Expression and effects of metabotropic CRF1 and CRF2 receptors in rat small intestine

Christophe Porcher,1 Aurélie Juhem,1 André Peinnequin,2 Valérie Sinniger,1 and Bruno Bonaz1

1Groupe d'Etude du Stress et des Interactions Neuro-Digestives, Equipe d’Accueil 3744, Department of Gastroenterology, Centre Hospitalier Universitaire de Grenoble, Grenoble; and 2Department of Radiobiology and Radiopathology, Centre de Recherches du Service de Santé des Armées, La Tronche, France

Submitted 9 July 2004 ; accepted in final form 4 January 2005


    ABSTRACT
 TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Corticotropin-releasing factor (CRF)-like peptides mediate their effects via two receptor subtypes, CRF1 and CRF2; these receptors have functional implication in the motility of the stomach and colon in rats. We evaluated expression and functions of CRF1 and CRF2 receptors in the rat small intestine (i.e., duodenum and ileum). CRF1–2-like immunoreactivity (CRF1–2-LI) was localized in fibers and neurons of the myenteric and submucosal ganglia. CRF1–2-LI was found in nerve fibers of the longitudinal and circular muscle layers, in the mucosa, and in mucosal cells. Quantitative RT-PCR showed a stronger expression of CRF2 than CRF1 in the ileum, whereas CRF1 expression was higher than CRF2 expression in the duodenum. Functional studies showed that CRF-like peptides increased duodenal phasic contractions and reduced ileal contractions. CRF1 antagonists (CP-154,526 and SSR125543Q) blocked CRF-like peptide-induced activation of duodenal motility but did not block CRF-like peptide-induced inhibition of ileal motility. In contrast, a CRF2 inhibitor (astressin2-B) blocked the effects of CRF-like peptides on ileal muscle contractions but did not influence CRF-like peptide-induced activation of duodenal motility. These results demonstrate the presence of CRF1–2 in the intestine and demonstrate that, in vitro, CRF-like peptides stimulate the contractile activity of the duodenum through CRF1 receptor while inhibiting phasic contractions of the ileum through CRF2 receptor. These results strongly suggest that CRF-like peptides play a major role in the regulatory mechanisms that underlie the neural control of small intestinal motility through CRF receptors.

astressin2-B; corticotropin-releasing factor receptors; urocortin


CORTICOTROPIN-RELEASING FACTOR (CRF) is a 41-amino acid peptide recognized as a major regulator of pituitary adrenocorticotropin (54). This neuropeptide is the principal mediator of a wide variety of physiological responses caused by sustained stresses (2, 9, 22, 51). The paraventricular nucleus (PVN) of the hypothalamus is the main source for CRF in the brain (49). Recently, the addition of three mammalian neuropeptides, urocortin (Ucn) 1, Ucn 2, and Ucn 3, have expanded the family of CRF-related peptides (18).

The CRF-like peptides mediate their effects via two receptor subtypes, CRF1 and CRF2{alpha}/CRF2{beta}, each encoded by distinct genes (6, 8, 27, 41). Both receptors belong to the seven-transmembrane domain family positively coupled to adenylate cyclase via G proteins (1, 3, 12). CRF1 receptor mRNA is predominantly expressed in the brain under basal conditions (4, 45) but is increased in the PVN in acute stress (4), somatovisceral pain (50), and colitis (42). The distribution of the CRF2 receptor variants differs by both tissues and species. Thus CRF2{alpha} is the predominant neuronal receptor subtype within the brain, whereas the CRF2{beta} form is localized in nonneuronal elements like the choroid plexus and cerebral blood vessels (28). In the periphery, CRF2{beta} receptor mRNA is expressed in both cardiac and skeletal muscles, with lower levels observed in both lung and intestine tissues in rat (28, 41). In humans, CRF2{alpha} is the major isoform in brain, heart, and skeletal muscle tissues (53). This heterogeneous distribution of CRF1 and CRF2 receptor mRNAs suggests distinctive functional roles for each receptor.

In vitro studies established that Ucn 1 binds with high affinity to both CRF receptor subtypes, whereas CRF exhibits higher affinity to CRF1 than CRF2 receptors (40). In situ hybridization studies showed that CRF2 mRNA was localized in the submucosal layer and over cells at the base of the villi in the mouse duodenum (41). CRF1 and CRF2 mRNA were also detected in lamina propria mononuclear cells of the human colonic mucosa (26). A recent immunochemical study indicates that both CRF1 and CRF2 are distributed throughout the oxyntic gland and submucosal blood vessels within the stomach corpus (7).

Functional studies demonstrated that central or peripheral injection of CRF or Ucn 1 inhibits gastric emptying in rats, mice, and dogs (27, 31, 37, 38, 56), influences small intestinal transit and motility in rats (21, 58), and stimulates colonic transit and defecation in rats and mice (29–31, 57). In vivo experiments on rats and mice showed that the selective CRF1 antagonists CP-154,526 and NBI-27914 prevented CRF action on the colon without influencing the gastric response (31, 57), whereas the CRF2 antagonists antisauvagine-30 and astressin2-B blocked CRF induced inhibition of gastric motility and had no effect on colonic response (32). These data suggest that CRF and Ucn 1 delay gastric emptying through CRF2, whereas the activation of CRF1 stimulates colonic motility. At the cellular level, intracellular recording of enteric neuronal activity in the guinea pig small intestine has demonstrated that CRF excites myenteric neurons by stimulation of adenylate cyclase and elevation of cAMP (15). In addition to their role in gut motility, CRF receptors are also involved in intestinal permeability and in stress-related abnormalities of mucosal functions (47, 52).

From these data, it appears that the distribution and functional implication of CRF receptors on intestinal motility remain complex and not fully understood. Consequently, the aims of this study were 1) to investigate the expression of CRF1 and CRF2 in the duodenum and ileum and 2) to examine their role in small intestinal motility.


