The intestinal epithelium is a major boundary with the outside world. Epithelial cells lining the surface of the intestinal epithelium are in direct contact with a luminal environment, the composition of which varies dramatically. It has long been recognised that the gut is capable of sensing changes in its luminal content and responding by releasing chemical signals. In 1902, Bayliss and Starling(Reference Bayliss and Starling1) noted that increasing the acidity in the lumen of the small intestine elicited pancreatic secretions, and that this was mediated, not via the nervous system, but by a humoral factor produced by the gut epithelium that they termed secretin.
Indeed, we now know that the nerve endings that transmit signals evoked by changes in the gut luminal contents do not reach the intestinal lumen and that information about the chemical nature of the luminal contents is transmitted to neurons via enteroendocrine cells (EEC) linking the gut, brain and peripheral tissues.
EEC, scattered amongst the cells lining the intestinal epithelium, are pivotal to the chemosensing pathways of the intestinal tract. They are flask-shaped, with the majority having open-type morphology with apically extended processes making direct contact with ingested nutrients and microbial products in the gut lumen. These cells respond to changes in luminal contents by releasing gut hormones into systemic circulation via their basolateral membrane domain. There are at least sixteen discrete cell types that make up the enteroendocrine family (generally named after letters of alphabet) and collectively they produce over twenty different hormones(Reference Helander and Fändriks2). These include cholecystokinin (CCK), peptide YY (PYY) and glucagon-like peptides 1 and 2 (GLP-1, GLP-2). CCK is released by I-cells predominantly located in the proximal intestine, whereas PYY, GLP-1 and GLP-2 are secreted mostly by L-cells residing frequently in the distal gut. However, L-cells have also been identified in the duodenum albeit in lower number than observed in jejunum and ileum(Reference Steinert, Gerspach and Gutmann3). Secretion of CCK, GLP-1 or PYY slows down gastric emptying, as well as reducing appetite and food intake. GLP-1 also functions as an incretin hormone, stimulating insulin secretion from pancreatic β-cells, improving meal-related glycaemia. GLP-2, coproduced with GLP-1, promotes intestinal epithelial cell growth and increased nutrient absorption(Reference Wölnerhanssen, Moran and Burdyga4,Reference Drucker and Yusta5) .
Although it was believed that gut hormone secretion was the result of direct EEC sensing of nutrients in the lumen of the intestine, until recently little was known about the initial molecular recognition events involved in the enteroendocrine luminal sensing.
G protein-coupled receptors and intestinal nutrient sensing
G protein-coupled receptors (GPCR) represent the largest family of cell-surface mediators of signal transduction(Reference Milligan and McGrath6). They are encoded by about 800 different genes in human subjects(Reference Takeda, Kadowaki and Haga7), enabling cells to respond to many diverse sensory inputs. Hence, GPCR are well-established targets for almost half of all therapeutic drugs, yet many are denominated as ‘orphan receptors’ whose physiological agonists remain unknown. As such these receptors have attracted significant attention in terms of continued identification and characterisation, with the recognition that they are potential targets for novel drug discovery. With more recent evidence that nutrient sensing in the gastrointestinal (GI) tract is accomplished by a number of GPCR(Reference Wellendorph, Johansen and Bräuner-Osborne8), the role of these receptors as important nutritional targets is becoming increasingly evident.
