AJP - GI Watch the video to see how APS reaches out to developing nations.
HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS
QUICK SEARCH:   [advanced]


     

Am J Physiol Gastrointest Liver Physiol 293: G461-G468, 2007. First published June 21, 2007; doi:10.1152/ajpgi.00424.2006
0193-1857/07 $8.00
This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow All Versions of this Article:
293/2/G461    most recent
00424.2006v1
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
Right arrow Citation Map
Services
Right arrow Email this article to a friend
Right arrow Similar articles in this journal
Right arrow Similar articles in ISI Web of Science
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Citing Articles
Right arrow Citing Articles via HighWire
Right arrow Citing Articles via ISI Web of Science (1)
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by McDonagh, S. C.
Right arrow Articles by Brubaker, P. L.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by McDonagh, S. C.
Right arrow Articles by Brubaker, P. L.

HORMONES AND SIGNALING

Role of glial cell-line derived neurotropic factor family receptor {alpha}2 in the actions of the glucagon-like peptides on the murine intestine

Sean C. McDonagh,1 Jenny Lee,1 Angelo Izzo,1 and Patricia L. Brubaker1,2

Departments of 1Physiology and 2Medicine, University of Toronto, Toronto, Ontario, Canada

Submitted 13 September 2006 ; accepted in final form 12 June 2007


   ABSTRACT
TOP
 ABSTRACT
METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
The intestinal glucagon-like peptides GLP-1 and GLP-2 inhibit intestinal motility, whereas GLP-2 also stimulates growth of the intestinal mucosa. However, the mechanisms of action of these peptides in the intestine remain poorly characterized. To determine the role of the enteric nervous system in the actions of GLP-1 and GLP-2 on the intestine, the glial cell line-derived neurotropic factor family receptor {alpha}2 (GFR{alpha}2) knockout (KO) mouse was employed. The mice exhibited decreased cholinergic staining, as well as reduced mRNA transcripts for substance P-ergic excitatory motoneurons in the enteric nervous system (ENS) (P < 0.05). Examination of parameters of intestinal growth (including small and large intestinal weight and small intestinal villus height, crypt depth, and crypt cell proliferation) demonstrated no differences between wild-type and KO mice in either basal or GLP-2-stimulated mucosal growth. Nonetheless, KO mice exhibited reduced numbers of synaptophysin-positive enteroendocrine cells (P < 0.05), as well as a markedly impaired basal gastrointestinal (GI) transit rate (P < 0.05). Furthermore, acute administration of GLP-1 and GLP-2 significantly inhibited transit rates in wild-type mice (P < 0.05–0.01) but had no effect in GFR{alpha}2 KO mice. Despite these changes, expression of mRNA transcripts for the GLP receptors was not reduced in the ENS of KO animals, suggesting that GLP-1 and -2 modulate intestinal transit through enhancement of inhibitory input to cholinergic/substance P-ergic excitatory motoneurons. Together, these findings demonstrate a role for GFR{alpha}2-expressing enteric neurons in the downstream signaling of the glucagon-like peptides to inhibit GI motility, but not in intestinal growth.

enteric nervous system; GFR{alpha}2; GLP-1; GLP-2; growth; motility

GLP-1 AND GLP-2 ARE SYNTHESIZED by posttranslational processing of proglucagon in the L cell of the distal gut and are coreleased into the circulation in response to enteral nutrition (11, 33). These peptide hormones act in concert to regulate nutrient assimilation; GLP-1 is highly insulinotropic and a potent inhibitor of gastrointestinal (GI) motility, effects that function in concert to limit postprandial glycemic excursions (5, 13, 24, 38), whereas GLP-2 promotes nutrient absorption by increasing the growth and function of the small intestinal epithelium (6, 13–15). However, although a GLP-1 receptor (GLP-1R) agonist is in the clinic for the treatment of Type 2 diabetes mellitus (4), and GLP-2 is in late-stage clinical trials for the treatment of short bowel syndrome (23), the mechanisms by which the glucagon-like peptides exert their effects on the gut remain poorly understood.

One major question regarding the actions of both GLP-1 and GLP-2 on the GI tract is the relative importance of the local enteric nervous system (ENS). GLP-1R expression has been demonstrated in the stomach and small and large intestine, as well as in the nodose ganglion of the vagus nerve (7, 30). Although the exact cellular localization of the GLP-1R within the GI tract has not been described, several studies have indicated that the effects of GLP-1 on gastric vs. intestinal motility are mediated through distinct mechanisms. Hence, vagal deafferentation prevents the inhibitory effect of GLP-1 on antral motility in rats (21), implicating the vagus in the gastric actions of GLP-1, although this does not appear to be the case in pigs (28). However, in the small intestine, adrenergic blockers antagonize the inhibitory actions of GLP-1 (19), indicative of local effects to inhibit excitatory cholinergic motoneurons. Nonetheless, one study has demonstrated that GLP-1 does not affect contractility in isolated gut segments (45), suggesting that, despite expression of the GLP-1R, this peptide does not directly modulate intestinal motility.

Unlike the GLP-1R, the glucagon-like peptide 2 receptor (GLP-2R) has been localized within the GI tract to a subset of enteroendocrine cells, as well as to subepithelial myofibroblast cells in the mucosa and neurons in the ENS (3, 20, 32, 48). Initial studies suggested a role for the ENS GLP-2R in intestinal growth through the demonstration that tetrodotoxin prevents GLP-2-mediated activation of c-fos in small intestinal crypt cells (3). However, the GLP-2R has recently been localized to ENS neurons expressing vasoactive intestinal peptide (VIP) and endothelial nitric oxide synthase, where it is suggested to mediate GLP-2-induced gut blood flow (20). Nonetheless, GLP-2 has recently been reported to inhibit gastric emptying in humans and pigs (29, 47), although the findings are controversial (39), possibly because of reduced potency compared with GLP-1 (29). Furthermore, although direct inhibitory effects of GLP-2 on large intestinal motility have been demonstrated (10), it has been suggested that its effects on the small intestine are mediated through GLP-1 (5). In addition, the exact localization within the GI tract of the GLP-2R mediating these effects has not been determined.