    MATERIALS AND METHODS
 TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Animals. Adult male Sprague-Dawley rats (Charles River Laboratory) weighing 200–250 g were used for the experiments. The rats were individually housed under controlled conditions of illumination (12:12-h light-dark cycle starting at 7 AM), humidity (60–70%), and temperature (22°C) with food and water available ad libitum. Animals were allowed a minimum of 7 days to adapt to housing conditions before any manipulation. Animal care and experiments were conducted in accordance with the National Institutes of Health Guide for the Care and Use of Laboratory Animals (NIH Pub. No. 80–23; revised 1978) and the legal French Ministry of Agriculture (authorization number 38 04 48). All procedures were performed according to protocols that were approved by the University of Grenoble Institutional Animal Care and Use Committee.

Tissue preparation and processing for immunohistochemistry. Animals were anesthetized with pentobarbital sodium (60 mg/kg ip; Sigma, St. Louis, MO). The abdomen was cut open, and the proximal duodenum and terminal ileum were removed for further dissection and washed of luminal contents with Krebs-Ringer buffer (KRB). KRB contained (in mM) 137.4 Na+, 5.9 K+, 2.5 Ca2+, 1.2 Mg2+, 134 Cl, 15.5 HCO3, 1.2 H2PO4, and 11.5 dextrose. This solution had a pH of 7.3–7.4 at 37.5°C when bubbled with 97% O2-3% CO2. Whole mount preparations (duodenum and ileum) and cryostat sections were generated to locate CRF receptors in the tissue by immunohistochemistry. After the removal of luminal contents, tissues to be sectioned were immediately fixed in cold acetone (for 20 min at 4°C) or in paraformaldehyde buffer (4% wt/vol in 0.1 M borax buffer, pH 7.4, overnight at 4°C). After fixation, tissues were washed in 0.1 M potassium PBS (KPBS, pH 7.4) and gradually infiltrated with increasing sucrose concentrations, 5–25%, for 1 h, each followed by 30% in KPBS overnight. Tissues were finally embedded in a 1:1 mixture of Tissue-Tek optimum cutting temperature compound (Miles, Elkhart, IN) and 30% sucrose in KPBS (vol/vol, pH 7.4) and frozen in isopentane precooled in liquid nitrogen. Tissues were cryostat cut (14-µm transverse sections) and mounted on poly-L-lysine-coated slides.

For whole mount preparations, fresh tissues were opened along the mesenteric border and pinned out in dishes coated with Sylgard silicone elastomere (Dow Corning) with the mucosal side facing up. The tissues were stretched to 110% of the original length and width, and the mucosa was removed by sharp dissection. The remaining tunica muscularis was fixed as described above. After fixation tissues were washed in KPBS (3 times for 10 min) before incubation in primary antibody.

Tissue sections and whole mount preparations were preincubated in a solution of KPBS containing 5% normal donkey serum (NDS), 0.3% Triton X-100, and 1% BSA for 1 h at room temperature before the addition of primary antibodies. For double immunostaining, tissues were incubated in each primary antibody for 12 h in a sequential manner at 4°C with a KPBS wash between steps. Incubations for double labeling were performed on tissues that were initially incubated in goat polyclonal antibodies directed against CRF1 (1:200; Santa Cruz Biotechnology) or CRF2 (1:400; Santa Cruz Biotechnology). Secondary incubations were carried out for an additional 12-h period at 4°C with mouse monoclonal antibodies directed against either 5-HT (1:200; DAKO, Glostrup, Denmark), or synaptophysin (Syn, 1:500; DAKO). Immunoreactivity for CRF1 and CRF2 was detected with donkey anti-goat secondary antibody coupled to Alexa Fluor 488 (1:200; Molecular Probes, Leiden, The Netherlands). Immunoreactivities for 5-HT and Syn were detected with Alexa Fluor 594-conjugated secondary antibodies (1:200; Molecular Probes). All the antisera were diluted with 1% NDS in 0.1 M KPBS (pH 7.4). Secondary antibody incubations were performed for 1 h at room temperature. Control tissues used to determine the level of nonspecific staining included tissues incubated without primary antibody and/or with the primary antibody blocked by overnight preincubation with the corresponding peptide (CRF1 sc-12383p and CRF2 sc-1826p, 10 µg/ml; Santa Cruz Biotechnology) before incubation of the antibody with the tissue. In both cases, no specific staining was observed (see Figs. 1B and 2B). Tissues were mounted on microscope slides in an aqueous mounting medium (Vector Laboratories, Burlingame, CA). Some cryostat sections were mounted in a mounting medium with 4',6-diamidino-2-phenylindole (DAPI) to label nuclei (Vector Laboratories). Tissues and sections were examined with a Zeiss confocal microscope (Zeiss LSM 510) and excitation wavelengths appropriate for Alexa Fluor 488 (495 nm), Alexa Fluor 594 (590 nm), and DAPI (365 nm). Confocal micrographs shown here are digital composites of Z-series scans of 5–15 optical sections through a depth of 4–20 µm. Final images were constructed with MetaMorph 4.6 software (Universal Imaging).



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Fig. 1. Corticotropin-releasing factor type 1 (CRF1) receptor-like immunoreactivity (CRF1-LI) in cryostat sections and whole mounts of the duodenum (A, D, G, and H) and ileum (E and F). CRF1-LI is found in the myenteric (arrows, A and E) and submucosal (arrowheads, E) ganglia, in the circular muscle layer and the deep muscular plexus (double arrow, A). Note also the presence of CRF1-LI in the septa (double arrow, E). CRF1-LI was observed in both nerve cell bodies, and processes occurred in the myenteric (arrowheads, C) and submucosal (arrowheads, F) ganglia. CRF1-LI of a Dogiel type II nerve cell bodies with a smooth contour labeled in the myenteric ganglia (D). CRF1-LI was also observed in mucosal cells of the duodenum (arrowheads, G and H). Preabsorption with the synthetic peptide abolished the CRF1 staining, demonstrating the specificity of the antibody to the tissue staining in cryostat sections (B). lm, Longitudinal muscle; cm, circular muscle; sm, submucosa. Scale bars: 100 (A, E, F), 40 (B, D, G, H), and 50 (C) µm.

