Neurotensin (NT) is a 13 amino acid peptide hormone initially isolated from bovine hypothalamus by Carraway and Leeman 1 and shortly thereafter from bovine small intestine 2. NT is primarily expressed in the brain and in the gastrointestinal (GI) tract but also in a variety of other peripheral organs such as in chromaffin cells of the adrenal medulla and in the pancreas 3,4. The majority of studies investigating NT physiology have focused on brain functions, whereas NT released from enteroendocrine cells has largely been neglected in the last decades. In the brain, NT regulates appetite, opioid-independent analgesia 5, hypothermia 6, and pituitary hormone secretion 7 and modulates several neurotransmitter systems, particularly the dopaminergic system 8. This review will, however, focus on gut-derived NT and, in particular, on NT coacting with the functionally related hormones glucagon-like peptide-1 (GLP-1) and peptide YY (PYY).
Neurotensin processing and degradation
NT (pyroGlu–Leu–Tyr–Glu–Asn–Lys–Pro–Arg–Arg–Pro–Tyr–Ile–Leu–OH) is synthesized as a large 169–170 amino acid long precursor and is highly conserved among species. In addition to NT, the precursor also contains neuromedin N (NN), a six amino acid peptide with strong homology to the C-terminal of NT. NT and NN are located in the C-terminal of the precursor and are separated and flanked by Lys–Arg cleavage sites, which are post-translationally processed by prohormone convertases (PCs) 1, 2, or 5A into active NT and NN dependent on the tissue. Further, a fourth Lys–Arg cleavage site is present in the middle of the precursor, but it is poorly processed likely because of its conformational environment 9. The processing occurs in a tissue-specific manner dependent on the availability of the PCs and their specificity toward the cleavage sites. For example, the major forms found in the brain are NT and NN, a processing pattern generated by PC2, whereas GI tract processing mainly gives rise to NT and the large form of NN, a processing pattern generated by PC1 (Fig. 1) 10–12. Little is known about the large forms of the peptides, but they are speculated to have biological functions, albeit with lower potency, but better plasma stability compared with NT and NN 9,13.
NT is rapidly degraded into inactive metabolites once released into the extracellular space. The major cleavage products in brain and gut tissue have been identified as NT1–8, NT1–10, and NT1–11. Three Zn-metallopeptidases are responsible for cleaving NT: endopeptidase 24.15 generates the NT1–8 fragment, and endopeptidase 24.11 generates the NT1–11 fragment, whereas both endopeptidase 24.11 and 24.16 can give rise to the NT1–10 fragment 14.
NT has a very short half-life in vivo when administered intravenously, in the order of 0.5–1.5 min in rodents and humans 15–18. When NT is incubated in human or rodent plasma or whole blood in vitro, however, the half-life of NT is markedly increased 15,19,20, suggesting that NT degradation occurs in the organs rather than in the circulation. Studies performed in rodents, dogs, sheep and humans suggest that NT clearance occurs in the kidney and intestine, some occurs in the brain, whereas the liver does not seem to contribute considerably to NT degradation 17,19,21–24. Interestingly, the active part of NT, which is recognized by the NT receptors, is the 8–13 C-terminal segment, and thus not one of the degradation products.
NT binds three known receptors, of which the neurotensin receptor 1 (NTS1) and neurotensin receptor 2 (NTS2) are G-protein coupled, whereas the neurotensin receptor 3 (NTS3) is a type 1 single-membrane spanning channel 25.
NTS1 is found both in the brain and in the GI tract 26,27. In the brain, it is widely distributed, and in-situ hybridization, quantitative RT-PCR, receptor autoradiography, and immunohistochemical studies showed high expression in the substantia nigra, ventral tegmental area, islands of Calleja, hypothalamus, diagonal band of Broca, amygdala, septal regions, and thalamic regions 28–32. Similarly, NTS1 is present throughout the GI tract, with the highest expression in the colon measured by quantitative RT-PCR 32. NTS1 binds NT with high affinity in the nanomolar range 26. It increases inositol phosphate, inhibits cyclic AMP formation, activates phospholipase C, and mobilizes intracellular calcium, indicating a preferential coupling to Gq/11, but has also been shown to couple to Gs and Gi/o in some cellular systems 25.
NTS2, which binds NT with lower affinity than NTS1 33, is found primarily in the brain, but NTS2 immunoreactivity has also been reported in the human GI tract 34. In-situ hybridization, immunohistochemistry, and receptor autoradiography showed widespread brain localization with high expression in the olfactory system, bed nucleus of the stria terminalis, cortex, hypothalamus, hippocampus, amygdala, and brainstem regions modulating descending control of nociceptive inputs 35–37. In the GI tract, high expression was observed in parietal cells of the gastric mucosa, which could indicate a role in the inhibition by NT of gastric acid secretion 34.
Although NTS1 is believed to mediate most of the effects of NT, there is evidence that NTS2 is responsible for the analgesic effect of NT 5 and for NT effects on the pancreas 38. NTS2 can couple to diverse signaling pathways and can also signal with constitutive activity in some cell systems 25. Coupling to Gq/11, Gi/o, and G12/13 has been shown to be dependent on the cell system studied and receptor species expressed, but in contrast to NTS1, no clear preference for a G-protein has been shown 25. However, contradictory data have been reported on the ability of NT to activate NTS2. Both agonisms, neutral antagonism and inverse agonism, have been observed depending on the cellular environment by this presumably endogenous ligand for the NTS2 33,39,40.
NTS3, the third well-known NT receptor, is identical to sortilin – a protein involved in receptor sorting and interactions with receptor-associated proteins. NT has a high binding affinity for NTS3 41; however, its exact function in relation to NT remains hypothetical. It has been suggested to be responsible for NT-induced migration of microglial cells 42. Because of its general role in receptor sorting and interaction, and the observation that it can form heterodimers with the other NT receptors 43,44, it has also been speculated to regulate the internalization and recycling process of NTS1 45 as well as modulating second messenger signaling caused by NT acting on NTS1 and NTS2 43. Furthermore, NTS3 may be important for NT effects on the pancreas 44.
Neurotensin in glucose homeostasis
NTS1, NTS2, and NTS3 are expressed in rodent β cell lines as well as in insulin-secreting islets and pancreatic tissue 32,44,46 and, in contrast to other tissues, NTS2 and NTS3 seem to be important for the effects of NT in the pancreas 38. Further, NT immunoreactivity has been found in the endocrine pancreas 47 and thus NT could act on the pancreas both in a paracrine manner, as a locally produced hormone, or in an endocrine manner, through NT released from the GI tract.
Early in-vivo studies showed conflicting results on the role of NT in insulin release, with some studies performed in dogs and in calves suggesting a positive association between increased endogenous or exogenous NT and insulin and glucagon release 48–50, whereas others found that NT produced hypoinsulinemia and hyperglucagonemia and consequently hyperglycemia in rat studies 51,52. More detailed studies carried out in rat islets and insulin-secreting cell lines showed that NT played a dual role in insulin secretion depending on the glucose concentration. NT stimulated insulin release at low glucose concentrations, whereas NT inhibited the glucose-induced insulin release 53,54.
NT levels in the pancreas are increased in diabetic rodents 47,55,56, although plasma concentrations are similar between diabetic rodents and their littermate controls 55. Clinical studies show conflicting results on associations between plasma NT levels and the diabetic state. One study found no association between NT plasma levels and the diabetic state 57, whereas another study found increased pro-NT levels associated with an increased diabetic risk in women, but not in men 58. A third study found increased pro-NT levels in obese and insulin-resistant patients and a positive association between pro-NT levels and the risk of developing obesity and diabetes later in life 59.
