The G-protein-coupled sweet taste receptor dimer T1R2/T1R3 is expressed in taste bud cells in the oral cavity. In recent years, its involvement in membrane glucose sensing was discovered in endocrine cells regulating glucose homeostasis. We investigated importance of extraorally expressed T1R3 taste receptor protein in age-dependent control of blood glucose homeostasis in vivo, using nonfasted mice with a targeted mutation of the Tas1r3 gene that encodes the T1R3 protein. Glucose and insulin tolerance tests, as well as behavioral tests measuring taste responses to sucrose solutions, were performed with C57BL/6ByJ (Tas1r3+/+) inbred mice bearing the wild-type allele and C57BL/6J-Tas1r3tm1Rfm mice lacking the entire Tas1r3 coding region and devoid of the T1R3 protein (Tas1r3-/-). Compared with Tas1r3+/+ mice, Tas1r3-/- mice lacked attraction to sucrose in brief-access licking tests, had diminished taste preferences for sucrose solutions in the two-bottle tests, and had reduced insulin sensitivity and tolerance to glucose administered intraperitoneally or intragastrically, which suggests that these effects are due to absence of T1R3. Impairment of glucose clearance in Tas1r3-/- mice was exacerbated with age after intraperitoneal but not intragastric administration of glucose, pointing to a compensatory role of extraoral T1R3-dependent mechanisms in offsetting age-dependent decline in regulation of glucose homeostasis. Incretin effects were similar in Tas1r3+/+ and Tas1r3-/- mice, which suggests that control of blood glucose clearance is associated with effects of extraoral T1R3 in tissues other than the gastrointestinal tract. Collectively, the obtained data demonstrate that the T1R3 receptor protein plays an important role in control of glucose homeostasis not only by regulating sugar intake but also via its extraoral function, probably in the pancreas and brain.
Citation: Murovets VO, Bachmanov AA, Zolotarev VA (2015) Impaired Glucose Metabolism in Mice Lacking the Tas1r3 Taste Receptor Gene. PLoS ONE 10(6): e0130997. https://doi.org/10.1371/journal.pone.0130997
Editor: Hiroaki Matsunami, Duke University, UNITED STATES
Received: October 21, 2014; Accepted: May 27, 2015; Published: June 24, 2015
Copyright: © 2015 Murovets et al. This is an open access article distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited
Data Availability: All relevant data are within the paper and its Supporting Information files.
Funding: This work was supported by the National Institutes of Health (NIH) (http://www.nih.gov/) grant# R03DC013526 to AAB and VAZ and National Institutes of Health (NIH) (http://www.nih.gov/) grant# R01DC00882 to AAB. The funders had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript.
Competing interests: The authors have declared that no competing interests exist.
The search for key regulators of blood or tissue glucose levels is relevant to treatments of diabetes, obesity, and metabolic syndrome. Sensing of glucose in viscera and brain is crucial for control of energy homeostasis. Cells involved in regulation of blood glucose concentration (the insulin-secreting β-cells of the pancreas, enteroendocrine L-cells of the small intestine, and glucose-excited neurons of ventromedial hypothalamus) share a metabolic mechanism of glucose sensing controlled by glucose transporters and glucokinase (hexokinase IV). This mechanism involves an increase of cytoplasmic glucose resulting in a series of intracellular events leading to a rise in the cytosolic ATP/ADP ratio and subsequent closure of ATP-sensitive potassium (KATP) channels, which leads to cell depolarization [1, 2]. However, several lines of evidence, including pharmacological blockage of glucokinase and gene knockout of the Kir6.2 subunit of the KATP channel, strongly suggest that glucosensing involves additional signaling pathways that do not require intracellular metabolic processing of glucose, that is, KATP-independent pathways [3–6]. In recent studies, membrane glucose-sensing mechanisms involving the T1R2/T1R3 heterodimeric complex of G-protein-coupled sweet taste receptor proteins [7, 8] and related intracellular transduction components, operating independently of cellular glucose transport and metabolism, were found in gastrointestinal, nervous, and endocrine cells regulating glucose homeostasis [9–17].
Results of in vitro and some in vivo studies confirm the role of T1R-related mechanisms in regulation of glucose metabolism. In cultures of enteroendocrine cells, these mechanisms involve insulinotropic hormones, or incretins: glucagon-like peptide 1 (GLP-1) and glucose-dependent insulinotropic peptide (GIP) [11–14]. Consistent with this, mice lacking Gα-gustducin or T1R3 demonstrated deficient incretin production and glucose tolerance after administration of glucose in the gastrointestinal tract [18–20].
In cultures of pancreatic islets or the glucose-responsive β-cell line MIN6, T1R-related mechanisms of glucose regulation involve insulin secretion [16, 17, 21]. However, physiological importance of pancreatic sweet taste receptors in control of blood glucose level in vivo was examined in only a few studies, which did not fully confirm it. In fasted mice, deletion of T1R2 or T1R3 did not affect glucose tolerance after systemic administration of glucose, which bypasses the intestinal lumen and thus does not induce secretion of incretins [19, 22]. This lack of consistency between the in vitro and in vivo studies may be due to differing nutrition status of cells in these two types of experiments. While in vitro studies use cell cultures supplied with nutrients, in vivo studies typically involve testing food-deprived mice. Overnight fasting (typically for 16–18 h) provokes in mice, which are nocturnal and eat during nighttime, a catabolic state and substantial reduction of incretins and insulin release, as well as changes in insulin sensitivity [23–25]. In contrast to humans, in rodents prolonged fast also enhances insulin-stimulated glucose utilization [26, 27]. Thus, overnight fasting is considered more useful for studies of glucose utilization (e.g., effects on muscle uptake of glucose), whereas reduced fast duration is better for assessing insulin action within a more physiological context .
Therefore, we compared glucose tolerance of nonfasted Tas1r3 knockout  and wild-type mice to examine the in vivo importance of the extraoral T1R3 taste receptor protein in controlling blood glucose homeostasis. To assess the role of T1R3 in the effect of incretins, we compared glucose clearance after intragastric or intraperitoneal administration of glucose. Additionally, there is substantial evidence showing that aging is associated with decreased glucose tolerance, primarily due to impairment of β-cell sensitivity to glucose, decreased insulin production, and increased tissue tolerance to insulin (for review see [29, 30]). To examine whether aging could affect involvement of extraoral sweet taste reception in glucose metabolism, we have studied effects of Tas1r3 deletion on glucose and insulin tolerance in mice of different ages. We confirmed the role of the oral T1R3 receptor in behavioral studies assessing taste responses to sucrose in Tas1r3 knockout and wild-type mice.
