Angiotensin receptors and actions in guinea pig enteric nervous system
Guo-Du Wang,
Xi-Yu Wang,
Hong-Zhen Hu,
Xiu-Cai Fang,
Sumei Liu,
Na Gao,
Yun Xia, and
Jackie D. Wood
Department of Physiology and Cell Biology, The Ohio State University, College of Medicine and Public Health, Columbus, Ohio
Submitted 18 March 2005
; accepted in final form 24 May 2005
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ABSTRACT
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Actions of ANG II on electrical and synaptic behavior of enteric neurons in the guinea pig small intestine were studied. Exposure to ANG II depolarized the membrane potential and elevated neuronal excitability. The number of responding neurons was small, with responses to ANG II in 32% of submucosal neurons and 25% of myenteric neurons. Hyperpolarizing responses were evoked by ANG II in 45% of the neurons. The hyperpolarizing responses were suppressed by
2-noradrenergic receptor antagonists, which suggested that the hyperpolarizing responses reflected stimulation of norepinephrine release from sympathetic neurons. Exposure to ANG II enhanced the amplitude and prolonged the duration of noradrenergic inhibitory postsynaptic potentials and suppressed the amplitude of both fast and slow excitatory postsynaptic potentials. The selective ANG II1 receptor (AT1R) antagonists, ZD-7115 and losartan, but not a selective AT2R antagonist (PD-123319), suppressed the actions of ANG II. Western blot analysis and RT-PCR confirmed expression of AT1R protein and the mRNA transcript for the AT1R in the enteric nervous system. No expression of AT2R protein or mRNA was found. Immunoreactivity for AT1R was expressed by the majority of neurons in the gastric antrum and small and large intestine. AT1R immunoreactivity was coexpressed with calbindin, choline acetyltransferase, calretinin, neuropeptide Y, and nitric oxide synthase in subpopulations of neurons. The results suggest that formation of ANG II might have paracrine-like actions in the enteric nervous system, which include alterations in neuronal excitability and facilitated release of norepinephrine from sympathetic postganglionic axons. The enhanced presence of norepinephrine is expected to suppress fast and slow excitatory neurotransmission in the enteric microcircuits and to suppress neurogenic mucosal secretion.
gastrointestinal tract; enteric nervous system; myenteric plexus; submucosal plexus; sympathetic nervous system; angiotensin II; inflammation; irritable bowel syndrome
ANG II IMMUNOREACTIVITY (IR) is expressed in the enteric nervous system (ENS; see Ref. 40). Two enzymes catalyze the conversion of ANG I to ANG II, which is the biologically active form in the intestine. One of the enzymes, angiotensin-converting enzyme (ACE), is expressed in a variety of tissues and organs. In the gastrointestinal tract, ACE is localized in the brush border of the small intestinal epithelium where it might be involved in digestion of peptides, as well as the generation of ANG II (54, 67). Synaptosomal fractions from dog ileal myenteric, deep muscular, and submucosal plexuses contain ACE (4). Mast cell
-kinases are a second set of converting enzymes.
-Chymase is the major non-ACE producer of ANG II in humans and dogs (24, 10, 53). Release of
-chymase accounts for the appearance of ANG II as one of the main products associated with degranulation of mast cells (10). Significantly elevated levels of ANG II are found in mucosal biopsies from patients with Crohns colitis, which suggests that elevated levels might be associated with inflammatory states, which include mast cell hyperplasia (36).
The predictable hypertensive action of systemically administered ANG II is well known. Systemic dosing with ANG II evokes vasoconstriction and reduced blood flow in the intestinal mesenteric vasculature in parallel with whole body hypertension. Elevated vascular resistance and decreased flow in the inferior mesenteric vascular bed leads to ischemic colitis in pigs receiving pathophysiological doses of ANG II (2).
ANG II alters intestinal absorption of Na+ and H2O. Low doses of ANG II (e.g., 060 ng·kg1·min1) stimulate Na+ and H2O absorption, and higher doses (e.g., 590 ng·kg1·min1) inhibit absorption in rodents (5, 40). Stimulation of absorption is secondary to elevated release of norepinephrine from intramural sympathetic nerves in concert with suppression of neuronal reuptake of norepinephrine (55). Intracerebroventricular administration of ANG II stimulates descending spinal pathways, which activate sympathetic outflow to the bowel (9). The inhibition of absorption in response to high doses of ANG II results from stimulation of prostaglandin production (12, 19, 42).
Inhibitory actions of norepinephrine on enteric secretomotor neurons explain the action of ANG II to induce an absorptive state. Secretomotor neurons are excitatory motor neurons in the submucosal division of the ENS that innervate the intestinal crypts of Lieberkühn (13). They are uniaxonal neurons with S-type electrophysiological behavior (8, 17). Firing of secretomotor neurons releases ACh and/or vasoactive intestinal polypeptide as neurotransmitters at their junctions in the crypts. Secretomotor axons also send collaterals to innervate submucosal arterioles (1, 6). Collateral innervation of the blood vessels links blood flow to secretion by releasing ACh simultaneously at neuroepithelial and neurovascular junctions. Once released, ACh acts at the blood vessels to dilate the vessels and increase blood flow in support of stimulated secretion. Secretomotor neurons have receptors that receive excitatory and inhibitory synaptic input from other neurons in the integrative circuitry of the ENS and from sympathetic postganglionic neurons. Activation of the excitatory receptors on secretomotor neurons stimulates the neurons to fire and release their transmitters at the junctions with the crypts and regional blood vessels. The overall result of secretomotor firing is stimulation of the secretion of H2O, electrolytes, and mucus from the crypts. Elevated firing of secretomotor neurons converts the intestine in situ from an absorptive state to a secretory state with increased liquidity of the luminal contents.
