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Endocrinology, doi:10.1210/en.2007-0561
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Endocrinology Vol. 149, No. 1 291-301
Copyright © 2008 by The Endocrine Society

Glucagon-Like Peptide-2 Activates β-Catenin Signaling in the Mouse Intestinal Crypt: Role of Insulin-Like Growth Factor-I

Philip E. Dubé, Katherine J. Rowland and Patricia L. Brubaker

Departments of Physiology (P.E.D., K.J.R., P.L.B.) and Medicine (P.L.B.), University of Toronto, Toronto, Ontario, Canada M5S 1A8

Address all correspondence and requests for reprints to: Dr. P. L. Brubaker, University of Toronto, Medical Sciences Building, Room 3366, 1 King’s College Circle, Toronto, Ontario, Canada M5S 1A8. E-mail: p.brubaker{at}utoronto.ca.


   Abstract
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 References
 
Chronic administration of glucagon-like peptide-2 (GLP-2) induces intestinal growth and crypt cell proliferation through an indirect mechanism requiring IGF-I. However, the intracellular pathways through which IGF-I mediates GLP-2-induced epithelial tropic signaling remain undefined. Because β-catenin and Akt are important regulators of crypt cell proliferation, we hypothesized that GLP-2 activates these signaling pathways through an IGF-I-dependent mechanism. In this study, fasted mice were administered Gly2-GLP-2 or LR3-IGF-I (positive control) for 0.5–4 h. Nuclear translocation of β-catenin in non-Paneth crypt cells was assessed by immunohistochemistry and expression of its downstream proliferative markers, c-myc and Sox9, by quantitative RT-PCR. Akt phosphorylation and activation of its targets, glycogen synthase kinase-3β and caspase-3, were determined by Western blot. IGF-I receptor (IGF-IR) and IGF-I signaling were blocked by preadministration of NVP-AEW541 and through the use of IGF-I knockout mice, respectively. We found that GLP-2 increased β-catenin nuclear translocation in non-Paneth crypt cells by 72 ± 17% (P < 0.05) and increased mucosal c-myc and Sox9 mRNA expression by 90 ± 20 and 376 ± 170%, respectively (P < 0.05–0.01), with similar results observed with IGF-I. This effect of GLP-2 was prevented by blocking the IGF-IR as well as ablation of IGF-I signaling. GLP-2 also produced a time- and dose-dependent activation of Akt in the intestinal mucosa (P < 0.01), most notably in the epithelium. This action was reduced by IGF-IR inhibition but not IGF-I knockout. We concluded that acute administration of GLP-2 activates β-catenin and proliferative signaling in non-Paneth murine intestinal crypt cells as well as Akt signaling in the mucosa. However, IGF-I is required only for the GLP-2-induced alterations in β-catenin.


   Introduction
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 References
 
GLUCAGON-LIKE PEPTIDE (GLP)-2 is an endogenous meal-induced gut hormone that stimulates intestinal growth, mainly through increased crypt cell proliferation and reduced epithelial apoptosis (1, 2, 3, 4). GLP-2 also enhances the functional capacity of the intestine by modulating digestive enzyme activity, nutrient absorption, epithelial barrier function, inflammatory processes, and intestinal blood flow (5, 6, 7, 8, 9). Combined, these actions suggest that GLP-2 may be clinically beneficial. Indeed, administration of human (h) (Gly2)GLP-2, a long-acting GLP-2 analog (10, 11), has been efficacious in numerous animal models of intestinal disease, and early results from human clinical trials are promising (7, 12, 13, 14, 15, 16, 17, 18). To better understand how GLP-2 may function in physiological or pathophysiological settings, it is therefore important to understand the cellular mechanisms underlying the biological actions of GLP-2.

The GLP-2 receptor (GLP-2R), a G protein-coupled receptor of the glucagon-secretin receptor superfamily, is expressed throughout the gastrointestinal tract, with highest expression in the proximal jejunum (19). Interestingly, the GLP-2R is not found in the intestinal epithelium, a main target of GLP-2 action in vivo; rather, the GLP-2R is expressed in intestinal subepithelial myofibroblasts (SEMFs), the enteric nervous system, and scattered enteroendocrine cells, suggesting that GLP-2 acts through multiple indirect paracrine mechanisms to produce its varied effects in the intestine (20, 21, 22, 23, 24). Indeed, within the enteric nervous system, nitric oxide and vasoactive intestinal polypeptide mediate the up-regulation of intestinal blood flow and the antiinflammatory activity of GLP-2, respectively (8, 9). Interestingly, the SEMFs express a wide variety of growth factors, which, given the coexpression of the GLP-2R, is suggestive of a role for these cells in the regulation of the epithelium. Consistent with this hypothesis, one study has indicated that keratinocyte growth factor mediates some of the effects of GLP-2 in the colon, although this effect appears to be limited to the mucous cell population (22). Our current model is that GLP-2 engages a variety of mediators to produce its multiple actions in the intestine and, specifically, that activation of the GLP-2R in SEMFs modulates the release of growth factors that act on the neighboring crypt and villus epithelial cells to ultimately induce tropic responses.

