D-Glucose is an important source of energy for the cells, and the only energy source for red blood cells. At rest, glucose provides ∼1/3 and fatty acids ∼2/3 of the energy used by the body. Hypoglycaemia can damage nervous system, blood vessels, kidneys, and eyes. Hyperglycaemia can lead to micro- and macrovascular and neurological complications. Endocrine system and autonomic nervous system regulate blood glucose level and energy homeostasis (López-Gambero et al., 2018).
Intracellular glucose is phosphorylated by hexokinase to form glucose-6-phosphate in the first step of glycolysis. In the following steps, this glucose-6-phosphate is broken down, leading to net production of two ATP molecules and two NADH molecules. Glycolytic intermediates can enter other metabolic pathways, such as pentose phosphate pathway. If oxygen is available, much more ATP (38 molecules of ATP per glucose molecule) can be produced in mitochondria in oxidative phosphorylation. Brain uses mostly glucose via oxidative phosphorylation, but can also use ketones. At resting state, skeletal muscle converts about half of its glucose uptake to lactate. If oxygen supply is limited, as is the case during tissue hypoxia, tissues need to rely anaerobic glycolysis in their ATP production, leading to increased glucose uptake.
Tp53-inducible glycolysis and apoptosis regulator (TIGAR) is an enzyme that has multiple functions in cells. TIGAR inhibits glycolytic ATP synthesis and directs glucose into pentose phosphate pathway directly by catalysing fructose-2,6-biphosphate and by binding and increasing activity of hexokinase-2 in mitochondria. TIGAR also increases the rate of gluconeogenesis. TIGAR transcription is activated by p53 tumour suppressor protein. TIGAR (and p53) usually decreases ROS and protects cells from apoptosis, but under severe cellular stress TIGAR blocking can reduce oxidative stress and apoptosis. TIGAR can directly reduce cellular ATP levels via its phosphatase activity. Low ATP level arrests cell division cycle, but TIGAR expression is increased in many cancer types.
Glucose uptake by organs can be measured invasively based on the difference of arterial and venous glucose concentrations (Fick principle). Glucose analogue [18F]2-fluoro-2-deoxy-D-glucose ([18F]FDG, FDG) is transported into tissues and can be phosphorylated, and can thus be used to measure regional glucose uptake non-invasively. [11C]3-O-methyl-D-glucose (Vyska et al., 1983) and [18F]6-fluoro-6-deoxy-D-glucose (6FDG) can be transported but not phosphorylated, and can be used to measure glucose transport specifically (Landau et al., 2007). [18F]FDG has high affinity for GLUTs, and low affinity for ATP-dependent sodium-glucose transporters SGLTs. The relative role of GLUTs and SGLTs can be studied with related tracers, for example with α-methyl-4-[18F]-fluoro-4-deoxy-D-glucopyranoside (Me-4FDG) and 4-deoxy-4-[18F]-fluoro-D-glucose (4-FDG); Me-4FDG has high affinity for SGLT1, medium affinity for SGLT2, and very low affinity for GLUTs; 4-FDG is a substrate for both GLUTs and SGLTs (Sala-Rabanal et al., 2016). Overexpression of GLUT1 is typical for tumours, and is the basis for using FDG PET in clinical cancer imaging. FDG PET does not detect reliably certain cancer types; increased glucose uptake via SGLTs may then be assessed using for instance Me-4FDG (Scafoglio et al., 2015).
Total body glucose uptake, when there is no endogenous glucose production, can be measured using clamp technique: when glucose uptake is constant, total body glucose uptake equals the amount of glucose (corrected for urinary excretion of glucose) that must be infused to maintain constant glucose concentration in the blood. Endogenous glucose production can be stopped by insulin (hyperinsulinemic euglycemic clamp), hyperglycemia, or somatostatin.
[18F]FDG study may be combined with hyperinsulinemic euglycemic clamp (DeFronzo et al., 1979) to assess the insulin sensitivity of specific organs (Nuutila et al., 1993; Johansson et al., 2017) and endogenous glucose production. The whole body insulin sensitivity can be measured simultaneously as M value by dividing the mean glucose infusion rate by the lean body mass. The clamp technique requires frequent blood sampling; venous, arterialized venous, and finger tip samples can be used (Nauck et al., 1996), or samples can be drawn from the foot (Seaquist, 1997).