 


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Fig. 2. CRF2 receptor-like immunoreactivity (CRF2-LI) in cryostat sections and whole mounts of the duodenum (E and F) and ileum (A, D, and C). CRF2-LI is found in the circular muscle layer and the deep muscular plexus (double arrows, A). CRF2-LI was observed in myenteric (arrow, A) and submucosal (arrowhead, A) ganglia, in nerve fibers (arrows, E), and sometimes throughout the cytoplasm of neuronal cells (arrow, D). CRF2-LI was observed in varicose nerve fibers running parallel to smooth muscle cells (F). In the duodenum, CRF2-LI was also detected in mucosa (arrowheads, C). Preabsorption with the synthetic peptide abolished the CRF2 staining, demonstrating the specificity of the antibody to the tissue staining in cryostat sections (B). Scale bars: 100 (A, D), 40 (B, E), and 50 (C, F) µm.

 
Total RNA extraction and reverse transcription reaction. Segments of duodenum and ileum were dissected and placed into KRB. Tissues were pinned to the Sylgard elastomer floor of a dissecting dish with the mucosa facing upward, and the mucosa and submucosa were removed by sharp dissection. Small strips of longitudinal and circular muscles and mucosa were placed in RNAlater (Qiagen) for 24 h at 4°C and stored in a –20°C freezer until being used. mRNA was isolated from tissue with a MagnaPure LC mRNA Isolation Kit II (Roche Applied Science, Mannheim, Germany). Seven milligrams of tissue was disrupted with 2-mm tungsten carbide beads in 300 µl of lysis buffer with a Mixer Mill MM300 (Rescht, Haan, Germany) for 40 s (30 Hz). Isolation was then carried out in a MagnaPure LC instrument per the manufacturer's instructions. mRNA was eluted in a 50-µl final volume. Reverse transcription was carried out in a 20-µl final volume from 8 liters of mRNA eluate with an avian myeloblastosis virus-based First Strand cDNA Synthesis kit (Roche Applied Science, Mannheim, Germany) with 1.6 µg oligo(dT)15 primer, according to the manufacturer's instructions.

Primer design. Oligonucleotide primers were synthesized at Eurogentec (Saraing, Belgium). Primer design and optimization regarding primer dimer, self-priming formation, and primer melting temperature were done with MacVector software (Accelrys, San Diego, CA). Specificities of the PCR amplification were documented with LightCycler melting curve analysis. Melting peaks obtained either from RT product or from specific recombinant DNA were identical. Selected forward (F) and reverse (R) primers are as follows: common region of CRF1A and CRF1B F: 5'-TGCCTGAGAAACATCATCCACTGG-3', R: 5'-TAATTGTAGGCGGCTGTCACCAAC-3', generating a 146-bp DNA fragment (bp 445–590 from GenBank L25438 [GenBank] ); common region of CRF2A and CRF2B F: 5'-AACGGCATCAAGTACAACACGAC-3', R: 5'-CGATTCGGTAATGCAGGTCATAC-3', generating a 142-bp DNA fragment (bp 420–561 from GenBank U16253 [GenBank] ); and cyclophylin A (CYCA) F: 5'-TATCTGCACTGCCAAGACTGAGTG-3', R: 5'-CTTCTTGCTGGTCTTGCCATTCC-3', generating a 127-bp DNA fragment (bp 381–507 from GenBank M19533 [GenBank] ).

Real-time quantitative RT-PCR. PCR was carried out with the LC Fast Start DNA Master SYBRgreen kit (Roche Applied Science), using 0.5 µl of cDNA (equivalent to 28 ng of organ) in a 20-µl final volume, 3 mM (CYCA) or 4 mM (CRF1–2) MgCl2, and each primer at 0.4 µM (final concentration). Quantitative PCR was performed with a LightCycler (Roche Applied Science) for 45 cycles of 95°C for 20 s, 58°C (CYCA, CRF2) or 60°C (CRF1) for 5 s, and a final step of 10 s at 72°C. The threshold cycle (CT) value, corresponding to the PCR cycle number at which fluorescence was detected above threshold, was calculated with LightCycler Software v.3.5 (Roche Applied Science), using the second derivative maximum method. Quantification was achieved by using a pool of cDNA samples as calibrator according to the comparative CT method (26) with CYCA as reference gene. CYCA was chosen because of the stability of its expression level among numerous tissues in control animals (10, 43). Relative mRNA values were calculated with RealQuant software (Roche Applied Science).

Isometric force measurements. Strips of duodenum and ileum muscles (~1 cm in length) were cut along the longitudinal axis of the circular muscle layer, and the mucosa was removed to avoid another source of endogenous CRF-like peptides. Muscles were attached to a fixed mount at one end and to a TRI 202P isometric strain gauge at the other (Letica, Barcelona, Spain) and immersed in 5-ml organ baths maintained at 37 ± 0.5°C with oxygenated KRB. A resting force of 1.0 g was applied to intestinal muscles to obtain the most reproducible responses. After the muscle strips were mounted, a 1.5-h equilibration period, during which the bath was constantly perfused with fresh KRB, was allowed before experiments began. After the 1.5-h equilibration period, the effects of various pharmacological reagents were investigated. Each addition of CRF or Ucn 1 was tested over a 5-min time period. Antagonists were added to the bath at least 10 min before application of agonists. After each experiment, tissues were constantly perfused with fresh KRB for 20 min to recover control spontaneous mechanical activities. If this latter condition was not reached, the muscle strip was not used for further experiments. Drugs were diluted to the desired concentrations in a reservoir before perfusion into the organ bath. Contractile data were digitized and stored in a Macintosh computer with Acknowledge software (MP 100; Biopac Systems).