Inhibitory inputs decrease the probability of secretomotor firing. The physiological effect of inhibiting secretomotor activity is suppression of mucosal secretion. Postganglionic neurons of the sympathetic nervous system are an important source of inhibitory input to the secretomotor neurons (43, 47). Norepinephrine released from sympathetic axons acts at
2a-noradrenergic receptors to inhibit secretomotor neurons. Inhibition of secretomotor firing reduces the release of excitatory neurotransmitters in the crypts. The end result is conversion to an absorptive state with reduced secretion of water and electrolytes. Suppression of secretion in this manner is postulated to be part of the mechanism by which low-dose ANG II appears to stimulate absorption in association with augmented intramural sympathetic nervous activity. We present electrophysiological results in the present report, which shows how ANG II enhances inhibitory sympathetic noradrenergic neurotransmission to enteric secretomotor neurons.
Administration of ANG II stimulates production of prostaglandins and might account for the inhibition of intestinal absorption by high doses of ANG II in vivo (19, 20, 21, 31, 41, 61). Prostaglandins, produced in the guinea pig intestine in vitro, excite submucosal secretomotor neurons and thereby stimulate mucosal secretion (22, 23). Inhibition of prostaglandin formation suppresses the secretory responses evoked by ANG II in guinea pig and rat intestine in vitro (15, 31). ANG II-evoked release of prostaglandins was reported to be enhanced, relative to noninflamed controls, in a rabbit model with experimentally induced colitis and secretory diarrhea (71).
Radiolabeled ANG II binding occurs at receptors in the small and large intestine. Binding is most dense in the colon, followed by the ileum, duodenum, and jejunum (18). Nevertheless, selective localization of the receptor subtypes in the ENS has not been studied in detail. ANG II type 1 (AT1R) and type 2 (AT2R) receptors are the receptor subtypes responsible for mediating the actions of ANG II in other tissues (11, 60). Most of the actions of ANG II are attributed to AT1R. Nevertheless, the action of low-dose ANG II to increase Na+ and H2O absorption appears to be mediated by the AT2R subtype in rat jejunum, and the action of high-dose ANG II to suppress absorption (i.e., increase secretion) and stimulate the production of prostaglandins is reported to be mediated by the AT1R receptor (37).
We tested how ANG II might act if released as a paracrine signal substance in the intestine by investigating actions of ANG II on electrical and synaptic behavior of morphologically and neurochemically identified neurons, including secretomotor and sympathetic neurons, in the guinea pig ENS. Identification of the receptor subtype responsible for the actions of ANG II was an aim of the study. A preliminary report has been published in abstract form (59).
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MATERIALS AND METHODS
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Immunohistochemistry, Western blot analysis, and RT-PCR studies were done with intestinal preparations from humans and guinea pigs. Only guinea pigs were involved in the electrophysiological studies. Preparations of human small intestinal myenteric and submucosal plexuses were obtained from: 1) subjects undergoing organ procurement after brain death resulting from trauma; 2) the adjacent discarded intestine of donor tissue at the time of kidney-pancreas transplantation; 3) adult subjects undergoing bowel resection resulting from trauma; and 4) the discarded small intestine after Roux-En-Y gastric bypass surgery. Human protocols were reviewed and approved by the Institutional Review Board of the Ohio State University Office of Research Risks Protection (Protocol 02H0208).
For the electrophysiological studies, preparations of submucosal or myenteric plexus were obtained from the ileum, jejunum, and colon of 52 male and 17 female Hartley-Dawley guinea pigs (0.30.6 kg). The guinea pigs were killed by rapid stunning and immediate exsanguination from the cervical vessels according to procedures reviewed and approved by the Ohio State University Laboratory Animal Care and Use Committee and United States Department of Agriculture veterinary inspectors. Flat-sheet preparations of myenteric and submucosal plexus (2.0 x 1.0 cm) were obtained by microdissection, as described earlier (64, 69). The preparations were pinned to Sylgard resin at the bottom of a 2.0-ml electrophysiological recording chamber that was perfused at a rate of 1015 ml/min with Krebs solution warmed to 37°C and gassed with 95% O2-5% CO2 to buffer at pH 7.37.4. The composition of the Krebs solution was (in mM) 120.9 NaCl, 5.9 KCl, 1.2 MgCl2, 1.2 NaH2PO4, 14.4 NaHCO3, 2.5 CaCl2, and 11.5 glucose. Nifedipine and scopolamine (1 µM) were added to the Krebs solution to suppress muscle movements during electrophysiological recording from the enteric neurons. Myenteric and submucosal ganglia in the electrophysiological studies were visualized microscopically with differential interference contrast optics and epilumination. Ganglia selected for microelectrode recording were immobilized with 100-µm-diameter L-shaped stainless steel wires placed on either side of the ganglion (64).
Electrophysiological recording.
Our methods for intracellular recording from neurons in the myenteric and submucosal plexuses are described in detail elsewhere (64, 69). Transmembrane electrical potentials were recorded with conventional "sharp" microelectrodes filled with 2% biocytin in 2 M KCl buffered with 0.05 M Tris at pH 7.4. Resistances of the electrodes ranged between 80 and 120 M
. The preamplifier (M-767; World Precision Instruments, Sarasota, FL) was equipped with a bridge circuit for intraneuronal injection of electrical current. Constant-current rectangular pulses were driven by a Grass SD9 stimulator (Grass Instrument Division, Astro-Med, Warwick, RI). Electrometer output was amplified and observed on an oscilloscope (Tektronics 5113; Tektronics, Beaverton, OR). Synaptic potentials were evoked by focal electrical stimulation of interganglionic fiber tracts in the myenteric or submucosal plexus with electrodes made from 20-µm Teflon-insulated Pt wire connected through stimulus-isolation units (Grass SIN5) to Grass S48 stimulators (Grass Instrument Division, Astro-Med). Stimulation of postganglionic sympathetic fibers in intestinal preparations with attached mesentery was used to release norepinephrine and evoke inhibitory postsynaptic potentials (IPSPs) in the myenteric and submucosal plexus, as described in detail elsewhere (34). Postsynaptic potentials were recorded with an expanded time scale and displayed in real-time on a digital oscilloscope (Tektronics TDS210; Tektronics) connected with a laserjet printer. Chart records were made on Astro-Med thermal recorders (Astro-Med). Amplitudes of action potentials on the chart records were blunted by the low-frequency response of the recorders. All data were recorded initially in digital format on magnetic tape for later analysis.