Materials and Methods
The described experimental procedures have been approved by the Institutional Animal Care and Use Committee (IACUC) at the Pavlov Institute of Physiology (Animal Welfare Assurance #A5952-01). The study was performed with 8- to 36-week-old male mice of two strains: C57BL/6ByJ bearing the wild-type Tas1r3 allele, used as control (Tas1r3+/+; Jackson Laboratory, Bar Harbor, ME), and C57BL/6J-Tas1r3tm1Rfm lacking the entire T1R3 coding region and devoid of T1R3 protein  (Tas1r3-/-; kindly provided by Dr. R. F. Margolskee, Monell Chemical Senses Center, Philadelphia, PA, USA). Separate groups of mice were used in different tests. (Numbers of mice are shown in the table and figures below.) During the study, animals were housed individually (taste tests) or by 4–5 in standard polycarbonate cages on a 12-h light-dark cycle (lights on at 8:00 a.m.) in a temperature- and humidity-controlled room. Throughout the study, mice were fed with a standard lab chow (PK-120, MEST Ltd., Moscow, Russia) containing 67% carbohydrates, 5% lipids, and 19% proteins, with an energy value of 13,000 kJ/kg; food and tap water were available ad libitum.
Behavioral taste responses to 0.03–0.93 mol/L sucrose (Sigma-Aldrich St. Louis, MO, USA) were assessed in separate groups of mice using the brief-access licking test (BALT) and the 48-h two-bottle preference test. The BALT was conducted during the light period using procedures similar to those described by Glendinning et al. . Before testing, mice had restricted access to water (1.5 mL for 22–23 h), while access to food remained unlimited. During the test session, an animal was exposed in gustometer Davis MS-160 (DiLog Instruments, Tallahassee, FL, USA) to three repetitive blocks of stimuli, each consisting of eight trials: six concentrations of sucrose presented in ascending order, and two presentations of distilled water as "washout," one before each of the two highest sucrose concentrations. Access to each solution lasted for 5 s, with 20-s interpresentation interval. Licking ratio was calculated as the percentage of licks to sucrose solution relative to the mean number of licks in the two preceding trials with water. In the 48-h two-bottle tests , mice in their home cages had free access to two tubes containing distilled water or sucrose solution. Positions of tubes was changed after 24 h. Sucrose preference was calculated as consumed sucrose solution as a percentage of total fluid intake.
Glucose and insulin tolerance tests
Glucose and insulin tolerance tests were started at 3–4 p.m. (7–8 h after the beginning of the light period). In the glucose tolerance test (GTT), matching specific recommendations of the Mouse Metabolic Phenotyping Center , nonfasted conscious animals received glucose (2 g/kg, 0.1 ml per 10 g body weight; Sigma-Aldrich St. Louis, MO, USA) either intraperitoneally (IP) or by intragastric gavage (IG). Aqueous solution was used in the IG GTT; in the IP GTT, glucose was dissolved in saline. In the insulin tolerance test (ITT), nonfasted mice were injected with insulin (2 U/kg, IP; insulin aspart, Novo Nordisk A/S, Bagsvaerd, Denmark). Blood was sampled by tail cut, and two measurements of glucose concentration for each time point were made 0–120 min after the infusion of glucose or insulin using the One Touch Ultra glucometer (LifeScan, Inc., USA). During the GTT, animals were gently held in custom-made restraint tubes, to which they were habituated during the preceding 2 days. During the ITT, mice were left unrestrained in their home cages.
Statistical analysis was performed using Statistica 7.0 software (StatSoft, Tulsa, OK, USA). Data from the behavioral taste tests and glucose clearance in the GTT and ITT were compared with two-way ANOVA. Concentration (for taste tests) and time (for GTT or ITT) were considered as within-subject factors, and strain was considered as a between-subject factor. Post hoc paired comparisons were made with Fisher’s least significant difference (LSD) test. Blood glucose area under the curve (AUC) was calculated using the trapezoidal rule, and differences between AUCs were assessed with one-way ANOVA. Between-strain comparisons of baseline glucose, body weight, and age were performed using the Student’s t-test. To quantify the correlation of physiological parameters within groups, the Pearson product-moment correlation coefficient was used. All data are presented as mean ± SEM; P values less than 0.05 were considered significant.
Body weight of Tas1r3-/- mice was about 6% greater than body weight of Tas1r3+/+ mice (t-test, p<0.01), and for both strains it increased with age (Table 1). Baseline blood glucose level was similar in both strains. There was no significant relationship between baseline glucose and body weight or age (Table 1).
Tas1r3-/- mice had a substantially reduced attraction to sucrose both in the BALT (Fig 1A) and in the 48-h two-bottle test (Fig 1B). Two-way ANOVA of the BALT data revealed significant effects of strain (F(1, 36) = 65.13, P<0.001) and concentration (F(5, 180) = 3.04, p<0.001), as well as strain × concentration interaction (F(5, 180) = 2.33, P<0.01). Mouse strains significantly differed in licking 0.23 and 0.46 mol/L sucrose solutions. A concentration-dependent increase in the licking ratio of sucrose to water was detected for Tas1r3+/+ mice at concentrations greater than 0.06 mol/L (P<0.01, Fisher LSD test); Tas1r3-/- mice licked all concentrations of sucrose at the same rate as they licked water. Similarly, in the 48-h two-bottle tests, two-way ANOVA also revealed strong strain differences in preference for 0.03–0.93 mol/L sucrose solutions (effect of strain: F(1, 28) = 640.42, P<0.0001; effect of concentration: F(5, 140) = 93.44, P<0.0001, strain × concentration interaction: F(5, 140) = 49.68, P<0.0001). Strain comparisons between preference scores at different concentrations confirmed concentration dependence of response. Tas1r3+/+ mice clearly preferred sucrose to water at concentrations of 0.03 mol/L and higher and showed maximal level of sucrose preference starting at 0.06 mol/L. Knockout mice were indifferent to 0.03–0.12 mol/L sucrose and preferred 0.23 mol/L and higher concentrations (P<0.001, Fisher LSD test).