In addition to regulating insulin and glucagon release, NT may affect the pancreas through other mechanisms. NT exerts a trophic effect on the pancreas, increasing pancreatic weight, DNA, and protein content 60,61. Furthermore, NT protects β cells from apoptosis in response to cytotoxic agents 44,46. An interesting idea proposed by Mazella et al. 38 is that NT may protect the endocrine pancreas from cell death. As NT is released after ingestion of a high-fat meal, it might serve as a protective agent from hyperlipidemia in a manner similar to GLP-1, protecting the endocrine pancreas from hyperglycemia 38.
Neurotensin and metabolic regulation
It is well established that brain NT regulates appetite as NT can inhibit food intake in rodents when injected into several brain regions including intracerebroventricularly 62–65, hypothalamic areas 62,66, the nucleus of the solitary tract 67, and the dopaminergic nuclei 64,68.
Obese Zucker rats and obese ob/ob mice have lower hypothalamic NT levels compared with their lean littermates 69–71 and treatment with leptin or α-melanocyte-concentrating hormone increases NT expression in a hypothalamic cell line 72, all pointing to a role of NT as an anorexigenic neuropeptide.
Brain NT is also an important mediator of other satiety signals such as the hormone leptin. NT antiserum as well as antagonism or deletion of the NTS1 in mice blunt the anorexigenic action of leptin 73,74. This is at least partly mediated by leptin action on NT-expressing neurons projecting from the lateral hypothalamus to the ventral tegmental area 75. Furthermore, mice with a specific deletion of the leptin receptor in NT-expressing neurons in the lateral hypothalamus show increased appetite, decreased activity, and early-onset obesity probably caused by an impaired control of lateral hypothalamus orexin neurons and dopaminergic neurons 76.
NTS1 knockout (ko) mice have increased appetite and body weight and are nonresponsive to the anorexigenic effects of NT 73,77. In contrast, a more recent study found decreased feeding and increased activity levels in NTS1 ko mice placed on a chow diet, with no overall effect on body weight and body composition 75. However, when placed on a palatable high-fat/high-sucrose diet, NTS1 ko mice ate more and gained more weight than controls and had higher sucrose preference, suggesting that the NTS1 regulates hedonic feeding behavior 75. Interestingly, a recent study found that NT-deficient mice were protected from high-fat diet-induced obesity because of decreased absorption of fat from the intestine 59.
Two studies have tested brain-penetrating NT analogues with improved plasma stability in relation to food intake and body weight in a chronic setting. The NT analogue NT69L decreased food intake and body weight gain dose dependently in chow-fed Sprague–Dawley rats and obese hyperphagic Zucker rats during testing periods ranging from 15 to 38 days 78. Similarly, the NTS1 agonist PD149163 decreased food intake and body weight in Brown Norway rats and obese ob/ob mice during a 10-day test period 79. Thus, there is ample evidence for a role of brain NT and brain-penetrating stable NT analogues in regulating energy metabolism, but the role of peripherally derived NT is less well established. Some studies find that peripheral NT can inhibit food intake acutely both in lean 63,80,81 and in obese rodents 80, but others do not 66. Peripherally injected NT has only been tested chronically in one study in mice where tachyphylaxis was observed 63. This could be because of internalization and degradation of the NTS1 as described in in-vitro studies upon multiple agonist stimulations 45,82.
Neurotensin in the gastrointestinal tract
NT is present throughout the GI tract with high levels in the ileum in pigs, rats, and humans and high levels in ileum and the proximal colon in mice 83–85. NT release is stimulated by endogenous fluids such as bile acids, gastric acid, and pancreatic juice 86,87 and by the ingestion of food, particularly fat 88,89, and to a lesser extent protein and carbohydrate 90. NT is released rapidly after food ingestion both through a direct contact of NT secreting cells to nutrients 86,91 as well as through feed-forward loops from the proximal GI tract including both neural and endocrine signals 88,91.
Once released, NT promotes absorptive processes through several mechanisms: NT increases ileal blood flow 89, pancreatic amylase secretion 60, and hepatic bile secretion 92. NT’s effect on bile acid secretion is mediated both directly by stimulating the contraction of the gall bladder and bile duct 93 and through reabsorption of bile in the ileum, thus upregulating enterohepatic bile recycling 94. In addition, NT stimulates electrolyte and fluid secretion 95,96 and regulates GI tract motility by slowing the transit time in the stomach and the small intestine and promoting movements that optimize absorption 97.
Because of the very short in-vivo half-life of NT, gut-derived NT was initially speculated to only work in a paracrine manner, but has been detected in plasma with increases in response to nutrient intake in humans 90 and been suggested to act as an endocrine hormone on for example, the pancreas in rats 94. Whether NT derived from the intestine can act as an endocrine hormone on the brain has similarly been questioned mainly because of the divergent effects observed after central and peripheral NT administration 98. We have recently observed that intraperitoneal administration of a high dose of NT increases the expression of c-Fos, a marker of neuronal activation, in brain areas with a leaky blood–brain barrier such as the arcuate nucleus and area postrema in mice (In press, Endocrinology). A recent study has directly examined the ability of NT to access the brain in mice and found NT capable of crossing the blood–brain barrier both from plasma to the brain and from the brain to plasma 19. Thus, the lack of centrally mediated effects upon peripheral administration of NT in some studies more likely reflects insufficiently high NT concentrations rather than incomplete penetration of the blood–brain barrier.
Coexpression with other hormones
NT was originally described to be present in enteroendocrine N cells 99. However, the ‘one cell one hormone’ dogma has been challenged and it is now established that many enteroendocrine cells coexpress a variety of different peptides in addition to the well-described coexpression of GLP-1 and PYY in L cells 100–104. It is believed that cells expressing one or more hormones derive from a common precursor cell line, which specializes into expressing mainly one or a few hormones, dependent on a temporal-specific and spatial-specific expression pattern of different transcription factors 105. Coexpression of gut hormones has been observed even in mature villi cells and is fairly common – for example, one mouse study found that out of all cholecystokinin (CCK)-expressing cells analyzed, 44% expressed at least two distinct peptides and 11% expressed at least three different peptides 100. Thus, NT is mainly expressed in N cells, but has been shown to be coexpressed with several other hormones such as GLP-1, PYY, and CCK 85,100–102,104,105. These hormones are also functionally related in for example their inhibition of appetite and gastric emptying; however, in relation to their expression pattern throughout the intestine, NT is likely most related to PYY and GLP-1, with all three hormones being abundant in the distal small intestine 85.
