Chemical Senses Advance Access originally published online on January 17, 2007
Chemical Senses 2007 32(3):255-262; doi:10.1093/chemse/bjl053
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Expression of T1Rs and Gustducin in Palatal Taste Buds of Mice
1 Rocky Mountain Taste and Smell Center, Aurora, CO 80045, USA 2 Department of Biomedical Sciences, Colorado State University, Fort Collins, CO 80523, USA 3 Department of Cell and Developmental Biology University of Colorado School of Medicine, Aurora, CO 80045, USA
Correspondence to be sent to: Leslie M. Stone, Department of Biomedical Sciences, Colorado State University, Fort Collins, Colorado, USA. e-mail: leslie.stone-roy{at}colostate.edu
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Abstract |
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The palatal region of the oral cavity in rodents houses 100–300 taste buds and is particularly sensitive to sweet and umami compounds; yet, few studies have examined the expression patterns of transduction-related molecules in this taste field. We investigated the interrelationships between members of the T1R family and between each T1R and gustducin in palatal taste buds. Similar to lingual taste buds, T1R1 and T1R2 are generally expressed in separate palatal taste cells. In contrast to lingual taste buds, however, T1R2 and T1R3-positive palatal taste cells almost always coexpress gustducin, suggesting that sweet taste transduction in the palate is almost entirely dependent on gustducin. T1R1-positive palate taste cells coexpress gustducin about half the time, suggesting that other G proteins may contribute to the transduction of umami stimuli in this taste field.
Key words: coexpression, gustatory, immunocytochemistry, taste transduction, umami
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Introduction |
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Taste buds, the transducing elements of gustatory sensation, are distributed throughout the oral cavity including the palate as well as the tongue. Lingual taste buds lie in 3 different types of connective tissue papillae residing in different areas of the tongue. Fungiform papillae, small, raised papillae on the anterior two-thirds of the tongue, each contain one or two taste buds that are innervated by the chorda tympani branch of the facial nerve. Vallate and foliate papillae are invaginations on the posterior and lateral surfaces of the tongue. These papillae contain hundreds of taste buds that are innervated primarily by the glossopharyngeal nerve. The taste buds of the palate are most plentiful in 3 locations: near and within the nasoincisor ducts, in a discrete Geschmacksstreifen, a stripe of taste buds located at the juncture of the soft and hard palate, and scattered within the soft palate just posterior to the Geschmacksstreifen. Palatal taste buds are innervated by the greater superficial petrosal branch of the facial nerve.
The chemical sensitivity of taste buds varies according to their location in the oral cavity (Frank et al. 1983
). The greater superficial petrosal branch of the facial nerve, which innervates taste buds on the palate, is highly responsive to sweet and umami taste stimuli (Krimm et al. 1987; Travers and Norgren 1991; Harada et al. 1997; Sako et al. 2000; Sollars and Hill 2005). In contrast, the glossopharyngeal nerve responds best to acidic and bitter stimuli (Danilova and Hellekant 2003), whereas the chorda tympani nerve responds to salty and acidic tastants and to a lesser extent to sweet and umami compounds (Frank et al. 1983; Sako et al. 2000; Danilova and Hellekant 2003).
Bitter, sweet, and umami taste stimuli are detected by G protein–coupled receptors, leading to activation of intracellular signaling cascades and transmission of taste information to associated nerve fibers. The T2Rs represent a family of about 30 receptors, which are activated by bitter compounds (Adler et al. 2000; Chandrashekar et al. 2000; Matsunami et al. 2000). Consistent with taste field specificity, T2Rs are more prevalent in the bitter-sensitive vallate papilla (Adler et al. 2000), although T2R expression in the palatal Geschmacksstreifen is similar to vallate and foliate indicating that the palate is also important for bitter taste transduction (Adler et al. 2000). Sweet and umami transduction are mediated through T1R receptors, a family consisting of 3 members, which combine to form primarily heteromeric receptors. T1R1 + T1R3 form an amino acid (umami) receptor, whereas T1R2 + T1R3 form a broadly tuned sweet receptor. The distributions of the T1Rs vary with location. T1R3 is expressed in all lingual papillae, as well as in taste buds of the palate (Kitagawa et al. 2001; Max et al. 2001; Montmayeur et al. 2001; Nelson et al. 2001; Sainz et al. 2001). In contrast, some reports indicate that T1R1 is expressed primarily in fungiform taste buds, whereas T1R2 is expressed primarily in vallate and foliate papillae (Hoon et al. 1999), but see Kim et al. (2003) who report more widespread expression of T1R1 and T1R2. The palate is the only taste field where both T1R1 and T1R2 are abundantly expressed, and it is also an area responsive to sweet and umami stimuli.