Nutrient-sensing GPCR are categorised as either class A or class C. GPCR belonging to class C, such as the calcium-sensing receptor (CaSR) and the taste 1 family receptors (T1R) comprise an N-terminal signal sequence, seven transmembrane domains coupled with a large extracellular domain (Venus flytrap module) and C-terminal cytoplasmic domain. Class A receptors such as fatty acid receptors also have seven transmembrane domains, but lack the Venus flytrap module(Reference Pongkorpsakol, Moonwiriyakit and Muanprasat9). In general, nutrient-sensing GPCR are classified based on their α-subunits and the corresponding downstream signalling pathways they recruit. They are grouped into families Gαs, Gαi, Gαq, Gα12/13, and gustducin. The physiological effects produced by these receptors in response to nutrients are mainly mediated via cyclic adenosine monophosphate (cAMP) and Ca2+ signalling cascades. Gαs stimulates adenylate cyclase leading to an increase in intracellular concentration of cAMP. Gαi inhibits adenylate cyclase resulting in decreased intracellular cAMP. Gαq stimulates phospholipase C resulting in the generation of diacylglycerol and inositol triphosphate, which respectively activate protein kinase C triggering Ca2+ release from intracellular stores. Gα12/13 couples to the activation of the small G-protein Rho. Gustducin, a heterotrimeric G protein and mainly a member of Gαi family, can stimulate phosphodiesterase resulting in cAMP degradation, but in parallel the co-released Gβγ subunits activate phospholipase C-β2 leading to inositol triphosphate-mediated Ca2+ release. The consequent elevation of cytoplasmic Ca2+ activates the Ca2+sensitive transient receptor potential channel M5 triggering membrane depolarisation and opening of voltage-gated Ca2+ channels(Reference Pongkorpsakol, Moonwiriyakit and Muanprasat9,Reference Chandrashekar, Hoon and Ryba10) . It has also been reported that gustducin activation stimulates adenylate cyclase, increasing cAMP directly or indirectly closing basolateral K+ channels and triggering membrane depolarisation(Reference Wong, Ruiz-Avila and Ming11).
Several nutrient-sensing GPCR have been identified in the intestinal epithelium. They are expressed mainly on the apical membrane domain of EEC and are directly activated by a variety of nutrients. These include receptors for glucose, amino acids, peptides, protein hydrolysates, calcium and both long-chain fatty acids and SCFA (Fig. 1)(Reference Reimann, Tolhurst and Gribble12). Nutrient sensing initiates a cascade of events involving hormonal and neural pathways. This culminates in functional responses that ultimately regulate vital physiological processes including food intake (appetite and satiety), nutrient digestion and absorption, intestinal barrier function, gut motility, and insulin secretion.
A schematic diagram of an enteroendocrine cell with luminal-nutrient sensing G protein-coupled receptors (GPR) and downstream signalling pathways. Taken from Reimann et al. (2012) with permission from Cell Press. CaSR, calcium-sensing receptor; FFAR, free fatty acid receptor; TGR, G-protein-coupled bile acid receptor, GPBAR1; T1R, taste 1 receptor; PLC, phospholipase C; AC, adenylate cyclase; TRPM5, transient receptor potential cation channel subfamily M member 5; IP, inositol phosphate; PKC, protein kinase C; Epac2, exchange protein directly activated by cAMP 2; PKA, Protein kinase A.
This review focuses on GPCR responsive to digestive products of macronutrients.
Carbohydrates
One of the primary functions of carbohydrates is to provide body energy. They comprise sugars, digestible polysaccharides (such as starch) and non-digestible carbohydrates consisting of plant-based fibres and non-starch polysaccharides. In the small intestine, digestible polysaccharides are hydrolysed by pancreatic amylase and brush border membrane disaccharidases to constituent monosaccharides, glucose, galactose and fructose(Reference Shirazi-Beechey13). Non-digestible carbohydrates, which escape digestion in the small intestine, reach the large intestine where they are fermented by gut microbiota, predominantly to SCFA.
Intestinal glucose (sweet) sensing
Glucose is an effective inducer of secretion of gut hormones such as GLP-1, GLP-2 and glucose-dependent insulinotropic peptide (GIP). It has been shown that there is a much greater insulin secretion from the pancreas after orally-ingested glucose than from intravenous injection of the same amount of glucose (the incretin effect)(Reference Creutzfeldt14), inferring the presence of an intestinal luminal glucose sensor responsible for glucose-induced gut peptide release.
In 2005, we reported, for the first time, that the heterodimeric sweet taste receptor T1R2-T1R3, previously characterised in the lingual epithelium, is expressed in gut EEC, and proposed that it acts as the intestinal glucose sensor(Reference Dyer, Salmon and Zibrik15).