Measurement of intestinal contractions. Intestinal motility was quantified by measuring the fraction of time occupied by contractions in 3-min periods. In some experiments, the amplitude of intestinal contractions was determined. Biopac Acknowledge software (Biopac Systems) was used to measure the amplitude of each contraction before and during drug perfusion. Amplitudes were added and divided by the total number of contractions. The mean amplitude was expressed as a percentage for each group of experiments, where 100% represented the maximum amplitude of contractions during drug exposure. Data are presented as means ± SE, and differences between groups were evaluated with Student's t-test. Differences were considered significant at P < 0.05.

CRF receptor-selective antagonists. CP-154,526-01 (gift from Pfizer R&D), SSR125543Q (gift from Sanofi-Synthélabo R&D) and astressin2-B (gift from Dr. Jean Rivier, Salk Institute, La Jolla, CA) were dissolved in DMSO made up in distilled water (pH 1.5) as a stock solution of 10–2 M. CP-154,526 and SSR125543Q are specific inhibitors of CRF1 (14, 20). Astressin2-B is a specific inhibitor of CRF2 (40). CP-154,526, SSR125543Q, and astressin2-B were added to the muscles at a final concentration of 1 µM. CRF and Ucn 1 (Sigma) were dissolved in distilled water at a concentration of 10 mM and diluted to 1 µM before being added to the organ bath. In some experiments, TTX (1 µM; Sigma) was added to the superfusion medium 10 min before CRF application.


    RESULTS
 TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Distribution of CRF1 immunoreactivity. CRF1-LI was found in a subpopulation of enteric nerve cell bodies and nerve terminals within myenteric and submucosal ganglia of the rat duodenum and ileum. CRF1-LI cell bodies were observed in the myenteric (Fig. 1, A and C) and submucosal (Fig. 1F) plexus of the intestine. In somas of enteric neurons, CRF1-LI was distributed throughout the cytoplasm; the nuclei were not stained. The number of CRF1-positive neurons in myenteric ganglia of the duodenum and ileum ranged from 3 to 15 and from 1 to 10, respectively; the average was 7.4 ± 3.5 and 5.8 ± 3.6, respectively (n = 15 ganglia in whole mount preparations from 3 animals). The number of CRF1-positive neurons in submucosal ganglia of duodenum and ileum ranged from 1 to 6 and from 1 to 5, respectively; the average was 3.7 ± 2.4 and 3.6 ± 2.1, respectively (n = 10 ganglia in whole mount preparations from 3 animals). The neuron cell bodies exhibited a strong and uniform cytoplasmic labeling and showed some processes similar to those displayed by peripheral choline acetyltransferase (36)-immunoreactive neurons (Fig. 1, C, D, and F). These cells were large and oval, with smooth surfaces and one or more long, smooth, tapering immunoreactive process (Fig. 1D).

In cross sections of the external muscle layers, CRF1-LI was present in nerve fibers of the duodenum and ileum. These nerve fibers were abundant in the circular muscle layer, whereas they were rare in the longitudinal layer (Fig. 1A). Some of these fibers were in nerve bundles emerging from the myenteric plexus and running toward the mucosa or the submucosal plexus (Fig. 1E). The CRF1-LI material showed a pattern of distribution in the proximal part of the small intestine similar to that observed in the distal part of the small intestine. In cross sections of the small intestine, CRF1-LI was detected in some mucosal cells (Fig. 1, G and H). Abolition of CRF1 immunostaining by preadsorption with an excess of the corresponding synthetic peptide demonstrated the specificity of the tissue staining (Fig. 1B).

Distribution of CRF2 immunoreactivity. CRF2-LI was found predominantly in nerve fibers and in enteric neurons within myenteric and submucosal ganglia of the duodenum and ileum (Fig. 2, A, D, and E). In the myenteric plexus of both regions studied, CRF2-LI was found in the soma of some neurons; the nuclei were not stained (Fig. 2D). Numerous CRF2-LI nerve fibers were observed in both myenteric and submucosal plexus, as well as in the interconnecting nerve strands. These processes were varicose and frequently encircled neuronal cell bodies of non-CRF2-LI neurons (Fig. 2E) and sometimes CRF2-LI neurons (Fig. 2D).

In the circular and longitudinal muscle layers of the duodenum and ileum, CRF2-LI was localized in varicose processes in nerve fibers that ran parallel to smooth muscle cells of the circular and longitudinal muscle layers (Fig. 2F). In cross sections of the small intestine, CRF2-LI was detected in the soma of cells distributed in the mucosa. CRF2-LI cells were intensely stained and were scattered among the epithelial cells along the villi (Fig. 2C). Abolition of CRF2 immunostaining by preadsorption with an excess of the corresponding synthetic peptide demonstrated the specificity of the tissue staining (Fig. 2B).

Double labeling of CRF1- or CRF2-like immunoreactivity with Syn or 5-HT. Further studies were done to chemically identify the cells and structures in the rat small intestine that display CRF1- or CRF2-LI, but labeling of CRF1 with the Q20 antibody requires fixation with acetone. Antibodies used in this study to colocalize CRF1 with 5-HT immunoreactivity did not perform well on acetone-fixed tissues.

Double-label immunofluorescence was used to determine whether or not CRF1- or CRF2-LI structures display markers expressed in nerve terminals, such as Syn. In addition, we examined the colocalization of CRF2 with 5-HT. CRF1- or CRF2-LI was visualized with Alexa Fluor 488, whereas the other markers were visualized with Alexa Fluor 594.

In the rat duodenum and ileum, Syn-like immunoreactive (Syn-LI) structures were abundant in enteric ganglia and localized on nerve processes that displayed CRF1- or CRF2-LI (Fig. 3, A–I). In the external muscle layers, colocalization of Syn-LI and CRF1- or CRF2-LI was found to occur in many of the nerve fibers. In whole mount preparations, colocalization of Syn-LI and CRF2-LI was observed in the varicosity of the internodal connecting strands (Fig. 3I).