Coexpression between NT and either GLP-1 or PYY has been estimated to be around 15% of the cells positive for any of the three hormones in the rat small intestine 85. Another study showed that NT colocalized with PYY in ∼50% of analyzed NT cells in the proximal mouse colon 106. We recently showed that cells mono-labeled for NT ranged from 20 to 40% along the crypt–villus axis in mouse ileal tissue 101. The degree of colocalization differed along the crypt–villus axis, with ∼20% of NT cells in the top villus costaining for PYY, 10% costaining with GLP-1, and 35% costaining for both PYY and GLP-1. A similar pattern was found in the lower villus, except a larger fraction of NT cells costained with GLP-1 compared with PYY. Finally, in the crypts, little colocalization of all three hormones was observed, whereas NT costained with GLP-1 to a large degree and only minor costaining with PYY was observed 101. NT was preferentially located in the upper villi, whereas PYY was mainly found in the middle villi and GLP-1 was predominantly found in the crypts (Fig. 2) 101. In the proximal mouse colon, NT has similarly predominantly been found in the surface epithelial cuff, whereas GLP-1 was most abundant in the lower crypt 106. This organization along the crypt–villus axis combined with the large degree of colocalization between NT, GLP-1, and PYY indicates that NT-expressing, PYY-expressing, and GLP-1-expressing cells derive from a common precursor cell line predominantly expressing GLP-1 in the crypts and increasingly starting to express NT and PYY as the cells move up the crypt–villus axis before being shed at the top villus. In support of this, transgenic mice with the diphtheria toxin receptor expressed under the proglucagon promoter have GLP-1-expressing, NT-expressing, and PYY-expressing cells ablated upon administration of the diphtheria toxin 101. This indicates that PYY and NT cells also express the proglucagon transcript. Furthermore, the first cells that reappeared were GLP-1 positive, whereas NT-positive and PYY-positive cells were only observed at later time points after ablation in concordance with the preferential expression of GLP-1 in the crypts and NT and PYY in the villi and thus in more mature cells 101. Despite this large degree of coexpression between NT and PYY and GLP-1, NT was found in separate secretory granules to PYY and GLP-1 101.
Finally, in addition to being colocalized, NT has also been shown to be co-released with PYY and GLP-1 in studies using perfused rat small intestine models and in mouse colonic crypt cultures in response to an array of neuropeptide, hormonal, and physiological metabolite stimuli 85,101. In isolated perfused rat small intestine, neuromedin C and glucose-dependent insulinotropic peptide (GIP) induced a fast and parallel release of NT, GLP-1, and PYY 101. Neuromedin C caused the largest release of NT on a molar basis compared with PYY and GLP-1, and GIP induced a larger release of NT and GLP-1 compared with PYY 101. In colonic crypt cultures, neuromedin C as well as agonists for the bile acid receptor TGR5, the 2-monoacylglycerol receptor GPR119, and the long-chain fatty acid receptor GPR40 stimulated NT, GLP-1, and PYY in a very similar pattern, with the TGR5 agonist being the most potent stimulator and the rank order of the different stimuli being identical for all three hormones 101. Although the secretion pattern in rats is very similar for NT, PYY, and GLP-1 in the distal small intestine, PYY release from the proximal small intestine is negligible, whereas both NT and GLP-1 are released from the proximal small intestine 85 and, in this respect, NT may be more related to GLP-1 than PYY. It is, however, not known whether this cosecretion occurs at a cellular level or whether it is purely physiological.
Coactions with glucagon-like peptide-1 and peptide YY
Additive or synergistic effects have been described previously between PYY and GLP-1 in relation to inhibition of food intake both in rodents and in humans 107–109, but not in the regulation of energy expenditure and glucose homeostasis 107,110. It is evident, from the above-mentioned studies, that NT is also closely intertwined with GLP-1 and PYY in relation to expression and release and that these hormones are functionally related. We recently found that NT acted synergistically with GLP-1 in inhibiting the intake of a palatable liquid diet and in slowing gastric emptying, but not during an oral glucose tolerance test. Further, NT and PYY inhibited gastric emptying in an additive manner (Fig. 3) 101.
Whether the synergistic and additive effects observed are a result of the hormones acting on the same cell type or on different cell types in the same target organ or through divergent overall mechanisms is not known and is further discussed below.
Effects on gastric emptying
Motility and emptying of the stomach are primarily controlled by the vagovagal reflex circuitry, which is integrated and controlled by the dorsal–vagal complex in the brainstem 111. Gut hormones modulating gastric emptying could either activate vagal afferents or act directly on targets in the brain controlling efferent vagal projections from the dorsal motor nucleus of the vagus (DMX) or by a direct paracrine action on the stomach. GLP-1, PYY, and NT receptors are present on vagal afferents in rats 112–114 and GLP-1 and PYY can modulate vagal afferent firing 114,115, suggesting that vagal afferents may be important for the inhibition of gastric emptying. There is, however, also evidence pointing to a direct role of these hormones in the dorsal–vagal complex. GLP-1, PYY, and NT receptors are expressed in the dorsal–vagal complex in rats 116–118. Further, GLP-1 and PYY directly regulate gastric motility when microinjected into the dorsal–vagal complex through a modulation of vagal efferent activity in rats 119,120. GLP-1 apparently acts by stimulating a nonadrenergic, noncholinergic vagal pathway inhibiting gastric tone rather than inhibiting cholinergic parasympathetic output 119, whereas PYY seems to target both pathways through Y2 receptor-dependent mechanisms 120,121.
To establish the relative contribution of the vagus nerve toward hormonal inhibition of gastric emptying, studies have been carried out using either full vagotomy or a selective afferent nerve vagotomy. Whereas the efferent vagal projections seem important for NT, PYY, and GLP-1 in their regulation of gastric emptying 120,122–124, discrepant results have been obtained on the contribution of the afferent nerve on the inhibition by GLP-1 of gastric emptying 125,126.
Finally, one study suggested that PYY can act directly on guinea pig stomach muscles to relax them 127, whereas a similar mechanism could not be found for GLP-1 in pigs 123 and has not been explored for NT. In conclusion, NT, PYY, and GLP-1 act synergistically or additively in their inhibition of gastric emptying, which is likely mediated through similar mechanisms involving the vagovagal reflex circuitry.
Effects on food intake
Because of the short half-life of many gut hormones once released, the afferent vagus nerve has also been suggested to be important for hormone signaling to the brain in the regulation of food intake. In relation to GLP-1-mediated decreased food intake, the relative contribution of the vagus nerve has, however, given rise to discrepant results, with some studies finding it important in humans and rodents, whereas another study performed in rats did not find the vagus nerve necessary for GLP-1-mediated decreased food intake 124,128,129. In support of a direct humoral action of GLP-1 in the brain are studies showing that peripherally administered GLP-1 can bind receptors in the area postrema and subfornical area in the rat 130 and induce c-Fos expression, a marker of neuronal activation, in a number of brainstem nuclei including the area postrema 131. Although GLP-1 receptors are present in the arcuate nucleus of the hypothalamus, these receptors do not seem to contribute toward GLP-1-induced satiety 132, although this site may be important for the action of long-acting GLP-1 analogues such as liraglutide 133.
The brainstem may similarly be important for NT effects on food intake as NT can modulate neuronal firing in the canine area postrema 134. Results from our group suggest that the vagus nerve is not necessary, but may contribute toward peripherally administered NT-mediated anorexia in mice (In press Endocrinology). In support of this, we found that peripherally administered NT increased c-Fos expression in mouse brainstem regions including the area postrema, similar to GLP-1 131. We also observed increased c-Fos expression in the arcuate nucleus after NT administration and increased expression of the anorexigenic neuropeptide proopiomelanocortin following NT treatment. In contrast, NT has previously been shown not to increase α-melanocyte-stimulating hormone release from hypothalamic explants 63; thus, the exact contribution of these neurons needs to be determined.
In conclusion, NT and GLP-1 can act synergistically in relation to inhibition of palatable food intake, which may be through a direct action on central receptors in the brainstem, but likely also involve vagal afferents projecting to the brainstem.
Effects on glucose homeostasis
Both NT and GLP-1 can directly act on pancreatic β cells to control insulin release 38,135. However, opposing actions between GLP-1 and NT on insulin levels have been observed during oral glucose tolerance tests in mice 101, and NT has been shown to delay the glucose excursion, likely through its inhibitory effects on gastric emptying 101. The hypoinsulinemic and thus opposite effect to GLP-1 of NT is not surprising considering previous studies showing that NT inhibits the glucose-induced insulin release 53,54. Therefore, NT and GLP-1 seem to, if anything, counteract each other in the control of glucose homeostasis. It is noteworthy, however, that GLP-1 almost completely abolished the hypoinsulinemic effect as well as effects on glucose excursions of NT when the hormones were coadministered 101.