We recently demonstrated that IGF-I is essential for the growth effects of GLP-2 in the murine small and large intestine, with a specific role in the modulation of small intestinal epithelial proliferation (25). IGF-I is a potent intestinal growth factor expressed locally within the intestine, largely in SEMFs and smooth muscle (26). Rodent studies have shown that the intestine is particularly sensitive to the growth effects IGF-I and that the IGF-I receptor (IGF-IR) is expressed throughout the intestinal epithelium (27, 28, 29, 30, 31). We have shown that mice lacking IGF-I are resistant to GLP-2-induced small intestinal growth, in terms of organ weight, crypt-villus height, and crypt cell proliferation. Furthermore, administration of GLP-2 increases intestinal IGF-I secretion in vitro and enhances intestinal IGF-I mRNA transcript levels both in vitro and in vivo (25). These data suggest that GLP-2 acts through intestinal IGF-I to induce intestinal growth and crypt cell proliferation (Fig. 1Go); however, the mechanisms through which GLP-2 affects the epithelium in an IGF-I-dependent manner are unknown.


Figure 1
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  		FIG. 1. Schematic representation of interactions between GLP-2 and IGF-I in the regulation of intestinal growth. After secretion by the intestinal L cell into the circulation, GLP-2 activates the G protein-coupled GLP-2 receptor in the subepithelial myofibroblast cells, which subtend the epithelium as a syncytium. This leads to release of IGF-I, which then acts in a paracrine fashion on the tyrosine kinase IGF-IR expressed in the proliferative compartment of the crypt.

 
There are several key signaling systems that are critical for intestinal epithelial homeostasis and that may play a role in the IGF-I-dependent tropic actions of GLP-2. These include not only the canonical wingless (Wnt)/β-catenin signaling pathway, which is important in the regulation of crypt cell proliferation and differentiation (32, 33) but also the phosphatidylinositol-3 kinase (PI3-K)/Akt pathway, a well-established regulator of epithelial cell proliferation, survival, and function (34, 35). GLP-2 has been demonstrated to activate Akt in the neonatal pig intestine (4, 36), but it is unknown whether IGF-I is required for this response or whether this effect can be generalized to other species or the mature intestine. However, several reports have demonstrated that the IGFs can activate β-catenin signaling through PI3-K/Akt-mediated inhibition of glycogen synthase kinase (GSK)-3β and activation of Ras, leading to both the stabilization of cytoplasmic β-catenin and its nuclear translocation (37, 38, 39, 40). We therefore hypothesized that GLP-2 induces both β-catenin and Akt signaling in the intestinal epithelium through an IGF-IR/IGF-I-dependent mechanism. To test this hypothesis, we developed a novel in vivo model to examine the acute tropic signaling effects of GLP-2 in the fasted mouse intestine and investigated the role of the IGF-I signaling pathway in these effects by pharmacologically inhibiting the IGF-IR (41, 42) and through the use of IGF-I knockout mice (25, 43).


   Materials and Methods
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 References
 
Animals and test agents
Animals consisted of adult female CD1 mice (Charles River Canada, St. Constant, Québec, Canada), adult male C57BL/6 mice (Charles River), and adult IGF-I knockout (Igf1–/–) mice. Igf1–/– mice, backcrossed on a CD1 background, were a kind gift of Drs. V. Han (University of Western Ontario, Ontario, Canada) and C. Bondy (National Institutes of Health, Bethesda, MD) (43) and were bred and genotyped as previously described (25, 43). Before experimental protocols, animals were given ad libitum access to water and standard rodent chow (Purina 5001; Purina Mills, St. Louis, MO) in a barrier animal facility at the University of Toronto with a 12-h light, 12-h dark cycle. All animal protocols were approved by the Animal Care Committee of the University of Toronto.

The degradation-resistant, long-acting GLP-2 analog, h(Gly2)GLP-2 (10, 11) (GLP-2), was from Peptidec Technologies (Pierrefonds, Québec, Canada). The identity and purity of the peptide were previously confirmed using mass spectrometry, amino acid analysis, and N-terminal Edman sequencing (25, 44). Long-R3-IGF-I (LR3IGF-I), an IGF-I analog with decreased affinity for IGF binding proteins (45), was from GroPep (Thebarton, South Australia, Australia). Wortmannin, a PI3-K inhibitor, was from Sigma-Aldrich (Oakville, Ontario, Canada). The IGF-IR inhibitor, NVP-AEW541 (41, 42), was a kind gift of Novartis Pharmaceuticals (Basel, Switzerland).

Experimental protocols
Before all experimental protocols, mice were fasted overnight in individual cages, with ad libitum access to water. Some animals were then pretreated with wortmannin [1.5 mg/kg in 4% (vol/vol) methanol in saline], NVP-AEW541 [50 mg/kg in 25 mM L(+)tartaric acid], or respective vehicles by orogastric gavage 30 min before subsequent treatments. Mice were then administered (ip) either h(Gly2)GLP-2 (0.05 ng/g to 5 µg/g; 1 ng to 100 µg total), LR3IGF-I (0.05–0.5 µg/g; 1–10 µg total), or saline (25 µl/g; control) and were killed 30 min to 4 h later. Blood glucose was measured using the One Touch Basic glucose meter (Lifescan Canada, Burnaby, British Columbia, Canada). LR3IGF-I treatment, but not h(Gly2)GLP-2, decreased blood glucose levels, compared with saline (P < 0.05). NVP-AEW541 did not alter basal blood glucose levels, compared with vehicle and saline-treated mice, and prevented the LR3IGF-I-induced decrease in blood glucose (P = 0.09). The jejunum (5–10 cm immediately proximal to the mid-small intestine) was removed and either fixed in 10% neutral buffered formalin (Sigma-Aldrich) for immunohistochemical analysis or rinsed gently with ice-cold saline and frozen on dry ice for subsequent biochemical analyses.