Histochemical methods.
The morphology of each of the neurons in the electrophysiological studies was determined by passing hyperpolarizing current (0.5 nA for 1030 min) to inject biocytin in the neuron from the microelectrode and later histochemical development of the intraneuronal biocytin. At the end of a recording session, the whole mount preparations were transferred to a disposable chamber filled with fixative containing 4% formaldehyde and 15% of a saturated solution of picric acid and stored at 4°C overnight. The preparations were cleared in three changes of DMSO and three 10-min washes with PBS and then reacted with fluorescein streptavidin (Vector, Burlingame, CA) diluted 1:200 for 30 min at 37°C and examined with a Nikon Eclipse-1000 fluorescence microscope.
Immunohistochemical studies were done on whole mounts that were incubated in 10% normal horse serum in PBS at pH 7.0 for 1 h at room temperature before exposure to the primary antisera diluted in hypertonic PBS containing 10% normal horse serum, 0.3% Triton X-100, and 0.1% sodium azide. The tissues were then placed in humidified chambers and processed for indirect single or double-immunofluorescence staining by incubating the tissue for 18 h at room temperature in rabbit or mouse anti-AT1 or anti-AT2 antiserum or a mixture of primary antibodies from different species to identify coexpression of ANG II receptors with known neurochemical codes for enteric neurons (Table 1 an Ref. 26). After incubation with the primary antibodies, the tissues were washed (3 x 10 min) in PBS, transferred to humidified chambers, and incubated at 37°C for 30 min with a single secondary antibody or a mixture of secondary antibodies conjugated with fluorescine isothiocyanate or Cy3, diluted in hypertonic PBS containing 10% normal horse serum, 0.3% Triton X-100, and 0.1% sodium azide. The tissue was then rinsed in PBS and covered with a cover slip with Vectorshield (Vector). All preparations were examined with an epifluorescence microscope (Nikon Eclipse-1000; Nikon, Melville, NY) and analyzed using filter combination that enabled separate visualization of multiple fluorophores. Digital images were obtained with a SPOT RT-cooled CCD digital camera (Diagnostic Instruments, Sterling Heights, MI) and analyzed with SPOT III software.
Western blotting.
Proteins in guinea pig and human small intestinal submucosal or myenteric plexus preparations were extracted with lysis buffer containing 50 mM Tris·HCl, pH 7.55, 5 mM EDTA, 250 mM NaCl, 1% Triton X-100, and a protease inhibitor cocktail consisting of 0.2 mM phenylmethlsulfonyl fluoride, 1 µg/ml aprotinin, 5 mM dithiothreitol, and 1 mM Na3VO4 on ice for 1 h. The ganglia were isolated by enzymatic digestion as described in detail elsewhere (65, 66). Equivalents of 50 µg extracted proteins were electrophoresed and then transferred to a 10% acrylamide gel nitrocellulose membrane (Amersham Pharmacia Biotech, Piscataway, NJ). The membranes were blocked with 5% nonfat milk in Tris·HCl-buffered saline (TBS) for 60 min. After being washed with TBS, the membranes were incubated in a solution containing primary antibodies for AT1R or AT2R for 24 h at room temperature. The membranes were then washed (3 x 10 min) with Tween 20 in TBS. Detection of AT1R or AT2R protein was done with enhanced chemiluminescence reagents from Amersham Pharmacia Biotech.
RT-PCR.
Total RNA from enzymatically dissociated ganglia was extracted with Trizol (Life Technology, Gaithersburg, MD). The tissue was homogenized in Trizol (1 ml/100 mg tissue). Chloroform (10% of the total volume) was added, and the sample was covered, shaken vigorously, and placed on ice for 10 min followed by centrifugation for 15 min at 13,000 g. The aqueous phase was then removed, and the RNA was precipitated with isopropanol (60% of the Trizol volume). The RNA pellet was then washed by resuspension in 70% ethanol. The suspension was centrifuged for 5 min (10,000 g), dried briefly, dissolved in diethyl pyrocarbonate-treated water, and divided into 1 µg/µl aliquots for further use. Reverse transcription to cDNA was started by adding the following and incubating at 42°C for 60 min: 2.0 µl of 10x RT buffer (0.1 M Tris and 0.5 M KCl, pH 8.3), 4.0 µl of 25 mM MgCl2, 2.0 µl Deoxynucleotide Mix (dATP, dCTP, dGTP, and dTTP each at 10 mM), 2.0 µl oligo-p(dT) primer (0.8 µg/µl), 50 units RNase inhibitor, 20 units of AMV RT (Roche Diagnostics, Indianapolis, IN), and 1 µl total RNA. Subsequent enzyme deactivation was accomplished by heating to 99°C for 5 min and then cooling to 4°C for 5 min. The following reaction mixture was added to 5 µl of the cDNA product: 25 µl PCR master mix (Roche Diagnostics), 2.0 µl primers, and 18 µl sterile water. AT1R or AT2R PCR primers were designed to span at least one intron. The primers used were as follows: 1) AT1R sense, 5'-CCT TGT GGT GGG AAT ATT TG-3'; 2) AT1 antisense, 5'-ATG ATG CAG GTG ACG TTG GC-3'; 3) AT2R sense, CTG AAC ATG TTT GCA AGC AT-3'; and 4) AT2R antisense, 5'-AGG TCA GAA CAT GGA AGG GA-3'.