Further work demonstrated that all signalling elements involved in sweet taste transduction in the gustatory buds of the tongue, T1R2-T1R3, phospholipase C-β2, transient receptor potential channel M5, α-gustducin and other associated signalling elements, are co-expressed in both L- and K-EEC in human and mouse intestine(Reference Jang, Kokrashvili and Theodorakis16,Reference Margolskee, Dyer and Kokrashvili17) . In mice in which the genes encoding for α-gustducin and T1R3 were deleted, there was a failure to secrete GLP-1 in response to luminal glucose(Reference Jang, Kokrashvili and Theodorakis16,Reference Margolskee, Dyer and Kokrashvili17) . These knockout mice also had abnormal insulin response and prolonged elevation of postprandial blood glucose, indicating that the sweet receptor expressed in intestinal L-cells coupled to α-gustducin sense luminal glucose leading to the secretion of GLP-1(Reference Jang, Kokrashvili and Theodorakis16,Reference Kokrashvili, Mosinger and Margolskee18) . More recent work(Reference Moran, Al-Rammahi and Batchelor19) has confirmed and extended these studies to demonstrate that in mouse small intestine, T1R2, T1R3, α-gustducin and GLP-2 are co-expressed in the same L-EEC and that mouse intestine secretes GLP-2 in response to glucose(Reference Moran, Al-Rammahi and Batchelor19). Moreover, this glucose-induced GLP-2 release was inhibited by gurmarin (a specific inhibitor of mouse T1R3)(Reference Imoto, Miyasaka and Ishima20,Reference Ninomyia and Imoto21) . Furthermore, the non-nutritive sweetener, sucralose, also induced GLP-2 release from mouse small intestine, which was again inhibited by gurmarin. However, the sweetener aspartame, which does not activate mouse T1R2-T1R3(Reference Li, Staszewski and Xu22), did not induce GLP-2 release, supporting the conclusion that the T1R2-T1R3 receptor, expressed in L-cells, senses luminal glucose and sweeteners to secrete GLP-2. A number of studies(Reference Steinert, Gerspach and Gutmann3,Reference Young, Sutherland and Pezos23,Reference Gerspach, Steinert and Schönenberger24) confirming the findings of previous reports(Reference Jang, Kokrashvili and Theodorakis16,Reference Margolskee, Dyer and Kokrashvili17) have demonstrated that transcripts for T1R2, T1R3, α-gustducin, transient receptor potential channel M5 and GLP-1 are expressed in the mucosa of the human proximal intestine. Young et al.(Reference Young, Sutherland and Pezos23) also reported that the expression of T1R2, at mRNA level, was reduced in the intestine of diabetic subjects with higher fasting blood glucose concentration(Reference Young, Sutherland and Pezos23). The magnitude of GLP-1, GLP-2 and GIP secretion has been reported to be diminished in patients with type 2 diabetes(Reference Sadry and Drucker25). A recent study has also shown that the number of EEC, including L-cells, is reduced significantly in the intestine of morbidly obese and diabetic individuals with type 2 diabetes compared to that in healthy controls(Reference Wölnerhanssen, Moran and Burdyga4). Thus, the reduction in T1R2 transcript level observed in diabetics(Reference Young, Sutherland and Pezos23) may be due to a reduced number of EEC expressing T1R2 and other signalling elements required for glucose-induced GLP-1 secretion. Moreover, it has been demonstrated that the intragastric administration of glucose, in healthy subjects, resulted in the secretion of GLP-1 and PYY, which was significantly reduced when lactisole, the specific inhibitor of human T1R3(Reference Winnig, Bufe and Meyerhof26) was co-administered(Reference Steinert, Gerspach and Gutmann3,Reference Gerspach, Steinert and Schönenberger24) . They have concluded that in the human intestine, T1R2-T1R3 is involved in glucose-induced secretion of GLP-1 and PYY, with potential consequences for reducing food intake, decreasing gut motility and increasing insulin secretion (the latter in response to GLP-1).
Mechanisms underlying intestinal sweet sensing and glucose transport regulation
One important manifestation of intestinal glucose sensing by T1R2-T1R3, expressed in L-cells, is the regulation of intestinal glucose transport.
The major route for transport of dietary glucose from the lumen of the intestine into absorptive enterocytes is via the brush border membrane protein, the sodium glucose co-transporter isoform 1 (SGLT1)(Reference Dyer, Wood and Palejwala27,Reference Gorboulev, Schürmann and Vallon28) . Absorption of glucose by SGLT1 also activates electrolyte (NaCl) and water absorption, the route used for oral rehydration therapy(Reference Hirschhorn and Greenough29–Reference Moran, Al-Rammahi and Daly31). SGLT1 activity and expression have been shown to be directly regulated by luminal glucose, including metabolisable, non-metabolisable and membrane-impermeable glucose analogues(Reference Ferraris and Diamond32–Reference Dyer, Vayro and King34). Furthermore, the pathway underlying monosaccharide-enhanced SGLT1 expression was via a luminal membrane glucose GPCR(Reference Dyer, Vayro and King34,Reference Dyer, Vayro and Shirazi-Beechey35) .
