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Am J Physiol Gastrointest Liver Physiol 292: G1420-G1428, 2007. First published February 8, 2007; doi:10.1152/ajpgi.00504.2006
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HORMONES AND SIGNALING

Phenotypic characterization of taste cells of the mouse small intestine

Kate Sutherland,1,3,* Richard L. Young,2,3,* Nicole J. Cooper,3 Michael Horowitz,2 and L. Ashley Blackshaw1,2,3

1Discipline of Physiology, School of Molecular and Biomedical Sciences and 2Discipline of Medicine, Faculty of Health Sciences, University of Adelaide, Australia; and 3Nerve Gut Research Laboratory, Hanson Institute, Department of Gastroenterology, Hepatology and General Medicine, Royal Adelaide Hospital, Australia

Submitted 29 October 2006 ; accepted in final form 6 February 2007


   ABSTRACT
TOP
 ABSTRACT
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Nutrient-evoked gastrointestinal reflexes are likely initiated by specialized epithelial cells located in the small intestine that detect luminal stimuli and release mediators that activate vagal endings. The G-protein {alpha}-gustducin, a key signal molecule in lingual taste detection, has been identified in mouse small intestine, where it may also subserve nutrient detection; however, the phenotype of {alpha}-gustducin cells is unknown. Immunohistochemistry was performed throughout the mouse small intestine for {alpha}-gustducin, enteroendocrine cell markers 5-HT and glucagon-like peptide-1 (GLP-1), and brush cell markers neuronal nitric oxide synthase and Ulex europaeus agglutinin-1 (UEA-1) lectin binding, singly, and in combination. {alpha}-Gustducin was expressed in solitary epithelial cells of the mid to upper villus, which were distributed in a regional manner with most occurring within the midjejunum. Here, 27% of {alpha}-gustducin cells colabeled for 5-HT and 15% for GLP-1; 57% of {alpha}-gustducin cells colabeled UEA-1, with no triple labeling. {alpha}-Gustducin cells that colabeled for 5-HT or GLP-1 were of distinct morphology and exhibited a different {alpha}-gustducin immunolabeling pattern to those colabeled with UEA-1. Neuronal nitric oxide synthase was absent from intestinal epithelium despite strong labeling in the myenteric plexus. We conclude that subsets of enteroendocrine cells in the midjejunum and brush cells (more generally distributed) are equipped to utilize {alpha}-gustducin signaling in mice. Intestinal taste modalities may be signaled by these enteroendocrine cells via the release of 5-HT, GLP-1, or coexpressed mediators or by brush cells via a nonnitrergic mediator in distinct regions of the intestine.

enteroendocrine cells; {alpha}-gustducin; glucagon-like peptide-1; 5-HT; Ulex europaeus agglutinin-1

SMALL INTESTINAL SENSORY FUNCTION is fundamental to effective modulation of gastrointestinal motor, secretory, and absorptive function as well as satiety by the chemical composition of the luminal content (4, 6, 29). Much of this feedback regulation is conveyed via neural reflexes, consequent upon activation of primary afferents within the gut wall. For example, the presence of carbohydrate and lipid in the proximal intestine inhibits gastric emptying via capsaicin-sensitive vagal afferents in the mucosa, thereby optimizing small intestinal absorption (49), which has been referred to as an "enterogastric reflex." Anterograde tracing studies have shown that vagal sensory endings are present within the parenchyma of mucosal villi, but these do not appear to penetrate the epithelial surface (1), precluding a direct action of luminal nutrient on afferent neurons. This suggests that an intermediary transduction event is required within the epithelium to signal nutrients to vagal sensory endings.

The intestinal epithelium provides many specialized enteroendocrine cell candidates for this role of primary chemosensor, each with direct access to the lumen via apical microvilli and equipped with neuroactive substances capable of activating sensory afferents. For example, luminal carbohydrate has been shown to stimulate the release of 5-hydroxytryptamine (5-HT) from enterochromaffin cells (22) and the incretin hormone glucagon-like peptide-1 (GLP-1) from L cells (33), signals that have been implicated in the regulation of gastric emptying (16, 30). Enteroendocrine cell types show distinct regional distributions throughout the gut that subserve nutrient sensing (32). These reflect their neuromodulator and hormone content and function at various points in the digestive process. Because a number of enteroendocrine mediators may be released in response to a specific nutrient, the chemosensory elements expressed in nutrient-sensing cells along the small intestine may vary. Hence the intestine may sense nutrient at multiple sites and recruit diverse and cascading sensing strategies depending on luminal concentration and length of intestinal exposure (20, 21). However, although paracrine mediators and vagal pathways involved in nutrient-activated gastrointestinal feedback are evident, the molecular identity of transduction mechanisms involved in nutrient detection in the small intestine remain poorly defined.