In addition to receptor proteins, downstream signaling effectors play important roles in taste transduction. One of these effectors is the G protein 
-gustducin, which is present in some taste cells in all gustatory fields (Boughter et al. 1997). Mice lacking the -gustducin protein are defective in their ability to detect bitter, sweet, and umami tastants (Wong, Gannon, et al. 1996; Wong, Ruiz-Avila, et al. 1996; Ruiz-Avila et al. 2001; Caicedo et al. 2003; Ruiz et al. 2003; Glendinning et al. 2005), suggesting that gustducin impacts on both the T2R and T1R transduction pathways. T2Rs and gustducin are coexpressed in many taste cells of the bitter-sensitive vallate region as expected. In contrast, T1Rs and gustducin are rarely coexpressed in this region, although some coexpression of T1Rs and gustducin was recently reported in fungiform taste buds (Kim et al. 2003). Palatal taste buds have not been examined for coexpression of gustducin and T1Rs, despite being the most sweet-sensitive region of the oral cavity.
To better understand sweet and umami taste transduction, the current study examines the expression patterns of each member of the T1R family and the relationship of each T1R with gustducin in palate taste buds. In addition, to facilitate comparisons between palatal and lingual taste buds, and because there is controversy in the literature, we reexamined the expression of T1Rs and gustducin in fungiform and circumvallate taste buds. A portion of these data was presented previously in abstract form (Stone et al. 2003).
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Animals
The tongues and palates from adult male and female C57/BL mice were used for these studies. Animals were housed and handled according to Animal Care and Use Committees at Colorado State University and the University of Colorado Health Sciences Center. For some experiments, mice were anesthetized with an overdose of sodium pentobarbital (or chloral hydrate) injection (55 mg/ml) and perfused transcardially with 4% paraformaldehyde (PFA) in 0.1 M phosphate buffer. For the remaining experiments, animals were first euthanized with carbon dioxide, followed by cervical dislocation. Tongues and palates were removed from the mice and fixed by immersion in 4% PFA.
In situ hybridization
In situ hybridization was carried out as previously described (Finger et al. 2003). Briefly, sense and antisense riboprobes were synthesized from cDNA plasmids for mouse T1R1 and T1R2 (gifts from C. Zuker, University of California at San Diego and N. Ryba, National Institutes of Health) incorporating digoxigenin or fluorescein-labeled uridine triphosphate (UTP) and unlabeled UTP and hydrolyzed to 
500 bp. Tongues and palates from perfused mice were postfixed in PFA with 20% sucrose overnight. Frozen cryostat sections (12–14 microns) were collected onto Fisher Superfrost Plus slides maintained at room temperature and stored at –20 °C until ready for hybridization. Sections were treated with proteinase K (20 µg/ml), washed, and treated with 0.5% H2O2 to quench endogenous peroxidase activity. After being incubated at 37–45 °C in prehybridization buffer for 30–60 min, the slides were incubated at 59 °C overnight in hybridization solution (same as prehybridization buffer with the addition of 5% dextran sulfate and 40–200 ng/ml of digoxigenin or fluorescein-labeled probe). Digoxigenin label was detected using biotin-labeled mouse antidigoxin antibody (Jackson Immunoresearch, West Grove, PA) diluted 1:200. After washing, avidin–biotin complex (Vector Laboratories, Burlingame, CA) was applied to the slides, followed by Alexa Fluor 568 tyramide (TSA, Molecular Probes, Eugene, OR) and reacted with 0.0015% H2O2. The slides then were washed, blocked with 1% normal serum (Jackson), and incubated overnight with rabbit anti-gustducin (Santa Cruz Biotechnology, Santa Cruz, CA) at 1:1000 or rabbit anti-T1R3 (gift from Robert Margolskee) at 1:500. The gustducin and T1R3 antisera were detected with Alexa Fluor 488 goat anti-rabbit antibody (Molecular Probes). The slides were washed and coverslipped with Fluoromount-G. Controls in which the probe or the primary antiserum was omitted showed no cross-reactivity.