Recent experimental evidence has demonstrated that T1R2-T1R3 expressed in L-cells senses dietary glucose (and other natural/artificial sweeteners) resulting in the secretion of GLP-2, which then, via a neuro-paracrine pathway involving the enteric nervous system, enhances the half-life of SGLT1 mRNA in neighbouring absorptive enterocytes. This leads to increased activity and expression of SGLT1, and enhanced intestinal glucose absorption(Reference Moran, Al-Rammahi and Batchelor19). Knocking out the genes for T1R2, T1R3 or GLP-2 receptor abolishes the ability of mouse intestine to up-regulate SGLT1 expression and activity in response to luminal glucose or sweeteners(Reference Margolskee, Dyer and Kokrashvili17,Reference Moran, Al-Rammahi and Batchelor19) .
It has been shown that the expression (and activity) of SGLT1 is enhanced in the intestine of human subjects with type 2 diabetes. This increase was shown to be independent of dietary carbohydrate intake level, or any changes in blood glucose or insulin concentration(Reference Dyer, Wood and Palejwala27), and proposed to be due to alterations in the mechanisms and signalling pathways involved in the regulation of SGLT1 activity and expression.
As noted earlier, the total number of EEC, the expression of T1R2 and levels of gut hormones including GLP-1, GLP-2 and GIP are all significantly reduced in the intestine of diabetic individuals(Reference Wölnerhanssen, Moran and Burdyga4,Reference Young, Sutherland and Pezos23,Reference Sadry and Drucker25) . Thus, it appears that in type 2 diabetes, deregulation of intestinal glucose sensing and downstream signalling may play a role in the observed overexpression of intestinal SGLT1.
Therapeutic potential of taste 1 receptor 2 and receptor 3
Post weaning intestinal disorders are major health problems for the young. Weaning-associated diarrhoea, dehydration and nutrient malabsorption result in high levels of mortality in farm animals worldwide. The finding that small concentrations of specific natural/artificial sweeteners are detected by the intestinal T1R2-T1R3 sweet receptor, activating the pathway leading to increased glucose, electrolyte and water absorption (oral rehydration therapy)(Reference Hirschhorn and Greenough29), has attracted worldwide uptake of these additive sweeteners in the diet of weaning animals. This innovation has improved the health and survival rate of young animals through avoidance of intestinal disorders, thereby increasing weight, enhancing immunity and optimising feed utilisation allowing the translation of scientific discoveries to animal health and welfare benefits(Reference Moran, Al-Rammahi and Daly31,Reference Moran, Al-Rammahi and Arora36,Reference Daly, Darby and Hall37) . Modulation of human intestinal T1R2-T1R3 activity may also have applications in human subjects by controlling glucose absorption(Reference Young, Sutherland and Pezos23).
Proteins
Dietary proteins are essential for growth, provision of energy and health maintenance. In the small intestine, proteins are digested by pancreatic and brush border membrane proteases to di-tri-oligopeptides and amino acids. There are these products that likely target EEC stimulating secretion of a range of gut hormones including CCK, GLP-1 and PYY(Reference Caron, Domenger and Dhulster38). The satiety effects associated with high-protein diets can also be mediated by sensing of the amino acid constituents of proteins.
Intestinal sensing of protein hydrolysis products
Amino acid sensing
A number of GPCR have been identified to respond to amino acids. They belong to a sub-group of C class GPCR and include CaSR, the heterodimeric umami receptor T1R1-T1R3, the goldfish 5⋅24 receptor and its mammalian orthologue GPCR6A, and the metabotropic glutamate receptors.