Nutrients are initially detected as distinct tastes on the surface of the tongue with the primary sensors, the taste cells, contained within taste buds in the lingual epithelium. Detection of tastants by the tongue may alter ingestive behavior by promoting consumption of sweet, energy-dense substances that are of nutritional benefit, while ensuring that bitter tasting, potentially toxic compounds are avoided. The molecular identity of the transduction mechanisms responding to sweet, L-amino acid (umami), and bitter taste on the tongue have been characterized with the discovery of mammalian taste receptors. The T1R and T2R taste receptors are unrelated G protein-coupled receptor families with T1R members that heterodimerize to detect sweet (T1R2+T1R3) and L-amino acid (T1R1+T1R3) tastants (19, 47), whereas T2R receptor family members are diverse and detect bitter taste as homomeric G protein-coupled receptors (25). The G protein gustducin is specifically expressed in taste cells of the tongue and mediates gustatory signals upon activation of T1R or T2R receptors on the apical membrane of these cells (39).

It has recently become apparent that key taste-recognition molecules are also specifically expressed in the mucosa of the gastrointestinal tract. Expression of T1R sweet receptors (T1R2+T1R3) as well as the alpha subunit of gustducin, {alpha}-gustducin, has been demonstrated in the rat and mouse intestinal mucosa as well as in STC-1 cells of a mouse enteroendocrine cell line (5, 14, 36). Bitter taste T2R receptor subtypes have also been reported in the stomach and duodenum of both rat and mouse and in STC-1 cells using RT-PCR-based approaches (46). These observations are consistent with the hypothesis that lingual taste transduction pathways are conserved throughout the alimentary tract and employed in primary intestinal chemosensing pathways.

It is not known which epithelial cell type(s) in the small intestinal mucosa utilize {alpha}-gustducin-mediated signaling. It has been shown by Höfer and colleagues (14) that {alpha}-gustducin expression in the rat duodenum is confined to brush cells, a specialized epithelial cell found scattered within the respiratory and gastrointestinal tract that possesses enough morphological similarities to lingual taste cells to suggest a chemosensory role. Although gastric brush cells are devoid of the secretory granules typical of enteroendocrine cells, they show a high level of immunolabeling for nitric oxide synthase (18), which may implicate nitric oxide as a means of signal transmission. In contrast, {alpha}-gustducin has been reported to be absent in rat duodenal enteroendocrine cells identified by chromogranin A (CgA) and 5-HT immunolabeling (14). However, it was reported that enteroendocrine cells equipped to release gastric inhibitory polypeptide, GLP-1, and cholecystokinin in the mouse small intestine do not express CgA (44) and that CgA is an incomplete marker of enteroendocrine cells in other species (2). Consequently, {alpha}-gustducin may be more widely expressed in mouse intestinal enteroendocrine cells than suggested. Because knockout mice studies are increasingly being used to dissect the signal elements of taste transduction, an accurate understanding of {alpha}-gustducin cell phenotype in mouse intestine is critical to interpreting potential signal role(s) of {alpha}-gustducin within the gastrointestinal tract.

The aim of these studies therefore was to establish a longitudinal and radial expression profile of {alpha}-gustducin throughout the mouse small intestine and to assess the phenotype of chemosensory cells that may use {alpha}-gustducin signaling pathways to detect nutrient. This was done by assessing colocalization of {alpha}-gustducin in 5-HT- and GLP-1-expressing enteroendocrine cells of the small intestine by dual-label immunohistochemistry. An assessment of {alpha}-gustducin cell phenotype was also extended to brush cells, which were identified within the small intestine by binding of the plant lectin Ulex europeaus agglutinin-1 (UEA-1), and by immunolabeling for neuronal nitric oxide synthase (nNOS).


   MATERIALS AND METHODS
TOP
 ABSTRACT
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Animal preparation. Experiments were performed on adult male C57BL/6 mice aged 7–10 wk. All studies were performed in accordance with the Australian code of practice for the care and use of animals for scientific purposes and with the approval of the Animal Ethics Committees of the Institute of Medical and Veterinary Science (Adelaide, Australia) and the University of Adelaide. Mice had free access to water and a standard rodent diet.

