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Review
. 2023 Apr 1;103(2):1423-1485.
doi: 10.1152/physrev.00025.2022. Epub 2022 Nov 24.

The elusive cephalic phase insulin response: triggers, mechanisms, and functions

Wolfgang Langhans  1 , Alan G Watts  2 , Alan C Spector  3
Affiliations
Review

The elusive cephalic phase insulin response: triggers, mechanisms, and functions

Wolfgang Langhans et al. Physiol Rev. .

Abstract

The cephalic phase insulin response (CPIR) is classically defined as a head receptor-induced early release of insulin during eating that precedes a postabsorptive rise in blood glucose. Here we discuss, first, the various stimuli that elicit the CPIR and the sensory signaling pathways (sensory limb) involved; second, the efferent pathways that control the various endocrine events associated with eating (motor limb); and third, what is known about the central integrative processes linking the sensory and motor limbs. Fourth, in doing so, we identify open questions and problems with respect to the CPIR in general. Specifically, we consider test conditions that allow, or may not allow, the stimulus to reach the potentially relevant taste receptors and to trigger a CPIR. The possible significance of sweetness and palatability as crucial stimulus features and whether conditioning plays a role in the CPIR are also discussed. Moreover, we ponder the utility of the strict classical CPIR definition based on what is known about the effects of vagal motor neuron activation and thereby acetylcholine on the β-cells, together with the difficulties of the accurate assessment of insulin release. Finally, we weigh the evidence of the physiological and clinical relevance of the cephalic contribution to the release of insulin that occurs during and after a meal. These points are critical for the interpretation of the existing data, and they support a sharper focus on the role of head receptors in the overall insulin response to eating rather than relying solely on the classical CPIR definition.

Keywords: glucose; metabolism; taste receptor; vagus; β-cell.

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Conflict of interest statement