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Fig. 3. CRF1 (A)- and CRF2 (D and G)-LI (green), synaptophysin (Syn)-like immunoreactivity (Syn-LI; red, B, E, and H), and their colocalization (yellow, C, F, and I) in cryostat sections and whole mounts of the small intestine. Colocalization of Syn-LI and CRF1–2-LI is found in the enteric nervous system, at the level of the myenteric (arrows, A–F) and submucosal (arrowheads, A–C) ganglia, in the circular muscle layer, and in the deep muscular plexus. Colocalization of CRF2-LI and Syn-LI varicose nerve fibers in the muscle layers is also shown (arrowheads, G–I). CRF2-LI is expressed in a subpopulation of mucosal cells (arrowheads, J–L) and in fibers (arrows, J and L) encircling the intestinal crypts. CRF2-LI (green, J), 5-HT-like immunoreactivity (5-HT-LI; red, K), and nuclear staining (DAPI; blue, L) in the same transverse section is also shown. None of the CRF2-LI cells (arrowheads, J) was immunopositive for 5-HT-LI endocrine cells (double arrowheads, K and L). Scale bars: 70 (A–F) and 35 (G–L) µm.

 
In the mucosa, CRF2-LI was detected in fibers encircling the intestinal crypts. In addition, mucosal cells were labeled with CRF2-LI throughout the small intestine. Nuclei, which were labeled with DAPI, were immunonegative (Fig. 3J). Enterochromaffin cells were identified by immunostaining with 5-HT. Nuclei, which were labeled with DAPI, were immunonegative (Fig. 3K). None of these enterochromaffin cells was immunopositive for CRF2-LI (Fig. 3L).

Relative quantitative expression of CRF1–2 transcripts. To confirm the expression of CRF1–2 mRNA by the rat small intestine, we used gene-specific primers and the highly sensitive technique of real-time RT-PCR (5). The LightCycler (Roche Applied Science) was used for accurate quantification of steady-state transcript levels by RT-PCR. Specificities of the PCR amplification were always documented with melting curve analysis with LightCycler Software v.3.5 (Roche Applied Science). Melting peaks obtained either from RT product or from specific recombinant DNA were identical. No product was detected when reverse transcription was not performed or when no template RNA was used. mRNA was prepared from isolated muscle layers and isolated mucosa of rat duodenum and ileum. The RNA was prepared from ~7 mg of tissue, with muscle layers and mucosa separated as described in MATERIALS AND METHODS. However, these preparations contain smooth muscle cells and other less numerous populations of cells (e.g., interstitial cells of Cajal, macrophages, fibroblasts) that would contribute to the quantitative measurement of CRF1–2 expression. RNA was reverse transcribed to cDNA, and steady-state transcripts were determined relative to an endogenous control housekeeping gene (CYCA) (13). Therefore, the data are expressed as CRF1/CYCA and CRF2/CYCA. The relative transcriptional expression of CRF1 and CRF2 is shown in Fig. 4. The results are expressed as means ± SE. In the rat duodenum and ileum, CRF1 expression relative to CYCA (relative unit) was 24.67 ± 4.23 and 15.37 ± 3.89 for external muscle layers, respectively (P < 0.05), and 12.45 ± 2.91 and 2,37 ± 0.64 for the mucosa, respectively (P < 0.001; n = 3 for all these data). In the rat duodenum and ileum, CRF2 expression relative to CYCA (relative unit) was 0.97 ± 0.31 and 24.20 ± 4.27 for external muscle layers, respectively (P < 0.001), and 9.64 ± 2.13 and 16.79 ± 3.58 for the mucosa, respectively (P < 0.001; n = 3 for all these data).



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Fig. 4. Relative expression of CRF1 and CRF2 receptors in rat small intestine: quantitative real-time RT-PCR performed on total RNA prepared from external muscle layers and mucosa of the duodenum and ileum. Values for CRF1 and CRF2 steady-state transcript relative to cyclophilin A (CYCA) in the same RNA preparation are shown. Results are expressed as means ± SE. Significant difference between CRF1-to-CRF2 ratios in duodenum and ileum muscle layer- and mucosa-derived RNA (n = 3): *P < 0.05; **P < 0.001.

 
Functional studies to demonstrate activity of CRF1 and CRF2. Isometric force measurements were performed on circular muscle strips of the rat small intestine to test the functional significance of CRF1 and CRF2 receptors. The first series of experiments tested the relative effects of CRF and Ucn 1, at 10–6 and 10–9 M, on the phasic contractions generated by the duodenum and ileum muscles. Circular muscle strips of rat duodenum and ileum were spontaneously active and generated periodic phasic contractions that averaged 319.26 ± 37.14 and 381.78 ± 37.31 mg (n = 16 animals) in amplitude and occurred at a frequency of 32.56 ± 4.64 and 31.77 ± 5.12 cycles/min, respectively. Perfusion of CRF (1 nM) and Ucn 1 (1 nM) in the bathing media produced 1) an increase in the amplitude of duodenal muscle contractions to 521.12 ± 48.05 and 631.56 ± 32.24 mg in force, respectively (P < 0.05 compared with control), and 2) a decrease in the amplitude of ileal muscle contractions to 216.41 ± 49.47 and 142.68 ± 53.15 mg in force, respectively (P < 0.05 compared with control). Perfusion of CRF (1 µM) and Ucn 1 (1 µM) in the bathing medium produced 1) an increase in the amplitude of duodenal muscle contractions to 535.34 ± 53.47 and 691.49 ± 30.46 mg in force, respectively (P < 0.05 compared with control; Fig. 5, A and B), and 2) a decrease in the amplitude of ileal muscle contractions to 209.13 ± 25.31 and 160.67 ± 36.72 mg in force, respectively (P < 0.05 compared with control; Fig. 5, C and D). In both regions studied, the frequency of contractions was not significantly affected by CRF or Ucn 1. After washout of CRF and Ucn 1, the amplitude of contractions returned to control levels. Data comparing the relative effects of CRF and Ucn 1, for both concentrations, on spontaneous contractions in the duodenum and ileum are summarized in Fig. 6. In the subsequent experiments, we used the dose of CRF and Ucn 1 showing maximal effects on intestinal motor activity (i.e., 1 µM).