Additivity/synergism on a molecular level
As mentioned above, it is not known whether NT acts on the same cellular targets as GLP-1 and PYY in the inhibition of gastric emptying and food intake. The GLP-1 receptor is coupled to Gs and the NTS1 couple to Gq/11, signaling pathways that are known to act in synergy in some intracellular effector pathways 136,137. Peripheral PYY3–36 mainly exerts its effects through the Y2 receptor, which couples to Gi. Signaling through Gi can activate other signaling pathways such as through βγ signaling, which could act in synergy with Gq-mediated signaling 137. Thus, theoretically, it is plausible that the functional synergistic/additive effects observed between NT and GLP-1 and PYY 101 could occur at the cellular level; however, further studies are needed to determine whether this is the case or whether the hormones act through separate pathways on the same target organ.
Past pharmacotherapy for the treatment of obesity has largely been withdrawn because of unacceptable side effects 138. Currently, a few anti-obesity agents are on the market including the GLP-1 analogue liraglutide but these only produce a modest weight loss of up to around 10% 139. The ineffectiveness of current pharmacotherapy, which generally targets only one physiological pathway, is not surprising considering the complexity and redundancy in energy balance regulation. The most effective obesity intervention today that produces a sustained and considerable weight loss is bariatric surgery, but it is associated with risks such as micronutrient deficiency, hypoglycemia, weight regain, and, in rare cases, mortality 140. Considerable effort has been expended to identify the physiological mechanisms of the metabolic improvements following bariatric surgery and altered gut hormone signaling through the gut–brain axis has received considerable attention as a number of anorexigenic hormones including GLP-1, PYY, and NT are increased in the circulation following surgery 141–143. The role of single hormones in mediating some of the beneficial effects of bariatric surgery – that is, weight loss and reduced appetite has, however, not yielded convincing results 141. It is more likely that the combined effects of enteroendocrine and other factors including the gut microbiome and bile acids are responsible for the weight loss and metabolic improvements 141. In line with this, recent focus in the development of antiobesity agents targets more than one signaling pathway often involving one or more gut hormones.
The success of already approved GLP-1-based treatments for diabetes and obesity has led to interest in developing combinatorial approaches with GLP-1 to enhance the rather modest weight loss observed with GLP-1 analogues alone. For example, the combination of GLP-1 analogues with either leptin, PYY, amylin, CCK, oxyntomodulin, GIP, or glucagon shows additive or synergistic body weight-lowering potential and improves a range of other metabolic parameters such as appetite, insulin sensitivity, glycemic control, energy expenditure, and adiposity in preclinical studies 144. In particular, the combination of GLP-1, GIP, and glucagon in a triagonist has been a very successful approach rivaling the weight loss observed after bariatric surgery in rodents 145, and currently, several combinatorial approaches based on GLP-1 receptor agonists and glucagon are in clinical development 144.
The recently demonstrated synergy between NT and GLP-1 in the inhibition of food intake 101 suggests that combining GLP-1 analogues with NT may also be a viable approach as an antiobesity intervention. In particular, the combination of stable NT analogues with GLP-1 analogues could be speculated to show enhanced effects on appetite and body weight. Stable NT analogues have previously been shown to inhibit appetite and lower body weight in chronic settings 78,79. Although GLP-1 and NT may have opposing physiological actions on glucoregulatory control during situations where the glucose concentration is high, the effect of GLP-1 on glucose metabolism would likely buffer the hyperglycemic effects exerted by NT, as has also been observed during GLP-1 and glucagon coadministration 146. Indeed, when NT and the GLP-1 receptor agonist liraglutide were coadministered during an oral glucose tolerance test, liraglutide, even at the low dose used, counteracted the hypoinsulinemic effect of NT as well as the delay in the glucose excursion caused by NT 101.
A crucial next step is the development of NT formulations with improved pharmacokinetic profiles that can be used in conjunction with GLP-1 analogues and potentially also other gut hormones. A better understanding of the mechanisms and molecular events responsible for the observed synergistic and additive effects between different gut hormones is also essential for the design of optimal combinatorial approaches.
In conclusion, NT is a metabolically active gut hormone with important effects related to energy balance regulation. The synergistic action between different hormones has been exploited recently in the development of novel antiobesity agents and shows promising results 144. The success of this approach, targeting several signaling pathways, is not surprising considering that obesity is a complex disease affecting multiple organs. The recently observed synergy between NT and GLP-1 and PYY 101 suggests that NT should also be considered and included among the anorexigenic hormones currently being studied for their combined effects on energy balance regulation.
This study was funded by the Novo Nordisk Foundation Center for Basic Metabolic Research.
Conflicts of interest
There are no conflicts of interest.
1. Carraway R, Leeman SE. The isolation of a new hypotensive peptide, neurotensin
, from bovine hypothalami. J Biol Chem 1973; 248:6854–6861.
2. Kitabgi P, Carraway R, Leeman SE. Isolation of a tridecapeptide from bovine intestinal tissue and its partial characterization as neurotensin
. J Biol Chem 1976; 251:7053–7058.
3. Goedert M, Emson PC. The regional distribution of neurotensin
-like immunoreactivity in central and peripheral tissues of the cat. Brain Res 1983; 272:291–297.
4. Goedert M, Sturmey N, Williams BJ, Emson PC. The comparative distribution of xenopsin- and neurotensin
-like immunoreactivity in Xenopus laevis
and rat tissues. Brain Res 1984; 308:273–280.
5. Dubuc I, Sarret P, Labbé-Jullié C, Botto JM, Honoré E, Bourdel E, et al.. Identification of the receptor subtype involved in the analgesic effect of neurotensin
. J Neurosci 1999; 19:503–510.
6. Popp E, Schneider A, Vogel P, Teschendorf P, Böttiger BW. Time course of the hypothermic response to continuously administered neurotensin
. Neuropeptides 2007; 41:349–354.
7. Stolakis V, Kalafatakis K, Botis J, Zarros A, Liapi C. The regulatory role of neurotensin
on the hypothalamic–anterior pituitary axons: emphasis on the control of thyroid-related functions. Neuropeptides 2010; 44:1–7.
8. Binder EB, Kinkead B, Owens MJ, Nemeroff CB. Neurotensin
and dopamine interactions. Pharmacol Rev 2001; 53:453–486.
9. Kitabgi P. Differential processing of pro-neurotensin
/neuromedin N and relationship to pro-hormone convertases. Peptides 2006; 27:2508–2514.
10. Shaw C, McKay D, Johnston CF, Halton DW, Fairweather I, Kitabgi P, Buchanan KD. Differential processing of the neurotensin
/neuromedin N precursor in the mouse. Peptides 1990; 11:227–235.
11. Carraway RE, Mitra SP. Differential processing of neurotensin
/neuromedin N precursor(s) in canine brain and intestine. J Biol Chem 1990; 265:8627–8631.
12. Rovère C, Barbero P, Kitabgi P. Evidence that PC2 is the endogenous pro-neurotensin
convertase in rMTC 6-23 cells and that PC1- and PC2-transfected PC12 cells differentially process pro-neurotensin
. J Biol Chem 1996; 271:11368–11375.