To determine the role of the ENS in the actions of GLP-1 and GLP-2 on the GI tract, we have employed a mouse model of enteric nervous deficit, the glial cell line-derived neurotropic factor family receptor {alpha}2 (GFR{alpha}2) knockout (KO) mouse. GFR{alpha}2 is a member of a family of co-receptors for RET signaling that are required for both colonization of the developing gut by neuronal precursor cells and subsequent tropic support of these neurons (1, 22). In contrast to the lethality of RET-null or other glial cell line-derived neurotropic factor-null mice (17), GFR{alpha}2 KO is associated with a partial deficit of parasympathetic innervation and a 35% reduction in cholinergic fiber and substance P (SP) soma density in the myenteric plexus, resulting in reduced but not abrogated GI transit rates in vivo (36, 37). We therefore hypothesized that these deficits in enteric innervation would result in an impaired tropic response to GLP-2 and attenuated inhibition of gut motility by GLP-1 and/or GLP-2.


   METHODS
TOP
 ABSTRACT
 METHODS
RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Animals. GFR{alpha}2 KO mice (C57Bl/6 x 129Sv/J) were a generous gift from Dr. J. Milbrandt (Washington University School of Medicine, St. Louis, MO). Briefly, these mice were generated by homologous recombination of the first coding exon with an enhanced green fluorescent protein cassette, thereby interrupting the GFR{alpha}2 coding sequence. GFR{alpha}2 genotype was determined by PCR with the following primers at an annealing temperature of 60°C: common forward, 5'-GGCGGGATCTCCGCATTGGATTT; wild-type (WT) reverse, 5'-AGAAGGCGTTTGCCAAGATCATGTTAA; KO reverse, 5'-CCCTGAGCATGATCTTCCATCACGTCG. Mice were housed in standard, pathogen-free conditions under a constant light-dark cycle and were fed Teklad rodent chow wet mash to prevent bowel obstruction (unpublished data) and promote normal growth, as reported for an independent line of GFR{alpha}2 KO mice (36). All experiments were approved by the Animal Care Committee of the University of Toronto.

Chronic GLP-2 administration study. Age- and sex- matched GFR{alpha}2 WT, heterozygous (HT), and KO littermates were administered 0.15 µg/g body wt human [Gly2]GLP-2 (Peptidec Technologies, Pierrefonds, QC, Canada) or an equivalent volume of PBS subcutaneously every 24 h for 14 days, and 3 h before euthanasia on day 15, as reported previously (6, 14, 15, 44). [Gly2]GLP-2 is a degradation-resistant analog of GLP-2 that has a longer half-life in vivo than the native peptide (44). Some animals were also injected with 100 mg/kg 5-bromo-2-deoxyuridine (BrdU) ip in PBS 1 h before death. Following anesthesia with isoflurane and exsanguination, the small and large intestines were cleaned and weighed, and whole jejunal segments, as well as microdissected jejunal tunica muscularis, were collected for further analysis. The presence of both the myenteric and the submucus plexus in the microdissected tissue was confirmed by demonstration of two morphometrically distinct layers of neurofilament-positive neurons and cell bodies (data not shown).

Morphometry, histochemistry, and immunohistochemistry. As the greatest effects of GLP-2 tropic actions are observed in the jejunum (46), all growth analyses were conducted in this tissue. Two-centimeter segments of midjejunum (12 cm distal to the pyloric sphincter) were cut into four equal segments, fixed in 10% neutral buffered formalin overnight and transferred to ethanol prior to paraffin embedding (to make n = 1). Embedded tissue was cut into crosssections which were mounted and stained with hematoxylin and eosin. Morphometric analysis was performed using a Zeiss microscope with Axio Vision software (Carl Zeiss Canada, Don Mills, ON, Canada). Villus heights and crypt depths were obtained by blinded measurements of the 4 tissue cross-sections (to make n = 1), with at least 50 and 30 determinations, respectively, per mouse (6, 14, 15). Cell width and height were determined for five cells per villus across ten villi per mouse (i.e., 50 cells per mouse to make n = 1).

Acetylcholinesterase (AChE) histochemistry was performed as described (42). Briefly, microdissected jejunal tunica muscularis was incubated in acetylthiocholine iodide for 30 min to generate a product with peroxidase activity within the tissue. Subsequently the tissue was incubated with 3,3'-diaminobenzidine and nickel ammonium sulfate for 5 min prior to addition of 0.003% H2O2 for 10 min to yield a dark precipitate on cholinergic neurons.

Ki-67 immunohistochemistry for proliferating cells was performed using a rat anti-mouse Ki-67 antibody (TEC-3, Dako, Glostrup, Denmark), as reported previously (15, 40). For proliferation, an average of 20 half-crypts per mouse were counted in a blinded manner (to make n = 1), with the cell at the base of the crypt designated cell 1. Half-crypts were defined as one side of the crypt compartment starting from the base of the crypt and were used to prevent redundant analysis of each crypt unit. Proliferation was determined as the proportion of Ki-67-positive cells relative to the total number of cells at each cell position. Uptake of BrdU was determined by antigen retrieval of tissue sections using 0.4% pepsin in 0.01 N HCl, followed by incubation with a mouse anti-BrdU antibody (1:1,000; Sigma Chemical, St. Louis, MO) and then with a biotinylated goat anti-mouse antibody (Vector Laboratories) and detection using 3,3'-diaminobenzidine. Positive cells were counted as described for Ki-67. Crypt and villus cell apoptosis was determined by using an antiserum directed against cleaved (active) caspase-3 (Cell Signaling, Danvers, MA) followed by a biotinylated goat anti-rabbit secondary (Vector Laboratories, Burlingame, CA) and detection using 3,3'-diaminobenzidine. In contrast to the proliferative cells, the number of apoptotic cells was low, and the total cells per crypt and per villus were therefore determined across four tissue sections from each mouse, and the results were expressed relative to the total number of crypts or villi counted.

Mucin histochemistry was performed with the Rapid Mucin Stain kit (Polysciences, Warrington, PA) according to the manufacturer's instructions. At least 20 half-villi were counted per mouse (to make n = 1). As for the half-crypts, half-villi were defined as one side of the villus and were used to prevent redundant analysis of the villus compartment. Synaptophysin immunofluorescence was conducted using polyclonal rabbit anti-human synaptophysin antibody (1:200 dilution, Dako) as the primary antibody, 5% normal donkey serum as the blocking solution, and Cy3-conjugated Affinipure donkey anti-rabbit IgG (Jackson Immunoresearch Laboratories, West Grove, PA) as the secondary antibody. At least 20 half-villi were counted per mouse (to make n = 1). Serotonin immunohistochemistry was performed using polyclonal rabbit serotonin antiserum as the primary antibody (Covance Research Products, Berkeley, CA) with biotinylated goat anti-rabbit secondary (Vector Laboratories), followed by detection with 3,3'-diaminobenzidine. At least 20 half-villi were counted per mouse (to make n = 1).

