Biochemistry or biological chemistry deals with the study of chemical processes that occur in animals, plants and microorganisms.
Biochemistry is, with inorganic chemistry, organic chemistry, physical chemistry and analytical chemistry one of the five main branches of chemistry. And, as the name suggests, it can also be considered a branch of biology.
It emerged as a discipline in its own right in the early 1900s when scientists combined biology, physiology and chemistry to study living beings, exploiting the knowledge accumulated in the previous century thanks to the work of scientists such as Justus von Liebig, Eduard Buchner, Louis Pasteur or Emil Fischer.
Since the mid-1900s, biochemists, also exploiting the advancement of knowledge in chemistry, physics and mathematics, have been able to analyze and clarify cellular metabolic pathways. Pathways that allow to obtain energy from food or from the sun, such as glycolysis, the citric acid cycle, lipolysis, oxidative phosphorylation, or photosynthesis. And how cells use the energy to produce what they need to live and reproduce. Therefore, light has been shed on what cellular metabolism is, namely, the anabolic and catabolic pathways that allow the synthesis and catabolism of molecules and macromolecules such as carbohydrates, proteins, lipids and nucleic acids have been clarified. The structure of these macromolecules has also been clarified, for example of DNA thanks to the work of Rosalind Elsie Franklin, James Dewey Watson and Francis Harry Compton Crick. And with metabolism and enzymology, structural biology is one of the three branches into which biochemistry can be divided.
Pyruvic acid, an alpha-ketoacid, is a molecule with a central role in cellular metabolism.[13][14]
It can be produced through various metabolic pathways, mostly cytosolic, among which glycolysis is usually the most important, while its fate depends on the cell type and the availability of oxygen, as it can be used for energy, biosynthetic, and anaplerotic purposes.[1]
Given its central role in cellular metabolism, mutations in the genes that encode the proteins involved in its metabolism cause, in humans, mild to severe diseases.[7]
Pyruvic acid or, according to the IUPAC nomenclature, 2-oxopropanoic acid, has a molecular weight of 88.06, molecular formula C3H4O3, and condensed formula CH3COCOOH.
It belongs to the group of alpha-ketoacids, that is, keto acids which have the carbonyl group adjacent to the carboxylic acid. Among the alpha-ketoacids, it has the simplest chemical structure.[13]
In purified form it appears as a colorless liquid, with a smell similar to that of acetic acid.
Its acid dissociation constant (pKa), at 25 °C, is equal to 2.45. It is therefore a very strong acid, and, at physiological pH, both in cells and in extracellular fluids, it is present almost entirely in its anionic form, pyruvate.
Its melting point is 13.8 °C (56.84 °F; 286.95 K), while its boiling point is 163.5 °C (329 °F; 438 K).[13]
Pyruvate metabolism
The biosynthetic pathways leading to pyruvic acid production, as well as its subsequent utilization, depend on the cell type and/or the availability of oxygen.[1]
In cells with mitochondria, pyruvate metabolism consists of a cytosolic and a mitochondrial phase. In the cytosol, several metabolic pathways lead to its formation, namely, glycolysis, the oxidation of lactate, the reaction catalyzed by the cytosolic malic enzyme, and the catabolism of at least six amino acids, the most important of which is alanine. Pyruvic acid can also be produced in the mitochondrial matrix from alanine and lactate.[10] The pyruvic acid produced in the cytosol, by means of specific transporters, enters the mitochondrion where it can be used for energy, entering the citric acid cycle, and/or biosynthetic/anaplerotic purposes, depending on the needs of the cell.[15]
If, instead, we consider cells without mitochondria, such as red blood cells, and, under hypoxic conditions, cells with mitochondria too, pyruvate is reduced to lactate and/or leaves the cell as such to be metabolized in other tissues, for example cardiac muscle.[11]
Glycolysis
Under physiological conditions, in most cells pyruvic acid is mainly derived from glycolysis, of which it is one of the three products, with ATP and NADH.
In the last step of the glycolytic pathway, pyruvate kinase (EC 2.7.1.40) catalyzes a substrate-level phosphorylation that leads to the transfer of the phosphoryl group from phosphoenolpyruvate to ADP. The reaction, essentially irreversible, produces an ATP and a pyruvate.
Phosphoenolpyruvate + ADP + H+ → Pyruvate + ATP
Therefore, during the glycolytic pathway, two molecules of pyruvate are produced from a glucose molecule.[12][14]
Lactate dehydrogenase
In the cytosol, pyruvic acid can be produced from lactic acid.
The enzyme lactate dehydrogenase (LDH) (EC 1.1.1.27) catalyzes the reversible conversion of pyruvate to lactate and of NADH to NAD+.
Pyruvate + NADH + H+ ⇄ Lactate + NAD+
The direction of the reaction depends on lactate dehydrogenase isozymes present and the NADH/NAD+ ratio in the cytosol.[1][10]
Isoenzymes in which the H subunit predominates, such as LDH-1, predominate in cardiac muscle, an exclusively aerobic tissue, where they catalyze the oxidation of lactate, formed in other tissues, to pyruvate, which is then used for energy.
The oxidation of lactate produced for example by red blood cells or skeletal muscle under hypoxic conditions, can also occur in hepatocytes, favored by the low NADH/NAD+ ratio in the cytosol, although LDH-5 is the main isoenzyme. In the hepatocyte, pyruvic acid can be enter gluconeogenesis, which in this case is part of the Cori cycle, or be oxidized for energy.[6]
Conversely, in skeletal muscle fiber under hypoxic conditions, in which the pyruvate dehydrogenase complex is inhibited and oxidative phosphorylation is blocked, in order for glycolysis to proceed pyruvate is reduced to lactate with the concomitant oxidation of NADH to NAD+.[9] Note that the conversion of glucose into lactate is defined as lactic fermentation. This reduction is favored by the fact that in skeletal muscle fiber isoenzymes with a prevalence of the M subunit predominate, such as LDH-4 and LDH-5.[4]
Alanine aminotransferase
Another source of pyruvic acid is alanine, an amino acid particularly abundant in muscle proteins.
The utilization of the carbon skeleton of amino acids for energy and/or anabolic purposes involves the removal of basic amino group, which occurs in reactions catalyzed by enzymes called transaminases (EC 2.6.1-), and the subsequent disposal of nitrogen in a non-toxic form through the urea cycle.[12][14]
The removal of the amino group of alanine is catalyzed by alanine aminotransferase (ALT) (EC 2.6.1.2). The reaction, reversible, leads to the formation of pyruvate and glutamate.
Two forms of ALT have been identified: ALT1, localized in the cytosol, and ALT2, localized in the mitochondrial matrix.[7] Hence, pyruvic acid can be produced by transamination of alanine also in the mitochondrial matrix.
Alanine is one of the main gluconeogenic precursors, and, through the glucose-alanine cycle, represents a link between the metabolism of carbohydrates and amino acids.[5]
In the cytosol, the carbon skeletons of five other amino acids, namely cysteine, glycine, serine, threonine and tryptophan, can be converted partly or entirely to pyruvate.[14]
Malic enzyme
Pyruvate can also be produced from malate.
In the reaction catalyzed by the cytosolic malic enzyme (EC 1.1.1.40), malate undergoes an oxidative decarboxylation to yield pyruvate.[1]
Malate + NADP+ → Pyruvate + CO2 + NADPH + H+
Malic enzyme plays an important role in the transport of intermediates of the citric acid cycle, such as, in addition to malate, oxaloacetate and citrate, between the cytosol and the mitochondrial matrix.[11]
Mitochondrial pyruvate carriers
In cells with mitochondria, most of the pyruvate produced in the cytosol enters the mitochondrial matrix passing through the outer and then the inner mitochondrial membrane. In the mitochondrial matrix pyruvate can then be used for both anabolic and catabolic purposes.
The passage through the outer mitochondrial membrane occurs by free diffusion through non-specific voltage-dependent anion channels (VDACs) or porins, the most abundant proteins of the outer mitochondrial membrane, whose function is to mediate the exchange of ions and small molecules, including, in addition to pyruvate, also ATP, NADH and others, between the cytosol and the intermembrane space of mitochondria.[3][16]
Conversely, the inner mitochondrial membrane is impermeable to charged molecules, which allows the maintenance of the proton gradient needed for oxidative phosphorylation to occur. The passage of pyruvate therefore occurs through specific transporters, the mitochondrial pyruvate carriers (MPC), a hetero-oligomeric complex of two subunits, indicated as MPC1 and MPC2. This transport is coupled to a symport of protons. MPC therefore links the cytosolic and mitochondrial metabolism of pyruvate.[2][8]
Pyruvate dehydrogenase complex
Once in the mitochondrial matrix, pyruvate is mostly oxidized to carbon dioxide in order to support ATP production.
The first step in this oxidation process involves the pyruvate dehydrogenase complex (PDC), one of the most important multienzyme complexes present in cells. The complex catalyzes the irreversible oxidative decarboxylation of pyruvate to form acetyl-coenzyme A (acetyl-CoA) and a carbon dioxide molecule, with the release of two electrons, which are carried by NAD.
Pyruvate + CoA + NAD+ → Acetyl-CoA + NADH + H+ + CO2
The acetyl group can enter the citric acid cycle to be completely oxidized to carbon dioxide, with the production of a molecule of GTP, 3 molecules of NADH, one of FADH2. The two reduced coenzymes, through oxidative phosphorylation, will allow the production of further ATP molecules.[14]
Alternatively, the acetyl group derived from pyruvic acid can be used for anabolic purposes, among which the synthesis of the synthesis of some lipids, such as of fatty acids, phospholipids and cholesterol, or for regulatory purposes through histone acetylation.
Since lipid biosynthesis occur in the cytosol and histone acetylation in the nucleus, acetyl-CoA must leave the mitochondrial matrix, a process that requires the formation of citrate through the condensation between the acetyl group and carbonyl of oxaloacetate, reaction catalyzed by citrate synthase (EC 2.3.3.1). Once citrate has reached the cytosol, through a citrate carrier, an integral protein of the inner mitochondrial membrane, it is cleaved to acetyl-CoA and oxaloacetate in the reaction catalyzed by citrate lyase (EC 4.1.3.6).[17]
Pyruvate carboxylase
In the mitochondrial matrix, pyruvic acid can be carboxylated to oxaloacetate in an irreversible reaction catalyzed by pyruvate carboxylase (PC) (EC 6.4.1.1).[12]
Pyruvate + HCO3– + ATP ⇄ Oxaloacetate + ADP + Pi
Several intermediates of the citric acid cycle are precursors for the synthesis of various molecules. It follows that each of these intermediates removed from the citric acid cycle must be reintegrated for the cycle to continue. Reactions that reintegrate the cycle are defined as anaplerotic. In this perspective, the reaction catalyzed by pyruvate carboxylase plays an anaplerotic function, catalyzing the formation of oxaloacetate.[15]
References
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^ Bricker D.K., Taylor E.B., Schell J.C., Orsak T., Boutron A., Chen Y.C., Cox J.E., Cardon C.M., Van Vranken J.G., Dephoure N., Redin C., Boudina S., Gygi S.P., Brivet M., Thummel C.S., Rutter J. A mitochondrial pyruvate carrier required for pyruvate uptake in yeast, Drosophila, and humans. Science 2012;337(6090):96-100. doi:10.1126/science.1218099
^ Colombini M. The VDAC channel: molecular basis for selectivity. Biochim Biophys Acta 2016;1863(10):2498-502. doi:10.1016/j.bbamcr.2016.01.019
^ Farhana A., Lappin S.L. Biochemistry, lactate dehydrogenase. [Updated 2023 May 1]. In: StatPearls [Internet]. Treasure Island (FL): StatPearls Publishing; 2024 Jan-. Available from: https://www.ncbi.nlm.nih.gov/books/NBK557536/
^ Felig P., Pozefsk T., Marlis E., Cahill G.F. Alanine: key role in gluconeogenesis. Science 1970;167(3920):1003-1004. doi:10.1126/science.167.3920.1003
^ Gleeson T.T. Post-exercise lactate metabolism: a comparative review of sites, pathways, and regulation. Annu Rev Physiol 1996;58:565-81. doi:10.1146/annurev.ph.58.030196.003025
^ ab Gray L.R., Tompkins S.C., Taylor E.B. Regulation of pyruvate metabolism and human disease. Cell Mol Life Sci. 2014 Jul;71(14):2577-604. doi:10.1007/s00018-013-1539-2
^ Herzig S., Raemy E., Montessuit S., Veuthey J.L., Zamboni N., Westermann B., Kunji E.R., Martinou J.C. Identification and functional expression of the mitochondrial pyruvate carrier. Science. 2012;337(6090):93-6. doi:10.1126/science.1218530
^ Li X., Yang Y., Zhang B., Lin X., Fu X., An Y., Zou Y., Wang J.X., Wang Z., Yu T. Lactate metabolism in human health and disease. Signal Transduct Target Ther 2022;7(1):305. doi:10.1038/s41392-022-01151-3
^ ab Markert C.L., Shaklee J.B., Whitt G.S. Evolution of a gene. Multiple genes for LDH isozymes provide a model of the evolution of gene structure, function and regulation. Science 1975;189(4197):102-14. doi:10.1126/science.1138367
^ ab McCommis K.S. and Finck B.N. Mitochondrial pyruvate transport: a historical perspective and future research directions. Biochem J 2015;466(3):443-454. doi:10.1042/BJ20141171
^ abc Moran L.A., Horton H.R., Scrimgeour K.G., Perry M.D. Principles of Biochemistry. 5th Edition. Pearson, 2012
^ abcd Nelson D.L., Cox M.M. Lehninger. Principles of biochemistry. 6th Edition. W.H. Freeman and Company, 2012
^ ab Owen O.E., Kalhan S.C., Hanson R.W. The key role of anaplerosis and cataplerosis for citric acid cycle function. J Biol Chem 2002;277(34):30409-12. doi:10.1074/jbc.R200006200
^ Young M.J., Bay D.C., Hausner G., Court D.A. The evolutionary history of mitochondrial porins. BMC Evol Biol 2007;7:31. doi:10.1186/1471-2148-7-31
^ Zara V., Assalve G., Ferramosca A. Multiple roles played by the mitochondrial citrate carrier in cellular metabolism and physiology. Cell Mol Life Sci 2022;79(8):428. doi:10.1007/s00018-022-04466-0
Lactate dehydrogenase or LDH (EC 1.1.1.27) is a family of oxidoreductases that catalyze the reversible conversion of pyruvate to lactate, with the concomitant interconversion of NADH and NAD+, which act as cofactors.
They are tetrameric enzymes, where each subunit has catalytic activity. The subunits, encoded by distinct genes, can be assembled in different combination to form isozymes with specific kinetic and regulatory properties.[3]
Lactate dehydrogenase is found in almost all animal tissues, plants, but also in microorganisms. Although it is mostly present in the cytosol, its presence has also been demonstrated in mitochondria, where it catalyzes the oxidation of lactate to pyruvate, and in peroxisomes.[14][20]
In humans, different isoenzymes have preferential tissue localizations, based on the specific metabolic phenotype of the tissue.
Lactate dehydrogenase is a important enzyme in cellular metabolism, as it is involved in energy production from carbohydrates under anaerobic conditions, in the synthesis of glucose from lactate, and utilization of the carbon skeleton of lactate for energy under aerobic conditions.[3][12]
In mammals, several genes encode the subunits of lactate dehydrogenase and are designated LDHA, LDHB, LDHC, LDHx, and LDHD. The first four genes encode enzymes that recognize as substrate the L-isomers of lactic acid, a molecule with a chirality center, the major enantiomeric form of the molecule present in vertebrates, and have NAD as a cofactor, whereas LDHD encodes an enzyme that recognizes as substrate the D-isomer of lactic acid and has FAD as a cofactor.[4][10] LDHA, located on chromosome 11p15.4, encodes the M subunit, named for its discovery in muscle, whereas LDHB, located on chromosome 12p12.2-p12.1, encodes the H subunit, named for its discovery in heart tissue.[11][14] LDHC, located on chromosome 11p15.5-p15.3 and probably derived from the duplication of the LDHA, encodes the C subunit.[23]
Finally, LDHx encodes the peroxisomal form of LDH.[14] LDHx is the readthrough form of the LDHB gene; in fact what happens is that when the ribosome reaches the stop codon of the mRNA for the H subunit, reads it as encoding an amino acid. Then the translation proceeds by adding another seven amino acids that constitute the peroxisomal targeting signal by which the proteins are imported into the peroxisome.[20]
Structure
The enzymes belonging to the lactate dehydrogenase family have an oligomeric structure, specifically they are tetramers resulting from the assembly of subunits, of approximately 35 kD, which can be of the same type or two different types. Each subunit has a catalytic site, hence, the tetramer has four active sites. However, the subunits, taken individually, are catalytically inactive.[11][17]
Considering the H and M subunits, their primary structures are approximately 75 percent identical, and consequently, their three-dimensional structure is also very similar, but with small differences at the at the substrate binding site that lead to significant differences in the kinetic properties of the proteins.[2][3] Another consequence of the differences in the primary structure concerns the net surface charge, which is -6 for the M subunit and +1 for the H subunit.[18]
The secondary structure is made up of approximately 40 percent alpha-helices and 23 percent beta sheets, forming beta-alpha-beta structures, with the parallel beta sheets as the main component.[1]
In humans, the prevalent isozymes are those formed by the H and M subunits. The assembly, in different combinations, of the two subunits leads to the formation of five isozymes, namely:
The C subunit forms a sixth isoform, a homotetramer, named as LDH-6 or C4. [23]
A seventh isoform derives from the assembly of the LDHx subunit.[20][23]
Reaction
Lactate dehydrogenase catalyzes the reversible conversion of pyruvate to lactate, which are the conjugate base of pyruvic acid and lactic acid, respectively, with the concomitant interconversion of NADH and NAD+. In both cases, the removal of two hydrogen atoms from the reducing agent occurs, followed by the transfer of a hydride ion, namely, a proton and two electrons, to the oxidizing agent, while the remaining proton is released into the aqueous medium as free H+ ion. The enzyme is able to increase the reaction rate by 14 times.[4]
When the reaction proceeds from pyruvate to lactate, the first step is the binding of NADH, the reducing agent, to the enzyme, which is followed by conformational changes that facilitate the formation of hydrogen bonds between specific residues surrounding the active site and the carbonyl carbon of pyruvate, namely C2, and the subsequent interaction between NADH and pyruvate.[6] At this point, a hydride ion is transferred from the nicotinamide ring of NADH to the C2 of pyruvate, which is therefore reduced to lactate. This causes the oxidation of the coenzyme and neutralization of the positive charge carried by the nitrogen of the nicotinamide ring, and reduction of pyruvate to lactate. Therefore, in this case the C2 of pyruvate is reduced from ketone to alcohol.
In the opposite direction, the hydride ion is transferred from the C2 of lactate, which in this case is the reducing agent, to the to the nicotinamide atom C-4. In this case the C2 of lactate is oxidized from alcohol to ketone.[15][16]
Active site
The active site of the H and M subunits has a highly conserved structure in different species, with the same amino acids participating in the reaction.[9] His-193, which is found near the binding site for the coenzyme, is among these, not only for human lactate dehydrogenase, but also for that of many other species.
However, the small differences in their primary structure lead to different biochemical properties. Among the differences, one is crucial for the catalytic properties: an alanine of the M subunit is replaced by a glutamine in the H subunit, namely, a non-polar amino acid is replaced by a polar one.[18] Below is a brief overview of the catalytic differences between the two subunits.
The H subunit binds substrate faster than the M subunit, but has about five times less catalytic activity than the M subunit.
The M subunit has a higher affinity for pyruvate, thus favoring the formation of lactate and NAD+, and is not inhibited by high concentrations of pyruvate. Conversely, the H subunit has a higher affinity for lactate, which favors the formation of pyruvate and NADH, and is inhibited by high concentrations of pyruvate.[5][8]
From all this it follows that the kinetic/catalytic properties of the different isozymes depend on the prevalence of one of the two subunits.[19]
Regulation by pyruvate and lactate
Lactate dehydrogenase isozymes are subject to inhibition by pyruvate and lactate.
The isozymes where the H subunit predominates are inhibited by lower pyruvate concentrations than those where the M subunit predominates. For example, H4 is inhibited by pyruvate concentrations of about 0.2 mM, while M4 is weakly inhibited by pyruvate concentrations up to 5 mM. Conversely, H4 is inhibited by lactate concentrations greater than 20-40 mM, whereas M4 is inhibited to a lesser extent by high lactate concentrations.[21]
Tissue distribution
In humans, the different lactate dehydrogenase isozymes have preferential tissue localizations, which generally reflect the metabolic phenotype of the tissue. In fact, tissues with a predominantly or exclusively aerobic metabolic phenotype, such as the heart, produce mainly isozymes in which the H subunit predominate.[22]
By contrast, tissues where anaerobic metabolism is important, such as muscle fibers during vigorous exercise, or hypoxic regions in solid tumors, produce mainly isozymes in which the M subunit predominate.[4]
However, there are exceptions, such as the liver, an organ with an aerobic metabolic phenotype where LDH-5 predominates, but where the oxidation of lactate to pyruvate, which can also be considered as a part of the hepatic branch of the Cori cycle, is favored by the low NADH/NAD+ ratio in the hepatocyte cytosol.[6]
Below is a brief overview.
LDH-1 or H4: cardiac muscle, kidney and red blood cells.
LDH-2 or M1H3 has a distribution similar to LDH-1.
LDH-3 or M2H2: spleen, brain, white blood cells, kidney and lung.
LDH-4 or M3H1: spleen, lung, skeletal muscle, red blood cells and kidney.
LDH-5 or M4: liver, skeletal muscle and lung.
LDH-6: sperm and testis.
In individual organs, LDH-1 and LDH-2 predominates in the heart, kidney and red blood cells, LDH-3 in the lung, LDH-4 and LDH-5 in skeletal muscle and LDH-5 in the liver.
The isozymes present in serum, whose dosage is used for diagnostic purposes, are always of tissue origin.[4]
Finally, in the same tissue/organ, different isozymes can be present in significant quantities in different cell types, as for example occurs in skeletal muscle, in the brain, but also in solid tumors.
Role
Lactate dehydrogenase is involved both in the synthesis and utilization/removal of lactate.[12]
Under hypoxic conditions, the cell obtains ATP from the anaerobic oxidation of glucose. Through the glycolytic pathway, the monosaccharide is oxidized to two molecules of pyruvate, yielding 2 ATP and two NADH. However, for glycolysis to proceed, the NADH produced must be reoxidized to NAD+, as the oxidized coenzyme is involved in the reaction catalyzed by glyceraldehyde-3-phosphate dehydrogenase (EC 1.2.1.12). As the pyruvate dehydrogenase complex is inhibited and oxidative phosphorylation is blocked, the oxidation of NADH occurs in the reaction catalyzed by lactate dehydrogenase, which then allows glycolysis to proceed even in hypoxic conditions. This metabolic pathway produces lactate from glucose, and is known as lactic acid fermentation.[6][13]
Lactic acid is often considered an end product of glucose metabolism, and its accumulation in the body is harmful as it may potentially causelactic acidosis.[13] Therefore, it must be rapidly removed from tissues and circulation. Considering for example the lactose produced by red blood cells or muscle fibers working in under hypoxic conditions, such as during vigorous exercise, or continuously by the red blood cell, through bloodstream, it can reach, among others, the liver and the heart. In the liver, although LDH-5 is the major isozyme, the low cytosolic NADH/NAD+ ratio shifts the equilibrium of the reaction toward pyruvate synthesis, which can be used for energy or enter into gluconeogenesis.[7] In the cardiomyocyte LDH-1 oxidizes lactate to pyruvate, that will be used for energy.
References
^ Auerbach G., Ostendorp R., Prade L., Korndörfer I., Dams T., Huber R., Jaenicke R. Lactate dehydrogenase from the hyperthermophilic bacterium thermotoga maritima: the crystal structure at 2.1 A resolution reveals strategies for intrinsic protein stabilization. Structure 1998;6(6):769-81. doi:10.1016/s0969-2126(98)00078-1
^ Bellamacina C.R. The nicotinamide dinucleotide binding motif: a comparison of nucleotide binding proteins. FASEB J 1996;10(11):1257-69. doi:10.1096/fasebj.10.11.8836039
^ abc Berg J.M., Tymoczko J.L., and Stryer L. Biochemistry. 5th Edition. W. H. Freeman and Company, 2002
^ abcd Farhana A., Lappin S.L. Biochemistry, lactate dehydrogenase. [Updated 2023 May 1]. In: StatPearls [Internet]. Treasure Island (FL): StatPearls Publishing; 2024 Jan-. Available from: https://www.ncbi.nlm.nih.gov/books/NBK557536/
^ Feng Y., Xiong Y., Qiao T., Li X., Jia L., Han Y. Lactate dehydrogenase A: a key player in carcinogenesis and potential target in cancer therapy. Cancer Med 2018;7(12):6124-6136. doi:10.1002/cam4.1820
^ abc Forkasiewicz A., Dorociak M., Stach K., Szelachowski P., Tabola R., Augoff K. The usefulness of lactate dehydrogenase measurements in current oncological practice. Cell Mol Biol Lett 2020;25:35. doi:10.1186/s11658-020-00228-7
^ Gleeson T.T. Post-exercise lactate metabolism: a comparative review of sites, pathways, and regulation. Annu Rev Physiol 1996;58:565-81. doi:10.1146/annurev.ph.58.030196.003025
^ Gray L.R., Tompkins S.C., Taylor E.B. Regulation of pyruvate metabolism and human disease. Cell Mol Life Sci. 2014 Jul;71(14):2577-604. doi:10.1007/s00018-013-1539-2
^ Holmes R.S., Goldberg E. Computational analyses of mammalian lactate dehydrogenases: human, mouse, opossum and platypus LDHs. Comput Biol Chem. 2009;33(5):379-85. doi:10.1016/j.compbiolchem.2009.07.006
^ Jin S., Chen X., Yang J., Ding J. Lactate dehydrogenase D is a general dehydrogenase for D-2-hydroxyacids and is associated with D-lactic acidosis. Nat Commun 2023;14(1):6638. doi:10.1038/s41467-023-42456-3
^ ab Krieg A.F., Rosenblum L.J., Henry J.B. Lactate dehydrogenase isozymes a comparison of pyruvate-to-lactate and lactate-to-pyruvate assays. Clin Chem 1967;13(3):196-203. doi:10.1093/clinchem/13.3.196
^ abc Laughton J.D., Charnay Y., Belloir B., Pellerin L., Magistretti P.J., Bouras C. Differential messenger RNA distribution of lactate dehydrogenase LDH-1 and LDH-5 isoforms in the rat brain. Neuroscience 2000;96(3):619-25. doi:10.1016/s0306-4522(99)00580-1
^ ab Li X., Yang Y., Zhang B., Lin X., Fu X., An Y., Zou Y., Wang J.X., Wang Z., Yu T. Lactate metabolism in human health and disease. Signal Transduct Target Ther 2022;7(1):305. doi:10.1038/s41392-022-01151-3
^ abc Markert C.L., Shaklee J.B., Whitt G.S. Evolution of a gene. Multiple genes for LDH isozymes provide a model of the evolution of gene structure, function and regulation. Science 1975;189(4197):102-14. doi:10.1126/science.1138367
^ Moran L.A., Horton H.R., Scrimgeour K.G., Perry M.D. Principles of Biochemistry. 5th Edition. Pearson, 2012
^ Nelson D.L., M. M. Cox M.M. Lehninger. Principles of biochemistry. 6th Edition. W.H. Freeman and Company, 2012
^ ab Read J.A., Winter V.J., Eszes C.M., Sessions R.B., Brady R.L. Structural basis for altered activity of M- and H-isozyme forms of human lactate dehydrogenase. Proteins 2001;43(2):175-85. doi:10.1002/1097-0134(20010501)43:2<175::AID-PROT1029>3.0.CO;2-%23
^ Rogatzki M.J., Ferguson B.S., Goodwin M.L., Gladden L.B. Lactate is always the end product of glycolysis. Front Neurosci 2015;9:22. doi:10.3389/fnins.2015.00022
^ abc Schueren F., Lingner T., George R., Hofhuis J., Dickel C., Gärtner J., Thoms S. Peroxisomal lactate dehydrogenase is generated by translational readthrough in mammals. Elife 2014;3:e03640. doi:10.7554/eLife.03640
^ Stambaugh R., Post D. Substrate and product inhibition of rabbit muscle lactic dehydrogenase heart (H4) and muscle (M4) isozymes. J Biol Chem 1966;241(7):1462-7. doi:10.1016/S0021-9258(18)96733-5
^ Wroblewski F, Gregory KF. Lactic dehydrogenase isozymes and their distribution in normal tissues and plasma and in disease states. Ann N Y Acad Sci 1961;94:912-32. doi:10.1111/j.1749-6632.1961.tb35584.x
^ abc Zinkham W.H., Blanco A., Clowry L.J. Jr. An unusual isozyme of lactic dehydrogenase in mature testes: localization, ontogeny, and kinetic properties. Ann N Y Acad Sci 1964;121:571-88. doi:10.1111/j.1749-6632.1964.tb14227.x
Keto acids or ketoacids are organic compounds containing two functional groups: a carboxyl acid group (−COOH) and a carbonyl group (˂C=O).
Based on the position of the carbonyl group relative to the carboxylic acid group, to which the IUPAC nomenclature rules assign the highest priority, ketoacids are classified as alpha-keto acids, beta-keto acids, and gamma-keto acids.[2]
Keto acids, and in particular alpha-keto acids, are very important in biochemistry, being involved in many metabolic pathways.[4]
Alpha-keto acids
They have the carbonyl group adjacent to the carboxylic acid. Many of these compounds, in the form of their conjugate bases, have important biological functions. Below are some examples. Pyruvic acid, the simplest alpha-keto acid, is the end metabolic product of glycolysis.
Oxaloacetic acid and alpha-ketoglutaric acid are intermediates of the citric acid cycle.
Alpha-keto acids can arise from transamination and oxidative deamination reactions of amino acids. In transamination reactions, the alpha amino group of the amino acid is transferred to an alpha-keto acid, usually alpha-ketoglutarate, with the formation of a new amino acid and an new alpha-keto acid. These reactions are catalyzed by enzymes called aminotransferases or transaminases (EC6.1.-).
alpha-Keto acid + Amino acid ⇄ New amino acid + New alpha-keto acid
In oxidative deaminations, amino acids are converted into the corresponding alpha-keto acids by removing the amino group, which is converted to ammonia and replaced by a carbonyl group. Since the reaction is reversible, ketoacids are also precursors of amino acids.
Note: ammonia is a toxic compound, and is converted into the safer compound urea via the urea cycle in the liver.[4]
Pyruvate, oxaloacetate and alpha-ketoglutarate, the latter via oxaloacetate, are the entry points into gluconeogenesis of the carbon skeleton of many glucogenic amino acids.[4]
It has also been observed that, in vitro, murine and human tumor cell lines secrete 2-chetoacids into the tumor microenvironment, such as α-ketoisocaproate, α-keto-β-methylvalerate and α-ketoisovalerate, which are capable to influence the anti-tumor activity of macrophages.[1]
Beta-keto acids
They have the carbonyl group at the second carbon from the carboxylic acid.
Examples of beta-keto acids are acetoacetic acid, the simplest one, and beta-hydroxybutyric acid, which are two of the three ketone bodies, together with acetone, produced by the hepatocyte when acetyl-CoA is produced in excess of the capacity of citrate synthase (EC 2.3.3.1), namely, of citric acid cycle to oxidize it fully, as during prolonged fasting or diets very low in carbohydrates.
Note that acetoacetyl-CoA and beta-hydroxybutyryl-CoA, namely, the activated forms of these beta-keto acids, are also intermediates in the butyric acid synthesis pathway which occurs in most butyrate-producing bacteria of the gut microbiota.[3][5][6]
Gamma-keto acids
They have the carbonyl group at the third carbon from the carboxylic acid.
An example is levulinic acid, the simplest one, which arises from the catabolism of cellulose.
References
^ Cai Z., Li W., Brenner M., Bahiraii S., Heiss E.H., Weckwerth W. Branched-chain ketoacids derived from cancer cells modulate macrophage polarization and metabolic reprogramming. Front Immunol 2022;13:966158. doi:10.3389/fimmu.2022.966158
^ Miller T.L., Wolin M.J. Pathways of acetate, propionate, and butyrate formation by the human fecal microbial flora. Appl Environ Microbiol 1996;62(5):1589-92. doi:10.1128/aem.62.5.1589-1592
^ abc Nelson D.L., Cox M.M. Lehninger. Principles of biochemistry. 6th Edition. W.H. Freeman and Company, 2012
^ Portincasa P., Bonfrate L.,Vacca M., De Angelis M., Farella I., Lanza E., Khalil M.,Wang D.Q.-H., Sperandio M., Di Ciaula A. Gut microbiota and short chain fatty acids: implications in glucose homeostasis. Int J Mol Sci 2022;23:1105. doi:10.3390/ijms23031105
^ Pryde S.E., Duncan S.H., Hold G.L., Stewart C.S., Flint H.J. The microbiology of butyrate formation in the human colon. FEMS Microbiol Lett 2002;217(2):133-9. doi:10.1111/j.1574-6968.2002.tb11467.x
A futile cycle, or substrate cycle, occurs when two non-equilibrium opposing reactions, catalyzed by different enzymes, or two opposite metabolic pathways run simultaneously with no other overall effect than the dissipation of energy.[4]
The name of futile cycle was coined because apparently these cycles seemed to confer no benefit to the cell, a sort of metabolic imperfection leading to energy expenditure.[1][3] However, recently, they have been recognized as important for the generation of heat, the amplification of metabolic signals, the redistribution of the energy, in form of triglycerides, between adipocytes and hepatocytes, and the modification of fatty acids stored as triglycerides in adipose tissue.[1][5][6][7]
In order to avoid uncontrolled dissipation of energy, futile cycles are strictly regulated.
Examples of futile cycles are glycolysis and gluconeogenesis when they proceed simultaneously at a high rate in the same cell, the Cori cycle, and the triglyceride/fatty acid cycle.[2]
Let us consider the conversion of A into B, which proceeds at a rate of 100, and B into A, which proceeds at a rate of 90. This results in a net flow of 10. Suppose that an effector increases the rate of conversion of A into B by 30 percent, to 130 percent, and reduces the rate of conversion of B into A by 30 percent, to 63 percent. The resulting net flux is equal to 130-63 = 67, namely, a 30 percent change in the rates of the opposing reactions has led to a 570 percent increase in the net flux.[1]
A mechanism of this type could, at least in part, explain the even 1000-fold increase in carbon flux down the glycolysis in the initial phase of intense exercise.
Regulation
In the course of evolution, the selection of different enzymes to catalyze irreversible and opposing reactions has made possible to avoid or put under strict control futile cycles. How? The selection of one enzyme that catalyzes the conversion of A into B, and another enzyme that catalyzes the opposing reaction, whose activities are regulated separately, allows the control of the net flux.[3][4] Enzymatic activities are controlled by:
allosteric mechanisms;
covalent modifications;
modifications in the concentration of the enzymes, due to variations in the ratio between their synthesis and/or degradation. A different mechanism regulates hepatic glucokinase (EC 2.7.1.2). During fasting, the enzyme is reversibly bound to GKPR, one of the liver-specific proteins, which anchors it inside the nucleus, separating it from the other glycolytic enzymes, and thus preventing the futile cycle between glycolysis and gluconeogenesis.
In this way, it is possible to obtain a coordinated regulation of the two opposing pathways, thus avoiding an uncontrolled futile cycle. Obviously, such a fine regulation could not be achieved if a single enzyme would operate in both directions.
Glycolysis and gluconeogenesis
If glycolysis, which converts glucose into pyruvate, the conjugate base of pyruvic acid, with the production of ATP, and gluconeogenesis, which converts pyruvate into glucose with the consumption of ATP, run simultaneously at high rate in the same cell, the net result would be a net consumption of ATP, therefore a futile cycle. This is avoided by the control of the irreversible steps of the two metabolic pathways, in particular the reactions catalyzed by phosphofructokinase-1 or PFK-1 (EC 2.7.1.11), and by fructose-1,6-bisphosphatase or FBPase (EC 3.1.3.11), mainly by the allosteric effector fructose 2,6-bisphosphate.[3][4]
It should be noted that in glycolysis, the control involves all the irreversible reactions, whereas in gluconeogenesis, the key regulatory points are the reactions catalyzed by pyruvate carboxylase (EC 6.4.1.1) and fructose 1,6-bisphosphatase.
Cori Cycle
In the Cori cycle, lactic acid produced from glucose in the muscle and other extrahepatic tissues reaches the liver, where it is converted back into glucose, which, released into the circulation, returns to the muscle and other extrahepatic tissues, thereby closing the cycle. From an energetic point of view, the Cori cycle can be considered a futile cycle because it results in a net consumption of 4 ATP with no other overall effect.[2] However, it allows many different types of extrahepatic cells to work at the expense of the liver.
Triglyceride/fatty acid cycle
In the triglyceride/fatty acid cycle, triglycerides in adipose tissue are partially or completely hydrolyzed to free fatty acids and glycerol, in a process called lipolysis; the released fatty acids are then used to resynthesize new molecules of triglycerides.[2][4][5] Four moles of ATP are consumed for every mole of triglycerides that completes the cycle.
This futile cycle can take place:
between adipose tissue, which releases fatty acids, and the liver, which re-esterified them to triglycerides, leading to a redistribution of stored energy;[6]
in adipocytes, where it may contribute to thermogenesis and modifications of the stored fatty acids.
Regarding the modifications, the cycle renders fatty acids accessible for re-arrangements such as elongations and desaturations, which allow saturated fatty acids to be converted to unsaturated fatty acids. However, the efficiency of the process seem to depend on the type of fatty acids.[7] For example, the metabolism of the released medium-chain fatty acids is faster than the conversion of palmitic acid, one of the long-chain fatty acids, to palmitoleic acid, oleic acid, and then, in hepatocytes, to arachidonic acid.
Note that the conversion of medium-chain fatty acids and palmitic acid to long-chain unsaturated fatty acids reduces the health risk associated to their accumulation in stored triglycerides.
Generation of heat
In some cases a futile cycle has the only function of producing heat through the hydrolysis of ATP. This occurs, for example, in the flight muscles of bumblebees, which, in order to fly, must maintain a thoracic temperature of about 30 °C, even when the external temperature is 10 °C.[1] The thoracic temperature is maintained at the optimal levels for flight thanks to the futile cycle between the reactions catalyzed by PFK-1 and FBPase. In fact, flight muscle FBAase is not inhibited by AMP, which suggests that, during evolution, this protein has been selected for the generation of heat.
Unlike bumblebees, flight muscles of honeybees contain almost no FBPase, and therefore these insects cannot fly when the external temperature is low.
References
^ abcd Berg J.M., Tymoczko J.L., and Stryer L. Biochemistry. 5th Edition. W. H. Freeman and Company, 2002
^ abc Brownstein A.J., Veliova M., Acin-Perez R., Liesa M., Shirihai O.S. ATP-consuming futile cycles as energy dissipating mechanisms to counteract obesity. Rev Endocr Metab Disord 2022;23(1):121-131. doi:10.1007/s11154-021-09690-w
Short-chain fatty acids or SCFAs are saturated fatty acids with a straight or branched carbon-chain made of 2-5 carbon atoms, and are acetic acid, propionic acid, butyric acid, isobutyric acid, valeric acid, isovaleric acid, and 2-methylbutyric acid.[1]
In humans, they are, along with secondary bile salts, the main metabolites produced by bacteria of the gut microbiota in the cecum and colon, and derive almost entirely from the anaerobic fermentation of non-digestible carbohydrates.[11] The most abundant are acetic acid, propionic acid and butyric acid, which represent 90-95 percent of the produced SCFAs.[6] The remaining percentage is made of the branched SCFAs.
They are the major anions present in the colon. Their concentration is higher in the cecum and in the proximal colon than in the distal part, where the substrates for their synthesis are depleted.[1][2][4] They are able to reduce colonic pH value and thus acidify the stool.
About 90-95 percent of the SCFAs are absorbed in the cecum and colon, whereas 5-10 percent are excreted with the feces.[13]
They are thought to provide about 70 percent of the energy needs of colonocytes.[4]
Short-chain fatty acids are able to modulate the physiology and composition of the gut microbiota.[7] Furthermore, a growing body of research suggests that they play a important role in maintaining human health.[4]
Like medium-chain fatty acids and long-chain fatty acids, short-chain fatty acids are present in animal and plant tissues mostly in the form of triglycerides, although in much lower amounts than long-chain ones.
In adults, the main food source is milk and dairy products, where butyric acid is the SCFA present with the highest concentration. Other sources are some vegetable oils, such as palm kernel oil and coconut oil.
In breastfed infants, the main source is breast milk.[10]
However, for humans, and most mammals, the most important source is the anaerobic fermentation of fibers and resistant starch, namely, indigestible carbohydrates, by the bacteria of the gut microbiota.[6] Approximately 500-600 mM of SCFAs are produced through this pathway per day. Acetic, propionic and butyric acids are present in a molar ratio of about 60:20:20, respectively, although the relative proportion of each depends on the microbiota composition, the substrate, and the intestinal transit time.[7][11]
Properties
Short-chain fatty acids have carbon chains made of 2-5 carbon atoms, a characteristic that strongly affects the physical properties.[9]
Acetic acid, propionic acid, butyric acid and valeric acid are straight-chain fatty acids, whereas isobutyric acid, isovaleric acid and 2-methylbutyric acid are branched-chain fatty acids.
They are small molecules, and are the smallest among all lipids.
They are liquid at room temperature, and are soluble in polar solvents such as water, unlike saturated fatty acids with longer carbon chains, whose solubility in polar solvents, considering those with straight chain, decreases as the length of the chain increases, as the hydrophobic part of the molecule is the carbon chain, whereas the carboxyl group is polar.[10]
Finally, it should also be noted that butyric acid and isobutyric acid, which have the molecular formula C4H8O2, are an example of chain isomerism, as well as valeric acid, isovaleric acid and 2-methylbutyric acid, which have the formula molecular C5H10O2.