Mice were anesthetized with pentobarbitone (60 mg/kg ip) and perfused transcardially with heparinized saline at 40°C, then 4% paraformaldehyde in 0.1 M phosphate-buffered saline (PBS), pH 7.4 at 4°C, and the tongue and entire small intestine were removed. The small intestine was cut longitudinally along the cephalocaudal axis, pinned, flushed with fresh fixative and fixed for 2–4 h in 4% paraformaldehyde-PBS at room temperature (RT). Tissues were then washed in PBS, cryoprotected in 30% sucrose at 4°C overnight, then embedded in cryomolds and rapidly frozen in liquid nitrogen. Frozen transverse sections at 14 µm were then cut serially from the tongue, and sections were cut from the small intestine perpendicular to the longitudinal axis on a cryostat (CRYOCUT 1800 Reichert-Jung) and thaw mounted directly onto slides coated with gelatin and chrome-alum. Intestinal segments were defined by distance from the pylorus and in reference to the ligament of Treitz, with duodenum (0–4 cm postpylorus), proximal jejunum (4–11 cm), midjejunum (11–17 cm), distal jejunum (17–23 cm), and ileum (23–27 cm).

Immunohistochemistry. Immunolabel for {alpha}-gustducin was antibody detected by a rabbit anti-rat {alpha}-gustducin polyclonal primary antibody raised against COOH-terminal sequence YVNPRSREDQQLLLS [working dilution 1:500 (38)]. 5-HT immunoreactivity was detected by a monoclonal mouse antibody (M0758, 1:100, DakoCytomation, Glopstrup, Denmark), GLP-1 by a goat polyclonal antibody (SC7782, 1:100, Santa Cruz Biotechnology), and nNOS by alternate rabbit polyclonal antibodies (AB1552, 1:200, Chemicon-Millipore, Boronia, Australia) and (61-7000, 1:500, Zymed Laboratories). Primary antibodies were visualized by species-specific Alexa Fluor conjugated secondary antibodies (1:200, Invitrogen, Mount Waverley, Australia).

Frozen sections were air dried at RT for 15 min and then rinsed in PBST (PBS+ 0.2% Triton X-100, Sigma-Aldrich, PBS-T, pH 7.4) to facilitate antibody penetration. Sections were incubated with blocking solution (2% normal goat serum, 1% BSA, 0.1% Triton X-100, 0.05% Tween 20, 0.1% gelatin, 1x PBS) for 1 h at RT then {alpha}-gustducin primary antibody was diluted in blocking solution and added to sections then incubated overnight at 4°C. Sections were then washed in PBS-T and subsequently incubated with the relevant secondary antibody (1:200 in PBS-T) for 1 h at RT. Sections were finally washed in PBS-T and mounted in ProLong Antifade reagent (Invitrogen), coverslipped, and dried to visualize single {alpha}-gustducin immunolabel or underwent sequential incubation with 5-HT or GLP-1 primary antibody and corresponding secondary antibody, as detailed above for dual-label immunohistochemistry. Tongue sections were included as positive controls for {alpha}-gustducin specificity, and sections preadsorbed with the immunizing agent, or lacking the primary antibody, served as negative controls and were devoid of labeling. Immunohistochemistry for nNOS was performed only as a single-label assay due to an absence of nNOS labeling in intestinal epithelium (see RESULTS).

Lectin labeling. Brush cells were identified by UEA-1 binding, a surrogate marker enriched in this cell type compared with enterocytes (9) and superior to cytokeratin-18 as a brush cell marker in mice (13). Labeling for UEA-1 was detected by biotinylated UEA-1 (B-1065, 1:500, Vector Laboratories, Burlingame, CA). Sections were dried, then washed in PBS-T and UEA-1 incubated on sections for 2–3 h at RT. Sections were then washed in PBS-T and biotinylated UEA-1 visualized by incubation of streptavidin Alexa Fluor 350 conjugate (S11249, 1:200, Invitrogen) for 1 h at RT before PBS washing. Sections then underwent immunolabeling for {alpha}-gustducin or dual labeling as described above.

Visualization and quantification. Epifluorescent images were obtained on a epifluorescence microscope (BX-51, Olympus) equipped with multiple excitation filters and images acquired on a monochrome charge-coupled device digital camera system (Photometrics CoolSNAPfx, Roper Scientific, Tucson, AZ). Fluorescence images were then imported unmodified into V++ Precision Digital Imaging System software (version 4.0, Digital Optics, Auckland, New Zealand), pseudo-colored, and merged for composite images; luminance intensity was not adjusted.

