser curvature, and in the longitudinal axis toward the antrum. In the region where ICC-IM were present, robust cholinergic and nitrergic neural responses were recorded. Cholinergic and nitrergic neural responses were not observed in the longitudinal muscle of the antrum that lacked ICC-IM.
The results of this study further emphasize the complexity of neural regulation of the pacemaker mechanism in gastric muscles. It has been previously demonstrated that vagal nerve stimulation in the gastric antrum evokes premature slow waves, and increases the frequency of slow waves. Simultaneous recordings of pacemaker potentials from ICC-MY and slow waves in the circular muscle layer showed that the site of initiation of rhythmical activity switched from ICC-MY to the circular layer, and it was hypothesized that the dominant pacemaker switches from ICC-MY to ICC-IM during vagal stimulation (Hirst et al. 2002b). Activation of muscarinic receptors of the gastric antrum is capable of increasing the frequency of antral slow waves (Beckett et al. 2003). Since this increase in frequency did not occur in the W/WV mutant mice, it was concluded from this study that the pacing of slow waves by cholinergic neurones occurs via ICC-IM. In the present study, we have observed that at higher frequencies, a noncholinergic neurokinin transmitter can also affect slow-wave frequency, and this can occur in the absence of ICC-IM. Thus, neurokinins, acting via NK1 and NK2 receptors, are capable of affecting pacemaker frequency either by depolarization of smooth muscle cells or by overflow onto nearby ICC-MY.
The fact that neurokinins released from enteric motor neurones can elicit responses in the absence of ICC-IM does not necessarily mean that extrajunctional receptors mediate the effects of neurokinin in wild-type muscles. Several studies have shown that substance-P-containing neurones form very close apposition with ICC-IM in the stomach, and in the deep muscular plexus of the small intestine (Lavin et al. 1998; Vannucchi et al. 1999; Ibba Manneschi et al. 2004; Iino et al. 2004). ICC-IM in the small bowel are typically referred to as ICC-DMP, and have similar morphological relationships with enteric motor neurones as ICC-IM in the stomach and colon. NK1 receptor internalization was studied in muscle stimulated with exogenous substance P or with neurokinins released from enteric motor neurones (Iino et al. 2004). NK1 receptor internalization occurred in smooth muscle cells and ICC-DMP in response to exogenous substance P, but receptors were internalized only in ICC-DMP in response to neurally released neurokinins (Iino et al. 2004). Thus when present, ICC-DMP (ICC-IM) may mediate the majority of neurokinin responses. It is also possible that ICC-IM might express enzymes to metabolize neurokinins released from nerve terminals, and loss of this mechanism could promote escape of neurokinins from the synaptic regions formed by ICC-IM and nerve varicosities to activate nonjunctional receptors.
In summary, there is considerable regional variation in the density of ICC-IM in the longitudinal muscle layer of the corpus and antral regions of the murine stomach. This variation in distribution and density occurs around the greater to lesser curvature and along the stomach wall. Cholinergic and nitrergic neural responses are present when ICC-IM are present in the corpus, but are absent in the antral region of the stomach when ICC-IM are absent. The regional variation in neural responses within the pacemaker region of the stomach could contribute to differences in contractile patterns during mixing of gastric contents.
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