13. Friry C, Feliciangeli S, Richard F, Kitabgi P, Rovere C. Production of recombinant large proneurotensin/neuromedin N-derived peptides and characterization of their binding and biological activity. Biochem Biophys Res Commun 2002; 290:1161–1168.
14. Kitabgi P. Inactivation of neurotensin
and neuromedin N by Zn metallopeptidases. Peptides 2006; 27:2515–2522.
15. Aronin N, Carraway RE, Ferris CF, Hammer RA, Leeman SE. The stability and metabolism of intravenously administered neurotensin
in the rat. Peptides 1982; 3:637–642.
16. Lee YC, Allen JM, Uttenthal LO, Walker MC, Shemilt J, Gill SS, Bloom SR. The metabolism of intravenously infused neurotensin
in man and its chromatographic characterization in human plasma. J Clin Endocrinol Metab 1984; 59:45–50.
17. Shulkes A, Bijaphala S, Dawborn JK, Fletcher DR, Hardy KJ. Metabolism of neurotensin
and pancreatic polypeptide in man: role of the kidney and plasma factors. J Clin Endocrinol Metab 1984; 58:873–879.
18. Holst Pedersen J, Fahrenkrug J. Neurotensin
-like immunoreactivities in human plasma: feeding responses and metabolism. Peptides 1986; 7:15–20.
19. Gevaert B, Wynendaele E, Stalmans S, Bracke N, D’Hondt M, Smolders I, et al.. Blood–brain barrier transport kinetics of the neuromedin peptides NMU, NMN, NMB and NT. Neuropharmacology 2016; 107:460–470.
20. Lee YC, Uttenthal LO, Smith HA, Bloom SR. In vitro degradation of neurotensin
in human plasma. Peptides 1986; 7:383–387.
21. Shulkes A, Englin I, Read D, Hardy KJ. Neurotensin
metabolism in the rat: contribution of the kidney. Peptides 1987; 8:961–965.
22. Shulkes A, Fletcher DR, Hardy KJ. Organ and plasma metabolism of neurotensin
in sheep. Am J Physiol 1983; 245 (Pt 1):E457–E462.
23. Checler F, Kostolanska B, Fox JA. In vivo inactivation of neurotensin
in dog ileum: major involvement of endopeptidase 24-11. J Pharmacol Exp Ther 1988; 244:1040–1044.
24. Barelli H, Fox-Threlkeld JE, Dive V, Daniel EE, Vincent JP, Checler F. Role of endopeptidase 220.127.116.11 in the catabolism of neurotensin
, in vivo, in the vascularly perfused dog ileum. Br J Pharmacol 1994; 112:127–132.
25. Pelaprat D. Interactions between neurotensin
receptors and G proteins. Peptides 2006; 27:2476–2487.
26. Tanaka K, Masu M, Nakanishi S. Structure and functional expression of the cloned rat neurotensin
receptor. Neuron 1990; 4:847–854.
27. Vita N, Laurent P, Lefort S, Chalon P, Dumont X, Kaghad M, et al.. Cloning and expression of a complementary DNA encoding a high affinity human neurotensin
receptor. FEBS Lett 1993; 317 :139–142.
28. Elde R, Schalling M, Ceccatelli S, Nakanishi S, Hökfelt T. Localization of neuropeptide receptor mRNA in rat brain: initial observations using probes for neurotensin
and substance P receptors. Neurosci Lett 1990; 120:134–138.
29. Alexander MJ, Leeman SE. Widespread expression in adult rat forebrain of mRNA encoding high-affinity neurotensin
receptor. J Comp Neurol 1998; 402:475–500.
30. Boudin H, Pélaprat D, Rostène W, Beaudet A. Cellular distribution of neurotensin
receptors in rat brain: immunohistochemical study using an antipeptide antibody against the cloned high affinity receptor. J Comp Neurol 1996; 373:76–89.
31. Nicot A, Rostene W, Berod A. Neurotensin
receptor expression in the rat forebrain and midbrain: a combined analysis by in situ hybridization and receptor autoradiography. J Comp Neurol 1994; 341:407–419.
32. Méndez M, Souazé F, Nagano M, Kelly PA, Rostène W, Forgez P. High affinity neurotensin
receptor mRNA distribution in rat brain and peripheral tissues. Analysis by quantitative RT-PCR. J Mol Neurosci 1997; 9:93–102.
33. Mazella J, Botto JM, Guillemare E, Coppola T, Sarret P, Vincent JP. Structure, functional expression, and cerebral localization of the levocabastine-sensitive neurotensin
/neuromedin N receptor from mouse brain. J Neurosci 1996; 16:5613–5620.
34. Schulz S, Röcken C, Ebert MP, Schulz S. Immunocytochemical identification of low-affinity NTS2 neurotensin
receptors in parietal cells of human gastric mucosa. J Endocrinol 2006; 191:121–128.
35. Sarret P, Beaudet A, Vincent JP, Mazella J. Regional and cellular distribution of low affinity neurotensin
receptor mRNA in adult and developing mouse brain. J Comp Neurol 1998; 394:344–356.
36. Asselin ML, Dubuc I, Coquerel A, Costentin J. Localization of neurotensin
NTS2 receptors in rat brain, using. Neuroreport 2001; 12:1087–1091.
37. Sarret P, Perron A, Stroh T, Beaudet A. Immunohistochemical distribution of NTS2 neurotensin
receptors in the rat central nervous system. J Comp Neurol 2003; 461:520–538.
38. Mazella J, Béraud-Dufour S, Devader C, Massa F, Coppola T. Neurotensin
and its receptors in the control of glucose homeostasis. Front Endocrinol (Lausanne) 2012; 3:143.
39. Holst B, Holliday ND, Bach A, Elling CE, Cox HM, Schwartz TW. Common structural basis for constitutive activity of the ghrelin receptor family. J Biol Chem 2004; 279:53806–53817.
40. Richard F, Barroso S, Martinez J, Labbé-Jullié C, Kitabgi P. Agonism, inverse agonism, and neutral antagonism at the constitutively active human neurotensin
receptor 2. Mol Pharmacol 2001; 60:1392–1398.
41. Vincent JP, Mazella J, Kitabgi P. Neurotensin
receptors. Trends Pharmacol Sci 1999; 20:302–309.
42. Martin S, Vincent JP, Mazella J. Involvement of the neurotensin
receptor-3 in the neurotensin
-induced migration of human microglia. J Neurosci 2003; 23:1198–1205.
43. Martin S, Navarro V, Vincent JP, Mazella J. Neurotensin
receptor-1 and -3 complex modulates the cellular signaling of neurotensin
in the HT29 cell line. Gastroenterology 2002; 123:1135–1143.
44. Béraud-Dufour S, Coppola T, Massa F, Mazella J. Neurotensin
receptor-2 and -3 are crucial for the anti-apoptotic effect of neurotensin
on pancreatic beta-TC3 cells. Int J Biochem Cell Biol 2009; 41:2398–2402.
45. Souazé F, Forgez P. Molecular and cellular regulation of neurotensin
receptor under acute and chronic agonist stimulation. Peptides 2006; 27:2493–2501.
46. Coppola T, Béraud-Dufour S, Antoine A, Vincent JP, Mazella J. Neurotensin
protects pancreatic beta cells from apoptosis. Int J Biochem Cell Biol 2008; 40:2296–2302.
47. Fernstrom MH, Mirski MA, Carraway RE, Leeman SE. Immunoreactive neurotensin
levels in pancreas: elevation in diabetic rats and mice. Metabolism 1981; 30:853–855.
48. Kaneto A, Kaneko T, Kajinuma H, Kosaka K. Effects of substance P and neurotensin
infused intrapancreatically on glucagon and insulin secretion. Endocrinology 1978; 102:393–401.