Epithelial cells immunopositive for {alpha}-gustducin were manually counted and averaged over a minimum of 10 intact transverse sections representative of each intestinal segment (duodenum, proximal, mid and distal jejunum, and ileum) in six mice. In dual-labeling assays {alpha}-gustducin-immunopositive cells were initially scored in intestinal segments and the section was then visualized under the matched filter for 5-HT, GLP-1, or UEA-1 detection to assess the degree of colabel. Epithelial cells colabeled for {alpha}-gustducin, 5-HT, GLP-1, and UEA-1 were represented as a proportion of {alpha}-gustducin-immunopositive cells averaged from 10 sections per intestinal segment and pooled from a minimum of four mice per gene target.

Differences in the number of {alpha}-gustducin-labeled epithelial cells in small intestinal regions were assessed by one-way ANOVA with Tukey's multiple comparison tests (Prism 3.02; Graphpad, San Diego, CA). A P value of <0.05 was considered significant, and results are expressed as means ± SE of the number of animals (n).


   RESULTS
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 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 GRANTS
 REFERENCES
 
Distribution of {alpha}-gustducin-immunopositive cells is region specific. Immunolabeling with a primary antibody directed against {alpha}-gustducin identified a specific population of epithelial cells in the mouse small intestine, as shown in Fig. 1. {alpha}-Gustducin-immunopositive cells occurred singularly and were dispersed throughout the surface epithelium of intestinal villi. Immunolabeled cells were generally columnar in shape and showed a homogenous label within the cytoplasm and a gradient to peak labeling at the apical tip. All immunopositive cells were of open cell type (possessing an apical tip that extended to, or beyond, the brush border membrane and had access to the lumen) and were located on the mid to upper portion of the villus, frequently at, or near, the villus tip. On rare occasions (1–2 cells/mouse), {alpha}-gustducin cells could be located within glandular epithelium within or near crypts in distal portions of the intestine (ileum).


Figure 1
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  		Fig. 1. {alpha}-Gustducin expression in the mouse small intestine. The specificity of {alpha}-gustducin labeling was confirmed in fungiform taste cells in the mouse tongue, which were labeled by {alpha}-gustducin antibody (A). {alpha}-Gustducin strongly labeled single cells in the villous epithelium (B and C). Immunopositive cells were more commonly observed near the upper villus region (B, proximal jejunum) or along the midvillus (C, distal jejunum). The midjejunum showed an increased frequency of {alpha}-gustducin-immunopositive epithelial cells; 3 {alpha}-gustducin cells are seen in close proximity (D, midjejunum). Scale bars = 50 µm.

 
{alpha}-Gustducin-immunopositive cells showed a distinct regional distribution pattern, shown in Fig. 2. Immunopositive cells were rare within the duodenum (0.3 ± 0.1 cells/section profile, range 0–2 cells/section profile), first became apparent within the proximal jejunum (1.4 ± 0.1 cells, range 0–5 cells), and reached a region of peak density in the midjejunum (7.4 ± 0.4 cells, range 0–25 cells). Immunopositive cells then decreased in density toward the distal jejunum (2.0 ± 0.2 cells, 0–9 cells) and ileum (2.7 ± 0.2 cells, range 0–10 cells) but were detected up to the ileocecal junction and in the colon (data not shown). The midjejunum contained the highest number of {alpha}-gustducin immunopositive epithelial cells (P < 0.05 compared with other regions).


Figure 2
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  		Fig. 2. The number of {alpha}-gustducin-immunopositive cells varies with region of small intestine. Mean {alpha}-gustducin cell count per section is shown for each small intestinal region. {alpha}-Gustducin-immunopositive cells were preferentially located in the midjejunum. *Significantly different from other intestinal regions (1-way ANOVA; P < 0.001). #Significantly different from duodenum (P < 0.05). ^Significantly different from proximal jejunum (P < 0.01).

 
5-HT and GLP-1 are expressed in different cells in the epithelium. Immunolabeling for 5-HT or GLP-1 was evident in single cells dispersed in villus epithelium and in crypts and glands throughout the mouse small intestine. Dual immunohistochemistry and cell counts indicated that 0% of epithelial cells colabeled 5-HT and GLP-1 in any region of the small intestine, as shown in Fig. 3. 5-HT-immunolabeled cells were more numerous than GLP-1 cells in the upper and midintestine. In all regions 5-HT or GLP-1 cells showed respective immunolabeling throughout the cytoplasm, often with strongest labeling in the basolateral portion of the cell, as shown in Figs. 3 and 4.