CaSR is a homodimeric receptor that predominantly couples to Gαq, activating phosphatidylinositol-specific phospholipase C and inducing mobilisation of intracellular Ca2+(Reference Conigrave and Brown39). However, it also couples to Gαs, Gαi and Gα12/13(Reference Saidak, Brazier and Kamel40). CaSR is a multimodal sensor for several key nutrients, notably Ca2+ ions and l-amino acids, and is expressed abundantly throughout the GI tract(Reference Conigrave and Brown39,Reference Brennan, Davies and Schepelmann41) . Although it acts as a sensor for Ca2+ in the gut lumen, it is allosterically activated by l-amino acids; responding to aromatic, aliphatic and polar, but not to branched or positively charged, amino acids(Reference Conigrave, Quinn and Brown42). CaSR is highly expressed in gastrin-secreting G-, somatostatin-secreting D-(Reference Haid, Widmayer and Breer43) and CCK-secreting I-cells(Reference Liou, Sei and Zhao44), and has been proposed to facilitate amino acid-induced secretion of these gut hormones. In studies using secretin tumour cell line (STC)-1 cells, it was shown that extracellular presence of l-phenylalanine induced mobilisation of intracellular Ca2+ and CCK secretion which was inhibited with the allosteric CaSR inhibitor NPS2143(Reference Hira, Nakajima and Eto45). Moreover, native intestinal I-cells from mice deficient in CaSR showed impaired l-phenylalanine mediated Ca2+ responses and CCK release(Reference Liou, Sei and Zhao44), indicating that CaSR plays a significant role in the chemosensing of amino acids in the GI tract.
GPRC6A is a Gq/11-coupled receptor widely expressed in human and rodent tissues. Being a promiscuous amino acid sensor, and expressed in the digestive system, it has been proposed to act as a candidate for sensing digested amino acids in the GI tract(Reference Haid, Widmayer and Breer43,Reference Haid, Jordan-Biegger and Widmayer46) . It has been reported by two groups that GPRC6A is involved in l-ornithine-induced GLP-1 release in the intestinal L-cell line GLUTag(Reference Rueda, Harley and Lu47,Reference Oya, Kitaguchi and Pais48) . However, Oya et al.(Reference Oya, Kitaguchi and Pais48) were unable to measure l-ornithine-induced GLP-1 release from mixed primary cultures of mouse small intestine(Reference Oya, Kitaguchi and Pais48). There are equally conflicting results using GPRC6A knockout mouse models. Alamshah et al.(Reference Alamshah, McGavigan and Spreckley49) demonstrated that l-arginine induced secretion of PYY from both wild-type and GPRC6A KO mouse primary colonic L-cells(Reference Alamshah, McGavigan and Spreckley49). Jørgensen & Bräuner-Osborne(Reference Jørgensen and Bräuner-Osborne50) addressing the in vivo relevance of these findings, administered l-ornithine and l-arginine orally to the full locus and exon VI GPRC6A KO mouse models(Reference Jørgensen and Bräuner-Osborne50). Whilst there was an immediate GLP-1 release that diminished over time, there were no overall differences in the ability of KO-mouse models and wild-type mice to secrete GLP-1 in response to these amino acids. The authors concluded that GPRC6A, in vivo, does not play a role in GLP-1 secretion in response to basic l-amino acids(Reference Jørgensen and Bräuner-Osborne50). Further work is required to unravel the precise role of GPRC6A in intestinal chemosensing.