Immunohistochemistry
Immunohistochemical analyses were performed on 4-µm transverse cross-sections of paraffin-embedded jejunum. Sections were rehydrated in a graded ethanol series, and heat-induced antigen retrieval was performed by microwave heating under pressure for 20 min in a 10-mM citrate buffer (pH 6.0). Blocking was performed by incubation in 5% (vol/vol) normal goat serum (Jackson ImmunoResearch, West Grove, PA) for 30 min at 37 C before incubation with primary antibodies overnight at 4 C. For β-catenin staining, the primary antibody was mouse anti-β-catenin (1:200 dilution; BD Biosciences, Mississauga, Ontario, Canada) and, in some sections, this was used in conjunction with either rabbit antilysozyme (1:300 dilution; Dako Canada, Inc., Mississauga, Ontario, Canada) or rat anti-Ki-67 (1:200 dilution; Dako Canada, Inc.). For β-catenin detection, endogenous peroxidase was quenched in 1% (vol/vol) hydrogen peroxide, followed by indirect immunoperoxidase detection using biotinylated goat antimouse secondary antibody (Vector Laboratories, Burlington, Ontario, Canada) and streptavidin-horseradish peroxidase (Vector Laboratories), with 3,3'-diaminobenzidine (Sigma-Aldrich) or tyramide-Cy3 (PerkinElmer, Waltham, MA) as the peroxidase substrate. Lysozyme was visualized using an Alexa 488-conjugated goat antirabbit secondary antibody (Invitrogen Canada Inc., Burlington, Ontario, Canada). For β-catenin and Ki-67 double immunofluorescence, β-catenin was visualized with a fluorescein-conjugated donkey antimouse secondary antibody (fluorescein isothiocyanate; Jackson ImmunoResearch), and Ki-67 was detected using indirect immunoperoxidase staining with a biotinylated donkey antirat secondary antibody (Vector Laboratories) and tyramide-Cy3, as above. For phosphorylated (p)-Akt staining, the primary antibody was rabbit anti-phospho-S473-Akt (1:50 dilution, immunohistochemistry-specific antibody; Cell Signaling Technology, Danvers, MA). Visualization of p-Akt staining was performed using a Cy3-conjugated goat antirabbit secondary antibody (Jackson ImmunoResearch). Nuclei were visualized by 4'-6-diamidino-2-phenylindole (DAPI) staining (Vector Laboratories). Antibody specificity was confirmed in negative control sections lacking primary antibodies (data not shown). Digital images were captured using a Zeiss AxioPlan epifluorescence microscope, with consistent light and exposure settings (Carl Zeiss Canada, Don Mills, Ontario, Canada).

Quantification of β-catenin nuclear translocation in jejunal crypt cells was performed after first excluding lysozyme-positive (i.e. Paneth) cells, which possessed constitutive nuclear β-catenin localization (Fig. 2Go). Positive β-catenin nuclear localization was therefore defined as a non-Paneth crypt cell exhibiting β-catenin and DAPI colocalization. Upper crypt and/or villus cells were used as negative controls for nuclear localization. For each mouse, an average of 48-well-oriented and intact half-crypts were scored. All analyses were performed in a blinded manner.


Figure 2
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  		FIG. 2. Localization of nuclear β-catenin and p-Akt in murine small intestinal crypt cells. A, Representative immunohistochemical staining for β-catenin in a mouse jejunal crypt, visualized by 3,3'-diaminobenzidine staining (brown). β-Catenin displayed membrane (black arrowheads) as well as nuclear (pink arrowheads) staining. B, Representative immunohistochemical staining for β-catenin in a mouse jejunal crypt, visualized by tyramide-Cy3 staining (red) as well as for lysozyme (green) and nuclear DNA (DAPI; blue); right panel shows the same intestinal crypt with β-catenin staining alone; scale bars, 16 µm. Intestinal crypt cells were classified according to the presence (pink arrowheads) or absence (blue arrowheads) of nuclear β-catenin staining. Paneth cells, which contain constitutive nuclear β-catenin (green arrowheads), were excluded from the cell count. C, Double immunofluorescence for β-catenin (green) and Ki-67 (red) in jejunal crypt of a mouse treated with saline (upper panels) or 0.05 µg/g h(Gly2)GLP-2 (lower panels) for 30 min; nuclei were identified by DAPI staining (blue); scale bar, 20 µm. Three different cell types were noted: nuclear β-catenin-positive proliferating cells (pink arrowheads); nuclear β-catenin-negative proliferating cells (blue arrowheads); and nuclear β-catenin-positive nonproliferating cells (green arrowheads). D, Representative immunohistochemical staining for phosphorylated S473 Akt (red) in jejunal villi (i and ii) and crypts (iii and iv) of mice treated with either saline (SAL, i and iii) or 0.05 µg/g h(Gly2)GLP-2 (GLP, ii and iv) for 30 min; scale bars, 30 µm. Samples were processed in parallel, and photomicrograph pairs (i and ii, iii and iv) were taken with identical microscope, light, and exposure settings.