The primers were degenerate primers. Primers 1 and 2 were designed according to human (Gene Bank no. S77410) and rat (Gene Bank no. S61896) mRNA sequence to amplify a 339-bp DNA for the AT1R. Primers 3 and 4 were designed according to human (Gene Bank no. NM_000686) and mouse (Gene Bank no. NM_007429) mRNA sequence to amplify a 454-bp DNA for the AT2R. Amplification was done with a iCycler (Bio-Rad Laboratories, Hercules, CA). PCR consisted of denaturation for 1 min at 94°C, annealing at 56°C, and extension at 72°C for 90 s. A total of 30 cycles was followed by extension for 7 min at 72°C. The PCR product (12 µl) was analyzed by electrophoresis on a 2% agarose gel. The negative control was reverse transcription without AMV RT.
Reagents and antibodies.
ANG II, 5-hydroxytryptamine, TTX, and hexamethonium were purchased from Sigma-Aldrich (St. Louis, MO). Losartan was a gift from Merck (Whitehouse Station, NJ). Saralasin, ZD-7155, and PD-123319 were from Tocris Cookson (Ellisville, MO). Fluorescein and Texas Red straptavidin were from Vector. Sources of antibodies and antisera are listed in Table 1.
Data analysis.
Results are expressed as means ± SE. Statistical significance was determined with paired Students t-test between control and experimental populations. Differences were considered significant at P < 0.05; n refers to the numbers of neurons examined. Concentration-response curves for drug-evoked responses were constructed using the following least-squares fitting routine: V = Vmax/1 + (EC50/C)
, where V is the observed response, Vmax is the maximal response, EC50 is the concentration that induces the half-maximal response, C is concentration, and nH is the apparent Hill coefficient. Concentration-response data were obtained in noncumulative fashion with washout and complete recovery preceding application of the next concentration.
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RESULTS
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Electrophysiological results were obtained with sharp intracellular microelectrodes for 117 submucosal and 87 myenteric neurons, each of which had stable resting membrane potentials more negative than 50 mV in the small intestine of 69 guinea pigs. The neurons were classified as S- or AH-type according to criteria generally applied for electrophysiological and morphological identification of enteric neurons (6, 62). S-type neurons were identified by 1) repetitive spike discharge during intraneuronal injection of depolarizing current pulses; 2) anodal-break excitation at the offset of hyperpolarizing current pulses; 3) fast nicotinic excitatory synaptic input; 4) absence of long-lasting postspike hyperpolarizing potentials; and 5) uniaxonal morphology. AH-type neurons were identified by 1) low excitability reflected by failure of depolarizing current pulses to evoke spikes or only one or a few spikes at the onset of the current pulse and absence of anodal-break excitation; 2) long-lasting postspike hyperpolarizing potentials; 3) a plateau-like "shoulder" at the onset of the repolarization phase of the action potential; and 4) multipolar Dogiel II morphology.
Application of ANG II in the superfusion solution evoked slowly activating depolarizing responses associated with decreased input resistance in 8 of 27 S-type neurons with uniaxonal morphology in the myenteric plexus and 12 of 25 neurons in the submucosal plexus (Fig. 1). The decreases in input resistance were sometimes small or undetectable. Desensitization phenomena and strong muscle contraction to concentrations >300 nM dictated that most of the work be done at concentrations between 100 and 300 nM. Nevertheless, concentration-response data over a range of 100 to 1,000 nM were obtained for 14 S-type neurons, most of which were in the submucosal plexus. The EC50 was 322.5 ± 41.1 nM for the S-type neurons (Fig. 2A).

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Fig. 1. Excitatory action of ANG II on uniaxonal neurons in the submucosal plexus of guinea pig small intestine was suppressed by ANG II type 1 receptor (AT1R) antagonists. A: application of ANG II depolarized the membrane potential without a significant change in input resistance. B: the presence of the selective AT1R antagonist, losartan, in the bathing solution suppressed the response to ANG II in the same neuron as A. C: washout reversed the action of losartan. D: morphology of the neuron from which the records in A-C were obtained. E: application of ANG II-evoked depolarization of the membrane potential in association with decreased input resistance and enhanced excitability reflected by repetitive spike discharge. F: the presence of the selective AT1R antagonist, ZD-7155, in the bathing solution abolished the ANG II-evoked response in the same neurons as E. G: after washout of ZD-7155, the presence of the selective ANG II type 2 receptor (AT2R) antagonist, PD-123319, did not suppress the ANG II-evoked response in the same neuron as E. H: morphology of the neuron from which the records in EG were obtained. Downward deflections on the voltage traces are electrotonic potentials evoked by repetitive intraneuronal injection of constant-current hyperpolarizing pulses. Decreased amplitude of the electrotonic potentials reflects decreased input resistance (i.e., increased ionic conductance).
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Fig. 2. Concentration-response relations for the action of ANG II on enteric neurons. A: concentration-response relation for the depolarizing responses to ANG II. B: concentration-response relation for the hyperpolarizing responses to ANG II.
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Exposure of AH-type myenteric neurons to 300 nM ANG II evoked slowly activating, depolarizing responses amounting to 11.64 ± 5.25 mV, which were associated with increased input resistance, augmented excitability, and suppression of the characteristic postspike hyperpolarizing potentials in 19 of 62 AH neurons in the myenteric plexus. All of the AH-type neurons had multipolar Dogiel Type II morphology.