Taste 1 receptor 1 and receptor 3
In taste cells of lingual epithelium, the heterodimeric combination of T1R1 and T1R3, members of the T1R family, has been identified as a broad-spectrum l-amino acid sensor responsible for mediating perception of the savoury umami taste of monosodium glutamate(Reference Li, Staszewski and Xu22,Reference Nelson, Chandrashekar and Hoon51) . In rodents and many other mammalian species, T1R1-T1R3 responds to a wide variety of l-amino acids in the millimolar range. However, the receptor is not activated by l-tryptophan(Reference Chandrashekar, Hoon and Ryba10). The human T1R1-T1R3 complex functions as a much more specific receptor, responding selectively to monosodium glutamate and aspartic acid (as well as to the glutamate analogue l-AP4)(Reference Li, Staszewski and Xu22,Reference Nelson, Chandrashekar and Hoon51,Reference Ikeda52) . The T1R1-T1R3 heterodimer, such as the sweet receptor T1R2-T1R3, is expressed in EEC(Reference Dyer, Salmon and Zibrik15) and is coupled to gustducin for the transmission of intracellular signals(Reference McLaughlin, McKinnon and Margolskee53). Using STC-1 cells and native mouse intestinal tissue, it has been shown that gut expressed T1R1-T1R3 serves as an intestinal l-amino acid sensor modulating amino acid-induced CCK release(Reference Daly, Al-Rammahi and Moran54). Using siRNA to inhibit the expression of T1R1 mRNA and protein in STC-1 cells, it was demonstrated that the inhibition of T1R1 expression had no effect on protein hydrolysate or peptide-induced CCK release, indicating that T1R1-T1R3 is not the intestinal sensor for peptones. However, in T1R1 knockdown STC-1 cells, there was a significant decline in phenylalanine-, leucine- and glutamate-induced CCK release. Conversely, tryptophan-induced CCK secretion was unaffected by inhibition of T1R1 expression, in agreement with tryptophan not being an agonist for T1R1-T1R3(Reference Daly, Al-Rammahi and Moran54).
Thus, both CaSR and T1R1-T1R3 have been recognised as intestinal l-amino acid sensors mediating CCK secretion in response to aromatic amino acids such as l-phenylalanine(Reference Liou, Sei and Zhao44,Reference Daly, Al-Rammahi and Moran54) . Using a range of agonists and antagonists of CaSR and T1R1-T1R3, it has been demonstrated that CaSR is an intestinal l-amino acid receptor specifically sensing aromatic amino acids, while T1R1-T1R3 responds to a broad spectrum of l-amino acids provoking CCK secretion from intestinal endocrine I-cells(Reference Daly, Al-Rammahi and Moran54).
Peptone receptor
The identity of the cell surface receptor(s) involved in peptone-induced CCK release remains unknown. GPCR92/93 is not a member of the C-class GPCR but has been proposed as a candidate sensor for peptones in STC-1 cells(Reference Choi, Lee and Shiu55). Further work is required to elucidate the peptone-sensing role of this GPCR, if any, in the intestine.
Fats
Fats play an important role in nutrition. As well as providing 30–40 % of total body energy, they also offer essential fatty acids such as linoleic (n-6) and α-linoleic (n-3) acid that cannot be de novo synthesised in the body. Like other macronutrients, fats must first be digested before triggering hormone secretion and are much more effective when administered into the gut lumen than into the circulation. Fat ingestion stimulates the secretion of a number of gut hormones, including CCK, GLP-1 and GIP(Reference Elliott, Morgan and Tredger56). It is reported that long-chain fatty acids inhibit gastric emptying and induce satiety(Reference Raybould57), with SCFA eliciting GLP-1 and PYY secretion(Reference Covasa, Stephens and Toderean58).
Intestinal fatty acid sensing
There are four principal GPCR, FFA1-FFA4, that have been officially classified as members of a NEFA receptor family. FFA1 (GPR40) and FFA4 (GPR120) are activated by both saturated and unsaturated medium-chain (carbon length 8–12) and longer chain (carbon chain length 14–22) fatty acids and are mainly Gαq-coupled(Reference Alvarez-Curto and Milligan59). The supporting evidence that long-chain fatty acid receptors contribute directly to intestinal fatty acid chemosensing is from the findings that their expression in the GI tract is largely limited to the EEC population. The pattern of expression of FFA4 in EEC appears to be similar to that of FFA1. This has highlighted the need for highly selective ligands to probe their functions. A number of preclinical and clinical development programmes have explored the therapeutic potential of agonists of FFA1(Reference Ghislain and Poitout60,Reference Li, Qiu and Geng61) . Indeed, some synthetic agonists of FFA1 have shown the capacity to improve glycaemic control in diabetes(Reference Watterson, Hudson and Ulven62). However, questions remain in terms of sustainability of effects during long-term treatment. There are conflicting experimental evidence relating to the roles of FFA1 and FFA4 and is not clear which one plays a more important role in enteroendocrine fatty acid sensing(Reference Mancini and Poitout63). Despite such concerns, the evidence suggests many positive reasons to promote FFA4 as a promising therapeutic target. They include the potential capacity to regulate GLP-1 secretion from L-cells to promote insulin release and to reduce insulin resistance via anti-inflammatory mechanisms. Thus, efforts have been made in medicinal chemistry for improving the selectivity of ligands between FFA1 and FFA4, and it is proposed that perhaps combined agonists of FFA1 and FFA4 may impart greater anti-diabetic efficacy, than targeting either receptor selectively(Reference Suckow and Briscoe64).