Health effects
Short-chain fatty acids appear to play a crucial role in maintaining human health.[4] Their activity seems to occur through direct and/or indirect effects on cellular processes such as proliferation, differentiation and gene expression, thus contributing to the regulation of processes such as glucose homeostasis, intestinal and immune function, and the regulation of the gut-brain axis.[6] Their health effects seem to be confirmed also by studies showing that intestinal dysbiosis appears to be implicated in metabolic pathologies, such as disorders involving glucose homeostasis, and behavioral and neurological pathologies, such as depression, and Alzheimer’s and Parkinson’s.[11]
Synthesis
In humans, the enzyme equipment carrying out carbohydrate digestion lacks the enzymes capable of digesting fiber and resistant starch, the latter so called precisely because it resists the action of alpha-amylase. On the contrary, the bacteria of the gut microbiota code for a large number of different glycoside hydrolases, more than 260, which also hydrolyze fibers and resistant starch, releasing the constituent monosaccharides.[7] Hexoses and deoxyhexoses enter glycolysis, and pentoses enter the pentose phosphate pathway, to give pyruvate, the conjugate base of pyruvic acid, which is the main precursor for the synthesis of short-chain fatty acids.[2][4][5]
The synthesis of SCFAs is affected by several factors; below are some examples.[4][11][13]
The fiber content of the diet. For example, a diet rich in fibers, such as the Mediterranean diet may influence their synthesis.
The composition of the gut microbiota.
The pH of the intestinal lumen, as the bacteria that produce butyric acid dominate at pH value around 5.5, while the bacteria that produce acetic and propionic acids dominate at pH value around 6.5.
The gut transit time.
The amount of oxygen in the intestinal lumen.
Acetic acid and propionic acid are mainly produced by species of the phylum Bacteroides, while butyric acid, for whose synthesis resistant starch is particularly important, by species of the phylum Firmicutes.[6]
Synthesis of acetic acid
Acetic acid, the most abundant SCFA in the colon, can be synthesized via the Wood-Ljungdahl pathway in the reductive direction, through the reduction of CO2 to acetate, or from acetyl-CoA, the most important metabolic pathway, responsible of the production of about two thirds of butyric acid present in the intestinal lumen.[4]
Synthesis of propionic acid
Propionic acid can be synthesized through three different metabolic pathways: the acrylate and succinate pathways, which use lactic acid produced by other bacteria, and the propanediol pathway, in which the precursors are deoxyhexoses.[2][3][10]
The acrylate pathway converts lactic acid to propionyl-CoA, via lactoyl-CoA. In the final step, propionyl-CoA is hydrolyzed to propionic acid.
In the succinate pathway, lactate is reduced to pyruvate, which is carboxylated to oxaloacetate, which, through a pathway that has malate, fumarate, succinate, and methylmalonyl-CoA as intermediates, is converted to propionyl-CoA, which in turn is hydrolyzed to propionic acid. The succinate pathway is thought to be the dominant pathway for propionic acid synthesis in the gut.
In the propanediol pathway, some deoxyhexoses, such as fucose and rhamnose, are converted via 1,2-propanediol to propionyl-CoA and then propionic acid.
Synthesis of butyric acid
The synthesis of butyric acid can follow two routes.[7][10]
In most butyric acid-producing bacteria, the short-chain fatty acid is synthesized through a pathway that begins with the condensation of two acetyl-CoA to acetoacetyl-CoA, which, through a pathway that has beta-hydroxybutyryl-CoA and crotonyl-CoA as intermediates, is converted to butyryl-CoA. The final step is the release of butyric acid from butyryl-CoA.[8]
In a small number of bacterial species, butyryl-CoA is converted to butyryl phosphate, from which butyric acid is released.[2]
Synthesis from amino acids
Acetic acid, propionic acid and butyric acid can also be produced from amino acids obtained from peptide and protein degradation, although the amount produced by these pathways is small.[7]
These synthesis occur in the distal part of the colon, often by non-commensal bacteria, as in the case of glutamate and lysine fermentation.[3] Different short-chain fatty acids are produced by the metabolism of different amino acids; below are some examples.[4]
Glutamic acid mainly produces acetic acid and butyric acid.
Aspartic acid mainly produces acetic acid and propionic acid.
The basic amino acids lysine, arginine and histidine produce acetic acid and butyric acid.
Cysteine produces acetic, propionic and butyric acids.
Methionine mainly produce propionic acid and butyric acid.
Branched-chain fatty acids derive from the branched-chain amino acids leucine, isoleucine and valine.
The pH of the intestinal lumen influences the metabolism of proteins by the gut microbiota; for example, their breakdown into amino acids is more likely at neutral or weakly alkaline pH.
It should be noted that potentially toxic compounds, such as ammonia, sulphites and phenols, are also produced from the intestinal metabolism of amino acids.
Endogenous synthesis
Mammals, and therefore humans, have the enzymatic equipment for the endogenous synthesis of short-chain fatty acids. The synthesis occurs mainly in the liver, by beta-oxidation cycles which lead to the formation of acyl-CoA with a shorter carbon chain than the starting fatty acid. The acil-CoA is then hydrolyzed to fatty acid and CoA by an acyl-CoA thioesterases (EC 3.1.2.20).[10][12]
Membrane receptors
Short-chain fatty acids can bind to specific receptors on the plasma membrane, the G protein-coupled receptors, including GPR41, GPR43 and GPR109A.[6]
The effects produced by the binding of the SCFAs on the receptors depend on the cell type. For example, binding to receptors on intestinal L cells is associated with the release of glucagon-like peptide-1, or GLP-1, and peptide YY, hormones that affect appetite and food intake. The binding to enterochromaffin cells induces the release of serotonin, which may affect intestinal motility. Finally, binding to receptors on pancreatic beta-cells increases insulin release.[11]
The different short-chain fatty acids have different ability to activate the receptors: GPR43 is more likely to be activated by acetic and propionic acids, GPR41 by propionic and butyric acids, while GPR109A by butyric acid.[4]
Absorption
About 90 percent of the short-chain fatty acids present in the intestinal lumen are absorbed by colonocytes. The passage across the plasma membrane can occur by passive diffusion or by active transport mediated by two types of membrane transporters: the H+-dependent and Na+-dependent monocarboxylate transporters, or MCTs and SMCTs, respectively.[6][7]
Passive transport affects protonated forms of SCFAs, so it is influenced by colonic pH value. A weak acidification of the intestinal lumen, which can be due to the metabolic activity of the microorganisms, increases the prevalence of the protonated form and therefore of passive transport.[10]
Role in colonocytes
In colonocytes, short-chain fatty acids have energy and regulatory function.
When used for energy purposes, acetic acid and butyric acid are converted to acetyl-CoA, and propionic acid to propionyl-CoA. Through the production of ATP, SCFAs contribute to the maintenance of cellular homeostasis, but also, for example, to the maintenance of the integrity of the tight junctions at the cell apex, and therefore of the integrity of the intestinal barrier.[11] Of the three major short-chain fatty acids produced by gut microbiota, butyric acid is the major source of energy for colonocytes, while acetic and propionic acids are poorly metabolized and mostly drained by the portal vein.[2][13]
Considering the regulatory role, SCFAs are, for example, capable of inhibiting histone deacetylases (EC 3.5.1.98), enzymes that catalyze the removal of acetyl groups from lysine residues of histone proteins, acetyl groups previously inserted by histone acetyltransferase (EC 2.3.1.48).[7] The R groups of deacetylated lysines have positive charges, which allows histone proteins to wrap the DNA more tightly. This makes the nucleosome more compact, and consequently more difficult to carry out transcription and gene expression. The different short-chain fatty acids have different abilities to inhibit histone deacetylases:
This mode of action on histone deacetylases has been observed not only in the gut and associated immune tissue, but also in the central and peripheral nervous systems.[11]
Transport
Short-chain fatty acids that have not been utilized by colonocytes leave the cell by passive diffusion and active transport across the basolateral membrane, and enter the portal circulation where acetic acid reaches the highest concentration, about 260 mM/L, while propionic and butyric acids reach concentrations of about 30 mM/L.[2]
In the rectum, a small amount of these lipids can pass directly into the systemic circulation, thus bypassing the liver, via the internal iliac vein.[13]
Unlike long-chain fatty acids, short- and medium-chains fatty acids are present in the circulation in free form, namely, as non-esterified fatty acids, and, bound to albumin, reach the liver. Subsequent cell uptake and intracellular transport do not require fatty acid transport proteins, plasma membrane fatty acid translocases, or cytosolic fatty acid binding proteins. Therefore, their oxidation may be much faster than that of long-chain fatty acids and the longest members of medium-chain fatty acids, namely, fatty acid with carbon chains longer than 8 carbon atoms.[10]
Hepatic and extrahepatic metabolism
The liver is an important site for short-chain fatty acids metabolism.[13]
It can to absorb about 40 percent of the acetic acid and 80 percent of the propionic acid from the portal vein. Propionic acid is mostly metabolized in the liver, where it can also be used as a substrate for gluconeogenesis.[7]
A small amount of the gut derived SCFAs, about 36 percent for acetic acid, 9 percent for propionic acid and only 2 percent for butyric acid, reach, through the systemic circulation, the peripheral tissues. In muscle, acetic acid can be used for lipid synthesis or be oxidized for energy production. Furthermore, it is thought that SCFA concentrations in the systemic circulation, even if small, are capable of influencing the metabolism and physiology of peripheral cells and tissues.
References
^ ab Abdul Rahim M.B.H., Chilloux J., Martinez-Gili L., Neves A.L., Myridakis A., Gooderham N., Dumas M.-E. Diet-induced metabolic changes of the human gut microbiome: importance of short-chain fatty acids, methylamines and indoles. Acta Diabetol 2019;56:493-500. doi:10.1007/s00592-019-01312-x
^ abcde Deleu S., Machiels K., Raes J., Verbeke K., Vermeire S. Short chain fatty acids and its producing organisms: an overlooked therapy for IBD? EBioMedicine 2021;66:103293. doi:10.1016/j.ebiom.2021.103293
^ ab Koh A., De Vadder F., Kovatcheva-Datchary P., Bäckhed F. From dietary fiber to host physiology: short-chain fatty acids as key bacterial metabolites. Cell 2016;165(6):1332-1345. doi:10.1016/j.cell.2016.05.041
^ abcdefghij Liu X.F., Shao J.H., Liao Y.T., Wang L.N., Jia Y., Dong P.J., Liu Z.Z., He D.D., Li C., Zhang X. Regulation of short-chain fatty acids in the immune system. Front Immunol 2023;14:1186892. doi:10.3389/fimmu.2023.1186892
^ Miller T.L., Wolin M.J. Pathways of acetate, propionate, and butyrate formation by the human fecal microbial flora. Appl Environ Microbiol 1996;62(5):1589-92. doi:10.1128/aem.62.5.1589-1592
^ abcdef Parada Venegas D., De la Fuente M.K., Landskron G., González M.J., Quera R., Dijkstra G., Harmsen H.J.M., Faber K.N., Hermoso M.A. Short chain fatty acids (SCFAs)-mediated gut epithelial and immune regulation and its relevance for inflammatory bowel diseases. Front Immunol 2019;10:1486. doi:10.3389/fimmu.2019.00277
^ abcdefgh Portincasa P., Bonfrate L., Vacca M., De Angelis M., Farella I., Lanza E., Khalil M.,Wang D.Q.-H., Sperandio M., Di Ciaula A. Gut microbiota and short chain fatty acids: implications in glucose homeostasis. Int J Mol Sci 2022;23:1105. doi:10.3390/ijms23031105
^ Pryde S.E., Duncan S.H., Hold G.L., Stewart C.S., Flint H.J. The microbiology of butyrate formation in the human colon. FEMS Microbiol Lett 2002;217(2):133-9. doi:10.1111/j.1574-6968.2002.tb11467.x
^ abcdefg Schönfeld P., Wojtczak L. Short- and medium-chain fatty acids in energy metabolism: the cellular perspective. J Lipid Res 2016;57(6):943-54. doi:10.1194/jlr.R067629
^ abcdefg Silva Y.P., Bernardi A., Frozza R.L. The role of short-chain fatty acids from gut microbiota in gut-brain communication. Front Endocrinol (Lausanne) 2020;11:25. doi:10.3389/fendo.2020.00025
^ abcde Xiong R.G., Zhou D.D., Wu S.X., Huang S.Y., Saimaiti A., Yang Z.J., Shang A., Zhao C.N., Gan R.Y., Li H.B. Health benefits and side effects of short-chain fatty acids. Foods 2022;11(18):2863. doi: 10.3390/foods11182863
Starch phosphorylase or alpha-glucan phosphorylase (EC 2.4.1.1) is a multimeric protein, with enzymatic and regulatory activity, that plays an important role in carbohydrate metabolism, both in prokaryotes and eukaryotes.[10]
The enzyme catalyzes the transfer of a glucosyl unit from glucose 1-phosphate to the non-reducing end of a nascent α-(1→4)-glucan, to which glucose is linked by an α-(1→4) glycosidic linkage.[6][8]The reaction is reversible and the direction depends on the phosphate/glucose-1-phosphate ratio present in vivo.[2] The enzyme, like starch synthase (EC 2.4.1.21), which is involved in the synthesis of amylose and amylopectin, the polysaccharides which make up starch granules, glycogen phosphorylase (EC 2.4.1.1), an enzyme involved in glycogenolysis, and glycogen synthase (EC 2.4.1.11), which is involved in glycogen synthesis, belongs to the family of glucosyltransferases (EC 2.4).[6]
Note that, while starch synthase uses ADP-glucose as glucosyl donor, and glycogen synthase uses UDP-glucose, starch phosphorylase uses glucose 1-phosphate.[2]
Starch phosphorylase appears to be involved in both the synthesis and degradation of amylose and amylopectin.[9]
Industrially, the phosphorolytic action of starch phosphorylase is used in the production of glucose-1-phosphate and in the preparation of carbohydrates such as glucans and modified starches.[6]
At least two isoforms of starch phosphorylase are present in plants, one in the stroma of plastids, named Pho1, and Pho2, with cytosolic localization. Both isoforms play a critical role in the synthesis and degradation of starch.[6][8]
Starch degradation
Although alpha-amylase (EC 3.2.1.1) is the first enzyme to intervene in the polysaccharide degradation during the early stages of germination, and beta-amylase (EC 3.2.1.2) is the first enzyme to intervene in the transient starch degradation in chloroplasts, other enzyme activities have also been implicated, such as alpha-glucan water dikinase (EC 2.7.9.4) and phospho-glucan water dikinase (EC 2.7.9.5), and the debranching enzyme.[10]
Of the two isoenzymes of starch phosphorylase, Pho1 seems to have an indirect or regulatory action capable of influencing the activity of the other enzymes involved in starch degradation, while Pho2 is capable of degrading starch granules and other branched glucans.
Starch synthesis
During starch biosynthesis, starch phosphorylase, particularly Pho1, appears to perform both an enzymatic and regulatory action.
Pho1 seems to be involved in the initial steps of starch synthesis, contributing to the elongation of the nascent chain.[1][8]
Starch synthesis involves a set of at least five classes of enzymes, namely ADP-glucose pyrophosphorylase (EC 2.7.7.27), starch synthases, starch branching enzymes (EC 2.4.1.18), starch debranching enzymes (EC 3.2.1.41) and starch phosphorylases.[5][10] To these must be added proteins with no catalytic activity but needed for the correct synthesis of the starch granule. In the endosperm of cereals, the formation of multienzyme complexes between starch synthases and branching enzymes depends not only on specific phosphorylations of the proteins, but also on the presence of Pho1.[1][3]
Starch phosphorylase is capable of forming a complex with disproportionating enzyme (EC 2.4.1.25).[1][10] Such complex seems capable of synthesizing short malto-oligosaccharides or MOS, which are α-(1→4)-glucans with a degree of polymerization between 2 and 7.[7] MOS act as primers for starch synthase IV and granule-bound starch synthase in the initial steps of amylopectin and amylose synthesis, respectively, a role which resembles that of glycogenin in glycogen synthesis.[4] MOS can also originate from the activity of starch debranching enzymes during the trimming of amylopectin molecules.
References
^ abc Crofts N., Abe N., Oitome N.F., Matsushima R., Hayashi M., Tetlow I.J., Emes M.J., Nakamura Y., Fujita N. Amylopectin biosynthetic enzymes from developing rice seed form enzymatically active protein complexes. J Exp Bot 2015;66(15):4469-82. doi:10.1093/jxb/erv212
^ ab Orzechowski S. Starch metabolism in leaves. Acta Biochim Pol 2008;55(3):435-45. doi:10.18388/abp.2008_3049
^ Pfister B., Zeeman S.C., Rugen M.D., Field R.A., Ebenhöh O., Raguin A. Theoretical and experimental approaches to understand the biosynthesis of starch granules in a physiological context. Photosynth Res 2020;145:55-70. doi:10.1007/s11120-019-00704-y
^ Qu J., Xu S., Zhang Z., Chen G., Zhong Y., Liu L., Zhang R., Xue J., Guo D. Evolutionary, structural and expression analysis of core genes involved in starch synthesis. Sci Rep 2018;8(1):12736. doi:10.1038/s41598-018-30411-y
^ abcd Rathore R.S., Garg N., Garg S., Kumar A. Starch phosphorylase: role in starch metabolism and biotechnological applications. Crit Rev Biotechnol 2009;29(3):214-24. doi:10.1080/07388550902926063
^ Tetlow I.J., Bertoft E. A review of starch biosynthesis in relation to the building block-backbone model. Int J Mol Sci 2020;21(19):7011. doi:10.3390/ijms21197011
^ abc Tickle P., Burrell M.M., Coates S.A., Emes M.J., Tetlow I.J., Bowsher C.G. Characterization of plastidial starch phosphorylase in Triticum aestivum L. endosperm. J Plant Physiol 2009;166(14):1465-78. doi:10.1016/j.jplph.2009.05.004
^ Voet D. and Voet J.D. Biochemistry. 4th Edition. John Wiley J. & Sons, Inc. 2011
^ abcd Yu G., Shoaib N., Xie Y., Liu L., Mughal N., Li Y., Huang H., Zhang N., Zhang J., Liu Y., Hu Y., Liu H., Huang Y. Comparative study of starch phosphorylase genes and encoded proteins in various Monocots and Dicots with emphasis on maize. Int J Mol Sci 2022;23:4518. doi:doi.org/10.3390/ijms23094518
Amylopectin is a highly branched polysaccharide made up of alpha-D-glucose units. Together with amylose, it is one of the two main constituents of starch granules, the means by which plants store energy and the most widespread and abundant form of carbohydrate storage on Earth.[12]
Glucose monomers are linked by α-(1→4) glycosidic bonds to form chains to which the branches are linked by α-(1→6) glycosidic bonds.[2]
Its synthesis requires the coordinated action of at least four distinct classes of enzymes: starch synthases (EC 2.4.1.21), starch branching enzymes (EC 2.4.1.18), starch debranching enzymes, and starch phosphorylase (EC 2.4.1.1).[17]
In starch granules, amylopectin is present in greater amount than amylose, and forms a semi-crystalline matrix within which amylose seems to be stored.
The amylose/amylopectin ratio significantly influences the physicochemical properties of starch, and consequently both its industrial applications, such as the production of food additives, and the possible health effects.[3]
Amylopectin is a highly branched polysaccharide, and has a molecular weight in the order of 107-108 Daltons, therefore much larger than amylose. It is formed of 104-105 glucose molecules which are linked by α-1,4 glycosidic bonds to form many relatively short chains, whose degree of polymerization is of 18-25 molecules.[17] The length of the chains vary depending on the source of the starch as well as the environmental and nutrient conditions during plant growth and seed formation.
The chains are interconnected by α-1,6 glycosidic bonds to form a tree-like architecture, with neighboring chains forming cluster-like structures.[1]
In most starches, α-1,6 glycosidic bonds account for about 5 percent of all glycosidic bonds, a lower percentage than that found in glycogen molecule, about 9 percent, where the branches are more evenly distributed. The length and distribution of the branches directly affect the physicochemical properties of amylopectin, such as solubility, viscosity, ease of retrogradation, and gelatinization and pasting temperature.[3] For example, glycogen is water soluble whereas amylopectin and starch are not.
Amylopectin chains
Amylopectin chains can be classified on the basis of their length or the presence or absence of branches.
The classification according to the length identifies two main types of chains: short and long chains. Short chains have a degree of polymerization of 6-36 glucosyl units, although the upper limit depends on the source of amylopectin, whereas long chains have a degree of polymerization greater than or equal to 36. In most starches, the molar distribution of long to short chains is about 19-6, and is generally higher in A-crystalline starches, such as cereal endosperm starches, than in B-crystalline starches, such as those in potatoes.[17]
The classification on the basis of their connections to other chains identifies three categories: A-chains, B-chains and C-chains.[7]
A-chains carry no branches, are short chains, with a degree of polymerization of about 13, and are the external chains.
B-chains contain at least one branch, namely, A- and/or B-chain, are longer than A-chains, and are present in the inner part of the molecule. B-chains are in turn divided into B1-chains, with a polymerization degree of about 22, B2-chains, with a polymerization degree of about 42, B3-chains, with a polymerization degree of about 69, B4 and so on.
C-chain is the B-chain which carries the sole reducing end.
A-chains and B1-chains participate in the formation of the clusters, while B2-, B3-, and B4-chains are thought to extend through two, three, and four clusters, respectively.[7][11]
A-type, B-type and C-type polymorphs
Within the clusters, neighboring linear chain segments form double helices, which run parallel to each other, with a period of 2.1 nm, and where each turn has six glucose molecules per chain.[10] The double helices form two types of crystalline structures called A-type polymorph, more dense and typical of cereal grains, and B-type polymorph, with a hexagonal structure, less dense, more hydrated, and typical of tuber and high amylose starches. Finally, C-type polymorph is a mixture of A-type and B-type polymorphs, and is found in the starches of root, legume and some fruits.[2]
This organization underlies the semi-crystalline nature of starch granules.
Growth rings
Although starch granules have different shapes, their internal architecture is remarkably conserved between different species. In fact, when viewed under a microscope, most starches exhibit a regular pattern of light and dark rings known as growth rings, so called because they resemble the growth rings of trees.[10]
The growth rings surround the hilum of the granule, namely, the core of the granule, whose exact structure is not known, although it appears to be formed by a relatively disordered alpha-glucan structure. The rings have a thickness of 200-400 nm, and are the result of alternating amorphous regions, less dense, and semi-crystalline regions, more dense.[10]
According to the cluster model, the semi-crystalline regions are due to the alternation of crystalline and amorphous lamellae, stacked with a periodicity of about 9-10 nm.[11] The crystalline lamellae are made up by the linear chains of amylopectin arranged to form the A-type, B-type or C-type polymorphs, and extend for about 6-7 nm, while the amorphous lamellae contain most of the branch points and extend for about 3nm.[5]
Amylopectin synthesis
The synthesis of amylopectin is believed to start from the hilum.[22]
The synthesis requires the coordinated activities of at least four distinct groups of enzymes: starch synthases, starch phosphorylase, starch branching enzymes and starch debranching enzymes.[12] Each group consists of several isoforms with distinct biochemical properties.
Like amylose synthesis, amylopectin synthesis requires short malto-oligosaccharides or MOS, α-(1→4)-glucans with a degree of polymerization of 2 to 7, which, acting as primers, are elongated by starch synthase, similarly to what happen with glycogenin in the initial steps of glycogen synthesis.[16]
MOS seem to have different origins, all of which require the activity of some enzymes involved in the synthesis of starch granules, namely:
starch synthase III and starch phosphorylase, the latter in combination with the disproportionating enzyme (EC 2.4.1.25), which use ADP-glucose and glucose-1-phosphate as substrates, respectively;
starch debranching enzymes, during the trimming of other amylopectin molecules.[16]
As MOS are poor solubility in aqueous environments, they seem to be able to evade the hydrolytic action of alpha-amylase (EC 3.2.1.1) and beta-amylase (EC 3.2.1.2).
The coordination, both spatial and temporal, of the involved enzymes, which in many cases also physically interact to form multienzyme complexes, is essential to allow the conversion of photosynthetic reaction products into the organized and insoluble structure of the polysaccharide. And, as in the case of amylose and glycogen synthesis, the polymerization of glucose into amylopectin, and more generally of osmotically active monosaccharides into osmotically inactive polysaccharides allows the storage of large amounts of monosaccharides inside the cell without any substantial increase in osmotic pressure.
Starch synthase
Six isoforms of starch synthase are known, all structurally related, of which five are involved in amylopectin synthesis, isoforms referred to as starch synthase I, II, III, IV and V or SSI, SSII, SSIII, SSIV and SSV, respectively, present in the stroma of plastids or portioned between the stroma and starch granules, whereas the sixth, the granule-bound starch synthase or GBSS (EC 2.4.1.242), is almost exclusively bound to granules and is involved in the synthesis of amylose.[10]
The first four isoforms have catalytic activity and belong, like GBSS, glycogen phosphorylase (EC 2.4.1.1) and glycogen synthase (EC 2.4.1.11), enzymes involved in glycogenolysis and glycogen synthesis, respectively, to the family of glycosyltransferases (EC 2.4). Conversely, starch synthase V has no catalytic activity.
During amylopectin synthesis, starch synthase catalyzes the adding of one glucose residue to the non-reducing end of a pre-existing α-(1→4) linked glucan chain. The monosaccharide is linked by a α-(1→4) glycosidic bond.[5]
Note that, unlike glycogen synthase, starch synthase uses ADP-glucose and not UDP-glucose as the glucosyl donor.
The mode of action of starch synthase I, II, III, and IV is different from that of GBSS in that they are able to catalyze the addition of only one glucose unit per substrate encounter, a mode of action defined as distributive, whereas GBSS is able to catalyze the addition of more than one glucose unit per substrate encounter, a mode of action defined as processive.[17]
Roles of starch synthases
The initial steps of amylopectin synthesis, as well as the formation of a normal starch granule, require the presence of SSIV, although SSIII also appears to play a role, overlapping its action with that of SSIV.[16]
Like GBSS, SSIV requires the presence of a protein of the PTST family, PTST2, which has no catalytic activity, but is able, thanks to the presence of a specific domain able to bind carbohydrates, to facilitate the binding of the enzyme to alpha-glucans. SSIV is also able to dimerize, an important feature both for the catalytic activity and the ability to interact with other proteins.
According to a model of PTST2 action, the protein, by means of the domain able to bind to carbohydrates, recognizes and forms a complex with MOS having a specific three-dimensional helical shape. In turn, the protein-MOS complex interacts with a SSIV dimer, which is now able to catalyze the elongation of the alpha-glucan, while PTST2 is released so as to allow it to bind another malto-oligosaccharide and facilitate its subsequent interaction with another SSIV dimer.[15]
The action of SSIV is followed by that of the other isoforms of starch synthase. SSI catalyzes the elongation of malto-oligosaccharides with a degree of polymerization of 6 to 7, to form oligosaccharides with a degree of polymerization of 8 to 12, which, in turn, are excellent substrates for SSII, which catalyzes the synthesizes of chain with a degree of polymerization of 12 to 30. These alpha-glucans are further elongated by SSIII, to give linear chains with a degree of polymerization greater than 30.[12] Thus, SSIII appears to act not only in the initial steps of starch granule synthesis, but also in the later steps.
SSIV and SSV appear to be necessary for the synthesis of a regular number of starch granules of normal morphology.[1][16]
Starch branching enzymes
Starch branching enzymes catalyze the formation of α-(1→6) glycosidic bonds, therefore creating branch points in the linear chain of alpha-glucans, of which the main ones are glycogen and amylopectin.[20] Their action increase the number of non-reducing ends, which act as acceptors of glucose units in the elongation reactions.[14]
SBEs catalyze the hydrolytic cleavage of an α-(1→4) bond within an alpha-glucan chain, releasing an oligosaccharide whose reducing end is then linked to the hydroxyl group at C6 position of a glucosyl unit of a alpha-glucan chain. Therefore, the two chains are linked by α-(1→6)-glycosidic bond.
The chain to which the oligosaccharide is linked can be the same one from which it was detached, and in this case we speak of intra-chain transfer, or a different chain, and in this case we speak of inter-chain transfer. Among the factors determining the type of transfer there seems to be the relative concentration of the linear α-(1→4) chains. In particular, it appears that closely associated chains, such as in the double helices in the clusters, promote inter-chain transfer.[19] Finally, it seems that the interaction between starch synthase I and starch branching enzyme is crucial in determining the bimodal chain length distribution observed in plant starches.
Starch branching enzyme isoforms
Two isoforms of the starch branching enzyme are present in plants, referred to as SBEI and SBEII.
Encoded by different genes, they have distinct biochemical properties, which suggests that they play different roles in determining the structure of amylopectin and amylose.[17]
SBEI appears to be expressed more in storage tissues, suggesting a significant role in determining the structural properties of storage starches, shows a substrate preference for amylose, and is able to transfer oligosaccharides with a degree of polymerization greater than 30, although most are between 10 and 13.[6] It also appears to be involved in the synthesis of super-long, or extra long, chains of amylopectin, whereas its other contributions to the structure of amylopectin appear to be less important. Not all plants express SBEI; for example, Arabidopsis and Canola (Brassica napus L.), which are oil-storing plants, have only SBEII and starch is present only in photosynthetic tissues.
SBEII is mostly expressed in grasses and cereals, and many other plants. Its loss causes important alterations in the architecture of amylopectin and reduce starch content. The enzyme shows a substrate preference for amylopectin and transfers oligosaccharides with a degree of polymerization of 6 to 14. In cereals and grasses, there are two tissue-specific isoforms encoded by distinct genes, termed SBEIIa, mainly present in the leaves, and SBEIIb, mainly present in the endosperm.[17]
Starch debranching enzymes
Starch debranching enzymes catalyze the hydrolysis of α-(1→6) glycosidic bonds, and are member of the alpha-amylase superfamily.
Two types of debranching enzymes are present in plants: isoamylases (EC 3.2.1.68) and pullulanases (EC 3.2.1.41).[8] Isoamylases acts on amylopectin and other polyglucans, whereas pullulanases debranch amylopectin and pullulan, a fungal polysaccharide. Isoamylase and pullulanase differ in substrate specificity as well, as they act on branches composed of at least three and two glycosidic residues, respectively.[18]
During starch granule formation, starch debranching enzymes play a crucial role in determining the water-insoluble properties and fine structure of amylopectin. Indeed, the enzyme activity is thought to allow the clustering of remaining branches, thus promoting interactions between adjacent chains and alpha-helix formation, which in turn appears to be important for the formation of the semi-crystalline structures of amylopectin, and then of starch.[17] In the semi-crystalline structure, the branches are presumably inaccessible to the action of starch debranching enzymes, alpha-amylase and beta-amylase.
Starch debranching enzymes are used industrially in the production of resistant starch and cyclodextrins, which are cyclic oligosaccharides.
Starch phosphorylase
The enzyme belongs, like starch synthases, to the family of glycosyltransferases, and is a phosphorylase which resembles glycogen phosphorylase. In plants, it is present in at least two isoenzymatic forms, Pho1, which is found in the stroma of plastids, and is thought to be the true starch phosphorylase involved in starch synthesis, and Pho2, isoform with cytosolic localization.[4]
Starch phosphorylase is thought to be involved in the initial steps of starch synthesis, catalyzing the reversible transfer of glucosyl units to an alpha-glucan, to which they are linked by a α-(1→4) glycosidic bond.[3] Unlike starch synthases, starch phosphorylase uses glucose-1-phosphate and not ADP-glucose as the glucosyl donor.[4]
Phosphorylation of amylopectin
Amylopectin, similarly to glycogen, binds phosphate groups in variable amounts depending on the botanical origin of the starch. For example, potato starch has a relatively high content of phosphate groups, with a degree of substitution of about 0.1 to 0.3 percent, whereas cereal endosperm starches have a phosphate content generally lower than 0.01 percent.[9]
The phosphorylations of amylopectin are catalyzed by two dikinases present in plastids: alpha-glucan water dikinase (EC 2.7.9.4) and phospho-glucan water dikinase (EC 2.7.9.5). These enzymes transfer the beta-phosphate group of ATP to a glucosyl unit of an alpha-glucan chain, while the gamma-phosphate group is transferred to water. Specifically, alpha-glucan water dikinase catalyzes the phosphorylation of the hydroxyl group at C6 position, whereas phospho-glucan water dikinase catalyzes the phosphorylation of the hydroxyl group at C3 position, generally of a prephosphorylated glucan chain.[13] About two thirds of the phosphate groups are bound at the C6 position, while about 20-30% at the C3 position. Phosphate groups are also present at the C2 position, although in a small percentage compared to the other positions. The enzyme which catalyze this phosphorylation in not known.
With regard to substrate specificity, it seems that phosphorylations accumulate more easily on longer chains. Furthermore, it seems to exist an inverse correlation between the total phosphate content and the frequency of amylopectin branching.
The negative charges carried by the phosphate groups cause the mutual repulsion between neighboring phosphorylated oligosaccharides. These repulsions appear to allow the opening and hydration of the chains, thus affecting the activity of the biosynthesis enzymes and making the chains more susceptible to attack by amylases as well.[21]
Amylose/amylopectin ratio
Starch granules are made up mostly of amylopectin and amylose.[12] The two polysaccharides are present in varying percentages, with amylose making up no more than 35 percent of the dry weight of the granule.[5] However, there are plants whose starch granules consist mostly, or almost exclusively, of amylopectin, and whose starches are defined as waxy starches, and plants whose starch granules consist mostly, or almost exclusively, of amylose.[19]
The amylose/amylopectin ratio influences the physicochemical properties of starch, such as the ability to absorb water, gelatinization, retrogradation, or resistance to enzymatic hydrolysis, the latter important in establishing the rate with which, during carbohydrate digestion, amylose and amylopectin are hydrolyzed to maltose and maltotriose by alpha-amylase.[3] Therefore, the amylose/amylopectin ratio influences the effects of the different types of starch on health, as well as their industrial uses.
References
^ ab Abt M.R., Pfister B., Sharma M., Eicke S., Bürgy L., Neale I., Seung D., Zeeman S.C. STARCH SYNTHASE5, a noncanonical starch synthase-like protein, promotes starch granule initiation in Arabidopsis. Plant Cell 2020;32(8):2543-2565. doi:10.1105/tpc.19.00946
^ ab Cornejo-Ramírez Y.I., Martínez-Cruz O., Del Toro-Sánchez C.L., Wong-Corral F.J., Borboa-Flores J. & Cinco-Moroyoqui F.J. The structural characteristics of starches and their functional properties. CYTA J Food 2018;16(1):1003-1017. doi:10.1080/19476337.2018.1518343
^ abcd Crofts N., Abe N., Oitome N.F., Matsushima R., Hayashi M., Tetlow I.J., Emes M.J., Nakamura Y., Fujita N. Amylopectin biosynthetic enzymes from developing rice seed form enzymatically active protein complexes. J Exp Bot 2015;66(15):4469-82. doi:10.1093/jxb/erv212
^ ab Cuesta-Seijo J.A., Ruzanski C., Krucewicz K., Meier S., Hägglund P., Svensson B., Palcic M.M. Functional and structural characterization of plastidic starch phosphorylase during barley endosperm development. PLoS One 2017;12(4):e0175488. doi:10.1371/journal.pone
^ abc Gous P.W., Fox G.P. Review: Amylopectin synthesis and hydrolysis – Understanding isoamylase and limit dextrinase and their impact on starch structure on barley (Hordeum vulgare) quality. Trends Food Sci Technol 2017;62:23-32. doi:10.1016/j.tifs.2016.11.013
^ Guan H.P., Preiss J. Differentiation of the properties of the branching isozymes from maize (Zea mays). Plant Physiol 1993;102(4):1269-1273. doi:10.1104/pp.102.4.1269
^ ab Li G., Yacine Y., Zhu F. Relationships between supramolecular organization and amylopectin fine structure of quinoa starch. Food Hydrocoll 2021;117:106685. doi:10.1016/j.foodhyd.2021.106685
^ Møller M.S., Henriksen A., Svensson B. Structure and function of α-glucan debranching enzymes. Cell Mol Life Sci. 2016;73(14):2619-41. doi:10.1007/s00018-016-2241-y
^ Nitschke F., Wang P., Schmieder P., Girard J.M., Awrey D.E., Wang T., Israelian J., Zhao X., Turnbull J., Heydenreich M., Kleinpeter E., Steup M., Minassian B.A. Hyperphosphorylation of glucosyl C6 carbons and altered structure of glycogen in the neurodegenerative epilepsy Lafora disease. Cell Metab 2013;17(5):756-67. doi:10.1016/j.cmet.2013.04.006
^ abcd Pfister B., Zeeman S.C. Formation of starch in plant cells. Cell Mol Life Sci 2016;73(14):2781-807. doi:10.1007/s00018-016-2250-x
^ ab Pfister B., Zeeman S.C., Rugen M.D., Field R.A., Ebenhöh O., Raguin A. Theoretical and experimental approaches to understand the biosynthesis of starch granules in a physiological context. Photosynth Res 2020;145:55-70. doi:10.1007/s11120-019-00704-y
^ abcd Qu J., Xu S., Zhang Z., Chen G., Zhong Y., Liu L., Zhang R., Xue J., Guo D. Evolutionary, structural and expression analysis of core genes involved in starch synthesis. Sci Rep 2018;8(1):12736. doi:10.1038/s41598-018-30411-y
^ Ritte G., Heydenreich M., Mahlow S., Haebel S., Kötting O., Steup M. Phosphorylation of C6- and C3-positions of glucosyl residues in starch is catalysed by distinct dikinases. FEBS Lett 2006;580(20):4872-6. doi:10.1016/j.febslet.2006.07.085
^ Sawada T., Nakamura Y., Ohdan T., Saitoh A., Francisco P.B. Jr, Suzuki E., Fujita N., Shimonaga T., Fujiwara S., Tsuzuki M., Colleoni C., Ball S. Diversity of reaction characteristics of glucan branching enzymes and the fine structure of α-glucan from various sources. Arch Biochem Biophys 2014;562:9-21. doi:10.1016/j.abb.2014.07.032
^ Seung D., Boudet J., Monroe J., Schreier T.B., David L.C., Abt M., Lu K.J., Zanella M., Zeeman S.C. Homologs of PROTEIN TARGETING TO STARCH control starch granule initiation in Arabidopsis leaves. Plant Cell 2017;29(7):1657-1677. doi:10.1105/tpc.17.00222
^ abcd Szydlowski N., Ragel P., Raynaud S., Lucas M.M., Roldán I., Montero M., Muñoz F.J., Ovecka M., Bahaji A., Planchot V., Pozueta-Romero J., D’Hulst C., Mérida A. Starch granule initiation in Arabidopsis requires the presence of either class IV or class III starch synthases. Plant Cell 2009;21(8):2443-57. doi:10.1105/tpc.109.066522
^ abcdefg Tetlow I.J., Bertoft E. A review of starch biosynthesis in relation to the building block-backbone model. Int J Mol Sci 2020;21(19):7011. doi:10.3390/ijms21197011
^ Xia W., Zhang K., Su L., Wu J. Microbial starch debranching enzymes: developments and applications. Biotechnol Adv 2021;50(3):107786. doi:10.1016/j.biotechadv.2021.107786
^ ab Wang J., Hu P., Lin L., Chen Z., Liu Q., Wei C. Gradually decreasing starch branching enzyme expression is responsible for the formation of heterogeneous starch granules. Plant Physiol 2018;176(1):582-595. doi:10.1104/pp.17.01013
^ Wilkens C., Svensson B., Møller M.S. Functional roles of starch binding domains and surface binding sites in enzymes involved in starch biosynthesis. Front Plant Sci 2018;9:1652. doi:10.3389/fpls.2018.01652
^ Zhou W., He S., Naconsie M., Ma Q., Zeeman S.C., Gruissem W. & Zhang P. Alpha-glucan, water dikinase 1 affects starch metabolism and storage root growth in Cassava (Manihot esculenta Crantz). Sci Rep 2017;7:9863 doi:10.1038/s41598-017-10594-6
^ Ziegler G.R., Creek J.A., Runt J. Spherulitic crystallization in starch as a model for starch granule initiation. Biomacromolecules 2005;6(3):1547-54. doi:10.1021/bm049214p
Starch synthase (EC 2.4.1.21) is an enzyme that catalyzes the transfer of glucose molecules from ADP-glucose to the non-reducing end of a pre-existing α-(1→4) linked glucan chain, to which the monosaccharides are linked by an α-(1→4) glycosidic bond.[4]
This enzymatic activity is involved in the synthesis of amylose and amylopectin, the two major constituents of starch, which is the main storage form of carbohydrates in plants.[7]
Starch synthase belongs to the family of glycosyltransferases (EC 2.4), like starch phosphorylase (EC 2.4.1.1), another enzyme involved in starch metabolism, and glycogen synthase (EC 2.4.1.11) and glycogen phosphorylase (EC 2.4.1.1), enzymes involved in glycogen synthesis and glycogen degradation or glycogenolysis, respectively.[6] However, while glycogen synthase uses UDP-glucose as the glucosyl donor, and starch phosphorylase uses glucose 1-phosphate, starch synthase uses ADP-glucose.[2][3]
Six isoforms of starch synthase are known in plants. They are structurally related proteins, with five involved in the synthesis of amylopectin, referred to as starch synthase I, II, III, IV and V, or SSI, SSII, SSIII, SSIV and SSV, respectively, and one involved in the synthesis of amylose, the granule-bound starch synthase (GBSS) (EC 2.4.1.242).[6] GBSS, SSI, SSII, SSIII, and SSIV have catalytic activity, whereas SSV lacks catalytic activity.[1]
GBSS is almost exclusively bound to starch granules and is located mostly within them, as evidenced by treatment of the granule surface with proteases. The other isoforms of starch synthase are present either only in the stroma of the plastids or portioned between starch granules and the stroma, and are called soluble starch synthases.[8]
Starch synthases I, II, III, and IV catalyze the adding of one glucose residue per substrate encounter, a distributive mode of action, whereas GBSS catalyze the addition of more than one glucosidic unit per substrate encounter, a processive mode of action.[11]
Starch synthase and MOS
Starch synthases involved in the early steps of amylose and amylopectin synthesis require short malto-oligosaccharides (MOS) to initiate de novo synthesis of the two polysaccharides.[11]
These small oligosaccharides, namely, α-(1→4)-glucans with a degree of polymerization of 2 to 7, act as primers and are elongated, a function analogous to that performed by glycogenin in glycogen synthesis.