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Fig. 5. Effects of CRF and urocortin (Ucn) 1 on circular muscle contractions of the rat duodenum and ileum. CRF (1 µM; A) and Ucn 1 (1 µM; B) increased the amplitude of duodenum spontaneous contractions. CRF (1 µM; C) and Ucn 1 (1 µM; D) decreased the amplitude of ileum spontaneous contractions.

 


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Fig. 6. A: summary of the relative changes in contractile force produced by circular muscles of the duodenum in response to CRF (10–9 and 10–6 M), plotted as % change in the maximal contraction. CRF caused 29% (1 nM) and 33% (1 µM) increase in force of spontaneous contractions. B: summary of the changes in mechanical force produced by circular muscles of the duodenum in response to the exogenous application of Ucn 1 (10–9 and 10–6 M), plotted as % change in the maximal contraction. Ucn 1 caused 31% (1 nM) and 38% (1 µM) increase in the force of spontaneous contractions of the circular muscle layer. C: summary of the relative changes in contractile force produced by circular muscles of the ileum in response to CRF (10–9 and 10–6 M), plotted as % change in the maximal contraction. CRF caused 22% (1 nM) and 28% (1 µM) decrease in force of spontaneous contractions. D: summary of the changes in mechanical force produced by circular muscles of the ileum in response to the exogenous application of Ucn 1 (10–9 and 10–6 M), plotted as % change in the maximal contraction. Ucn 1 caused 42% (1 nM) and 40% (1 µM) decrease in the force of spontaneous contractions of the circular muscle layer. *Significance level of P < 0.05.

 
The second series of experiments tested the relative effects of CRF (1 µM) and Ucn 1 (1 µM) on the phasic contractions generated by the duodenum and ileum muscles in the presence of specific CRF1 inhibitors (CP-154,526 and SSR125543Q). Perfusion of CP-154,526 (1 µM) or SSR125543Q (1 µM) in the bathing medium did not change the force of spontaneous contractions of duodenal (i.e., 344.37 ± 17.25 mg after CP-154,526 and 314.28 ± 17.21 mg after SSR125543Q) and ileal (i.e., 357.82 ± 20.73 mg after CP-154,526 and 376.01 ± 18.94 mg after SSR125543Q) muscles. In the continued presence of CP-154,526 or SSR125543Q, addition of CRF (1 µM) and Ucn 1 (1 µM) in the bathing medium 1) did not significantly change the force of spontaneous contractions of duodenal muscles (i.e., 359.75 ± 23.26 mg in the presence of CP-154,526 and CRF and 358.33 ± 22.33 mg in the presence of CP-154,526 and Ucn 1; 332.14 ± 20.42 mg in the presence of SSR125543Q and CRF and 341.89 ± 17.61 mg in the presence of SSR125543Q and Ucn 1) and 2) did produce a decrease in the amplitude of ileal muscle contractions to 297.09 ± 15.58 and 259.71 ± 7.66 mg in force, respectively, in the continued presence of CP-154,526 (P < 0.05; compared with control activity) and to 288.43 ± 18.63 mg in the presence of SSR125543Q and Ucn 1 (P < 0.05; compared with control activity). There was also a slight decrease in the ileal muscle contractions from a control value of 376.05 ± 18.92 mg in the presence of SSR125543Q to 328.36 ± 20.61 mg after CRF, but this change did not reach a level of significance (P > 0.05). Data from experiments with CP-154,526 and SSR125543Q are summarized graphically in Figs. 7 and 8.



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Fig. 7. Summary of the effects of CRF1 inhibitors (CP-154,526 and SSR125543Q) on contractile force produced by circular muscles of the duodenum in response to the application of CRF or Ucn 1. A and D: effects of CP-154,526 (1 µM) and SSR125543Q (1 µM) on circular muscle contractile activity. CRF1 inhibitors did not influence basal duodenal motility. B, E, C, and F: summary of the effects of CRF (1 µM) and Ucn 1 (1 µM) in the presence of CP-154,526 or SSR125543Q. CRF and Ucn 1 did not produce a significant increase in the force of contractions of circular muscles from the duodenum.

 


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Fig. 8. Summary of the effects of CRF1 inhibitors (CP-154,526 and SSR125543Q) in contractile force produced by circular muscles of the ileum in response to the application of CRF or Ucn 1. A and B: effects of CP-154,526 (1 µM) and SSR125543Q (1 µM) on circular muscle contractile activity. CRF1 inhibitors did not influence basal ileal motility. C–F: summary of the effects of CRF (1 µM) and Ucn 1 (1 µM) in the presence of CP-154,526 or SSR125543Q. CRF and Ucn 1 had a decreased effect in the force of contractions of circular muscles from the ileum. *Significance level of P < 0.05.

 
We also tested the relative effects of CRF and Ucn 1 on the phasic contractions generated by the duodenum and ileum muscles in the presence of a selective CRF2 inhibitor (astressin2-B). Perfusion of astressin2-B (1 µM) in the bathing medium did not change the force of spontaneous contractions of duodenal (i.e., 339.14 ± 40.38 mg) and ileal (i.e., 359.30 ± 14.88 mg) muscles. In the continued presence of astressin2-B, addition of CRF (1 µM) and Ucn 1 (1 µM) in the bathing medium 1) produced an increase in the amplitude of duodenal muscle contractions to 425.86 ± 33.71 and 471.62 ± 38.68 mg in force, respectively (P < 0.05; compared with control activity), and 2) did not significantly change the force of spontaneous contractions of ileal muscles (i.e., 333.87 ± 31.43 mg in the presence of astressin2-B and CRF and 336.17 ± 26.91 mg in the presence of astressin2-B and Ucn 1). Data from experiments with astressin2-B are summarized graphically in Fig. 9.