A) Licking ratio (%) as a function of sucrose concentration in the brief-access licking test (mean±SEM); n(Tas1r3+/+) = 15, n(Tas1r3-/-) = 29. B) Sucrose preference scores (%) in the 48-h two bottle test; n(Tas1r3+/+) = 18, n(Tas1r3-/-) = 12. Post hoc comparisons with Fisher LSD test (Tas1r3 +/+ vs. Tas1r3-/-): *—p<0.05, ***—p<0.001
After IP load with glucose, a significant Pearson’s correlation between AUC of the time course of blood glucose concentration and age was found for Tas1r3-/- mice (r = 0.59, P<0.05), while Tas1r3+/+ mice demonstrated only a nonsignificant tendency of age dependence (Fig 2A). Based on this result, we divided animals of each strain into two age-matched groups (9–21 and 22–34 weeks old) and analyzed within-group differences. Tas1r3-/- mice of both ages had significantly impaired glucose tolerance compared with Tas1r3+/+ mice (Fig 2B and 2C). Although initial peaks of glucose concentrations (15 min after IP administration of glucose) were similar in Tas1r3-/- and Tas1r3+/+ mice, the subsequent decrease of blood glucose level was much slower in Tas1r3-/- mice, particularly in the older group. For 9- to 21-week-old mice, two-way ANOVA revealed significant effects of strain (F(1, 35) = 8.80, P<0.01), time (F(7, 245) = 112.11, P<0.000001), and strain × time interaction (F(7, 245) = 6.72, P<0.000001); for 22- to 34-week-old animals two-way ANOVA revealed significant effects of strain (F(1, 20) = 12.60, P<0.01), time (F(7, 170) = 59.95, P<0.000001), and strain × time interaction (F(7, 140) = 6.95, p<0.000001). In the 9- to 21-week-old group, blood glucose AUC of Tas1r3-/- mice was about 25% greater than in age-matched Tas1r3+/+ mice (Fig 2B; F(1, 35) = 9.38, P<0.01, one-way ANOVA); in the 22- to 34-week-old group it was about 75% greater (Fig 2C; F(1, 20) = 15.00, P<0.001, one-way ANOVA).
A) Relationship between glucose AUC and age. Pearson’s coefficient of correlation was calculated; n(Tas1r3+/+) = 29, n(Tas1r3-/-) = 30. B, C) Blood glucose concentration (left) and glucose AUC (right) in 9- to 21-week-old (B) and 22- to 34-week-old (C) mice. B) n(Tas1r3+/+) = 19, n(Tas1r3-/-) = 18; C) n(Tas1r3+/+) = 10; n(Tas1r3-/-) = 12. Post hoc comparisons with Fisher LSD test (Tas1r3 +/+ vs. Tas1r3-/-): *—p<0.05, **—p<0.01, ***—p<0.001.
In the IG GTT, no significant correlations between blood glucose concentration and age were found for either Tas1r3+/+ or Tas1r3-/- animals (Fig 3A). There was a marked augmentation of blood glucose levels in both age groups of knockout mice compared with Tas1r3+/+ mice (Fig 3B and 3C). For 9- to 21-week-old and 22- to 34-week-old mice, respectively, two-way ANOVA showed significant effects of strain (F(1, 26) = 16.20, P<0.001; and F(1, 19) = 4.71, p<0.05), time (F(7, 182) = 103.10, P<0.000001; and F(7, 133) = 47.69, p<0.000001), and their interaction (F(7, 182) = 3.76, P<0.001; and F(7, 133) = 4.42, P<0.001). In both age groups, blood glucose AUC was about 30% greater in Tas1r3-/- mice than in age-matched Tas1r3+/+ mice, as confirmed by one-way ANOVA (for 9- to 21- and 22- to 34-week-old mice, respectively: F(1, 26) = 13.52, P<0.01; and F(1, 19) = 4.26, p<0.05; Fig 3B and 3C).
A) Relationship between glucose AUC and age. Pearson’s coefficient of correlation was calculated. n(Tas1r3+/+) = 23, n(Tas1r3-/-) = 26. B, C) Blood glucose concentration (left) and glucose AUC (right) in 9- to 21-week-old (B) and 22- to 34-week-old (C) mice. B) n(Tas1r3+/+) = 13, n(Tas1r3-/-) = 15; C) n(Tas1r3+/+) = 10; n(Tas1r3-/-) = 11. Post hoc comparisons with Fisher LSD test (Tas1r3 +/+ vs. Tas1r3-/-): *—p<0.05, ***—p<0.001.
Additional analysis of the data showed an impact of the route of glucose administration, which was similar in both strains (S1 Fig). In general, blood glucose utilization occurred faster after IG infusion than after IP load in mice of both age groups. For 9- to 21- and 22- to 34-week-old Tas1r3+/+ mice, respectively, two-way ANOVA showed a significant influence of the route of glucose administration (F(1, 19) = 6.40, p<0.05; and F(1, 13) = 15.23, p<0.002), time (F(7, 133) = 35.99, p<0.0001; and F(7, 91) = 40.01, p<0.0001), and their interactions (F(7, 133) = 0.96, p>0.46; and F(7, 91) = 3.60, p<0.002). Glucose AUC was greater in the IP GTT groups than in the IG GTT groups of Tas1r3+/+ mice (one-way ANOVA: F(1, 19) = 4.88, p<0.05; and F(1, 13) = 12.03, p<0.01, respectively). For 9- to 21- and 22- to 34-week-old Tas1r3-/- mice, respectively, two-way ANOVA also showed an effect of the route of glucose administration (F(1, 26) = 5.05, p<0.05; and F(1, 21) = 9.42, p<0.01), time (F(7, 182) = 109.72, p<0.0001; and F(7, 147) = 73.49, p<0.0001), and their interactions (F(7, 182) = 0.94, p>0.47; and F(7, 147) = 2.70, p<0.05). Glucose AUC after IP administration was significantly larger than after IG administration only in the older group of Tas1r3-/- mice (one-way ANOVA: F(1, 21) = 8.33, p<0.01) but not in the younger group (F(1, 26) = 3.51, p>0.07).
Injection of insulin (2 U/kg, IP) caused a rapid reduction of blood glucose concentration, reaching a minimum 15 min after injection (Fig 4B and 4C). In both strains, basal glucose level was completely restored within 120 min. Hypoglycemia induced by insulin did not depend on age (Fig 4A) or body weight in either strain (data not shown); therefore, calculations were made for the combined group with ages ranging from 8 to 36 weeks. Analysis of both absolute data (Fig 4B) and percentage of basal glucose level (Fig 4C) demonstrated that Tas1r3-/- mice had impaired sensitivity to insulin. For absolute values, the two-way ANOVA revealed a significant effect of time (F(3, 108) = 211.16, P<0.000001) and strain × time interaction (F(3, 108) = 11.34, P<0.01); effect of strain was nonsignificant. Fisher’s LSD post hoc test revealed a difference in glucose levels between Tas1r3-/- and Tas1r3+/+ mice at 60 min (Fig 4B; P<0.05, n = 20–31). Comparisons of values normalized relative to baseline showed significant effects of strain (F(1, 36) = 4.79, P<0.05), time (F(2, 72) = 167.10, P<0.000001), and strain × time interaction (F(2, 72) = 8.63, P<0.001). Post hoc tests confirmed difference in normalized glucose levels between Tas1r3-/- and Tas1r3+/+ mice at 15 and 60 min (Fig 4C; Fisher’s LSD, P<0.05).
