1Department of Internal Medicine-Gastroenterology, and 3Department of Physiology, University of Michigan, Ann Arbor, Michigan 48109; and 2Laboratory of Autonomic Neuroscience, Pennington Biomedical Research Center, Baton Rouge, Louisiana 70808
Submitted 9 July 2003 ; accepted in final form 19 August 2003
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ABSTRACT |
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brain stem; electrophysiology; gastrointestinal-receptive relaxation
In the present study, we aimed to determine at the cellular level the effects of NE on identified gastric-projecting DMV neurons.
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MATERIALS AND METHODS |
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Electrophysiology. Brain stems were removed as described previously (6, 26). Briefly, the rats were anesthetized with halothane. When a deep level of anesthesia was induced, the rat was killed by severing the major blood vessels in the chest. The brain stem was then removed and placed in oxygenated Krebs solution at 4°C (see Solution composition). The site of DiI labeling in the stomach was confirmed by visual inspection of the organ. The brain stem was used only from those animals in which the glue covering the site of DiI application was still in place at the time of the experiment. With the use of a vibratome, six to eight coronal sections (200-µm thick) containing the dorsal vagal complex (DVC) were cut and stored in oxygenated Krebs solution at 30°C for at least 1 h before use. A single slice was transferred to a custom-made perfusion chamber (volume 500 µl) and kept in place by using a nylon mesh. The chamber was maintained at 35 ± 1°C by perfusion with warmed, oxygenated Krebs solution at a rate of 2.5-3.0 ml/min.
Before electrophysiological recording, gastric-projecting DMV neurons were identified by using a Nikon E600-FN microscope equipped with epifluorescent filters suitable for visualizing DiI. Once the identity of a labeled neuron was confirmed, whole cell recordings were made under bright-field illumination by using DIC (Nomarski) optics.
Whole cell recordings were made with patch pipettes (3-8 M resistance) filled with a potassium gluconate solution (see Solution composition) by using an Axoclamp 2B single-electrode voltageclamp amplifier (Axon Instruments, Union City, CA). Recordings were made only from neurons unequivocally labeled with DiI. Data were sampled every 100 µs and filtered at 2 kHz, digitized via a Digidata 1200C interface (Axon Instruments) and acquired, stored, and analyzed on an IBM PC utilizing pClamp 8 software (Axon Instruments). Recordings were accepted only if the series resistance was <15 M
. In addition, the action potential evoked after injection of depolarizing current must have had an amplitude of at least 60 mV and the membrane potential had to return to the baseline value after the action potential afterhyperpolarization (AHP). Drugs were made fresh immediately before use and were applied to the bath via a series of manually operated valves.
Concentration-response curves were constructed from fundus-, corpus-, and antrum/pylorus-projecting neurons in which at least three concentrations of NE were tested. For measurements of the NE-induced depolarization or hyperpolarization, neurons were hyperpolarized to -65 mV via injection of direct current; for measurements of the effects of NE on the spontaneous action potential discharge rate, neurons were held at their resting potential (I = 0). When tested on spontaneously active DMV neurons, the NE-induced variation in firing rate was measured as the number of action potentials counted during the 20 s preceding the administration of the drug and during the 20 s of maximal firing rate variation after drug superfusion. Cells were defined as responders if NE (10 µM) induced a change in the membrane potential of at least 3 mV or a 20% variation in the action potential discharge rate.
Morphological reconstructions. At the end of the electrophysiological recordings, before removal of the pipette, Neurobiotin (2.5% wt/vol) was injected into the DMV (0.4 nA, 600 ms on/1,200 ms off for 10 min). After injection of Neurobiotin, the pipette was retrieved from the cell, which was allowed to seal for 10-20 min before overnight fixation at 4°C in Zamboni's fixative (see Solution composition). The diaminobenzidine-horseradish peroxidase technique used to develop the Neurobiotin stain and the protocol used for neuronal reconstruction have been described previously (6).
Immunohistochemistry. Five rats were injected with Fluoro-Gold (20 µg·ml saline-1·rat-1 ip) (Fluorochrome, Englewood, CO) 3 days before brain stem removal to label preganglionic neurons innervating the subdiaphragmatic viscera, allowing delineation of the boundaries of the DMV (11, 19, 32). Rats were anesthetized deeply (abolition of foot pinch withdrawal reflex) with halothane and perfused with 200 ml saline followed by 200 ml Zamboni's fixative.