MOS can originate from the activity of starch synthase III, starch phosphorylase, or starch debranching enzymes.
Being poorly water soluble, MOS seem able to evade the hydrolytic action of alpha-amylase (EC 3.2.1.1) and beta-amylase.[8]
Starch synthase and amylopectin synthesis
The synthesis of amylopectin requires the temporally coordinated action of at least four classes of enzymes, namely, starch synthase isoenzymes, starch phosphorylase, starch branching enzymes (EC 2.4.1.18), and starch debranching enzymes.[2][7] It is also believed that, in many cases, these enzymes physically interact with each other to form multienzyme complexes, which are structures capable of increasing the efficiency of a metabolic pathway.[11]
It is generally accepted that the growth of the starch granule occurs from a central core called the hilum, whose precise structure is unknown, although it appears to be formed by a disordered structure of α-glucans.[13] The initiation of the hilum, the formation of a normal starch granule morphology, and the degree of starch accumulation require the presence of SSIV, although it has been suggested that SSIII and SSV may also play a role, overlapping their activity with that of SSIV.[10]
GBSS and amylose synthesis
Granule-bound starch synthase is involved in the synthesis of amylose.
This enzyme was first reported by Luis Federico Leloir in the early 60s, the same researcher who in 1948 had discovered the main pathway for galactose metabolism, the Leloir pathway.[5]
Its catalytic action is not entirely simultaneous with that of the other starch synthases, as it requires the presence of an amylopectin-matrix.[6]
In grasses, the granule-bound starch synthase is present in two isoforms, encoded by distinct genes, and referred to as GBSSI and GBSSII.[12]
GBSS requires, for its catalytic activity, the presence of a protein of the PTST family, PTST1, which allows its binding to the starch granule, and whose action seems to be more important in chloroplasts than in amyloplasts.[9]
PTST1 appears to associate, in the plastid stoma, with GBSS. The complex binds to the nascent starch granule, the protein dissociates from the enzyme, that begins to catalyze the elongation of the malto-oligosaccharides, while the protein returns to the stroma where it recruits another GBSS.
References
^ Abt M.R., Pfister B., Sharma M., Eicke S., Bürgy L., Neale I., Seung D., Zeeman S.C. STARCH SYNTHASE5, a noncanonical starch synthase-like protein, promotes starch granule initiation in Arabidopsis. Plant Cell 2020;32(8):2543-2565. doi:10.1105/tpc.19.00946
^ ab Crofts N., Abe N., Oitome N.F., Matsushima R., Hayashi M., Tetlow I.J., Emes M.J., Nakamura Y., Fujita N. Amylopectin biosynthetic enzymes from developing rice seed form enzymatically active protein complexes. J Exp Bot 2015;66(15):4469-82. doi:10.1093/jxb/erv212
^ Cuesta-Seijo J.A., Ruzanski C., Krucewicz K., Meier S., Hägglund P., Svensson B., Palcic M.M. Functional and structural characterization of plastidic starch phosphorylase during barley endosperm development. PLoS One 2017;12(4):e0175488. doi:10.1371/journal.pone
^ Gous P.W., Fox G.P. Review: Amylopectin synthesis and hydrolysis – Understanding isoamylase and limit dextrinase and their impact on starch structure on barley (Hordeum vulgare) quality. Trends Food Sci Technol 2017;62:23-32. doi:10.1016/j.tifs.2016.11.013
^ Leloir L.F., de Fekete M.A., Cardini C.E. Starch and oligosaccharide synthesis from uridine diphosphate glucose. J Biol Chem 1961;236:636-41. doi:10.1016/S0021-9258(18)64280-2
^ abc Pfister B., Zeeman S.C. Formation of starch in plant cells. Cell Mol Life Sci 2016;73(14):2781-807. doi: 10.1007/s00018-016-2250-x
^ ab Qu J., Xu S., Zhang Z., Chen G., Zhong Y., Liu L., Zhang R., Xue J., Guo D. Evolutionary, structural and expression analysis of core genes involved in starch synthesis. Sci Rep 2018;8(1):12736. doi:10.1038/s41598-018-30411-y
^ ab Seung D., Boudet J., Monroe J., Schreier T.B., David L.C., Abt M., Lu K.J., Zanella M., Zeeman S.C. Homologs of PROTEIN TARGETING TO STARCH control starch granule initiation in Arabidopsis leaves. Plant Cell 2017;29(7):1657-1677. doi:10.1105/tpc.17.00222
^ Seung D., Soyk S., Coiro M., Maier B.A., Eicke S., and Zeeman S.C. PROTEIN TARGETING TO STARCH is required for localising GRANULE-BOUND STARCH SYNTHASE to starch granules and for normal amylose synthesis in Arabidopsis. PLoS Biol 2015;13:e1002080. doi:10.1371/journal.pbio.1002080
^ Szydlowski N., Ragel P., Raynaud S., Lucas M.M., Roldán I., Montero M., Muñoz F.J., Ovecka M., Bahaji A., Planchot V., Pozueta-Romero J., D’Hulst C., Mérida A. Starch granule initiation in Arabidopsis requires the presence of either class IV or class III starch synthases. Plant Cell 2009;21(8):2443-57. doi:10.1105/tpc.109.066522
^ abc Tetlow I.J., Bertoft E. A review of starch biosynthesis in relation to the building block-backbone model. Int J Mol Sci 2020;21(19):7011. doi:10.3390/ijms21197011
^ Vrinten P.L., Nakamura T. Wheat granule-bound starch synthase I and II are encoded by separate genes that are expressed in different tissues. Plant Physiol 2000;122(1):255-64. doi:10.1104/pp.122.1.255
^ Ziegler G.R., Creek J.A., Runt J. Spherulitic crystallization in starch as a model for starch granule initiation. Biomacromolecules 2005;6(3):1547-54. doi:10.1021/bm049214p
Amylose is a polysaccharide made up of α-D-glucose units linked by α-(1→4) glycosidic bonds, with few branches connected to the main chain by α-(1→6) glycosidic bonds.[17]
Together with amylopectin, it is one of the two main constituents of the starch, the major storage form of energy and carbohydrates in the Biosphere.[12]
Its synthesis is catalyzed by the enzyme granule-bound starch synthase or GBSS (EC 2.4.1.242), and requires the presence of a second protein, named protein targeting to starch 1 or PTST1, with no catalytic activity.[15]
Inside the starch granule, amylose is embedded within the semi-crystalline matrix formed by amylopectin.[13] Unlike amylopectin, amylose is not necessary for the formation of starch granules, is present in smaller amounts, but it has a great influence on the physicochemical properties of the starch.[7]
Plants have been selected whose starch granules hold negligible amounts or, conversely, very high amounts of amylose. These phenotypes have both industrial applications and potential health benefits.[4][20]
Amylose molecules have a molecular weight of about 106 daltons, are mostly linear and made up of α-D-glucose units, hereinafter referred to as glucose, linked by α-(1→4)-glycosidic bonds, namely, covalent bonds between C-1 of one unit and the hydroxyl group on the C-4 of the next unit.[1] The linear chains are made up of a number of monosaccharides ranging from a few hundred to several thousand; therefore, they are much longer than amylopectin chains.[17]
The few branches are connected to the linear chain by α-(1→6) glycosidic bonds, such as in amylopectin and glycogen, the storage form of carbohydrates in animals. An α-(1→6) glycosidic bond is a covalent bonds between C-1 of one unit and the hydroxyl group on the C-6 of another glucose unit. The number of branches is between 5 and 20, depending on the botanical origin of the starch, and the branches, compared to amylopectin, are not grouped.[6] Studies on the length of the amylose branches have shown a bimodal distribution, with the two fractions termed as:
AM1, which includes the shorter chains, with a degree of polymerization between 100 and 700 daltons;
AM2, which includes the longer chains, with a degree of polymerization between 700 and 40,000 daltons.[19]
A similar bimodal distribution of the length of the branches is also observed for amylopectin, whose fractions are indicated as AP1, shorter and more abundant, and AP2.
The intraspecies variation of the distribution of the AM1 and AM2 fractions is relatively small, whereas it is large between different species, variation that has a genetic basis.[19]
Amylose location in the starch granules
The precise location of amylose in the starch granule is not known, although it is believed that most are found in the amorphous regions. However, some studies have suggested that its localization is not restricted to the amorphous regions, but is also present between amylopectin chains and on the surface of the granules.[13] Hence, amylose could have several locations within the granule.
Synthesis
In plants, the synthesis of amylose is catalyzed by GBSS, one of the six isoforms of starch synthase.[11] The enzyme, whose action is the major determinant of amylose content of starch granules, requires the presence of the PTST1 protein.[15] Amylose branching appears to be carried out by starch branching enzyme or SBE (EC 2.4.1.18).
As in the case of amylopectin synthesis, it is believed that the enzymes involved physically interact to form multienzyme complexes, which are structures able to optimize the efficiency of the process.[17]
Since amylose synthesis requires a pre-existent amylopectin matrix in order to target the granule-bound starch synthase to the starch granules, the synthesis of the two polysaccharides is not entirely simultaneous.[11]
In the initial steps, the granule-bound starch synthase, like the other starch synthases, requires short malto-oligosaccharides or MOS, α-(1→4) glucans with a degree of polymerization of 2 to 7, which act as a primer and are elongated.[17] MOS can originate from various sources, such as the trimming process of nascent amylopectin molecules by starch debranching enzyme, or from the activity of starch phosphorylase (EC 2.4.1.1), another enzyme involved in starch metabolism.[16] As MOS are poorly water soluble, they seem able to evade the hydrolytic action of alpha-amylase (EC 3.2.1.1) and beta-amylase (EC 3.2.1.2), and diffuse within the starch granule matrix where they are elongated by GBSS.[14] Once malto-oligosaccharides are elongated beyond seven glucose residues, they cannot easily diffuse out of the granule, and are further elongated.
Requiring a primer, the early stage of the synthesis of amylose and amylopectin resembles that of glycogen. However, the primer required during glycogen synthesis is the self-glucosylating protein glycogenin.
It should be noted that the polymerization of glucose to amylopectin, amylose, glycogen, and more generally, of osmotically active monosaccharides into osmotically inactive polysaccharides allows the storage of large amounts of osmotically active molecules inside the cell without an increase in osmotic pressure.
Granule-bound starch synthase
GBSS, the other isoforms of starch synthase, and starch phosphorylase belong to the glycosyltransferase family (EC 2.4).[11]
GBSS, discovered by the group of Louis Federico Leloir, who had previously discovered the main pathway for the metabolism of galactose, the Leloir pathway, is the most abundant among proteins associated with starch granules.[9] It is present almost exclusively bound to the granule, unlike the other isoforms of starch synthase which are mostly present in the stroma of the plastids or portioned between starch granules and the stroma. Moreover, the treatment of starch granule surface with proteases showed that the majority of the granule bound starch synthase is present within the granule rather than on its surface, a location consistent with amylose synthesis within the nascent granules.[15]
In grasses, GBSS is present in two isoforms, encoded by distinct genes. The isoforms are known as GBSSI, present in the amyloplasts of storage tissues, therefore non-photosynthetic tissues, and GBSSII, present in the chloroplasts, therefore in photosynthetic tissues, where it takes part in the synthesis of transient starch.[18]
GBSS catalyzes the transfer of a glucose residue from ADP-glucose to the non-reducing end of an α-(1→4)-glucan, to which the glucose residue is joined by an α-(1→4) glycosidic bond.[5]
Notee that starch synthases use ADP-glucose as the glucosyl donor, whereas glycogen synthase, an enzyme involved in glycogen synthesis, uses UDP-glucose.[3]
Granule-bound starch synthase is capable of adding more than one glucose monomer per substrate encounter, a processive mode of action, which differentiates it from the other isoforms of starch synthase, which are capable of adding only one glucose unit per substrate encounter, a distributive mode of action. The processive action allows the synthesis of long linear chains, and seems to be strongly increased by the presence of amylopectin.[17]
PTST1
Amylose synthesis requires the presence of a protein of the PTST family, namely PTST1, which was discovered more than fifty years after GBSS.[15]
PTST1 has no catalytic activity, but allows the binding of GBSS to the starch granule, an activity that seems more important in chloroplasts, and therefore for the synthesis of transient starch, than in amyloplasts. It was proposed that PTST1 associates with the synthase in the plastid stroma, the complex binds to the nascent starch granule, PTST1 dissociates from the enzyme which initiates the synthesis of amylose, while the PTST1 returns to the plastid stroma to recruit another GBSS molecule.
The importance of PTST1 is underlined by the fact that it is conserved throughout the plant kingdom, and its loss causes the enzyme to detach from the nascent starch granule.
Starch branching enzymes
The enzyme that generate the few α-(1→6) linkages present in amylose molecules is not known, although an isoform of the starch branching enzyme, SBEI, may be involved.[13]
SBEI is present mostly in the plastid stroma, whereas only a small proportion is bound to starch granule, and is mostly expressed in the storage tissues. The low branch frequency of the amylose could be due to its synthesis inside the granules, where SBEI is scarcely present, which would protect the nascent molecule from the action of the enzyme.[8]
Amylose/amylopectin ratio
In the starch granules of land plants, amylose is almost always present, in variable percentages, generally between 5-35%.[5] The variability occurs not only between different species, but also within the same species based on the organ or tissue considered, and, in tubers and seeds, based on the stage of development, being the content generally low in the early stages, then increasing until the final value is reached, a pattern consistent with the synthesis of amylose within an amylopectin matrix.[13]
However, there are plants whose amylose content is very low, or even absent. Their starch is referred to as waxy, due to the appearance of the endosperm of the raw grains which resembles wax. Conversely, there are plants with starch granules containing mostly, or entirely, amylose.[20]
Although its role has not yet been clarified, its near-constant presence seems to indicate this polysaccharide plays an important structural role in the starch granule, and provides the plant some advantage in the growth and development phase.[13]
The amylose/amylopectin ratio strongly influences the physicochemical properties of starch, such as its ability to absorb water, which influences processes such as starch retrogradation and gelatinization, or its resistance to enzymatic hydrolysis, which determinates, for example, the rate at which maltose and maltotriose are released during starch degradation by alpha-amylase or beta-amylase.[10] These properties, in turns, are able to affect the industrial applications of starch as well as its effects on health.
High-amylose starches
High-amylose cereals are obtained through enhancing GBSSI gene expression, or suppressing or eliminating the genes encoding for starch branching enzyme, SSIIa, or other enzymes and proteins involved in amylopectin biosynthesis. However, at least in cereal endosperm, the most effective method is the suppression or elimination of one or more starch branching enzyme isoforms.[4][20]
High amylose starches have peculiar physicochemical properties, such as a high gelling strength, an ease retrogradation, and an excellent ability to form films, properties which make them suitable for industrial applications such as the production of biodegradable plastics, paper, and adhesives.[7][10]
High amylose starches are high in resistant starch, which is a starch that escape digestion in the small intestine by alpha-amylase, one of the hydrolases involved in carbohydrate digestion. Studies conducted using foods enriched with resistant starch have shown an improvement in insulin and glycemic responses, as well as a reduction in the risk of developing cardiovascular disease, obesity, and type II diabetes mellitus.[2] How do they work?
Resistant starch, being able to escape intestinal digestion, may low the glycemic index of the foods in which it is found, thus contributing to the regulation of blood glucose levels. Furthermore, once reached the colon, it can be fermented by the bacteria of the gut microbiota, which is part of the human microbiota, with production of short-chain fatty acids, mainly butyric acid, acetic acid, and propionic acid, fatty acids essential for intestinal health.
^ Birt D.F., Boylston T., Hendrich S., Jane J.L., Hollis J., Li L., McClelland J., Moore S., Phillips G.J., Rowling M., Schalinske K., Scott M.P., Whitley E.M. Resistant starch: promise for improving human health. Adv Nutr 2013;4(6):587-601. doi:10.3945/an.113.004325
^ Crofts N., Abe N., Oitome N.F., Matsushima R., Hayashi M., Tetlow I.J., Emes M.J., Nakamura Y., Fujita N. Amylopectin biosynthetic enzymes from developing rice seed form enzymatically active protein complexes. J Exp Bot 2015;66(15):4469-82. doi:10.1093/jxb/erv212
^ ab Funnell-Harris D.L., Sattler S.E., O’Neill P.M., Eskridge K.M., and Pedersen J.F. Effect of waxy (low amylose) on fungal infection of Sorghum grain. Phytopathology 2015;105(6):716-846. doi:10.1094/PHYTO-09-14-0255-R
^ ab Gous P.W., Fox G.P. Review: amylopectin synthesis and hydrolysis – Understanding isoamylase and limit dextrinase and their impact on starch structure on barley (Hordeum vulgare) quality. Trends Food Sci Technol 2016;62:23-32. doi:10.1016/j.tifs.2016.11.013
^ Hizukuri S., Takeda Y., Yasuda M., Suzuki A. Multi-branched nature of amylose and the action of debranching enzymes. Carbohydr Res 1981;94:205-213. doi:10.1016/S0008-6215(00)80718-1
^ ab Jobling S. Improving starch for food and industrial applications. Curr Opin Plant Biol 2004;7(2):210-8. doi:10.1016/j.pbi.2003.12.001
^ Kram A.M., Oostergetel G.T., Van Bruggen E. Localization of branching enzyme in potato tuber cells with the use of immunoelectron microscopy. Plant Physiol 1993;101(1):237-243. doi:10.1104/pp.101.1.237
^ Leloir L.F., de Fekete M.A., Cardini C.E. Starch and oligosaccharide synthesis from uridine diphosphate glucose. J Biol Chem 1961;236:636-41. doi:10.1016/S0021-9258(18)64280-29
^ ab Magallanes-Cruz P.A., Flores-Silva P.C., Bello-Perez L.A. Starch structure influences its digestibility: a review. J Food Sci 2017;82(9):2016-2023. doi:10.1111/1750-3841.13809
^ abc Pfister B., Zeeman S.C. Formation of starch in plant cells. Cell Mol Life Sci 2016;73(14):2781-807. doi:10.1007/s00018-016-2250-x
^ Qu J., Xu S., Zhang Z., Chen G., Zhong Y., Liu L., Zhang R., Xue J., Guo D. Evolutionary, structural and expression analysis of core genes involved in starch synthesis. Sci Rep 2018;8(1):12736. doi:10.1038/s41598-018-30411-y
^ abcde Seung D. Amylose in starch: towards an understanding of biosynthesis, structure and function. New Phytol 2020;228:1490-1504. doi:10.1111/nph.16858
^ Seung D., Boudet J., Monroe J., Schreier T.B., David L.C., Abt M., Lu K.J., Zanella M., Zeeman S.C. Homologs of PROTEIN TARGETING TO STARCH control starch granule initiation in Arabidopsis leaves. Plant Cell 2017;29(7):1657-1677. doi:10.1105/tpc.17.00222
^ abcd Seung D., Soyk S., Coiro M., Maier B.A., Eicke S., Zeeman S.C. PROTEIN TARGETING TO STARCH is required for localising GRANULE-BOUND STARCH SYNTHASE to starch granules and for normal amylose synthesis in Arabidopsis. PLOS Biol 2015;13(2):e1002080. doi:10.1371/journal.pbio.1002080
^ Szydlowski N., Ragel P., Raynaud S., Lucas M.M., Roldán I., Montero M., Muñoz F.J., Ovecka M., Bahaji A., Planchot V., Pozueta-Romero J., D’Hulst C., Mérida A. Starch granule initiation in Arabidopsis requires the presence of either class IV or class III starch synthases. Plant Cell 2009;21(8):2443-57. doi:10.1105/tpc.109.066522
^ abcde Tetlow I.J., Bertoft E. A review of starch biosynthesis in relation to the building block-backbone model. Int J Mol Sci 2020;21(19):7011. doi:10.3390/ijms21197011
^ Vrinten P.L., Nakamura T. Wheat granule-bound starch synthase I and II are encoded by separate genes that are expressed in different tissues. Plant Physiol 2000;122(1):255-64. doi:10.1104/pp.122.1.255
^ ab Wang K., Hasjim J., Wu A.C., Henry R.J., Gilbert R.G. Variation in amylose fine structure of starches from different botanical sources. J Agric Food Chem 2014;62(19):4443-53. doi:10.1021/jf5011676
^ abc Wang J., Hu P., Chen Z., Liu Q., Wei C. Progress in high-amylose cereal crops through inactivation of starch branching enzymes. Front Plant Sci 2017;8:469. doi:10.3389/fpls.2017.00469
The Leloir pathway is the main pathway for galactose metabolism.
Discovered by Leloir L.F. and colleagues in 1948, it leads to the conversion of galactose into glucose 1-phosphate, namely, to the inversion of the configuration of hydroxyl group at C4 of galactose, one of the chirality centers of the monosaccharide.[9]
The metabolic intermediates involved in this isomerization are building blocks in many metabolic pathways, such as glycosylation reactions of proteins and lipids or glycogen synthesis, depending on the stage of development, the metabolic state of the cell, and the type of tissue.[6]
Except for the first step, the other reactions of the Leloir pathway may proceed in both directions, depending on substrate levels and metabolic demands of the tissue. This allows the interconversion of galactose and glucose.[4]
The importance of the Leloir pathway, and therefore of galactose, is emphasized by the fact that it is highly conserved in nature, from bacteria to plants and animals, and, in humans, by the severity of the consequences due to mutations in one of the genes encoding the enzymes of the pathway, mutations that cause the genetic metabolic disorder known as galactosemia.[2][8]
Galactose, together with glucose and fructose, is one of the monosaccharides that can be absorbed in the intestine. The main dietary source of galactose is lactose, which, with maltose, trehalose and sucrose, is one of the disaccharides found in food. Since there are no transporters for disaccharides, in the last stage of carbohydrate digestion their glycosidic bonds are hydrolyzed with the release of the constituent monosaccharides, which for lactose are glucose and galactose. Then follows the absorption of monosaccharides which, through the portal system, reach the liver, which is the main site for the metabolism of galactose and absorbs, through passive diffusion mediated by the GLUT2 transporter, most of it, about 88%.[3] The residual circulating quantity reaches other organs and tissues, such as the mammary gland that, during the lactation phase, uses it for the production of lactose and the glycosylation of milk proteins and lipids.[4]
The steps of the Leloir pathway
The Leloir pathway consists of four reactions catalyzed by the enzymes galactose mutarotase or aldose 1-epimerase (EC 5.1.3.3), galactokinase (EC 2.7.1.6), galactose 1-phosphate uridylyltransferase or GALT (EC 2.7.7.12) and UDP-galactose 4-epimerase or GALE (EC 5.1.3.2).[8]
The Leloir Pathway
Step 1: galactose mutarotase
The cleavage of the β-(1→4) glycosidic bond of lactose leads to the release of glucose and beta-galactose.
Galactokinase, the enzyme that catalyzes the second step of the Leloir pathway, is specific for the alpha anomer of galactose. The conversion of beta-galactose to alpha-anomer is catalyzed by galactose mutarotase. The enzyme is also able to interconvert alpha and beta-configurations of glucose, xylulose, maltose and lactose, albeit with variable efficiency.[12]
Step 2: galactokinase
In the second step, alpha-galactose is phosphorylated to galactose 1-phosphate, in a reaction catalyzed by galactokinase.[8] The phosphorylation of galactose has some functions.
Galactose 1-phosphate cannot diffuse out of the cell, as it is negative charged and there are no transporters for phosphorylated sugars in the plasma membrane.
Phosphorylation increases the energy content of the monosaccharide, and this starts to destabilize the molecule facilitating its further metabolism.
The phosphorylation of galactose maintains a low intracellular concentration of the monosaccharide, thus favoring its facilitated transport into the cell.
The reaction catalyzed by galactokinase is the irreversible step of the Leloir pathway.[4] Unlike hexokinase and glucokinase (EC 2.7.1.1), which phosphorylate the hydroxyl group at C6 of glucose, galactokinase and fructokinase (EC 2.7.1.4) phosphorylate the hydroxyl group at C1 of galactose and fructose, respectively.[11]
The conversion of galactose 1-phosphate to glucose 1-phosphate requires two reactions, the third and fourth of the Leloir pathway, respectively.
Step 3: galactose 1-phosphate uridylyltransferase
In the third step, galactose 1-phosphate uridylyltransferase catalyzes the transfer to galactose 1-phosphate of the UMP group of UDP-glucose, to form UDP-galactose and glucose 1-phosphate. The reaction proceeds with a ping-pong mechanism with the formation of a covalent intermediate between the enzyme and UMP.[7]
Step 4: UDP-galactose 4-epimerase
In the last step, UDP-galactose is converted to UDP-glucose in a reaction catalyzed by UDP-galactose 4-epimerase. The enzyme inverts the configuration of the hydroxyl group at C4, and is responsible for the interconversion of UDP-galactose and UDP-glucose, hence between galactose and glucose.[8] UDP-galactose 4-epimerase requires NAD+ as a cofactor and the reaction proceeds through the formation of a ketonic intermediate at C4 with simultaneous reduction of NAD+ to NADH. Subsequently, the ketonic intermediate rotates presenting the opposite face of the sugar to NADH, and a hydride ion is transferred back from NADH to C4 but in the opposite configuration.[13] Since NAD is first reduced and then oxidized, no net oxidation or reduction of the coenzyme occurs, which therefore does not appear in the reaction equation. In mammals, UDP-galactose 4-epimerase also catalyzes the interconversion between UDP-N-acetylgalactosamine and UDP-N-acetylglucosamine.[3] It seems that UDP-galactose 4-epimerase is the rate-limiting enzyme of the Leloir pathway.
UDP-glucose produced is recycled in the reaction catalyzed by UDP-glucose pyrophosphorylase, with the release of glucose 1-phosphate.[6]
Note that UDP-galactose and UDP-glucose, or more generally galactose and glucose, have the same molecular formula. Therefore, they are isomers, and differ only in the configuration of the fourth carbon atom. Therefore they are an example of optical isomerism.
What is the role of the Leloir pathway?
The Leloir pathway allows the cell to use galactose or the derived glucose in many metabolic pathways, both anabolic and catabolic, depending on the metabolic state of the cell or tissue. Furthermore, since the reaction catalyzed by UDP-galactose 4-epimerase is reversible, conversion of glucose to galactose and nucleotide derivatives is possible.[4][6]
UDP-galactose can be used:
in the glycosylation of proteins and lipids, such as galactocerebrosides which are the major glycolipids components in myelin, and for this reason galactose was initially called cerebrose;
in the lactating mammary gland for the production of lactose in a reaction catalyzed by the lactose synthase system. Moreover, as the reaction catalyzed by UDP-galactose 4-epimerase may proceed in both directions, glucose can be converted, after activation to UDP-glucose in a reaction catalyzed by UDP-glucose pyrophosphorylase (EC 2.7.7.9), into UDP-galactose for the synthesis of lactose.
In the liver and skeletal muscle, UDP-glucose derived from UDP-galactose, can be used:[4]
in the glycosylation of lipids and proteins;
in glycogen synthesis, when the energy demand of the cell is low; and, with respect to glucose and fructose, galactose is preferentially incorporated in hepatic glycogen rather than being directed to oxidative metabolism;
following conversion to glucose 1-phosphate, in a reaction catalyzed by UDP-glucose pyrophosphorylase, and isomerization to glucose 6-phosphate in a reaction catalyzed by phosphoglucomutase (EC 5.4.2.2), it may enter different metabolic pathways, such as glycolysis, the pentose phosphate pathway or gluconeogenesis.[3]
Note: UDP-galactose, discovered during the studies on the Leloir pathway, was the first nucleotide sugar to be identified.[1]
Leloir pathway and galactosemia
Glycosylations are post-translational modifications that play a key role in enabling and regulating many biological processes. Glycosylation defects have been related to many pathological conditions such as cancer, diabetes, and congenital inborn errors of metabolism such as congenital disorders of glycosylation, which are mainly autosomal recessive monogenic disorders.[5] Congenital disorders of glycosylation include galactosemia, which was first described by von Reuss A. in 1908.[14] Galactosemia is due to mutations in one of the genes encoding the four enzymes of the Leloir pathway, and four types have been identified:
type I, due to deficiency of galactose 1-phosphate uridylyltransferase, the most common form, also called classical galactosemia;
type II, due to galactokinase deficiency;
type III, due to galactose 4-epimerase deficiency;
type IV, due to galactose mutarotase deficiency.[15]
To date, the therapeutic standard of care for galactosemia is a galactose-restricted diet.
Galactosemia and cataracts
The accumulation of galactose fuels alternative metabolic pathways such as the synthesis of galactitol and galactonate.
The symptoms of galactosemia include the early onset of cataracts, usually within the first two years of life, and in the most severe cases, brain, liver, and kidneys damages, too.[12]
It seems that one of the factors triggering cataracts is the reduction of galactose, accumulated in the lens of the eye, to galactitol, in a reaction catalyzed by aldose reductase (EC 1.1.1.21).[10] Galactitol, which is poorly metabolized, does not diffuse through cell membranes due to its poor lipophilicity and, being osmotically active, increases the intracellular osmotic pressure causing a net flow of water inside the cell. Moreover, its synthesis, depleting NADPH levels, may reduce glutathione reductase (EC 1.8.1.7) activity and lead to free radical accumulation.[3]
The osmotic effect and the build-up of free radicals may ultimately damage cell integrity and cause cell death. Furthermore, it has been reported that galactitol is a galactose mutarotase inhibitor, so its accumulation may lead to further accumulation of unmetabolized galactose.[12]
References
^ Cardini C.E., Paladini A.C., Caputto R., Leloir L.F. Uridine diphosphate glucose: the coenzyme of the galactose-glucose phosphate isomerization. Nature 1950;165:191-192. doi:10.1038/165191a0
^ Coelho A.I., Berry G.T., Rubio-Gozalbo M.E. Galactose metabolism and health. Curr Opin Clin Nutr Metab Care 2015;18(4):422-427. doi:10.1097/MCO.0000000000000189
^ abcd Coelho A.I., Rubio-Gozalbo M.E., Vicente J.B., Rivera I. Sweet and sour: an update on classic galactosemia. J Inherit Metab Dis 2017;40(3):325-342. doi:10.1007/s10545-017-0029-3
^ abcde Conte F., van Buuringen N., Voermans N.C., Lefeber D.J. Galactose in human metabolism, glycosylation and congenital metabolic diseases: time for a closer look. Biochim Biophys Acta Gen Subj 2021;1865(8):129898. doi:10.1016/j.bbagen.2021.129898
^ abc Frey P.A. The Leloir pathway: a mechanistic imperative for three enzymes to change the stereochemical configuration of a single carbon in galactose. FASEB J 1996;10(4):461-70. doi:10.1096/fasebj.10.4.8647345
^ abcd Holden H.M., Rayment I., Thoden J.B. Structure and function of enzymes of the Leloir pathway for galactose metabolism. J Biol Chem 2003;278(45):43885-43888. doi:10.1074/jbc.R300025200
^ Leloir L.F., de Fekete M.A., Cardini C.E. Starch and oligosaccharide synthesis from uridine diphosphate glucose. J Biol Chem 1961;236:636-41. doi:10.1016/S0021-9258(18)64280-2
^ Pintor J. Sugars, the crystalline lens and the development of cataracts. Biochem Pharmacol 2012;1:4. doi:10.4172/2167-0501.1000e1190
^ Rosenthal M.D., Glew R.H. Medical Biochemistry – Human Metabolism in Health and Disease. John Wiley J. & Sons, Inc., 2009
^ abc Thoden J.B., Timson D.J., Reece R.J., and M. Holden H.M. Molecular structure of human galactose mutarotase. J Biol Chem 2004;279(22):23431-23437. doi:10.1074/jbc.M402347200
^ Thoden J.B., Wohlers T.M., Fridovich-Keil J.L., Holden H.M. Human UDP-galactose 4-epimerase. Accommodation of UDP-N-acetylglucosamine within the active site. J Biol Chem 2001;276(18):15131-6. doi:10.1074/jbc.M100220200
^ Wada Y., Kikuchi A., Arai-Ichinoi N., Sakamoto O., Takezawa Y., Iwasawa S., Niihori T., Nyuzuki H., Nakajima Y., Ogawa E., Ishige M., Hirai H., Sasai H., Fujiki R., Shirota M., Funayama R., Yamamoto M., Ito T., Ohara O., Nakayama K., Aoki Y., Koshiba S., Fukao T., Kure S. Biallelic GALM pathogenic variants cause a novel type of galactosemia. Genet Med 2019;21(6):1286-1294. doi:10.1038/s41436-018-0340-x
In solution, solvent molecules tend to move from a region of higher concentration to one of lower concentration. When two different solutions are separated by a semipermeable membrane, namely, a membrane that allows certain ions or molecules to pass, in this case the solvent molecules, a net flow of solvent molecules from the side with higher concentration to the side with lower concentration will occur. This net flow through the semipermeable membrane produces a pressure called osmotic pressure, indicated as Π, that can be defined as the force that must be applied to prevent the movement of the solvent molecules through a semipermeable membrane.
Osmotic pressure, together with boiling point elevation, freezing point depression, and vapor pressure lowering, is one of the four colligative properties of solutions, properties that do not dependent of the chemical properties of the solute particles, namely ions, molecules or supramolecular structures, but depend only on the number of solute particles present in solution.
For a solutions of n solutes, the equation that describes osmotic pressure is the sum of the contributions of each solute:
Π = RT(i1c1 + i2c2 + … + incn)
The equation is known as the van ’t Hoff equation, where:
The van ‘t Hoff factor is a measure of the degree of dissociation of solutes in solution, and is described by the equation:
i = 1 + α(n-1)
where:
α is the degree of dissociation of the solute molecules, equal to the ratio between the moles of the solute molecules that have dissociated and the number of the original moles, and is comprised between 0, for substances that do not ionize or dissociate in solution, and 1, for substances that completely dissociate or ionize;
n is the number of ions formed from the dissociation of the solute molecule.
For non ionizable compounds, such as glucose, glycogen or starch, n = 1, and i = 1, whereas for compounds that completely dissociate, such as strong acids and strong bases or salts, the van ‘t Hoff factor is a whole number greater than one, as α = 1 and n is equal to at least 2. For example, if we consider sodium chloride, NaCl, potassium chloride, KCl, or calcium chloride, CaCl2, in dilute solution:
So in the first two cases i = 2, whereas with calcium chloride, i = 3.
Finally, for substances that do not completely ionize, such as weak acids and weak bases, i is not an integer.
The product of the van ’t Hoff factor and the molar concentration of the solute particles, ic, is the osmolarity of the solution, namely, the concentration of the solute particles osmotically active per liter of solution.
Osmotic pressure, osmosis, and plasma membranes
Osmosis can be defined as the net movement or flow of solvent molecules through a semipermeable membrane driven by osmotic pressure differences across the membrane, to try to equal the concentration of the solute on the two sides of the membrane itself. In biological systems, water is the solvent and plasma membranes are the semipermeable membranes.
Plasma membranes allow water molecules to pass, due to protein channels, known as aquaporins, as well as small non-polar molecules that diffuse rapidly across them, whereas they are impermeable to ions and macromolecules.
Inside the cell there are macromolecules, such as nucleic acids, proteins, glycogen, and supramolecular aggregates, for example multienzyme complexes, but also ions in a higher concentration than that of the extracellular environment. This causes osmotic pressure to drive water from outside to inside the cell. If this net flow of water toward the inside of the cell is not counterbalanced, cell swells, and plasma membrane is distended until the cell bursts, that is, an osmotic lysis occurs. Under physiological conditions, this does not happen because during evolution several mechanisms have been developed to oppose, and in some cases even exploit, these osmotic forces. Two of these are energy-dependent ion pumps and, in plants, bacteria and fungi, the cell wall.
Energy dependent ion pumps
Ion pumps reduce, at the expense of ATP, the intracellular concentrations of specific ions with respect to their concentrations in the extracellular environment, thereby creating an unequal distribution of the ions across the plasma membrane, namely, an ion gradient. In this way the cell counterbalances the osmotic forces due to the ions and macromolecules trapped inside it. An example of energy-dependent ion pump is Na+/K+ ATPase, which reduces the concentration of Na+ inside the cell relative to the outside.
Cell wall
Plant cells are surrounded by an extracellular matrix, the cell wall, that, being non expandable and positioned next to the plasma membrane, allows cell to resist osmotic forces that would cause its swelling and finally the lysis. How?
Inside mature plant cells, the vacuoles are the largest organelles, occupying about 80% of the total cell volume. Large quantities of solutes, for the most part organic and inorganic acids, are accumulated within them and osmotically draw water, causing their swelling. In turn, this causes the tonoplast, the membrane that surrounds the vacuole, to press the plasma membrane against the cell wall, that mechanically opposes to these forces and avoids the osmotic lysis. This osmotic pressure is called turgor pressure, and can reach up 2 MPa, that is, 20 atmospheres, a value about 10 times higher than the air pressure in tires. It is responsible for the rigidity of non woody parts of plants, is involved in plant growth, as well as in:
wilting of vegetables, due to its reduction;
plants movements, such as:
the circadian movements of the leaves;
the movements of the leaves of Dionaea muscipula, the Venus flytrap, or of the leaves of the sensitive plants such as Mimosa pudica.
Even in bacteria and fungi, the plasma membrane is surrounded by a cell wall that stably withholds the internal pressure, then preventing osmotic lysis of the cell.
Isotonic, hypotonic, and hypertonic solutions
By comparing the osmotic pressure of two solutions separated by a semipermeable membrane, it is possible to define three types of solutions, briefly described below.
The solutions are isotonic when they have the same osmotic pressure.
If the solutions have different osmotic pressures, that with the higher osmotic pressure is defined hypertonic with respect to the other.
If the solutions have different osmotic pressures, that with the lower osmotic pressure is defined hypotonic with respect to the other.
In biological systems, the cytosol is the reference solution; then, if we place a cell in a:
isotonic solution, no net flow of water occurs into or out of the plasma membrane;
hypertonic solution, there is a net flow of water out of the cell, therefore the cell loses water and shrinks;
hypotonic solution, there is a net flow of water into the cell, the cell swells and can burst, i.e., an osmotic lysis can occur.
In addition to ion pumps and the cell wall, in the course of evolution multicellular organisms have developed another mechanism to oppose the osmotic forces: to surround the cells with an isotonic solution or close to isotonicity that prevents, or at least limits, a net inflow or outflow of water. An example is plasma, that is, the liquid component of blood, which, due to the presence of salts and proteins, primarily albumin in humans, has an osmolarity similar to that of the cytosol.
Osmotic pressure, starch and glycogen
Living organisms store glucose in the form of polymers, glycogen in animals, fungi, bacteria, and starch in photoautotrophs, but not in the free form. In this way, they avoid that the osmotic pressure exerted by the stores of carbohydrates becomes too high. Indeed, since osmotic pressure, like the other colligative properties, depends only on the number of solute molecules, storing millions of glucose units in the form of a significantly lower number of polysaccharides allows to avoid an excessive pressure. Here are some examples.
A gram of polysaccharide, e.g. glycogen or starch, composed of 1000 glucose units has an effect on osmotic pressure lower than that of a milligram of free glucose.
In hepatocyte, if the glucose stored in the form of glycogen was present in the free form, its concentration would be about 0.4 M, whereas the polysaccharide concentration of about 0.04 μM, and this would cause a net flow of water inside the cell such as to lead to osmotic lysis.
Furthermore, even if osmotic lysis could be avoided, there would be problems with the transport of glucose into the cell. In humans, under physiological conditions, blood glucose levels range from 3.33 to 5.56 mmol/L (60-100 mg/dL); if glucose was present in the free form, its intracellular concentration would be 120 to 72 times greater than that of the blood, and its transport into the hepatocyte would entail a large energy expenditure.
Osmotic diarrhea
In the presence of diseases that cause non-absorbed and osmotically active solutes accumulation in the distal portion of the small intestine and in the colon, a condition known as osmotic diarrhea occurs.
Causes can be, for example, bacterial infections, pancreatic diseases, celiac disease, an autoimmune enteropathy due to gluten intake in genetically predisposed subjects, or a congenital deficiency of one of the disaccharidases of the brush border of enterocytes, such as in lactose intolerance. In these conditions, an incomplete carbohydrate digestion can occur as a consequence of a deficit of alpha-amylase and/or of one or more disaccharidases. Moreover, unabsorbed osmotically active solutes pass into the colon where they can be fermented by bacteria of gut microbiota, which is part of the human microbiota, resulting in production of excessive gas, such as hydrogen, carbon dioxide and methane, and short-chain fatty acids, mainly butyric acid, acetic acid and propionic acid. This causes a condition known as osmotic-fermentative diarrhea.
Osmotic diarrhea can also result from the use of osmotic laxatives such as polyethylene glycol or PEG, and magnesium sulfate.
Osmotically active solutes deriving from incomplete digestion, and osmotic laxatives lead to an increase in intraluminal osmotic pressure and inhibit the normal absorption of water and electrolytes, causing a reduction in the consistency of the stool and an increase in intestinal motility.
Osmotic pressure, galactosemia, and cataracts
Galactose, glucose and fructose are the monosaccharides that are absorbed in the intestine.
Galactose is metabolized mostly in the liver, and to a lesser extent by other organs and tissues. After conversion to UDP-glucose and UDP-galactose through the Leloir pathway, it can be used for both anabolic and catabolic purposes.