A) Relationship between glucose AUC and age. Pearson’s correlation coefficients (r) were calculated. B) Absolute values of blood glucose concentration. C) Percentage relative to baseline level. Insulin (2 U/kg, IP) was injected at zero time point; n(Tas1r3+/+) = 18, n(Tas1r3-/-) = 20. Post hoc comparisons with Fisher LSD test (Tas1r3 +/+ vs. Tas1r3-/-): *—p<0.05
Our results support the important role of cell membrane sensing of sweeteners with T1R3 taste receptor protein at different levels of control of carbohydrate ingestion and homeostasis. Mice lacking the Tas1r3 gene demonstrated reduced behavioral taste responses to sucrose: they had totally suppressed attraction to sucrose when they were presented with different concentrations for 5-s periods in the BALT (Fig 1A) and had lower preferences for sucrose than Tas1r3+/+ mice in the 48-h two-bottle test. However, unlike during the BALT, during the long-term preference tests Tas1r3-/- mice preferred higher concentrations of sucrose (≥0.23 mol/L) over water (Fig 1B). These data are consistent with earlier studies of Tas1r3-/- mice, which revealed a residual behavioral preference of caloric sugars but not nonnutritive sweeteners such as sucralose, acesulfame K, SC-45647, and saccharin, in long-term preference tests [28, 33]. Both Tas1r3-independent pre- and postingestive mechanisms could be involved in the observed residual preference of sucrose. The Tas1r3-independent preingestive effects are supported by results from a recent study with Tas1r3-/-, TRPM5-/-, and Gα-gustducin-knockout mice, in which chorda tympani nerve activity in response to sucrose was reduced by roughly 80% relative to wild-type mice but responses to glucose were reduced only by 40% . This suggests that in taste cells, sugars such as sucrose and glucose can activate an additional taste transduction pathway that does not require T1R3, gustducin’s Gα subunit, or TRPM5 and was proposed to be metabolic and KATP channel dependent . The Tas1r3-independent postingestive factors likely include conditioned flavor preference reinforced by nutritive value of sucrose .
Our results demonstrate that in nonfasted mice Tas1r3 deficiency markedly worsens glucose tolerance, regardless of whether the route of glucose administration is intragastric or intraperitoneal (Figs 2 and 3), indicating possible involvement of T1R3-mediated glucose sensing in intestinal enteroendocrine, pancreatic, and/or brain mechanisms controlling glucose metabolism. It is well established that T1R3 is expressed in a variety of tissues beyond the tongue and gut mucosa (e.g., 9–15); however, it is still not clear to what extent these extraoral taste receptors are involved in control of carbohydrate metabolism. In early studies in the human pancreas, T1R3 was immunolabeled in excretory ducts and centroacinar cells, but the endocrine portion of the gland was immunonegative . Later, RT-PCR showed expression of the TAS1R3 gene in human pancreatic islets  and in MIN6 cells, a glucose-responsive β-cell line . Mouse islets  and MIN6 cells  express elements of intracellular taste signal transduction cascade as well. The sweet taste receptor system of mouse pancreatic β-cells and MIN6 cells seems functional since artificial sweeteners are able to stimulate insulin secretion, which was attenuated by gurmarin, an inhibitor of the mouse sweet taste receptor [16, 22]. In human pancreatic islets, potentiation of insulin release induced by fructose was suppressed by lactisole, an allosteric inhibitor of human T1R3. Further, in vitro, genetic ablation of T1R2 or T1R3 led to substantial reduction of the effect of sweeteners on insulin output from mouse islets [19, 22].
In contrast with these results of in vitro studies, recent in vivo studies in food-deprived mice revealed that the lack of T1R2  or T1R3  had no significant effect on the blood glucose level after IP administration of glucose, although after IG glucose administration Tas1r3-/- mice had higher blood glucose and lower plasma insulin levels than did wild-type controls . A likely explanation for this discrepancy between in vitro and in vivo results is the difference in nutrition status of cells. In cultured mouse islets, positive effects of fructose or noncaloric sweeteners on insulin secretion require presence of an optimal glucose level in the medium. For instance, a sharp reduction of glucose concentration in islet media abolished the potentiating effect of fructose  and stimulated activity of noncaloric sweeteners  in MIN6 cells. Therefore, pre-experimental fasting can also influence results of in vivo experiments. Overnight fasting provokes a catabolic state in mice, which have a unique metabolic response to prolonged fasting that differs from the response to fasting seen in humans. Specifically, fasting impairs insulin-stimulated glucose utilization in humans but enhances it in normal mice [26, 27]. In mice and rats, fasting, or even mild caloric deprivation, leads to the increase in insulin binding in the tissues [38, 39]. Earlier, we found out that effect of T1R3 ablation on glucose utilization was more pronounced in euglycaemic state than after fasting . The present data show that in mice in the nonfasted state, when β-cells are already partially depolarized due to KATP-dependent mechanisms [22, 35] and maintain basal levels of insulin secretion, deletion of T1R3 causes a significant impairment of glucose tolerance in both IP GTT and IG GTT. Thus, the apparent discrepancy between our data and these previous results is likely due to prolonged fasting used in these other studies, which likely caused marked changes in blood glucose regulation mechanisms.
Potentially, T1R3-signalling could be also involved in regulation of glycogenolysis and/or gluconeogenesis. In the adipose tissue, T1R3-signalling induced by non-caloric sweeteners stimulates adipogenesis and suppresses lipolysis . T1R3 is expressed in excretory ducts of the liver , where it probably does not interact with glycogenolysis. However, an involvement of T1R3-dependent mechanisms in fat, liver and other tissues in direct or indirect control of glycogen breakdown and gluconeogenesis in our study is unlikely because animals in all our experiments were in non-fasting state, in which synthesis of glucose from polysaccharides and non-carbohydrates is suppressed.
Additionally, our results demonstrate that in the IP (but not IG) GTT, the effect of T1R3 deletion was age related (Fig 2A), suggesting that normal T1R3-mediated extraoral sensing of sweeteners somewhat prevents deterioration of glucose tolerance with age. Decreased insulin secretion due to the loss of β-cell mass or impaired β-cell function and increased insulin resistance are considered two major factors leading to impaired glucose tolerance in the elderly [42–44].