 
Western blot
Mucosa and muscularis fractions were obtained by thawing jejunal sections on ice and then gently scraping the tissue with a glass slide to separate the layers. Whole jejunum and isolated mucosa and muscularis were homogenized in ice-cold buffer containing 50 mM β-glycerol-phosphate, 10 mM HEPES, 1% (vol/vol) Triton X-100, 70 mM sodium chloride, 2 mM EGTA, 1 mM sodium orthovanadate, 1 mM sodium fluoride (pH 7.4), supplemented with complete protease inhibitor (Roche Diagnostics, Laval, Québec, Canada). Protein concentration was determined using the Bradford method (Bio-Rad Laboratories, Hercules, CA), and 50 µg protein were subjected to SDS-PAGE and transferred to polyvinylidene difluoride membranes (Bio-Rad). Western blot was carried out as previously described (46). Briefly, membranes were blocked with 5% (wt/vol) nonfat dried milk, followed by overnight incubation with primary antibody at 4 C, and detection with horseradish peroxidase-linked secondary antibodies and enhanced chemiluminescence (Amersham Pharmacia Biotech, Piscataway, NJ). Membranes were then stripped in buffer containing 62.5 mM Tris, 2% (wt/vol) sodium dodecyl sulfate and 100 mM β-mercaptoethanol at 50 C for 20 min before probing with additional primary antibodies. Primary antibodies for Western blot analysis consisted of Akt (Cell Signaling), phospho-Ser473-Akt (p-Akt; Cell Signaling), phospho-Ser9-GSK-3β (Cell Signaling), caspase-3 (Cell Signaling), and actin (Sigma-Aldrich).

Real-time RT-PCR
Expression of mRNA transcripts in whole jejunum was quantified as previously described (25). In brief, total RNA was subjected to reverse transcription using SuperScript II and random hexamers (Invitrogen Canada), followed by real-time PCR using TaqMan gene expression assays (Applied Biosystems, Foster City, CA) for c-myc (no. Mm00487803_m1), Sry-type high-mobility-group box (Sox)-9 (no. Mm00448840_m1), and 18S (no. Hs99999901_s1). Relative quantification of c-myc and Sox9 mRNA expression was calculated using the {Delta}{Delta} cycle threshold [{Delta}{Delta}C(t)] method (47). 18S RNA was used as the endogenous control because its expression is unaffected by GLP-2 treatment (25).

Statistical analysis
All data are expressed as mean ± SEM. Statistical significance was established using the Student t test, one-factor ANOVA, followed by post hoc analysis (Bonferroni adjustment for multiple comparisons), or two-factor ANOVA followed by one-factor analysis or pair-wise Bonferroni comparisons, as appropriate (GraphPad Software, San Diego, CA). Statistical significance was defined as P < 0.05. Factors included in ANOVA analyses consisted of the effect of h(Gly2)GLP-2/LR3IGF-I for one-factor analyses and the effects of h(Gly2)GLP-2/LR3IGF-I and wortmannin, NVP-AEW541, or genotype (wild-type vs. Igf1–/–) for two-factor analyses.


   Results
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 References
 
In all crypt cells, β-catenin staining displayed intense membrane localization, particularly bordering adjacent crypt cells, consistent with the role of β-catenin in adherens junction formation (33) (Fig. 2AGo). However, some cells also demonstrated nuclear β-catenin staining. Because the identity of these cells could not be determined from analysis of β-catenin alone, sections were then costained for both β-catenin and lysozyme, a marker of Paneth cells (Fig. 2BGo). Paneth cells, which are known to possess constitutively active β-catenin signaling (48, 49), demonstrated intense nuclear localization of β-catenin. However, the occasional non-Paneth crypt cell also displayed enriched nuclear localization of β-catenin, accompanied by diffuse cytoplasmic staining, whereas the majority of non-Paneth crypt cells lacked nuclear β-catenin.

To determine whether GLP-2 and/or IGF-I induce β-catenin signaling in non-Paneth crypt cells, a novel in vivo model was developed in which fasted mice were assessed for responses to acute administration of saline, 0.05 µg/g h(Gly2)GLP-2, or 0.5 µg/g LR3IGF-I. In mice treated for 30 min with saline, 1.8 ± 0.2 non-Paneth crypt cells per half-crypt were positive for nuclear β-catenin (Fig. 3AGo); h(Gly2)GLP-2 treatment increased this to 3.1 ± 0.3 cells per half-crypt (P < 0.05, n = 3). Treatment with 0.5 µg/g LR3IGF-I similarly increased nuclear β-catenin localization (42 ± 17% increase vs. saline, P = 0.10). To further characterize the cells responding to h(Gly2)GLP-2 treatment with nuclear β-catenin, double immunofluorescence was performed with β-catenin and the proliferative marker, Ki-67 (Fig. 2CGo). In both control and h(Gly2)GLP-2-treated mice, nuclear β-catenin-positive non-Paneth crypt cells were also found to be positive for Ki-67. However, the number of Ki-67-positive cells was greater than that of the nuclear β-catenin-positive cells (8.3 ± 0.4, compared with 1.8–3.1 cells per half-crypt), and thus, many Ki-67-positive cells did not appear to express detectable levels of nuclear β-catenin. Furthermore, the number of Ki-67-positive cells was not changed by acute treatment with h(Gly2)GLP-2. Nonetheless, the colocalization of Ki-67 and nuclear β-catenin is consistent with the known role of β-catenin to induce crypt cell proliferation through the transcriptional activation of a number of genes, including the immediate early gene c-myc (50, 51).