49. Blackburn AM, Bloom SR, Edwards AV. Pancreatic endocrine responses to physiological changes in plasma neurotensin
concentration in the calf. J Physiol 1981; 318:407–412.
50. Blackburn AM, Bloom SR, Edwards AV. Pancreatic endocrine responses to exogenous neurotensin
in the conscious calf. J Physiol 1981; 314:11–21.
51. Brown M, Vale W. Effects of neurotensin
and substance P on plasma insulin, glucagon and glucose levels. Endocrinology 1976; 98:819–822.
52. Carraway RE, Demers LM, Leeman SE. Hyperglycemic effect of neurotensin
, a hypothalamic peptide. Endocrinology 1976; 99:1452–1462.
53. Béraud-Dufour S, Abderrahmani A, Noel J, Brau F, Waeber G, Mazella J, Coppola T. Neurotensin
is a regulator of insulin secretion in pancreatic beta-cells. Int J Biochem Cell Biol 2010; 42:1681–1688.
54. Dolais-Kitabgi J, Kitabgi P, Brazeau P, Freychet P. Effect of neurotensin
on insulin, glucagon, and somatostatin release from isolated pancreatic islets. Endocrinology 1979; 105:256–260.
55. Sheppard MC, Bailey CJ, Flatt PR, Swanston-Flatt SK, Shennan KI. Immunoreactive neurotensin
in spontaneous syndromes of obesity and diabetes in mice. Acta Endocrinol (Copenh) 1985; 108:532–536.
56. Berelowitz M, Frohman LA. Immunoreactive neurotensin
in the pancreas of genetically obese and diabetic mice. A longitudinal study. Diabetes 1983; 32:51–54.
57. Service FJ, Jay JM, Rizza RA, O’Brien PC, Go VL. Neurotensin
in diabetes and obesity. Regul Pept 1986; 14:85–92.
58. Melander O, Maisel AS, Almgren P, Manjer J, Belting M, Hedblad B, et al.. Plasma proneurotensin and incidence of diabetes, cardiovascular disease, breast cancer, and mortality. JAMA 2012; 308:1469–1475.
59. Li J, Song J, Zaytseva YY, Liu Y, Rychahou P, Jiang K, et al.. An obligatory role for neurotensin
in high-fat-diet-induced obesity. Nature 2016; 533:411–415.
60. Wood JG, Hoang HD, Bussjaeger LJ, Solomon TE. Effect of neurotensin
on pancreatic and gastric secretion and growth in rats. Pancreas 1988; 3:332–339.
61. Feurle GE, Müller B, Rix E. Neurotensin
induces hyperplasia of the pancreas and growth of the gastric antrum in rats. Gut 1987; 28 (Suppl):19–23.
62. Hawkins MF, Barkemeyer CA, Tulley RT. Synergistic effects of dopamine agonists and centrally administered neurotensin
on feeding. Pharmacol Biochem Behav 1986; 24:1195–1201.
63. Cooke JH, Patterson M, Patel SR, Smith KL, Ghatei MA, Bloom SR, Murphy KG. Peripheral and central administration of xenin and neurotensin
suppress food intake in rodents. Obesity (Silver Spring) 2009; 17:1135–1143.
64. Hawkins MF. Aphagia in the rat following microinjection of neurotensin
into the ventral tegmental area. Life Sci 1986; 38:2383–2388.
65. Luttinger D, King RA, Sheppard D, Strupp J, Nemeroff CB, Prange AJ Jr. The effect of neurotensin
on food consumption in the rat. Eur J Pharmacol 1982; 81:499–503.
66. Stanley BG, Hoebel BG, Leibowitz SF. Neurotensin
: effects of hypothalamic and intravenous injections on eating and drinking in rats. Peptides 1983; 4:493–500.
67. de Beaurepaire R, Suaudeau C. Anorectic effect of calcitonin, neurotensin
and bombesin infused in the area of the rostral part of the nucleus of the tractus solitarius in the rat. Peptides 1988; 9:729–733.
68. Vaughn AW, Baumeister AA, Hawkins MF, Anticich TG. Intranigral microinjection of neurotensin
suppresses feeding in food deprived rats. Neuropharmacology 1990; 29:957–960.
69. Beck B, Burlet A, Nicolas JP, Burlet C. Hyperphagia in obesity is associated with a central peptidergic dysregulation in rats. J Nutr 1990; 120:806–811.
70. Wilding JP, Gilbey SG, Bailey CJ, Batt RA, Williams G, Ghatei MA, Bloom SR. Increased neuropeptide-Y messenger ribonucleic acid (mRNA) and decreased neurotensin
mRNA in the hypothalamus of the obese (ob/ob) mouse. Endocrinology 1993; 132:1939–1944.
71. Williams G, Cardoso H, Lee YC, Ghatei MA, Flatt PR, Bailey CJ, Bloom SR. Reduced hypothalamic neurotensin
concentrations in the genetically obese diabetic (ob/ob) mouse: possible relationship to obesity. Metabolism 1991; 40:1112–1116.
72. Cui H, Cai F, Belsham DD. Anorexigenic hormones leptin, insulin, and alpha-melanocyte-stimulating hormone directly induce neurotensin
(NT) gene expression in novel NT-expressing cell models. J Neurosci 2005; 25:9497–9506.
73. Kim ER, Leckstrom A, Mizuno TM. Impaired anorectic effect of leptin in neurotensin
receptor 1-deficient mice. Behav Brain Res 2008; 194:66–71.
74. Sahu A, Carraway RE, Wang YP. Evidence that neurotensin
mediates the central effect of leptin on food intake in rat. Brain Res 2001; 888:343–347.
75. Opland D, Sutton A, Woodworth H, Brown J, Bugescu R, Garcia A, et al.. Loss of neurotensin
receptor-1 disrupts the control of the mesolimbic dopamine system by leptin and promotes hedonic feeding and obesity. Mol Metab 2013; 2:423–434.
76. Leinninger GM, Opland DM, Jo YH, Faouzi M, Christensen L, Cappellucci LA, et al.. Leptin action via neurotensin
neurons controls orexin, the mesolimbic dopamine system and energy balance. Cell Metab 2011; 14:313–323.
77. Remaury A, Vita N, Gendreau S, Jung M, Arnone M, Poncelet M, et al.. Targeted inactivation of the neurotensin
type 1 receptor reveals its role in body temperature control and feeding behavior but not in analgesia. Brain Res 2002; 953:63–72.
78. Boules M, Cusack B, Zhao L, Fauq A, McCormick DJ, Richelson E. A novel neurotensin
peptide analog given extracranially decreases food intake and weight in rodents. Brain Res 2000; 865:35–44.
79. Feifel D, Goldenberg J, Melendez G, Shilling PD. The acute and subchronic effects of a brain-penetrating, neurotensin
-1 receptor agonist on feeding, body weight and temperature. Neuropharmacology 2010; 58:195–198.
80. Bailey CJ, Flatt PR. Anorectic effect of fenfluramine, cholecystokinin and neurotensin
in genetically obese (ob/ob) mice. Comp Biochem Physiol A Comp Physiol 1986; 84:451–454.
81. Sandoval SL, Kulkosky PJ. Effects of peripheral neurotensin
on behavior of the rat. Pharmacol Biochem Behav 1992; 41:385–390.
82. Hermans E, Maloteaux JM. Mechanisms of regulation of neurotensin
receptors. Pharmacol Ther 1998; 79:89–104.