Mutations in one of the genes encoding the enzymes of the Leloir pathway leads to an accumulation of galactose and causes galactosemia, a genetic metabolic disorder whose only treatment is a galactose-restricted diet.
The accumulation of the monosaccharide fuels alternative metabolic pathways that lead to the synthesis of the galactose-related chemicals galactitol and galactonate.
The symptoms of galactosemia include cataracts, and the synthesis and accumulation of galactitol in the lens of the eye seems to be one of the triggers. Why? Galactitol is a poorly metabolized polyol, and, due to its poor lipophilicity does not diffuse through cell membranes and accumulates inside the cell. Being osmotically active, its accumulation causes an increase in intracellular osmotic pressure and then a net flow of water into the cell. Such osmotic effect seems to be one of the mechanisms through which galactitol contributes to the development of galactosemic cataracts.
References
Beauzamy L., Nakayama N., and Boudaoud A. Flowers under pressure: ins and outs of turgor regulation in development. Ann Bot 2014;114(7):1517-1533. doi:10.1093/aob/mcu187
Berg J.M., Tymoczko J.L., and Stryer L. Biochemistry. 5th Edition. W. H. Freeman and Company, 2002
Coelho A.I., Berry G.T., Rubio-Gozalbo M.E. Galactose metabolism and health. Curr Opin Clin Nutr Metab Care 2015;18(4):422-427. doi:10.1097/MCO.0000000000000189
Coelho A.I., Rubio-Gozalbo M.E., Vicente J.B., Rivera I. Sweet and sour: an update on classic galactosemia. J Inherit Metab Dis 2017;40(3):325-342. doi:10.1007/s10545-017-0029-3
Conte F., van Buuringen N., Voermans N.C., Lefeber D.J. Galactose in human metabolism, glycosylation and congenital metabolic diseases: time for a closer look. Biochim Biophys Acta Gen Subj 2021;1865(8):129898. doi:10.1016/j.bbagen.2021.129898
Multifunctional enzymes are proteins in which two or more enzymatic activities, that catalyze consecutive steps of a metabolic pathway, are located on the same polypeptide chain. It seems likely they have arisen by gene fusion events, and represent, like multienzyme complexes, a strategy of evolution to maximize catalytic efficiency, providing advantages that you wouldn’t have if such enzymatic activities were present on distinct proteins dissolved in the cytosol.
What advantages do multifunctional enzymes provide?
Living organisms fight against the natural processes of decay that, if not counteracted, leads to an increasing disorder, until death. At the molecular level, the maintenance of life is made possible by the extraordinary effectiveness achieved by enzymes in accelerating chemical reactions and avoiding side reactions. The rate of ATP turnover in a mammalian cell gives us an idea of the rate at which cellular metabolism proceeds: every 1-2 minutes, the entire ATP pool is turned over, namely, hydrolyzed and restored by phosphorylation. This corresponds to the turnover of about 107 molecules of ATP per second, and, for the human body, to the turnover of about 1 gram of ATP every minute. Some enzymes have even achieved catalytic perfection, that is, they are so efficient that nearly every collision with their substrates results in catalysis. And one of the factors limiting the rate of an enzymatic reaction is just the frequency with which substrates and enzymes collide. The simplest way to increase the frequency of collisions would be to increase the concentration of substrates and enzymes. However, due to the large number of different reactions that take place in the cell, this route is not feasible. In other words, there is a limit to the concentration that substrates and enzymes can reach, concentrations that are in the micromolar range for substrates, and even lower for enzymes. Exceptions are the enzymes of glycolysis in muscle cells and erythrocytes, present in concentrations of the order of 0.1 mM and even higher.
Metabolic channeling
One of the routes taken by evolution to increase the rate at which enzymatic reactions proceed is to select molecular structures, such as multifunctional enzymes and multienzyme complexes, that allow, through the optimization of the spatial organization of the enzymes of a metabolic pathway, to minimize the distance that the product of reaction A must travel to reach the active site that catalyzes the reaction B in the sequence, and so on, thus obtaining the substrate or metabolic channeling of the pathway itself. For some multifunctional enzymes and multienzyme complexes the channeling is obtained through intramolecular channels.
Metabolic channeling increases the catalytic efficiency, and then the reaction rate, in various ways, briefly described below.
It minimizes the diffusion of substrates in the bulk solvent, then their dilution; this allows to obtain high local concentrations, even when their concentration in the cell is low, thus increasing the frequency of enzyme-substrate collisions.
It minimizes the time required by substrates to diffuse from one active site to the next.
It minimizes the probability of side reactions.
It minimizes the probability that labile intermediates are degraded.
Multifunctional enzymes offer advantages with regard to the regulation of their synthesis, too: being encoded by a single gene, it is possible to coordinate the synthesis of all the enzymatic activities.
Finally, like multienzyme complexes, multifunctional enzymes allow the coordinated control of their catalytic activities. And, because the enzyme that catalyzes the committed step of the sequence often catalyzes the first reaction, this prevents the synthesis of unneeded molecules, which would be produced if the control point were downstream of the first reaction, as well as a waste of energy and the removal of metabolites from other metabolic pathways.
Examples of multifunctional enzymes
Like multienzyme complexes, multifunctional enzymes, too, are very common and involved in many metabolic pathways, both anabolic and catabolic.
Here are some examples.
Acetyl-CoA carboxylase
Acetyl-CoA carboxylase or ACC (EC 6.4.1.2), a biotin-dependent carboxylase, is composed of two enzymes, a biotin carboxylase (EC 6.3.4.14) and a carboxyltransferase, plus a biotin carboxyl-carrier protein or BCCP. ACC catalyzes the synthesis of malonyl-CoA by the carboxylation of acetyl-CoA. The reaction, which is the committed step of the synthesis of fatty acids, proceeds in two steps. In the first step, biotin carboxylase catalyzes, at the expense of ATP, the carboxylation of a nitrogen atom of biotin, that acts as a carbon dioxide (CO2) carrier, while the source of CO2 is bicarbonate ion. In the second step, carboxyltransferase catalyzes the transfer of the carboxyl group from carboxybiotin to acetyl-CoA to form malonyl-CoA. Malonyl-CoA is the donor of an activated two carbon unit to fatty acid synthase (EC 2.3.1.85) during fatty acid elongation.
In mammals and birds, acetyl-CoA carboxylase is a multifunctional enzyme, as biotin carboxylase activity and carboxyltransferase activity, plus BCCP, are located on the same polypeptide chain.
Conversely, in bacteria it is a multienzyme complex made up of three distinct polypeptide chains, namely, the two enzymes plus BCCP.
Both forms are present in higher plants.
Type I fatty acid synthase
Fatty acid synthase or FAS catalyzes the synthesis of palmitic acid using malonyl-CoA, the product of the reaction catalyzed by acetyl-CoA carboxylase, as a donor of two-carbon units.
There are two types of FAS.
In fungi and animals, it is a multifunctional enzyme, and is called type I. The animal enzyme is an homodimer, and each polypeptide chain contains all seven enzymatic activities plus acyl carrier protein or ACP. In yeast and fungi FAS consists of two multifunctional subunits, called α and β, arranged in an α6β6 heterododecameric structure.
In most prokaryotes and in plants, fatty acid synthase, called type II, it is not a multifunctional enzyme but a multienzyme complex, being composed of distinct enzymes plus ACP.
PRA-isomerase:IGP synthase
The synthesis of the amino acid tryptophan from chorismate involves several steps, briefly described below.
In the first step, glutamine donates a nitrogen to the indole ring of chorismate, that is converted to anthranilate, and glutamine to glutamate; the reaction is catalyzed by anthranilate synthase (EC 4.1.3.27). Anthranilate is phosphoribosylated to form N-(5’-phosphoribosyl)-anthranilate or PRA, in a reaction catalyzed by anthranilate phosphoribosyltransferase (EC 2.4.2.18); in the reaction 5-phosphoribosyl-1-pyrophosphate or PRPP acts as a donor of a 5-phosphoribosyl group. In the next step, catalyzed by PRA isomerase (EC 5.3.1.24), PRA is isomerized to enol-1-o-carboxyphenylamino-1-deoxyribulose phosphate or CdRP. PRA e CdRP are an example of structural isomerism.
CdRP is converted to indole-3-glycerol phosphate or IGP, in a reaction catalyzed by indole-3-glycerol phosphate synthase or IGP synthase (EC 4.1.1.48). Finally, tryptophan synthase (EC 4.2.1.20) catalyzes the last two steps of the pathway: the conversion of IGP to indole, a hydrolysis, and the reaction of indole with a serine to form tryptophan.
In E. coli, PRA isomerase and IGP synthase are located on a single polypeptide chain, which is therefore a bifunctional enzyme. In other microorganisms, such as Bacillus subtilis, Salmonella typhimurium and Pseudomonas putida, the two catalytic activities located on distinct polypeptide chains.
Conversely, tryptophan synthase is a classic example of a multienzyme complex, and one of the best characterized examples of metabolic channeling.
Glutamine-PRPP amidotransferase
Glutamine-PRPP amidotransferase or GPATase (EC 2.4.2.14) catalyzes the first of ten steps leading to de novo synthesis of purine nucleotides, namely, the formation of 5-phosphoribosylamine through the transfer of the glutamine amide nitrogen to PRPP. Note that glutamine acts as a nitrogen donor.
The reaction proceeds in two steps, which take place on different active sites, an N-terminal active site and a C-terminal active site.
In the first step, the N-terminal active site catalyzes the hydrolysis of glutamine amide nitrogen to form glutamate and ammonia.
During the second step, catalyzed by the C-terminal active site, which has phosphoribosyltransferase activity, the released ammonia is attached at the C-1 of PRPP to form 5-phosphoribosylamine. In this step the inversion of configuration about the C-1 position of the ribose, from α to β, occurs, then establishing the anomeric form of the future nucleotide.
There are three control points that cooperate in the regulation of de novo synthesis of purine nucleotides, and the reaction catalyzed by glutamine-PRPP amidotransferase, the first committed step of the pathway, is the first control point.
Like in bacterial carbamoyl phosphate synthetase complex (EC 6.3.4.16), the active sites of this multifunctional enzyme are connected through an intramolecular channel. However, this channel is shorter, being about 20 Å long, and lined by conserved nonpolar amino acids, then, it is highly hydrophobic. Lacking hydrogen-bonding groups, it does not impede the diffusion of the ammonia to the other active site.
CAD
The de novo synthesis of pyrimidine nucleotides occurs through a series of enzymatic reactions that, unlike de novo synthesis of purine nucleotides, begins with the formation of the pyrimidine ring, which is then bound to ribose 5-phosphate. The first three steps of the pathway are catalyzed sequentially by carbamoyl phosphate synthetase, aspartate transcarbamoylase (EC 2.1.3.2), and dihydroorotase (EC 3.5.2.3), and are common to all species.
In the first step, carbamoyl phosphate synthetase, which has two enzymatic activities, namely, a glutamine-dependent amidotransferase and a synthase, catalyzes the synthesis of carbamoyl phosphate from glutamine, bicarbonate ion and ATP. In the second step, which is the committed step of the metabolic pathway and is catalyzed by aspartate transcarbamoylase, carbamoyl phosphate reacts with aspartate to form N-carbamoyl aspartate. Finally, dihydroorotase, catalyzing the removal of H2O from N-carbamoyl aspartate, leads to the closure of the pyrimidine ring to form of L-dihydroorotate.
In eukaryotes, particularly in mammals, in Drosophila and Dictyostelium, a genus of amoebae, the three enzymatic activities are located on a single polypeptide chain, encoded by a gene derived from a gene fusion event occurred at least 100 million years ago. The multifunctional enzyme, known by the acronym CAD, is a homomultimer of three subunits or more.
Conversely, in prokaryotes, the three enzymes are distinct, and carbamoyl phosphate synthase is an example of a multienzyme complex.
In yeasts the dihydroorotase is present on a distinct protein.
Studies on enzyme activity have revealed the existence of a substrate channeling, more effective in yeast protein, with respect to the first two steps, than in that of mammals.
References
Alberts B., Johnson A., Lewis J., Morgan D., Raff M., Roberts K., Walter P. Molecular Biology of the Cell. 6th Edition. Garland Science, Taylor & Francis Group, 2015
Eriksen T.A., Kadziola A., Bentsen A-K., Harlow K.W. & Larsen S. Structural basis for the function of Bacillus subtilis phosphoribosyl-pyrophosphate synthetase. Nat Struct Biol 2000:7;303-308. doi:10.1038/74069
Hyde C.C., Ahmed S.A., Padlan E.A., Miles E.W., and Davies D.R. Three-dimensional structure of the tryptophan synthase multienzyme complex from Salmonella typhimurium. J Biol Chem 1988:263(33);17857-17871. doi:10.1016/S0021-9258(19)77913-7
Hyde C.C., Miles E.W. The tryptophan synthase multienzyme complex: exploring structure-function relationships with X-ray crystallography and mutagenesis. Nat Biotechnol 1990:8;27-32. doi:10.1038/nbt0190-27
Michal G., Schomburg D. Biochemical pathways. An atlas of biochemistry and molecular biology. 2nd Edition. John Wiley J. & Sons, Inc. 2012
Muchmore C.R.A., Krahn J.M, Smith J.L., Kim J.H., Zalkin H. Crystal structure of glutamine phosphoribosylpyrophosphate amidotransferase from Escherichia coli. Protein Sci 1998:7;39-51. doi:10.1002/pro.5560070104
Nelson D.L., Cox M.M. Lehninger. Principles of biochemistry. 6th Edition. W.H. Freeman and Company, 2012
Pareek V., Sha Z., He J., Wingreen N.S., Benkovic SJ. Metabolic channeling: predictions, deductions, and evidence. Mol Cell 2021;81(18):3775-3785. doi:10.1016/j.molcel.2021.08.030
Yon-Kahn J., Hervé G. Molecular and cellular enzymology. Springer, 2009
Multienzyme complexes are discrete and stable structures composed of enzymes associated noncovalently that catalyze two or more sequential steps of a metabolic pathway.[7]
They can be considered a step forward in the evolution of catalytic efficiency as they provide advantages that individual enzymes, even those that have achieved catalytic perfection, would not have alone.[1][12]
During evolution, some enzymes evolved to reach a virtual catalytic perfection, namely, for such enzymes nearly every collision with their substrate results in catalysis. Examples are:
fumarase (EC 4.2.1.2), which catalyzes the seventh reaction of the citric acid cycle, the reversible hydration/dehydration of fumarate’s double bond to form malate;
acetylcholinesterase (EC 3.1.1.7), which catalyses the hydrolysis of acetylcholine, a neurotransmitter, to choline and acetic acid, which in turn dissociates to form an hydrogen ion and acetate;
superoxide dismutase (EC 1.15.1.1), which catalyzes the conversion, and then the inactivation, of the highly reactive superoxide radical, (O2.-), to hydrogen peroxide (H2O2) and water;
catalase (EC 1.11.1.6), which catalyzes the degradation of H2O2 to water and oxygen.
Then, the rate at which an enzymatic reaction proceeds is partly determined by the frequency with which enzymes and their substrates collide. Hence, a simple way to increase it is to increase the concentrations of enzymes and substrates. However, their concentrations cannot be high because of the enormous number of different reactions that occur within the cell. And in fact in the cells most reactants are present in micromolar concentrations (10-6 M), whereas most enzymes are present in much lower concentrations.[1][12]
So, evolution has taken different routes to increase the reaction rate, one of which has been to optimize the spatial organization of enzymes with the formation of multienzyme complexes and multifunctional enzymes, that is, structures that allow minimizing the distance that the product of a reaction must travel to reach the active site that catalyzes the subsequent step in the sequence, being active sites close to each other.[2][7] In other words, what happens is the substrate or metabolic channeling,[8][9][14] that can also occur through intramolecular channels connecting the active sites, as in the case of, among the multienzyme complexes, tryptophan synthase complex (EC 4.2.1.20),[3][4] whose tunnel was the first to be discovered, and bacterial carbamoyl phosphate synthase complex (EC 6.3.4.16).[5][13]
Metabolic channeling can increase the reaction rate, but more generally, the catalytic efficiency, in several ways, briefly described below.[8][12]
The diffusion of substrates and products in the bulk solvent is minimized, then their dilution and decrease of concentration, too. This leads to the production of high local concentrations, even when their intracellular concentration is low. In turn this leads to an increase in the frequency of enzyme-substrate collisions.
The time required by substrates to diffuse between successive active sites is minimized.
The probability of side reactions is minimized.
Chemically labile intermediates are protected from degradation by the solvent.
Another metabolic advantage of multienzyme complexes, similarly to what happens with multifunctional enzymes, is that they allow to control coordinately the catalytic activity of the enzymes that compose them. And taking into account that the enzyme that catalyzes the first reaction of a pathway is often the regulatory enzyme, it is possible to avoid:
the synthesis of unneeded intermediates, which would be produced if the sequence of reactions were regulated downstream of the first reaction;
the removal of metabolites from other pathways as well as a waste of energy.
Examples
From what was said above, it is not surprising that, especially in eukaryotes, the multienzyme complexes, like multifunctional enzymes, are common and involved in different metabolic pathways, both anabolic and catabolic, whereas there are few enzymes that diffuse freely in solution. Below are some examples.
2-Ketoacid dehydrogenase family
A classic example of multienzyme complexes are the three complexes belonging to the 2-ketoacid dehydrogenase family, also called 2-oxoacid dehydrogenase family, namely:
the branched-chain alpha-keto acid dehydrogenase complex (BCKDH);
the alpha-ketoglutarate dehydrogenase complex, also called 2-oxoglutarate dehydrogenase (OGDH).[2][6][7][10]
Alpha-Ketoglutarate Dehydrogenase Complex
These complexes are similar both from structural and functional points of view.
For example, PDC is composed of multiple copies of three different enzymes:
pyruvate dehydrogenase or E1 (EC 1.2.4.1);
dihydrolipoyl transacetylase or E2 ;(EC 2.3.1.12);
dihydrolipoyl dehydrogenase or E3 (EC 1.8.1.4).
Then, PDC, both in prokaryotes and eukaryotes, has the basic E1-E2-E3 structure, a structure also found in the other two complexes.[15] Moreover, within a given species:
dihydrolipoyl dehydrogenase is identical;
pyruvate dehydrogenase and dihydrolipoyl transacetylase are homologous.
And, although these enzymes are specific for their substrates, they use the same cofactors, namely, coenzyme A, NAD, thiamine pyrophosphate, FAD, and lipoamide.
In order to differentiate them, for the pyruvate dehydrogenase complex, the alpha-ketoglutarate dehydrogenase complex, and the branched-chain alpha-ketoacid dehydrogenase complex, they are indicated, respectively:
E1p, E1o and E1b (EC 1.2.4.4);
E3p, E3o, and E3b (EC 1.8.1.4).
Note: the eukaryotic PDC is the largest multienzyme complex known, larger than a ribosome, and can be visualized with the electron microscope.[2]
The pyruvate dehydrogenase complex is the bridge between glycolysis and the citric acid cycle, and catalyzes the irreversible oxidative decarboxylation of pyruvate, which is the conjugate base of pyruvic acid and belongs to the group of keto acids, in particular it is an alpha-ketoacid. During the reactions the carboxyl group of pyruvate is released as carbon dioxide (CO2) and the resulting acetyl group is transferred to coenzyme A to form acetyl-coenzyme A. Furthermore, two electrons are released and transferred to NAD+.
Even during the reactions catalyzed by the alpha-ketoglutarate dehydrogenase complex and the branched-chain alpha-keto acid dehydrogenase complex, respectively, the fourth reaction of the citric acid cycle, the oxidation of alpha-ketoglutarate to succinyl-CoA, and the oxidation of alpha-keto acids deriving from the catabolism of the branched-chain amino acids valine, leucine and isoleucine, it occurs:
the release of the carboxyl group of the α-keto acid as CO2;
the transfer of the resulting acyl group to coenzyme A to form the acyl-CoA derivatives;
the reduction of NAD+ to NADH.
The remarkable similarity between protein structures, required cofactors and reaction mechanisms undoubtedly reflect a common evolutionary origin.
Tryptophan synthase complex
The tryptophan synthase complex is one of the best-studied examples of substrate channeling.[3][12][14] Present in bacteria and plants, but not in animals, in bacteria it is composed of two α and two β subunits associated as αβ dimers, which are considered the functional unit of the complex, in turn associated to form an αββα tetramer.[4][5][8]
The complex catalyzes the final two steps of the synthesis of tryptophan. In the first step, indole-3-glycerol phosphate undergoes an aldol cleavage, catalyzed by a lyase (EC 4.1.2.8) present on the α subunits, to yield indole and a molecule of glyceraldehyde 3-phosphate. Indole then reaches the active site of the β subunit via a about 30 Å long hydrophobic tunnel that, in each αβ dimers, connects the two active sites. In the second step, in the presence of pyridoxal 5-phosphate, a condensation between indole and a serine forms tryptophan.
Acetyl-CoA carboxylase
Acetyl-CoA carboxylase (ACC) (EC 6.4.1.2), a member of the biotin-dependent carboxylase family, catalyzes the committed step of de novo synthesis of fatty acids, namely, the carboxylation of acetyl-CoA to malonyl-CoA, which, in turn, serves as a donor of two-carbon units for the elongation process leading to the synthesis of palmitic acid, catalyzed by fatty acid synthase (EC 2.3.1.85).[9][11][12]
In bacteria, ACC is an multienzyme complex composed of two enzymes, biotin carboxylase (EC 6.3.4.14) and a carboxytransferase, plus a biotin carboxyl-carrier protein or BCCP.
Conversely, in mammals and birds, it is a multifunctional enzyme, as the two enzymatic activities, and BCCP, are present on the same polypeptide chain.[2]
In higher plants both forms are present.
Carbamoyl phosphate synthetase complex
Another well-characterized example of substrate channeling is the bacterial carbamoyl phosphate synthetase complex,[9][12][14] which catalyzes the synthesis of carbamoyl phosphate, needed for pyrimidine and arginine synthesis. The complex has a about 100 Å long tunnel that connects the three active sites.[13]
The first active site catalyzes the release of the amide nitrogen of glutamine as ammonium ion, that enters the tunnel and reaches the second active site where, at the expense of ATP, is combined with bicarbonate to yield carbamate, that, in the last active site, is phosphorylated to carbamoyl phosphate.
References
^ ab Alberts B., Johnson A., Lewis J., Morgan D., Raff M., Roberts K., Walter P. Molecular biology of the cell. 6th Edition. Garland Science, Taylor & Francis Group, 2015
^ ab Hilario E., Caulkins B.G., Huang Y-M. M., You W., Chang C-E. A., Mueller L.J., Dunn M.F., and Fan L. Visualizing the tunnel in tryptophan synthase with crystallography: insights into a selective filter for accommodating indole and rejecting water. Biochim Biophys Acta 2016;1864(3):268-279. doi:10.1016/j.bbapap.2015.12.006
^ ab Hyde C.C., Ahmed S.A., Padlan E.A., Miles E.W., and Davies D.R. Three-dimensional structure of the tryptophan synthase multienzyme complex from Salmonella typhimurium. J Biol Chem 1988;263(33):17857-17871. doi:10.1016/S0021-9258(19)77913-7
^ ab Hyde C.C., Miles E.W. The tryptophan synthase multienzyme complex: exploring structure-function relationships with X-ray crystallography and mutagenesis. Nat Biotechnol 1990:8;27-32. doi:10.1038/nbt0190-27
^ Koolman J., Roehm K-H. Color atlas of Biochemistry. 2nd Edition. Thieme, 2005
^ abc Michal G., Schomburg D. Biochemical pathways. An atlas of biochemistry and molecular biology. 2nd Edition. John Wiley J. & Sons, Inc. 2012
^ abc Moran L.A., Horton H.R., Scrimgeour K.G., Perry M.D. Principles of Biochemistry. 5th Edition. Pearson, 2012
^ abc Nelson D.L., Cox M.M. Lehninger. Principles of biochemistry. 6th Edition. W.H. Freeman and Company, 2012
^ Perham R.N., Jones D.D., Chauhan H.J., Howard MJ. Substrate channeling in 2-oxo acid dehydrogenase multienzyme complexes. Biochem Soc Trans 2002;30(2):47-51. doi:10.1042/bst0300047
The pyruvate dehydrogenase complex (PDC) is one of the mitochondrial multienzyme complexes, and is composed of three different enzymes:
pyruvate dehydrogenase or E1 (EC 1.2.4.1);
dihydrolipoyl transacetylase or dihydrolipoamide acetyltransferase or E2 (EC 2.3.1.12);
dihydrolipoyl dehydrogenase or dihydrolipoamide dehydrogenase or E3 (EC 1.8.1.4).
Each of these enzymes is present in multiple copies whose number, and then the size of the complex itself, varies from species to species, with molecular masses ranging from 4×106 to 1×107 daltons.
The multienzyme complex contains additional subunits:
five different coenzymes;
in plants, fungi, and among animals, aves and mammals, two enzymes with regulatory properties: a Mg2+-dependent pyruvate dehydrogenase kinase (EC 2.7.1.99) and a Ca2+-activated pyruvate dehydrogenase phosphatase(EC 3.1.3.43);
in eukaryotes, a binding protein called E3BP.
The pyruvate dehydrogenase complex catalyzes, through five sequential reactions, the oxidative decarboxylation of pyruvate, the conjugate base of pyruvic acid, to form a carbon dioxide molecule (CO2) and the acetyl group of acetyl-coenzyme A or acetyl-CoA, with the release of two electrons, carried by NAD.
PDC Reaction
The overall reaction is essentially irreversible, with a ΔG°’ of -8.0 kcal/mol (-33.4 kJ/mol), and requires the intervention of the three enzymes, whose activities are sequentially coordinated. During the reactions, the intermediate products remain bound to the enzymes and, at the end of the reaction sequence, the multienzyme complex is ready for the next cycle.
The Five Reactions Catalyzed by the PDC
Note: the pyruvate dehydrogenase complex catalyzes the same reactions through similar mechanisms in all organisms.
Five coenzymes are used in the pyruvate dehydrogenase complex reactions. They are thiamine pyrophosphate (TPP), flavin adenine dinucleotide (FAD), coenzyme A (CoA), nicotinamide adenine dinucleotide (NAD), and lipoic acid.
Thiamine pyrophosphate is the active form of thiamine or vitamin B1. TPP is the coenzyme of pyruvate dehydrogenase, to which it is strictly bound through noncovalent interactions. It is involved in the transfer of hydroxyethyl or “activated aldehyde” groups.
Flavin adenine dinucleotide is one the active forms of riboflavin or vitamin B2; the other is flavin mononucleotide (FMN).
Reduced and Oxidized Form of Flavin Adenine Dinucleotide
FAD is the coenzyme of dihydrolipoyl dehydrogenase, to which it is strictly bound. Like NAD, it participates in electron transfer, or hydride ion (:H– or H+ + 2e–) transfer.
Coenzyme A consists of a β-mercaptoethylamine group connected to pantothenic acid or vitamin B5 through an amide linkage, which, in turn, is bonded to 3′-phosphoadenosine moiety, through a pyrophosphate bridge.
CoA is involved in the reaction catalyzed by dihydrolipoyl transacetylase, and acts as a carrier of acyl groups.
Coenzyme A and Acetyl-coA
The β-mercaptoethylamine moiety terminates with a sulfhydryl group (–SH), a reactive thiol crucial for the role played by the coenzyme, because the acyl groups bonded to it through a thioester bond have a high standard free energy of hydrolysis. This provides acyl groups with a high transfer potential, equal to -31.5 kcal/mol (-7.5 kJ/mol), slightly more exergonic [1 kJ/mol (0.2 kcal/mol )] than that for the hydrolysis of ATP to ADP and Pi. Thioesters have therefore a high transfer potential of the acyl group and can donate it to a variety of molecules, that is, such acyl group can be considered as an activated group ready for a transfer. It is also possible to state that the formation of the thioester bond allows to conserve a portion of the free energy derived from the oxidation of the metabolic fuel. It should be noted that coenzyme A is also abbreviated as CoA-SH to emphasize the role played by the thiol group.
Note: in the thioester bond a sulfur atom sits in the position where an oxygen atom is in the ester bond.
Ester and Thioester Bonds
Nicotinamide adenine dinucleotide can be synthesized from tryptophan, an essential amino acid, or from niacin or vitamin B3 or vitamin PP, from Pellagra-Preventing, the source of the nicotinamide moiety.
Reduced and Oxidized Form of Nicotinamide Adenine Dinucleotide Phosphate
NAD is involved in the reaction catalyzed dihydrolipoyl dehydrogenase, and, like FAD, participates in electron transfer, or hydride transfer.
Unlike the other coenzymes of the pyruvate dehydrogenase complex, lipoic acid does not derive, directly or indirectly, from vitamins and/or essential amino acids, that is, from building blocks that cannot be synthesized de novo by the organism and must be supplied by the diet.
It is the coenzyme of dihydrolipoyl transacetylase, to which it is covalently bound, through an amide linkage, to the ɛ-amino group of a lysine residue to form a lipoyl-lysine or lipoamide, the so-called lipoyllysyl arm. It couples electron transfer to acyl group transfer.
Lipoamide or Lipoyl-lysine
Lipoic acid has two thiol groups that can undergo a reversible intramolecular oxidation to form a disulfide bridge (-S-S-), a reaction analogous to that between two cysteine (Cys) residues of a protein.
Because the disulfide bridge (note: a cyclic disulfide) is capable of undergoing redox reactions, during the reactions catalyzed by the pyruvate dehydrogenase complex, it is first reduced to dihydrolipoamide, a dithiol or the reduced form of the prosthetic group, and then, reoxidized to the cyclic form.
Note
Many enzymes require small non protein components, called cofactors, for their catalytic activity. Cofactors can be metal ions or small organic or metalloorganic molecules, and are classified as coenzymes and prosthetic groups.
A prosthetic group is cofactor that binds tightly to an enzyme by non-covalent or covalent bond, namely, it is permanently bound to the protein.
A coenzyme is cofactor that is not permanently bound to the enzyme.
Where is the pyruvate dehydrogenase complex located?
In eukaryotes, the pyruvate dehydrogenase complex, like the enzymes for citric acid cycle and oxidation of fatty acids, is located in the mitochondrion, where is associated with the surface of the inner membrane facing the matrix.
In prokaryotes, it is located in the cytosol.
Functions of the pyruvate dehydrogenase complex
The main functions of the pyruvate dehydrogenase complex are to produce acetyl-CoA and NADH.
The acetyl group linked to coenzyme A, an activated acetate, depending on the metabolic conditions within the cell and/or the cell type, can be:
oxidized to two carbon dioxide molecules via the citric acid cycle reactions to harvest a portion of the potential energy stored in the form of ATP or GTP;
utilized for the synthesis of fatty acids, cholesterol, steroids, isoprenoids, ketone bodies and acetylcholine.
It is therefore possible to state that, depending on the metabolic conditions and/or cell type, the pyruvate dehydrogenase complex commits carbon intermediates from amino acid and glucose catabolism to:
citric acid cycle, and then to the production of energy, e.g. in skeletal muscle in aerobic conditions, and, always, in cardiac muscle;
synthesis of lipids and acetylcholine.
In aerobic organisms, NADH can be oxidized to NAD+ via hydride ion transfer to the mitochondrial electron transport chain that, in turn, carries the two electrons to molecular oxygen (O2), allowing the production of 2.5 ATP molecules per pair of electrons.Note: in anaerobic organisms there are electron acceptors alternative to oxygen, such as sulfate or nitrate.
Conceptually, the pyruvate dehydrogenase complex is the bridge between glycolysis and the citric acid cycle. However, due to the irreversibility of the overall reaction catalyzed by the multienzyme complex, it acts as an one-way bridge: pyruvate can be decarboxylated, oxidized and the remaining acetyl unit linked to the CoA, but it is not possible to carry out the opposite reaction, namely, to convert acetyl-CoA into pyruvate.
The irreversibility of this reaction, and the absence of alternative pathways, explain why it is not possible to use acetyl-CoA, and therefore fatty acids, as a substrate for gluconeogenesis.
Other sources of acetyl-CoA
Other than pyruvate, the acetyl group of acetyl-CoA can derive from the oxidation of fatty acids and the catabolism of many amino acids. However, regardless of its origin, acetyl-CoA represents an entry compound for new carbon units into the citric acid cycle. And, it is also possible to state that the acetyl group of acetyl-CoA represents the form in which most of the carbon enters the cycle.
Mitochondrial pyruvate transport
In eukaryotes, glycolysis occurs in the cytosol whereas all the subsequent steps of the aerobic metabolism, that is, the reactions catalyzed by the pyruvate dehydrogenase complex, the citric acid cycle, the electron transport chain, and the oxidative phosphorylation occur in the mitochondria.
Similarly to most other metabolites and anions, the transport of pyruvate across the outer mitochondrial membrane is probably mediated by a relatively non-specific, voltage-dependent anion channel. Conversely, its transport across the inner mitochondrial membrane occurs through a specific transporter made up of two proteins named MPC1 and MPC2, acronym of mitochondrial pyruvate carrier, which form a hetero-oligomeric complex in the membrane.
Structure of the pyruvate dehydrogenase complex
Although the pyruvate dehydrogenase complex is composed of multiple copies of three different enzymes and catalyzes the same reactions by similar mechanisms in all the organisms in which it is present, it has a very different quaternary structure.
The structure of E. coli multienzyme complex, which has a weight of ∼4,600 kD and a diameter of ∼300 Å, was the first to be characterized, thanks to the work of Lester Reed. In this complex, 24 units of dihydrolipoyl transacetylase form a structure with cubic symmetry, namely, the enzymes, associated as trimers, are placed at the corners of a cube. Dimers of pyruvate dehydrogenase are associated with dihydrolipoyl transacetylase core, at the center of each edge of the cube, for a total of 24 units. Finally, dimers of dihydrolipoyl dehydrogenase are located at the center of each of the six faces of the cube, for a total of 12 units. Note that the entire complex is composed of 60 units. A similar structure with cubic symmetry is also found in most other Gram-negative bacteria.
In some Gram-positive bacteria and in eukaryotes, the pyruvate dehydrogenase complex has a dodecahedral form, namely, that of a regular polyhedron with 20 vertices, 12 pentagonal faces, and 30 edges, with icosahedral symmetry, also called I symmetry. Considering for example the multienzyme complex present in mitochondria, it is the largest known multienzyme complex, with a weight of ∼10,000 kD and a diameter of ∼500 Å, so more than 5 times the size of a ribosome. Noteworthy, it can be visualized with the electron microscope. The complex is composed of a dodecahedral core, with a diameter of about 25 nm, formed, as in Gram-negative bacteria, by dihydrolipoyl transacetylase, but consisting of 20 trimers of the enzyme, for a total of 60 units, located at the vertices of the structure. Its core is surrounded by 30 units of pyruvate dehydrogenase, one centered on each edge, and 12 units of dihydrolipoyl dehydrogenase, one centered on each face. The entire complex is therefore composed of 102 units.
The quaternary structure of the complex is further complicated, as mentioned previously, by the presence of three additional subunits: a pyruvate dehydrogenase kinase, a pyruvate dehydrogenase phosphatase, and the E3-binding protein.
Kinase and phosphatase are bound to the dihydrolipoyl transacetylase core.
E3BP is bound to each of the 12 pentagonal faces, and therefore is present in about 12 copies. It is required to bind dihydrolipoyl dehydrogenase to the core of dihydrolipoyl transacetylase, as demonstrated by the fact that its partial proteolysis decreases the binding ability of the dehydrogenase. In E3-binding protein it is possible to identify a C-terminal domain, that has no catalytic activity, and a lipoamide-containing domain, similar to that of dihydrolipoyl transacetylase, capable of accepting an acetyl group, too. However, the removal of this domain does not cause any reduction of the catalytic activity of the multienzyme complex.
Structure of pyruvate dehydrogenase or E1
Pyruvate dehydrogenase of eukaryotes and some Gram-positive bacteria is composed of two different polypeptide chains, called α and β, associated to form a 2-fold symmetric α2β2 heterotetramer. Conversely, in E. coli and other Gram-negative bacteria the two subunits are fused to form a single polypeptide chain, and the enzyme is a homodimer.
The enzyme has two active sites.
Considering the heterotetrameric structure of Bacillus stearothermophilus pyruvate dehydrogenase, a Gram-positive bacteria, each thiamine pyrophosphate binds between the N-terminal domains of an α and a β subunit, at the end of a ∼21 Å deep funnel-shaped channel leading to the active site, with its reactive group, the thiazole ring, closest to the channel entrance. At the entrance this channel there are also two conserved loops, essential both for the catalytic activity of the enzyme and for its regulation. The X-ray analysis of B. stearothermophilus enzyme, when it binds both TPP and the peripheral subunit-binding domain (PSBD) of dihydrolipoyl transacetylase, which binds to the C-terminal domain of the β subunits, has revealed that, in addition to a heterotetramer with a very tight structure, the two active sites have a different structure, in particular regarding the arrangement of the two conserved loops. In fact, in one enzyme subunit, in the presence of the activated form of thiamine pyrophosphate, the inner loop is ordered in a way that it blocks the entrance to the active site, whereas the loop at the entrance of the other active site is disordered and does not block the entrance. This explains, from a structural point of view, the observed differences in the rate of substrate binding exhibited by the two active sites. A similar arrangement and asymmetry have been observed in all thiamine pyrophosphate-dependent enzymes of which the structure has been solved.
In addition to TPP and a magnesium ion (Mg2+), located in each of the two active sites, a third Mg2+ is located at the centre of the tetramer, within a ∼20 Å deep solvent-filled tunnel that connects the two active sites. The tunnel is largely lined by 10 conserved amino acid residues from all four subunits, in particular glutamate (Glu) and aspartate (Asp), six and four, respectively, plus other acidic residues around the TPP aminopyrimidine ring. And it should be underlined the absence of basic residues to neutralize them. Similar tunnels have been found in all thiamine pyrophosphate-dependent enzymes with known crystalline structure, with dimeric or tetrameric structure, for example in transketolase, an enzyme of the pentose phosphate pathway.
Note: B. stearothermophilus belongs to the phylum Firmicutes and is recently renamed Geobacillus stearothermophilus.
What is the function of the acidic tunnel?
Through mutagenesis experiments on B. stearothermophilus pyruvate dehydrogenase, the tunnel has been shown to play a role in the catalytic mechanism.
The change of some of the aforementioned acidic residues to neutral amino acids does not alter, compared with the wild-type pyruvate dehydrogenase:
the efficiency of incorporation of the modified enzyme into the multienzyme complex;
the structure of active sites;
the quaternary structure of the enzyme.
However, rate of decarboxylation is reduced by over 70 percent compared to the wild-type enzyme, as well as, once the multienzyme complex is assembled with the mutant pyruvate dehydrogenase, the PDC activity, which is reduced by over 85 percent compared to the wild-type complex. But how does this occur?
Because the distance between the substituted amino acids and the active sites is ≥7 Å, that is, these amino acids are remote from the active sites of pyruvate dehydrogenase, they cannot directly influence its catalytic activity. Then, the catalytic mechanism described below was proposed.
Considering the apoenzyme, thiamine pyrophosphate binds fast and strongly to the first active site, is activated, and the active site is closed, thus protecting the zwitterionic thiazolium from the external environment.
Conversely, in the second active site TPP binds, but is not activated, and the active site remains in an open conformation.
In the first active site, pyruvate reacts with the thiazolium C-2, and thiamine pyrophosphate of the second active site, which is a general acid, donates a proton to the first site. The result is a decarboxylation in the first site and the activation of the coenzyme in the second site, which is then closed.
It should be noted that while the activation of the first thiamine pyrophosphate is the result of the binding to the active site, the activation of the second coenzyme, and therefore of the second active site, is coupled to the decarboxylation of pyruvate in the first active site. Or, from another point of view, while an active site requires a general acid, the other requires a general base.
Protons are needed for the catalytic activity, and their transport between the two active sites occurs via the acidic tunnel. They are reversibly shuttled along a chain of donor-acceptor groups provided by glutamate and aspartate residues and the entrained water, that act as a proton wire.
It seems, therefore, that unlike many other enzymes, in which the communication between the active sites occurs through conformational changes and subunit rearrangements, in pyruvate dehydrogenase and in the other TPP-dependent enzymes, the proton wire is the molecular basis of such communication.
At this point, the holoenzyme has been formed and the active sites are in a dynamic equilibrium, each exchanging between the dormant and the activated state. This seems to be the state in which the enzyme is found in vivo at the start of each catalytic cycle.
A consequence of such a mechanism is that, as the catalytic cycles occur, the two active sites are out of phase with each other, namely, when an active site requires a general acid, the other requires a general base, and vice versa.
Finally, it should be noted that this mechanism allows the switching of the loops that close the active sites so as to:
coordinate substrate uptake and product release;
explain the asymmetry existing between the two active sites.
Note: an apoenzyme is an enzyme that lacks the association of its cofactors. Conversely, an holoenzyme is an apoenzyme together with its cofactors. The apoenzyme is a catalytically inactive enzyme, whereas the holoenzyme is a catalytically active enzyme.
Structure of dihydrolipoyl transacetylase or E2
Three functionally distinct domains can be identify in the structure of dihydrolipoyl transacetylase: an N-terminal lipoyl domain, a peripheral subunit-binding domain, and a C-terminal catalytic domain or acyltransferase domain. These domains are connected by 20- to 40 amino acid residues rich in alanine and proline, hydrophobic amino acids that are interspersed with charged residues. These linkers are highly flexible and largely extended, that allows the three domains to kept away from each other.
E2 Domains
Note: flexible linkers are present in E3BP, too.
The N-terminal lipoyl domain is composed of ∼80 amino acid residues, and is so called because it binds lipoic acid. The number of these domains depend on the species, ranging from one to three. For example, there is one domain in B. stearothermophilus and in yeasts, two in Streptococcus faecalis and in mammals, and three in Azotobacter vinelandii and E. coli.