Figure 3
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  		FIG. 3. GLP-2 activates small intestinal crypt cell β-catenin signaling in an IGF-I-dependent manner. A, Quantification of nuclear β-catenin localization in non-Paneth jejunal crypt cells of C57BL/6 mice treated with saline, 0.05 µg/g h(Gly2)GLP-2 (GLP-2), or 0.5 µg/g LR3IGF-I (IGF-I) for 30 min. *, P < 0.05 vs. saline controls. B and C, Real-time RT-PCR quantification of the β-catenin target genes, c-myc (B), and Sox9 (C) mRNA transcript expression in jejunal extracts of CD1 mice treated with saline, 0.05 µg/g h(Gly2)GLP-2 (GLP-2), or 0.5 µg/g LR3IGF-I (IGF-I) for 90 min. Expression was normalized for expression of 18S RNA, and cycle threshold C(t) values for control mice were as follows: c-Myc and 18S, 21.0 and 16.4, respectively; and Sox9 and 18S, 19.9 and 18.5, respectively. *, P < 0.05; **, P < 0.01 vs. saline controls. D, Quantification of GLP-2-induced nuclear β-catenin localization in CD1 mice with IGF-IR inhibition. Mice were pretreated with 25 mM L(+)tartaric acid (vehicle) or IGF-IR inhibitor (NVP-AEW541) 30 min before treatment and quantification of nuclear β-catenin localization in non-Paneth jejunal crypt cells, as in A. ***, P < 0.001 vs. vehicle/saline controls. E, Quantification of GLP-2-induced nuclear β-catenin localization in CD1 IGF-I knockout mice. Nuclear β-catenin localization was determined in non-Paneth jejunal crypt cells of Igf1–/– mice treated as in A. ***, P < 0.001 vs. saline controls.

 
No change in the expression of c-myc mRNA transcripts could be detected 30 min after treatment with either h(Gly2)GLP-2 or LR3IGF-I (data not shown). However, there was a 90 ± 20% increase in c-myc mRNA transcript expression after 90 min in h(Gly2)GLP-2-treated mice (P < 0.05, n = 6) and a 110 ± 40% increase in c-myc mRNA in LR3IGF-I-treated mice (P < 0.01, n = 3) vs. saline-treated animals (Fig. 3BGo). Furthermore, expression of Sox9, a transcription factor that requires β-catenin for its expression in the crypt (52) was also increased by 90 min treatment with either h(Gly2)GLP-2 (by 376 ± 170%, P < 0.01, n = 8) or LR3IGF-I (by 181 ± 91%, P < 0.05, n = 7; Fig. 3CGo). These data indicate that both GLP-2 and IGF-I induce β-catenin signaling in non-Paneth crypt cells of the murine small intestine.

We have previously shown that intestinal growth and, more specifically, crypt cell proliferation induced by chronic GLP-2 treatment requires the actions of IGF-I (25). We therefore tested the hypothesis that the effect of acute GLP-2 treatment on β-catenin in the intestinal crypt requires IGF-IR/IGF-I signaling. Two different and complementary approaches were used: pharmacological inhibition of the IGF-IR and genetic elimination of IGF-I. First, mice were pretreated for 30 min with the IGF-IR kinase inhibitor, NVP-AEW541, at a dose previously shown to inhibit IGF-I signaling in vivo (41, 42), or vehicle, followed by a 30 min treatment with either saline or 0.05 µg/g h(Gly2)GLP-2, and β-catenin signaling in non-Paneth crypt cells was assessed by quantification of nuclear translocation (Fig. 3DGo). In vehicle-pretreated mice, h(Gly2)GLP-2 treatment produced a robust 90 ± 10% increase in nuclear β-catenin-positive intestinal crypt cells (P < 0.001 vs. saline controls, n = 3–4). NVP-AEW541 pretreatment did not affect basal levels of β-catenin nuclear localization; however, NVP-AEW541 pretreatment completely abrogated this response to GLP-2 (P = 0.23). Next, IGF-I signaling was eliminated by using the Igf1–/– mouse, which we have shown to be resistant to the growth effects of chronic GLP-2 treatment (25). Adult Igf1–/– littermates were treated for 30 min with saline, 0.05 µg/g h(Gly2)GLP-2, or 0.5 µg/g LR3IGF-I (positive control), and non-Paneth crypt cell β-catenin signaling was assessed by quantification of nuclear translocation (Fig. 3EGo). In Igf1–/– mice, GLP-2 treatment did not affect the levels of β-catenin nuclear translocation (P = 0.59 vs. saline-treated Igf1–/– mice, n = 4). Nonetheless, these mice demonstrated a significant response to IGF-I treatment, with a 77 ± 3% increase in nuclear β-catenin localization (P < 0.001 vs. saline-treated Igf1–/– animals, n = 4).

The PI3-K/Akt signaling pathway is an important regulator of growth and metabolic responses of the intestinal epithelium and has previously been implicated in the in vivo biological activities of GLP-2 (4, 36, 53). We therefore also examined our mouse model for the effects of h(Gly2)GLP-2 treatment on Akt signaling. Fasted mice were treated with saline, 0.05 ng/g to 5 µg/g h(Gly2)GLP-2, or 0.5 µg/g LR3IGF-I for time periods of 30 min to 4 h, and phosphorylation of Akt at S473, which is known to be critical for Akt activity (54), was assessed by Western blot (Fig. 4Go, A and B). h(Gly2)GLP-2 treatment induced a dose- and time-dependent stimulation of Akt phosphorylation, with maximal effects observed at 1 µg/g h(Gly2)GLP-2 and at t = 30 min (P < 0.001, n = 3–18). LR3IGF-I treatment, as a positive control, also increased Akt phosphorylation, by up to 3060 ± 520%, compared with saline-treated animals.