According to the classical concept, the oral ingestion of glucose stimulates more insulin release than does intravenous infusion while causing a similar elevation of the plasma glucose level . This phenomenon, known as the incretin effect, is largely attributable to two insulinotropic hormones released in response to food ingestion from intestinal enteroendocrine K-cells (GIP) or L-cells (GLP-1). Both GIP and GLP-1 have direct stimulatory effects on pancreatic β-cells (for review see ). The combined action of incretins is believed to account for about 50% of the total insulin secretory response after a meal . In recent years, sweet taste molecules, including T1R3, as well as intracellular taste signal transduction machinery in the gut enteroendocrine cells were described among the regulators of incretin production. Immunolabeling has revealed taste signal transducing elements in a number of intestinal L-cells, ranging from 15% in mouse jejunum up to 90% in the human duodenum [11, 48], whereas K-cells likely express only marginal levels of sweet taste protein transcripts . The artificial sweetener sucralose administrated to the mouse enteroendocrine GLUTag cell line or to the human L-cell line NCI-H716 enhances GLP-1 output that could be blocked by species-specific inhibitors of the sweet taste receptors [14, 18]. Knockout mice lacking T1R3, or ileum explants from these mice, showed markedly reduced GLP-1 release in response to luminal infusion of glucose [19, 20]. Consistent with this and with our results (Fig 3), Tas1r3-/- mice had higher blood glucose and lower plasma insulin levels during an oral glucose challenge compared with wild-type controls . However, in mouse or rat duodenum and jejunum, only a small number of taste proteins are colocalized with enteroendocrine cells [13, 49], and there is still no convincing evidence that T1R3-dependent intestinal endocrine mechanisms are potent enough to control blood glucose levels in vivo.
In our study, like in classical investigations , the involvement of T1R3 in regulation of intestinal secretion of incretins could be evident by comparing blood glucose clearance after administration of the same dose of glucose by different routes. We show that both Tas1r3+/+ and Tas1r3-/- mice demonstrated similar incretin effects (S1 Fig): in both types of mice blood glucose clearance was more active after IG glucose administration than after IP administration. Pancreatic β-cells and gut enteroendocrine cells use a common metabolic mechanism of glucose sensing, which requires glucose transporter GLUT2, the glycolytic enzyme glucokinase, and the KATP channel [50–52]. Therefore, because the route of glucose administration affected blood glucose clearance in Tas1r3-/- mice, we suggest that in the euglycemic state KATP-dependent metabolic mechanisms predominantly determine gut regulation of the glucose homeostasis.
Impaired glucose tolerance is usually associated with reduced insulin sensitivity, which was also demonstrated for Tas1r3-/- mice in our study (Fig 4A). Higher body mass of Tas1r3-/- mice could have contributed to their lower insulin sensitivity, but the difference in body weight was small (about 6%, Table 1), and body weight did not correlate with glucose level. Reduction of insulin tolerance also did not correlate with age (Fig 4B) and body weight. Therefore, higher body weight of Tas1r3-/- mice seems insufficient to explain their reduced insulin sensitivity. Another possible cause of decreased insulin sensitivity of Tas1r3-/- mice could be chronic elevation of postprandial glucose level, which was shown in our glucose tolerance experiments. In particular, raised blood glucose levels cause overactivity of the hexosamine biosynthesis pathway of glycolysis via modulation of transcriptional factors by O-N-acetylglucosamine, including transcriptional factors of the insulin receptor substrate and probably GLUT4 (for review see ), which may lead to reduced insulin sensitivity observed in Tas1r3-/- mice.
There is evidence that in addition to the gastrointestinal tract and pancreas, the central nervous system may have sweet taste signaling mechanisms that play an important role in regulating glucose homeostasis and therefore may be involved in effects of T1R3 deficiency found in this study. The fall of central glucose levels causes a sequence of neurohormonal reactions known as feedback response launched mainly by activation of glucose-sensing neurons in ventromedial hypothalamic nuclei, orexin neurons in perifornical area, and neurons in the brainstem [54–56]; this includes sympathoadrenal activation followed by increases of plasma epinephrine, norepinephrine, and glucagon, which in turn leads to hepatic gluconeogenesis and inhibition of pancreatic insulin secretion . An acute increase in central glucose, which likely occurs in our experimental protocol, results in an opposite response: an increase in insulin levels and suppression of hepatic glucose production through reduction of gluconeogenesis and glycogenolysis . Several mechanisms of glucose sensing, which do not require intracellular glucose metabolism or glucokinase/KATP pathways, have been demonstrated in the hypothalamus (for review see ). It is quite plausible that glucosensing neurons could use a sweet taste receptor. Ren et al.  have reported that T1Rs and α-gustducin are highly expressed in neurons of mouse hypothalamus compared with cortex and hippocampus. Strong expression of T1R2 and T1R3 was found in arcuate and paraventricular nuclei of the hypothalamus, as well as in the medial habenula and the epithelial cells of the choroid plexus. Importantly, the arcuate nucleus is a key region detecting peripheral metabolic status and then relaying this information to other hypothalamic nuclei, including the ventromedial nucleus and paraventricular nuclei 53].
Most of glucose-sensing neurons are glucose inhibited (GI) and reduce their activity during elevation of blood glucose above the euglycemic level. Both glucose-excited (GE) and GI neurons of the hypothalamus are extremely sensitive to glucose changes when extracellular concentrations are less than 2 mM, which occurs at euglycemic blood levels , and they have minimal response when glucose levels in the hypothalamus exceed 2 mM, suggesting that these glucose-sensing neurons primarily sense glucose deficit [5, 61]. An additional smaller population of glucose-sensing units is present in the arcuate nucleus, which include high-GE and high-GI neurons responding to an increase in extracellular glucose from 5 to 20 mM; however, it is still not clear whether these neurons play role in regulating hyperglycemic states. It is interesting that glucose sensing of the high-GE and high-GI neurons is KATP independent . Collectively, there is evidence that glucose-metabolism-independent pathways in the central nervous system may involve T1Rs and take place under control of peripheral glucose homeostasis in the hyperglycemic state; however, their role needs to be elucidated. Additional investigations of neurotransmitter and hormonal specificity of hypothalamic neurons expressing T1R3 possibly will shed light on their physiological relevance.