The link between the ɛ-amino group of a lysine residue and lipoic acid leads to the formation of a flexible arm, the lipoyl-lysine, which has a maximum extended length of ∼14 Å. Adding the polypeptide segment which connects the N-terminal domain to the adjacent domain, whose length is greater than 140 Å, the resulting flexible tether is able to swings the lipoyl group between the active sites of pyruvate dehydrogenase and dihydrolipoyl dehydrogenase, as well as to interact with neighboring dihydrolipoyl transacetylases of the core.
It should be noted that the number of these tethers is 3 x 24 = 72 in E. coli, whereas in mammals 2 x 60 = 120, based on the number of N-terminal domains and the units of dihydrolipoyl transacetylase.
One pyruvate dehydrogenase can therefore acetylate numerous dihydrolipoyl transacetylases, and one dihydrolipoyl dehydrogenase can reoxidize many dihydrolipoamide groups.
Moreover, it also occurs:
an interchange of the acetyl groups between the lipoyl groups of the dihydrolipoyl transacetylase core;
the exchange of both acetyl groups and disulfides between the tethered arms.
PSBD is composed of ∼35 amino acid residues arranged to form a globular structure that binds to both pyruvate dehydrogenase and dihydrolipoyl dehydrogenase, that is, it holds the multienzyme complex together.
The C-terminal catalytic domain, which, of course, contains the active site, is composed of ∼250 amino acid residues arranged to form a hollow cage-like structure containing channels large enough to allow substrates and products to diffuse in and out. For example, CoA ad lipoamide, the two substrate of dihydrolipoyl transacetylase, bind, in their extended conformation, at the opposite ends of a channel located at the interface between each pair of subunits in each trimers.
Structure of dihydrolipoyl dehydrogenase or E3
The structure of dihydrolipoyl dehydrogenase was deduced from studies of the enzyme in several microorganisms. It has a homodimeric structure, with each ∼470 amino acid residue chain folded into four domains, from the N-terminal to the C-terminal end: a FAD-binding domain, a NAD+-binding domain, a central domain, and an interface domain. All domains participate in the formation of the active site.
FAD is almost completely hidden inside the protein because, unlike thiol or NADH, it is easily oxidizable and must therefore be protected from the surrounding solution, namely, from O2. In fact, in the absence of the nicotinamide coenzyme, the phenol side chain of a tyrosine residue (Tyr), for example Tyr181 in the Gram-negative bacteria Pseudomonas putida, covers the NAD+-binding pocket so as to protect FADH2 from the contact with the surrounding solution.
Conversely, when NAD+ is located in the active site, the phenol side chain of the aforementioned tyrosine residue is interposed between the nicotinamide ring and the flavin ring.
In the active site of the enzyme’s oxidized form is also present a redox-active disulphide bridge. It forms between two cysteine residues located in a highly conserved segment of the polypeptide chain, e.g., Cys43 and Cys48 in P. putida, and is located on opposite side of the flavin ring with respect to the nicotinamide ring. The disulphide bridge links consecutive turns in a segment of a distorted α-helix, and, noteworthy, in the absence of such distortion, Cα atoms of the two cysteine residues would be too distant to allow the disulfide bridge to form.
Dihydrolipoyl dehydrogenase has therefore two electron acceptors: FAD and the redox-active disulphide bridge.
Note: the heterocyclic rings of NAD and FAD are parallel and in contact through van der Waals interactions; S48 is also in contact through van der Waals interactions with the flavin ring, on the opposite side of it from the NAD ring.
Reaction of pyruvate dehydrogenase or E1
In the reaction sequence catalyzed by components of the pyruvate dehydrogenase complex, pyruvate dehydrogenase catalyzes the first two steps, namely:
the decarboxylation of pyruvate to form CO2 and the hydroxyethyl-TPP intermediate;
the reductive acetylation of the lipoyl group of dihydrolipoyl transacetylase.
The first reaction is essentially identical to pyruvate decarboxylase reaction (EC 4.1.1.1), which carries out a non-oxidative decarboxylation in glucose fermentation to ethanol. What differs is the fate of the hydroxyethyl group bound to thiamine pyrophosphate that, in the reaction catalyzed by pyruvate dehydrogenase is transferred to the next enzyme in the sequence, dihydrolipoyl transacetylase, whereas in the reaction catalyzed by pyruvate decarboxylase is converted into acetaldehyde.
Catalytic mechanism of pyruvate dehydrogenase or E1
In thiamine pyrophosphate-dependent enzymes, the thiazolium ring is the active center, but only as dipolar carbanion or ylid, namely, as a dipolar ion, or zwitterion (German for “hybrid ion”), with positive charge on the N-3 and negative charge on C-2. Conversely, the positively charged thiazolium ring, that is, positive charged nitrogen and no charge on C-2, can be defined as “dormant” or inactive form.
The reaction begins with the nucleophilic attack by C-2 carbanion to the carbonyl carbon of pyruvate, which has the oxidation state of an aldehyde, and leads to the formation of a covalent bond between coenzyme and pyruvate.
Then, the cleavage of C-1–C-2 bond of pyruvate occurs. This leads to the release of the carboxyl group, namely, of the C-1 as CO2, while the remaining carbon atoms, C-2 and C-3, stay bound to the thiamine pyrophosphate as hydroxyethyl group. The cleavage of the C-1–C-2 bond, and therefore the decarboxylation of pyruvate, is favored by the fact that the negative charge on the C-2 carbon, that is unstable, is stabilized by the presence in the thiazolium ring of the positively charged N-3, a imine nitrogen (C=N+), that is, due to the presence of an electrophilic or electron deficient structure that acts as an electron sink or electron trap, in which the carbanion electrons can be delocalized by resonance.
At this point, the intermediate stabilized by resonance can be protonated to form hydroxyethyl-TPP.
Note: this first reaction catalyzed by pyruvate dehydrogenase is that in which the private dehydrogenase complex exercises its substratum specificity; furthermore, it is the slowest of the five reactions, hence limiting the rate of the overall reaction.
Catalytic mechanism of pyruvate dehydrogenase
Pyruvate dehydrogenase then catalyzes the oxidation of the hydroxyethyl group to an acetyl group, and its transfer on lipoyllysyl arm of dihydrolipoyl transacetylase. The reaction begins with the formation of a carbanion on the hydroxylic carbon of the hydroxyethyl-TPP, by the removal of the carbon-linked proton by an enzyme base.
The carbanions carry out a nucleophilic attack on the lipoamide disulfide, with the formation of a high-energy acetyl-thioester bond with one of the two -SH groups. In this reaction, the oxidation of the hydroxyethyl group to an acetyl group occurs with the concomitant reduction of the lipoamide disulfide bond: the two electrons removed from the hydroxyethyl group are used to reduce the disulfide. This reaction is therefore a reductive acetylation accompanied by the regeneration of the active form of pyruvate dehydrogenase, namely, the enzyme with the thiazolium C-2 in the deprotonated form, the ylid or dipolar carbanion form.
Note that the energy derived from the oxidation of the hydroxyethyl group to an acetyl group drives the formation of the thioester bond between the acetyl group and coenzyme A.
Note: as previously said, the lipoyllysyl arm, arranged in an extended conformation in the channel where TPP is also found, allows the transfer of hydroxyethyl from hydroxyethyl-TPP to CoA, that is, it can move from the active site of pyruvate dehydrogenase to the active sites of dihydrolipoyl transacetylase, and then of dihydrolipoyl dehydrogenase.
A deeper look on thiamine pyrophosphate
Thiamine pyrophosphate molecule consists of three chemical moieties, from which its chemistry and enzymology depend: a thiazolium ring, a 4-aminopyrimidine ring, and the diphosphate side chain.
The diphosphate side chain binds the cofactor to the enzyme via the formation of electrostatic bonds between the negative charges carried by its phosphoryl groups and the positive charges carried by Ca2+ and Mg2+ ions, in turn, bound to highly conserved sequences, GlyAspGly (GDG) and GlyAspGly-X26-AsnAsn (GDG-X26-NN), respectively.
The thiazolium ring plays a central role in catalysis, due to its ability to form the C-2 carbanion, that is, a nucleophilic center on the C-2 atom.
Note: as mentioned previously in this article, once bound to the enzyme, thiamine pyrophosphate locates in the active site so that the thiazolium ring is positioned close to the channel entrance leading to the active site.
The aminopyrimidine ring has a dual function:
it anchors the coenzyme holding it in place;
it has a specific catalytic role, participating in acid/base catalysis, as evidenced by studies with thiamine pyrophosphate analogs in which each of the three nitrogen atoms of the ring were replaced in turn. These studies demonstrated that the N-1’ atom and the N-4’-amino group are required, whereas the other nitrogen atom of the ring, the N-3’ atom, is required to a lesser extent.
How is the dipolar carbanion of thiamine pyrophosphate formed?
Three tautomeric forms of the aminopyrimidine ring can be identified in the enzyme-bound coenzyme not involved in the reaction:
the canonical 4’-aminopyrimidine tautomer;
the N-1 protonated form, that is, 4-aminopyrimidinium ion;
the 1’,4’-iminopyrimidine tautomer.
It seems that the 1′,4′-imino tautomer is the tautomer that undergoes deprotonation, before the entry of the substrate into the active site. The C-2 of the thiazolium ring is “much more acidic than most =C-H groups found in other molecules”. The higher acidity, i.e., the fact that the C-2 proton is easily dissociable, is due to the presence of the quaternary nitrogen on the thiazolium ring, a positively charged nitrogen atom able to electrostatically stabilize the resulting carbanion. In the deprotonation reaction, the amino group of the aminopyrimidine ring seems to play an essential role: it acts as a base and is suitably positioned to accept the proton. However, in the 4’-aminopyrimidine tautomer one of its protons sterically collides with the C-2 proton; in addition, its pK is too low to carry out the deprotonation efficiently. A mechanism was therefore proposed, in which the side chain of a conserved glutamate residue, for example βGlu59 in B. stearothermophilus, or Glu51 in Saccharomyces uvarum (brewer’s yeast) pyruvate decarboxylase, donates a proton to the aminopyrimidine, converting it to its 1′,4′-iminopyrimidine tautomer that, accepting the C-2 proton, returns to the canonical 4′-aminopyrimidinic form and allows the formation of the carbanion.
Note: carbanion formation on C-2 is a consequence of an intramolecular proton transfer.
Deprotonation of thiamine pyrophosphate and closure of pyruvate dehydrogenase active site
The loss of the C-2 proton of the thiazolium ring leads, from a positively charged ring, to a dipolar ion, or zwitterion. This change in state of charge triggers a conformational change in one of the two conserved loops at the entrance of the active site channel, specifically, the inner of these loops, that, in turn, leads to the closure of the channel to the surrounding water environment. In this closed conformation the thiazolium carbanion is protected against electrophiles.
To sum up: the deprotonation of thiamine pyrophosphate leads to the closure of the active site and the protection of the newly formed dipolar carbanion, that is, TPP-dependent enzymes would be only active in closed conformation.
Conversely, in the other active site, thiamine pyrophosphate is not in the ylid form, the channel is open, and the site is inactive.
Reaction of dihydrolipoyl transacetylase or E2
In the reaction sequence catalyzed by components of the pyruvate dehydrogenase complex, dihydrolipoyl transacetylase catalyzes the third step, namely, the transfer of the acetyl group from acetyl-dihydrolipoamide to CoA to form acetyl-CoA and dihydrolipoamide, the fully reduced form of lipoamide, the dithiol.
It should be noted that the acetyl group, initially bound by ester linkage to one of the –SH group of lipoamide is next bound to the –SH group of coenzyme A, again by ester bond, hence the term transesterification.
Catalytic mechanism of dihydrolipoyl transacetylase or E2
During the reaction, the sulfhydryl group of coenzyme A carries out a nucleophilic attack on the carbonyl carbon of the acetyl group of acetyl dihydrolipoamide-dihydrolipoyl transacetylase to form a transient tetrahedral intermediate, that “decomposes” to dihydrolipoamide-dihydrolipoyl transacetylase and acetyl-CoA.
E2: Catalytic Mechanism
As previously said, the mobility of the lipoyllysyl arm plays a central role in the reaction mechanism.
Reaction of dihydrolipoyl dehydrogenase or E3
In the reaction sequence catalyzed by components of the pyruvate dehydrogenase complex, dihydrolipoyl dehydrogenase catalyzes the fourth and fifth steps.
The enzyme catalyzes electron transfers needed to regenerate the disulfide bridge of the lipoyl group of dihydrolipoyl transacetylase, that is, to regenerate the oxidized form of the prosthetic group, and thus completing the catalytic cycle of the transacetylase.
The reaction has a ping-pong catalytic mechanism: it occurs in two successive half-reaction, in which each of the two substrates, NAD+ and dihydrolipoamide, reacts in the absence of the other. Moreover during the first half-reaction, the release of the first product and the formation of an enzyme intermediate complex occur before the second substrate binds, while the enzyme underogoes a structural change, whereas in the second half-reaction the release of the second product and the return of the enzyme to its starting state, again, via a structural change, occur.
Considering the ping-pong kinetic mechanism of dihydrolipoyl dehydrogenase:
in the first half-reaction the oxidation of dihydrolipoamide to lipoamide occurs;
in the second half-reaction the reduction of NAD+ to NADH occurs.
Catalytic mechanism of dihydrolipoyl dehydrogenase or E3
Below, the reaction mechanism of P. putida dihydrolipoyl dehydrogenase is described.
In the first half-reaction, the oxidized dihydrolipoyl dehydrogenase (E), i.e., the enzyme with the disulfide bridge between Cys43 and Cys48, binds dihydrolipoamide (LH2) to form the enzyme-dihydrolipoamide complex (E●LH2). At this point, a sulfur atom of dihydrolipoamide carries out a nucleophilic attack on the sulfur of Cys43, to form the disulfide bridge lipoamide-Cys43 (E-S-S-L), while the sulfur of Cys48 is released as a thiolate ion (S–48).
The proton on the second thiol group of lipoamide is then abstracted by histidine (Hys) 451, that acts as a general acid-base catalyst, leading to the formation of a second thiolate ion, this time on the lipoamide (E-S-S-L ●S–), that, through a nucleophilic attack, displaces the sulfur of Cys43, S43, aided in this by general acid catalysis by Hys451 which donates a proton to S43. The catalytic action of Hys451 is essential, as demonstrated by mutagenesis studies in which its substitution with a glutamine residue causes the enzyme to retain ∼ 0.4% of the wild-type catalytic activity.
Then, thiolate anion S–48 contacts, through non-covalent interactions, the flavin ring near 4a position, i.e., an electron pair of S–48, which acts as electron donor, is partially transferred to the oxidized flavin ring, which, in turn, is the electron acceptor. The resulting structure is called charge-transfer complex.
Meanwhile, the phenolic side chain of the Tyr181 continues to hinder access to the flavin ring, thus protecting it from oxidation by O2.
Dihydrolipoamide Oxidation via E3
To sum up, what occurs is an interchange reaction of disulfide bridges leading to the formation of the oxidized form of lipoamide, the first product, which is released, and the reduced form of the dihydrolipoyl dehydrogenase.
The second half-reaction involves the reduction of NAD+ to NADH + H+ by electron transfer from the reactive disulfide of the enzyme via FAD.
Oxidation of Reduced E3
It begins with the entry of NAD+ into the active site and its binding to form the EH2●NAD+ complex. It should be noted that the entry of the coenzyme causes the phenolic side chain of the Tyr181 to be pushed aside by the nicotinamide ring.
Following the collapse of the charge-transfer complex, a covalent bond is formed between the flavin atom C-4a and S48, to which the extraction of a proton from S43 by the flavin atom N-5 is accompanied, with the formation of the corresponding thiolate anion, S–43.
S–43carries out a nucleophilic attack on S48, leading to the formation of the redox-active disulphide bridge between Cys43 and Cys48, followed by the breakdown of the covalent bond between S48 and the flavin atom C-4a to form reduced FADH anion, FADH–, with negative charge on atom N-1. It should be noted that dihydrolipoyl dehydrogenase is in the oxidized form (E).
FADH– has a transient existence because the proton bound to its N-5 is instantly transferred, as hydride ion, to the nicotinamide atom C-4, that is juxtaposed to flavin atom N-5. This leads to the formation of FAD and of the second product of the reaction, NADH, which is released.
To sum up, what occurs is that the electrons removed from the hydroxyethyl group, which derives from pyruvate, pass, via FAD, to NAD+. The catalytic cycle of dihydrolipoyl dehydrogenase is therefore completed, being the enzyme and its coenzymes in their oxidized form. At this point, the catalytic cycle of the entire pyruvate dehydrogenase complex is completed, too, and the complex is ready for a new reaction cycle.
Note: unlike the thiazolium ring of thiamine pyrophosphate, FAD does not acts as electron trap or electron sink, but rather as an electron conduit between the redox-active disulphide, in its reduced form, and NAD+.
Note: the catalytic mechanism of dihydrolipoyl dehydrogenase has been determined in analogy with that of glutathione reductase (EC 1.8.1.7), at 33% identical and whose structure is more extensively characterized. It should, however, be noted that although the two enzymes catalyze similar reactions, these usually occur in opposite direction:
dihydrolipoyl dehydrogenase uses NAD+ to oxidize two –SH groups to a disulfide (–S–S–);
glutathione reductase uses NADPH to reduce a –S–S– to two thiol groups.
Nevertheless, their active sites are closely superimposable.
Regulation of the pyruvate dehydrogenase complex
In mammals, the regulation of the activity of the pyruvate dehydrogenase complex is essential, both in the fed and fasted states. In fact, the multienzyme complex plays a central role in metabolism because, catalyzing the irreversible oxidative decarboxylation of pyruvate, represents the entry point of the carbon flux from all carbohydrate sources as well as from ∼50% of carbon skeletons of glucogenic amino acids, that, as a whole, correspond to ∼60 percent of the daily calorie intake, into:
the citric acid cycle, and therefore to the full oxidation to CO2;
the synthesis of lipids (fed state) and acetylcholine.
The importance of the regulation of the conversion of pyruvate into acetyl-CoA is also underlined by the fact that mammals, although able to produce glucose from pyruvate, cannot synthesize it from acetyl-CoA, because of the irreversibility of pyruvate dehydrogenase reaction and the absence of alternative pathways. Then, the inhibition of the activity of the complex allows to spare glucose and the amino acids that can be converted into pyruvate, such as alanine, when other fuels, for example acetyl-CoA from fatty acid oxidation, are available.
This explains why the activity of the complex is carefully regulated by:
feedback inhibition;
nucleotides;
covalent modifications, namely, phosphorylation and dephosphorylation of specific target proteins.
Regulation by feedback inhibition and energy status of the cell
The activity of the dephosphorylated form of the pyruvate dehydrogenase complex is regulated by feedback inhibition.
Acetyl-CoA and NADH allosterically inhibit the enzymes that catalyze their formation, dihydrolipoyl transacetylase and dihydrolipoyl dehydrogenase, respectively.
In addition, CoA and acetyl-CoA, as well as NAD+ and NADH, compete for binding sites on E2 and E3, respectively, that catalyze reversible reactions. This means that, in the presence of high ratios of [Acetyl-CoA]/[CoA] and [NADH]/ [NAD+], the reactions of transacetylation and dehydrogenation work in reverse; therefore, dihydrolipoyl transacetylase cannot accept the hydroxyethyl group from TPP because it is maintained in the acetylated form. This cause thiamine pyrophosphate to remain bound to pyruvate dehydrogenase in its hydroxyethyl form, which, in turn, decreases the rate of pyruvate decarboxylation. Hence, high ratios of [Acetyl-CoA]/[CoA] and [NADH]/[NAD+] indirectly influence pyruvate dehydrogenase activity.
PDC Activiy: Regulation by Feedback Inhibition
Acetyl-CoA and NADH are also produced by fatty acid oxidation, which takes place, like the reactions of the pyruvate dehydrogenase complex, within the mitochondrion. This means that the cell, by regulating the activity of the multienzyme complex, preserves carbohydrate stores when fatty acids are available for energy. For example, during the fasted state, liver, skeletal muscle and many other organs and tissues rely primarily on fatty acid oxidation for energy. Conversely, the activity of the multienzyme complex is increased in the fed state, when many different types of cells and tissues mainly use glucose as a fuel.
More generally, when the production of NADH and/or acetil-CoA exceeds the capacity of the cell to use them for ATP production, the activity of the pyruvate dehydrogenase complex is inhibited. The same is true when there is no need for additional ATP to be produced. Infact, the activity of multienzyme complex is also sensitive to the energy charge of the cell. Through allosteric mechanisms, high ATP levels inhibit the activity of the pyruvate dehydrogenase component of the complex, whereas high ADP levels, that signs that the energy charge of the cell may become low, activate it, thus committing the carbon skeleton of carbohydrates and some amino acids to energy production.
Note: in the skeletal muscle, the activity of the pyruvate dehydrogenase complex increases with increased aerobic activity, resulting in a in greater dependence on glucose as a fuel source.
Regulation by phosphorylation/dephosphorylation
Unlike prokaryotes, in mammals the activity of the pyruvate dehydrogenase complex is also regulated by covalent modifications, i.e., phosphorylation and dephosphorylation of three specific serine residues of the α subunit of pyruvate dehydrogenase, the enzyme that catalyzes the first, irreversible step of the overall reaction sequence.
Note: as mammalian pyruvate dehydrogenase is an heterotetramer, there are six potential phosphorylation sites.
PDC Activity: Regulation by Covalent Modifications
Phosphorylation, which inactivates pyruvate dehydrogenase, and then blocks the overall reaction sequence, is catalyzed by pyruvate dehydrogenase kinase. Two of the aforementioned serine residues are located on the more C-terminal loop, at the entrance of the substrate channel leading to the respective active site, and the phosphorylation of only one of them inactivates the pyruvate dehydrogenase, hence demonstrating the out of phase coupling between its active sites.
Conversely, in the dephosphorylated state the complex is active. Dephosphorylation is catalyzed by a specific protein phosphatase, the pyruvate dehydrogenase phosphatase.
The activities of pyruvate dehydrogenase kinase and pyruvate dehydrogenase phosphatase are in turn subject to allosteric regulation by several modulators.
Regulation of pyruvate dehydrogenase kinase
The activity of pyruvate dehydrogenase kinase depends on the ratios of [NADH]/[NAD+], [acetyl-CoA]/[CoA], and [ATP]/[ADP] , as well as on the pyruvate concentration, in the mitochondrial matrix.
High ratios of [NADH]/[NAD+] and [acetyl-CoA]/[CoA], as during the oxidation of fatty acids and ketone bodies, activate the kinase, pyruvate dehydrogenase is phosphorylated, and the pyruvate dehydrogenase complex is inhibited. This allows tissues, such as cardiac muscle, to preserve glucose when fatty acids and/or ketone bodies are utilized for energy, because acetyl-CoA synthesis from pyruvate, and hence from carbohydrates (and some amino acids) is turned off.
Conversely, when the concentrations of NAD+ and coenzyme A are high the activity of the kinase is inhibited and the multienzyme complex is active.
Therefore, acetyl-CoA and NADH, two of the three end products of the reactions catalyzed by the pyruvate dehydrogenase complex, allosterically control their synthesis by regulating directly and indirectly, by regulating the activity of pyruvate dehydrogenase kinase, the activity of the complex.
A high ratio of [ATP]/[ADP] activates the kinase, and then inhibits the pyruvate dehydrogenase complex.
Note: unlike many other kinases, such as those involved in the control of glycogen metabolism, pyruvate dehydrogenase kinase is not regulated by cAMP levels, but by molecules that signal changes in energy status of the cell and in the availability of biosynthetic intermediates: ATP and NADH, and acetyl-CoA, respectively.
Pyruvate allosterically inhibits pyruvate dehydrogenase kinase.
When its levels are high, it binds to kinase and inactivates it, pyruvate dehydrogenase is not phosphorylated, and the pyruvate dehydrogenase complex remains active.
Pyruvate dehydrogenase kinase is also activated by interaction with dihydrolipoyl transacetylase in its acetylated form, i.e. when acetyl-dihydrolipoamide is present.
Other activators of the kinase is potassium and magnesium ions.
Regulation of pyruvate dehydrogenase phosphatase
The activity of pyruvate dehydrogenase phosphatase depends on the ratios of [NADH]/[NAD+] and [acetyl-CoA]/[CoA], as well as on [Ca2+], in the mitochondrial matrix.
Low ratios of [NADH]/[NAD+] and [acetyl-CoA]/[CoA] activate the phosphatase, pyruvate dehydrogenase is dephosphorylated, and the pyruvate dehydrogenase complex is activated.
Conversely, when the aforesaid ratios are high, phosphatase activity is reduced, kinase activity is increased, and the multienzyme complex is inhibited.
Calcium ion activates pyruvate dehydrogenase phosphatase.
Ca2+ is an important second messenger that signals the cell requires more energy. Therefore, when its levels are high, as in cardiac muscle cells after epinephrine stimulation or in skeletal muscle cells during the muscular contraction, the phosphatase is active, the complex is dephosphorylated, and then active.
Insulin, too, is involved in the control of the activity of the pyruvate dehydrogenase complex through the activation of pyruvate dehydrogenase phosphatase. The hormone, in response to increases in blood glucose, stimulates glycogen synthesis and the synthesis of acetyl-CoA, a precursor in the synthesis of lipids.
Fasting and subsequent refeeding, too, affect the activity of the multienzyme complex.
In tissues such as skeletal muscle, cardiac muscle or kidney, fasting significantly decreases the activity of the complex, whereas refeeding reverses the inhibition of fasting.
In the brain, however, these variations are not observed because the activity of pyruvate dehydrogenase complex is essential for ATP production.
References
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Gray L.R., Tompkins S.C., Taylor E.R. Regulation of pyruvate metabolism and human disease. Cell Mol Life Sci 2014;71(14):2577-2604. doi:10.1007/s00018-013-1539-2
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Zhou Z.H., McCarthy D.B., O’Connor C.M., Reed L.J., and J.K. Stoops. The remarkable structural and functional organization of the eukaryotic pyruvate dehydrogenase complexes. Proc Natl Acad Sci USA 2001;98(26):14802-14807. doi:10.1073/pnas.011597698
The pentose phosphate pathway, also called the phosphogluconate pathway, is a metabolic pathway, common to all living organisms, for the oxidation of glucose alternative to glycolysis, from which it branches downstream of glucose 6-phosphate synthesis, and whose main function is the production, in variable ratios, of NADPH, a reduced coenzyme, and ribose 5-phosphate, a five-carbon phosphorylated sugar, namely, a pentose phosphate, hence the name pentose phosphate pathway.[14]
The phosphogluconate pathway, branching from glycolysis, is also called the hexose monophosphate shunt.
Conceptually, this pathway can be viewed as consisting of two phases: the oxidative phase, in which NADPH is produced, and the non-oxidative phase, which produces ribose 5-phosphate and other phosphorylated carbohydrates.[16]
Summary of the Reactions of Pentose Phosphate Pathway
It has been estimated that more than 10 percent of glucose is shuttled through this metabolic pathway that, noteworthy, although it oxidizes the monosaccharide, does not involve any direct production or consumption of ATP.[7][13]
The first evidence of the existence of the pentose phosphate pathway was obtained in the 1930s by the studies of Otto Warburg, Nobel Prize in Physiology or Medicine in 1931, who discovered NADP during studies on the oxidation of glucose 6-phosphate to 6-phosphogluconate.[22]
Further indications came from the observation that glucose continued to be metabolized in tissues even in the presence of glycolysis inhibitors, such as fluoride and iodoacetate ions, inhibitors of enolase (EC 4.2.1.11) and glyceraldehyde 3-phosphate dehydrogenase (EC 1.2.1.12), respectively.
However, the pathway was fully elucidated only in 1950s thanks to the work of several researchers and primarily of Efraim Racker, Fritz Lipmann, Nobel Prize in Physiology or Medicine in 1953 thanks to the discovery of coenzyme A, Bernard Horecker and Frank Dickens.[8]
Main function
The main function of the pentose phosphate pathway is the production of NADPH and ribose 5-phosphate.[22]
NADPH is needed for reductive biosynthesis, such as the synthesis of fatty acids, cholesterol, steroid hormones and of two non-essential amino acids, proline and tyrosine, from glutamate and phenylalanine, respectively, as well as for the reduction of oxidized glutathione.[14] In such reactions the reduced coenzyme acts as an electron donor, or rather as a donor of a hydride ion (:H–), namely, a proton and two electrons.
Reduced and Oxidized Form of Nicotinamide Adenine Dinucleotide Phosphate
Note: In vertebrates, about half of the NADPH necessary for the reductive steps of fatty acid synthesis derives from the pentose phosphate pathway, and the rest from the malic enzyme (EC 1.1.1.40) reaction.[13]
Malate + NADP+ ↔ Pyruvate + NADPH + H+ + HCO3–
Ribose 5-phosphate is used for the synthesis of nucleotides and nucleic acids, DNA and RNA, of ATP, coenzymes such as coenzyme A, NAD, NADP and FAD, and of the essential amino acids tryptophan and histidine.[5] This five-carbon phosphorylated is not used as such; it is activated to 5-phosphoribosyl 1-pyrophosphate (PRPP), in the reaction catalyzed by ribose phosphate pyrophosphokinase or PRPP synthase (EC 2.7.6.1).[13]
Ribose 5-phosphate + ATP → 5-Phosphoribosyl 1-pyrophosphate + AMP
Additional functions
In addition to the production of NADPH and ribose 5-phosphate, the pentose phosphate pathway has other functions, both anabolic and catabolic.[18]
In yeasts and many bacteria it is involved in the catabolism of the five carbon sugars ribose, xylose and arabinose.
In humans too, it is involved in catabolism of the aforementioned pentoses and of the less common sugars with three, four and seven carbon atoms derived from diet, as well as of:
pentoses derived from the catabolism of structural carbohydrates;
ribose 5-phosphate derived from nucleotide catabolism.
In photosynthetic organisms it contributes to carbon dioxide (CO2) fixation during the Calvin cycle.[18]
In addition to ribose 5-phosphate, it also provides other intermediates for various biosynthetic processes, such as:
erythrose 4-phosphate, used for the synthesis of phenylalanine, tryptophan, and tyrosine, the three aromatic amino acids;
ribulose 5-phosphate, used for riboflavin synthesis;
sedoheptulose 7-phosphate which, in Gram-negative bacteria, is used for the synthesis of heptose units in the lipopolysaccharide layer of the outer membrane.[19]
Where does the pentose phosphate pathway occur?
In animal cells and bacteria, the hexose monophosphate shunt, as well as glycolysis, fatty acid synthesis, and most of the reactions of gluconeogenesis, occurs in the cytosol.[5] And, considering glycolysis, gluconeogenesis and the pentose phosphate pathway we can state that these three metabolic pathways are interconnected through several shared enzymes and/or intermediates.
In plant cells the pentose phosphate pathway occurs in plastids, and its intermediates can reach the cytosol through membrane pores of these organelles.[13]
In humans, the level of expression of the enzymes of the pathway varies widely from tissue to tissue. Relatively high levels are found in the liver, adrenal cortex, testicles and ovaries, thyroid, mammary glands during lactation, and in red blood cells.[14] In all these sites, constant supply of NADPH is required to support reductive biosynthesis and/or to counteract the effects of reactive oxygen species (ROS) on sensitive cellular structures, such as DNA, membrane lipids, and proteins by the reduction of oxidized glutathione (GSSG) to reduced glutathione (GSH), which is the major intracellular antioxidant in erythrocytes, as in most other cells, in the reaction catalyzed by glutathione reductase (EC 1.8.1.7).[1]
GSSG + NADPH + H+ → 2 GSH + NADP+
High levels of the of the phosphogluconate pathway enzymes are also present in rapidly dividing cells such as enterocytes, skin cells, bone marrow cells, those of the early embryo and, in pathological conditions, cancer cells. Indeed, these cell types require a constant supply of ribose 5-phosphate for nucleic acid synthesis.[16]
Conversely, these enzymes are present in very low levels in skeletal muscle, in which the pentose phosphate pathway is virtually absent and glucose 6-phosphate is primarily used for energy production via glycolysis and the citric acid cycle.[5]
Oxidative phase
The oxidative phase of the pentose phosphate pathway consists of two irreversible oxidations, the first and third reactions, and a hydrolysis.
In this phase, glucose 6-phosphate is converted to ribulose 5-phosphate, a five-carbon phosphorylated sugar and the starting substrate for the reactions of the non-oxidative phase, with the concomitant formation of two molecules of NADPH and the release of C-1 of glucose as CO2.[7] The overall equation is:
Oxidation of glucose 6-phosphate to 6-phosphoglucono-delta-lactone
In the first step of the oxidative phase, glucose 6-phosphate dehydrogenase catalyzes the oxidation of glucose 6-phosphate to 6-phosphoglucono-δ-lactone, an intramolecular ester, via the transfer of a hydride ion from carbon 1 of glucose 6-phosphate to NADP+, that acts as oxidizing agent.
Note: This reaction yields the first molecule of NADPH of the pentose phosphate pathway.
The reaction catalyzed by glucose 6-phosphate dehydrogenase or G6PD (EC 1.1.1.49) is unique to the pathway.[1] And, similarly to what happens in most metabolic pathways, also in this case the first reaction unique to the pathway, generally known as a committed step, is an essentially irreversible step, with a ΔG in the liver of -17.6 kJ/mol (-4.21 kcal/mol), and is highly allosterically regulated.[22] And the enzyme is indeed the major control point for the flow of metabolites through the pathway.
In humans, the highest levels of G6PD are found in neutrophils and macrophages, phagocytic cells in which, during inflammation, NADPH is used for to produce superoxide radicals (O2-.) from molecular oxygen in the reaction catalyzed by NADPH oxidase (EC 1.6.3.1).[2][16]
2 O2 + NADPH → 2 O2-. + NADP+ + H+
In turn, superoxide radicals can be used for the synthesis for defensive purposes, namely, to kill phagocytized microorganisms, of other ROS but also of reactive nitrogen species (RNS), such as:
hydrogen peroxide (H2O2), in the reaction catalyzed by superoxide dismutase or SOD (EC 1.15.1.1)
2 O2-. + 2 H+ → H2O2 + O2
peroxynitrite (O=N–O–O), in the reaction with nitric oxide (•NO)
O2–. + •NO → O=N–O–O–
hydroperoxide radical (HOO•)
O2-. + H+ → HOO•
Catalytic mechanism of glucose 6-phosphate dehydrogenase
The catalytic mechanism of the enzyme has been studied in great detail in the microorganism Leuconostoc mesenteroides, whose glucose 6-phosphate dehydrogenase has the peculiar characteristic of being able to use NAD+ and/or NADP+ as coenzyme.[12]
Catalytic Mechanism of Glucose 6-Phosphate Dehydrogenas
The enzyme does not require metal ions for its activity; one of the amino acids in the active site acts as a general base being able to abstract a hydride ion from the hydroxyl group bound to C1 of glucose 6-phosphate.[4]
In the bacterial enzyme this is carried out by the atom Nɛ2 of the imidazole ring of a histidine side chain. This nitrogen atom has a lone pair of electrons able to make a nucleophilic attack. This causes glucose 6-phosphate, a cyclic hemiacetal with carbon 1 in the aldehyde oxidation state, to be oxidized to a cyclic ester, namely, a lactone, and the transfer of an hydride ion from C1 of glucose to C4 of the nicotinamide ring of NADP+ to form NADPH.
Because such histidine is conserved in many of the glucose 6-phosphate dehydrogenases sequenced, it is likely that this catalytic mechanism can be generalized to all glucose 6-phosphate dehydrogenases.
Regulation of glucose 6-phosphate dehydrogenase
Glucose 6-phosphate dehydrogenase is the major control point of carbon flow through the pentose phosphate pathway, and then the major control point for the rate of NADPH synthesis.[2]
In humans, the enzyme exists in two forms: the inactive monomeric form, and the active form that exists in a dimer-tetramer equilibrium.[1]
One of the main modulators of its activity is the cytosolic NADP+/NADPH ratio.[15] High levels of NADPH inhibit enzyme activity, because NADPH is a potent competitive inhibitor of G6PD, whereas NADP+ is required for the catalytic activity and for the maintenance of the active conformation. In fact, the binding of the oxidized coenzyme to a specific site close to the dimer interface, but distant from the active site, is required to maintain its dimeric conformation.
Regulation of G6PD Activity
Under most metabolic conditions the NADP+/NADPH ratio is low, less NADP+ is available to bind to the enzyme, and hence enzyme activity is reduced, regardless of gene expression levels. Under these conditions the oxidative phase is virtually inactive.
Conversely, in cells in which metabolic pathways and/or reactions using NADPH are particularly active, the reduction of cytosolic NADPH concentration, and hence the increase in NADP+ concentration occurs. This leads to an increase in glucose 6-phosphate dehydrogenase activity, and to the activation of the oxidative phase of the hexose monophosphate pathway.
Therefore it is possible to state that the fate of glucose 6-phosphate also depends on the current needs for NADPH.[15]
A second mechanism for the regulation of G6PD activity calls into question the accumulation of acyl-CoAs, intermediates in fatty acid synthesis.[5] These molecules, by binding to the dimeric form of the enzyme, lead to dissociation into the constitutive monomers, and then to the loss of the catalytic activity.
Insulin up-regulates the expression of the genes for glucose 6-phosphate dehydrogenase and 6-phosphogluconate dehydrogenase. Therefore, in the well-fed state, the hormone increases carbon flow through the pentose phosphate pathway and then the production of NADPH.[7]
Note: Insulin also promotes the synthesis of fatty acids.
Hydrolysis of 6-phosphoglucono-delta-lactone to 6-phosphogluconate
In the second step of the oxidative phase 6-phosphoglucono-δ-lactone is hydrolyzed to 6-phosphogluconate, a linear molecule.
6-Phosphoglucono-δ-lactone is hydrolytically unstable and undergoes a nonenzymatic ring-opening, a reaction that occurs at a significant rate.[22] However, in the cell this ring-opening reaction, an hydrolysis, is accelerated by the catalytic action of 6-phosphogluconolactonase (EC 3.1.1.31).[3]
Oxidative decarboxylation of 6-phosphogluconate to ribulose 5-phosphate
In the last step of the oxidative phase, 6-phosphogluconate undergoes an oxidative decarboxylation to form ribulose 5-phosphate, a keto pentose, CO2, and a molecule of NADPH. The reaction is catalyzed by 6-phosphogluconate dehydrogenase (EC 1.1.1.44), enzyme that requires the presence of magnesium ions, Mg2+.
6-Phosphogluconate + NADP+ → Ribulose 5-phosphate + NADPH + CO2
Note: This reaction yields the second molecule of NADPH of the pentose phosphate pathway.[15]
Catalytic mechanism of 6-phosphogluconate dehydrogenase
The catalytic mechanism of the enzyme is similar to that of isocitrate dehydrogenase (EC 1.1.1.41), an enzyme of the citric acid cycle.[22] It consists of an acid-base catalysis proceeding through a three step mechanism in which two strictly conserved residues, a lysine (Lys), and a glutamate (Glu), are involved; in humans, Lys185 and the Glu192. Lysine acts as acid/base group, whereas glutamate as an acid.[6][17]
Catalytic Mechanism of 6-Phosphogluconate Dehydrogenase
In the first step, the oxidative step, 6-phosphogluconate is oxidized to a beta-keto acid, the 3-keto-6-phosphogluconate.
In this step the ε-amino group of the aforementioned lysine acts as a general base, as a nucleophile, abstracting a proton from the hydroxyl group bound to C-3. Then, the transfer of a hydride ion from C-3 to C-4 of the nicotinamide ring of the NADP+ occurs. This leads to the formation of the 3-keto intermediate and a molecule of NADPH that leaves the active site.
In the second step, the decarboxylation step, 3-keto-6-phosphogluconate, that is very susceptible to decarboxylation, is converted to the cis-1,2-enediol of ribulose 5-phosphate, a high energy intermediate.In this step the aforementioned lysine acts as a general acid donating an H+ at the C-3 carbonyl oxygen, and the C-1 of glucose 6-phosphate is lost as CO2.
Finally, 6-phosphogluconate dehydrogenase catalyzes a stereospecific keto-enol conversion leading to the formation of ribulose 5-phosphate. In this step, the aforementioned glutamic acid residue acts as a general acid donating an H+ to the C-1 of cis-1,2-enediol intermediate, while the ε-amino group of the lysine accepts a proton from the hydroxyl group bound to the C-2. The result is the formation of ribulose 5-phosphate.
Non-oxidative phase
In the non-oxidative phase of the pentose phosphate pathway, several phosphorylated carbohydrates are produced, whose fate depends on the relative needs for NADPH, ribose 5-phosphate, and ATP of the cell.[7] This phase consists of five steps, all freely reversible, in which a series of interconversions of phosphorylated sugars occurs. It begins with two reactions: the isomerization and epimerization of ribulose 5-phosphate to form ribose 5-phosphate and xylulose 5-phosphate, respectively.[14] Note: enzymatic isomerizations and epimerizations play an important role in carbohydrate metabolism.
Epimerases (EC 5.1), a subclass of Isomerases (EC 5.), catalyze the configurational reversal at an asymmetric carbon atom, usually by a deprotonation/protonation mechanism.
In isomerization reactions, the interchange of chemical groups occurs between carbon atoms.
Isomerization of ribulose 5-phosphate to ribose 5-phosphate
In the first step of the nonoxidative phase of the pentose phosphate pathway, an isomerization reaction, ribulose 5-phosphate, a ketose, is converted to the corresponding aldose, ribose 5-phosphate. The reaction is catalyzed by phosphopentose isomerase or ribose 5-phosphate isomerase (EC 5.3.1.6).
Ribulose 5-phosphate ⇄ Ribose 5-phosphate
These molecules are an example of functional group isomerism.
Catalytic mechanism of phosphopentose isomerase
The catalytic mechanism of phosphopentose isomerase is similar to that of phosphohexose isomerase (EC 5.3.1.9), a glycolytic enzyme, and leads to the formation of the high energy intermediate cis-1,2-enediol of ribulose 5-phosphate via a proton-transfer mechanism common to the aldose-ketose isomerizations.