Quantitative and RT-PCR. One-centimeter sections of proximal jejunum (11 cm from the pyloric sphincter) were snap-frozen in liquid nitrogen and stored at –80°C until homogenization in RLT RNA preservation buffer (Qiagen, Mississauga, ON, Canada) and RNA extraction. Strips of tunica muscularis were sonicated in RLT buffer and stored at –80°C until use. RNA was isolated with the RNeasy kit (Qiagen) according to the manufacturer's instructions. Some tissue samples were analyzed by RT-PCR using a One-Step kit (from Qiagen). The RT-PCR primers and conditions have been reported previously (27, 40). Other samples were reverse-transcribed to cDNA with Superscript II Reverse Transcriptase (Invitrogen, Burlington, ON, Canada). Quantitative RT-PCR (qRT-PCR) was performed in a Chromo4 Continuous Fluorescence Detection unit with Opticon Monitor 3 software (Bio-Rad Laboratories, Mississauga, ON, Canada) using Taqman Gene Expression Assays (Applied Biosystems, Foster City, CA), as listed in Table 1 (because the data for the GLP-2R exons 3-4 was not different than that for exons 11-12, only the latter is shown). All reactions were performed in triplicate, and control reactions were performed without RT enzyme and/or without template. The linearity of amplification of the Taqman primer-probe sets was verified over four orders of magnitude. Expression was calculated relative to the internal control, 18S rRNA, which was selected because we have previously shown that it is not affected by chronic GLP-2 administration (15). Relative transcript expression was calculated according to the {delta}{delta}CT method (34), whereby the difference in cycle threshold ({delta}CT) of the transcript of interest and the internal control were divided for each animal to make n = 1.


View this table:
[in this window]
[in a new window]

  		Table 1. Taqman gene expression assays

 
Functional analyses. To examine total GI transit, overnight-fasted, age- and sex-matched GFR{alpha}2 WT and KO littermates were injected intraperitoneally with 0.15 µg/g body wt [Gly2]GLP-2, 0.06 µg/g body wt GLP-1, or an equivalent volume of PBS; these peptide doses were selected based on the results of preliminary studies. Five minutes after injection, mice were gavaged (10 ml/kg) with 5% activated charcoal in 10% gum arabic (12). Twenty minutes later the animals were euthanized and the transit of the charcoal solution along the length of the GI tract was assessed visually with a ruler under constant tension. Gastric emptying was determined by spectrophotometric analysis (at 405 nm) of phenol red remaining in the stomach of fasted mice 20 min after oral gavage of 0.1 mg/ml phenol red, as per the protocol of Crowell et al. (8). Exendin-4 (0.1 µg ip; Bachem California, Torrance, CA)-induced suppression of 8-h food intake was determined in fasted mice, as previously reported (43).

Statistical analysis. All statistical analyses were conducted with Prism 4.00 software (GraphPad Software, San Diego, CA). Student's t-test or one- or two-way ANOVA was performed, as appropriate. ANOVA was followed, when significant, by post hoc comparisons using Student's t-test or one-way ANOVA.


   RESULTS
TOP
 ABSTRACT
 METHODS
 RESULTS
DISCUSSION
 GRANTS
 REFERENCES
 
Characterization of the ENS in GFR{alpha}2 KO mice. GFR{alpha}2 WT, HT, and KO animals were identified by PCR of tail DNA (Fig. 1A), and qRT-PCR of microdissected tunica muscularis confirmed the absence of GFR{alpha}2 mRNA in KO animals (Fig. 1B). Histochemical staining of the tunica muscularis for AChE also demonstrated decreased cholinergic fiber staining in KO mice relative to WT controls (Fig. 1C).


Figure 1
View larger version (45K):
[in this window]
[in a new window]

  		Fig. 1. Characterization of glial cell line-derived neurotropic factor family receptor {alpha}2 (GFR{alpha}2) knockout (KO) mice. A: mice were genotyped by PCR (MW, molecular weight ladder; HT, heterozygous). B: results of the genotyping were confirmed by quantitative RT-PCR (qRT-PCR) (n = 10, ***P < 0.001). C: histochemistry for acetylcholinesterase in microdissected jejunal tunica muscularis from wild type (WT) and KO animals.

 
The presence of mRNA transcripts for both the GLP-1 and the GLP-2 receptor in microdissected tunica muscularis was demonstrated by RT-PCR (Fig. 2A). The same tissues did not express mRNA for an epithelial endocrine cell marker, proglucagon (control), indicating the specificity of the separation technique. Knockout of GFR{alpha}2 was not associated with changes in mRNA transcript levels for the parasympathetic cholinergic marker vesicular acetylcholine transporter (VAChT), the more general cholinergic marker AChE, or the general neural marker beta-tubulin 3 (BT3) in microdissected tunica muscularis (Fig. 2B). However, expression of mRNA transcripts for the excitatory neurotransmitter SP was significantly reduced in KO mice (by 37 ± 11%, P < 0.05). No difference was observed in mRNA transcript levels for VIP (Fig. 2C), nor was there differential expression of GLP-1R or GLP-2R mRNA transcripts in tunica muscularis from the KO animals (Fig. 2D).


Figure 2
View larger version (14K):
[in this window]
[in a new window]

  		Fig. 2. mRNA transcript expression in tunica muscularis. A: microdissected jejunal tunica muscularis (TM) and whole jejunum (WJ) were examined by RT-PCR for GLP-1 receptor (GLP-1R), GLP-2 receptor (GLP-2R), and the epithelial-specific marker, proglucagon (proG) mRNA transcripts (MW, molecular weight ladder; neg, negative control). B–D: qRT-PCR of mRNA transcripts for cholinergic (VAChT and AChE) and general [beta-tubulin 3 (BT3)] neural markers (n = 5 for VAChT, n = 11 for AChE and BT3) (B), excitatory [substance P (SP)] and inhibitory (VIP) neurotransmitters (n = 10 each) (C), and glucagon-like peptide receptors (n = 11 each) (D), in KO mice (open bars) relative to WT animals (hatched bars). *P < 0.05.