Application of ANG II evoked, in concentration-dependent manner, slowly activating hyperpolarizing responses amounting to 14.23 ± 6.79 mV, which were associated with decreased input resistance in 26 of 87 neurons in the myenteric plexus of the small intestine and 11 of 25 submucosal neurons (Figs. 2B and 3B). Micropressure application of norepinephrine mimicked the hyperpolarizing responses to ANG II, and the
2-noradrenergic receptor antagonists, yohimbine or idazoxan, suppressed the hyperpolarizing responses (Fig. 3, AE).
Neurotransmission.
Electrical stimulation of mesenteric nerve trunks or interganglionic connectives evoked characteristic slow IPSPs in S- and AH-type neurons in the submucosal plexus (Fig. 3, G-M). The presence of the selective
2-noradrenergic receptor antagonists yohimbine or idazoxan in the bathing solution suppressed or abolished the slow IPSPs in each of 15 neurons (Figs. 3J and 4A). Application of ANG II significantly enhanced the amplitude and prolonged the duration of the slow IPSPs, and selective ATR1, but not ATR2 antagonists, suppressed or abolished potentiation of the IPSPs by ANG II (Figs. 3, GM, and 4A).

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Fig. 4. The presence of ANG II in the bathing medium enhanced stimulus-evoked noradrenergic IPSPs and suppressed stimulus-evoked slow excitatory postsynaptic potentials (EPSPs). A: ANG II enhanced the amplitude and prolonged the duration of the IPSPs and at the same time suppressed the slow EPSP. The selective AT1R antagonist, ZD-7155, but not the selective AT2R antagonist, PD-123319, suppressed the facilitative action of ANG II on the IPSPs and inhibitory action on the slow EPSPs. The 2 receptor antagonist, idazoxan, abolished the IPSPs and left the EPSPs intact. B: records with an expanded time base show stimulus-evoked fast EPSPs in the same neuron. Exposure to ANG II suppressed the fast EPSPs, and the selective AT1R antagonist, ZD-7155, reversed the action of ANG II. C: morphology of the neuron from which the results in A and B were obtained.
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Focal electrical stimulation of interganglionic fiber tracts also evoked fast excitatory postsynaptic potentials (fast EPSPs) and slow excitatory postsynaptic potentials (slow EPSPs) in addition to IPSPs. Application of 300 nM ANG II suppressed the fast EPSPs in 10 of 15 submucosal neurons (Fig. 4B). Nicotinic-like depolarizing responses to microejection pulses of ACh were unaffected by 300 nM ANG II, whereas stimulus-evoked fast EPSPs were suppressed. The presence of the AT1R antagonist ZD-7155 (13 µM) reversed the inhibitory action of ANG II on the fast EPSPs (Fig. 4B). Application of 0.3 µM ANG II also suppressed the amplitude of stimulus-evoked slow EPSPs in six of eight submucosal neurons (Fig. 4A). Suppression of the slow EPSPs occurred in parallel with enhancement of the slow IPSPs (Fig. 4A). The AT1R antagonist ZD-7155 suppressed the action of ANG II for both the slow and fast EPSPs (Fig. 4, A and B). The presence of the
2-noradrenergic antagonist idazoxan (10 µM) prevented suppression of fast EPSPs by ANG II in each of three neurons and suppression of the slow EPSPs by ANG II in five of nine neurons.
Pharmacology.
All morphologically identified neurons with depolarizing or hyperpolarizing responses to ANG II expressed IR for the AT1R ANG II receptor (see Fig. 7). Presence of the selective AT1R antagonists ZD-7155 (5 µM) or losartan (5 µM) in the bathing solution suppressed or abolished the depolarizing or hyperpolarizing responses to ANG II in each of 12 neurons, respectively (Fig. 5, A and B). A nonselective antagonist, saralasin (210 µM), also suppressed or abolished the depolarizing responses to ANG II in 16 neurons (Fig. 5A). The selective AT2R receptor agonist CGD-42112A (3 µM) did not alter the membrane potential in nine neurons. The selective AT2R antagonist PD-123319 (5 µM) did not suppress the depolarizing or hyperpolarizing responses to ANG II in 24 neurons.

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Fig. 7. Expression of AT1R-immunoreactivity (IR) in the enteric nervous system of guinea pig. A17, regional expression of AT1R-IR in the myenteric plexus of the gastrointestinal tract. B16, regional expression of AT1R-IR in the submucosal plexus of the gastrointestinal tract. C13, coexpression of AT1R-IR with anti-Hu-IR reveals expression of AT1R-IR by a majority of neurons in a submucosal ganglion. C46, coexpression of AT1R-IR with anti-Hu-IR reveals expression of AT1R-IR by a majority of neurons in a myenteric ganglion. D13, a uniaxonal neuron in a submucosal ganglion, which was filled with biocytin from the microelectrode, expressed AT1R-IR. D46, a neuron with multipolar Dogiel II morphology in a myenteric ganglion, which was filled with biocytin from the microelectrode, expressed AT1R-IR. D79, a uniaxonal neuron in a myenteric ganglion, which was filled with biocytin from the microelectrode, expressed AT1R-IR. Calibration bars are 20 µm.
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Fig. 5. Pharmacology of the depolarizing and hyperpolarizing actions of ANG II. A: the selective AT1R antagonists, losartan or ZD-7155, or the nonselective AT1R antagonist, sarlasin, each suppressed the depolarizing action of ANG II. B: the selective AT1R antagonists, losartan or ZD-7155, or the nonselective AT1R antagonist, sarlasin, each suppressed the hyperpolarizing action of ANG II. N values represent nos. of neurons studied; mean ± SE change in membrane potential evoked by 0.3 µM ANG II alone and in the presence of the various pharmacological agents is given above each bar; *significant difference (P < 0.05) from responses to ANG II alone.