For double in situ hybridization labeling, one probe was labeled with digoxigenin, the other with fluorescein. The tissue was prepared in the same manner as the single in situ labeled tissue except that the fluorescein-labeled probe was detected with rabbit anti-fluorescein (Zymed Laboratories, San Francisco, CA) before the digoxigenin-labeled probe was detected. The fluorescein antibody was detected using biotin donkey anti-rabbit (Jackson), avidin–biotin complex, and Alexa Fluor 488 TSA. The sections were then retreated with 0.5% H2O2 and blocked with avidin–biotin blocking reagent (Vector), and the digoxigenin-detecting procedure described above was performed for detection of the second probe. Controls in which one of the two probes was omitted showed no cross-reactivity.
Immunocytochemistry
Tissue was collected and postfixed for 1–2 h in 4% PFA. Both immersion-fixed and perfused tissues were transferred into 20% sucrose in 0.1 M phosphate buffer and put at 4 °C overnight. Next, tissue was trimmed and frozen in O.C.T. Compound (Electron Microscopy Sciences, Fort Washington, PA). Fifteen-micron cryosections were collected on Superfrost Plus slides maintained at room temperature (VWR, West Chester, PA) and washed in 0.1 M phosphate-buffered saline (PBS).
To evaluate the relationship between T1R3 and gustducin expression, an immunocytochemical protocol similar to that developed by Shindler and Roth (1996) was used. This technique allows immunocytochemical evaluation of 2 antigens using antibodies raised in the same species. To prepare taste tissue for immunolabeling, endogenous peroxidases were inactivated by incubating tissue sections in 0.5% hydrogen peroxide in 0.1 M phosphate buffer for 15 min. This was followed by washes in 0.1 M PBS, and then the slides were incubated in blocking solution (0.3% triton X-100 [US Biochemical Corporation, Cleveland, OH], 1% bovine serum albumin, and 1% normal goat serum [Jackson Immunoresearch Laboratories] in 0.1 M PBS), for 1–2 h at room temperature. The sections then were incubated in rabbit antigustducin antibodies (Santa Cruz #sc-395; 1:20 000 to 1:30 000) overnight. Due to the dilute concentration of primary antibody, subsequent application of fluorescent anti-rabbit secondary antibodies using standard protocols produces no visible label. However, amplification protocols result in detectable labeling. Tyramide amplification of the signal was used for the current studies. For this protocol, first the sections were washed in 0.1 M PBS and incubated in biotinylated goat anti-rabbit antibodies (Jackson; 1:1000 in blocking solution) for 2–3 h. Next, the slides were washed in PBS and incubated with streptavidin–horseradish peroxidase (Molecular Probes, TSA kit, component C) for 1 h at room temperature. Following one 5-min wash in PBS, the slides were washed twice in wash buffer (0.1 M Tris, 150 mM NaCl, 0.1% Tween20). Tyramide working solution was then applied to the slides. This consisted of the labeled tyramide (with Alexa Fluor 488 or Alexa Fluor 647, Molecular Probes kit component A) dissolved in amplification buffer (kit component E) with 0.0015% hydrogen peroxide (made from kit component F). The labeled tyramide solution was left on the slide for 5–10 min, then rinsed off with PBS plus 0.1% tween (3 washes of 15 min each), resulting in tyramide precipitate indicative of gustducin. Labeling for the second protein was done according to standard indirect immunofluorescence procedures. The tyramide-labeled sections were incubated in blocking solution for 1 h, and the second primary antibody was added at a normal dilution (rabbit anti-T1R3 1:200 to 1:300). The second primary antibody was left on overnight, and then the slides were washed in PBS and the second secondary antibody applied (Cy5 [1:400] or fluorescein isothiocyanate [1:100] goat anti-rabbit). After 2 h, the slides were washed in PBS and coverslipped with fluoromount-G. For each experiment, 2 controls were done: one lacking primary antibodies and one omitting the first secondary antibody and the second primary antibody to insure that the second secondary antibody did not cross-react with the first primary antibody. The controls indicated that there was no cross-reactivity. Unless noted, chemicals were purchased from Sigma.
Imaging and analysis
Processed tissue was viewed with an Olympus Fluoview FV300 confocal microscope or a Zeiss LSM 510 meta confocal microscope. Z stacks of images from labeled taste buds were collected with each channel being acquired separately to avoid bleedthrough of signal from one channel to the other. Acquisition settings were kept constant for experimental samples and control samples. The individual images of each Z series were combined prior to counting T1R3-immunoreactive (IR) cells, gustducin-IR, and double-labeled cells. Only labeled cells with visible nuclei were included in the cell counts. Images were saved as tif files, and identifying labels were applied using Photoshop 6.0 or 7.0 software. If necessary, brightness and contrast were adjusted using Photoshop 6.0 or 7.0 software.