 
Effects of chronic GLP-2 administration. The body weight of age- and sex-matched KO mice was significantly lower than that of WT animals (18.2 ± 0.6 vs. 16.0 ± 0.8, P < 0.05), likely due to general parasympathetic denervation as reported for an independent line of GFR{alpha}2 KO mice (37). Subcutaneous administration of [Gly2]GLP-2 daily for 14 days significantly increased small intestinal wet weight relative to body weight in WT, HT, and GFR{alpha}2 KO mice (Fig. 3A, P < 0.001). A similar trend was seen in the large intestine (Fig. 3B). Small intestinal villus height (P < 0.001, Fig. 3C) was also increased by chronic administration of [Gly2]GLP-2, although only a trend toward increased crypt depth was observed (Fig. 3D). However, no effects of GFR{alpha}2 KO were observed in any of these parameters of epithelial growth, and no differential effect of GLP-2 administration was seen between groups. Surprisingly, despite the increased small intestinal weight and crypt-villus height, no effect of GLP-2 was seen on small intestinal crypt cell proliferation in WT or KO animals, as determined by Ki-67 staining (Fig. 3, E and F) and, again, no differences between WT and KO mice were noted. Furthermore, analysis of BrdU uptake by the crypt cells revealed a similar profile as found for Ki-67 (with peak values of 0.55 ± 0.04 cells at positions 9–11) and, again, did not reveal any differences between saline- and GLP-2-treated WT or KO mice (data not shown). Determination of changes in crypt and villus cell apoptosis similarly did not reveal any differences due to GLP-2 treatment, although it is noted that the number of cells detectable in the normal mouse intestine was extremely low (only 1.1 ± 0.3% of all crypts and 9.8 ± 3.0% of all villi were found to contain any caspase-3-positive cells), making quantitative demonstration of any differences difficult. Finally, analysis of cell size did not demonstrate any significant differences in either cell width or length (5.1 ± 0.3 and 20.0 ± 0.9 µm, respectively, in WT-PBS mice) in response to GLP-2 treatment or in KO animals.


Figure 3
View larger version (25K):
[in this window]
[in a new window]

  		Fig. 3. Effect of GFR{alpha}2 KO and chronic GLP-2 administration on parameters of intestinal growth. A–D: WT, HT, and KO mice were administered 0.15 µg/g body wt GLP-2 daily for 14 days, followed by tissue collection on day 15. Small (A) and large (B) intestinal wet weight per body weight, and jejunal villus height (C) and crypt depth (D) (n= 10 for all groups). E–F: WT and KO mice were administered 0.15 µg/g body wt GLP-2 daily for 14 days, followed by tissue collection on day 15. Small intestinal crypt cell Ki-67 positivity was determined along the length of the crypt, with the cell at the base defined as position 1. *P < 0.05 and ***P < 0.001 for GLP-2 vs. saline treatment within the same genotype.

 
To determine the effects of GLP-2 and GFR{alpha}2 KO on lineage differentiation of the intestinal epithelium, histochemistry for the mucous cell product, mucin, and immunohistochemistry for both the enteroendocrine cell marker, synaptophysin, and enterochromaffin cells was performed. A significant increase in the number of mucin cells per villus was seen with GLP-2 treatment in WT mice (P < 0.05), and a similar trend was observed in synaptophysin-positive staining, concomitant with the increases observed in villus length (Fig. 4, A and B). Unexpectedly, significantly fewer synaptophysin-positive cells were observed in the villi of PBS-treated GFR{alpha}2 KO mice relative to PBS-WT animals (P < 0.05), and GLP-2 treatment partially reversed this deficit (P < 0.05). Serotonin-positive cell numbers in the jejunum equaled ~70% of the total synaptophysin-positive enteroendocrine cells (Fig. 4C). Furthermore, in keeping with the changes in synaptophysin positivity, the number of serotonin cells was increased by GLP-2 treatment of WT mice (P < 0.05), and there was a similar trend to decreased cell numbers in the KO animals.


Figure 4
View larger version (10K):
[in this window]
[in a new window]

  		Fig. 4. Effect of GFR{alpha}2 KO and acute GLP administration on villus cell numbers. Mucin positivity (A), synaptophysin positivity (B), and serotonin positivity (C) (n = 4–5 for all groups). *P < 0.05 for GLP-2 vs. saline treatment within the same genotype; +P < 0.05 for KO vs. WT with saline treatment.

 
Effects of acute administration of the glucagon-like peptides on GI transit and food intake. In WT mice, intraperitoneal administration of 0.15 µg/g [Gly2]GLP-2 caused a significant reduction in GI transit of a charcoal solution relative to treatment with vehicle alone (by 29 ± 8%, P < 0.05). GLP-1 (0.06 µg/g) administration also significantly impaired transit of the gavaged charcoal solution in WT animals (by 55 ± 4%, P < 0.001). GFR{alpha}2 KO mice exhibited decreased basal transit with vehicle administration relative to WT controls (by 37 ± 6%, P < 0.01); however, in contrast to their effects in WT animals, neither GLP-2 nor GLP-1 further reduced transit rates in KO mice (Fig. 5A). To determine whether the impairment in GI transit in KO animals was due to alterations in gastric emptying and/or intestinal motility, the retention of phenol red in the stomach was determined following oral gavage. No significant differences were observed between WT and KO mice, suggesting that the changes in GI transit observed in KO animals occurred consequent to altered intestinal motility (Fig. 5B). To further determine whether general vagal tone was altered in the KO animals, mice were administered a long-acting GLP-1 agonist, exendin-4, that we have previously demonstrated to inhibit food intake in mice through a vagal-dependent mechanism (43). No differences in exendin-4-induced reductions in food intake were observed between WT and KO animals (Fig. 5C).


Figure 5
View larger version (11K):
[in this window]
[in a new window]

  		Fig. 5. Effect of GFR{alpha}2 KO and acute GLP administration on gastrointestinal (GI) function and food intake. A: fasted WT and KO littermates were administered 0.06 µg/g body wt GLP-1, 0.15 µg/g body wt GLP-2, or an equivalent volume of saline ip at t = –5 min, followed by oral gavage with a charcoal solution at t = 0 min, and determination of GI transit at t = 20 min (n = 4–10). B: fasted WT and KO mice were given phenol red by orogastric gavage and the content remaining in the stomach after 20 min was determined spectrophotometrically (n = 4–6). C: fasted WT and KO mice were treated with or without 0.1 µg of exendin-4, and cumulative food intake was determined over an 8 h period (n = 3). *P < 0.05 for GLP vs. saline treatment within the same genotype; +P < 0.05 for KO vs. WT with saline treatment.