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Fig. 9. Summary of the effects of CRF2 inhibitor astressin2-B in contractile force produced by circular muscles of the duodenum and ileum in response to the application of CRF or Ucn 1. A–C: summary of activities from the duodenum. A: effects of astressin2-B on circular muscle contractile activity. Astressin2-B did not influence basal duodenal motility. B and C: summary of the effects of CRF (1 µM) and Ucn 1 (1 µM) in the presence of Astressin2-B. CRF and Ucn 1 had an increased effect in the force of contractions of circular muscles from the duodenum. D–F: summary of activities from the ileum. D: effects of astressin2-B on circular muscle contractile activity. Astressin2-B did not influence basal ileal motility. E and F: summary of the effects of CRF (1 µM) and Ucn 1 (1 µM) in the presence of astressin2-B. CRF and Ucn 1 did not produce a significant decrease in the force of contractions of circular muscles from the ileum. *Significance level of P < 0.05.

 
To investigate whether the responses of the duodenum and ileum circular muscles to CRF and Ucn 1 applications were mediated by a direct effect of these compounds on the smooth muscle cells or originated from an indirect response through actions on the enteric nervous system, 1 µM CRF or Ucn 1 applications were repeated in the presence of 1 µM TTX. In the presence of TTX, CRF or Ucn 1 did not affect the amplitude of spontaneous contractions.


    DISCUSSION
 TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
This study provides the first comprehensive description of the expression and effects of CRF1–2 receptors in the small intestine (i.e., duodenum and ileum). The immunofluorescence and molecular experiments performed provide evidence that CRF receptors are expressed in the tunica muscularis of the rat small intestine. CRF1–2 receptors are expressed in neuronal elements within the intestinal wall of the rat. Functional studies show that, in vitro, CRF and Ucn 1 induce a stimulation of the contractile activity of the duodenum mediated by the activation of CRF1 receptor, whereas they inhibit phasic contractions of the ileum by activating CRF2 receptor.

CRF1-LI was mainly located in enteric neurons that have Dogiel type II morphology, which have been shown to be sensory neurons in the guinea pig small intestine (11). Dogiel type II neurons have smooth cell bodies with relatively large diameters and one or more axonal processes. We observed CRF1-LI in the cell bodies and axons of neurons of Dogiel type II neurons located in both myenteric and submucosal ganglia. The cell body labeling with CRF1 antibodies was mainly cytoplasmic and, although the receptor occurred at the membrane, the resolution of the immunohistochemistry is not sufficient to determine that the receptor is an integral membrane protein. However, preliminary intracellular recordings obtained in the guinea pig small intestine indicate that Dogiel type II myenteric neurons with AH-type electrophysiological behavior do have functional CRF1 receptor in their soma membranes (25). In the myenteric plexus, AH type II neurons exhibited depolarizations that were induced by Ucn 1 and abolished by the CRF1 receptor antagonist NBI-27914, but not the CRF2 receptor antagonist antisauvagine-30.

CRF2-LI was predominantly expressed by nerve varicosities, although in some neurons CRF2-LI was also detected in the cytoplasm. The mucosa contained CRF2-LI nerve fibers; however, the innervation of the mucosa was sparse compared with the dense CRF2 innervation of ganglia and muscle. Our morphological data obtained by the use of polyclonal antibodies raised against an internal region of CRF1 and the amino terminus region of CRF2 show that CRF1 and CRF2 were present in nerve elements supplying the tunica muscularis of the rat small intestine. By contrast, a recent immunohistochemical study using polyclonal antisera directed against the carboxy terminus of the CRF receptor proteins failed to localize CRF1 or CRF2 in the muscle layers of the rat small intestine (7). Consequently, the discrepancy between our results and the above-mentioned immunohistochemical study could be explained by different degrees of immunoreactivity depending on the protein sequences used for the generation of these antibodies. In addition to neuronal cell bodies and nerve fibers in the enteric nervous system, CRF1–2-LI was found in cells scattered among the epithelial cells along the villi of the mucosa. The identity of these cells was not defined, as demonstrated by the lack of colocalization of CRF2 receptor with 5-HT-like immunoreactivity. Further studies are needed to precisely identify the cell types expressing CRF receptors in the mucosa.

Although we did not observe significant differences in the density of CRF1 and CRF2 immunoreactivities in the regions studied, quantitative analysis of CRF1 and CRF2 transcriptional expression indicates that both CRF receptor subtypes are differentially expressed in the rat small intestine with a distinct pattern and regional differences. CRF1 showed a greater expression in the duodenum than in the ileum, whereas CRF2 was expressed more in the ileum than in the duodenum.

Functional experiments showed that, in vivo, central or peripheral administration of CRF or Ucn 1 inhibited gastric emptying, delayed small intestinal transit, and stimulated colonic transit (21, 23, 31, 57, 58). In those experiments, injection of CRF or Ucn 1 induced an increase in the contractile activity of the duodenum but inhibited the percentage of propagated duodenal contractions (21, 23). These effects of CRF or Ucn 1 on the duodenal contractile activity are in agreement with the present results, in which CRF or Ucn 1 elicited an increase of the duodenal smooth muscle contractions. Both present and previous studies show that the motility changes associated with CRF administration are region specific. A direct action of CRF in the intestinal wall is supported by the demonstration that CRF increases peristaltic activity in isolated rat distal colon and that functional CRF binding sites are present in cecal smooth muscle cells of guinea pig (19, 30, 31, 57). In addition, it was demonstrated that intraperitoneal injection of CRF activated myenteric neurons, increased colonic motility, and induced diarrhea through CRF-CRF1 pathways (17, 32, 52). A direct action within the enteric nervous system is supported by the induction of Fos expression in myenteric neurons after intraperitoneal injection of CRF (33). Furthermore, intracellular recording on myenteric neurons from the guinea pig ileum showed that CRF evoked a prolonged excitatory response (15). These results are consistent with the present study, in which TTX treatment resulted in a loss of effects induced by CRF or Ucn 1 on the contractile activity in the rat small intestine.