Measures of glucose metabolism used in this study may have been affected by direct effects of absence of the T1R3 protein in extraoral cells, as well as by indirect effects from these cells to other tissues (e.g., mediated by metabolic, hormonal or paracrine effects). The goal of our ongoing studies is to find out, which of these mechanisms are involved in impaired glucose metabolism in Tas1r3-/- mice. If genetic variants of the Tas1r3 gene alter glucose metabolism in mice, then similar relationships may also exist in humans. Human T1R genes are polymorphic , and some of these polymorphisms are associated with taste functions [63–67]. Our study suggests that these functional polymorphisms of human T1R genes may also be associated with glucose metabolism and related diseases in humans. This emphasizes importance of human studies of T1R genes as potential new targets for diagnostics, prevention and treatment of metabolic diseases.
In conclusion, we have shown that the lack of attraction to sucrose demonstrated in Tas1r3-/- mice, compared with Tas1r3+/+ mice, is associated with reduced glucose tolerance in these mice. In nonfasted mice, the deletion of the T1R3 subunit of the sweet taste receptor results in substantial impairment of blood glucose clearance after both intragastric and intraperitoneal glucose administration. This clearly indicates involvement of the extraorally expressed T1R3 protein in control of glucose homeostasis in hyperglycemic states. Deletion of T1R3 had minor impact on the incretin effect, which suggests that intestinally expressed T1R3 protein is less important for regulation of blood glucose level compared with other extraoral sites, such as pancreas or brain. Reduced glucose tolerance after T1R3 deletion was associated with impaired insulin sensitivity. We have also demonstrated a marked age dependence of the effect of T1R3 receptor protein on blood glucose levels in the intraperitoneal glucose tolerance test. Altogether, our results suggest that further investigation of visceral reception of sugars with the T1R3 protein may lead to therapeutic approaches in the treatment of carbohydrate homeostasis disorders.
S1 Fig. Effect of intraperitoneal (IP) versus intragastric (IG) administration of glucose (2 g/kg) on blood glucose concentration (left) and glucose AUC (right).
Nonfasted Tas1r3+/+ (A, B) and Tas1r3-/- (C, D) mice 9–21 weeks of age (A, C) and 22–34 weeks of age (B, D). Post hoc comparisons with Fisher LSD test (IP vs. IG): *p<0.05, **p<0.001.
We thank Ms. Maria Theodorides and Mr. Serge V. Travnikov for technical assistance, and Dr. R. F. Margolskee for providing C57BL/6J-Tas1r3tm1Rfm mice. The partial data for taste responses to sucrose in Tas1r3+/+ and Tas1r3-/- mice (presented here in Figs 1A and 4B) have been published previously .
Conceived and designed the experiments: VOM AAB VAZ. Performed the experiments: VOM. Analyzed the data: VOM AAB VAZ. Contributed reagents/materials/analysis tools: VOM AAB VAZ. Wrote the paper: VOM AAB VAZ.
- 1. Mountjoy PD, Rutter GA. Glucose sensing by hypothalamic neurons and pancreatic islet cells: AMPle evidence for common mechanisms? Exp Physiol. 2007; 92(2): 311–3199. pmid:17158178
- 2. Reimann F, Habib AM, Tolhurst G, Parker HE, Rogers GJ, Gribble FM. Glucose sensing in L cells: a primary cell study. Cell Metab. 2008; 8(6): 532–9. pmid:19041768
- 3. Gembal M, Gilon P, Henquin JC. Evidence that glucose can control insulin release independently from its action on ATP—sensitive K+ channels in mouse β-cells. J Clin Invest. 1992; 89(4): 1288–1295. pmid:1556189
- 4. Straub SG, Sharp GW. Glucose-stimulated signaling pathways in biphasic insulin secretion. Diabetes Metab Res Rev. 2002; 18(6): 451–463. pmid:12469359
- 5. Wang R, Liu X, Hentges ST, Dunn-Meynell AA, Levin BE, Wang W, et al. The regulation of glucose-excited neurons in the hypothalamic arcuate nucleus by glucose and feeding-relevant peptides. Diabetes. 2004; 53: 1959–1965. pmid:15277373
- 6. Fioramonti X, Lorsignol A, Taupignon A, Pénicaud L. A new ATP-sensitive K+ channel-independent mechanism is involved in glucose-excited neurons of mouse arcuate nucleus. Diabetes. 2004; 53(11): 2767–2775. pmid:15504956
- 7. Bachmanov AA, Li X, Reed DR, Ohmen JD, Li S, Chen Z, et al. Positional cloning of the mouse saccharin preference (Sac) locus. Chem Senses. 2001; 26: 925–933. pmid:11555487
- 8. Nelson G, Hoon MA, Chandrashekar J, Zhang Y, Ryba NJ, Zuker CS. Mammalian sweet taste receptors. Cell. 2001; 106: 381–390. pmid:11509186
- 9. Hoefer D, Pueschel B, Drenckhahn D. (1996) Taste receptor-like cells in the rat gut identified by expression of alpha-gustducin. Proc Natl Acad Sci U SA. 1996; 93: 6631–6634.
- 10. Wu SV, Rozengurt N, Yang M, Young SH, Sinnett-Smith J, Rozengurt E. Expression of bitter taste receptors of the T2R family in the gastrointestinal tract and enteroendocrine STC-1 cells. Proc Natl Acad Sci USA. 2002; 99: 2392–2397. pmid:11854532
- 11. Rozengurt N, Wu SV, Chen MC, Huang C, Sternini C, Rozengurt E. Colocalization of the alphasubunit of gustducin with PYY and GLP-1 in L cells of human colon. Am J Physiol Gastrointest Liver Physiol. 2006; 291:G792–802. pmid:16728727
- 12. Bezençon C, le Coutre J, Damak S. Taste-signaling proteins are coexpressed in solitary intestinal epithelial cells. Chem Senses. 2007; 32(1): 41–49. pmid:17030556
- 13. Sutherland K, Young RL, Cooper NJ, Horowitz M, Blackshaw LA. Phenotypic characterization of taste cells of the mouse small intestine. Am J Physiol Gastrointest Liver Physiol. 2007; 292(5): G1420–1428. pmid:17290008
- 14. Jang HJ, Kokrashvili Z, Theodorakis MJ, Carlson OD, Kim BJ, Zhou Z, et al. Gut-expressed gustducin and taste receptors regulate secretion of glucagon-like peptide-1. Proc Natl Acad Sci U S A. 2007; 104(38): 15069–15074. pmid:17724330
- 15. Ren X, Zhou L, Terwilliger R, Newton SS, de Araujo IE. Sweet taste signaling functions as a hypothalamic glucose sensor. Front Integr Neurosci. 2009; 3:12. pmid:19587847
- 16. Nakagawa Y, Nagasawa M, Yamada S, Hara A, Mogami H. Sweet taste receptor expressed in pancreatic beta-cells activates the calcium and cyclic AMP signaling systems and stimulates insulin secretion. PLoS One. 2009; 4: e5106. pmid:19352508
- 17. Oya M, Suzuki H, Watanabe Y, Sato M, Tsuboi T. Amino acid taste receptor regulates insulin secretion in pancreatic β-cell line MIN6 cells. Genes Cells. 2011; 16(5): 608–616. pmid:21470345
- 18. Margolskee RF, Dyer J, Kokrashvili Z, Salmon KS, Ilegems E, Daly K, et al. T1R3 and gustducin in gut sense sugars to regulate expression of Na+-glucose cotransporter 1. Proc Natl Acad Sci U S A. 2007; 104(38): 15075–15080. pmid:17724332
- 19. Geraedts MC, Takahashi T, Vigues S, Markwardt ML, Nkobena A, Cockerham RE, et al. Transformation of postingestive glucose responses after deletion of sweet taste receptor subunits or gastric bypass surgery. Am J Physiol Endocrinol Metab. 2012; 303(4): E464–474. pmid:22669246
- 20. Kokrashvili Z, Mosinger B, Margolskee RF. Taste signaling elements expressed in gut enteroendocrine cells regulate nutrient-responsive secretion of gut hormones. Am J Clin Nutr. 2009; 90(3): 822S–825S. pmid:19571229
- 21. Kojima I, Nakagawa Y. The Role of the Sweet Taste Receptor in Enteroendocrine Cells and Pancreatic β-Cells. Diabetes Metab J. 2011; 35(5): 451–457.