After extraction, brain stems were fixed in Zamboni's fixative overnight, rinsed with PBS containing Triton X-100 (PBS-TX) and stored in PBS-TX overnight at 4°C. The brain stem was then placed in a 2.5% sucrose-in-PBS solution before coronal sections of 40-µm thickness were cut by using a cryostat. Every third slice was mounted onto gelatin-coated coverslips. The portion of the DMV located caudal to the obex was defined as the caudal DMV and the portion of DMV located rostral to the anterior tip of the area postrema as the rostral DMV. The area comprising the extension of the area postrema was defined as the intermediate DMV.
Slices were rinsed with fresh PBS-TX-BSA solution and incubated at 37°C for 2 h with the primary antibody (mouse--TH; 1:500 dilution in PBS-TX containing 0.1% BSA). Slices were rinsed with PBS-TX-BSA and incubated again at 37°C for 30 min with secondary antibody (goat-
-mouse FITC; Sigma, St. Louis, MO) 1:100 dilution in PBS containing 0.1% BSA. Specimens were again rinsed with PBS-TX-BSA solution before being allowed to air dry and were mounted with Fluoromount-G (Southern Biotechnology Associated, Birmingham, AL). Control experiments were carried out to ensure that the antibody labeling was selective, namely 1) incubation of primary or secondary antibodies only and 2) reaction of primary antibody with inappropriate secondary antibody. All tests proved negative, indicating that the secondary antibodies were selective for their primary antibodies and that the antibodies themselves exhibited neither nonspecific binding nor excessive autofluorescence.
Optical density measurements. Tissue sections were examined and photographed with a Nikon E400 microscope equipped with epifluorescent filters for FITC and UV, SPOT camera, and software (Diagnostic Instruments, Sterling Heights, MI). The settings of the acquisition parameters were kept constant throughout the experiments. The optical density measurements of TH-IR within the DMV were analyzed by using Image J software (developed at the National Institutes of Health and available from the internet at http://rsb.info.nih.gov/ij). Background subtraction was conducted on each specimen by measuring the optical density of an area within the visual field that did not contain TH-IR somata, usually in the NTS area 100-200 µm dorsal to the DMV. Optical density measurements were calculated by subtracting the background measurement from the DMV area analyzed and are expressed in arbitrary units of fluorescence. In each group (caudal, intermediate, and rostral DMV), a minimum of 10 slices from each rat was averaged.
Statistical analysis. Results are expressed as means ± SE. Intergroup comparisons were analyzed with one-way ANOVA followed by the conservative Bonferroni test for individual post hoc comparisons, Student's paired t-test, or 2 test. Significance was defined as P < 0.05.
Drugs and chemicals. All drugs were purchased from Sigma (St. Louis, MO); stock solutions were freshly prepared and diluted to the final concentration in Krebs solution just before use. Permount was purchased from Fisher Scientific (Pittsburgh, PA), DiI was purchased from Molecular Probes (Eugene, OR), and Neurobiotin was purchased from Vector Labs (Burlingame, CA).
Solution composition. Krebs solution was (in mM) 126 NaCl, 25 NaHCO3, 2.5 KCl, 1.2 MgCl2, 2.4 CaCl2, 1.2 NaH2PO4, and 11 dextrose, maintained at pH 7.4 by bubbling with 95% O2-5% CO2. Intracellular solution was (in mM) 128 potassium gluconate, 10 KCl, 0.3 CaCl2, 1 MgCl2, 10 HEPES, 1 EGTA, 2 ATP, and 0.25 GTP, adjusted to pH 7.35 with KOH. Zamboni's fixative was 1.6% (wt/vol) paraformaldehyde, 19 mM KH2PO4, and 100 mM Na2HPO4·7H2O in 240 ml saturated picric acid-1,600 ml H2O, adjusted to pH 7.4 with HCl. PBS-TX was (in mM) 115 NaCl, 75 Na2HPO4·7H2O, 7.5 KH2PO4, and 0.15% Triton X-100. Avidin D-horseradish peroxidase solution was 0.05% diaminobenzidine in PBS containing 0.5% gelatin supplemented with 0.025% CoCl2 and 0.02% NiNH4SO4.