The proposed catalytic mechanism for phosphopentose isomerase from E. coli, in the direction of ribulose 5-phosphate formation from ribose 5-phosphate, as in the Calvin cycle of photosynthesis, is described below.[23]
Catalytic Mechanism of Phosphopentose Isomerase
In the first step, the furanose ring of the substrate is opened, opening induced by the interaction with an aspartic acid residue (Asp81) that accepts a proton from the hydroxyl group bound to C-1, whereas it is likely that water is the proton donor.
Note: The opening of the furanose ring is quite rare in solution (<0.5 percent).
Once the chain is opened, a glutamic acid residue (Glu103) acts as a general base, as a nucleophile, abstracting a proton bound to the C-2, whereas Asp81 donates a proton. As a result, cis-1,2-enediol intermediate is produced.
Finally, the protonated Glu103 acts as a general acid and donates an H+ at C-1 of the cis-1,2-enediol intermediate, while Asp81 acts as a general base accepting a proton from the hydroxyl group bound to C-2. The result is the formation of ribulose 5-phosphate.
During the synthesis of ribose 5-phosphate from ribulose 5-phosphate phosphopentose isomerase works in reverse.
Epimerization of ribulose 5-phosphate to xylulose 5-phosphate
The other metabolic fate of ribulose 5-phosphate in the pentose phosphate pathway is to be epimerized to xylulose 5-phosphate, a ketose like ribulose 5-phosphate, in the reaction catalyzed by phosphopentose epimerase (EC 5.1.3.1).
Ribulose 5-Phosphate ⇄ Xylulose 5-Phosphate
Note: Xylulose 5-phosphate is a regulatory molecule that inhibits gluconeogenesis and stimulates glycolysis by controlling the levels of fructose 2,6-bisphosphate in the liver.[10]
Catalytic mechanism of phosphopentose epimerase
Also this reaction, like those catalyzed by 6-phosphogluconate dehydrogenase and ribose 5-phosphate isomerase, proceeds through the formation of an enediol intermediate, but with the double bond between C-2 and C-3 and not between C-1 and C-2.
During the reaction an amino acid residue present in the active site of the enzyme acts as a general base, as a nucleophile, and abstracts a proton bound to the C-3, leading to the formation of the cis-2,3-enediol intermediate. Then, an acidic amino acid residue donates a proton to C-3, but from the opposite side, hence, with an inversion at C-3 to form xylulose 5-phosphate.[5][9]
Catalytic Mechanism of Phosphopentose Epimerase
To this point, the hexose monophosphate shunt has generated for each molecule of glucose 6-phosphate metabolized:
a pool of three pentose 5-phosphates, namely, ribulose 5-phosphate, ribose 5-phosphate and xylulose 5-phosphate, that coexist at equilibrium;
2 molecules of NADPH.
In the following three steps, from the sixth to the eighth, transketolase (EC 2.2.1.1) and transaldolase (EC 2.2.1.2), two enzymes unique to the pentose phosphate pathway, catalyze a series of rearrangements of the carbon skeletons leading to the formation of three-, four-, six-, and seven carbon units, that can be used for various metabolic purposes, depending on the needs of the cell.[2]
Analyzing the flow of metabolites through the different metabolic pathways, the concerted action of transketolase and transaldolase allows the interaction of the pentose phosphate pathway, in particular of its non-oxidative phase, with glycolysis, and gluconeogenesis, as well as with the pathways leading to the formation of numerous vitamins, coenzymes and nucleic acid precursors.
Transketolase
Transketolase is the rate-limiting enzyme of the nonoxidative phase of the pentose phosphate pathway, and the first enzyme that acts downstream of ribose 5-phosphate isomerase and phosphopentose epimerase.[11]
Discovered independently in 1953 by Horecker and Racker, and named by Racker, it catalyzes in the sixth and eighth steps, the transfer of a two carbon unit from a ketose, the donor substrate, namely, xylulose 5-phosphate, sedoheptulose 7-phosphate or fructose 6-phosphate, to an aldose, the acceptor substrate, ribose 5-phosphate, glyceraldehyde 3-phosphate or erythrose 4-phosphate.[8]
Reactions Catalyzed by Transketolase
Taking as an example the forward reactions, in the sixth step, the ketose donor is xylulose 5-phosphate, whereas the aldose acceptor is ribose 5-phosphate, to form glyceraldehyde 3-phosphate, the remaining three-carbon fragment from xylulose 5-phosphate, and sedoheptulose 7-phosphate, a seven-carbon sugar that will be used in the next step, the seventh.
In the eighth step, the ketose donor is xylulose 5-phosphate, whereas the aldose acceptor is erythrose 4-phosphate, to form another glyceraldehyde 3-phosphate and a fructose 6-phosphate.
It should be noted that three of the four products of the reactions catalyzed by this enzyme, two molecules of glyceraldehyde 3-phosphate and one of fructose 6-phosphate, are also intermediates of glycolysis.[11] In addition to xylulose 5-phosphate, sedoheptulose 7-phosphate and fructose 6-phosphatetransketolase can use as substrates other 2-keto sugars, as well as a variety of different aldose phosphates.
Transketolase and thiamine pyrophosphate
Transketolase is an enzyme that requires thiamine pyrophosphate (TPP) as a cofactor.
Thiamine pyrophosphate is the biologically active form of thiamin or vitamin B1, and is tightly bound to the enzyme.
Thiamine Pyrophosphate
Other enzymes that require TPP as a cofactor are:
pyruvate decarboxylase (EC 4.1.1.1), that is involved in alcoholic fermentation;
alpha-keto acid dehydrogenase or E1 component (EC 1.2.4.4) of the branched-chain alpha-ketoacid dehydrogenase complex, which catalyzes the oxidative decarboxylation of branched-chain alpha-keto acids;
alpha-ketoglutarate dehydrogenase or E1 component (EC 1.2.4.2) of the alpha-ketoglutarate dehydrogenase complex, an enzyme of the citric acid cycle.
TPP is involved in the transfer of activated aldehyde intermediates by stabilizing the two-carbon carbanions formed during the reaction.
Catalytic mechanism of transketolase
The carbon atom between the sulfur and nitrogen atoms of the thiazolium ring of thiamine pyrophosphate, namely, the C-2 atom, is much more acidic than most =CH groups found in other molecules because of adjacent positively charged nitrogen atom that electrostatically stabilizes the carbanion resulting from dissociation of the proton. This causes the C-2 proton to be easily dissociable to form a carbanion, i.e. a carbon atom with a negative charge. Such proton abstraction is catalyzed by transketolase.[5][14][22]
The carbanions attack the carbonyl carbon of the substrate, in the step 6, xylulose 5-phosphate or, in the reverse reaction, sedoheptulose 7-phosphate, whereas in the step 8, xylulose 5-phosphate or, in the reverse reaction, fructose 6-phosphate.
Taking as an example the forward reaction of step 6, the covalent adduct between thiamine pyrophosphate and xylulose 5-phosphate undergoes fragmentation, via the cleavage of the C2-C3 bond of xylulose 5-phosphate, to form glyceraldehyde 3-phosphate, that is released, and a two carbon unit, a negatively charged hydroxyethyl group, that remains bound to C-2 of the thiazolium ring.
Catalytic Mechanism of Transketolase
The negative charge on the hydroxyethyl intermediate is stabilized by the thiazolium ring of thiamine pyrophosphate because of the positively charged nitrogen atom that acts as an electron trap. Therefore, thiazolium ring provides an electron deficient or electrophilic structure that can delocalize by resonance the carbanion electrons.
Then, the condensation occurs between the hydroxyethyl group and the ribose 5-phosphate, the acceptor aldehyde substrate, via carbanion attack on the aldehyde carbon of ribose 5-phosphate, to form a covalent adduct bound to thiamine pyrophosphate.
Finally, the cleavage of the adduct leads to the release of sedoheptulose 7-phosphate, and regenerates the TPP carbanion.
Transaldolase
Discovered in 1953 by Horecker and Smyrniotis in the brewer’s yeast, assigned to the species Saccharomyces cerevisiae, it catalyzes, in the seventh step of the pentose phosphate pathway, the transfer of a three carbon unit from a donor substrate, sedoheptulose 7-phosphate, to an acceptor substrate, glyceraldehyde 3-phosphate, to form fructose 6-phosphate and erythrose 4-phosphate.[18]
Note: Like in transketolase catalyzed reactions, the carbon unit donor is a ketose while the acceptor is an aldose.
In the reverse reaction, the donor substrate is fructose 6-phosphate, while the acceptor substrate is erythrose 4-phosphate.
Catalytic mechanism of transaldolase
Unlike transketolase, transaldolase does not require a cofactor for activity.
The reaction occurs in two step, an aldol cleavage and an aldol condensation. Below, the catalytic mechanism of E. coli transaldolase B is analyzed, taking as an example the forward reaction leading to erythrose 4-phosphate and fructose 6-phosphate synthesis.
In the first step an ε-amino group of a lysine residue (Lys132) in the active site, after a proton transfer to a glutamic acid residue (Glu96) mediated by a water molecule, performs a nucleophilic attack on the carbonyl carbon of sedoheptulose 7-phosphate, that is, on the C-2 atom. The result is the formation of a carbinolamine with sedoheptulose 7-phosphate.[5][14][18][22]
Catalytic Mechanism of Transaldolase
In the second step, the removal of a water molecule from carbinolamine leads to the formation of an enzyme-bound imine or Schiff base intermediate; this step, too, involves the transfer of a proton from Glu96 to the “catalytic” water molecule.
Note: This enzyme-substrate covalent intermediate is quite similar to that formed in the reaction catalyzed by aldolase (EC 4.1.2.13) in the fourth step of glycolysis.
In the next step, the carboxylic group of an aspartic acid residue (Asp17) extracts a proton from the hydroxyl group bound to C-4, leading to the cleavage of the C–C bond between C-3 and C-4. This reaction is an aldol cleavage and releases the first product, erythrose 4-phosphate, an aldose, whereas a three-carbon carbanion remains bound to the enzyme and is stabilized by resonance, like in transketolase catalyzed reactions. In fact, like the nitrogen atom in the thiazolium ring of thiamine pyrophosphate, the nitrogen atom with a positive charge of the Schiff base acts as an electron trap stabilizing the negative charge carried by the carbanion.
Once the acceptor substrate glyceraldehyde 3-phosphate is in the active site, the carbanion performs a nucleophilic attack on the carbonyl carbon of glyceraldehyde 3-phosphate to form, by aldol condensation, a new C–C bond and an enzyme-bound ketose.
Then, the hydrolysis of the Schiff base releases fructose 6-phosphate, a ketose and the second product of the reaction. At this point, a new reaction cycle can start.
Finally, as seen previously, in the eighth step of the pentose phosphate pathway, transketolase catalyzes the synthesis of fructose 6-phosphate and glyceraldehyde 3-phosphate from erythrose 4-phosphate and xylulose 5-phosphate.
Cell’s need for NADPH, ribose 5-phosphate and ATP
From the molecular point of view, the fate of glucose 6-phosphate depends, to a large extent, on the relative activities of the enzymes that metabolize it in glycolysis and in the pentose phosphate pathway, in particular phosphofructokinase 1 (PFK-1) (EC 2.7.1.11) and glucose 6-phosphate dehydrogenase, whose activities that are highly regulated.
PFK-1 is inhibited when ATP and/or citrate concentrations increase, namely, when the energy charge of the cell is high, whereas it is activated when AMP and/or fructose 2,6-bisphosphate concentrations increase, namely, when the energy charge of the cell is low. Thus, when the energy charge of the cell is high, the carbon flow, and therefore the flow of glucose 6-phosphate, through the glycolytic pathway decreases.[20][21]
Glucose 6-phosphate dehydrogenase is inhibited by NADPH and acyl-CoAs, intermediates in fatty acid biosynthesis.[15] Thus, when the cytosolic levels of NADPH increases, the flow of glucose 6-phosphate through the pentose phosphate pathway is inhibited, whereas if NADPH levels drop, the inhibition disappears, the pathway switches on again, and NADPH and ribose 5-phosphate are synthesized.
Therefore, depending on the cell’s need for ATP, NADPH and ribose 5-phosphate, some reactions of glycolysis and the pentose phosphate pathway can be combined in novel ways to emphasize the synthesis of needed metabolites, also exploiting the fact that the non-oxidative phase of the hexose monophosphate shunt is essentially controlled by the availability of the substrates.
The four principal possibilities are described below.[2]
The need for NADPH is much greater than that for ribose 5-phosphate and ATP
When much more NADPH than ribose 5-phosphate is needed, and there is no need for additional ATP to be produced, namely, the energy charge of the cell is high, glucose 6-phosphate enters the pentose phosphate pathway and is completely oxidized to CO2. Such metabolic conditions are found, for example, in the adipose tissue during fatty acid synthesis.
Through a combination of the reactions of the non-oxidative phase and of some reactions of gluconeogenesis, namely, those catalyzed by triose phosphate isomerase (EC 5.3.1.1), aldolase (EC 4.1.2.13), phosphohexose isomerase (EC 5.3.1.9), and fructose 1,6-bisphosphatase (EC 3.1.3.11), six molecules of ribulose 5-phosphate are converted into five molecules of glucose 6-phosphate. Thus, it is possible to state that the reactions of the non-oxidative phase allow the reactions of the oxidative phase to proceed.
Three groups of reactions can be identify.
In the first group there are the reactions catalyzed by the enzymes of the oxidative phase, leading to the formation of two molecules of NADPH and one molecule of ribulose 5-phosphate.
In the second group there are the reactions catalyzed by the enzymes phosphopentose epimerase, ribose 5-phosphate isomerase, transketolase and transaldolase, namely, those of the non-oxidative phase of the pathway, that lead to the conversion of ribulose 5-phosphate to fructose 6-phosphate and glyceraldehyde 3-phosphate.
Finally, fructose 6-phosphate and glyceraldehyde 3-phosphate can be recycled to glucose 6-phosphate via some reactions of gluconeogenesis, so that the cycle can begin again.
The sum of the last two reactions shows that six molecules of ribulose 5-phosphate are converted to five molecules of glucose 6-phosphate.
6 Ribulose 5-phosphate+ H2O → 5 Glucose 6-phosphate + Pi
The sum of the reactions of the first, second and third group gives the overall reaction:
Glucose 6-phosphate + 12 NADP+ + 7 H20 → 6 CO2 + 12 NADPH + 12 H+ + Pi
Therefore, one molecule of glucose 6-phosphate, via six cycles of the pentose phosphate pathway coupled with some reactions of gluconeogenesis, is converted to six molecules of CO2, with the concomitant production of 12 molecules of NADPH, and without net production of ribose-5-phosphate.
The need for NADPH and ATP is much greater than that for ribose 5-phosphate
When much more NADPH than ribose 5-phosphate is needed, and the energy charge of the cell is low, that is, there is a need for ATP, ribulose 5-phosphate formed in the oxidative phase of the pentose phosphate pathway is converted to fructose 6-phosphate and glyceraldehyde 3-phosphate through the reactions of the non-oxidative phase. These two intermediates, through the reactions of glycolysis, are oxidized to pyruvate, the conjugate base of pyruvic acid, with concomitant ATP production.
The net reaction is:
If the cell requires more ATP, the pyruvate produced can be oxidized through the citric acid cycle.
Conversely, if there is no need for additional ATP to be produced, the carbon skeleton of pyruvate can be used as a building block in several biosynthetic pathways.
Note: As in the previous case, there is no net production of ribose 5-phosphate.
The need for ribose 5-phosphate is much greater than that for NADPH
When much more ribose 5-phosphate than NADPH is needed, as in rapidly dividing cells in which there is a high rate of synthesis of nucleotides, precursors of DNA, the reactions of the oxidative phase of the pentose phosphate pathway are bypassed, and there is no synthesis of NADPH. Conversely, because the reactions of the non-oxidative phase are easily reversible, the drop in ribose 5-phosphate levels, due to its rapid use, stimulates its synthesis.
What happens is that, through the glycolytic pathway, most of the glucose 6-phosphate is converted to fructose 6-phosphate and glyceraldehyde 3-phosphate. Then, transaldolase and transketolase lead to the synthesis of ribose 5-phosphate and xylulose 5-phosphate. Xylulose 5-phosphate, through the reactions catalyzed by phosphopentose epimerase and ribose 5-phosphate isomerase, is converted to ribose 5-phosphate.
The net reaction is:
Under this metabolic conditions therefore, what happens is an interplay between reactions of glycolysis and of the non-oxidative phase of the phosphogluconate pathway, with the latter in the direction of ribose 5-phosphate synthesis.
It should be noted no metabolites return to glycolysis.
The needs for ribose 5-phosphate and NADPH are balanced
If one molecule of ribose 5-phosphate and two molecules of NADPH per molecule of glucose 6-phosphate metabolized satisfy the metabolic needs of the cell, the reactions that predominate are those of the oxidative phase and that catalyzed by ribose 5-phosphate isomerase.
The net reaction is:
Under this metabolic conditions, too, no metabolites return to glycolysis.
References
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^ abcd Berg J.M., Tymoczko J.L., and Stryer L. Biochemistry. 5th Edition. W. H. Freeman and Company, 2002
^ Brodie A.F., Lipmann F. Identification of a gluconolactonase. J Biol Chem 1955;212(2):677-85. doi:10.1016/S0021-9258(18)71006-5
^ Cosgrove M.S., Naylor C., Paludan S., Adams M.J., Richard Levy H. On the mechanism of the reaction catalyzed by glucose 6-phosphate dehydrogenase. Biochemistry 1998;37(9);2759-2767. doi:10.1021/bi972069y
^ Hanau S., Montin K., Cervellati C., Magnani M., Dallocchio F. 6-Phosphogluconate dehydrogenase mechanism: evidence for allosteric modulation by substrate. J Biol Chem 2010;285(28):21366-21371. doi:10.1074/jbc.M110.105601
^ ab Horecker B.L. The pentose phosphate pathway. J Biol Chem 2002;277(50):47965-47971. doi:10.1074/jbc.X200007200
^ Jelakovic S., Kopriva S., Süss K-H, Schulz G.E. Structure and catalytic mechanism of the cytosolic D-ribulose-5-phosphate 3-epimerase from rice. J Mol Biol 2003;326:127-135. doi:10.1016/S0022-2836(02)01374-8
^ Kabashima T., Kawaguchi T., Wadzinski B.E., Uyeda K. Xylulose 5-phosphate mediates glucose-induced lipogenesis by xylulose 5-phosphate-activated protein phosphatase in rat liver. Proc Natl Acad Sci USA 2003;100:5107-5112. doi:10.1073/pnas.0730817100
^ ab Kochetov G.A., Solovjeva O.N. Structure and functioning mechanism of transketolase. Biochim Biophys Acta 2014;1844(9):1608-1618. doi:10.1016/j.bbapap.2014.06.003
^ Levy H.R. Glucose-6-phosphate dehydrogenase from Leuconostoc mesenteroides. Biochem Soc Trans. 1989 Apr;17(2):313-5. doi:10.1042/bst0170313
^ abcd Michal G., Schomburg D. Biochemical pathways. An atlas of biochemistry and molecular biology. 2nd Edition. John Wiley J. & Sons, Inc. 2012
^ abcdef Nelson D.L., Cox M.M. Lehninger. Principles of biochemistry. 6th Edition. W.H. Freeman and Company, 2012
^ abcd Patra K.C., Hay N. The pentose phosphate pathway and cancer. Trends Biochem Sci 2014;39(8):347-354. doi:10.1016/j.tibs.2014.06.005
^ abc Rosenthal M.D., Glew R.H. Medical Biochemistry – Human Metabolism in Health and Disease. John Wiley J. & Sons, Inc., 2009
^ Ruda G.F., Campbell G., Alibu V.P., Barrett M.P., Brenk R., Gilbert I.H. Virtual fragment screening for novel inhibitors of 6-phosphogluconate dehydrogenase. Bioorg Med Chem 2010;18(14):5056-62. doi:10.1016/j.bmc.2010.05.077
^ abcd Samland A.K., Sprenger G.A. Transaldolase: from biochemistry to human disease. Int J Biochem Cell Biol 2009;41(7):1482-1494. doi:10.1016/j.biocel.2009.02.001
^ Sprenger G.A. Genetics of pentose-phosphate pathway enzymes of Escherichia coli K-12. Arch Microbiol 1995;164(5):324-30. doi:10.1007/BF02529978
^ Van Schaftingen E., Hers H-G. Inhibition of fructose-1,6-bisphosphatase by fructose-2,6-bisphosphate. Proc Natl Acad Sci USA 1981;78(5):2861-2863. doi:10.1073/pnas.78.5.2861
^ Van Schaftingen E., Jett M-F., Hue L., Hers H-G. Control of liver 6-phosphofructokinase by fructose 2,6-bisphosphate and other effectors. Proc Natl Acad Sci USA 1981;78(6):3483-3486. doi:10.1073/pnas.78.6.3483
^ abcdefg Voet D. and Voet J.D. Biochemistry. 4th Edition. John Wiley J. & Sons, Inc. 2011
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Glycolysis, from Greek word glykys, meaning “sweet”, and lysis, meaning “dissolution or breakdown”, can be defined as the sequence of enzymatic reactions that, in the cytosol, also in the absence of oxygen, leads to the conversion of one molecule of glucose, a six carbon sugar, to two molecules of pyruvate, a three carbon compound, with the concomitant production of two molecules of ATP, the universal energy currency in biological systems.[5]
The Glycolytic Pathway
Glycolysis, which evolved before a substantial amount of oxygen had accumulated in the atmosphere, is the metabolic pathway with the largest flux of carbon in most living cells, and is present in almost all organisms.
This pathway, not requiring oxygen, played a crucial role in metabolic processes during the first 2 billion years of evolution of life, and probably represents the most ancient biological mechanism for extracting energy from organic molecules when oxygen availability is low. Moreover, glycolysis is a source of precursors for aerobic catabolism and for various biosynthetic processes.
Note: glycolysis is also known as the Embden-Meyerhof pathway, named after Gustav Embden and Otto Meyerhof, the two researchers who elucidated the entire pathway in the muscle.[2][5]
The development of biochemistry has gone hand in hand with the elucidation of glucose metabolism, especially glycolysis, the first metabolic pathway to have been elucidated.[2][5]
Though the elucidation of this metabolic pathway was worked out in the ‘40 of the last century, the key discovery about glucose metabolism was made in 1897, quite by accident, following a problem arose a year earlier, when a German chemist, M. Hahn, in attempting to obtain and preserve cell-free protein extracts of yeast, encountered difficulties in its conservation. A colleague, Hans Buchner, remembering a method for preserving jams, suggested to add sucrose to the extract.
Eduard Buchner, Hans’s brother, put the idea of Hans into practice, and observed that the solution produced bubbles. This prompted Eduard to conclude that a fermentation was occurring, a quite surprising discovery. Indeed fermentation, according to Pasteur’s assertion in 1860, was inextricably tied to living cells, whereas it was now demonstrated that it could also occur outside them. Briefly, these two researchers refuted the vitalist dogma and had a pivotal role in starting modern biochemistry.
Eduard Buchner was awarded the Nobel Prize in Chemistry in 1907 for this research, and was the first of several researchers who won the award for their discoveries concerning glycolysis.
It was later demonstrated, working with muscle extracts, that many of the reactions of lactic fermentation were the same of those of alcoholic fermentation, thus revealing the underlying unity in biochemistry.
As previously mentioned, glycolysis was then fully elucidated in the first half of the last century largely due to the work of researchers such as Gerty and Carl Cori, Carl Neuberg, Arthur Harden, William Young, Jacob Parnas, Otto Warburg, Hans von Euler-Chelpin, Gustav Embden and Otto Meyerhof. In particular, Warburg and von Euler-Chelpin elucidated the whole pathway in yeast, and Embden and Meyerhof in muscle in the 30’s.
Why is it so important?
Glycolysis is essential to most living cells both from the energy point of view and as a source of precursors for many other metabolic pathways. And the rate of carbon flow through glycolysis, namely, the amount of glucose converted to pyruvate per unit time, is regulated to meet these two basic needs for the cell.[7]
From the energetic point of view, although glycolysis is a relatively inefficient pathway, it can occur in the absence of oxygen, the condition in which life evolved on Earth and that many contemporary cells, both eukaryotic and prokaryotic, experience. Here are some examples.
In most animals, muscles exhibit an activity-dependent anaerobiosis, namely, they can work anaerobically for short periods. For example, when animals, but also athletes, perform high intense exercises, their need for ATP exceeds body’s ability to supply oxygen to the muscle. In such situation, muscles function, albeit for a short period of time, anaerobically.
Another example is the cornea of the eye, a poorly vascularized tissue.
Many microorganisms live in environments where oxygen is low or absent, such as deep water, soil, but also skin pores. And a variety of microorganisms called obligate anaerobes cannot survive in the presence of oxygen, a highly reactive molecule. Examples are Clostridium perfringens, Clostridium tetani, and Clostridium botulinum, that cause gangrene, tetanus and botulism, respectively.
It should also be underlined that glycolysis also plays a key role in those cells and tissues in which glucose is the sole source of energy, such as:
red blood cells, lacking mitochondria,
sperm cells;
the brain, which can also use ketone bodies for fuel in times of low glucose;
the adrenal medulla.
A similar situation is also found in the plant world where many aquatic plants and some plant tissues specialized in starch accumulation, such as potato tubers, use glucose as the main source of energy.
Note: There are organisms that are facultative anaerobes, namely organisms that can survive in the presence and in the absence of oxygen, acting aerobically or anaerobically, respectively. Examples are animals belonging to the genus Mytilus, which display a habitat-dependent anaerobiosis, a condition similar to the activity-dependent anaerobiosis seen in muscle.
Finally, it should not be forgotten that under aerobic conditions, in cells with mitochondria, glycolysis constitutes the upper part of the metabolic pathway leading to the complete oxidation of glucose to carbon dioxide (CO2) and water for energy purposes.
Glycolysis as a Source of Building Blocks
Some glycolytic intermediates, for example glucose 6-phosphate (G-6-P), fructose 6-phosphate (F-6-P) or dihydroxyacetone phosphate (DHAP), may be used as building blocks in several metabolic pathways, such as those leading to glycogen synthesis, and the synthesys of fatty acids, triglycerides, nucleotides, some amino acids, and 2,3-bisphosphoglycerate (2,3-BPG).
Sequence of reactions
The 10 steps that make up glycolysis can be divided into two phases.
The first, called the preparatory phase, consists of 5 steps and starts with the conversion of glucose to fructose 1,6-bisphosphate (F-1,6-BP) through three enzymatic reactions, namely, a phosphorylation at C-1, an isomerization, and a second phosphorylation, this time at C-6, with consumption of 2 ATP. Fructose 1,6-bisphosphate is then cleaved into two phosphorylated three-carbon compounds, glyceraldehyde 3-phosphate and dihydroxyacetone phosphate. Finally, the isomerization of DHAP to a second molecule of glyceraldehyde-3- phosphate occurs. In the preparatory phase therefore a glucose is split into two molecules of glyceraldehyde 3-phosphate, and two ATP are consumed.[2][5]
In the second phase, called the payoff phase, consisting of the remaining 5 steps of the pathway, the two molecules of glyceraldehyde 3-phosphate are converted into two molecules of pyruvate, the conjugate base of pyruvic acid, with the concomitant production of 4 ATP. So, in this phase, part of the energy present in the chemical bonds of glucose is extracted and conserved in the form of ATP. Furthermore, reducing equivalents are extracted and conserved in the form of the reduced coenzyme NADH. The metabolic fate of NADH will depend on the cell type and aerobic or anaerobic conditions.[2][5]
Note: Glucose metabolized in the glycolytic pathway derives both from glucose that enters the cell through specific membrane transporters, that in turn derives from the bloodstream, and glucose 6-phosphate produced by glycogenolysis.
Reaction 1
In the first step of the glycolytic pathway glucose is phosphorylated to glucose 6-phosphate at the expense of one ATP.
Glucose + ATP → Glucose 6-phosphate + ADP + H+
In most cells this reaction is catalyzed by hexokinase (EC 2.7.1.1), enzyme present in the cells of all organisms, and in humans with four isozyme.
Hexokinase and pyruvate kinase, the other kinase of the glycolysis, like many other kinases, require the presence of magnesium ion, Mg2+, or of another bivalent metal ion such as manganese, Mn2+, for their activity. Mg2+ binds to the ATP to form the complex MgATP2-, and in fact the true substrate of the enzyme is not ATP but this complex. It should be emphasized that the nucleophilic attack by a hydroxyl group (-OH) of glucose at the terminal phosphorus atom of the ATP is facilitated by the action of Mg2+ that interacts with the negative charges of the phosphoryl groups of the nucleoside triphosphate.
The formation of the phosphoester bond between a phosphoryl group and the hydroxyl group at C-6 of glucose is thermodynamically unfavorable and requires energy to proceed, energy that is provided by the ATP. Indeed, while the phosphorylation of glucose at C-6 by inorganic phosphate has a ΔG°’ of 13.8 kJ/mol (3.3 kcal/mol), namely, it is an endergonic reaction, the hydrolysis of ATP to ADP and Pi has ΔG°’ of -30.5 kJ/mol (-7.3 kcal/mol), namely, it is an exergonic reaction. The net reaction has a ΔG°’ of (-30.5 + 13.8) = -16.7 kJ/mol (-7.3 + 3.3 = -4.0 kcal/mol). Under cellular conditions the reaction is even more favorable, with a ΔG equal to -33.5 kJ/mol (-8.0 kcal/mol).
Therefore, this is an essentially irreversible reaction.
Note: In biochemistry, phosphorylations are fundamental reactions catalyzed by enzymes called kinases, a subclass of transferases. Kinases catalyze the transfer of the terminal phosphoryl group, or γ-phosphoryl group, of a nucleoside triphosphate to an acceptor nucleophile to form a phosphoester bond. Specifically, hexokinase catalyzes the transfer of the γ-phosphoryl group of ATP to a variety of hexoses, that is, sugars with six carbons, such as fructose and mannose, in addition to glucose.
The importance of glucose phosphorylation
The phosphorylation of glucose has some functions.[2]
Glucose 6-phosphate, due to its negative charge and because there are no transporters for phosphorylated sugars in the plasma membrane, cannot diffuse out of the cell. Thus, after the initial phosphorylation, no further energy is needed to keep the phosphorylated molecule within the cell, despite the large difference between its intra- and extracellular concentrations.
Similar considerations are valid for each of the eight phosphorylated intermediates between glucose 6-phosphate and pyruvate.
The rapid phosphorylation of glucose maintains a low intracellular concentration of the hexose, thus favoring its facilitated diffusion into the cell.
Phosphorylation causes an increase in the energy content of the molecule, that is, it starts to destabilize it, thus facilitating its further metabolism.
Other possible fates of glucose 6-phosphate
Glucose 6-phosphate is a key metabolite of glucose metabolism. In fact, in addition to be metabolized in the glycolytic pathway, in anabolic conditions it can have other fates. Here are some examples.
It can be used in the synthesis of:
glycogen, a polysaccharide stored mainly in the liver and muscle;
complex polysaccharides present in the extracellular matrix; galactose;
glucosamine and other sugars used for protein glycosylation.
NADPH, needed for reductive biosynthesis, such as fatty acid, cholesterol, steroid hormone, and deoxyribonucleotide biosynthesis, and for preventing oxidative damage in cells such as erythrocytes;
ribose 5-phosphate, used in nucleotide synthesis but also in NADH, FADH2 and coenzyme A synthesis.
Reaction 2
In the second step of the glycolysis, the isomerization of glucose 6-phosphate, an aldose, to fructose 6-phosphate, a ketose, occurs. This reaction is catalyzed by phosphoglucose isomerase, also known as phosphohexose isomerase or glucose phosphate isomerase (EC 5.3.1.9).
Glucose 6-phosphate ⇄ Fructose 6-phosphate
These molecules are an example of functional group isomerism.
Like hexokinase, phosphoglucose isomerase requires the presence of Mg2+.
The ΔG°’ of the reaction is 1.7 kJ/mol (0.4 kcal/mol), while the ΔG is -2.5 kJ/mol (-0.6 kcal/mol). These small values indicate that the reaction is close to equilibrium and is easily reversible.
The reaction essentially consists in the shift of the carbonyl group at C-1 of the open-chain form of glucose 6-phosphate to C-2 of the open-chain form of fructose 6-phosphate.
Phosphoglucose Isomerase Reaction
The enzymatic reaction can be divided at least into three steps. Since in aqueous solution both hexoses are primarily present in the cyclic form, the enzyme must first open the ring of G-6P, catalyze the isomerization, and, finally, the formation of the five-membered ring of F-6-P.
This isomerization is a critical step for glycolytic pathway, as it prepares the molecule for the subsequent two steps.
Why?
The phosphorylation that occurs in the third step requires the presence of an alcohol group at C-1, and not of a carbonyl group.
In the fourth step, the covalent bond between C-3 and C-4 is cleaved, and this reaction is facilitated by the presence of the carbonyl group at C-2.
Reaction 3
In the third step of the glycolytic pathway, a second phosphorylation reaction occurs. Phosphofructokinase 1 or PFK-1 (EC 2.7.1.11) catalyzes the phosphorylation of fructose 6-phosphate at C-1 to form fructose 1,6-bisphosphate, at the expense of one ATP.
PFK-1 is so named to distinguish it from phosphofructokinase 2 or PFK-2 (EC 2.7.1.105), the enzyme that catalyzes the phosphorylation of fructose 6-phosphate to fructose 2,6-bisphosphate.
Like the reaction catalyzed by hexokinase/glucokinase, this phosphorylation, too, is an essentially irreversible step, irreversibility, once again, achieved by coupling, by phosphofructokinase 1, with the hydrolysis of ATP. In fact, phosphorylation of fructose 6-phosphate by inorganic phosphate is endergonic, with a ΔG°’ of 16.3 kJ/mol (3.9 kcal/mol), whereas, when the reaction is coupled to the hydrolysis of ATP, the overall equation becomes exergonic, with a ΔG°’ of -14.2 kJ/mol (-3.4 kcal/mol) and a ΔG of -22.2 kJ/mol (-5.3 kcal/mol).
While hexokinase allows to trap glucose inside the cell, phosphofructokinase 1 prevents glucose to be used for glycogen synthesis or the production of other sugars, but is instead metabolized in the glycolytic pathway. In fact, unlike glucose 6-phosphate, fructose 1,6-bisphosphate cannot be used directly in other metabolic pathways than glycolysis/gluconeogenesis, that is, phosphofructokinase 1 catalyzes the first “committed” step of the glycolytic pathway. Such reactions are usually catalyzed by enzymes regulated allosterically, that prevent the accumulation of both intermediates and final products. PFK-1 is no exception, being subject to allosteric regulation by positive and negative effectors that signal the energy level and the hormonal status of the organism.
Some protists and bacteria, and perhaps all plants, have a phosphofructokinase that uses pyrophosphate (PPi) as a donor of the phosphoryl group in the synthesis of F-1,6-BP. This reaction has a ΔG°’ of -2.9 kJ/mol (-12.1 kcal/mol).
Fructose 6-phosphate + PPi → Fructose 1,6-bisphosphate + Pi
Note: The prefix bis– in bisphosphate, as fructose 1,6-bisphosphate, indicates that there are two phosphoryl groups are bonded to different atoms.
The prefix di– in diphosphate, as in adenosine diphosphate, indicates that there are two phosphoryl groups connected by an anhydride bond to form a pyrophosphoryl group, namely, they are directly bonded to one another.
Similar rules also apply to the nomenclature of molecules that have three phosphoryl groups standing apart, such as inositol 1,4,5-trisphosphate, or connected by anhydride bonds, such as ATP or guanosine triphosphate or GTP.
Reaction 4
In the fourth step of the glycolysis, fructose 1,6-bisphosphate aldolase, often called simply aldolase (EC 4.1.2.13), catalyzes the reversible cleavage of fructose 1,6-bisphosphate into glyceraldehyde 3-phosphate, an aldose, and dihydroxyacetone phosphate, a ketose. The enzyme cleaves the bond between C-3 and C-4.
All glycolytic intermediates downstream to this reaction are three-carbon molecules, instead of six-carbon molecules as the previous ones.
The ΔG°’ of the reaction in the direction of glyceraldehyde 3-phosphate and dihydroxyacetone phosphate production is of 23.8 kJ/mol (5.7 kcal/mol), and the Km is approximately 10-4 M, values that would indicate that the reaction does not proceed as written from left to right. However, under normal cellular conditions, due to the lower concentrations of the reactants, the ΔG is -1.3 kJ/mol (-0.3 kcal/mol), a very small value, thus the reaction is easily reversible, that is, essentially to equilibrium.
Note: The name “aldolase” derives from the nature of the reverse reaction, from right to left as written, that is, an aldol condensation.
Reaction 5
Of the two products of the previous reaction, glyceraldehyde 3-phosphate goes directly into the second phase of the glycolytic pathway. Conversely, DHAP is not on the direct pathway of glycolysis and must be converted, isomerized, to glyceraldehyde 3-phosphate to continue through the pathway. This isomerization is catalyzed by triose phosphate isomerase (EC 5.3.1.1).
Triose phosphate isomerase, in converting dihydroxyacetone phosphate into glyceraldehyde 3-phosphate, catalyzes the transfer of a hydrogen atom from C-1 to C-2, that is, catalyzes an intramolecular oxidation-reduction. And in essence, after the enzyme reaction, the carbons C-1, C-2 and C-3 of the starting glucose to become equivalent, chemically indistinguishable, from the carbons C-6, C-5 and C-4, respectively.
Therefore, the net result of the last two steps of glycolysis is the production of two molecules of glyceraldehyde 3-phosphate.
The ΔG°’ of the reaction is of 7.5 kJ/mol (1.8 kcal/mol), while the ΔG is 2.5 kJ/mol (0.6 kcal/mol). Although at equilibrium dihydroxyacetone phosphate represent about 96 percent of the triose phosphates, the reaction proceeds readily towards the formation of glyceraldehyde 3-phosphate because of the subsequent step of the glycolytic pathway that removes the glyceraldehyde 3-phosphate produced.
One of the distinguishing features of triose phosphate isomerase is the great catalytic efficiency. The enzyme is in fact considered kinetically perfect. Why? The enzyme enhances the isomerization rate by a factor of 1010 compared with that obtained with a catalyst such as acetate ion. Indeed, the Kcat/KM ratio for the isomerization of glyceraldehyde 3-phosphate is equal to 2×108 M-1s-1, value close to the diffusion-controlled limit. Thus, the rate-limiting step in the reaction catalyzed by triose phosphate isomerase is diffusion-controlled encounter of enzyme and substrate.
From the energetic point of view, the last two steps of glycolysis are unfavorable, with ΔG°’ of 31.3 kJ/mol (7.5 kcal/mol), whereas the net ΔG°’ of the first five reactions is of 2.1 kJ/mol (0.5 kcal/mol), with a Keq of about 0.43. And it is the free energy derived from the hydrolysis two ATP that, under standard-state conditions, makes the value of the overall equilibrium constant close to one. If instead we consider ΔG, it is quite negative, -56.8 kJ/mol (-13.6 kcal/mol).
Notice that dihydroxyacetone phosphate may also be reduced to glycerol 3-phosphate in the reaction catalyzed by cytosolic glycerol 3-phosphate dehydrogenase (EC 1.1.1.8).
The enzyme acts as a bridge between glucose and lipid metabolism because the glycerol 3-phosphate produced is used in the synthesis of lipids such as triacylglycerols.
This reaction is an important source of glycerol 3-phosphate in adipose tissue and small intestine.
Reaction 6
In the sixth step of the glycolysis, the first step of the second phase, the payoff phase, glyceraldehyde 3-phosphate dehydrogenase (EC 1.2.1.12) catalyses the oxidation of glyceraldehyde 3-phosphate to 1,3-bisphosphoglycerate (1,3-BPG), with the concomitant reduction of NAD+ to NADH.
This is the first of the two glycolytic reactions in which the chemical energy needed for the subsequent synthesis of ATP is harvested and made available; the other reaction is catalyzed by enolase (EC 4.2.1.11). Why?
This reaction is the sum of two processes.
In the first reaction, the oxidation of the aldehyde group to a carboxyl group occurs, step in which NAD+ is used as oxidizing agent. The reaction is quite exergonic, with a ∆G’° of -43 kJ/mol (-10.3 kcal/mol).
In the second reaction, the formation of the bond between the carboxylic group at C-1 of 1,3-bisphosphoglycerate and orthophosphate occurs, to form an anhydride called acyl phosphate. The reaction is quite endergonic, with a ∆G’° of 49.3 kJ/mol (11.8 kcal/mol).
These two chemical processes must not take place in succession but must be coupled in order to allow the formation of the acyl phosphate because the oxidation of the aldehyde group is used to drive the formation of the anhydride, with an overall ΔG°’ of 6.3 kJ/mol (1.5 kcal/mol), and a ΔG of 2.5 kJ/mol (0.6 kcal/mol), both slightly endergonic.
Therefore, the free energy that might be released as heat is instead conserved by the formation of the acyl phosphate.
Note: The reversible reduction of the nicotinamide ring of NAD+ or NADP+ is due to the loss of two hydrogen atoms by another molecule, in this case the aldehyde group of glyceraldehyde 3-phosphate, that undergoes oxidation, and to the subsequent transfer of a hydride ion, the equivalent of two electrons and a proton, to the nicotinamide ring. The other proton removed from the substrate is released to the aqueous solution. Below, the half reactions for both coenzymes.
NAD+ + 2 e– + 2 H+ → NADH + H+
NADP+ + 2 e– + 2 H+ → NADH + H+
Reaction 7
In the seventh step of the glycolytic pathway, phosphoglycerate kinase (EC 2.7.2.3) catalyzes the transfer of the high-energy phosphoryl group from the acyl phosphate of 1,3-BPG to ADP to form ATP and 3-phosphoglycerate (3-PG).