83. Wewer Albrechtsen NJ, Kuhre RE, Toräng S, Holst JJ. The intestinal distribution pattern of appetite- and glucose regulatory peptides in mice, rats and pigs. BMC Res Notes 2016; 9:60.
84. Evers BM, Rajaraman S, Chung DH, Townsend CM Jr, Wang X, Graves K, Thompson JC. Differential expression of the neurotensin
gene in the developing rat and human gastrointestinal tract. Am J Physiol 1993; 265 (Pt 1):G482–G490.
85. Svendsen B, Pedersen J, Albrechtsen NJ, Hartmann B, Toräng S, Rehfeld JF, et al.. An analysis of cosecretion and coexpression of gut hormones from male rat proximal and distal small intestine. Endocrinology 2015; 156:847–857.
86. Rökaeus A, Al-Saffar A. The importance of bile and pancreatic juice for fat-induced release of neurotensin
-like immunoreactivity (NTLI) from the small intestine of the rat. Acta Physiol Scand 1983; 119:33–37.
87. Dakka T, Dumoulin V, Chayvialle JA, Cuber JC. Luminal bile salts and neurotensin
release in the isolated vascularly perfused rat jejuno-ileum. Endocrinology 1994; 134:603–607.
88. Drewe J, Mihailovic S, D’Amato M, Beglinger C. Regulation of fat-stimulated neurotensin
secretion in healthy subjects. J Clin Endocrinol Metab 2008; 93:1964–1970.
89. Hammer RA, Matsumoto BK, Blei AT, Pearl G, Ingram H. Local effect of neurotensin
on canine ileal blood flow, and its release by luminal lipid. Scand J Gastroenterol 1988; 23:449–457.
90. Rosell S, Rökaeus A. The effect of ingestion of amino acids, glucose and fat on circulating neurotensin
-like immunoreactivity (NTLI) in man. Acta Physiol Scand 1979; 107:263–267.
91. Miyashita T, Hashimoto T, Gomez G, Townsend CM Jr, Greeley GH Jr, Thompson JC. Neurotensin
secretion in response to intraduodenal and intraileal administration of fat in dogs. Biol Signals 1992; 1:275–281.
92. Gui X, Degolier TF, Duke GE, Carraway RE. Neurotensin
elevates hepatic bile acid secretion in chickens by a mechanism requiring an intact enterohepatic circulation. Comp Biochem Physiol C Toxicol Pharmacol 2000; 127:61–70.
93. Yamasato T, Nakayama S. Effects of neurotensin
on the motility of the isolated gallbladder, bile duct and ampulla in guinea-pigs. Eur J Pharmacol 1988; 148:101–106.
94. Gui X, Carraway RE. Enhancement of jejunal absorption of conjugated bile acid by neurotensin
in rats. Gastroenterology 2001; 120:151–160.
95. Miller RJ, Kachur JF, Field M, Rivier J. Neurohumoral control of ileal electrolyte transport. Ann N Y Acad Sci 1981; 372:571–593.
96. Wiklund B, Liljeqvist L, Rökaeus A. Neurotensin
increases net fluid secretion and transit rate in the small intestine of man. Regul Pept 1984; 8:33–39.
97. Kalafatakis K, Triantafyllou K. Contribution of neurotensin
in the immune and neuroendocrine modulation of normal and abnormal enteric function. Regul Pept 2011; 170 (1–3):7–17.
98. Nemeroff CB, Bissette G, Prange AJ Jr, Loosen PT, Barlow TS, Lipton MA. Neurotensin
: central nervous system effects of a hypothalamic peptide. Brain Res 1977; 128:485–496.
99. Leeman SE, Carraway RE. Neurotensin
: discovery, isolation, characterization, synthesis and possible physiological roles. Ann N Y Acad Sci 1982; 400:1–16.
100. Egerod KL, Engelstoft MS, Grunddal KV, Nohr MK, Secher A, Sakata I, et al.. A major lineage of enteroendocrine cells coexpress CCK, secretin, GIP, GLP-1, PYY, and neurotensin
but not somatostatin. Endocrinology 2012; 153:5782–5795.
101. Grunddal KV, Ratner CF, Svendsen B, Sommer F, Engelstoft MS, Madsen AN, et al.. Neurotensin
is coexpressed, coreleased, and acts together with GLP-1 and PYY in enteroendocrine control of metabolism. Endocrinology 2016; 157:176–194.
102. Habib AM, Richards P, Cairns LS, Rogers GJ, Bannon CA, Parker HE, et al.. Overlap of endocrine hormone expression in the mouse intestine revealed by transcriptional profiling and flow cytometry. Endocrinology 2012; 153:3054–3065.
103. Habib AM, Richards P, Rogers GJ, Reimann F, Gribble FM. Co-localisation and secretion of glucagon-like peptide 1 and peptide YY
from primary cultured human L cells. Diabetologia 2013; 56:1413–1416.
104. Sykaras AG, Demenis C, Cheng L, Pisitkun T, Mclaughlin JT, Fenton RA, Smith CP. Duodenal CCK cells from male mice express multiple hormones including ghrelin. Endocrinology 2014; 155:3339–3351.
105. Engelstoft MS, Egerod KL, Lund ML, Schwartz TW. Enteroendocrine cell types revisited. Curr Opin Pharmacol 2013; 13:912–921.
106. Roth KA, Kim S, Gordon JI. Immunocytochemical studies suggest two pathways for enteroendocrine cell differentiation in the colon. Am J Physiol 1992; 263 (Pt 1):G174–G180.
107. Schmidt JB, Gregersen NT, Pedersen SD, Arentoft JL, Ritz C, Schwartz TW, et al.. Effects of PYY3-36 and GLP-1 on energy intake, energy expenditure, and appetite in overweight men. Am J Physiol Endocrinol Metab 2014; 306:E1248–E1256.
108. Neary NM, Small CJ, Druce MR, Park AJ, Ellis SM, Semjonous NM, et al.. Peptide YY3-36 and glucagon-like peptide-17-36 inhibit food intake additively. Endocrinology 2005; 146:5120–5127.
109. De Silva A, Salem V, Long CJ, Makwana A, Newbould RD, Rabiner EA, et al.. The gut hormones PYY 3-36 and GLP-1 7-36 amide reduce food intake and modulate brain activity in appetite centers in humans. Cell Metab 2011; 14:700–706.
110. Tan TM, Salem V, Troke RC, Alsafi A, Field BC, De Silva A, et al.. Combination of peptide YY3-36 with GLP-1 (7-36) amide causes an increase in first-phase insulin secretion after IV glucose. J Clin Endocrinol Metab 2014; 99:E2317–E2324.
111. Berthoud HR. The vagus nerve, food intake and obesity. Regul Pept 2008; 149:15–25.
112. Vahl TP, Tauchi M, Durler TS, Elfers EE, Fernandes TM, Bitner RD, et al.. Glucagon-like peptide-1
(GLP-1) receptors expressed on nerve terminals in the portal vein mediate the effects of endogenous GLP-1 on glucose tolerance in rats. Endocrinology 2007; 148:4965–4973.
113. Kessler JP, Beaudet A. Association of neurotensin
binding sites with sensory and visceromotor components of the vagus nerve. J Neurosci 1989; 9:466–472.
114. Koda S, Date Y, Murakami N, Shimbara T, Hanada T, Toshinai K, et al.. The role of the vagal nerve in peripheral PYY3-36-induced feeding reduction in rats. Endocrinology 2005; 146:2369–2375.