Figure 3
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  		Fig. 3. Coexpression of {alpha}-gustducin with 5-HT in mouse midjejunum. A proportion of {alpha}-gustducin-immunopositive cells in midjejunum (red fluorescence; A and D) colabeled with 5-HT (green fluorescence; B and E). A composite image (yellow; C and F) confirms localization within the same cell. A 5-HT-positive cell not colabeled with {alpha}-gustducin is shown in close proximity to the dual-labeled cell (C). The majority of {alpha}-gustducin-immunopositive cells (G), however, were immunonegative for 5-HT (H and I). The proportion of {alpha}-gustducin+5-HT colabeled cells in midjejunum was 27 ± 2% of total {alpha}-gustducin cells (J). Scale bars = 50 µm.

 

Figure 4
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  		Fig. 4. Coexpression of {alpha}-gustducin with glucagon-like peptide-1 (GLP-1) in mouse midjejunum. A proportion of {alpha}-gustducin-immunopositive cells in midjejunum (red fluorescence; A and D) colabeled with GLP-1 (green fluorescence; B and E). A composite image (yellow; C and F) confirms localization within the same cell. Labeling patterns of the two antibodies often appeared polar with {alpha}-gustducin strongest in the apical regions of the cell and GLP-1 strongest in the basolateral portion. The majority of {alpha}-gustducin-immunopositive cells (G), however, were immunonegative for GLP-1 (H and I). The proportion of {alpha}-gustducin+GLP-1 colabeled cells in midjejunum was 15 ± 2% of total {alpha}-gustducin cells (J). Scale bars = 50 µm.

 
Subsets of {alpha}-gustducin-immunopositive epithelial cells in the midjejunum contain 5-HT or GLP-1. In the midjejunum a population of epithelial cells that colabeled for {alpha}-gustducin+5-HT (Fig. 3) and another for {alpha}-gustducin+GLP-1 were identified (Fig. 4). Cell counts showed that 27 ± 2% of midjejunal {alpha}-gustducin cells contained 5-HT, whereas 15 ± 2% contained GLP-1. No {alpha}-gustducin-labeled cells in the midjejunum triple labeled ({alpha}-gustducin+5-HT+GLP-1). Conversely, of the total number of 5-HT labeled cells within the midjejunum 25 ± 9% labeled with {alpha}-gustducin, whereas 8 ± 2% of total GLP-1 cells colabeled with {alpha}-gustducin. In all other regions of the small intestine {alpha}-gustducin did not colabel with either enteroendocrine cell marker, indicating that {alpha}-gustducin signaling via 5-HT and GLP-1 is likely to occur only within the midjejunum in mice.

{alpha}-Gustducin is expressed in different small intestinal cell populations. Dual-labeled {alpha}-gustducin+5-HT and {alpha}-gustducin+ GLP-1 epithelial cells consistently displayed a distinct morphology compared with single-labeled {alpha}-gustducin cells. Dual-labeled {alpha}-gustducin cells were largely spindle shaped, did not possess an apical process that extended beyond the brush-border membrane, showed punctate labeling for {alpha}-gustducin within the cytoplasm (Fig. 5), and were consistently negative for the lectin UEA-1 (see GoFig. 7D). In contrast, single-labeled {alpha}-gustducin cells (i.e., immunonegative for 5-HT or GLP-1) were mainly columnar in shape, possessed an apical process that extended beyond the brush-border membrane, showed homogenate labeling within the cytoplasm for {alpha}-gustducin, and were largely UEA-1 positive throughout the small intestine, suggesting that they were brush cells (Fig. 5).


Figure 5
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  		Fig. 5. {alpha}-Gustducin epithelial cell types exhibit different morphology and immunolabeling patterns in mouse small intestine. Immunopositive cells colabeled with {alpha}-gustducin and 5-HT or GLP-1 in midjejunum showed a different morphology and {alpha}-gustducin labeling pattern (arrowhead) compared with those that were not colabeled (arrow). Scale bar = 50 µm.