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Presence of ZD-7155, a selective AT1R antagonist, offset the action of ANG II to augment stimulus-evoked slow noradrenergic IPSPs (Figs. 3 and 4). The selective AT2R antagonist PD-123319 did not alter the action of ANG II on the IPSPs (Figs. 3 and 4). Suppression of the stimulus-evoked IPSPs by idazoxan or yohimbine was supporting evidence that the IPSPs were mediated by the synaptic release of norepinephrine at
2-noradrenergic synapses (Figs. 3 and 5).
In view of evidence that actions of ANG II on intestinal secretion and motility are secondary to stimulation and formation of prostaglandins (15, 31, 71), we tested the hypothesis that ANG II stimulates prostaglandin production in the ENS. Piroxicam, a cyclooxygenase inhibitor, was used to test the possibility that the excitatory actions of ANG II were secondary to the release of prostaglandins. Piroxicam (60 µM) is known to prevent stimulation of prostaglandin formation by bradykinin and the secondary excitatory action of the released prostaglandins on neurons in the guinea pig small intestinal myenteric and submucosal plexuses (32, 33). We treated submucosal and myenteric plexus preparations with 60 µM piroxicam before application of ANG II and found no significant effects on the excitatory or hyperpolarizing actions of ANG II or on the action of ANG II to augment characteristic stimulus-evoked slow IPSPs (data not shown).
Western blot and RT-PCR.
The results of pharmacological analysis of the actions of ANG II on electrical and synaptic behavior in both myenteric and submucosal plexuses suggested that the actions were mediated by AT1R. This hypothesis was tested with Western blot analysis, which identified the presence of AT1R protein in extracts from both guinea pig and human myenteric and submucosal plexuses and from the guinea pig spinal cord and thalamus (Fig. 6A). A rabbit anti-A1R antibody (sc-1173; Santa Cruz Biotechnology, Santa Cruz, CA) recognized a protein band of 50 kDa in protein extracts from whole mount preparations of guinea pig and human myenteric and submucosal plexuses (Fig. 6A). The same antibody also identified A1R protein in extracts from guinea pig spinal cord and thalamus and in extracts from enzymatically dissociated myenteric ganglia from the guinea pig (Fig. 6A). A goat anti-A2R antibody (sc-7420; Santa Cruz Biotechnology) did not detect A2R protein in extracts from human myenteric or submucosal plexuses nor from extracts of guinea pig myenteric or submucosal plexuses.

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Fig. 6. AT1R expression in the intestine. A: Western blot analysis found evidence for expression of the receptor as a 50-kDa band in whole mount preparations of the guinea pig and human myenteric and submucosal plexuses and for dissociated myenteric ganglia, spinal cord, and thalamus from the guinea pig. B: RT-PCR analysis identified the appropriate 339-bp band for AT1R. Lane 1 is a DNA calibration ladder. Lanes 2 and 3 show the RT-PCR product obtained with specific primers for AT1R for the myenteric plexus (lane 2) and submucosal plexus (lane 3). Lane 4 is a negative control.
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RT-PCR analysis detected a 339-bp band, which identified expression of A1R receptor mRNA in extracts from enzymatically dissociated myenteric and submucosal ganglia from the guinea pig and human whole mount preparations of the myenteric and submucosal plexuses (Fig. 6B). No A2R mRNA was detected in total mRNA extracts from either the myenteric or submucosal plexus of the guinea pig or human intestine.
Immunohistochemistry.
Results for immunohistochemical localization of ANG II receptors and colocalization with common neurochemical codes for ENS neurons were obtained for 157 preparations from 68 guinea pigs. The results for localization of the AT2R receptor were consistent with the pharmacological, Western blot, and RT-PCR findings, which suggested that the AT2R is not expressed by enteric neurons. We found no IR for AT2R anywhere in the ENS with a goat anti-AT2R antibody (Table 1).
Both rabbit and mouse anti-AT1R antibodies were used to identify AT1R-IR expressed by neurons in the submucosal and myenteric plexuses. AT1R-IR was found distributed exclusively in ganglia of the myenteric and submucosal plexuses of the small and large intestine and myenteric plexus of the stomach (Fig. 7, A and B). Expression of AT1R-IR was found in 70.5% of submucosal neurons and in 65.0% of myenteric neurons in grouped data for the small and large intestine (Table 2). The proportion of the total number of neurons that expressed AT1R-IR in each of the plexuses was determined by double labeling with anti-HuC/HuD neuronal protein mouse monoclonal antibody, which labels all enteric neurons (Fig. 7C). AT1R-IR was localized to neurons with multipolar Dogiel Type II morphology and AH-type electrophysiological behavior and neurons with uniaxonal morphology and S-type electrophysiological behavior in both myenteric and submucosal plexuses of the small and large intestine (Fig. 7D).
The chemical codes for ENS neurons with AT1R-IR were studied by double labeling AT1R-IR with IR for nitric oxide synthase (NOS), neuropeptide Y (NPY), choline acetyltransferase (ChAT), calbindin, or calretinin (Figs. 8 and 9). Results for the small and large intestine were not significantly different and are combined in the presentation of the data in Table 2.

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Fig. 8. Coexpression of AT1R-IR with expression of chemical codes for guinea pig enteric neurons. A13, coexpression of AT1R-IR with calbindin in a submucosal ganglion. A46, coexpression of AT1R-IR with calretinin in a myenteric ganglion. B13, coexpression of AT1R-IR with calretinin in a submucosal ganglion. B46, coexpression of AT1R-IR with calretinin in a myenteric ganglion. Calibration bars are 20 µm.
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Fig. 9. Coexpression of AT1R-IR with expression of chemical codes for guinea pig enteric neurons. A13, coexpression of AT1R-IR with NOS in a myenteric ganglion. B13, coexpression of AT1R-IR with neuropeptide Y (NPY) in a submucosal ganglion. B46, coexpression of AT1R-IR with NPY in a myenteric ganglion. Calibration bars are 20 µm.