The SCFA receptors FFA2 (GPR43) and FFA3 (GPR41) have been shown, by immunohistochemistry, to be expressed in colonic L-cells. They selectively bind to and are activated by SCFA (carbon chain length 1–6), particularly acetate (C2), propionate (C3) and butyrate (C4). FFA2 responds to C2–C3 fatty acids and couples to Gαi/o as well as Gαq, whereas FFA3 preferentially binds C3–C5 and couples only to Gαi/o. These SCFA are generated predominantly in the distal gut by microbial fermentation of non-digestible carbohydrates, such as fibre and NSP. It has been reported that non-digestible and fermentable dietary fibre and starch, as well as SCFA themselves, enhance GLP-1 secretion(Reference Dagbasi, Lett and Murphy65). Moreover, the SCFA-induced release of GLP-1 from EEC appears to be mediated by FFA2(Reference Hudson, Due-Hansen and Christiansen66).
Although also activated by the same group of SCFA as FFA2, and with a broadly similar expression profile, FFA3 is less well characterised than FFA2. To date, there have been no reports of highly selective synthetic ligands for FFA3 that target the same binding site as SCFA, and as such, detailed understanding of the function of this receptor lags behind(Reference Milligan, Ulven and Murdoch67). There is also significant species orthologue variation in the pharmacology of SCFA receptors in respect to their endogenous ligands, which can be translated to species selectivity of synthetic ligands targeting these receptors.
As alterations in population and diversity of gut microbiota are associated with dysbiosis, there is considerable interest in both prebiotic and probiotic strategies to modulate microbial populations and hence the effectiveness of SCFA production(Reference Pekmez, Dragsted and Brahe68). Thus, the physiological role of SCFA receptors, and their relative importance, compared with other possible targets of prebiotic supplementation remains to be established.
Other NEFA-related receptors
GPR84 is recognised as a receptor responsive to medium-chain fatty acids. However, it is by far the least studied and understood of the currently described receptors for fatty acids. GPR119 is predominantly coupled to Gαs and is responsive to monoacylglycerols, products of TAG hydrolysis. It is proposed that small-molecule ligands of GPR119 increase GLP-1, GIP and insulin release(Reference Chu, Carroll and Alfonso69), however studies have shown these ligands have limited glucose lowering and incretin activity in subjects with type 2 diabetes(Reference Katz, Gambale and Rothenberg70).
Conclusion
The nutrient-sensing GPCR, expressed in EEC, play important roles in sensing the gut luminal environment, transmitting nutrient-evoked signals leading to coordination of various physiological functions such as nutrient digestion, absorption, insulin secretion and food intake.
Targeting the gut-expressed sweet receptor, T1R2-T1R3, with dietary sweetener additives has made a significant contribution to veterinary medicine, through enhancing absorption of glucose, electrolytes and water (oral rehydration therapy) in young animals, thereby preventing weaning-induced intestinal disorders. This strategy may also have applications for the prevention of digestive disorders in premature or newborn human infants. As many GPCR are targets for numerous drugs used in clinical practice, a number of GPCR expressed in gut chemosensory cells are currently under assessment as potential new pathways for treating diabetes and obesity. However, a number of these receptors remain poorly or incompletely characterised. It is envisaged that access to high-quality and well-defined agonists/antagonists, appropriate animal models, closer collaborations between different disciplines and true ligand selectivity/specificity will allow further expansion of this GPCR repertoire. It is predicted that with these basic criteria in place, the potential for much more convincing target validation of nutrient-sensing GPCR will be possible.
With the major role that gut nutrient sensing plays in the control of GI physiology and gut–brain communication, it is expected that further therapeutic successes and nutritional recommendations will arise from research in this area.
Financial Support
The work in the laboratory of S. P. S.B. is supported by Pancosma/ADM and the UK Cystic Fibrosis Trust.
Conflict of Interest
None.
Authorship
The authors had joint responsibility for preparation of this paper.
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