A. C. Spector is a member of the Scientific Advisory Board of Gila Therapeutics.

Figures

None
Graphical abstract
FIGURE 1.
FIGURE 1.
A hypothetical timeline of carbohydrate (CHO) processing (i), and the dynamics (not shown to scale) of oropharyngeal-gastrointestinal (Oro-GI) glucose (ii), blood glucose (iii), and blood insulin (iv) after food is either ingested via the mouth (A) or given directly into the stomach (B). Note that in this diagram the cephalic stimulation of insulin release is not a transient event only seen at the beginning of a meal but instead it lasts throughout.
FIGURE 2.
FIGURE 2.
Depiction of a simplified hypothetical framework for how signals arising from the interaction of a ligand (glucose, red hexagon) with two different receptors, T1R2 + T1R3 and an unidentified specific glucosensor (perhaps a GLUT or SGLT, see FIGURE 3), expressed in taste receptor cells might be differentially channeled into brain circuits subserving different taste functions. In the example, a second ligand (fructose, green pentagon) can only bind one (T1R2+T1R3) of the two receptor types, whereas glucose can activate both. Thus, glucose and fructose (and other potential ligands binding with the T1R2 + T1R3) activate similar neural circuits leading to identical outcomes in terms of perception and motivation and affect (with the exception that differences in binding affinity would lead to differences in relative intensity at isomolar concentrations). However, only glucose can stimulate the second chemospecific receptor, the output of which is channeled into circuits that give rise to a cephalic phase insulin response. Some support for this model derives from the facts that glucose and fructose: 1) appear to lead to identical qualitative taste perceptions, and 2) have a similar unconditioned positive taste valence that is virtually eliminated by genetic deletion of one or both subunits of the T1R2 + T1R3 receptor without affecting the ability of glucose to elicit a CPIR. The colored lines within the Integrative Circuits box represent the potential interconnections between the different neural circuits that underlie these taste functions. Importantly, this model makes no assumptions about the brain site(s) of these processes. Such interconnectivity would allow for conditioning to play a role in modulating the CPIR to a normally ineffective ligand and could also support the development of a hedonic discrimination between glucose and other T1R2 + T1R3 ligands such as fructose. T1R2, taste type 1 receptor 2; T1R3, taste type 1 receptor 3; SGLT, sodium-glucose cotransporter. The generic pink-shaded protein with a ? in the potential glucosensors signifies the possible contribution of some other receptor type that remains to be identified. See text for further discussion.
FIGURE 3.
FIGURE 3.
A: an idealized type II taste receptor cell that expresses the T1R2 + T1R3 heterodimer, which binds with sugars and other sweeteners, is depicted. The intracellular signaling cascades leading to the release of ATP through a CALHM1/3 channel are also illustrated. The ATP binds with P2X2/X3 receptors found on nearby afferent fibers. This form of neurotransmission involves an unconventional synapse. B: the various T1R-independent proteins in the apical membrane of some taste receptor cells that have been proposed to serve in the potential specific sensing of glucose in the oral cavity. The generic pink-shaded protein with a ? in the apical membrane signifies the possible contribution of some other receptor type that remains to be identified. The corresponding possible transduction pathways leading to putative neurotransmitter release are also depicted. α-Amylase in saliva and α-glucosidases expressed by some taste receptor cells and found in the taste pore are thought to hydrolyze various glucose polymers, and di-, and tri-saccharides providing a source of glucose moieties to be transported either through GLUTs or SGLT1 into the cell. There is evidence that glucokinase expressed in some type III cells is involved. The ATP produced by glucose catabolism is thought to close a KATP channel causing depolarization of the cell membrane. Additionally, the sodium cations co-transported with glucose through the SGLT1 are thought to contribute to membrane depolarization. The neurotransmitter and the mechanism of its release are unclear because there is evidence that P2X2/X3 knockout mice still display a cephalic phase insulin release to glucose. As discussed in the text, there are caveats for many of these proposed glucose-sensing mechanisms that remain to be resolved. CALHM1, calcium homeostasis modulator 1; DAG, diacylgycerol, GLUTs, glucose transporter(s); IP3, inositol triphosphate; Kir6.1, inward rectifying potassium channel subtype 6.1; P2X2/X3R, purinergic receptor 2, subtype 2, and subtype 3; PIP2, phosphatidylinositol 4,5-bisphosphate; PLCβ2, phospholipase C-β2; SGLT1, sodium-glucose cotransporter-1; SUR1, sulfonylurea receptor subtype-1; T1R2, taste type 1 receptor 2, T1R3, taste type 1 receptor 3; TRPM4, transient receptor potential melastatin type 4; TRPM5, transient receptor potential melastatin type 5; ↑ Vm, change in membrane potential. See text for more discussion.
FIGURE 4.
FIGURE 4.
A hypothetical schematic of the neural circuitry that can influence insulin release from β-cells (red text and arrows) in response to gustatory stimulation. The primary neural drive for cephalic phase insulin release (CPIR) consists of preganglionic parasympathetic neurons [vagal motor neurons (VMN)] in the dorsal motor nucleus of the vagus (DMX) that project to the pancreas to increase insulin release (brown text and arrows). Please see text and FIGURES 5–7 for more detailed representations of the intrapancreatic processes. How the neural control of these DMX neurons is organized for CPIR is still unclear, but this schema (1: green text and arrows) is a reasonable starting point based on the literature. The light tan box highlights this circuitry, which is also represented in FIGURE 5. Glucose in the oropharynx interacts with receptors expressed in taste receptor cells (see FIGURE 3) and generates sensory signals in the taste buds that are conveyed to the rostral nucleus of the solitary tract (rNTS) by the chorda tympani nerve (CT), the