Finally, the finding that CRF1–2-LI was in neurons and nerve fibers supported these data, suggesting that CRF-related peptides can directly influence intestinal function by acting on myenteric neurons. Furthermore, the morphological evidence showing the presence of Ucn 1, but not CRF, mRNA in the enteric nervous system of the rat small intestine (16) suggests that Ucn 1 could be the main endogenous ligand for CRF1–2 receptors in the regulation of intestinal motor function. This finding concurs well with our results showing that Ucn 1 has a more potent effect than CRF on intestinal motility.

Previous reports have shown that CRF and Ucn 1 delay gastric emptying by activating CRF2 receptor and stimulate colonic motility by activation of CRF1 receptor. The selective CRF2 receptor antagonists astressin2-B and antisauvagine-30 prevented CRF- and Ucn 1- induced inhibition of gastric emptying; in contrast, the stimulation of the distal colonic transit was not modified (31, 34). Conversely, the selective CRF1 receptor antagonists CP-154,526 and NBI-27914 blocked CRF or Ucn 1 stimulatory effect on colonic motor function and Fos expression while not altering the delayed gastric emptying (29, 31, 33, 34, 48). In the present study, the selective CRF1 antagonists CP-154,526 and SSR125543Q blocked CRF- or Ucn 1-induced activation of duodenal contractile activity but did not block CRF- or Ucn 1-induced inhibition of ileal contractile activity. In contrast, a specific CRF2 inhibitor (astressin2-B) blocked the effects of CRF and Ucn 1 on the ileal muscle contractions but did not influence CRF- or Ucn 1-induced activation of duodenal contractile activity. The use of these antagonists confirmed that the observed effects of CRF and Ucn 1 on intestinal motility are receptor mediated. In these experiments, astressin2-B and both CRF1-selective antagonists CP-154,526 and SSR125543Q did not influence basal intestinal motility; these results indicate that peripheral CRF or Ucn 1 does not contribute to the basal regulation of intestinal motor function.

Our findings indicate that CRF and Ucn 1 at 10–6 M produce a similar level of activation on duodenal contractile activity (33% and 38%, respectively), whereas Ucn 1 (40% relaxation) has a significantly more potent effect than CRF (28% relaxation) on the relaxation of ileal muscles. Mancinelli et al. (30) showed that CRF in vitro increases colonic motor activity in a dose-dependent manner. Indeed, the excitatory effect of CRF in vitro appeared at 10–10 M, peaked at 10–8 M, decreased at 10–7 M, and disappeared at 10–6 M. In our study, data obtained at a concentration of 10–9 M produced comparable actions of CRF and Ucn 1 on intestinal motility; however, both compounds were slightly more potent at 10–6 M. Thus the dose-dependent effect of CRF-like peptides on the mechanical activity of the rat small intestine was not observed in the present study for both concentrations tested.

One important characteristic of CRF receptor subtypes is their distinct affinity for mammalian CRF family ligands. CRF has higher affinity (10- to 40-fold) for CRF1 than CRF2 receptor, whereas Ucn 1 binds with equal, and higher than CRF, affinity to both receptors (24, 44, 57). These data provide convergent evidence that CRF1 is predominantly expressed in the duodenum, whereas CRF2 is the major receptor for CRF and Ucn 1 in the ileum. Together, these data support the idea that CRF and Ucn 1 inhibitory effects on the gastrointestinal tract are mediated through CRF2, whereas excitatory effects of these compounds are mediated through CRF1.

In summary, we have characterized the distribution of CRF1 and CRF2 receptors in the rat small intestine and shown that CRF and Ucn 1 activate contractile activity in the duodenum through CRF1 receptor while inhibiting ileal contractions through CRF2 receptor. Pharmacological experiments showed that the selective antagonist SSR125543Q is a relevant tool to assess the role of peripheral CRF1 as observed for CP-154,526. Quantitative analysis of mRNA expression indicates that CRF1 is preferentially expressed in duodenum, whereas CRF2 is more represented in the ileum. On the basis of our findings, it seems likely that CRF and Ucn 1 play a significant role in modulating gastrointestinal function through CRF1 and CRF2 receptors. These data have pharmacological applications in that selective CRF receptor subtype antagonists may be used to regulate motor activity of the gastrointestinal tract in patients with motility disorders.


    ACKNOWLEDGMENTS
 
We thank Dr. J. Rivier (Clayton Foundation Laboratories for Peptide Biology, Salk Institute, La Jolla, CA) for the generous gift of the CRF2 antagonist astressin2-B, Dr. Daniel Gully (Sanofi-Synthélabo Recherche, Toulouse, France) for the donation of the CRF1 antagonist SSR125543Q, and Dr. Donnie W. Owens (Global Research & Development, Pfizer, Groton, CT) for the supply of CRF1 antagonist CP-154,526. Confocal microscopy was performed at the Institut Albert Bonniot of the University Joseph Fourier, Grenoble, France. We give special thanks to Alexeï Grichine and Jacques Mazzega for help with confocal microscopy.


    FOOTNOTES
 

Address for reprint requests and other correspondence: C. Porcher, GESIND EA 3744, Pavillon Neurologie B.P. 217, Centre Hospitalier Universitaire Michallon, 38043 Grenoble Cedex 09, France (E-mail: christophe.porcher{at}ujf-grenoble.fr)

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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