Glucose can be released from the glycogen storages (glycogenolysis) of the liver, or made from other substrates via gluconeogenesis in the liver, kidneys, and gut. Insulin and a decrease in plasma concentration of FFAs suppresses both mechanisms. Glucagon increases endogenous glucose production (EGP) and GLP-1 inhibits it (Seghieri et al., 2013).
Abnormal regulation of endogenous glucose production (EGP) is typical in patients with diabetes and insulin resistance. EGP represents the net production of glucose, as opposed to the total output of glucose, which includes EGP and the glucose cycling; glucose is constantly taken up by organs and circulated through cycling between glucose-6-phosphate and fructose-6-phosphate, and released back to blood (Iozzo et al., 2006). Glucose tracers that are labelled in position 3 or 6 do not loose the label in the isomerase reaction, and can be used to measure EGP. Tracers labelled in position 2 loose the label, and can be used to measure total glucose output (Iozzo et al., 2006). In [18F]FDG molecule fluorine-18 substitutes the hydroxyl group of glucose at position 2; [18F]FDG can be phosphorylated, but resulting [18F]FDG-6-phosphate cannot be transported out of the tissue, and in most cells further metabolism and dephosphorylation of [18F]FDG-6-phosphate is slow; in effect, phosphorylated [18F]FDG is trapped inside those cells. Plasma kinetics of [18F]FDG during hyperinsulinemic euglycemic clamp can be used to estimate whole-body EGP, and the metabolic clearance rate of glucose (Iozzo et al., 2006). FDG plasma clearance is adjusted with the FDG lost in urine to determine the rate of glucose disappearance, and EGP is calculated by subtracting glucose infusion rate from the rate of disappearance.
The superior mesenteric artery supplies 0.6-1.8 L/min blood to the proximal small intestine (jejunum and ileum) where glucose is mainly absorbed. Splanchnic circulation channels the absorbate via portal vein to the liver.
The sodium dependent glucose transporter SGLT1 transports glucose in the small intestine. If SGLT1 is inhibited or deficient, glucose is not absorbed in the small intestine, but fermented in the large intestine. Glucose absorption can be enhanced by apical GLUT2, when the enterocyte and submucosal glucose concentrations are lower than in the intestinal lumen. Otherwise, as a passive transporter, GLUT2 will reduce net absorption. Intestinal capillary network is an essential component of glucose transport as it maintains the glucose concentration gradient. SGLT1 can be studied using α-methyl-D-glucoside, which is a specific substrate for SGLT1. 3-O-methylglucose is a substrate for all SGLTs, but not GLUTs.
In the kidneys, D-glucose is reabsorbed almost fully, mostly already in the proximal tubules by SGLT2. SGLT2 inhibitors, such as canagliflozin and empaglifozin, can be used to lower plasma glucose levels in insulin-independent manner (de Albuquerque Rocha et al., 2018) and to reduce the risk for cardiovascular events (Åkerblom et al., 2019). Dapagliflozin reduces liver fat and visceral adipose tissue volume, but in 8 weeks no changes in insulin sensitivity in the major organs studied with FDG PET (Latva-Rasku et al., 2019).
Since SGLT2 in the proximal tubules is mainly responsible for the reabsorption of glucose, but 2-deoxy-D-glucose is not transported via SGLTs, reabsorption of 2-deoxy-D-glucose and 2-deoxy-D-glucose analogue FDG is low, and it happens in all of the tubular system, including collecting ducts. Knight et al (1977) reported that reabsorption of 3-O-methyl-D-glucose in rats is also low, and that it takes place mainly in the proximal tubules. Previously, Woosley & Huang (1968) reported that 3-O-methyl-D-glucose in rats is was predominantly reabsorbed, but in dogs excretion of 3-O-methyl-D-glucose was found to be the predominant process. Woosley et al (1970) measured higher but not full reabsorption of 2-deoxy-D-glucose in rat and dog kidneys, mainly in proximal tubules. In also humans, FDG is partially excreted into urine.