glossopharyngeal nerve (GL), the greater superficial petrosal nerve (GSP), and the superior laryngeal nerve (SLN) (purple text and arrows; see text for more details). Output from the rNTS is directed in two ways: first, either directly or indirectly to the DMX by as yet undefined projections (green dashed arrow); and second, to integrative regions in the caudal brainstem, primarily in the reticular formation and, in rodents, the parabrachial nucleus. In turn, these integrative regions project to the DMX (green arrows), but they also have bidirectional projections with various forebrain regions, including the hypothalamus, that may influence CPIR by direct and indirect projections to the DMX. DMX control of CPIR may also be influenced by glucagon-like peptide-1 (GLP-1) released from L-cells (2: blue text and arrows) in response to increasing glucose in the gut (see FIGURE 5). GLP-1 is detected by receptors (small blue circles) expressed by vagal sensory nerves that are located close to L-cells and in the hepatic portal vein wall (dark gray box). These vagal sensory nerves (VSN) then project to the medial part of the NTS (mNTS). Finally, glucose in the posthepatic circulation acts directly on β-cells to stimulate insulin secretion (3: black text and arrows).
FIGURE 5.
FIGURE 5.
Three mechanisms are activated by glucose as food from a meal passes from the oropharynx to the small intestine. 1: Cephalic phase (brown text and arrows) is the first of three consecutively engaged mechanisms that glucose uses to increase the release of insulin from β-cells in pancreatic islets (red text and arrows). When a meal begins, the action of glucose on taste receptors in the oropharynx leads via central integrative mechanisms (green dashed line and arrow) to the activation of vagal parasympathetic preganglionic neurons in the dorsal motor nucleus of the vagus (DMX). These vagal motor neurons (VMN) innervate intra-pancreatic ganglia (IPG) whose postganglionic neurons then innervate pancreatic islets and release the acetylcholine that potentiates the actions of ambient glucose to increase insulin release from β-cells (brown text and arrows). The light tan box indicates the brain circuitry shown in more detail in FIGURE 4. 2: The actions of incretins constitute the second mechanism (blue text and arrows) that is initiated as food transits within the small intestine where the increasing amounts of glucose that result from digestion act on enteroendocrine K-cells to release glucose-dependent insulinotropic peptide (GIP) and on L-cells to release glucagon-like peptide-1 (GLP-1). GIP enters the hepatic portal vein and then the posthepatic circulation to act as an incretin on β-cells. GLP-1 mainly acts on GLP-1 receptors (solid blue circles) on vagal sensory nerve (VSN) endings found in close proximity to L-cells and in the wall of the hepatic portal vein wall (dark gray box). These VSNs project to the medulla. Most of the GLP-1 released into the hepatic portal vein during normal meals is degraded by the liver and is presumably not the major stimulus for β-cell insulin secretion. 3: The final mechanism (black text and arrows) is the direct action on β-cells of the increased blood glucose concentrations that result from digestion and absorption.
FIGURE 6.
FIGURE 6.
A schematic to illustrate how parasympathetic neural drive, incretins, and glucose can engage intracellular signaling pathways in β-cells to increase insulin release. When a meal begins food is digested as it passes from the oropharynx to the stomach and then into the small intestine. Insulin release is stimulated by β-cell signaling pathways associated with three distinct but interacting mechanisms that originate in the oropharynx and small intestine. 1: Cephalic phase insulin release (CIPR) is stimulated by the interaction of glucose with taste receptors in the oropharynx (also see FIGURE 4). These mechanisms then stimulate the release of acetylcholine from vagal preganglionic nerves (i), and then in pancreatic islets from postganglionic parasympathetic motor fibers (brown text and arrows). Some cholinergic terminals are found distal to β-cells (ii), while in some species these terminals are found more proximal and in close apposition to β-cells (iii). Acetylcholine then binds to M3 muscarinic G-protein-coupled receptors (GPCRs) in the β-cell plasma membrane (brown square), which in turn activate membrane-bound phospholipase C (PLC) to catalyze the conversion of phosphatidylinositol 4,5-bisphosphate (PIP2) to diacylglycerol (DAG) and inositol trisphosphate (IP3). DAG-dependent mechanisms then activate protein kinase C (PKC) to increase the phosphorylation of proteins involved with insulin packaging, trafficking, and release (red text and arrows). IP3 increases the release of calcium (Ca2+) from intracellular stores thereby increasing insulin release into the hepatic portal vein. Note that acetylcholine-dependent mechanisms in β-cells can potentiate the actions of glucose on insulin secretion without concomitant increases in glucose (also see FIGURE 7). 2: As food is digested in the small intestine increased secretion of incretins from enteroendocrine cells is stimulated by elevated free glucose concentrations (see FIGURES 1 AND 5). During a meal glucose-dependent insulinotropic peptide (GIP) is the principal incretin (blue text and arrows) that binds to GPCRs in the β-cell plasma membrane (blue square) to activate adenyl cyclase (AC) and, via cAMP, protein kinase A (PKA). Downstream signaling from PKA can, like PKC, increase the phosphorylation of proteins involved with insulin packaging, trafficking, and release. This emphasizes the interactions between acetylcholine and incretins to potentiate the actions of glucose on insulin release (also see FIGURE 7). 3: Glucose absorption from the small intestine increases glucose in the blood and pancreatic islets. Glucose (black text and arrows) enters the β-cell via glucose transporters (GLUT) in its plasma membrane; primarily GLUT 1 in humans, GLUT 2 in mice. Glucose, together with certain amino acids and nonesterified fatty acids, is oxidized to release the energy that is used to synthesize ATP. The increasing ATP/ADP ratio then closes KATP channels in the β-cell plasma membrane, which helps to depolarize the β-cell and opens Ca2+ channels, thereby rapidly increasing intracellular Ca2+ concentrations (small gray box with red border) and insulin release.