 

Figure 6
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  		Fig. 6. Coexpression of {alpha}-gustducin with lectin Ulex europeaus agglutinin 1 (UEA-1) in mouse small intestine. The majority of {alpha}-gustducin-immunopositive cells throughout the mouse small intestine did not colabel with GLP-1 or 5-HT but bound the lectin UEA-1, a brush cell marker in mouse intestine (blue fluorescence, B and C). The proportion of {alpha}-gustducin+UEA-1 colabeled cells in the midjejunum from (n = 2) mice was 57% of total {alpha}-gustducin cells (D). Not all UEA-1 labeled cells expressed {alpha}-gustducin (arrowhead).

 

Figure 7
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  		Fig. 7. {alpha}-Gustducin is expressed in different populations of epithelial cell in mouse small intestine. The majority of {alpha}-gustducin-immunopositive cells throughout the mouse small intestine did not colabel with GLP-1 or 5-HT but bound the lectin UEA-1 (blue fluorescence, arrows; C and D). A minority of {alpha}-gustducin-immunopositive cells colabeled with 5-HT (green fluorescence, arrowhead; D). Although rare 5-HT-expressing cells colabeled with UEA-1 (inset in B), cells triple labeled for these markers ({alpha}-gustducin+ 5-HT+UEA-1) were not found (D). Scale bars = 50 µm.

 
UEA-1 labeling was widespread in mouse small intestine with single epithelial cells seen labeled at the glycocalyx and apex and often within the cytoplasm, as shown in Fig. 6. UEA-1 cells occasionally colabeled with 5-HT, although no triple labeled cells ({alpha}-gustducin+UEA-1+5-HT) were seen (Fig. 7). In the midjejunum 57% of {alpha}-gustducin cells colabeled with UEA-1, fewer than in other intestinal regions. However, even here {alpha}-gustducin cells represented only a small proportion of the total UEA-1 epithelial cell population, with cell counts showing that 17 ± 2% of UEA-1 cells colabeled with {alpha}-gustducin.

Mouse small intestinal epithelium is immunonegative for nNOS. Immunolabeling for nNOS was performed to further classify the brush cell phenotype of {alpha}-gustducin cells colabeled by UEA-1. Intense labeling for nNOS was observed within neural cell bodies and fibers in the myenteric plexus of the small intestine with both antibodies, confirming the specificity of labeling, as shown in Fig. 8. Neither nNOS antibody labeled the intestinal epithelium, indicating that epithelial cells of the mouse small intestine do not express significant amounts of nNOS. As a consequence, dual immunolabeling for {alpha}-gustducin+nNOS was not performed in these studies.


Figure 8
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  		Fig. 8. Neuronal nitric oxide synthase (nNOS) expression in the mouse small intestine. nNOS immunoreactivity was localized to the myenteric plexus of the mouse small intestine but was absent from the epithelium (A). nNOS immunoreactivity was absent from negative control sections (inset in A). B: higher power image of nNOS immunoreactivity in the myenteric plexus of the midjejunum. Scale bars = 50 µm (A, inset) and 10 µm (B).

 

   DISCUSSION
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
GRANTS
 REFERENCES
 
Our data provide the first evidence that the taste-specific G protein {alpha}-gustducin is expressed in distinct epithelial cell populations of the mouse small intestine. We have further demonstrated that these fall into three distinct groups: enteroendocrine cells that contain 1) 5-HT or 2) GLP-1, localized to the midjejunum, or 3) brush cells that are more generally distributed and bind the lectin UEA-1. Because these cell types are involved in initiating gastrointestinal reflexes and behavioral responses to luminal nutrients (41, 48), our observations suggest that {alpha}-gustducin signaling pathways may potentially provide a selective therapeutic target in disordered gastrointestinal function and nutrient detection.

We mapped immunolabeling for {alpha}-gustducin throughout the mouse small intestine and assessed the phenotype of {alpha}-gustducin immunopositive epithelial cells. {alpha}-Gustducin was confined to solitary epithelial cells scattered throughout the upper villus epithelium, and preferentially in those located within the midjejunum. Outside the midjejunum, {alpha}-gustducin cells were less common within the duodenal epithelium and were detected at low to moderate levels in the proximal jejunum, distal jejunum, and ileum). Our findings are in agreement with previous {alpha}-gustducin expression reported in rodent small intestine by RT-PCR and Western blotting (5). In addition, we provide the first details of {alpha}-gustducin immunolabeling in single intestinal epithelial cells in mice and their location at key sites in the upper gastrointestinal tract.

Immunolabeling for {alpha}-gustducin was prominent in the midjejunal epithelium, whereas relatively few chemosensory cells of the proximal small intestine expressed {alpha}-gustducin in mouse. This appears to differ from the situation in human gastrointestinal tract, where {alpha}-gustducin immunolabeling has been described in duodenal and colonic mucosa (37), although the relative distribution is not known. This may suggest that intestinal {alpha}-gustducin-dependent signaling shows distinct species differences.