 

   DISCUSSION
TOP
 ABSTRACT
 METHODS
 RESULTS
 DISCUSSION
GRANTS
 REFERENCES
 
Despite more than a decade of research, the sites and mechanisms of action of the glucagon-like peptides within the intestine remain controversial and poorly understood. GLP-1 exerts a profound inhibitory effect on GI motility, yet the role of the GLP-1R in the intestine has never been explicitly studied. Meanwhile, several locations have been demonstrated for the GLP-2R within the intestinal mucosa and ENS, but the contributions of each to the biological effects of GLP-2 have not been confirmed. To better understand the actions of the glucagon-like peptide receptors in the ENS, we employed a model of partial enteric nervous deficit, the GFR{alpha}2 KO mouse. In this context, we hypothesized that GFR{alpha}2 KO mice would exhibit reduced mucosal growth in response to GLP-2 and impaired inhibition of gut motility by GLP-1 and/or GLP-2. The results of this study demonstrate that, although loss of GFR{alpha}2 abrogates the ability of both glucagon-like peptides to inhibit GI transit, there is no differential intestinotropic response to GLP-2 in GFR{alpha}2 KO mice.

Histochemical analysis of acetylcholinesterase distribution in microdissected tunica muscularis revealed a dramatic reduction in staining for small fibers in GFR{alpha}2 KO mice, as previously reported for a similar mouse model (37). Characterization of mRNA expression in the tunica muscularis by qRT-PCR further demonstrated that SP transcript levels were decreased by 35%, consistent with the results of immunohistochemical analyses showing reduced substance P-positive fiber density in other GFR{alpha}2-deficient animals (37). In contrast, mRNA transcript levels for the more general markers of cholinergic innervation, including VAChT, AChE, and BT3, were not altered by GFR{alpha}2 KO. These findings accord with our current understanding of the ENS, in that substance P-ergic neurons represent only a small proportion (~12–25%) of the total cholinergic neurons in the plexus (16). Thus, GFR{alpha}2 KO would not be expected to result in a profound reduction in either general cholinergic or neural markers of the entire ENS. Furthermore, the reduced cholinergic fiber staining did not appear to be associated with a demonstrable loss of cholinergic soma, where most gene expression occurs, which accounts for the apparent discrepancy between the histochemical and gene expression analyses. Finally, as VIP and SP or ACh have not been colocalized in the ENS (16), no change in VIP expression was expected in the GFR{alpha}2 KO mouse, nor was any such change observed. Taken together, therefore, the results of the present study confirm that GFR{alpha}2 KO results in a highly selective deficit of the cholinergic/substance P fibers that constitute one component of the excitatory enteric neural input to the longitudinal muscle of the small intestine (16, 25). These findings are consistent with reduced basal intestinal transit rates in these animals, as found in the present study and previously reported for other GFR{alpha}2 KO mice (36).

Administration of GLP-1 prior to oral gavage of a charcoal solution significantly reduced GI transit in WT mice, in keeping with the well-characterized ability of GLP-1 to inhibit both gastric emptying and intestinal motility (5, 19, 21, 45). However, GLP-1 did not reduce GI transit below baseline in the GFR{alpha}2 KO mice. Studies using vagal deafferentation and vagotomy have demonstrated that the vagus is an important site at which GLP-1 inhibits gastric emptying (21, 47). However, GFR{alpha}2 KO mice did not demonstrate any reductions in gastric emptying, nor did they exhibit an impaired response to vagal-dependent exendin-4-induced reductions in food intake. It is therefore likely that the loss of the inhibitory effect of GLP-1 in the KO mice is due to their deficiency in excitatory myenteric innervation in the small intestine. Finally, the localization of GLP-1R expression to the jejunal tunica muscularis suggests a local role for GLP-1 in the inhibition of gut motility, whereas the absence of an inhibitory response in KO mice demonstrates that GLP-1 does not act directly on the smooth muscle.

Given the lack of effect of GLP-1 on GI transit in GFR{alpha}2 KO mice, it was somewhat unexpected that no decrease in GLP-1R mRNA transcript levels in the ENS could be detected in these animals. However, prior studies have demonstrated that adrenergic blockers abolish the effect of GLP-1 on intestinal motility (19), suggesting that GLP-1 may act via presynaptic inhibition of cholinergic transmission. One model to explain the absence of a GLP-1 effect in the GFR{alpha}2 KO mice is that the GLP-1R is located on postganglionic sympathetic efferents in the gut that inhibit descending cholinergic motoneurons. GLP-1 signaling in this model would thus prevent cholinergic stimulation of excitatory GFR{alpha}2-expressing cholinergic/substance P neurons, with loss of these fibers resulting in abrogation of GLP-1-mediated transit inhibition. Future studies using animals that lack selective components of this signaling pathway will be required to confirm this mechanism of GLP-1-mediated inhibition of intestinal motility.

Similar to the findings made with GLP-1, administration of GLP-2 was found to inhibit GI transit in WT mice. Interestingly, the dose of [Gly2]GLP-2 that inhibited transit by 29% in these animals was markedly higher than that of GLP-1, which resulted in 55% inhibition (0.15 vs. 0.06 µg/g body wt, respectively). This difference in potency was further magnified by our use of a degradation-resistant analog of GLP-2 (44) compared with the very rapidly degraded native form of GLP-1 (9). Consistent with our findings, previous studies have demonstrated that the effects of GLP-2 on antral motility are markedly less potent than those of GLP-1 (29). However, the only prior study of GLP-2 and small intestinal motility in rodents has shown an additive effect of GLP-2 to augment GLP-1 inhibition of motility but no independent actions of GLP-2 (5). Furthermore, as also found for GLP-1, GLP-2 failed to inhibit GI transit in GFR{alpha}2 KO mice, despite the presence of GLP-2R mRNA transcripts in the tunica muscularis. Although Guan et al. (20) have colocalized the GLP-2R to enteric neurons expressing VIP, hallmarks of inhibitory motoneurons (16), no reduction in VIP-expressing neurons was detected in the present study. However, it is possible that GLP-2 mediates its anti-transit effect by inhibiting the release of a factor such as serotonin, thereby reducing excitatory cholinergic transmission of the GI tract. This is consistent with the demonstration of GLP-2R expression on serotonin-expressing enterochromaffin cells in the mucosa (48) that are known to stimulate intestinal motility in response to luminal stimuli (18). Interestingly, the majority of synaptophysin-expressing enteroendocrine cells in the mouse jejunum were represented by enterochromaffin cells, as also reported by others (35), and these were reduced in number in the GFR{alpha}2 KO mice. The mechanisms underlying these changes are not known but may provide an explanation for the inability of GLP-2 to inhibit motility in the GFR{alpha}2-deficient animals. A relationship between GLP-2, enterochromaffin cells, and intestinal transit has been further suggested by a recent study demonstrating that ileitis increases the numbers of colonic GLP-2- and serotonin-expressing cells in association with colonic hyperexcitability (31). Nonetheless, it remains possible that GLP-2 exerted its effects through a GLP-1R-dependent pathway in the gut, as suggested by others (5). For example, the ability of the structurally related peptide, glicentin, to inhibit colonic smooth muscle motility is blocked by the GLP-1R antagonist exendin9–39 (2). Moreover, Drucker and coworkers (26) have demonstrated that GLP-2 signaling in the central nervous system (CNS) is potentiated in the GLP-1R KO mouse, suggesting that GLP-2 can signal upstream of GLP-1, at least in central neurons. Thus, although the present study clearly identifies GLP-2 as an inhibitor of intestinal transit in mice, the physiological significance of these findings remains unclear.