1,3-Bisphosphoglycerate + ADP + H+ ⇄ 3-Phosphoglycerate + ATP
ΔG°’ is of -18.5 kJ/mol (-4.4 kcal/mol), namely, it is an exergonic reaction, whereas ΔG is 1.3 kJ/mol (0.3 kcal/mol).
The high phosphoryl-transfer potential of the acyl phosphate is used to phosphorylate ADP. The production of ATP in this manner is called substrate-level phosphorylation. In other words, part of the energy released during the oxidation of the aldehyde group in the sixth step is now conserved by the synthesis of ATP from the ADP and Pi.
The reaction catalyzed by phosphoglycerate kinase is the first reaction of glycolysis in which part of the chemical energy present in glucose molecule is conserved as ATP. And, because the reactions catalyzed by aldolase and triose phosphate isomerase, step 4 and 5, respectively, lead to the formation of two molecules of glyceraldehyde 3-phosphate per molecule of glucose, in this step two ATP are produced and the ATP debt created by the preparatory phase, steps 1 and 3, respectively, is “paid off”.
It should be noted that the enzyme is named for the reverse reaction, from right to left as written, that is, the phosphorylation of 3-phosphoglycerate to form 1,3-bisphosphoglycerate at the expense of one ATP.
Indeed, this enzyme, like all other enzymes, is able to catalyze the reaction in both directions. And the direction leading to the synthesis of 1,3-bisphosphoglycerate occurs during the photosynthetic CO2 fixation and gluconeogenesis.
The sixth and seventh reactions of glycolysis, are, as a whole, an energy-coupling process in which the common intermediate is 1,3-bisphosphoglycerate. While the reaction leading to the synthesis of 1,3-BPG is endergonic, with a ΔG°’ of 6.3 kJ/mol (1.5 kcal/mol), the second reaction is strongly exergonic, with a ΔG°’ of -18.5 kJ/mol (-4,4 kcal/mol). The overall ΔG°’ is -12.2 kJ/mol (-2.9 kcal/mol), namely, the reaction catalyzed by phosphoglycerate kinase is sufficiently exergonic to pull even the previous one, too, making the overall reaction exergonic.
Glyceraldehyde 3-phosphate + ADP + Pi + NAD+ ⇄ 3-Phosphoglycerate + ATP + NADH + H+
In reality, phosphoglycerate kinase reaction is sufficiently exergonic to pull also the reactions catalyzed by aldolase and triose phosphate isomerase.
What is substrate-level phosphorylation?
Substrate-level phosphorylation is defined as the production of ATP by the transfer of a phosphoryl group from a substrate to ADP, a process involving chemical intermediates and soluble enzymes.
There is also a second type of phosphorylation for the synthesis of ATP called oxidative phosphorylation, a process involving not chemical intermediates and soluble enzymes but transmembrane proton gradients and membrane-bound enzymes.
Because the standard free energy of hydrolysis of the phosphoryl group of 3-phosphoglycerate is equal to 12.5 kJ/mol (-3 kcal/mol), it is not sufficient to produce ATP by phosphoryl group transfer. In the two subsequent reactions of glycolysis, 3-phosphoglycerate is converted to phosphoenolpyruvate (PEP), a molecule with a phosphoryl group transfer potential sufficiently elevated to allow the synthesis of ATP.
Reaction 8
In the eighth step of the glycolysis, 3-phosphoglycerate is converted into 2-phosphoglycerate (2-PG), in a reversible reaction catalyzed by phosphoglycerate mutase (EC 5.4.2.1). The reaction requires Mg2+, and has a very small ΔG, equal to about 0.8 kJ/mol (0.2 kcal/mol) and a ΔG°’ of 4.4 kJ/mol (1.1 kcal/mol).
Phosphoglycerate mutase is a mutase, enzymes that catalyze intramolecular group transfers, in this case the transfer of a phosphoryl group from C-3 to C-2 of the 3-phosphoglycerate. Mutases, in turn, are a subclass of isomerases.
The mechanism by which this reaction takes place depends on the type of organism studied. For example, in yeast or in rabbit muscle the reaction occurs in two steps and involves the formation of phosphoenzyme intermediates. In the first step, a phosphoryl group bound to a histidine residue in the active site of the enzyme is transferred to the hydroxyl group at C-2 of 3-PG to form 2,3-bisphosphoglycerate. In the next step, the enzyme acts as a phosphatase converting 2,3-BPG into 2-phosphoglycerate; however, the phosphoryl group at C-3 is not released but linked to the histidine residue of the active site to regenerate the intermediate enzyme-His-phosphate. Schematically:
Notice that the phosphoryl group of 2-phosphoglycerate is not the same as that of the substrate 3-phosphoglycerate.
Approximately once in every 100 catalytic cycles, 2,3-BPG dissociates from the active site of the enzyme, leaving it unphosphorylated, that is, in the inactive form. The inactive enzyme may be reactivated by binding 2,3-bisphosphoglycerate, which must, therefore, be present in the cytosol to ensure the maximal activity of the enzyme. And 2,3-BPG is present in small, but sufficient amounts in most cells, except for red blood cells, where it acts as an allosteric inhibitor, too, reducing the affinity of hemoglobin for oxygen, and has a concentration of 4-5 mM.
Note: 3-Phosphoglycerate can also be used for the biosynthesis of serine, from which glycine and cysteine derive. The biosynthesis of serine begins with the reaction catalyzed by phosphoglycerate dehydrogenase (EC 1.1.1.95). The enzyme catalyzes the oxidation of 3-phosphoglycerate to 3-phosphohydroxypyruvate and the concomitant reduction of NAD+ to NADH. This reaction is also the rate-limiting step of this biosynthetic pathway, because serine inhibits the activity of the enzyme.
Synthesis of 2,3-bisphosphoglycerate and the Rapoport-Luebering pathway
1,3-Bisphosphoglycerate can be also converted into 2,3-bisphosphoglycerate.
In red blood cells this reaction is catalyzed by the bisphosphoglycerate mutase,[6] one of the three isoforms of phosphoglycerate mutase found in mammals. The enzyme requires the presence of 3-phosphoglycerate as it catalyzes the intermolecular transfer of a phosphoryl group from C-1 of 1,3-bisphosphoglycerate to the C-2 of 3-phosphoglycerate. Therefore, 3-phosphoglycerate becomes 2,3-BPG, while 1,3-BPG is converted into 3-phosphoglycerate. The mutase enzyme activity has EC number 5.4.2.4.
Synthesis of 2,3-BPG
2,3-Bisphosphoglycerate can then be hydrolyzed to 3-phosphoglycerate in the reaction catalyzed by the phosphatase activity of bisphosphoglycerate mutase, that removes the phosphoryl group at C-2. This activity has EC number 3.1.3.13. The enzyme is also able to catalyze the interconversion of 2-phosphoglycerate and 3-phosphoglycerate, therefore, it is a trifunctional enzyme. 3-Phosphoglycerate can then re-enter the glycolytic pathway. This detour from glycolysis, also called Rapoport-Luebering pathway, that leads to the synthesis of 3-phosphoglycerate without any ATP production.
The other two isoforms of phosphoglycerate mutase, phosphoglycerate mutase 1 or type M, present in the muscle, and phosphoglycerate mutase 2 or type B, present in all other tissues, are able to catalyze, in addition to the interconversion of the 2-phosphoglycerate and 3-phosphoglycerate, the two steps of Rapoport-Luebering pathway, although with less efficacy than the glycolytic reaction. Therefore, they are trifunctional enzymes.
Reaction 9
In the ninth step of the glycolytic pathway, 2-phosphoglycerate is dehydrated to form phosphoenolpyruvate, an enol, in a reversible reaction catalyzed by enolase.
2-Phosphoglycerate ⇄ Phosphoenolpyruvate + H2O
The reaction requires Mg2+ that stabilizes the enolic intermediate that is formed during the process.
The ΔG°’ of the reaction is 7.5 kJ/mol (1.8 kcal/mol), while ΔG -3.3 kJ/mol (-0.8 kcal/mol).
Like 1,3-BPG, phosphoenolpyruvate has a phosphoryl group transfer potential high enough to allow ATP formation. Why does this phosphoryl group have a high free energy of hydrolysis?
Although phosphoenolpyruvate and 2-phosphoglycerate contain nearly the same amount of metabolic energy with respect to decomposition to CO2, H20 and Pi, 2-PG dehydration leads to a redistribution of energy such that the standard free energy of hydrolysis of the phosphoryl groups vary as described below:
-17.6 kJ/mol (-4.2 kcal/mol) for 2-phosphoglycerate, a phosphoric ester;
-61.9 kJ/mol (-14.8 kcal/mol) for phosphoenolpyruvate, an enol phosphate.
What happens is that the phosphoryl group traps PEP in its unstable enol form. When, in the last step of glycolysis, phosphoenolpyruvate donates the phosphoryl group to ADP, ATP and the enol form of pyruvate are formed. The enol form of pyruvate is unstable and tautomerizes rapidly and nonenzymatically to the more stable keto form, that predominates at pH 7. So, the high phosphoryl-transfer potential of PEP is due to the subsequent enol-keto tautomerization of pyruvate.
Reaction 10
In the final step of the glycolysis, pyruvate kinase (EC 2.7.1.40) catalyzes the transfer of the phosphoryl group from phosphoenolpyruvate to ADP to form pyruvate and ATP. This is the second substrate-level phosphorylation of glycolysis.
Phosphoenolpyruvate + ADP + H+ → Pyruvate + ATP
The enzyme is a tetramer and, like PFK-1, is a highly regulated. Indeed, it has binding sites for numerous allosteric effectors. Moreover, in vertebrates, there are at least three isozymes of pyruvate kinase, of which the M type predominates in muscle and brain, while the L type in liver. These isozymes have many properties in common, whereas differ in the response to hormones such as glucagon, epinephrine and insulin.
The enzyme activity is stimulated by potassium ion (K+) and some other monovalent cations.
The reaction is essentially irreversible, with a ΔG°’ of -31.4 kJ/mol (-7.5 kcal/mol), and a ΔG of -16.7 kJ/mol (-4.0 kcal/mol), largely due, as anticipated in the previous paragraph, to the tautomerization of the pyruvate from the enol form to the more stable keto form.
Spontaneous Tautomerization of Pyruvate
And, of the -61.9 kJ/mol (14.8 kcal/mol) released from the hydrolysis of the phosphoryl group of PEP, nearly half is conserved in the formation of the phosphoanhydride bond between ADP and Pi, whose ΔG°’ is of -30.5 kJ/mol (-7.3 kcal/mol). The remaining energy, -31.4 kJ/mol (-7.5 kcal/mol), is the driving force that makes the reaction proceed towards ATP production.
While the reaction catalyzed by phosphoglycerate kinase, in the seventh step of the glycolytic pathway, pays off the ATP debt of the preparatory phase, the reaction catalyzed by pyruvate kinase allows a net gain of two ATP.
Fate of NADH and pyruvate
Glycolysis produces 2 NADH, 2 ATP, and 2 pyruvate molecules per molecule of glucose.
NADH must be reoxidized to NAD+ to allow glycolysis to proceed.[2] NAD+, a coenzyme that is produced from the vitamin B3, also known as niacin, is present in limited amounts in the cytosol, ≤ 10-5M, a value well below than that of glucose metabolized in a few minutes, and must be continuously regenerated. Therefore, the final step of the glycolytic pathway is the regeneration of NAD+ from NADH through aerobic or anaerobic pathways, each of which involves pyruvate. Such pathways allow, therefore, maintenance of the redox balance of the cell.
Pyruvate is a versatile metabolite that can enter several metabolic pathways, both anabolic and catabolic, depending on the type of cell, the energy state of the cell and the availability of oxygen.[5]
Catabolic Fates of Pyruvate
With the exception of some variations encountered in bacteria, exploited, for example, in food industry for the production of various foods such as many cheeses, there are essentially three pathways in which pyruvate may enter:
reduction to lactate, through lactic acid fermentation;
reduction to ethanol or ethyl alcohol, through alcoholic fermentation;
aerobic oxidation.
This allows glycolysis to proceed in both anaerobic and aerobic conditions.
It is therefore possible to state that the catabolic fate of the carbon skeleton of glucose is influenced by the cell type, the energetic state of the cell, and the availability of oxygen.
Lactic acid fermentation
In animals, with few exceptions, and in many microorganisms when oxygen availability is insufficient to meet the energy requirements of the cell, or if the cell is without mitochondria, the pyruvate produced by glycolysis is reduced to lactate in the cytosol, in a reaction catalyzed by lactate dehydrogenase (EC 1.1.1.27).
Pyruvate + NADH + H+ ⇄ Lactate + NAD+
In the reaction, pyruvate, by accepting electrons from NADH, is reduced to lactate, while NAD+ is regenerated. And the overall equilibrium of the reaction strongly favors the formation of lactate, as shown by the value of ΔG°’ of -25.1 kJ/mol (-6 kcal/mol).
The conversion of glucose to lactate is called lactic acid fermentation. The overall equation of the process is:
Glucose + 2 Pi + 2 ADP + 2H+ → 2 Lactate + 2 ATP + 2 H2O
Notice that fermentation, discovered by Louis Pasteur who defined it “la vie sans l’air”, is a metabolic pathway that:
extracts energy from glucose and stores it as ATP;
does not consume oxygen;
does not change the concentration of NAD+ or NADH.
With regard to coenzymes, neither NAD+ nor NADH appears in the overall equation, although both are crucial in the process, that is, no net oxidation-reduction occurs. In other words, in the conversion of glucose, C6H12O6, to lactate, C3H6O3, the ratio of hydrogen to carbon atoms of the reactants and products does not change.
From an energy point of view, it should however be emphasized that fermentation extracts only a small amount of the chemical energy of glucose.
In humans, much of the lactate produced enters the Cori cycle for glucose production via gluconeogenesis. We can also state that lactate production shifts part of the metabolic load from the extrahepatic tissues, such as skeletal muscle during intense bouts of exercise, like a 200-meter, when the rate of glycolysis can almost instantly increase 2,000-fold, to the liver.
In contrast to skeletal muscle that releases lactate into the venous blood, the heart muscle is able to take up and use it for fuel, due to its completely aerobic metabolism and to the properties of the heart isozyme of lactate dehydrogenase, referred to as H4. Therefore, portion of the lactate released by skeletal muscle engaged in intense exercise is used by the heart muscle for fuel.
Note: Lactate produced by microorganisms during lactic acid fermentation is responsible for both the scent and taste of sauerkraut, namely, fermented cabbage, as well as for the taste of soured milk.
Alcoholic fermentation
In microorganisms such as brewer’s and baker’s yeast, in certain plant tissues, and in some invertebrates and protists, pyruvate, under hypoxic or anaerobic conditions, may be reduced in two steps to ethyl alcohol or ethanol, with release of CO2.
The first step involves the non-oxidative decarboxylation of pyruvate to form acetaldehyde, an essentially irreversible reaction catalyzed by pyruvate decarboxylase (EC 4.1.1.1), an enzyme that requires Mg2+ and thiamine pyrophosphate, a coenzyme derived from vitamin thiamine or vitamin B1. The enzyme is absent in vertebrates and in other organisms that perform lactic acid fermentation.
In the second step, acetaldehyde is reduced to ethanol in a reaction catalyzed by alcohol dehydrogenase (EC 1.1.1.1), an enzyme that contains a bound zinc atom in its active site. In the reaction, NADH supplies the reducing equivalents and is oxidized to NAD+. At neutral pH, the equilibrium of the reaction lies strongly toward ethyl alcohol formation.
The conversion of glucose to ethanol and CO2 is called alcoholic fermentation. The overall reaction is:
Glucose + 2 Pi + 2 ADP + 2 H+ → 2 Ethanol + 2 CO2 + 2 ATP + 2 H2O
And, as for lactic fermentation, even in alcoholic fermentation no net oxidation-reduction occurs.
Alcoholic fermentation is the basis of the production of beer and wine. Notice that the CO2 produced by brewer’s yeast is responsible for the characteristics “bubbles” in beer, champagne and sparkling wine, while that produced by baker’s yeast causes dough to rise.
Fate of pyruvate and NADH under aerobic conditions
In cells with mitochondria and under aerobic conditions, the most common situation in multicellular and many unicellular organisms, the oxidation of NADH and pyruvate catabolism follow distinct pathways.
Pyruvate, in the mitochondrial matrix, is first converted to acetyl-CoA in the reactions catalyzed by the pyruvate dehydrogenase complex, one of the mitochondrial multienzyme complexes. In the reaction, an oxidative decarboxylation, pyruvate loses a carbon atom as CO2, and the remaining two carbon units iare bound to Coenzyme A to form acetyl-coenzyme A or acetyl-CoA.
Pyruvate + NAD+ + CoA → acetyl-CoA + CO2 + NADH + H+
The acetyl group of acetyl-CoA is then completely oxidized to CO2 in the citric acid cycle, with production of NADH and FADH2. The pyruvate dehydrogenase complex therefore represents a bridge between glycolysis, which occurs in the cytosol, and the citric acid cycle, which occurs in the mitochondrial matrix.
In turn, electrons derived from oxidations that occur during glycolysis are transported into mitochondria via the reduction of cytosolic intermediates. In this way, in the cytosol NADH is oxidized to NAD+, while the reduced intermediate, once in the mitochondrial matrix, is reoxidized through the transfer of its reducing equivalents to Complex I of the mitochondrial electron transport chain. Here the electrons flow to oxygen to form H2O, a transfer that supplies the energy needed for the synthesis of ATP through the process of oxidative phosphorylation. Of course, also the electrons carried by NADH formed by pyruvate dehydrogenase complex reactions and citric acid cycle and by FADH2 formed by citric acid cycle meet a similar fate.
Note: FADH2 transfers its reducing equivalents not to Complex I but to Complex II.
Anabolic fates of pyruvate
Under anabolic conditions, the carbon skeleton of pyruvate may have fates other than complete oxidation to CO2 or conversion to lactate. In fact, after its conversion to acetyl-CoA, it may be used, for example, for the synthesis of fatty acids, or of the amino acid alanine.
Glycolysis and ATP production
In the glycolytic pathway the glucose molecule is degraded to two molecules of pyruvate.
During the first phase, the preparatory phase, two ATP are consumed per molecule of glucose in the reactions catalyzed by hexokinase and PFK-1. In the second phase, the payoff phase, 4 ATP are produced through substrate-level phosphorylation in the reactions catalyzed by phosphoglycerate kinase and pyruvate kinase. So there is a net gain of two ATP per molecule of glucose used. In addition, in the reaction catalyzed by glyceraldehyde 3-phosphate dehydrogenase, two molecules of NADH are produced for each glucose molecule.[5]
Energy Changes of Glycolytic Reactions
The overall ΔG°’ of glycolysis is -85 kJ/mol (-20.3 kcal/mol), value resulting from the difference between the ΔG°’ of the conversion of glucose into two pyruvate molecules, -146 kJ/mol (-34,9 kcal/mol), and the ΔG°’ of the formation of ATP from ADP and Pi, 2 x 30.5 kJ/mol = 61 kJ / mol (2 x 7.3 kcal/mol = 14.6 kcal/mol). Here are the two reactions.
Glucose + 2 NAD+ → 2 Pyruvate + 2 NADH + 2 H+
2 ADP + 2 Pi → 2 ATP + 2 H2O
The sum of the two reactions gives the overall equation of glycolysis.
Thus, under standard conditions, the amount of released energy stored within ATP is (61/146) x 100 = 41.8 percent.
Notice that the overall equation of glycolysis can also be derived by considering all the reagents, ATP, NAD+, ADP, and Pi and all the products.
Cancelling the common terms on both sides of the equation, we obtain the overall equation shown above.
ATP production under anaerobic conditions
Under anaerobic conditions, regardless of what is the metabolic fate of pyruvate, conversion to lactate, ethanol or other molecules, there is no additional production of ATP downstream of glycolysis.
Therefore, under these conditions, glycolysis extracts only a small fraction of the chemical energy of the glucose molecule, energy equal to 2840 kJ/mol (679 kcal/mol) released as a result of its conversion to CO2 and H2O. Indeed, only 146 kJ/mol are released in the conversion of a glucose molecule to two pyruvate molecules, equal to 5 percent, [(146/2,840) x 100], of the available chemical energy. Therefore, pyruvate still contains most of the chemical energy of the hexose.
Similarly, the 4 electrons carried by NADH produced in step 6 of glycolysis cannot be used for ATP production.
In lactic acid fermentation, the ΔG°’ associated with the conversion of a glucose molecule to two molecules of lactate is -183.6 kJ/mol (-43.9 kcal/mol), and 33.2 percent of such free energy, [(61/183.6) x 100] is stored within ATP, whereas it is 41.8 percent in the conversion of a glucose molecule to two molecules of pyruvate.
It should be noted that under actual conditions the amount of free energy required for the synthesis of ATP from ADP and Pi is much higher than that required under standard conditions, namely, approximately 50 percent of the energy released is stored within ATP.
ATP production under aerobic conditions
Under aerobic conditions, in cells with mitochondria, the amount of chemical energy that can be extracted from glucose and stored within ATP is much greater than under anaerobic conditions.
If we consider the two NADH produced during glycolysis, the flow of their 4 reducing equivalents along the mitochondrial electron transport chain allows the production of 2-3 ATP per electron pair through oxidative phosphorylation. Therefore, 6 to 8 ATP are produced when one molecule of glucose is converted into two molecules of pyruvate, 2 from glycolysis and 4-6 from oxidative phosphorylation.
Note: The amount of ATP produced from the reducing equivalents of NADH depends upon the mechanism by which they are shuttled into mitochondria.
On the other hand, if we analyze the coordinated and consecutive action of glycolysis, the pyruvate dehydrogenase complex, citric acid cycle, mitochondrial electron transport chain and oxidative phosphorylation, much more energy can be extracted from glucose and stored within ATP. In this case, according to what reported by Lehninger, 30 to 32 ATP are produced for each glucose molecule, although recent estimates suggest a net production equal to 29.85 ATP/glucose, or 29.38 ATP/glucose if also ATP formed from GTP, in turn produced by the citric acid cycle, is exported. Considering both estimates, the production of ATP is about 15 times greater than under anaerobic condition.
Feeder pathways
Other carbohydrates besides glucose, both simple and complex, can be catabolized via glycolysis, after enzymatic conversion to one of the glycolytic intermediates.[2][7]
Among the most important are:
the monosaccharides galactose, fructose, and the less common mannose.
Feeder Pathways for Glycolysis
At the intestinal level, starch and disaccharides encounter the enzymes responsible for carbohydrate digestion. Then, the absorption of monosaccharides released or derived from diet occur. Once in the venous circulation, monosaccharides reach the liver through the portal vein. This organ is the main site where they are metabolized.
Glycogen and starch
Regarding the phosphorolytic breakdown of starch and endogenous glycogen refer to the corresponding articles.
Fructose
Under physiological conditions, the liver removes much of the ingested fructose from the bloodstream before it can reach extrahepatic tissues.
The hepatic pathway for the conversion of the monosaccharide to intermediates of glycolysis consists of several steps.
In the first step fructose is phosphorylated to fructose 1-phosphate at the expense of one ATP. This reaction is catalyzed by fructokinase (EC 2.7.1.4), and requires the presence of Mg2+.
Fructose + ATP → Fructose 1-phosphate + ADP + H+
As for glucose, fructose phosphorylation traps the molecule inside the cell.
In the second step fructose 1-phosphate aldolase catalyzes the hydrolysis, an aldol cleavage, of fructose 1-phosphate to dihydroxyacetone phosphate and glyceraldehyde.
Dihydroxyacetone phosphate is an intermediate of the glycolytic pathway and, after conversion to glyceraldehyde 3-phosphate, may flow through the pathway. Conversely, glyceraldehyde is not an intermediate of the glycolysis, and is phosphorylated to glyceraldehyde 3-phosphate at the expense of one ATP. The reaction is catalyzed by triose kinase (EC 2.7.1.28), and requires the presence of Mg2+.
Glyceraldehyde + ATP → Glyceraldehyde 3-phosphate + ADP + H+
In hepatocytes, therefore, a molecule of fructose is converted to two molecules of glyceraldehyde 3-phosphate, at the expense of two ATP, as for glucose.
In extrahepatic sites, such as skeletal muscle, kidney or adipose tissue, fructokinase is not present, and fructose enters the glycolytic pathway as fructose 6-phosphate. In fact, as previously seen, hexokinase can catalyze the phosphorylation of fructose at C-6.
Fructose + ATP → Fructose 6-phosphate + ADP + H+
However, the affinity of the enzyme for fructose is about 20 times lower than for glucose, so in the hepatocyte, where glucose is much more abundant than fructose, or in the skeletal muscle under anaerobic conditions, that is, when glucose is the preferred fuel, little amounts of fructose 6-phosphate are formed.
Conversely, in adipose tissue, fructose is more abundant than glucose, so that its phosphorylation by hexokinase is not competitively inhibited to a significant extent by glucose. In this tissue, therefore, fructose 6-phosphate synthesis is the entry point into glycolysis for the monosaccharide.
With regard to the metabolic effects of fructose, it is important to underline that in the liver the monosaccharide, being phosphorylated at C-1, enters glycolysis at triose phosphate level, thus downstream to the reaction catalyzed by PFK-1, an enzyme that plays a key role in the regulation of the flow of carbon through this metabolic pathway. Conversely, when fructose is phosphorylated at C-6, it enters the glycolytic pathway upstream of PFK-1.
Sorbitol
Fructose is the entry point into glycolysis for sorbitol, a sugar present in many fruits and vegetables, and used as a sweetener and stabilizer, too. In the liver, sorbitol dehydrogenase (EC 1.1.99.21) catalyzes the oxidation of sorbitol to fructose. The enzyme is also part of the polyol pathway, for the conversion of glucose to fructose.
Sorbitol + NAD+ → Fructose + NADH + H+
The reaction requires the presence of zinc ion, and occurs in the cytosol.
Galactose
Galactose, for the most part derived from intestinal digestion of the lactose, once in the liver is converted, via the Leloir pathway, to glucose 1-phosphate.
The metabolic fate of glucose 1-phosphate depends on the energy status of the cell.
Under conditions promoting glucose storage, glucose 1-phosphate can be channeled to glycogen synthesis. Conversely, under conditions that favor the use of glucose as fuel, glucose 1-phosphate is isomerized to glucose 6-phosphate in the reversible reaction catalyzed by phosphoglucomutase (EC 5.4.2.2).
Glucose 1-phosphate ⇄ Glucose 6-phosphate
In turn, glucose 6-phosphate can be channeled to glycolysis and be used for energy production, or dephosphorylated to glucose in the reaction catalyzed by glucose 6-phosphatase, and then released into the bloodstream.
Mannose
Mannose is present in various dietary polysaccharides, glycolipids and glycoproteins. In the intestine, it is released from these molecules, absorbed, and, once reached the liver, is phosphorylated at C-6 to form mannose 6-phosphate, in the reaction catalyzed by hexokinase.
Mannose + ATP → Mannose 6-phosphate + ADP + H+
The sugar phosphate is then isomerized to fructose 6-phosphate in the reaction catalyzed by mannose 6-phosphate isomerase (EC 5.3.1.8.).
Mannose 6-phosphate ⇄ Fructose 6-phosphate</p
Regulation of glycolysis
The flow of carbon through the glycolytic pathway is regulated in response to metabolic conditions, both inside and outside the cell, essentially to meet two needs: the production of ATP and the supply of precursors for biosynthetic reactions.
And in the liver, to avoid wasting energy, glycolysis and gluconeogenesis are reciprocally regulated so that when one pathway is active, the other slows down. During evolution this was achieved by selecting different enzymes to catalyze the essentially irreversible reactions of the two pathways, whose activity are regulated separately. Indeed, if these reactions proceeded simultaneously at high speed, they would create a futile cycle or substrate cycle.[7] A such fine regulation could not be achieved if a single enzyme operates in both directions.
The control of the glycolysis involves essentially the reactions catalyzed by hexokinase, PFK-1, and pyruvate kinase, whose activity is regulated through:
allosteric modifications, that occur on a timescale of milliseconds and are instantly reversible;
covalent modifications, that is, phosphorylations and dephosphorylations, that occur on a timescale of seconds;
changes in enzyme concentrations, resulting from changes in the rate of their synthesis and/or degradation, that occur on a timescale of hours.
Note: The main regulatory enzymes of gluconeogenesis are pyruvate carboxylase (EC 6.4.1.1) and fructose 1,6-bisphosphatase (EC 3.1.3.11).
Hexokinase
In humans, hexokinase has four tissue specific isozymes, designated as hexokinase I, II, III, and IV, encoded by as many genes.[2][5]
Hexokinase I is the predominant isozyme in the brain, whereas in skeletal muscle hexokinase I and II are present, accounting for 70-75 percent and 25-30 percent of the isozymes, respectively.
Hexokinase IV, also known as glucokinase (EC 2.7.1.2), is mainly present in hepatocytes and beta cells of the pancreas, where it is the predominant isozyme. In the liver it catalyzes, with glucose 6-phosphatase, the substrate cycle between glucose and glucose 6-phosphate. Glucokinase differs from the other hexokinase isozymes in kinetic and regulatory properties.[1]
Note: Isoenzymes or isozymes are different proteins that catalyze the same reaction, and that generally differ in kinetic and regulatory properties, subcellular distribution, or in the cofactors used. They may be present in the same species, in the same tissue or even in the same cell.
Comparison of the kinetic properties of hexokinase isozymes
The kinetic properties of hexokinase I, II, and III are similar.
Hexokinase I and II have a Km for glucose of 0.03 mM and 0.1 mM, respectively. Therefore, these isoenzymes work very efficiently at normal blood glucose levels, 4-5 mM.[7]
Conversely, glucokinase has a high Km for glucose, approximately 10 mM; this means that the enzyme works efficiently only when blood glucose concentration is high, for example after a meal rich in carbohydrates with a high glycemic index.
Regulation of the activity of hexokinases I-III
Hexokinases I-III are allosterically inhibited by glucose 6-phosphate, the product of their reaction. This ensures that glucose 6-phosphate does not accumulate in the cytosol when glucose is not needed for energy, for glycogen synthesis, for the pentose phosphate pathway, or as a source of precursors for biosynthetic pathways, leaving, at the same time, the monosaccharide in the blood, available for other organs and tissues. For example, when PFK-1 is inhibited, fructose 6-phosphate accumulates and then, due to phosphoglucose isomerase reaction, glucose 6-phosphate accumulates. Therefore, inhibition of PFK-1 leads to inhibition of hexokinases I-III.
In skeletal muscle, the activity of hexokinase I and II is coordinated with that of GLUT4, a low Km glucose transporter (5mM), whose translocation to the plasma membrane is induced by both insulin and physical activity. The combined action of GLUT4 on plasma membrane and hexokinase in the cytosol maintains a balance between glucose uptake and its phosphorylation. Because blood glucose concentration is between 4 and 5 mmol/L, its entry into the myocyte through GLUT4 may cause an increase in its concentration sufficient to saturate, or near saturate the enzyme, which therefore operates at or near its Vmax.
Regulation of the activity of hepatic glucokinase
Glucokinase differs in three respects from hexokinases I-III, and is particularly suitable for the role that the liver plays in the regulation of blood glucose levels.[1] Why?
As previously said, glucokinase has a Km for glucose of about 10 mM, much higher than the Km for glucose of hexokinases I-III, and higher than the value of fasting blood glucose levels (4-5 mM) as well. In the liver, where it is the predominant hexokinase isoenzyme, its role is to provide glucose 6-phosphate for the synthesis of glycogen and fatty acids. The activity of glucokinase is linked to that of GLUT2, the major glucose transporter in hepatocytes, with a high Km for glucose, approximately 10 mM. Hence, GLUT2 is very active when blood glucose concentration is high, rapidly equilibrating sugar concentrations in cytosol of hepatocytes and blood. Under such conditions glucokinase is active and converts glucose to glucose 6-phosphate, and, due to high Km for glucose, its activity continues to increase even when the intracellular concentration of the monosaccharide reaches or exceeds 10 mM. Therefore, the rate at which glucose uptake and phosphorylation occurs are determined by the value of blood glucose level itself. On the other hand, when glucose availability is low, its concentration in the cytosol of hepatocytes is just as low, much lower than the Km for glucose of glucokinase, so that glucose produced through gluconeogenesis and/or glycogenolysis is not phosphorylated and can leave the cell.
A similar situation also occurs in pancreatic beta cells, where the GLUT2/glucokinase system causes the intracellular G-6-P concentration to equalize with glucose concentration in the blood, allowing the cells to detect and respond to hyperglycemia.
Unlike hexokinases I-III, glucokinase is not inhibited by glucose 6-phosphate, that is, is not product inhibited, and catalyzes its synthesis even when it accumulates.
Glucokinase is inhibited by the reversible binding of glucokinase regulatory protein or GKRP, a liver-specific regulatory protein.[4] The mechanism of inhibition by GKRP occurs via the anchorage of glucokinase inside the nucleus, where it is separated from the other glycolytic enzymes.
Regulation of Glucokinase Activity
The binding between glucokinase and GKRP is much tighter in the presence of fructose 6-phosphate, whereas it is weakened by glucose and fructose 1-phosphate.
In the absence of glucose, glucokinase is in its super-opened conformation that has low activity. The rise in cytosolic glucose concentration causes a concentration dependent transition of glucokinase to its close conformation, namely, its active conformation that is not accessible for glucokinase regulatory protein. Hence, glucokinase is active and no longer inhibited.
Notice that fructose 1-phosphate is present in the hepatocyte only when fructose is metabolized. Hence, fructose relieves the inhibition of glucokinase by glucokinase regulatory protein. Example
After a meal rich in carbohydrates, blood glucose levels rise, glucose enters the hepatocyte through GLUT2, and then moves inside the nucleus through the nuclear pores. In the nucleus glucose determines the transition of glucokinase to its close conformation, active and not accessible to GKRP, allowing glucokinase to diffuse in the cytosol where it phosphorylates glucose.
Conversely, when glucose concentration declines, such as during fasting when blood glucose levels may drop below 4 mM, glucose concentration in hepatocytes is low, and fructose 6-phosphate binds to GKRP allowing it to bind tighter to glucokinase. This results in a strong inhibition of the enzyme. This mechanism ensures that the liver, at low blood glucose levels, does not compete with other organs, primarily the brain, for glucose.
In the cell, fructose 6-phosphate is in equilibrium with glucose 6-phosphate, due to phosphoglucose isomerase reaction. Through its association with GKRP, fructose 6-phosphate allows the cell to decrease glucokinase activity, so preventing the accumulation of intermediates.
To sum up, when blood glucose levels are normal, glucose is phosphorylated mainly by hexokinases I-III, whereas when blood glucose levels are high glucose can be phosphorylated by glucokinase as well.
Regulation of phosphofructokinase 1 activity
Phosphofructokinase 1 is the key control point of carbon flow through the glycolysis.
The enzyme, in addition to substrate binding sites, has several binding sites for allosteric effectors.
ATP, citrate, and hydrogen ions are allosteric inhibitors of the enzyme, whereas AMP, Pi and fructose 2,6-bisphosphate are allosteric activators.[8][9]
Regulation of PFK 1 and Fructose 1,6-bisphosphatase
It should be noted that ATP, an end product of glycolysis, is also a substrate of phosphofructokinase 1. Indeed, the enzyme has two binding sites for the nucleotide: a low-affinity regulatory site, and a high affinity substrate site.
What do allosteric effectors signal?
ATP, AMP and Pi signal the energy status of the cell.
The activity of PFK-1 increases when the energy charge of the cell is low, namely, when there is a need for ATP, whereas it decreases when the energy charge of the cell is high, namely when ATP concentration in the cell is high. How?
When the nucleotide is produced faster than it is consumed, its cellular concentration is high. Under such condition ATP, binding to its allosteric site, inhibits PFK-1 by reducing the affinity of the enzyme for fructose 6-phosphate. From the kinetic point of view, the increase in ATP concentration modifies the relationship between enzyme activity and substrate concentration, chancing the hyperbolic fructose 6-phosphate velocity curve into a sigmoidal one, and then, increasing Km for the substrate. However, under most cellular conditions, ATP concentration does not vary much. For example, during a vigorous exercise ATP concentration in muscle may lower of about 10 percent compared to the resting state, whereas glycolysis rate varies much more than would be expected by such reduction.
When ATP consumption exceeds its production, ADP and AMP concentrations rise, in particular that of AMP, due to the reaction catalyzed by adenylate kinase (EC 2.7.4.3), that form ATP from ADP.
ADP + ADP ⇄ ATP + AMP
The equilibrium constant, Keq, of the reaction is:
Keq = [ATP][AMP]/[ADP]2= 0.44
Under normal conditions, ADP and AMP concentrations are about 10 percent and often less than 1 percent of ATP concentration, respectively. Therefore, considering that the total adenylate pool is constant over the short term, even a small reduction in ATP concentration leads, due to adenylate kinase activity, to a much larger relative increase in AMP concentration. In turn, AMP acts by reversing the inhibition due to ATP.
Therefore, the activity of phosphofructokinase 1 depends on the cellular energy status:
when ATP is plentiful, enzyme activity decreases;
when AMP levels increase and ATP levels fall, enzyme activity increases.
Why isn’t ADP a positive effector of PFK-1? There are two reasons.
When the energy charge of the cell falls, ADP is used to regenerate ATP, in the reaction catalyzed by adenylate kinase Moreover, as previously said, a small reduction in ATP levels leads to larger-percentage changes in ADP levels and, above all, in AMP levels.
Hydrogen ions inhibit PFK-1. Such inhibition prevents, by controlling the rate of glycolysis, excessive lactate buildup and the consequent fall of blood pH.
Citrate is an allosteric inhibitor of PFK-1 that acts by enhancing the inhibitory effect of ATP.
It is the product of the first step of the citric acid cycle, a metabolic pathway that provides building blocks for biosynthetic pathways and directs electrons into mitochondrial electron transport chain for ATP synthesis via oxidative phosphorylation. High citrate levels in the cytosol mean that, in the mitochondria, an overproduction of building blocks is occurring and the current energy are met, namely, the citric acid cycle has reached saturation. Under such conditions glycolysis, that feeds the cycle under aerobic condition, can slow down, sparing glucose.
So, it should be noted that PFK-1 couples glycolysis and the citric acid cycle.
In the liver, the central control point of glycolysis and gluconeogenesis is the substrate cycle between F-6-P and F-1,6-BP, catalyzed by PFK-1 and fructose 1,6-bisphosphatase.
The liver plays a pivotal role in maintaining blood glucose levels within the normal range.
When blood glucose levels drop, glucagon stimulates hepatic glucose synthesis, via both glycogenolysis and gluconeogenesis, and at the same time signals the liver to stop consuming glucose to meet its needs.
Conversely, when blood glucose levels are high, insulin causes the liver to use glucose for energy, and to synthesize glycogen, and triglycerides.
In this context, the regulation of glycolysis and gluconeogenesis is mediated by fructose 2,6-bisphosphate, a molecule that allows the liver to play a major role in regulating blood glucose levels, and whose levels are controlled by insulin and glucagon.
As a result of binding to its allosteric site on PFK-1, fructose 2,6-bisphosphate increases the affinity of the enzyme for fructose 6-phosphate, its substrate, while decreases its affinity for the allosteric inhibitors citrate and ATP. It is remarkable to note that under physiological concentrations of the substrates and positive and negative allosteric effectors, phosphofructokinase 1 would be virtually inactive in the absence of fructose 2,6-bisphosphate.
On the other hand, the binding of fructose 2,6-bisphosphate to fructose 1,6-bisphosphatase inhibits the enzyme, even in the absence of AMP, another allosteric inhibitor of the enzyme.
Due to these effects, fructose 2,6-bisphosphate increases the net flow of glucose through glycolysis.
Another metabolite involved in the control of the flow of carbon through glycolysis and gluconeogenesis is xylulose 5-phosphate,[3] a product of the pentose phosphate pathway, whose concentration in hepatocytes rises after ingestion of a carbohydrate-rich meal. The molecule, by activating protein phosphatase 2A, finally leads to an increase in the concentration of fructose 2,6-bisphosphate, and then to an increase in the flow of carbon through glycolysis and to a reduction in the flow of carbon through gluconeogenesis.
Regulation of pyruvate kinase activity
A further control point of carbon flow through glycolysis and gluconeogenesis is the substrate cycle between phosphoenolpyruvate and pyruvate, catalyzed by pyruvate kinase for glycolysis, and by the combined action of pyruvate carboxylase and phosphoenolpyruvate carboxykinase (EC 4.1.1.32) for gluconeogenesis.[5][7]
All isozymes of pyruvate kinase are allosterically inhibited by high concentrations of ATP, long-chain fatty acids, and acetyl-CoA, all signs that the cell is in an optimal energy status. Alanine, too, that can be synthesized from pyruvate through a transamination reaction, is an allosteric inhibitor of pyruvate kinase; its accumulation signals that building blocks for biosynthetic pathways are abundant.
Regulation of Pyruvate Kinase Activity
Conversely, pyruvate kinase is allosterically activated by fructose 1,6-bisphosphate, the product of the first committed step of glycolysis. Therefore, F-1,6-BP allows pyruvate kinase to keep pace with the flow of intermediates. It should be underlined that, at physiological concentration of PEP, ATP and alanine, the enzyme would be completely inhibited without the stimulating effect of F-1,6-BP.
The hepatic isoenzyme, but not the muscle isoenzyme, is also subject to regulation through phosphorylation by:
protein kinase A or PKA, activated by the binding of glucagon to the specific receptor or epinephrine to β-adrenergic receptors;
calcium/calmodulin dependent protein kinase or CAMK, activated by the binding of epinephrine to α1-adrenergic receptors.
Phosphorylation of the enzyme decreases its activity, by increasing the Km for phosphoenolpyruvate, and slows down glycolysis.