Figure 4
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  		FIG. 4. GLP-2 activates Akt signaling in the small intestinal mucosa. Characterization of mucosal Akt signaling in jejuna of mice treated with saline (control), h(Gly2)GLP-2 (GLP-2), or LR3IGF-I (IGF-I), as determined by Western blot and normalized as indicated. Representative blots are shown corresponding to the columns on the graphs (A–D). A, Dose-dependent effect of GLP-2 on Akt phosphorylation. Relative quantification of S473 Akt phosphorylation (p-Akt) in mucosal extracts of CD1 and C57BL/6 mice treated with saline, 0.001–100 µg GLP-2, or 10 µg IGF-I for 30 min. ***, P < 0.001 vs. saline-treated control mucosa. B, Time-dependent effect of GLP-2 on Akt phosphorylation. Relative quantification of p-Akt in mucosal extracts of C57BL/6 mice treated with saline for 30 min, 1 µg GLP-2 for 30–240 min, or 10 µg IGF-I for 30 min. *, P < 0.05; ***, P < 0.001 vs. saline-treated control mucosa. C, S9 phosphorylation of GSK-3β in mucosal extracts of CD1 mice treated with saline, 1 µg GLP-2, or 10 µg IGF-I for 30 min. D, Relative quantification of caspase-3 cleavage in mucosal extracts of CD1 mice treated with saline for 30 min, 1 µg GLP-2 for 30–240 min, or 10 µg IGF-I for 30 min, as determined by Western blot for full-length (35 kDa; i.e. inactive) and cleaved (17 kDa; i.e. active) caspase-3. *, P < 0.05 vs. saline-treated control mucosa. E, Effect of GLP-2 on mucosal vs. muscularis p-Akt in CD1 mice. Relative quantification of p-Akt in whole intestinal extracts as well as adjacent mucosal and muscularis extracts of the same mice treated with saline, 1 µg GLP-2, or 10 µg IGF-I for 30 min. Values are presented as the relative mucosal and muscularis contributions to whole tissue Akt phosphorylation, normalized for respective tissue protein content. **, P < 0.01; ***, P < 0.001 vs. saline-treated control mucosa; §§, P < 0.01 vs. saline-treated muscularis.

 
To confirm that increased Akt phosphorylation represented Akt activation, the levels of phospho-Ser9-GSK-3β, a known Akt substrate (55), were determined in mucosal extracts of mice treated for 30 min with saline, 0.05 µg/g h(Gly2)GLP-2, or 0.5 µg/g LR3IGF-I (Fig. 4CGo). There was a trend toward increased GSK-3β phosphorylation (230 ± 90% vs. saline-treated controls; P = 0.08, n = 3). Because Akt also opposes apoptosis through the inhibition of cascades leading to caspase-3 cleavage (56), we also determined whether h(Gly2)GLP-2 treatment affected the levels of cleaved (active) caspase-3 in mucosal extracts of these mice (Fig. 4DGo). The ratio of cleaved to full-length caspase-3 in saline-treated mice was 1.5 ± 0.3, and treatment with h(Gly2)GLP-2 for 30 min decreased this ratio to 0.9 ± 0.1 (P < 0.05, n = 5–8). Finally, because Akt is ubiquitously distributed, we ascertained the relative effect of GLP-2 on Akt phosphorylation in the isolated intestinal mucosa, compared with the muscularis compartment (Fig. 4EGo). Whereas LR3IGF-I treatment increased Akt phosphorylation in both the mucosa (P < 0.001, n = 3) and the muscularis (P < 0.01, n = 3), h(Gly2)GLP-2 treatment affected only Akt phosphorylation in the mucosa (P < 0.01 vs. saline-treated mice for the mucosa only). Furthermore, immunostaining for phosphorylated S473 Akt confirmed the presence of this activated enzyme throughout the intestinal mucosa, and particularly in villus and crypt epithelial cells, in both saline- and h(Gly2)GLP-2-treated animals (Fig. 2DGo), demonstrating that Akt phosphorylation in response to GLP-2 is not specific to the intestinal crypt cell. Therefore, in contrast to β-catenin, Akt phosphorylation may not be a suitable marker for the proliferative actions of GLP-2 in this bioassay, given that the proliferative cells are restricted to the crypt compartment.

Finally, to test the hypothesis that IGF-IR/IGF-I signaling is required for this effect of GLP-2 on Akt signaling in the mouse intestine, we used a similar strategy as in the prior β-catenin studies. First, as a control study to verify that the effects of GLP-2 or IGF-I on Akt phosphorylation could be blocked, mice were pretreated with wortmannin, a well-established inhibitor of PI3-K (Fig. 5AGo). Wortmannin completely abrogated the phosphorylation of Akt in response to either h(Gly2)GLP-2 or LR3IGF-I treatment, whereas normal responses were maintained in vehicle-treated animals (P < 0.05, n = 4). Pretreatment with the IGF-IR kinase inhibitor, NVP-AEW541, markedly reduced the Akt response to both h(Gly2)GLP-2 and LR3IGF-I (Fig. 5BGo; ANOVA effect of NVP-AEW541, P < 0.001, n = 4–10). However, there was a nonsignificant trend toward increased Akt phosphorylation in the NVP-AEW541/h(Gly2)GLP-2-treated mice (160 ± 70% greater than NVP-AEW541/saline controls, P = 0.19), and although NVP-AEW541 prevented the effects of a low dose of LR3IGF-I on Akt, it did not block the effects of the higher dose of LR3IGF-I (P < 0.01; Fig. 5BGo). Therefore, to ascertain whether this reflected a true role for IGF-I in the Akt response to GLP-2, Igf1–/– and wild-type mice were also assessed for Akt phosphorylation 30 min after treatment with saline, 0.05 µg/g h(Gly2)GLP-2, or 0.5 µg/g LR3IGF-I (Fig. 5CGo). In contrast to the results with IGF-IR inhibition, the effect of GLP-2 on Akt phosphorylation was completely retained in Igf1–/– mice (P < 0.05, compared with saline-treated animals, n = 3–4). The effect of LR3IGF-I treatment was also observed in the Igf1–/– mice, with a 570 ± 100% (P < 0.001) increase in Akt phosphorylation, compared with saline-treated mice.