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In the myenteric plexus, 36% of the AT1R-IR neurons were immunopositive for calbindin, whereas only 0.51% of the AT1R-IR neurons in the submucosal plexus expressed calbindin. Of the calbindin-positive neurons in the myenteric plexus, 51.3% expressed AT1R-IR, and 6.7% of calbindin-positive neurons in submucosal ganglia expressed AT1R-IR (Fig. 8A). Calretinin-IR was found in 24.2% of the AT1R-IR neurons in the myenteric plexus and 14.4% of the AT1R-IR neurons in the submucosal plexus. AT1R-IR was localized to 89.6% of calretinin-IR neurons in the myenteric plexus and 91.79% of calretinin-IR in the submucosal plexus (Fig. 8B). In the myenteric plexus, 10.5% of the AT1R-IR neurons expressed NOS-IR (Fig. 9A). NOS-IR was found in 2% of the AT1R-IR submucosal neurons (Fig. 9A). NPY-IR was coexpressed with AT1-IR for 53% of submucosal neurons and 77% of myenteric neurons (Fig. 9B).
Most of the ChAT-IR neurons in both plexuses expressed IR for the AT1R (Fig. 10B and Table 2). AT1R-IR was expressed in 98.7% of neurons in the myenteric plexus and 91.5% of neurons in the submucosal plexus. AT1R-IR was found in 98.7% of ChAT-IR neurons in the myenteric plexus and in 91.5% of ChAT-IR neurons in the submucosal plexus.

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Fig. 10. Coexpression of AT1R-IR with expression of chemical codes for guinea pig enteric neurons. A13, coexpression of AT1R-IR with choline acetyltransferase (ChAT) in a submucosal ganglion. A46, coexpression of AT1R-IR with ChAT in a myenteric ganglion. Calibration bars are 20 µm.
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DISCUSSION
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Evidence that ACE is expressed in the brush borders of the small intestinal epithelium and together with enteric mast cell proteases catalyzes the formation of ANG II, which when released might act as a paracrine mediator in the ENS and thereby alter secretory and motility functions at the organ level, was the underlying rationale for the study (10, 24, 52, 54, 67). Release of
-chymase accounts for the appearance of ANG II as one of the main products associated with degranulation of mast cells (10). Significantly elevated levels of ANG II are found in mucosal biopsies from patients with Crohns colitis, which suggests that elevated levels might be associated with inflammatory changes that include mast cell hyperplasia (36). Furthermore, mast cell hyperplasia and elevated numbers of degranulated mast cells are found in close apposition to enteric nerves in biopsies from patients with the irritable bowel syndrome (3, 51, 53). Mast cell degranulation in these cases is expected to release chymases that catalyze formation of ANG II in close proximity to the neural elements of the ENS. Our findings suggest that ANG II, if formed in this manner, would alter the electrophysiological and synaptic behavior of the neurons that make up the integrative neural networks of the ENS and might be an underlying factor in the disordered defecation, disordered motility, and visceral sensory sensitivity associated with inflammatory states, food allergies, and subsets of the irritable bowel syndrome.
ANG II receptors.
A primary action of ANG II was excitation of neurons in both the myenteric and submucosal plexuses. The evidence supports the AT1R as the exclusive mediator of the neuronal excitatory responses. No evidence for involvement of the AT2R was found. Evidence for AT1R involvement included blockade of ANG II-evoked responses by selective AT1R antagonists and lack of effect of selective AT2R antagonists together with immunohistochemical, Western blot, and RT-PCR evidence for expression of AT1R, but not AT2R. Others reported expression of IR and mRNA transcripts for both AT1R and AT2R in mucosal samples from human colon (29). We cannot deny expression of AT2R in the mucosa because our work was focused on the ENS and the mucosa was always removed during dissection and never analyzed for receptor expression.
A discrepancy exists between the numbers of neurons expressing AT1R-IR and the numbers of neurons responding to application of ANG II. The immunohistochemical results showed expression of AT1R in 6570% of the neurons in the myenteric and submucosal plexuses. A much smaller proportion of the neurons in the electrophysiological studies (<12%) responded to application of ANG II. The discrepancy between the larger numbers of neurons expressing AT1R-IR and the smaller numbers showing electrophysiological responses might be explained by desensitization processes. In our experience, the neuronal responses to ANG II desensitized rapidly, and tachyphylaxis was often present for as long as 1 h after washout.
Desensitization of responses mediated by AT1R and AT2R is a well-known phenomenon that arises as an issue in pharmacological analysis of concentration-response relations in organ bath studies. The mechanisms of ANG II receptor internalization and thereby desensitization of AT1R- and AT2R-mediated responses include activation of protein kinase C and phosphorylation of the cytoplasmic tail of the receptor (27, 49, 56, 57).
Norepinephrine release.
ANG II acts at AT1R on sympathetic postganglionic axons to facilitate the release of norepinephrine in the heart, spleen, blood vessels, and the small intestine (16, 55, 70). Our results are consistent with the same action at sympathetic postganglionic terminals in the ENS. Most of the sympathetic innervation of the digestive tract outside of the sphincters and blood vessels enters the ENS (46, 35). Varicose sympathetic fibers project for distances of several millimeters without branching through ganglia and interganglionic connectives of the myenteric and submucosal plexuses. Dense pericellular rings of varicose noradrenergic fibers, which might reflect specific input to a specialized type of neuron, are found around a small proportion of nerve cell bodies in the myenteric plexus of the guinea pig small intestine (44).