- 22. Kyriazis GA, Soundarapandian MM, Tyrberg B. Sweet taste receptor signaling in beta cells mediates fructose-induced potentiation of glucose-stimulated insulin secretion. Proc Natl Acad Sci U S A. 2012; 109(8): E524–532. pmid:22315413
- 23. Andrikopoulos S, Blair AR, Deluca N, Fam BC, Proietto J. Evaluating the glucose tolerance test in mice. Am J Physiol Endocrinol Metab. 2008; 295(6): E1323–1332. pmid:18812462
- 24. Ayala JE, Samuel VT, Morton GJ, Obici S, Croniger CM, Shulman GI, et al. Standard operating procedures for describing and performing metabolic tests of glucose homeostasis in mice. Dis Model Mech. 2010; 3(9–10): 525–534. pmid:20713647
- 25. McGuinness OP, Ayala JE, Laughlin MR, Wasserman DH. NIH experiment in centralized mouse phenotyping: the Vanderbilt experience and recommendations for evaluating glucose homeostasis in the mouse. Am J Physiol Endocrinol Metab. 2009; 297(4): E849–855. pmid:19638507
- 26. Ayala JE, Bracy DP, McGuinness O P, Wasserman DH. Considerations in the design of hyperinsulinemic-euglycemic clamps in the conscious mouse. Diabetes. 2006; 55: 390–397. pmid:16443772
- 27. Heijboer AC, Donga E, Voshol PJ, Dang ZC, Havekes LM, Romijn JA, et al. Sixteen hours of fasting differentially affects hepatic and muscle insulin sensitivity in mice. J Lipid Res. 2005; 46: 582–588. pmid:15576835
- 28. Damak S, Rong M, Yasumatsu K, Kokrashvili Z, Varadarajan V, Zou S, et al. Detection of sweet and umami taste in the absence of taste receptor T1r3. Science. 2003; 301: 850–853. pmid:12869700
- 29. Shih SR, Tseng CH. The Effects of Aging on Glucose Metabolism. Taiwan Geriatrics & Gerontology. 2009; 4(1): 27–38.
- 30. Elahi D, Muller DC. Carbohydrate metabolism in the elderly. European Journal of Clinical Nutrition. 2000; 54(Suppl 3): S112–S120 pmid:11041082
- 31. Glendinning JI, Gresack J, Spector AC. A high-throughput screening procedure for identifying mice with aberrant taste and oromotor function. Chem Senses. 2002; 27(5): 461–474. pmid:12052783
- 32. Bachmanov AA, Reed DR, Beauchamp GK, Tordoff MG. Food intake, water intake, and drinking spout side preference of 28 mouse strains. Behavior Genetics. 2002; 32: 435–443. pmid:12467341
- 33. Zhao GQ, Zhang Y, Hoon MA, Chandrashekar J, Erlenbach I, Ryba NJ, et al. The Receptors for mammalian sweet and umami taste. Cell. 2003; 115: 255–266. pmid:14636554
- 34. Ohkuri T, Yasumatsu K, Horio N, Jyotaki M, Margolskee RF, Ninomiya Y. Multiple sweet receptors and transduction pathways revealed in knockout mice by temperature dependence and gurmarin sensitivity. Am J Physiol Regul Integr Comp Physiol. 2009; 296(4): 960–971.
- 35. Yee KK, Sukumaran SK, Kotha R, Gilbertson TA, Margolskee RF. Glucose transporters and ATP-gated K+ (KATP) metabolic sensors are present in type 1 taste receptor 3 (T1r3)-expressing taste cells. Proc Natl Acad Sci U S A. 2011; 108(13): 5431–5436. pmid:21383163
- 36. Sclafani A, Glass DS, Margolskee RF, Glendinning JI. Gut T1R3 sweet taste receptors do not mediate sucrose-conditioned flavor preferences in mice. Am J Physiol Regul Integr Comp Physiol. 2010; 299(6): R1643–1650. pmid:20926763
- 37. Taniguchi K. Expression of the sweet receptor protein, T1R3, in the human liver and pancreas. J Vet Med Sci. 2004; 66(11): 1311–1314. pmid:15585941
- 38. Olefsky JM, Reaven GM. Effects of age and obesity on insulin binding to isolated adipocytes. Endocrinology. 1975; 96: 1486–1490. pmid:165063
- 39. Soll AH, Goldfine ID, Roth J, Kahn CR, Neville D M Jr. Thymic lymphocytes in obese (ob/ob) mice. A mirror of the insulin receptor defect in liver and fat. J Biol Chem. 1974; 249: 4127–4131. pmid:4369380
- 40. Murovets VO, Bachmanov AA, Travnikov SV, Churikova AA, Zolotarev VA. The involvement of the T1R3 receptor protein in the control of glucose metabolism in mice at different levels of glycemia. J Evol Biochem Physiol. 2014; 50(4): 334–344. pmid:25983343
- 41. Simon BR, Learman BS, Parlee SD, Scheller EL, Mori H, Cawthorn WP, et al. Sweet taste receptor deficient mice have decreased adiposity and increased bone mass. PLoS One. 2014; 9(1): e86454. pmid:24466105
- 42. Chang AM, Smith MJ, Galecki AT, Bloem CJ, Halter JB. Impaired beta-cell function in human aging: response to nicotinic acid-induced insulin resistance. J Clin Endocrinol Metab. 2006; 91: 3303–3309. pmid:16757523
- 43. Maedler K, Schumann DM, Schulthess F, Oberholzer J, Bosco D, Berney T, et al Aging correlates with decreased beta-cell proliferative capacity and enhanced sensitivity to apoptosis: a potential role for Fas and pancreatic duodenal homeobox-1. Diabetes. 2006; 55: 2455–2462. pmid:16936193
- 44. Szoke E, Shrayyef MZ, Messing S, Woerle HJ, van Haeften TW, Meyer C, et al. Effect of aging on glucose homeostasis: accelerated deterioration of beta cell function in individuals with impaired glucose tolerance. Diabetes Care. 2008; 31: 539–543. pmid:18083793
- 45. McIntyre N, Holdsworth CD, Turner DS. New interpretation of oral glucose tolerance. Lancet. 1964; 2: 20–21.