115. Bucinskaite V, Tolessa T, Pedersen J, Rydqvist B, Zerihun L, Holst JJ, Hellström PM. Receptor-mediated activation of gastric vagal afferents by glucagon-like peptide-1
in the rat. Neurogastroenterol Motil 2009; 21:978–e978.
116. Kessler JP, Moyse E, Kitabgi P, Vincent JP, Beaudet A. Distribution of neurotensin
binding sites in the caudal brainstem of the rat: a light microscopic radioautographic study. Neuroscience 1987; 23:189–198.
117. Göke R, Larsen PJ, Mikkelsen JD, Sheikh SP. Distribution of GLP-1 binding sites in the rat brain: evidence that exendin-4 is a ligand of brain GLP-1 binding sites. Eur J Neurosci 1995; 7:2294–2300.
118. Hernandez EJ, Whitcomb DC, Vigna SR, Taylor IL. Saturable binding of circulating peptide YY
in the dorsal vagal complex of rats. Am J Physiol 1994; 266 (Pt 1):G511–G516.
119. Holmes GM, Browning KN, Tong M, Qualls-Creekmore E, Travagli RA. Vagally mediated effects of glucagon-like peptide 1: in vitro and in vivo gastric actions. J Physiol 2009; 587 (Pt 19):4749–4759.
120. Chen CH, Rogers RC. Central inhibitory action of peptide YY
on gastric motility in rats. Am J Physiol 1995; 269 (Pt 2):R787–R792.
121. Putnam WS, Liddle RA, Williams JA. Inhibitory regulation of rat exocrine pancreas by peptide YY
and pancreatic polypeptide. Am J Physiol 1989; 256 (Pt 1):G698–G703.
122. Hellström PM. Vagotomy inhibits the effect of neurotensin
on gastrointestinal transit in the rat. Acta Physiol Scand 1986; 128:47–55.
123. Wettergren A, Wojdemann M, Holst JJ. Glucagon-like peptide-1
inhibits gastropancreatic function by inhibiting central parasympathetic outflow. Am J Physiol 1998; 275 (Pt 1):G984–G992.
124. Plamboeck A, Veedfald S, Deacon CF, Hartmann B, Wettergren A, Svendsen LB, et al.. The effect of exogenous GLP-1 on food intake is lost in male truncally vagotomized subjects with pyloroplasty. Am J Physiol Gastrointest Liver Physiol 2013; 304:G1117–G1127.
125. Imeryüz N, Yeğen BC, Bozkurt A, Coşkun T, Villanueva-Peñacarrillo ML, Ulusoy NB. Glucagon-like peptide-1
inhibits gastric emptying via vagal afferent-mediated central mechanisms. Am J Physiol 1997; 273 (Pt 1):G920–G927.
126. Nagell CF, Wettergren A, Ørskov C, Holst JJ. Inhibitory effect of GLP-1 on gastric motility persists after vagal deafferentation in pigs. Scand J Gastroenterol 2006; 41:667–672.
127. Wiley JW, Lu YX, Chung OY. Mechanism of action of peptide YY
to inhibit gastric motility. Gastroenterology 1991; 100:865–872.
128. Abbott CR, Monteiro M, Small CJ, Sajedi A, Smith KL, Parkinson JR, et al.. The inhibitory effects of peripheral administration of peptide YY
(3-36) and glucagon-like peptide-1
on food intake are attenuated by ablation of the vagal–brainstem–hypothalamic pathway. Brain Res 2005; 1044:127–131.
129. Rüttimann EB, Arnold M, Hillebrand JJ, Geary N, Langhans W. Intrameal hepatic portal and intraperitoneal infusions of glucagon-like peptide-1
reduce spontaneous meal size in the rat via different mechanisms. Endocrinology 2009; 150:1174–1181.
130. Orskov C, Poulsen SS, Møller M, Holst JJ. Glucagon-like peptide I receptors in the subfornical organ and the area postrema are accessible to circulating glucagon-like peptide I. Diabetes 1996; 45:832–835.
131. Baumgartner I, Pacheco-López G, Rüttimann EB, Arnold M, Asarian L, Langhans W, et al.. Hepatic-portal vein infusions of glucagon-like peptide-1
reduce meal size and increase c-Fos expression in the nucleus tractus solitarii, area postrema and central nucleus of the amygdala in rats. J Neuroendocrinol 2010; 22:557–563.
132. Sandoval DA, Bagnol D, Woods SC, D’Alessio DA, Seeley RJ. Arcuate glucagon-like peptide 1 receptors regulate glucose homeostasis but not food intake. Diabetes 2008; 57:2046–2054.
133. Secher A, Jelsing J, Baquero AF, Hecksher-Sørensen J, Cowley MA, Dalbøge LS, et al.. The arcuate nucleus mediates GLP-1 receptor agonist liraglutide-dependent weight loss. J Clin Invest 2014; 124:4473–4488.
134. Carpenter DO, Briggs DB, Strominger N. Responses of neurons of canine area postrema to neurotransmitters and peptides. Cell Mol Neurobiol 1983; 3:113–126.
135. Lee YS, Jun HS. Anti-diabetic actions of glucagon-like peptide-1
on pancreatic beta-cells. Metabolism 2014; 63:9–19.
136. Hauge M, Vestmar MA, Husted AS, Ekberg JP, Wright MJ, Di Salvo J, et al.. GPR40 (FFAR1) – combined Gs and Gq signaling in vitro is associated with robust incretin secretagogue action ex vivo and in vivo. Mol Metab 2015; 4:3–14.
137. Werry TD, Wilkinson GF, Willars GB. Mechanisms of cross-talk between G-protein-coupled receptors resulting in enhanced release of intracellular Ca2+. Biochem J 2003; 374 (Pt 2):281–296.
138. Shukla AP, Buniak WI, Aronne LJ. Treatment of obesity in 2015. J Cardiopulm Rehabil Prev 2015; 35:81–92.
139. Nuffer W, Trujillo JM, Megyeri J. A comparison of new pharmacological agents for the treatment of obesity. Ann Pharmacother 2016; 50:376–388.
140. Hammer HF. Medical complications of bariatric surgery: focus on malabsorption and dumping syndrome. Dig Dis 2012; 30:182–186.
141. Lutz TA, Bueter M. The physiology underlying Roux-en-Y gastric bypass: a status report. Am J Physiol Regul Integr Comp Physiol 2014; 307:R1275–R1291.
142. Dirksen C, Jørgensen NB, Bojsen-Møller KN, Kielgast U, Jacobsen SH, Clausen TR, et al.. Gut hormones, early dumping and resting energy expenditure in patients with good and poor weight loss response after Roux-en-Y gastric bypass. Int J Obes (Lond) 2013; 37:1452–1459.
143. Holdstock C, Zethelius B, Sundbom M, Karlsson FA, Edén Engström B. Postprandial changes in gut regulatory peptides in gastric bypass patients. Int J Obes (Lond) 2008; 32:1640–1646.
144. Finan B, Clemmensen C, Müller TD. Emerging opportunities for the treatment of metabolic diseases: glucagon-like peptide-1
based multi-agonists. Mol Cell Endocrinol 2015; 418 (Pt 1):42–54.
145. Finan B, Yang B, Ottaway N, Smiley DL, Ma T, Clemmensen C, et al.. A rationally designed monomeric peptide triagonist corrects obesity and diabetes in rodents. Nat Med 2015; 21:27–36.
146. Day JW, Gelfanov V, Smiley D, Carrington PE, Eiermann G, Chicchi G, et al.. Optimization of co-agonism at GLP-1 and glucagon receptors to safely maximize weight reduction in DIO-rodents. Biopolymers 2012; 98:443–450.