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RESULTS |
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Analysis of the effects of NE on identified DMV neurons. Seventy-five percent of neurons (i.e., 170 of 228) responded to NE (1-100 µM) in a concentration-dependent manner. Response to NE was either an excitation (111 of 228, i.e., 49% of total or 65% of responsive neurons), an inhibition (59 of 228, i.e., 26% of total or 35% of responsive neurons), or an inhibition followed by an excitation (10 of 228, i.e., 4% of total or 6% of responsive neurons. These neurons have been included in the group defined as inhibited by NE). The remaining 58 (i.e., 25%) neurons did not respond to NE.
Interestingly, a similar proportion of antrum/pylorus- and corpus-projecting neurons were excited (49 and 59% of the total neurons, respectively) or inhibited by NE (20% of the total neurons for both groups). A lower percentage of fundus-projecting neurons, however, was excited (37% of the total neurons), and a larger percentage of neurons was inhibited by NE (36% of the total neurons, P < 0.05 vs. corpus- or antrum/pylorus-projecting neurons). Results are summarized in Table 1.
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In all gastric-projecting groups, the threshold for the response to NE was 1 µM and maximum current (IMAX) was obtained at 100 µM; the data have thus been pooled.
To ascertain whether the NE-induced membrane effects were due to a direct effect of NE on DMV neurons, we compared the amplitude of the NE-induced membrane depolarization in the absence and presence of the synaptic transmission blocker TTX (1 µM). In seven neurons, NE (30 µM) induced a 7.1 ± 1.05-mV depolarization that recovered to baseline on washout. After 10 min of perfusion with TTX, reapplication of NE in the presence of TTX induced a 5.4 ± 0.95-mV depolarization (i.e., 78 ± 12% of control, P > 0.05 vs. NE alone). Similarly, in five other neurons, NE induced a 11.4 ± 1.3-mV hyperpolarization that recovered to baseline on washout. After 10 min of perfusion with TTX, reapplication of NE in the presence of TTX induced a 8.8 ± 1.3-mV hyperpolarization (i.e., 81 ± 16% of control, P > 0.05 vs. NE alone; data not shown).
In eight cells in which NE (30 µM) had an excitatory effect, 10-min pretreatment with the selective 1-adrenoceptor antagonist prazosin (100 nM), which per se did not affect the basal firing rate (0.91 ± 0.17 and 0.88 ± 0.19 spikes/s in control and in prazosin, respectively; P > 0.05), antagonized the NE-induced increase in action potential firing rate from 380 ± 81% of control in NE alone to 98 ± 12% of control in prazosin + NE (P < 0.05 vs. NE alone). Similarly, in another group of neurons, pretreatment with prazosin (100 nM) attenuated the NE-induced depolarization from 9.1 ± 1.5 mV in control to 2 ± 0.5 mV after prazosin (n = 7; P < 0.05; Fig. 1). The NE-mediated depolarization was mimicked by perfusion with the selective
1-adrenoceptor agonist phenylephrine (10 µM) that induced a depolarization of 4.7 ± 0.6 mV (n = 12).
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In four neurons in which NE (30 µM) decreased the discharge rate to 2.8 ± 2.8% of control, 10-min pretreatment with the selective 2-adrenoceptor antagonist yohimbine (10 µM), which per se did not affect the basal firing rate (0.81 ± 0.13 and 0.97 ± 0.09 spikes/s in control and in yohimbine, respectively; P > 0.05), restored the firing rate to 123 ± 16% of control (P < 0.05 vs. NE alone). Similarly, in three different neurons in which NE (30 µM) induced a hyperpolarization, a 10-min pretreatment with yohimbine (10 µM) attenuated the NE-induced hyperpolarization from 9.6 ± 4 mV in NE to 2 ± 3 mV in NE + yohimbine (P < 0.05 vs. NE alone; Fig. 2). In four cells in which NE induced a hyperpolarization, perfusion with the selective
2-adrenoceptor agonist UK-14304 (1 µM) also induced a hyperpolarization of 4 ± 1.4 mV.