Figure 5
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  		FIG. 5. IGF-I is not strictly required for GLP-2-induced small intestinal Akt phosphorylation. A–C, CD1 mice were treated with saline, 0.05 µg/g h(Gly2)GLP-2 (GLP-2), or 0.5 µg/g LR3IGF-I (IGF-I) for 30 min; relative quantification of phosphorylated S473 Akt (p-Akt), normalized for actin, was determined by Western blot of jejunal extracts (left); representative blots are shown on the right. A, Effect of PI3-K inhibition on GLP-2-induced Akt phosphorylation. Mice were administered 4% methanol in saline (vehicle) or the PI3-K inhibitor, wortmannin (1.5 mg/kg), 30 min before treatment. B, Effect of IGF-IR inhibition on GLP-2-induced Akt phosphorylation. Mice were administered 25 mM L(+)tartaric acid (vehicle) or the IGF-IR inhibitor, NVP-AEW541 (50 mg/kg), 30 min before treatment. C, Effect of IGF-I knockout on GLP-2-induced Akt phosphorylation. Wild-type or Igf1–/– mice were treated with saline, GLP-2, or IGF-I for 30 min. *, P < 0.05; **, P < 0.01; ***, P < 0.001 vs. respective saline-treated controls.

 

   Discussion
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
References
 
The mechanisms through which GLP-2 affects intestinal growth are unclear but likely involve the coordinated action of paracrine growth factors released locally in the intestine. Importantly, it remains unknown how such factors mediate growth of the intestinal epithelium in response to GLP-2. In part, this is due to a paucity of appropriate in vitro models that recapitulate the apparent complexity of GLP-2 biology. Nonetheless, our recent in vivo studies have demonstrated an essential role for IGF-I in mediating GLP-2-induced small and large intestinal growth as well as in GLP-2-induced small intestinal epithelial proliferation (25). In the current study, we therefore developed a novel in vivo model to characterize in more detail the tropic signaling events that occur in the murine small intestine on GLP-2 administration. Using this model, we identified a novel action of GLP-2, namely acute activation of the β-catenin signaling pathway in non-Paneth mouse intestinal crypt cells, and have shown that this β-catenin response to GLP-2 requires IGF-IR/IGF-I signaling. Furthermore, we present data that GLP-2 activates Akt signaling in the mouse intestinal mucosa, including the crypt and villus epithelium but not in the muscularis and, furthermore, that this effect is not strictly dependent on IGF-I.

Although β-catenin was initially identified as a contributor to adherens junction formation (33), it is now well recognized to play a central role in the regulation of intestinal epithelial proliferation and differentiation (32, 48, 49, 50, 51). Normally, cytoplasmic β-catenin is bound to the adenomatous polyposis coli complex, which marks it for ubiquitination. Activation of the Wnt signaling cascade inhibits the adenomatous polyposis coli complex, thus increasing the half-life of cytoplasmic β-catenin and allowing its translocation to the nucleus (57). β-Catenin and the Wnt signaling cascade are key regulators of cell fate in the intestinal crypt; by maintaining an undifferentiated phenotype in these cells, β-catenin allows for epithelial proliferation (50). In fact, c-myc, a direct transcriptional target of β-catenin, is required for the maintenance of crypt cell proliferation (51). Furthermore, β-catenin signaling also allows for the differentiation of epithelial cells into the secretory cell lineages, such as Paneth cells (49, 50).

In this paper, we show for the first time that GLP-2 enhances β-catenin signaling in the mouse intestine by causing β-catenin nuclear translocation in some, although not all, Ki-67-positive, proliferative crypt cells and, ultimately, increasing c-myc mRNA expression. GLP-2 also enhanced expression of Sox9, a transcription factor that has been suggested to prevent differentiation of crypt cells, thereby maintaining the progenitor cell compartment (52). These findings suggest that GLP-2-mediated β-catenin regulation is one mechanism through which GLP-2 modulates crypt cell proliferation in the murine jejunum. Although GLP-2 did not directly alter the number of Ki-67-positive cells, this is likely due to the very brief time course used in the present study (e.g. 30 min) because detectable effects of GLP-2 on proliferation in the porcine intestine have been reported to require several days (36). Nonetheless, a relationship among GLP-2, nuclear β-catenin, and crypt cell proliferation is further supported by the fact that elimination of IGF-IR/IGF-I signaling not only prevented β-catenin translocation but also prevented the effect of GLP-2 on epithelial proliferation (25). The activation of β-catenin by GLP-2 may suggest a caution regarding the potential clinical use of GLP-2 because overactivation or dysregulation of β-catenin is a feature common to many types of colorectal cancer and, indeed, a majority of early colorectal adenomas in humans possess one or more activating mutations in the Wnt/β-catenin signaling pathway (32). Because one report shows that GLP-2 can promote chemically induced intestinal tumor growth in mice (58), the ability of this growth factor to activate β-catenin, at least in murine jejunal crypt cells, should be considered when determining the relative risk of GLP-2-based therapies.