Glucosensor cells react to changes in glucose concentration, and are present in the central nervous system and periphery to maintain glucose homeostasis. Glucosensors are found in high densities in the hypothalamus, and in gastrointestinal tract (including oral cavity, small intestine, pancreas, and portal vein). Gut-brain axis keeps the brain informed about glucose concentration changes in the gastrointestinal tract, and controls metabolism in the peripheral organs. Hypothalamus contains neurons that excited (in the lateral part of arcuate nucleus) or inhibited (in the medial part of arcuate nucleus and in the ventromedial nuclei) by D-glucose. Glucose-excited neurons rely on GLUT2 in glucose sensing, but GLUT4 and SGLT1 may also play a role; increased glucose leads to increased [ATP] and ROS, which lead to membrane depolarization and release of neurotransmitters. Glucose-inhibited neurons also rely on GLUT2: decreased ATP/ADP-ratio increases NO production, which inhibits Cl--channels, affecting membrane depolarization. Glial cells, including astrocytes, can also take part in glucosensing, via NO signalling and regulating lactate shuttle. GLUT1 is present at the blood-brain barrier and choroid plexus.
Glucose sensing involves active and passive glucose transporters and GPCRs. Paracrine hormones, including GLP-1 and glucagon, can alter the glucose sensing, starting from the taste receptor cells. Enteric nervous system (ENS) controls intestinal motility, secretion, and absorption, but is also involved in glucosensing. Epithelial cells in the small intestine include brush cells, a distinct type of enterocytes, that participate in glucose sensing, triggering afferent nerves. Enteroendocrine cells (EECs) in small intestine secrete incretin peptides GLP-1 and GIP. Gut itself can produce glucose through gluconeogenesis, although less than the liver and kidneys.
In pancreatic islets of Langerhans, glucose is transported into β-cells via GLUT2, phosphorylated by hexokinase IV, and used to produce ATP in mitochondria. Increased cytosolic [ATP] and production of ROS leads to depolarization of β-cell membranes, increased cytosolic [Ca2+], and exocytosis of a subset of insulin containing vesicles. Parasympathetic nervous system is also involved in the control of insulin release. Pancreatic α-cells do not express GLUT2, and while glucagon secretion is mainly controlled by hypothalamus via sympathetic nervous system and signalling from other islet cell types, α-cells can still themselves sense the glucose level. Glucagon activates glycogenolysis and gluconeogenesis in the liver and lipolysis in adipose tissue.
Hepatoportal glucosensing is dependent on GLUT2 and GLP-1R. SGLT3 may be important when glucose concentrations are very low. Increasing D-glucose concentration (but not D-fructose or other carbohydrates that are not metabolized by nerve cells) in portal vein decreases the firing rate of hepatic afferent nerves. Signal is then transmitted to other organs via vagus nerve and autonomic nervous system. Activation of sympathetic nervous system stimulates glucose uptake and glycogenesis in skeletal muscle, and synthesis and storage of lipids in white adipose tissue. Hypothalamus can regulate hepatic glucose production via parasympathetic nervous system. Liver can also detect glucose concentration gradients between the portal vein and hepatic artery. In type 2 diabetes hepatoportal glucose sensing is altered.
Glycosylation is the strictly regulated enzymatic addition of glucose and other sugars to proteins and lipids, which is essential to their function and regulation.
Glycation is the spontaneous reaction of glucose and other hexoses with the amine groups of proteins; long-term high blood glucose level leads to accumulation of glycation products () and is involved in complications of diabetes, such as peripheral neuropathy and renal failure. Measurement of the amount of glycated haemoglobin in blood sample is used to assess three-month average plasma glucose concentration. Glucose has relatively stable cyclic forms, and therefore its glycation rate is lower than that of many other hexoses.
Alvarsson A, Stanley SA. Remote control of glucose-sensing neurons to analyze glucose metabolism. Am J Physiol Endocrinol Metab. 2018; 315: E327-E339. doi: 10.1152/ajpendo.00469.2017.