FIGURE 7.
FIGURE 7.
There are three mechanisms that act in pancreatic islets and β-cells to control insulin release before and during a meal. This schematic illustrates the interactions between these mechanisms to control insulin release (red text and arrows), and how they change during a meal. They are as follows: 1: the parasympathetic motor system and the intra-pancreatic ganglia (IPG) (brown text and arrows); 2: incretins (blue text and arrows), primarily glucose-dependent insulinotropic peptide (GIP); and 3: the direct action of glucose on β-cells (black text and arrows). Mechanisms 2 and 3 are activated postabsorptively. The different text and arrow sizes denote relative concentrations and activation levels. Note, this schematic omits the details of signaling mechanisms, together with their molecular components, found in islets and β-cells. See FIGURE 6 and the text for more details of these mechanisms. A: in the absence of significant challenges to blood glucose homeostasis, insulin is released from β-cells primarily in response to ambient glucose to help maintain euglycemia. Influences from the parasympathetic motor system and incretins are minimal. B: as food enters the oral cavity it is chewed and digestion begins. Free glucose is detected by taste receptors in the oropharynx (purple circle) whose cognate primary sensory neurons project to the rostral part of the nucleus of the solitary tract (see FIGURE 4). This process involves integrative circuits, mainly in medulla (green arrow), that rapidly stimulates the parasympathetic motor system to release acetylcholine into pancreatic islets. Acetylcholine then potentiates the ability of glucose to stimulate insulin release, which occurs before the glucose produced from digestion enters the circulation. C: as the concentration of digested glucose and other nutrients increase in the gut, stimulate the release of GIP and other incretins, and move into the hepatic portal vein. The actions of increasing glucose concentrations on β-cells are potentiated by GIP. Because CPIR may well be maintained throughout the meal, the continuing potentiating actions of acetylcholine on β-cells continue to help maintain insulin secretion.
FIGURE 8.
FIGURE 8.
Section 5 discusses nine critical points and open questions about cephalic phase insulin release (CPIR). FIGURE 8 shows how these nine topics (sects 5.2 to 5.10) relate to the processes that enable the information flow from the oropharyngeal sensory systems (purple) that begins with increased amounts of glucose in the oropharynx during a meal (black), and ends with the initiation of insulin release from the β-cell (red) in response to cholinergic stimulation from parasympathetic innervation (brown). The color scheme used here is the same as that shown in FIGURE 4 and others in this review.
FIGURE 9.
FIGURE 9.
Plasma concentrations of glucose (black), insulin (red), and glucose-dependent insulinotropic peptide (GIP; blue) before, during, and after test meals in six adult rhesus macaques infused with saline (top) or atropine (bottom) from 20 min prior to 60 min after the onset of a test meal consisting of one banana, half of an apple cut into four pieces, and three monkey biscuits. During saline infusion, plasma levels of insulin, glucose, and GIP increased from baseline levels continuously throughout the meal and remained elevated thereafter. As indicated by the bigger symbols and asterisks, the increase in insulin was already significant at 2 min, whereas the increases in glucose and GIP did not reach significance before 10 min into the meal. During atropine infusion, the prandial increases in glucose and GIP were similar as during saline infusion (both increases over baseline significant after 15 min), but the plasma level of insulin remained virtually unchanged and did not increase significantly over baseline during the 60 min of the infusion, indicating that a substantial part of the overall insulin response to the meal was mediated by cephalic stimulation. See text for further details. Data are from Ref. .
FIGURE 10.
FIGURE 10.
Effect of oral sensory stimulation on plasma levels (mean ± SE) of glucose (black), insulin (red), and C-peptide (green) of eight normal-weight men after administration of 75 g of glucose intragastrically, with and without oral sensory stimulation. Time 0 indicates onset of glucose administration (beige bar) and sensory stimulation. Dashed lines represent control condition (no sensory stimulation), and solid lines represent sensory stimulation condition. Inset: areas under the curve (AUC) for conditions 1 and 2 (hatched bar, control condition; solid bar, sensory stimulation condition). *P < 0.05, **P < 0.03, significant differences between conditions. The oral stimulation led to significantly higher plasma insulin levels during the first 75 min after glucose administration load administration and subsequently to significantly lower glucose concentrations, indicating better glucose tolerance compared with controls. Figure is modified from Ref. , with permission from the American Physiological Society.
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References

    1. Moran TH. Gut peptides in the control of food intake: 30 years of ideas. Physiol Behav 82: 175–180, 2004. doi:10.1016/j.physbeh.2004.04.048. - DOI - PubMed
    1. Langhans W, Holst JJ. Afferent endocrine control of eating. In: Neuroendocrinology of Appetite, edited by Dickson SL, Mercer JG.. Chichester, UK: John Wiley & Sons, Ltd., 2016, p. 24–54.
    1. de Jong A, Strubbe JH, Steffens AB. Hypothalamic influence on insulin and glucagon release in the rat. Am J Physiol Endocrinol Metab 233: E380–E388, 1977. doi:10.1152/ajpendo.1977.233.5.E380. - DOI - PubMed
    1. Eales JG. The influence of nutritional state on thyroid-function in various vertebrates. Am Zool 28: 351–362, 1988. doi:10.1093/icb/28.2.351. - DOI
    1. Steffens AB, Van der Gugten J, Godeke J, Luiten PG, Strubbe JH. Meal-induced increases in parasympathetic and sympathetic activity elicit simultaneous rises in plasma insulin and free fatty acids. Physiol Behav 37: 119–122, 1986. doi:10.1016/0031-9384(86)90393-8. - DOI - PubMed
-