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The distribution of T1R family members and the relationships between specific T1Rs and between each T1R and
-gustducin in palate and lingual taste buds was examined using immunocytochemistry and in situ hybridization. The primary focus of the current study was the evaluation of palatal taste buds. Examination of lingual taste buds was done to allow comparisons between different taste cell types and because there are differing reports in the literature concerning the distributions of T1Rs in lingual taste buds. We found that all 3 T1R subunits are expressed in palate taste buds, as reported by Nelson et al. (2001). In addition, and in agreement with previous work, T1Rs also were expressed in taste buds from fungiform, vallate, and foliate papillae (Hoon et al. 1999; Kitagawa et al. 2001; Max et al. 2001; Montmayeur et al. 2001; Nelson et al. 2001; Sainz et al. 2001; Kim et al. 2003).
Of all palatal taste cells labeled by T1R1 or T1R2 probes, 40% expressed T1R1 only, 50% expressed T1R2 only, and 10% expressed both subunits (Figures 1 and 2A, Table 1). Similar to previous studies on lingual taste buds (Hoon et al. 1999), we found that about 10% of circumvallate and foliate taste cells coexpress T1R1 and T1R2. Thus, palatal taste buds are similar to taste buds in other regions in that T1R1 and T1R2 are generally expressed in different taste cells. Although the percentage of cells coexpressing both T1R subunits was similar in all taste fields, the proportions of cells expressing T1R1 only or T1R2 only differed in some regions. Palate and fungiform taste buds were similar to each other but differed from taste buds in vallate and foliate fields. Palate and fungiform taste buds expressed roughly equal numbers of T1R1 and T1R2 cells, whereas in circumvallate and foliate taste buds, T1R2 expression predominated, with T1R2-only cells representing over 70% of the T1R-expressing population (Figure 1, Table 1).
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Both T1R receptors and the G protein
-gustducin play roles in sweet and umami transduction (Wong et al. 1996; Nelson et al. 2001; Ruiz-Avila et al. 2001; Li et al. 2002; Caicedo et al. 2003; Damak et al. 2003; Ruiz et al. 2003; Zhao et al. 2003; Glendinning et al. 2005); yet, T1R receptors and gustducin are usually not coexpressed in the same taste cells in vallate or foliate papillae (Hoon et al. 1999; Kim et al. 2003). To test whether T1Rs and gustducin are coexpressed in taste buds of the sweet-sensitive palate, we examined the expression of each T1R relative to -gustducin in these taste buds. We found that T1Rs and gustducin were commonly expressed in the same cells in palate taste buds (Figure 2B,C and Table 2). The highest incidence of coexpression was with T1R3 and gustducin; 100% of gustducin-IR cells also expressed T1R3 (Figures 3 and 4); 38% of gustducin cells expressed T1R2, and 12% expressed T1R1 (Figure 5). In contrast to the palate, the foliate and vallate papillae contained a large percentage of gustducin-positive cells that did not express a member of the T1R family (Figures 3 and 5). Presumably the non-T1R gustducin-positive cells in these lingual fields express T2R receptors as reported elsewhere (Adler et al. 2000; Kim et al. 2003).
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Evaluation of the data in terms of the percentage of T1R cells coexpressing gustducin illustrates that in the palate, T1R3 and T1R2 cells almost always express gustducin and half of T1R1 cells express gustducin (Figure 6, Table 2). Specifically, 100% of T1R3+ palate cells and 91% of T1R2+ palate cells coexpressed gustducin with T1R1 cells coexpressing gustducin 50% of the time (Figure 6). Examination of fungiform taste buds revealed the expression of T1Rs relative to gustducin was similar to palate taste buds. However, fewer T1R2-expressing cells were present in the fungiform papillae (Figure 6). Taste buds located in the vallate and foliate papillae contained few cells expressing both a known T1R member and gustducin (Figures 2D and 6) as reported previously (Hoon et al. 1999
; Adler et al. 2000; Montmayeur et al. 2001; Kim et al. 2003).
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Discussion |
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The palate is the most sweet-sensitive region of the oral cavity and is also responsive to umami stimuli. Despite this, few studies have focused on palatal taste buds in terms of the expression patterns of molecules implicated in sweet and umami taste transduction. The principal finding in this study is that T1Rs are expressed only by gustducin-containing cells in the palate. This contrasts with circumvallate and foliate taste buds where these molecules are rarely found in the same cells and helps explain behavioral and physiological data that indicate that both T1Rs and gustducin are involved in the transduction of sweet and umami stimuli.