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To further support the selective activation of 1-adrenoceptor and
2-adrenoceptor in the excitatory and inhibitory effects of NE, respectively, we conducted the following experiments. In four neurons in which NE (100 µM) induced a depolarization, a 10-min pretreatment with yohimbine (10 µM) did not attenuate the NE-induced depolarization, which was 10.5 ± 0.5 mV in NE and 9.3 ± 1.5 mV in NE + yohimbine (P > 0.05 vs. NE alone, data not shown). Likewise, in three neurons in which NE (30 µM) induced a hyperpolarization, 10-min pretreatment with prazosin (100 nM) did not attenuate the NE-induced hyperpolarization, which was 6.4 ± 1.4 mV in NE and 6.0 ± 1.1 mV in NE + prazosin (P > 0.05 vs. NE alone, data not shown).
In an effort to investigate the possible mechanism of action, we tested the effects of NE (30 µM) on the shape of the action potential on neurons depolarized by NE. Cells were clamped at -55 mV by injection of direct current and then injected with a 10-ms-long pulse of depolarizing current sufficient to evoke a single action potential at its offset. In control conditions, the amplitude of the AHP was 16.6 ± 0.71 mV (n = 13). After NE (30 µM) perfusion, current was injected to return the membrane potential to baseline values. In the presence of NE, the amplitude of the action potential AHP was reduced to 14.6 ± 0.7 mV (P < 0.05 vs. control). Similarly, the AHP kinetic of decay () was 69 ± 9 ms in control and 55 ± 7 ms in the presence of NE (n = 13, P < 0.05 vs. control; Fig. 3). Conversely, in neurons in which application of NE (30 µM) hyperpolarized the DMV membrane, the AHP and its
were unaltered by NE (16.4 ± 1 mV and 66 ± 16 ms and 15.3 ± 1 mV and 54 ± 10 ms in control and NE, respectively; n = 7; P > 0.05 vs. control; Fig. 3).
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Analysis of the location of TH-positive neurons. Distribution of TH-IR within the DMV was analyzed by using optical density measurements. Compact neuronal organization and the presence of TH-IR-positive somata prevented a compartmentalized analysis of the caudal portion of the DMV. In fact, the optical density of TH-IR yielded similar values throughout the caudal DMV (20 ± 2 arbitrary units, n = 5). Conversely, in both the intermediate and rostral portions of the DMV, areas of different optical density could be distinguished throughout the rostrocaudal extent of the DMV. An area of low optical density (14 ± 1.5 and 10.4 ± 2.2 arbitrary units in the intermediate and rostral DMV, respectively) was present in the central portion of the DMV; an area of high optical density (22.7 ± 2 and 20.6 ± 1.1 arbitrary units in the intermediate and rostral DMV, respectively) was present close to the NTS, hypoglossal nucleus, and central canal (Fig. 4).
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A schematic representation of the DMV was then drawn in which we divided the nucleus according to the high- and low-optical-density areas; the localization of the DMV cells in relation to their electrophysiological responses to NE (i.e., excitation, inhibition, or not responsive) was then superimposed on the schematic. We found no apparent relationship between the optical density and a particular response to NE (Fig. 5).
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DISCUSSION |
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The following evidence supports our data. Perfusion with NE induced a direct membrane response in the vast majority of DMV neurons. The most common response, observed in 60% of the responsive neurons, consisted of a concentration-dependent excitation. A large subgroup of DMV neurons,
35% of the responsive ones, was hyperpolarized by perfusion with NE, whereas only a limited fraction of the responsive neurons,
5% of the total, responded to NE with an initial hyperpolarization followed by a depolarization. Interestingly, we observed that DMV neurons projecting to either the antrum/pylorus or the corpus had a similar incidence of neurons responding to NE with either excitatory or inhibitory responses. Fundus-projecting neurons, however, had a significantly lower percentage of excitatory responses and a higher percentage of inhibitory responses compared with neurons projecting to antrum/pylorus or corpus. Our data confirm and extend the previous observation reported by Fukuda et al. (9), who described similar percentages of NE-responsive neurons.
It is interesting to note that the concentration-response curve of the NE-induced decrease in firing rate shifted to the left relative to that of the NE-induced membrane hyperpolarization. We take this shifted concentration response curve as an indication of the major sensitivity of DMV neurons to inhibitory synaptic inputs utilizing catecholamines and, in general, other inhibitory neurotransmitters.