A key finding in this study was that the majority of {alpha}-gustducin cells along the longitudinal axis of the small intestine colabeled with the plant lectin UEA-1, particularly at the glycocalyx and apex. UEA-1 has been validated as a selective marker for brush cells in the mouse small intestine at both confocal light and electron microscopic levels (7, 9), although UEA-1 does colabel a subset of enteroendocrine cells in the mouse cecum that contain 5-HT or peptide YY (PYY) (8). In the present study we found only rare colabel between 5-HT and UEA-1 in epithelial cells, but importantly in the context of {alpha}-gustducin signaling no triple-labeled UEA-1+5-HT+{alpha}-gustducin cells were seen. This finding indicates that UEA-1+5-HT cells do not signal nutrient via {alpha}-gustducin. Conversely, the dual-labeled {alpha}-gustducin+UEA-1 cells identified represent a distinct subset of mouse intestinal brush cells that may use {alpha}-gustducin pathways to signal intestinal nutrient. Brush cells in the intestine have been proposed to have chemosensory functions because they possess a prominent glycocalyx, an anatomical feature shared by chemosensory epithelial cells in other sensory organs, including the nasal cavity and tongue (43, 45). In the rat stomach and pancreas, brush cells have been shown to express high levels of nitric oxide synthase by immunolabeling (18) but lack neural synapses and secretory granules. As a consequence, nitric oxide has been proposed as a gaseous molecule for signaling to afferent nerves or other intermediary cell types within the epithelium (12). In the present study we did not detect nNOS by immunolabeling in mouse epithelium, despite strong nNOS immunoreactivity in the myenteric plexus. It appears therefore that, unlike brush cells of the rat stomach, brush cells in mouse small intestine are not equipped to synthesize nitric oxide for release by established mechanisms and that other signaling mechanism(s) may be responsible for transmission of chemosensory signals in this cell type.

Intestinal enteroendocrine cells represent less than 1% of intestinal epithelial cells but collectively form the largest endocrine organ of the body, releasing more than 14 known hormones (32) that have key gastrointestinal and systemic roles. Of these, enterochromaffin cells constitute the largest enteroendocrine cell type and are equipped to release 5-HT in vitro (17) and in vivo (reviewed in Ref. 29). We confirmed a large population of 5-HT containing cells in the mouse small intestine in the present study. There is persuasive evidence that 5-HT has a role in gastrointestinal reflex responses to nutrients in that 5-HT released from enteroendocrine cells in response to carbohydrate can activate vagal afferent endings in the rat intestine (48). The 5-HT3 receptor antagonist tropisetron has also been shown to abolish the inhibition of gastric emptying in rats induced by small intestinal glucose (30). Other studies have shown that 5-HT3 receptor-immunopositive nerve fibers run within duodenal villi in rats (10), and retrograde tracing from this site has confirmed that vagal afferents innervating this region express 5-HT3 receptors (30). It emerges from our data that 5-HT may be released via both {alpha}-gustducin-dependent and -independent pathways. It also emerges that its role as a "taste" mediator is probably confined to the midjejunum in mice.

The incretin hormone GLP-1 modulates intestinal nutrient-evoked reflexes. GLP-1 is expressed in L cells throughout the intestine, but predominantly in the ileum and colon, and is released postprandially in response to luminal carbohydrate when it acts primarily to inhibit gastrointestinal motor function and to delay gastric emptying (15). These effects are mediated by capsaicin-sensitive vagal afferents (16). Indeed, GLP-1 receptors have recently been identified in the rat nodose ganglion (28), indicating that vagal afferent terminals in the periphery receive chemosensory information via peripheral GLP-1 receptors. However, a central site of action of GLP-1 on gastric emptying has also been shown (16, 27), and GLP-1 and GLP-1 receptor expression have been reported in the rat brain, notably within the dorsal vagal complex (11, 23). GLP-1 also acts centrally to inhibit gastric acid secretion (16), whereas both central and peripheral sites of action have been described for suppression of energy intake by GLP-1 (34) and stress-induced colonic motility responses (26). A central humoral role for endogenous GLP-1 is less likely, however, because of the rapid degradation of GLP-1 that occurs at the site of peripheral release and within the liver and circulation (3). These diverse actions of GLP-1 highlight the complex roles of GLP-1 in nutrient-evoked gastrointestinal reflexes. Importantly, our {alpha}-gustducin colabeling results indicate that, like 5-HT, at least two different mechanisms are likely to be involved in the triggering of GLP-1 release from the midjejunal epithelium, one involving {alpha}-gustducin and others as yet unidentified.