Stimulatory effects of IGF-I on β-catenin signaling have been reported in a number of different cell types, including intestinal epithelial cells (37), and the docking protein insulin receptor substrate-1 has been reported to serve as a link among IGF-I, nuclear β-catenin, and proliferation in mouse fibroblasts (40). However, the current studies provide the first evidence for a role of IGF-I in the modulation of intestinal crypt β-catenin signaling in vivo. Using the complementary approaches of pharmacological inhibition of the IGF-IR as well as IGF-I knockout, we have shown that IGF-IR/IGF-I signaling is required for the effect of GLP-2 on β-catenin translocation in the proliferative compartment of the intestinal epithelium. These data also therefore provide important insight into the role of IGF-I as a mediator for the growth effects of GLP-2, consistent with our previous findings demonstrating an essential role for IGF-I in GLP-2-induced intestinal growth (25). This supports our model in which GLP-2 induces paracrine secretion of IGF-I from the intestinal SEMFs, which acts on IGF-IR-expressing epithelial crypt cells to activate β-catenin signaling and, ultimately, increase epithelial proliferation (Fig. 1Go). However, the specific cellular mechanism through which GLP-2 stimulates the expression or secretion of IGF-I from SEMFs is unknown; such knowledge will be necessary to conclusively define how GLP-2 induces IGF-I-dependent intestinal growth and is a major unanswered question in the field.

The pleiotropic serine-threonine kinase, Akt, plays a critical role in the regulation of cellular metabolism, survival, and growth in many different cell types, including the intestinal epithelial cell (34, 35). Although IGF-I is well known to activate Akt, this enzyme is stimulated by many other growth factors and cytokines through class 1A PI3-K as well as by ligands for G protein-coupled receptors through class 1B PI3-K{gamma} (59, 60). Previous studies have demonstrated that GLP-2 activates Akt in the neonatal pig intestinal epithelium, in association with a reduction in apoptotic signaling (4, 36). Furthermore, PI3-K is required for GLP-2-induced glucose uptake through the sodium-dependent glucose transporter-1 in the rat jejunum (53). We have now confirmed that GLP-2 activates Akt in the mouse intestinal mucosa. Of some interest, the dose-response curve for activation of Akt was an inverted bell shape, suggestive of desensitization. Similar shaped curves have been reported for GLP-2 activation of cAMP in multiple different primary cell models (25, 44, 61).

Unexpectedly, although GLP-2-induced activation of Akt was reduced by preadministration of the IGF-IR inhibitor, NVP-AEW541, there was no abrogation of this response in IGF-I null mice. However, the mechanism of action of NVP-AEW541 would include inhibition of signaling by both of the intestinal IGF-IR ligands, IGF-I and IGF-II. These studies therefore suggest that IGF-I specifically is not required for effect of GLP-2 on Akt. Nonetheless, the findings do not exclude a role for other Akt-activating growth factors in the nonproliferative actions of GLP-2 in the intestinal epithelium, such as the regulation of cell survival, nutrient absorption, and/or barrier function. Indeed, we have previously demonstrated a role for IGF-II in GLP-2-induced alterations in mucosal surface area (25). Furthermore, a robust activation of Akt was observed in the nonproliferative villus epithelium, and in addition to IGF-I, there are numerous additional growth factors and hormones that may also induce Akt-mediated epithelial responses, including keratinocyte growth factor (21, 22), epidermal growth factor and related ligands (30, 62, 63), and neurotensin (64, 65). Finally, at least three different isoforms of Akt are known (66), and the antiserum used in the present study recognized all three isoforms. Thus, the possibility of cell-specific roles of these isoforms in the intestinal epithelium must also be considered (67).

To fully understand the mechanism of action of GLP-2 on the intestine, it will be important to determine which factors interact with GLP-2 and the relative importance of each to the effects of GLP-2 on gut growth. Indeed, the relationship between GLP-2 and intestinal growth factors other than IGF-I may be particularly significant, most notably GH (30, 68), given its potential relationship with IGF-I, and R-Spondin-1 (25, 69), a potent intestinal growth factor and activator of the β-catenin signaling pathway. Further study is clearly required to fully characterize the mechanisms through which GLP-2 induces intestinal growth.


   Acknowledgments
 
The authors thank Drs. C. Bondy (National Institutes of Health) and V. Han (University of Western Ontario, Ontario, Canada) for the Igf1–/– mice, Novartis (Geneva, Switzerland) for the generous gift of NVP-AEW541, and Mr. A. Izzo for technical assistance.


   Footnotes
 
This work was supported by an Operating Grant MOP-9940 from the Canadian Institutes of Health Research and an equipment grant from the Banting and Best Diabetes Centre, University of Toronto. P.E.D. was supported by a Doctoral Research Award from the Canadian Institutes of Health Research in partnership with the Canadian Digestive Health Foundation and P.L.B. by the Canada Research Chairs Program.

Disclosure Statement: The authors have nothing to disclose.

First Published Online September 20, 2007

Abbreviations: DAPI, 4'-6-Diamidino-2-phenylindole; GLP, glucagon-like peptide; GLP-2R, GLP-2 receptor; GSK, glycogen synthase; h, human; Igf1–/–, IGF-I knockout; IGF-IR, IGF type 1 receptor; LR3IGF-I, long-R3-IGF-I; p, phosphorylated; PI3-K, phosphatidylinositol-3 kinase; SEMF, subepithelial myofibroblast; Sox, Sry-type high-mobility-group box; Wnt, wingless.

Received April 30, 2007.

Accepted for publication September 12, 2007.


   References
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
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