DeFronzo RA, Tobin JD, Andres R. Glucose clamp technique: a method for quantifying insulin secretion and resistance. Am J Physiol. 1979; 237(3): E214-E223. doi: 10.1152/ajpendo.1979.237.3.E214.
Fournel A, Marlin A, Abot A, Pasquio C, Cirillo C, Cani PD, Knauf C. Glucosensing in the gastrointestinal tract: Impact on glucose metabolism. Am J Physiol Gastrointest Liver Physiol. 2016; 310(9): G645-G658. doi: 10.1152/ajpgi.00015.2016.
Iozzo P, Gastaldelli A, Järvisalo MJ, Kiss J, Borra R, Buzzigoli E, Viljanen A, Naum G, Viljanen T, Oikonen V, Knuuti J, Savunen T, Salvadori PA, Ferrannini E, Nuutila P. 18F-FDG assessment of glucose disposal and production rates during fasting and insulin stimulation: a validation study. J Nucl Med. 2006; 47(6): 1016-1022. PMID: 16741312.
Jouvet N, Estall JL. The pancreas: Bandmaster of glucose homeostasis. Exp Cell Res. 2017; 360: 19-23. 10.1016/j.yexcr.2017.03.050.
Lassen N.A., Gjedde A. (1983): Kinetic Analysis of the Uptake of Glucose and Some of its Analogs in the Brain Using the Single Capillary Model: Comments on Some Points of Controversy. In: Lambrecht R.M., Rescigno A. (eds.) Tracer Kinetics and Physiologic Modeling. Lecture Notes in Biomathematics, vol 48. Springer, Berlin, Heidelberg. doi: 10.1007/978-3-642-50036-7_8.
López-Gambero AJ, Martínez F, Salazar K, Cifuentes M, Nualart F. Brain glucose-sensing mechanism and energy homeostasis. Mol Neurobiol. 2018 (in press). doi: 10.1007/s12035-018-1099-4.
Lund-Andersen. Transport of glucose from blood to brain. Phys Rev. 1979; 59(2): 305-352. doi: 10.1152/physrev.19184.108.40.2065.
Knight T, Sansom S, Weinman EJ. Renal tubular absorption of D-glucose, 3-O-methyl-D-glucose, and 2-deoxy-D-glucose. Am J Physiol. 1977; 233(4): F274-F277. doi: 10.1152/ajprenal.1977.233.4.F274.
Kowalski GM, Bruce C. The regulation of glucose metabolism: Implications and considerations for the assessment of glucose homeostasis in rodents. Am J Physiol Endocrinol Metab. 2014; 307: E859-E871. doi: 10.1152/ajpendo.00165.2014.
Petersen MC, Vatner DF, Shulman GI. Regulation of hepatic glucose metabolism in health and disease. Nat Rev Endocrinol. 2017; 13(10): 572-587. doi: 10.1038/nrendo.2017.80.
Poretsky L (ed.): Principles of Diabetes Mellitus, 3rd ed., Springer, 2017. doi: 10.1007/978-3-319-18741-9.
Verberne AJ, Sabetghadam A, Korim WS. Neural pathways that control the glucose counterregulatory response. Front Neurosci. 2014; 8: 38. doi: 10.3389/fnins.2014.00038.
Vyska K, Profant M, Schuier F, Freudlieb C, Höck A, Thal H-U, Becker V, Feinendegen LE. The use of 11C-methyl-D-glucose for assessment of glucose transport in the human brain: theory and application. In: Lambrecht R.M., Rescigno A. (eds.) Tracer Kinetics and Physiologic Modeling. Lecture Notes in Biomathematics, vol 48. Springer, Berlin, Heidelberg, 1983. doi: 10.1007/978-3-642-50036-7_9.
Zierler K. Whole body glucose metabolism. Am J Physiol. 1999; 276(3 Pt 1): E409-E426. 10.1152/ajpendo.1999.276.3.E409.
Updated at: 2019-03-30
Created at: 2018-08-28
Written by: Vesa Oikonen