Although GLP-2 has been demonstrated to activate tetrodotoxin-dependent c-fos expression in crypt cells, leading to the suggestion that the enteric GLP-2R is important for the intestinotropic effects of GLP-2 (3), no differential growth responses to GLP-2 administration were found in GFR{alpha}2 KO mice. The simplest interpretation for this is that the growth effects of GLP-2 are not directly mediated by receptor signaling in the ENS. Several lines of evidence suggest that this may be the case. First, the GLP-2R has recently been sublocalized to vasodilatory and/or inhibitory motoneurons that do not project to the intestinal epithelium (4, 16). Secondly, recent studies have demonstrated that the intestinotropic actions of GLP-2 are mediated by growth factors, such as insulin-like growth factor-1, insulin-like growth factor-2, and keratinocyte growth factor, that arise from the intestinal mucosa (15, 32), rather than by enteric neurotransmitters/neuropeptides. Nonetheless, because there was no decrease in GLP-2R expression in the tunica muscularis of KO animals, it remains possible that the ENS may mediate some of the intestinotropic effect of GLP-2 indirectly. Furthermore, a very recent study has implicated the submucosal GLP-2R in GLP-2 effects on crypt cell proliferation in rodent models of inflammation (41). This finding suggests that the present model of myenteric deficit may not have been optimal to investigate the tropic effects of GLP-2.

Studies in other murine models have demonstrated that GLP-2 stimulates intestinal growth in the small intestine, as well as in the colon, although greatest effects are typically observed in the jejunum (46). These actions of GLP-2 in the normal mouse intestine are thought to be mediated, at least in part, through enhancement of crypt cell proliferation (14, 15). However, despite the increase in both intestinal weight and villus height observed in response to treatment with GLP-2, no enhancement of crypt cell proliferation could be detected in the present study, as determined by both Ki-67 staining and BrdU uptake. Interestingly, the GFR{alpha}2 mice were studied on a mixed C57Bl/6 x 129Sv/J background, and a similar absence of effect of GLP-2 to increase Ki-67 positivity has been observed in mice on a pure 129Sv/J background, compared with the stimulatory effects of GLP-2 that were observed in identically treated CD1 animals (15). Collectively, these findings provide further support for the notion of strain-specific differences in proliferation in response to chronic administration of GLP-2 in mice (15). As to the mechanisms underlying the GLP-2-induced small increase in villus height in both the WT and KO animals, no effect to reduce apoptosis or increase cell width was detected. However, the effects of GLP-2 on detachment-induced cell anoikis, as well as on villus cell necrosis remain unknown, and changes in these parameters may well contribute to the increased jejunal villus height observed in GLP-2-treated mice. When taken together, these findings suggest that 129Sv/J mice may be an interesting model whereby to study nonproliferative growth effects of GLP-2. Finally, all of the GLP-2-induced changes in differentiated cell numbers were found to occur in parallel with the increases in villus height, suggesting that GLP-2 had no effect on cell lineage differentiation. Similar results have been reported by others for the effects of GLP-2 on small intestinal mucus cell numbers (3).

In summary, the results of the present study demonstrate a role for GFR{alpha}2-expressing enteric neurons in the downstream signaling of the glucagon-like peptides to inhibit GI motility but do not show a role for these neurons in either basal or GLP-2-stimulated intestinal growth. A better understanding of the mechanism of action of the glucagon-like peptides in the intestine may lead to novel therapeutic approaches to treat patients with disordered intestinal motility.


   GRANTS
TOP
 ABSTRACT
 METHODS
 RESULTS
 DISCUSSION
 GRANTS
REFERENCES
 
This work was supported by an operating grant from the Canadian Institutes of Health Research and an equipment grant from the Banting and Best Diabetes Centre, University of Toronto. P. L. Brubaker was supported by the Canada Research Chairs Program.


   ACKNOWLEDGMENTS
 
GFR{alpha}2 mice were a kind gift from Dr. J. Milbrandt (Washington University School of Medicine, St. Louis, MO).


   FOOTNOTES
 

Address for reprint requests and other correspondence: P. L. Brubaker, Rm. 3366 Medical Sciences Bldg., Univ. of Toronto, 1 King's College Circle, Toronto, ON M5S 1A8 Canada (e-mail: p.brubaker{at}utoronto.ca)

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.


   REFERENCES
TOP
 ABSTRACT
 METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 