Our data also confirm previous studies showing largely independent expression of T1R1 and T1R2 in lingual taste buds. Previous reports indicated that T1R1 and T1R2 are not only expressed in different taste cells but also mostly expressed in different taste fields; T1R1 expression is prevalent in fungiform taste buds but rare in vallate and the reverse for T1R2 (Hoon et al. 1999; Adler et al. 2000; Nelson et al. 2001). However, a more recent report by Kim et al. (2003) suggests that the expression of T1R1 and T1R2 are more widespread and that T1R1 and T1R2 are coexpressed in some taste cells. In the current study, we find that T1R1 and T1R2 are expressed in vallate and fungiform taste fields, although we did not find T1R expression to be as widespread as that reported by Kim et al. (2003). Furthermore, similar to studies by Hoon, Adler, and Nelson, we found that the amino acid–sensitive subunit T1R1 and the sweet-sensitive subunit T1R2 generally occur in separate cells in both lingual and palatal taste buds.
This is interesting in light of data from single fiber recordings, which show that gustatory afferents that respond best to sucrose often also respond to umami stimuli (Formaker et al. 2004). Also, behavioral data suggest mice have difficulty distinguishing sucrose from monosodium glutamate (MSG) when amiloride is added to block the sodium component of MSG taste (Yamamoto et al. 1991; Chaudhari et al. 1996; Stapleton et al. 1999; Heyer et al. 2003, 2004). The single fiber and behavioral data are consonant with the hypothesis that a subset of taste receptor cells responds to both sweet and umami stimuli. However, because T1R1 and T1R2 are usually not expressed in the same taste cells, the behavioral and electrophysiological overlap between sweet and umami qualities may indicate that umami-sensitive receptors other than the T1R1/T1R3 heteromer may be expressed in T1R2 (sweet responsive) taste cells. Consistent with this hypothesis, previously published data indicate the possibility of additional umami receptors including a truncated form of mGluR4 (Chaudhari et al. 2000; Maruyama et al. 2006). Functional studies of T1R3 knockout mice have produced different conclusions about whether T1R1/T1R3 is the only umami receptor. Damak et al. (2003) and Maruyama et al. (2006) noted residual responses to umami stimuli in T1R3 knockout mice and concluded that receptors other than T1R1/T1R3 must be responsible for the remaining response. In contrast, studies by Zhao et al. (2003) indicated that T1R1 and T1R3 knockout mice completely lose their ability to detect umami tastants, and therefore, the T1R1/T1R3 receptor is the only receptor responsible for mediating umami taste. Our data are consonant with the hypothesis of multiple umami receptors, some of which are expressed in T1R2-expressing, sweet-sensitive receptor cells.
The idea that G proteins other than gustducin play a role in umami taste is also supported by the present study. Previous studies of vallate and foliate taste buds indicated that the sweet-sensitive T1R2/T1R3 receptor is expressed in different taste cells than gustducin (Hoon et al. 1999; Adler et al. 2000; Montmayeur et al. 2001; Kim et al. 2003). Yet, gustducin knockout mice are compromised in their ability to detect sweet compounds (Wong et al. 1996). Thus, it was unclear how gustducin could impact on the sweet transduction pathway as mediated by the T1R2/T1R3 heteromer. We found that in the palate, most T1R2 and all T1R3 taste cells coexpress gustducin. Our data from palate and fungiform taste buds expand on the description of gustducin expression in T1R2-expressing cells of fungiform taste buds (Kim et al. 2003), suggesting that an -gustducin–mediated pathway does exist in T1R2/T1R3-positive cells in these highly sweet-responsive taste fields. This is consistent with the behavioral data that support a role for gustducin in sweet taste transduction. Furthermore, because T1R2 and T1R3 are present in vallate and foliate taste buds, but are not expressed in gustducin cells, another G protein, perhaps Gi or Gs, may be involved in the limited sweet responsiveness in those taste fields. We also found that T1R1 is coexpressed with gustducin only half of the time, even in the palate. This is consistent with the idea that additional G proteins are involved in umami transduction as reported by Ruiz et al. (2003) and He et al. (2004).
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This work was supported by National Institutes of Health grants RO1 DC006021 and P30 DC04657. The authors thank Dr Nicholas Ryba and Dr Charles Zuker for the cDNA plasmids for mouse T1R1 and T1R2 and Dr Robert Margolskee for the T1R3 antibody. In addition, we thank Elodie Chaplain for technical assistance.
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References |
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Accepted 8 December 2006
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