Our observation of separate 5-HT or GLP-1 enteroendocrine cell populations support earlier findings that 5-HT and GLP-1 cells are distinct cell lineages in mouse intestine (35). However, it is likely that other mediators may be released via {alpha}-gustducin signaling in these enteroendocrine cells. We found that 99% of {alpha}-gustducin cells in the midjejunum were labeled by 5-HT, GLP-1, or UEA-1 markers. This suggests that if other enteroendocrine mediators are released by {alpha}-gustducin signals they are likely to be coexpressed in the {alpha}-gustducin+5-HT or {alpha}-gustducin+GLP-1 cells we identified. Dual GLP-1+PYY cell phenotypes have recently been described in the porcine, rat, and human intestine (24). Moreover, there is potential for corelease of substance P from {alpha}-gustducin+5-HT cells and corelease of PYY, cholecystokinin, or neurotensin from {alpha}-gustducin+GLP-1 cells (35), in response to intestinal nutrient.

Despite clear evidence for roles of both 5-HT and GLP-1 in nutrient signaling and paracrine activation of vagal afferents, our observations suggest that a minority of enteroendocrine cells use {alpha}-gustducin signaling and are equipped to release 5-HT, GLP-1, or other mediators. Accordingly, we speculate that these {alpha}-gustducin+5-HT and {alpha}-gustducin+GLP-1 cell subpopulations subserve specific nutrient-evoked gastrointestinal reflexes. In contrast, the proportionally larger {alpha}-gustducin+UEA-1 brush cell population identified is consistent with the concept that small intestinal brush cells contribute significantly to nutrient-evoked signaling via {alpha}-gustducin and release of a nonnitrergic mediator. It is also important to note that {alpha}-gustducin signal pathways also subserve nonnutrient (bitter taste) detection; {alpha}-gustducin-equipped enteroendocrine or brush cells identified in this study may equally signal the presence of bitter nutrients, drugs, or toxins in the small intestine (reviewed in Ref. 42).

Interestingly, respiratory tract brush cells in humans are infrequent compared with those in rodents (31). If this species difference is reflected in gastrointestinal brush cell populations, then {alpha}-gustducin-dependent signaling from this cell type may be underrepresented in humans. Alternatively, {alpha}-gustducin may be expressed in different intestinal cell populations in humans. Indeed, {alpha}-gustducin immunolabeling has recently been shown in human colonic epithelial cells that colabel with PYY and GLP-1 (37), the first report of a human intestinal cell type that may use {alpha}-gustducin-dependent signaling. However, no report has directly compared brush cell populations in human and mouse gastrointestinal tract, and specifically, no details have yet emerged on the phenotype of {alpha}-gustducin cells in human proximal small intestine, a potentially more relevant site for initiating nutrient-based motility reflexes that modify gastric emptying (40).

In conclusion, this study provides the first direct evidence of small intestinal enteroendocrine and brush cell subpopulations expressing {alpha}-gustducin and their potential paracrine neurotransmitters in mice. The identification of these distinct {alpha}-gustducin cell subpopulations has substantial implications for understanding the nature of small intestinal nutrient signaling. It may be possible to exploit tastant-induced release of gastrointestinal neurotransmitters or novel drugs that target the {alpha}-gustducin signal pathway to alter nutrient-evoked signaling in the small intestine. Such strategies would potentially provide a novel approach to therapy in patients with upper gastrointestinal motility disorders.


   GRANTS
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
REFERENCES
 
This work was supported by The University of Adelaide.


   ACKNOWLEDGMENTS
 
We thank Professor Robert F. Margolskee and Zaza Kokrashvili (Department of Neuroscience, Mount Sinai School of Medicine, New York, NY) for valuable comments and provision of mouse {alpha}-gustducin antibody.


   FOOTNOTES
 

Address for reprint requests and other correspondence: R. L. Young, Nerve-Gut Research Laboratory, Level 1 Hanson Institute, Frome Rd., Adelaide SA 5000, Australia (e-mail: richard.young{at}adelaide.edu.au)

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.

* K. Sutherland and R. L. Young contributed equally to this work. Back


   REFERENCES
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 

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