  1. Airaksinen MS, Saarma M. The GDNF family: signaling, biological functions and therapeutic value. Nat Rev Neurosci 3: 383–394, 2002.[CrossRef][Web of Science][Medline]
  2. Ayachi SE, Borie F, Magous R, Sasaki K, Le Nguyen D, Bali JP, Millat B, Jarrousse C. Contraction induced by glicentin on smooth muscle cells from the human colon is abolished by exendin (9–39). Neurogastroenterol Motil 17: 302–309, 2005.[CrossRef][Web of Science][Medline]
  3. Bjerknes M, Cheng H. Modulation of specific intestinal epithelial progenitors by enteric neurons. Proc Natl Acad Sci USA 98: 12497–12502, 2001.[Abstract/Free Full Text]
  4. Blonde L, Klein EJ, Han J, Zhang B, Mac SM, Poon TH, Taylor KL, Trautmann ME, Kim DD, Kendall DM. Interim analysis of the effects of exenatide treatment on A1C, weight and cardiovascular risk factors over 82 weeks in 314 overweight patients with type 2 diabetes. Diabetes Obes Metab 8: 436–447, 2006.[CrossRef][Web of Science][Medline]
  5. Bozkurt A, Naslund E, Holst JJ, Hellstrom PM. GLP-1 and GLP-2 act in concert to inhibit fasted, but not fed, small bowel motility in the rat. Regul Pept 107: 129–135, 2002.[CrossRef][Web of Science][Medline]
  6. Brubaker PL, Izzo A, Hill M, Drucker DJ. Intestinal function in mice with small bowel growth induced by glucagon-like peptide-2. Am J Physiol Endocrinol Metab 272: E1050–E1058, 1997.[Abstract/Free Full Text]
  7. Campos RV, Lee YC, Drucker DJ. Divergent tissue-specific and developmental expression of receptors for glucagon and glucagon-like peptide-1 in the mouse. Endocrinology 134: 2156–2164, 1994.[Abstract]
  8. Crowell MD, Mathis C, Schettler VA, Yunus T, Lacy BE. The effects of tegaserod, a 5-HT receptor agonist, on gastric emptying in a murine model of diabetes mellitus. Neurogastroenterol Motil 17: 738–743, 2005.[CrossRef][Web of Science][Medline]
  9. Deacon CF, Knudsen LB, Madsen K, Wiberg FC, Jacobsen O, Holst JJ. Dipeptidyl peptidase IV resistant analogues of glucagon-like peptide-1 which have extended metabolic stability and improved biological activity. Diabetologia 41: 271–278, 1998.[CrossRef][Web of Science][Medline]
  10. Demchyshyn LL, Wang H. Teduglutide (ALX-0600), a glucagon-like peptide-2 (GLP-2) analogue inhibits contractility in the colon ex vivo. Gastroenterology 130 Suppl 2: W1605, 2006.
  11. Dhanvantari S, Seidah NG, Brubaker PL. Role of prohormone convertases in the tissue-specific processing of proglucagon. Mol Endocrinol 10: 342–355, 1996.[Abstract/Free Full Text]
  12. Dong DL, Wang QH, Chen W, Fan JJ, Mu JW, Ke J, Yang BF. Contrasting effects of tetraethylammonium and 4-aminopyridine on the gastrointestinal function of mice. Eur J Pharmacol 509: 179–185, 2005.[CrossRef][Web of Science][Medline]
  13. Drucker DJ. Biologic actions and therapeutic potential of the proglucagon-derived peptides. Nat Clin Pract Endocrinol Metab 1: 22–31, 2005.[CrossRef][Web of Science][Medline]
  14. Drucker DJ, Ehrlich P, Asa SL, Brubaker PL. Induction of intestinal epithelial proliferation by glucagon-like peptide 2. Proc Natl Acad Sci USA 93: 7911–7916, 1996.[Abstract/Free Full Text]
  15. Dube PE, Forse CL, Bahrami J, Brubaker PL. Essential role of insulin-like growth factor-1 in the intestinal tropic effects of glucagon-like peptide-2 in mice. Gastroenterology 131: 589–605, 2006.[CrossRef][Web of Science][Medline]
  16. Furness JB. Types of neurons in the enteric nervous system. J Auton Nerv Syst 81: 87–96, 2000.[CrossRef][Web of Science][Medline]
  17. Gershon MD. Lessons from genetically engineered animal models. II Disorders of enteric neuronal development: insights from transgenic mice. Am J Physiol Gastrointest Liver Physiol 277: G262–G267, 1999.[Abstract/Free Full Text]
  18. Gershon MD. Review article: roles played by 5-hydroxytryptamine in the physiology of the bowel. Aliment Pharmacol Ther 13, Suppl 2: 15–30, 1999.
  19. Giralt M, Vergara P. Sympathetic pathways mediate GLP-1 actions in the GI tract of the rat. Regul Pept 74: 19–25, 1998.[CrossRef][Web of Science][Medline]
  20. Guan X, Karpen HE, Stephens J, Bukowski JT, Niu S, Zhang G, Stoll B, Finegold MJ, Holst JJ, Hadsell D, Nichols BL, Burrin DG. GLP-2 receptor localizes to enteric neurons and endocrine cells expressing vasoactive peptides and mediates increased blood flow. Gastroenterology 130: 150–164, 2006.[CrossRef][Web of Science][Medline]
  21. Imeryüz N, Yegen BÇ, Bozkurt A, Coskun T, Villanueva-Peñacarrillo ML, Ulusoy NB. Glucagon-like peptide-1 inhibits gastric emptying via vagal afferent-mediated central mechanisms. Am J Physiol Gastrointest Liver Physiol 273: G920–G927, 1997.[Abstract/Free Full Text]
  22. Jain S, Naughton CK, Yang M, Strickland A, Vij K, Encinas M, Golden J, Gupta A, Heuckeroth R, Johnson EM Jr, Milbrandt J. Mice expressing a dominant-negative Ret mutation phenocopy human Hirschsprung disease and delineate a direct role of Ret in spermatogenesis. Development 131: 5503–5513, 2004.[Abstract/Free Full Text]
  23. Jeppesen PB. Glucagon-like peptide-2: update of the recent clinical trials. Gastroenterology 130: S127–S131, 2006.[CrossRef][Web of Science][Medline]
  24. Kreymann B, Ghatei MA, Williams G, Bloom SR. Glucagon-like peptide-1 7–36: a physiological incretion in man. Lancet 2: 1300–1304, 1987.[Web of Science][Medline]
  25. Llewellyn-Smith IJ, Costa M, Furness JB, Bornstein JC. Structure of the tertiary component of the myenteric plexus in the guinea-pig small intestine. Cell Tissue Res 272: 509–516, 1993.[CrossRef][Web of Science][Medline]
  26. Lovshin J, Estall J, Yusta B, Brown TJ, Drucker DJ. Glucagon-like peptide (GLP)-2 action in the murine central nervous system is enhanced by elimination of GLP-1 receptor signaling. J Biol Chem 276: 21489–21499, 2001.[Abstract/Free Full Text]
  27. Lovshin JA, Huang Q, Seaberg R, Brubaker PL, Drucker DJ. Extrahypothalamic expression of the glucagon-like peptide-2 receptor is coupled to reduction of glutamate-induced cell death in cultured hippocampal cells. Endocrinology 145: 3495–3506, 2004.[Abstract/Free Full Text]
  28. Nagell CF, Wettergren A, Orskov C, Holst JJ. Inhibitory effect of GLP-1 on gastric motility persists after vagal deafferentation in pigs. Scand J Gastroenterol 41: 667–672, 2006.[CrossRef][Web of Science][Medline]