Electrical stimulation of the noradrenergic axons in the ENS, either by placement of the stimulating electrodes on sympathetic nerves in the mesentery or on interganglionic connectives in the myenteric or submucosal plexuses, evokes slow IPSPs in the cell somas of subsets of myenteric and submucosal neurons (30, 43, 47, 68). Eighty percent of the neurons in the submucosal plexus receive inhibitory noradrenergic input, which is a marker for secretomotor neurons (48). Electrical stimulation in the myenteric plexus evokes slow IPSPs in only
9% of the neurons (38). We found that application of ANG II increased the amplitude of noradrenergic slow IPSPs in both plexuses. The finding suggests that ANG II acts at presynaptic AT1R on sympathetic axons in the ENS to enhance the release of norepinephrine. This presynaptic facilitatory action of ANG II at sympathetic synapses is reminiscent of presynaptic facilitation by CCK, which enhances the release of ACh at nicotinic synapses in the ENS of the gallbladder (45).
Neurons in the submucosal plexus, which display S-type electrophysiological behavior, uniaxonal morphology, and IR for vasoactive intestinal peptide and/or ChAT, function as secretomotor neurons (6, 8). Secretomotor neurons make up the majority of neurons in the submucosal plexus of guinea pig small intestine. Inhibition of secretomotor neurons by sympathetic noradrenergic input suppresses mucosal secretion (13). Presynaptic facilitation of norepinephrine release by ANG II would therefore facilitate suppression of mucosal secretion and "lock" the intestine in an absorptive state in parallel with suppression of intestinal blood flow. The resulting decrease in liquidity of the intestinal contents in such cases might lead to constipation.
Suppression of fast and slow EPSPs by ANG II behaved as if this action reflected activation of presynaptic inhibitory receptors, which suppressed the release of neurotransmitters for the synaptic events. Failure of ANG II to suppress responses to exogenous application of neurotransmitters (e.g., micropressure application of ACh at fast nicotinic synapses), while suppression of the stimulus-evoked synaptic event was in effect, fulfills criteria for a presynaptic inhibitory action. Blockade of the inhibitory action of ANG II on both forms of excitatory neurotransmission by
2-noradrenergic antagonists is consistent with ANG II-evoked elevation of norepinephrine in the extracellular milieu and a presynaptic inhibitory action at the excitatory synapses. Norepinephrine is known to act at presynaptic
2 inhibitory receptors to suppress both fast and slow EPSPs in the ENS (63).
Immunohistochemistry.
The results for colocalization of the AT1R in relation to common immunoreactive chemical codes for enteric neurons are not readily reconciled with the electrophysiological findings in terms of anticipated functional outcomes. Aside from inhibition of neuronal excitability by presynaptic facilitation and stimulation of norepinephrine release, the action of ANG II was excitatory, as reflected by membrane depolarization and stimulation of action potential discharge. Yet, the excitatory action was restricted to a relative small percentage of the total number of neurons that were evaluated and was complicated by a strong tendency for tachyphylaxis to occur. Approximately one-half of the neurons that expressed calbindin-IR also expressed AT1R-IR. Calbindin is a chemical code for neurons with AH-type electrophysiology and multipolar Dogiel II morphology (8, 25). AH-type neurons are believed to be interneurons synaptically connected one with another to form positive feedforward networks that connect with and drive the behavior of motor neurons to the musculature and secretory epithelium (58, 63). Excitation in members of this population of neurons would be expected to have ramifications for the functional state of the intestine that differ from stimulation of excitability in motor neurons alone.
Neurons with uniaxonal morphology, S-type electrophysiological behavior, and expression of IR for calretinin, NOS, NPY, or ChAT also expressed AT1R-IR. Selective excitation of neurons belonging to this grouping of chemical codes has broad implications for function/malfunction at the organ level. Regrettably, too little is known about specific integrative functions involving these neurons at the microcircuit level of organization to enable prediction of outcomes in terms of total organ performance. Nevertheless, some predictions can be made. For example, myenteric ganglion cells, which express NOS-IR and have their single axon projecting in the aboral direction, are inhibitory motor neurons to the circular musculature (8). Elevation of excitability in members of this group of neurons would be expected to suppress contractile activity of the musculature. Likewise, some of the uniaxonal neurons that express ChAT are excitatory cholinergic musculomotor neurons, which when activated can selectively stimulate contractile activity in the circular or longitudinal muscle coats, or can be secretomotor neurons that stimulate mucosal secretion when activated (6, 8). Any suggestion that ANG II stimulates cholinergic motor neurons to release ACh and thereby evoke muscle contraction is unsupported because contractile actions of ANG II in the guinea pig small intestine reflect direct action at the musculature and not stimulation of release of ACh (50).
Prostaglandins.
ANG II in high doses inhibits intestinal absorption, and this is mediated by stimulation of enteric prostaglandin production (19, 42, 12). In view of known excitatory actions of prostaglandins on enteric neurons and recent findings that bradykinin acts to stimulate enteric neurons to synthesize and release prostaglandins, we addressed the question of whether the excitatory actions of ANG II might also reflect stimulation of formation of prostaglandins (32, 33). Based on the finding that inhibition of prostaglandin synthesis by piroxicam failed to suppress excitatory actions of ANG II, we concluded that, unlike bradykinin, ANG II actions were not secondary to release of prostaglandins.
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GRANTS
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This work was supported by National Institute of Diabetes and Digestive and Kidney Diseases Grants RO1 DK-37238 and RO1 DK-57075 (to J. D. Wood) and KO8 DK-60468 (to Y. Xia) and by a Pharmaceutical Manufacturers of America Foundation Postdoctoral Fellowship (to S. Liu).
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FOOTNOTES
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Address for reprint requests and other correspondence: J. D. Wood, Dept. of Physiology and Cell Biology, 304 Hamilton Hall, 1645 Neil Ave., Columbus, OH 43210 (e-mail: wood.13{at}osu.edu)
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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