With the use of agonists and antagonists selective for subtypes of -adrenoceptors, we demonstrated that the excitatory response to NE is mediated by interaction with
1-adrenoceptors and that the inhibitory response to NE is mediated by interaction with postsynaptic
2-adrenoceptors. Our data confirm the conclusions reached by Fukuda et al. (9), which suggested
1- and
2-adrenoceptors as the receptors responsible for the actions of NE on unidentified DMV neurons.
Our data suggest that, at least in part, the excitatory effects of NE are due to a reduction of the amplitude and kinetics of decay of the action potential AHP. An excitatory effect on vagal motor neurons determined by a reduction of the AHP is similar to our recent reports on the effects of corticotropin-releasing factor and thyrotropin-releasing hormone in DMV neurons (15, 27) and similar to other reports of the effects of NE in cardiopulmonary vagal motor neurons (18) or in hypothalamic neurons (30). Our data also suggest that the calcium-dependent potassium current underlying the AHP might contribute to the resting membrane potential. Interestingly, although Fukuda et al. (9) suggested that the NE-mediated hyperpolarization of DMV neurons was mediated by an increase in a potassium-sensitive current, we did not observe any significant variation of the action potential AHP in cells that were inhibited by NE. It is reasonable to assume that, given that the complement of membrane currents present in DMV neurons is dependent on the peripheral projection area (6) and that Fukuda et al. (9) conducted their recordings on unidentified DMV neurons, the cells from which Fukuda et al. (9) recorded projected to areas other than the gastric areas targeted in the present investigation.
DMV is a brain stem area that contains a very pronounced network of TH-IR fiber terminals (3, 14, 20) and is possibly the nucleus with the highest density of TH-IR-positive fibers within the brain stem (20). Although we cannot say with certainty that all of the TH-IR-positive terminals are catecholaminergic, relatively recent data would suggest a 1:1 distribution of TH and dopamine--hydroxylase (31), thus suggesting that NE is the neurotransmitter involved in these synaptic contacts. Optical density measurements we report in the present study clearly demonstrate that within the DMV, distribution of the TH-IR-positive fibers is nonhomogeneous. In fact, a careful examination of the figures presented in the work by previous investigators (3, 14) clearly shows areas of lesser density of TH-IR-positive fibers located along the same mediolateral extension reported here. Given that these areas represent approximately one-fourth of the DMV, and a similar percentage of DMV neurons did not respond to NE, we investigated whether the NE-nonresponsive neurons were localized in the areas that showed a low density of TH-IR-positive fibers. Our data, however, show that this is not the case. In fact, the type of response, or lack thereof, to NE did not correlate to any particular location of the DMV neurons.
Physiological significance. Several authors (9, 14, 17, 24) have put forward the idea of a possible role of catecholamines in the modulation of brain stem vagal circuits. The majority of local synaptic inputs onto DMV neurons arises from the adjacent NTS, which contains the A2 catecholaminergic group (14). With the noticeable exception of the esophageal-mediated gastric relaxation (21, 28), the vast majority of vagovagal reflexes induce a powerful inhibition of the DMV firing rate via activation of inhibitory NTS neurons, which then inhibit DMV vagal output (4, 7, 8, 10, 13, 16, 22, 28, 29). An inhibitory effect on gastric tone and motility, however, can also be attained by stimulation of DMV neurons forming inhibitory nonadrenergic, noncholinergic pathways (12, 28). Indeed, we have shown that repetitive esophageal stimulation induces gastric inhibition via activation of neurons in the subnucleus centralis of the NTS (cNTS) and both excitation and inhibition of DMV neurons (21). More recently, we (23) have shown that the vast majority of the cNTS neurons are TH-IR positive and that administration on the surface of the fourth ventricle of both 1- and
2-antagonists significantly attenuates the esophageal distension-induced gastric relaxation. We would therefore like to suggest that part of gastric relaxation induced by esophageal distension is mediated by NE acting on
1- and
2-adrenoceptors present on the membrane of gastric-projecting DMV neurons and that part of the excitatory response observed following NE perfusion is due to an effect on the action potential AHP.
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ACKNOWLEDGMENTS |
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GRANTS
This work is supported by National Institute of Diabetes and Digestive and Kidney Diseases Grants DK-55530 and DK-56373.
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FOOTNOTES |
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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. Section 1734 solely to indicate this fact.
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REFERENCES |
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