  29. Nagell CF, Wettergren A, Pedersen JF, Mortensen D, Holst JJ. Glucagon-like peptide-2 inhibits antral emptying in man, but is not as potent as glucagon-like peptide-1. Scand J Gastroenterol 39: 353–358, 2004.[CrossRef][Web of Science][Medline]
  30. Nakagawa A, Satake H, Nakabayashi H, Nishizawa M, Furuya K, Nakano S, Kigoshi T, Nakayama K, Uchida K. Receptor gene expression of glucagon-like peptide-1, but not glucose-dependent insulinotropic polypeptide, in rat nodose ganglion cells. Auton Neurosci 110: 36–43, 2004.[CrossRef][Web of Science][Medline]
  31. O'Hara JR, Lomax AE, Mawe GM, Sharkey KA. Ileitis alters neuronal and enteroendocrine signaling in guinea pig distal colon. Gut 56: 186–194, 2007.[Abstract/Free Full Text]
  32. Orskov C, Hartmann B, Poulsen SS, Thulesen J, Hare KJ, Holst JJ. GLP-2 stimulates colonic growth via KGF, released by subepithelial myofibroblasts with GLP-2 receptors. Regul Pept 124: 105–112, 2005.[CrossRef][Web of Science][Medline]
  33. Orskov C, Holst JJ, Knuhtsen S, Baldissera FGA, Poulsen SS, Nielsen OV. Glucagon-like peptides GLP-1 and GLP-2, predicted products of the glucagon gene, are secreted separately from pig small intestine but not pancreas. Endocrinology 119: 1467–1475, 1986.[Abstract/Free Full Text]
  34. Pfaffl MW. A new mathematical model for relative quantification in real-time RT-PCR. Nucleic Acids Res 29: e45, 2001.[Abstract/Free Full Text]
  35. Portela-Gomes GM, Stridsberg M, Johansson H, Grimelius L. Co-localization of synaptophysin with different neuroendocrine hormones in the human gastrointestinal tract. Histochem Cell Biol 111: 49–54, 1999.[CrossRef][Web of Science][Medline]
  36. Rossi J, Herzig KH, Voikar V, Hiltunen PH, Segerstrale M, Airaksinen MS. Alimentary tract innervation deficits and dysfunction in mice lacking GDNF family receptor alpha2. J Clin Invest 112: 707–716, 2003.[CrossRef][Web of Science][Medline]
  37. Rossi J, Luukko K, Poteryaev D, Laurikainen A, Sun YF, Laakso T, Eerikainen S, Tuominen R, Lakso M, Rauvala H, Arumae U, Pasternack M, Saarma M, Airaksinen MS. Retarded growth and deficits in the enteric and parasympathetic nervous system in mice lacking GFR alpha2, a functional neurturin receptor. Neuron 22: 243–252, 1999.[CrossRef][Web of Science][Medline]
  38. Schirra J, Nicolaus M, Roggel R, Katschinski M, Storr M, Woerle HJ, Goke B. Endogenous glucagon-like peptide 1 controls endocrine pancreatic secretion and antro-pyloro-duodenal motility in humans. Gut 55: 243–251, 2006.[Abstract/Free Full Text]
  39. Schmidt PT, Naslund E, Gryback P, Jacobsson H, Hartmann B, Holst JJ, Hellstrom PM. Peripheral administration of GLP-2 to humans has no effect on gastric emptying or satiety. Regul Pept 116: 21–25, 2003.[CrossRef][Web of Science][Medline]
  40. Shin ED, Estall JL, Izzo A, Drucker DJ, Brubaker PL. Mucosal adaptation to enteral nutrients is dependent on the physiologic actions of glucagon-like peptide-2 in mice. Gastroenterology 128: 1340–1353, 2005.[CrossRef][Web of Science][Medline]
  41. Sigalet DL, Wallace LE, Holst JJ, Martin GR, Kaji T, Tanaka H, Sharkey KA. Enteric neural pathways mediate the anti-inflammatory actions of glucagon-like peptide 2. Am J Physiol Gastrointest Liver Physiol doi:10.1152/ajp.00530.2006.
  42. Tago H, Kimura H, Maeda T. Visualization of detailed acetylcholinesterase fiber and neuron staining in rat brain by a sensitive histochemical procedure. J Histochem Cytochem 34: 1431–1438, 1986.[Abstract]
  43. Talsania T, Anini Y, Siu S, Drucker DJ, Brubaker PL. Peripheral exendin-4 and peptide YY(3-36) synergistically reduce food intake through different mechanisms in mice. Endocrinology 146: 3748–3756, 2005.[Abstract/Free Full Text]
  44. Tavares W, Drucker DJ, Brubaker PL. Enzymatic- and renal-dependent catabolism of the intestinotropic hormone glucagon-like peptide-2 in rats. Am J Physiol Endocrinol Metab 278: E134–E139, 2000.[Abstract/Free Full Text]
  45. Tolessa T, Gutniak M, Holst JJ, Efendic S, Hellström PM. Glucagon-like peptide-1 retards gastric emptying and small bowel transit in the rat: effect mediated through central or enteric nervous mechanisms. Dig Dis Sci 43: 2284–2290, 1998.[CrossRef][Web of Science][Medline]
  46. Tsai CH, Hill M, Asa SL, Brubaker PL, Drucker DJ. Intestinal growth-promoting properties of glucagon-like peptide-2 in mice. Am J Physiol Endocrinol Metab 273: E77–E84, 1997.[Abstract/Free Full Text]
  47. Wojdemann M, Wettergren A, Hartmann B, Holst JJ. Glucagon-like peptide-2 inhibits centrally induced antral motility in pigs. Scand J Gastroenterol 33: 828–832, 1998.[CrossRef][Web of Science][Medline]
  48. Yusta B, Huang L, Munroe D, Wolff G, Fantaske R, Sharma S, Demchyshyn L, Asa SL, Drucker DJ. Enteroendocrine localization of GLP-2 receptor expression in humans and rodents. Gastroenterology 119: 744–755, 2000.[CrossRef][Web of Science][Medline]



This article has been cited by other articles:


Home page
 J. Neurosci.Home page
M. A. Brantley Jr, S. Jain, E. E. Barr, E. M. Johnson Jr, and J. Milbrandt
Neurturin-Mediated Ret Activation Is Required for Retinal Function
J. Neurosci., April 16, 2008; 28(16): 4123 - 4135.
[Abstract] [Full Text] [PDF]

This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow All Versions of this Article:
293/2/G461    most recent
00424.2006v1
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
Right arrow Citation Map
Services
Right arrow Email this article to a friend
Right arrow Similar articles in this journal
Right arrow Similar articles in ISI Web of Science
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Citing Articles
Right arrow Citing Articles via HighWire
Right arrow Citing Articles via ISI Web of Science (1)
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by McDonagh, S. C.
Right arrow Articles by Brubaker, P. L.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by McDonagh, S. C.
Right arrow Articles by Brubaker, P. L.

HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS
Visit Other APS Journals Online
Copyright © 2007 by the American Physiological Society.