- 46. Drucker DJ. Incretin action in the pancreas: potential promise, possible perils, and pathological pitfalls. Diabetes. 2013; 62(10): 3316–3323. pmid:23818527
- 47. Elrick H, Stimmler L, Hlad CJ Jr, Arai Y. Plasma insulin response to oral and intravenous glucose administration. J Clin Endocrinol Metab. 1964; 24: 1076–1082. pmid:14228531
- 48. Steinert RE, Gerspach AC, Gutmann H, Asarian L, Drewe J, Beglinger C. The functional involvement of gut-expressed sweet taste receptors in glucose-stimulated secretion of glucagon-like peptide-1 (GLP-1) and peptide YY (PYY). Clin Nutr. 2011; 30(4): 524–532.
- 49. Fujita Y, Wideman RD, Speck M, Asadi A, King DS, Webber TD, et al. Incretin release from gut is acutely enhanced by sugar but not by sweeteners in vivo. Am J Physiol Endocrinol Metab. 2009; 296(3): E473–479. pmid:19106249
- 50. Gribble FM, Williams L, Simpson AK, Reimann F. A novel glucose-sensing mechanism contributing to glucagon-like peptide-1 secretion from the GLUTag cell line. Diabetes. 2003; 52: 1147–1154. pmid:12716745
- 51. Preitner F, Ibberson M, Franklin I, Binnert C, Pende M, Gjinovci A, et al. Gluco-incretins control insulin secretion at multiple levels as revealed in mice lacking GLP-1 and GIP receptors. J Clin Invest. 2004; 113: 635–645. pmid:14966573
- 52. Reimann F, Gribble FM. Glucose-sensing in glucagon-like peptide-1-secreting cells. Diabetes. 2002; 51: 2757–2763. pmid:12196469
- 53. Buse MG. Hexosamines, insulin resistance, and the complications of diabetes: current status. Am J Physiol Endocrinol Metab. 2006; 290(1): E1–E8. pmid:16339923
- 54. Borg WP, Sherwin RS, During MJ, Borg MA, Shulman GI. Local ventromedial hypothalamus glucopenia triggers counterregulatory hormone release. Diabetes. 1995; 44(2): 180–184. pmid:7859938
- 55. Yi CX, Serlie MJ, Ackermans MT, Foppen E, Buijs RM, Sauerwein HP, et al. A major role for perifornical orexin neurons in the control of glucose metabolism in rats. Diabetes. 2009; 58(9): 1998–2005. pmid:19592616
- 56. Ritter S, Bugarith K, Dinh TT. Immunotoxic destruction of distinct catecholamine subgroups produces selective impairment of glucoregulatory responses and neuronal activation. J Comp Neurol. 2001; 432(2): 197–216. pmid:11241386
- 57. Nonogaki K. New insights into sympathetic regulation of glucose and fat metabolism. Diabetologia. 2000; 43(5): 533–549. pmid:10855527
- 58. Lam TK, Gutierrez-Juarez R, Pocai A, Rossetti L. Regulation of blood glucose by hypothalamic pyruvate metabolism. Science. 2005; 309(5736): 943–947. pmid:16081739
- 59. Burdakov D, Luckman SM, Verkhratsky A. Glucose-sensing neurons of the hypothalamus. Philos Trans R Soc Lond B Biol Sci. 2005; 360(1464): 2227–2235. pmid:16321792
- 60. Silver IA, Erecinska M. Extracellular glucose concentration in mammalian brain: continuous monitoring of changes during increased neuronal activity and upon limitation in oxygen supply in normo-, hypo-, and hyperglycemic animals. J Neurosci. 1994; 14, 5068–5076. pmid:8046468
- 61. Song Z, Routh VH. Differential effects of glucose and lactate on glucosensing neurons in the ventromedial hypothalamic nucleus. Diabetes. 2005; 54: 15–22. pmid:15616006
- 62. Kim UK, Wooding S, Riaz N, Jorde LB, Drayna D. Variation in the human TAS1R taste receptor genes. Chem Senses. 2006; 31: 599–611. pmid:16801379
- 63. Fushan AA, Simons CT, Slack JP, Manichaikul A, Drayna D. Allelic polymorphism within the TAS1R3 promoter is associated with human taste sensitivity to sucrose. Curr Biol. 2009; 19: 1288–93. pmid:19559618
- 64. Eny KM, Wolever TM, Corey PN, El-Sohemy A. Genetic variation in TAS1R2 (Ile191Val) is associated with consumption of sugars in overweight and obese individuals in 2 distinct populations. Am J Clin Nutr. 2010; 92: 1501–10. pmid:20943793
- 65. Chen QY, Alarcon S, Tharp A, Ahmed OM, Estrella NL, Greene TA, et al. Perceptual variation in umami taste and polymorphisms in TAS1R taste receptor genes. Am J Clin Nutr. 2009; 90: 770S–779S. pmid:19587085
- 66. Raliou M, Boucher Y, Wiencis A, Bézirard V, Pernollet JC, Trotier D, et al. Tas1R1-Tas1R3 taste receptor variants in human fungiform papillae. Neurosci Lett. 2009; 451: 217–21. pmid:19146926
- 67. Shigemura N, Shirosaki S, Sanematsu K, Yoshida R, Ninomiya Y. Genetic and molecular basis of individual differences in human umami taste perception. PLoS One. 2009; 4: e6717. pmid:19696921