For example, when the blood glucose levels are low, glucagon-induced phosphorylation decreases pyruvate kinase activity. The phosphorylated enzyme is also less readily stimulated by fructose 1,6-bisphosphate but more readily inhibited by alanine and ATP. Conversely, the dephosphorylated form of pyruvate kinase is more sensitive to fructose 1,6-bisphosphate, and less sensitive to ATP and alanine. In this way, when blood glucose levels are low, the use of glucose for energy in the liver slows down, and the sugar is available for other tissues and organs, such as the brain. However, it should be noted that pyruvate kinase does not undergo glucagon-induced phosphorylation in the presence of fructose 1,6-bisphosphate.
An increase in the insulin/glucagon ratio, on the other hand, leads to dephosphorylation of the enzyme and then to its activation. The dephosphorylated enzyme is more readily stimulated by its allosteric activators F-1,6-BP, and less readily inhibited by allosteric inhibitors alanine and ATP.
References
ab de la Iglesia N., Mukhtar M., Seoane J., Guinovart J.J., & Agius L. The role of the regulatory protein of glucokinase in the glucose sensory mechanism of the hepatocyte. J Biol Chem OK 2000;275(14):10597-10603. doi: 10.1074/jbc.275.14.10597
^ Kabashima T., Kawaguchi T., Wadzinski B.E., Uyeda K. Xylulose 5-phosphate mediates glucose-induced lipogenesis by xylulose 5-phosphate-activated protein phosphatase in rat liver. Proc Natl Acad Sci USA 2003;100:5107-5112. doi:10.1073/pnas.0730817100
^ Kaminski M.T., Schultz J., Waterstradt R., Tiedge M., Lenzen S., Baltrusch S. Glucose-induced dissociation of glucokinase from its regulatory protein in the nucleus of hepatocytes prior to nuclear export. BBA – Molecular Cell Research 2014;1843(3):554-564. doi:10.1016/j.bbamcr.2013.12.002
^ abcdefghi Nelson D.L., M. M. Cox M.M. Lehninger. Principles of biochemistry. 6th Edition. W.H. Freeman and Company, 2012
^ Oslund R.C., Su X., Haugbro M., Kee J-M., Esposito M., David Y., Wang B., Ge E., Perlman D.H., Kang Y., Muir T.W., & Rabinowitz J.D. Bisphosphoglycerate mutase controls serine pathway flux via 3-phosphoglycerate. Nat Chem Biol 2017;13:1081-1087. doi:10.1038/nchembio.2453
^ abcdefg Stipanuk M.H., Caudill M.A. Biochemical, physiological, and molecular aspects of human nutrition. 3rd Edition. Elsevier health sciences, 2012
^ Van Schaftingen E., and Hers H-G. Inhibition of fructose-1,6-bisphosphatase by fructose-2,6-bisphosphate. Proc Natl Acad Sci USA 1981;78(5):2861-2863. doi:10.1073/pnas.78.5.2861
^ Van Schaftingen E., Jett M-F., Hue L., and Hers H-G. Control of liver 6-phosphofructokinase by fructose 2,6-bisphosphate and other effectors. Proc Natl Acad Sci USA 1981;78(6):3483-3486. doi:10.1073/pnas.78.6.3483
Gluconeogenesis is a metabolic pathway that leads to the synthesis of glucose from pyruvate, the conjugate base of pyruvic acid, and other non-carbohydrate precursors, even in non-photosynthetic organisms.[1][6]
It occurs in all microorganisms, fungi, plants and animals, and the reactions are essentially the same, leading to the synthesis of one glucose molecule from two pyruvate molecules. Therefore, it is in essence glycolysis in reverse, which instead goes from glucose to pyruvate, and shares seven enzymes with it.[5]
Gluconeogenesis and Glycolysis Pathways
Glycogenolysis is quite distinct from gluconeogenesis: it does not lead to de novo production of glucose from non-carbohydrate precursors, as shown by its overall reaction:
Glycogen or (glucose)n → n glucose molecules
The following discussion will focus on gluconeogenesis that occurs in higher animals, and in particular in the liver of mammals.
Maintaining blood glucose within the normal range, 3.3 to 5.5 mmol/L (60 and 99 mg/dL), is essential because many cells and tissues depend, largely or entirely, on glucose to meet their ATP demands; examples are red blood cells, neurons, skeletal muscle working under low oxygen conditions, the medulla of the kidney, the testes, the lens and the cornea of the eye, and embryonic tissues. For example, glucose requirement of the brain is about 120 g/die that is equal to:
over 50 percent of the total body stores of the monosaccharide, about 210 g, of which 190 g are stored as liver and muscle glycogen, and 20 g are found in free form in body fluids;
about 75 percent of the daily glucose requirement, about 160 g.[1]
During fasting, as in between meals or overnight, the blood glucose levels are maintained within the normal range due to hepatic glycogenolysis, and to the release of fatty acids from adipose tissue and ketone bodies by the liver. Fatty acids and ketone bodies are preferably used by skeletal muscle, thus sparing glucose for cells and tissues that depend on it, primarily red blood cells and neurons. However, after about 18 hours of fasting or during intense and prolonged exercise, glycogen stores are depleted and may become insufficient. At that point, if no carbohydrates are ingested, gluconeogenesis becomes important.
And, the importance of gluconeogenesis is further emphasized by the fact that if the blood glucose levels fall below 2 mmol/L, unconsciousness occurs.
The excretion of pyruvate would lead to the loss of the ability to produce ATP through aerobic respiration, i.e. more than 10 molecules of ATP for each molecule of pyruvate oxidized.
Where does it occur?
In higher animals, gluconeogenesis occurs in the liver, kidney cortex and epithelial cells of the small intestine, that is, the enterocytes.[5]
Quantitatively, the liver is the major site of gluconeogenesis, accounting for about 90 percent of the synthesized glucose, followed by kidney cortex, with about 10 percent.[1] The key role of the liver is due to its size; in fact, on a wet weight basis, the kidney cortex produces more glucose than the liver.
In the kidney cortex, gluconeogenesis occurs in the cells of the proximal tubule, the part of the nephron immediately following the glomerulus. Much of the glucose produced in the kidney is used by the renal medulla, while the role of the kidney in maintaining blood glucose levels becomes more important during prolonged fasting and liver failure. It should, however, be emphasized that the kidney has no significant glycogen stores, unlike the liver, and contributes to maintaining blood glucose homeostasis only through gluconeogenesis and not through glycogenolysis.
Part of the gluconeogenesis pathway also occurs in the skeletal muscle, cardiac muscle, and brain, although at very low rate.
In adults, muscle is about 18 the weight of the liver; therefore, its de novo synthesis of glucose might have quantitative importance. However, the release of glucose into the circulation does not occur because these tissues, unlike liver, kidney cortex, and enterocytes, lack glucose 6-phosphatase (EC 3.1.3.9), the enzyme that catalyzes the last step of gluconeogenesis.[6] Therefore, the production of glucose 6-phosphate, including that from glycogenolysis, does not contribute to the maintenance of blood glucose levels, and only helps to restore glycogen stores, in the brain small and limited mostly to astrocytes. For these tissues, in particular for skeletal muscle due to its large mass, the contribution to blood glucose homeostasis results only from the small amount of glucose released in the reaction catalyzed by enzyme debranching (EC 3.2.1.33) of glycogenolysis.
With regard to the cellular localization, most of the reactions occur in the cytosol, some in the mitochondria, and the final step) within the endoplasmic reticulum cisternae.
Irreversible steps
As previously said, gluconeogenesis is in essence glycolysis in reverse. And, of the ten reactions that constitute gluconeogenesis, seven are shared with glycolysis; these reactions have a ΔG close to zero, therefore easily reversible. However, under intracellular conditions, the overall ΔG of glycolysis is about -63 kJ/mol (-15 kcal/mol) and of gluconeogenesis about -16 kJ/mol (-3.83 kcal/mol), namely, both the pathways are irreversible.[5]
The irreversibility of the glycolytic pathway is due to three strongly exergonic reactions, that cannot be used in gluconeogenesis, and listed below.
The phosphorylation of glucose to glucose 6-phosphate, catalyzed by hexokinase (EC 2.7.1.1) or glucokinase (EC 2.7.1.2).
ΔG = -33.4 kJ/mol (-8 kcal/mol)
ΔG°’ = -16.7 kJ/mol (-4 kcal/mol)
The phosphorylation of fructose 6-phosphate to fructose 1,6-bisphosphate, catalyzed by phosphofructokinase-1 or PFK-1 (EC 2.7.1.11)
ΔG = -22.2 kJ/mol (-5.3 kcal/mol)
ΔG°’ = -14.2 kJ/mol (-3.4 kcal/mol)
The conversion of phosphoenolpyruvate or PEP to pyruvate, catalyzed by pyruvate kinase (EC 2.7.1.40)
ΔG = -16.7 kJ/mol (-4.0 kcal/mol)
ΔG°’ = -31.4 kJ/mole (-7.5 kcal/mol)
In gluconeogenesis, these three steps are bypassed by enzymes that catalyze irreversible steps in the direction of glucose synthesis.[6] This ensures the irreversibility of the metabolic pathway.
Below, such reactions are analyzed.
From pyruvate to phosphoenolpyruvate
The first step of gluconeogenesis that bypasses an irreversible step of glycolysis, namely the reaction catalyzed by pyruvate kinase, is the conversion of pyruvate to phosphoenolpyruvate.[8]
Phosphoenolpyruvate is synthesized through two reactions catalyzed, in order, by the enzymes:
pyruvate carboxylase (EC 6.4.1.1);
phosphoenolpyruvate carboxykinase or PEP carboxykinase (EC 4.1.1.32).
Pyruvate → Oxaloacetate → Phosphoenolpyruvate
Pyruvate carboxylase catalyzes the carboxylation of pyruvate to oxaloacetate, with the consumption of one ATP. The enzyme requires the presence of magnesium or manganese ions
Pyruvate + HCO3–+ ATP → Oxaloacetate + ADP + Pi
The enzyme, discovered in 1960 by Merton Utter, is a mitochondrial protein composed of four identical subunits, each with catalytic activity. The subunits contain a biotin prosthetic group, covalently linked by amide bond to the ε-amino group of a lysine residue, that acts as a carrier of activated CO2 during the reaction. An allosteric binding site for acetyl-CoA is also present in each subunit.
It should be noted that the reaction catalyzed by pyruvate carboxylase, leading to the production of oxaloacetate, also provides intermediates for the citric acid cycle or Krebs cycle.
Phosphoenolpyruvate carboxykinase is present, approximately in the same amount, in mitochondria and cytosol of hepatocytes. The isoenzymes are encoded by separate nuclear genes.
The enzyme catalyzes the decarboxylation and phosphorylation of oxaloacetate to phosphoenolpyruvate, in a reaction in which GTP acts as a donor of high-energy phosphate. PEP carboxykinase requires the presence of both magnesium and manganese ions. The reaction is reversible under normal cellular conditions.
Oxaloacetate + GTP ⇄ PEP + CO2 + GDP
During this reaction, a CO2 molecule, the same molecule that is added to pyruvate in the reaction catalyzed by pyruvate carboxylase, is removed. Carboxylation-decarboxylation sequence is used to activate pyruvate, since decarboxylation of oxaloacetate facilitates, makes thermodynamically feasible, the formation of phosphoenolpyruvate.
More generally, carboxylation-decarboxylation sequence promotes reactions that would otherwise be strongly endergonic, and also occurs in the citric acid cycle, in the pentose phosphate pathway, also called the hexose monophosphate pathway, and in the synthesis of fatty acids.
The levels of PEP carboxykinase before birth are very low, while its activity increases several fold a few hours after delivery. This is the reason why gluconeogenesis is activated after birth.
The sum of the reactions catalyzed by pyruvate carboxylase and phosphoenolpyruvate carboxykinase is:
Pyruvate + ATP + GTP + HCO3– → PEP + ADP + GDP + Pi + CO2
ΔG°’ of the reaction is equal to 0.9 kJ/mol (0.2 kcal/mol), while standard free energy change associated with the formation of pyruvate from phosphoenolpyruvate by reversal of the pyruvate kinase reaction is + 31.4 kJ/mol (7.5 kcal/mol).
Although the ΔG°’ of the two steps leading to the formation of PEP from pyruvate is slightly positive, the actual free-energy change (ΔG), calculated from intracellular concentrations of the intermediates, is very negative, -25 kJ/mol (-6 kcal/mol). This is due to the fast consumption of phosphoenolpyruvate in other reactions, that maintains its concentration at very low levels. Therefore, under cellular conditions, the synthesis of PEP from pyruvate is irreversible.[5]
It is noteworthy that the metabolic pathway for the formation of phosphoenolpyruvate from pyruvate depends on the precursor: pyruvate or alanine, or lactate.
Phosphoenolpyruvate precursors: pyruvate or alanine
The bypass reactions described below predominate when alanine or pyruvate is the glucogenic precursor.
Pyruvate carboxylase is a mitochondrial enzyme, therefore pyruvate must be transported from the cytosol into the mitochondrial matrix. This is mediated by transporters located in the inner mitochondrial membrane, referred to as MPC1 and MPC2. These proteins, associating, form a hetero-oligomer that facilitates pyruvate transport.[4]
Pyruvate can also be produced from alanine in the mitochondrial matrix by transamination, in the reaction catalyzed by alanine aminotransferase (EC 2.6.1.2), while the amino group is then converted into urea through the urea cycle.
Conversion of Pyruvate and Alanine to Phosphoenolpyruvate
Since the enzymes involved in the later steps of gluconeogenesis, except glucose-6-phosphatase, are cytosolic, the oxaloacetate produced in the mitochondrial matrix is transported into the cytosol. However, there are no oxaloacetate transporters in the inner mitochondrial membrane. The transfer to the cytosol occurs as a result of its reduction to malate, that, on the contrary, can cross the inner mitochondrial membrane. The reaction is catalyzed by mitochondrial malate dehydrogenase (EC 1.1.1.37), an enzyme also involved in the citric acid cycle, where the reaction proceeds in the reverse direction. In the reaction NADH is oxidized to NAD+.
Oxaloacetate + NADH + H+ ⇄ Malate + NAD+
Although ΔG°’ of the reaction is highly positive, under physiological conditions, ΔG is close to zero, and the reaction is easily reversible.
Malate crosses the inner mitochondrial membrane through a component of the malate-aspartate shuttle, the malate-α-ketoglutarate transporter. Once in the cytosol, the malate is re-oxidized to oxaloacetate in the reaction catalyzed by cytosolic malate dehydrogenase. In this reaction NAD+ is reduced to NADH.
Malate + NAD+ → Oxaloacetate + NADH + H+
Note: Malate-aspartate shuttle is the most active shuttle for the transport of NADH-reducing equivalents from the cytosol into the mitochondria. It is found in mitochondria of liver, kidney, and heart.
The reaction enables the transport into the cytosol of mitochondrial reducing equivalents in the form of NADH.[8] This transfer is needed for gluconeogenesis to proceed, as in the cytosolic the NADH, oxidized in the reaction catalyzed by glyceraldehydes 3-phosphate dehydrogenase (EC 1.2.1.12), is present in very low concentration, with a [NADH]/[NAD+] ratio equal to 8×10-4, about 100,000 times lower than that observed in the mitochondria.[5]
Finally, the oxaloacetate is converted to phosphoenolpyruvate in the reaction catalyzed by PEP carboxykinase.
Phosphoenolpyruvate precursor: lactate
Lactate is one of the major gluconeogenic precursors. It is produced for example by:
red blood cells, that are completely dependent on anaerobic glycolysis for ATP production;
skeletal muscle during intense exercise, that is, under low oxygen condition, when the rate of glycolysis exceeds the rate of the citric acid cycle and oxidative phosphorylation.[5]
When lactate is the gluconeogenic precursor, PEP synthesis occurs through a different pathway than that previously seen. In the hepatocyte cytosol NAD+ concentration is high and the lactate is oxidized to pyruvate in the reaction catalyzed by the liver isoenzyme of lactate dehydrogenase (EC 1.1.1.27). In the reaction NAD+ is reduced to NADH.
Lactate + NAD+ → Pyruvate + NADH + H+
The production of cytosolic NADH makes unnecessary the export of reducing equivalents from the mitochondria.
Pyruvate enters the mitochondrial matrix to be converted to oxaloacetate in the reaction catalyzed by pyruvate carboxylase. In the mitochondria, oxaloacetate is converted to phosphoenolpyruvate in the reaction catalyzed by mitochondrial pyruvate carboxylase. Phosphoenolpyruvate exits the mitochondria through an anion transporter located in the inner mitochondrial membrane, and, once in the cytosol, continues in the gluconeogenesis pathway.
Note: the synthesis of glucose from lactate may be considered as the part of the Cori cycle that takes place in the liver.
From fructose 1,6-bisphosphate to fructose 6-phosphate
The second step of gluconeogenesis that bypasses an irreversible step of the glycolytic pathway, namely the reaction catalyzed by PFK-1, is the dephosphorylation of fructose 1,6-bisphosphate to fructose 6-phosphate.
This reaction, catalyzed by fructose 1,6-bisphosphatase or FBPasi-1 (EC 3.1.3.11), a Mg2+ dependent enzyme located in the cytosol, leads to the hydrolysis of the C-1 phosphate of fructose 1,6-bisphosphate, without production of ATP.
Fructose 1,6-bisphosphate + H2O → Fructose 6-phosphate + Pi
The ΔG°’ of the reaction is -16.3 kJ/mol (-3.9 kcal/mol), therefore an irreversible reaction.
From glucose 6-phosphate to glucose
The third step of gluconeogenesis that bypasses an irreversible step of the glycolytic pathway, namely the reaction catalyzed by hexokinase or glucokinase, is the dephosphorylation of glucose 6-phosphate to glucose.
This reaction is catalyzed by the catalytic subunit of glucose 6-phosphatase, a protein complex located in the membrane of the endoplasmic reticulum of hepatocytes, enterocytes and cells of the proximal tubule of the kidney.[8] Glucose 6-phosphatase complex is composed of a glucose 6-phosphatase catalytic subunit and a glucose 6-phosphate transporter called glucose 6-phosphate translocase or T1.
Glucose 6-phosphatase catalytic subunit has the active site on the luminal side of the organelle. This means that the enzyme catalyzes the release of glucose not in the cytosol but in the lumen of the endoplasmic reticulum.
Glucose 6-phosphate, both resulting from gluconeogenesis, produced in the reaction catalyzed by glucose 6-phosphate isomerase or phosphoglucose isomerase (EC 5.3.1.9), and glycogenolysis, produced in the reaction catalyzed by phosphoglucomutase (EC 5.4.2.2), is located in the cytosol, and must enter the lumen of the endoplasmic reticulum to be dephosphorylated. Its transport is mediated by glucose-6-phosphate translocase.
The catalytic subunit of glucose 6-phosphatase, a Mg2+-dependent enzyme, catalyzes the last step of both gluconeogenesis and glycogenolysis. And, like the reaction catalyzed by fructose 1,6-bisphosphatase, this reaction leads to the hydrolysis of a phosphate ester.
Glucose 6-phosphate + H2O → Glucose + Pi
It should also be underlined that, due to orientation of the active site, the cell separates this enzymatic activity from the cytosol, thus avoiding that glycolysis, that occurs in the cytosol, is aborted by enzyme action on glucose 6-phosphate.
The ΔG°’ of the reaction is -13.8 kJ/mol (-3.3 kcal/mol), therefore it is an irreversible reaction. If instead the reaction were that catalyzed by hexokinase/glucokinase in reverse, it would require the transfer of a phosphate group from glucose 6-phosphate to ADP. Such a reaction would have a ΔG equal to +33.4 kJ/mol (+8 kcal/mol), and then strongly endergonic. Similar considerations can be made for the reaction catalyzed by FBPase-1.
Glucose and Pi group seem to be transported into the cytosol via different transporters, referred to as T2 and T3, the last one an anion transporter.
Finally, glucose leaves the hepatocyte via the membrane transporter GLUT2, enters the bloodstream and is transported to tissues that require it. Conversely, under physiological conditions, as previously said, glucose produced by the kidney is mainly used by the medulla of the kidney itself.
Gluconeogenesis: energetically expensive
Like glycolysis, much of the energy consumed is used in the irreversible steps of the process.[5]
Six high-energy phosphate bonds are consumed: two from GTP and four from ATP. Furthermore, two molecules of NADH are required for the reduction of two molecules of 1,3-bisphosphoglycerate in the reaction catalyzed by glyceraldehyde 3-phosphate dehydrogenase. The oxidation of NADH causes the lack of production of 5 molecules of ATP that are synthesized when the electrons of the reduced coenzyme are used in oxidative phosphorylation.
Also these energetic considerations show that gluconeogenesis is not simply glycolysis in reverse, in which case it would require the consumption of two molecules of ATP, as shown by the overall glycolytic equation.
At least in the liver, ATP needed for gluconeogenesis derives mostly from the oxidation of fatty acids or of the carbon skeletons of the amino acids, depending on the available “fuel”.
Coordinated regulation of gluconeogenesis and glycolysis
If glycolysis and gluconeogenesis were active simultaneously at a high rate in the same cell, the only products would be ATP consumption and heat production, in particular at the irreversible steps of the two pathways, and nothing more.
For example, considering PFK-1 and FBPasi-1:
ATP + Fructose 6-phosphate → ADP + Fructose 1,6-bisphosphate
Fructose 1,6-bisphosphate + H2O → Fructose 6-phosphate + Pi
The sum of the two reactions is:
ATP + H2O → ADP + Pi + Heat
Two reactions that run simultaneously in opposite directions result in a futile cycle or substrate cycle.
The modulation of the activity of involved enzymes occurs through:
allosterical mechanisms;
covalent modifications, such as phosphorylation and dephosphorylation;
changes in the concentration of the involved enzymes, due to changes in their synthesis/degradation ratio.
Allosteric mechanisms are very rapid and instantly reversible, taking place in milliseconds. The others, triggered by signals from outside the cell, such as hormones, like insulin, glucagon, or epinephrine, take place on a time scale of seconds or minutes, and, for changes in enzyme concentration, hours.
This allows a coordinated regulation of the two pathways, ensuring that when pyruvate enters gluconeogenesis, the flux of glucose through the glycolytic pathway slows down, and vice versa.
Regulation of gluconeogenesis
The regulation of gluconeogenesis and glycolysis involves the enzymes unique to each pathway, and not the common ones.[1]
While the major control points of glycolysis are the reactions catalyzed by PFK-1 and pyruvate kinase, the major control points of gluconeogenesis are the reactions catalyzed by fructose 1,6-bisphosphatase and pyruvate carboxylase.
The other two enzymes unique to gluconeogenesis, glucose-6-phosphatase and PEP carboxykinase, are regulated at transcriptional level.
Pyruvate carboxylase
In the mitochondrion, pyruvate can be converted to:
acetyl-CoA, in the reactions catalyzed by pyruvate dehydrogenase complex, reaction that connects glycolysis to the Krebs cycle;
oxaloacetate, in the reaction catalyzed by pyruvate carboxylase, to continue in the gluconeogenesis pathway.
The metabolic fate of pyruvate depends on the availability of acetyl-CoA, that is, by the availability of fatty acids in the mitochondrion.
When fatty acids are available, their beta-oxidation leads to the production of acetyl-CoA, that enters the Krebs cycle and leads to the production of GTP and NADH. When the energy needs of the cell are met, oxidative phosphorylation slows down, the [NADH]/[NAD+] ratio increases, NADH inhibits the citric acid cycle, and acetyl-CoA accumulates in the mitochondrial matrix.[1][5]
Acetyl-CoA is a positive allosteric effector of pyruvate carboxylase, and a negative allosteric effector of pyruvate kinase. Moreover, it inhibits pyruvate dehydrogenase complex both through feedback inhibition and phosphorylation through the activation of a specific kinase.
Fates for Pyruvate
This means that when the energy charge of the cell is high, the formation of acetyl-CoA from pyruvate slows down, while the conversion of pyruvate to glucose is stimulated. Therefore acetyl-CoA is a molecule that signals that additional glucose oxidation for energy is not required and that glucogenic precursors can be used for the synthesis and storage of glucose.
Conversely, when acetyl-CoA levels decrease, the activity of pyruvate kinase and of the pyruvate dehydrogenase complex increases, and therefore also the flow of metabolites through the citric acid cycle. This supplies energy to the cell.
Summarizing, when the energy charge of the cell is high pyruvate carboxylase is active, and that the first control point of gluconeogenesis determines what will be the fate of pyruvate in the mitochondria.
Fructose 1,6-bisphosphatase
The second major control point in gluconeogenesis is the reaction catalyzed by fructose 1,6-bisphosphatase. The enzyme is allosterically inhibited by AMP. Therefore, when AMP levels are high, and consequently ATP levels are low, gluconeogenesis slows down. This means that, as previously seen, FBPase-1 is active when the energy charge of the cell is sufficiently high to support de novo synthesis of glucose.
Conversely, PFK-1, the corresponding glycolytic enzyme, is allosterically activated by AMP and ADP and allosterically inhibited by ATP and citrate, the latter resulting from the condensation of acetyl-CoA and oxaloacetate. Therefore:
when AMP levels are high, gluconeogenesis slows down, and glycolysis accelerates;
when ATP levels are high or when acetyl-CoA or citrate are present in adequate concentrations, gluconeogenesis is promoted, while glycolysis slows down.
The increase in citrate levels indicates that the activity of the citric acid cycle can slow down; in this way, pyruvate can be used in glucose synthesis.
PFK-1, FBPase-1 and fructose 2,6-bisphosphate
The liver plays a key role in maintaining blood glucose homeostasis: this requires regulatory mechanisms that coordinate glucose consumption and production. Two hormones are mainly involved: glucagon and insulin.[6] They act intracellularly through fructose 2,6-bisphosphate or F2,6BP, an allosteric effector of PFK-1 and FBPase-1. This molecule is structurally related to fructose 1,6-bisphosphate, but is not an intermediate in glycolysis or gluconeogenesis.[1]
It was discovered in 1980 by Emile Van Schaftingen and Henri-Gery Hers, as a potent activator of PFK-1. In the subsequent year, the same researchers showed that it is also a potent inhibitor of FBPase-1.[9][10]
Fructose 2,6-bisphosphate, by binding to the allosteric site on PFK-1, reduces the affinity of the enzyme for ATP and citrate, allosteric inhibitors, and at the same time increases the affinity of the enzyme for fructose 6-phosphate, its substrate. PFK-1, in the absence of fructose 2,6-bisphosphate, and in the presence of physiological concentrations of ATP, fructose 6-phosphate, and of allosteric effectors AMP, ATP and citrate, is practically inactive. Conversely, the presence of fructose 2,6-bisphosphate activates PFK-1, thus stimulating glycolysis in the hepatocytes. At the same time fructose 2,6-bisphosphate slows down gluconeogenesis by inhibiting fructose 1,6-bisphosphatase, even in the absence of AMP. However, the effects of fructose-2,6-bisphosphate and AMP on FBPase-1 activity are synergistic.
F2,6BP: Regulation of Glycolysis and Gluconeogenesis
Fructose-2,6-bisphosphate concentration is regulated by the relative rates of synthesis and degradation. It is synthesized from fructose 6-phosphate in the reaction catalyzed by phosphofructokinase-2 or PFK-2 (EC 2.7.1.105), and is hydrolyzed to fructose 6-phosphate in the reaction catalyzed by fructose 2,6-bisphosphatase or FBPasi-2 (EC 3.1.3.46). These two enzymatic activities are located on a single bifunctional enzyme or tandem enzyme. In the liver, the balance of these two enzymatic activities is regulated by insulin and glucagon, as described below.
Glucagon
It is released into the circulation when blood glucose levels drop, signaling the liver to reduce glucose consumption for its own needs and to increase de novo synthesis of glucose and its release from glycogen stores.
After binding to specific membrane receptors, glucagon stimulates hepatic adenylate cyclase (EC 4.6.1.1) to synthesize 3′,5′-cyclic AMP or cAMP, that activates cAMP-dependent protein kinase or protein kinase A or PKA (EC 2.7.11.11). The kinase catalyzes the phosphorylation, at the expense of one molecule of ATP, of a specific serine residue (Ser32) of PFK-2/FBPase-2. As a result of the phosphorylation, phosphatase activity increases while kinase activity decreases. Such reduction, due to the increase in the Km for fructose 6-phosphate, causes a decrease in the levels of fructose 2,6-bisphosphate, that, in turn, inhibits glycolysis and stimulates gluconeogenesis. Therefore, in response to glucagon, hepatic production of glucose increases, enabling the organ to counteract the fall in blood glucose levels.
Note: glucagon, like adrenaline, stimulates gluconeogenesis also by increasing the availability of substrates such as glycerol and amino acids.
Insulin
After binding to specific membrane receptors, insulin activates a protein phosphatase, the phosphoprotein phosphatase 2A or PP2A, that catalyzes the removal of the phosphate group from PFK-2/FBPase-2, thus increasing PFK-2 activity and decreasing FBPase-2 activity. (At the same time, insulin also stimulates a cAMP phosphodiesterase that hydrolyzes cAMP to AMP). This increases the level of fructose 2,6-bisphosphate, that, in turn, inhibits gluconeogenesis and stimulates glycolysis.
In addition, fructose 6-phosphate allosterically inhibits FBPase-2, and activates PFK-2. It should be noted that the activities of PFK-2 and FBPase-2 are inhibited by their reaction products. However, the main effectors are the level of fructose 6-phosphate and the phosphorylation state of the enzyme.
Glucose 6-phosphatase
Unlike pyruvate carboxylase and fructose-1,6-bisphosphatase, the catalytic subunit of glucose-6-phosphatase is not subject to allosteric or covalent regulation. The modulation of its activity occurs at the transcriptional level.[7] Low blood glucose levels and glucagon, namely, factors that lead to increased glucose production, and glucocorticoids stimulate its synthesis, that, conversely, is inhibited by insulin.
Also, the Km for glucose 6-phosphate is significantly higher than the range of physiological concentrations of glucose 6-phosphate itself. This is why it is said that the activity of the enzyme is almost linearly dependent on the concentration of the substrate, that is, enzyme is controlled by the level of substrate.
PEP carboxykinase
The enzyme is regulated mainly at the level of synthesis and degradation. For example, high levels of glucagon or fasting increase protein production through the stabilization of its mRNA and the increase in its transcription rate. High blood glucose levels or insulin have opposite effects.
Xylulose 5-phosphate
Xylulose 5-phosphate, a product of the pentose phosphate pathway, is a recently discovered regulatory molecule. It stimulates glycolysis and inhibits gluconeogenesis by controlling the levels of fructose 2,6-bisphosphate in the liver.
When blood glucose levels increase, e.g. after a meal high in carbohydrates, the activation of glycolysis and hexose monophosphate pathway occurs in the liver. Xylulose 5-phosphate produced activates protein phosphatase 2A, that, as previously said, dephosphorylates PFK-2/FBPase-2, thus inhibiting FBPase-2 and stimulating PFK-2. This leads to an increase in the concentration of fructose 2,6-bisphosphate, and then to the inhibition of gluconeogenesis and stimulation of glycolysis, resulting in increased production of acetyl-CoA, the main substrate for the synthesis of lipids.[2] At the same time, an increase in flow through the hexose monophosphate shunt occurs, leading to the production of NADPH, a source of electrons for lipid synthesis. Finally, PP2A also dephosphorylates carbohydrate-responsive element-binding protein or ChREBP, a transcription factor that activates the expression of hepatic genes for lipid synthesis. Therefore, in response to an increase in blood glucose levels, lipid synthesis is stimulated.
It is therefore evident that xylulose 5-phosphate is a key regulator of carbohydrate and fat metabolism.
Precursors
Besides the aforementioned pyruvate, the major gluconeogenic precursors are lactate, glycerol, the majority of the amino acids, and, more generally, any compound that can be converted to pyruvate or oxaloacetate.[1]
Glycerol
Glycerol is released by lipolysis in adipose tissue. With the exception of propionyl-CoA, it is the only part of the lipid molecule that can be used for de novo synthesis of glucose in animals.
Glycerol enters gluconeogenesis, or glycolysis, depending on the cellular energy charge, as dihydroxyacetone phosphate or DHAP, whose synthesis occurs in two steps.
In the first step, glycerol is phosphorylated to glycerol 3-phosphate, in the reaction catalyzed by glycerol kinase (EC 2.7.1.30), with the consumption of one ATP.
Glycerol + ATP → Glycerol 3-phosphate + ADP + Pi
The enzyme is absent in adipocytes but present in the liver; this means that glycerol needs to reach the liver to be further metabolized.
Glycerol 3-phosphate is then oxidized to dihydroxyacetone phosphate, in the reaction catalyzed by glycerol 3-phosphate dehydrogenase (EC 1.1.1.8). In this reaction NAD+ is reduced to NADH.
During prolonged fasting, glycerol is the major gluconeogenic precursor, accounting for about 20 percent of glucose production.[3]
Glucogenic amino acids
Pyruvate and oxaloacetate are the entry points for the glucogenic amino acids, i.e. those whose carbon skeleton or part of it can be used for de novo synthesis of glucose.
Amino acids result from the catabolism of proteins, both food and endogenous proteins, like those of skeletal muscle during the fasting state or during intense and prolonged exercise.
The catabolic processes of each of the twenty amino acids that made up the proteins converge to form seven major products, acetyl-CoA, acetoacetyl-CoA, α-ketoglutarate, succinyl-CoA, fumarate, oxaloacetate, and pyruvate.
Except acetyl-CoA, acetoacetyl-CoA , the other five molecules can be used for gluconeogenesis. This means that gluconeogenic amino acids may also be defined as those whose carbon skeleton or part of it can be converted to one or more of the above molecules.
Below, the entry points of the gluconeogenic amino acids are shown.
Pyruvate: alanine, cysteine, glycine, serine, threonine and tryptophan.
Oxaloacetate: aspartate and asparagine.
α-Ketoglutarate: glutamate, arginine, glutamine, histidine and proline.
Succinyl-CoA: isoleucine, methionine, threonine and valine.
Fumarate: phenylalanine and tyrosine.
Glucogenic and Ketogenic Amino Acids
α-Ketoglutarate, succinyl-CoA and fumarate, intermediates of the citric acid cycle, enter the gluconeogenic pathway after conversion to oxaloacetate.
The utilization of the carbon skeletons of the amino acids requires the removal of the amino group. Alanine and glutamate, the key molecules in the transport of amino groups from extrahepatic tissues to the liver, are major glucogenic amino acids in mammals. Alanine is the main gluconeogenic substrate for the liver; this amino acid is shuttled to the liver from muscle and other peripheral tissues through the glucose-alanine cycle.
Ketogenic amino acids
Acetyl-CoA and acetoacetyl-CoA cannot be used for gluconeogenesis and are precursors of fatty acids and ketone bodies. The stoichiometry of the citric acid cycle elucidates why they cannot be used for de novo synthesis of glucose.
Acetyl-CoA, in the reaction catalyzed by citrate synthase, condenses with oxaloacetate to form citrate, a molecule with 6 carbon atoms instead of 4 as oxaloacetate. However, although the two carbon atoms from acetyl-CoA become part of the oxaloacetate molecule, two carbon atoms are oxidized and removed as CO2, in the reactions catalyzed by isocitrate dehydrogenase (EC 1.1.1.42) and α-ketoglutarate dehydrogenase complex. Therefore, acetyl-CoA does not yield any net carbon gain for the citric acid cycle.
Furthermore, the reaction leading to the formation of acetyl-CoA from pyruvate, catalyzed by the pyruvate dehydrogenase complex, that is the bridge between glycolysis and the Krebs cycle, is irreversible, and there is no other pathway to convert acetyl-CoA to pyruvate.
For this reason, amino acids whose catabolism produces acetyl-CoA and/or acetoacetyl-CoA, are termed ketogenic.
Only leucine and lysine are exclusively ketogenic.
Note: Plants, yeasts, and many bacteria can use acetyl-CoA for de novo synthesis of glucose as they do have the glyoxylate cycle. This cycle has four reactions in common with the citric acid cycle, two unique enzymes, isocitrate lyase (EC 4.1.3.1) and malate synthase (EC 2.3.3.9), but lacks the decarboxylation reactions. Therefore, organisms that have such pathway are able to use fatty acids for gluconeogenesis.
Five amino acids, isoleucine, phenylalanine, tyrosine, threonine and tryptophan, are both glucogenic and ketogenic, because part of their carbon backbone can be used for gluconeogenesis, while the other gives rise to ketone bodies.
Propionate
Propionate, which belongs to the group of short-chain fatty acids, is a gluconeogenic precursor because, as propionyl-CoA, the active molecule, can be converted to succinyl-CoA.
Below, the different sources of propionate are analyzed.
It may arise from beta-oxidation of odd-chain fatty acids such as margaric acid, a saturated fatty acid with 17 carbon atoms. Such fatty acids are rare compared to even-chain fatty acids, but present in significant amounts in the lipids of some marine organisms, ruminants, and plants. In the last pass through the β-oxidation sequence, the substrate is a five carbon fatty acid. This means that, once oxidized and cleaved to two fragments, it produces an acetyl-CoA and propionyl-CoA.
Another source is the oxidation of branched-chain fatty acids, with alkyl branches with an odd number of carbon atoms. An example is phytanic acid, produced in ruminants by oxidation of phytol, a breakdown product of chlorophyll.
In ruminants, propionate is also produced from glucose. Glucose is released from breakdown of cellulose by bacterial cellulase (EC 3.2.1.4) in the rumen, one of the four chambers that make up the stomach of these animals. These microorganisms then convert, through fermentation, glucose to propionate, which, once absorbed, may be used for gluconeogenesis, synthesis of fatty acids, or be oxidized for energy.[5]
In ruminants, in which gluconeogenesis tends to be a continuous process, propionate is the major gluconeogenic precursor.
Propionate may also result from the catabolism of valine, leucine, and isoleucine.
In the colon, propionate is produced by bacteria of the gut microbiota through anaerobic fermentation of non-digestible carbohydrates, such as resistant starch and fibers. Absorbed by the enterocytes, the unmetabolized portion passes into the portal circulation and reaches the liver where it can enter gluconeogenesis.
The oxidation of propionyl-CoA to succinyl-CoA involves three reactions that occur in the liver and other tissues.
In the first reaction, propionyl-CoA is carboxylated to D-methylmalonyl-CoA in the reaction catalyzed by propionyl-CoA carboxylase (EC 6.4.1.3), a biotin-requiring enzyme. This reaction consumes one ATP.
Propionyl-CoA + HCO3– + ATP → D-methylmalonyl-CoA+ ADP + Pi
In the subsequent reaction, catalyzed by methylmalonyl-CoA epimerase (EC 5.1.99.1), D-methylmalonyl-CoA is epimerized to its L-stereoisomer.
D-Methylmalonyl-CoA ⇄ L-Methylmalonyl-CoA
Finally, L-methylmalonyl-CoA undergoes an intramolecular rearrangement to succinyl-CoA, in the reaction catalyzed by methylmalonyl-CoA mutase (EC 5.4.99.2). This enzyme requires 5-deoxyadenosylcobalamin or coenzyme B12, a derivative of cobalamin or vitamin B12, as a coenzyme.
^ Kabashima T., Kawaguchi T., Wadzinski B.E., Uyeda K. Xylulose 5-phosphate mediates glucose-induced lipogenesis by xylulose 5-phosphate-activated protein phosphatase in rat liver. Proc Natl Acad Sci USA 2003;100:5107-5112. doi:10.1073/pnas.0730817100
^ Kuriyama H. et all. Coordinated regulation of fat-specific and liver-specific glycerol channels, aquaporin adipose and aquaporin 9. Diabetes 2002;51(10):2915-2921. 10.2337/diabetes.51.10.2915
^ McCommis K.S. and Finck B.N. Mitochondrial pyruvate transport: a historical perspective and future research directions. Biochem J 2015;466(3):443-454. doi:10.1042/BJ20141171
^ abcdefghi Nelson D.L., M. M. Cox M.M. Lehninger. Principles of biochemistry. 6th Edition. W.H. Freeman and Company, 2012
^ abcde Rosenthal M.D., Glew R.H. Medical biochemistry – Human metabolism in health and disease. John Wiley J. & Sons, Inc., Publication, 2009
^ Soty M., Chilloux J., Delalande F., Zitoun C., Bertile F., Mithieux G., and Gautier-Stein A. Post-Translational regulation of the glucose-6-phosphatase complex by cyclic adenosine monophosphate is a crucial determinant of endogenous glucose production and is controlled by the glucose-6-phosphate transporter. J Proteome Res 2016;15(4):1342-1349. doi:10.1021/acs.jproteome.6b00110
^ abc Stipanuk M.H., Caudill M.A. Biochemical, physiological, and molecular aspects of human nutrition. 3rd Edition. Elsevier health sciences, 2012
^ Van Schaftingen E., and Hers H-G. Inhibition of fructose-1,6-bisphosphatase by fructose-2,6-bisphosphate. Proc Natl Acad Sci USA 1981;78(5):2861-2863. doi:10.1073/pnas.78.5.2861
^ Van Schaftingen E., Jett M-F., Hue L., and Hers H-G. Control of liver 6-phosphofructokinase by fructose 2,6-bisphosphate and other effectors. Proc Natl Acad Sci USA 1981;78(6):3483-3486. doi:10.1073/pnas.78.6.3483
The glucose-alanine cycle, or Cahill cycle, proposed for the first time by Mallette, Exton and Park, and Felig et al. between 1969 and 1970, consists of a series of steps through which extrahepatic tissues, for example the skeletal muscle, export pyruvate, the conjugate base of pyruvic acid, and amino groups as alanine to the liver, and receive glucose from the liver via the bloodstream.[1][4]
The main steps of the glucose-alanine cycle are summarized below.[5][6]
When in extrahepatic tissues amino acids are used for energy, pyruvate, derived from glycolysis, is used as amino group acceptor, forming alanine, a nonessential amino acid.
Alanine diffuses into the bloodstream and reaches the liver.
In the liver, the amino group of alanine is transferred to alpha-ketoglutarate to form pyruvate and glutamate, respectively.
The amino group of glutamate mostly enters the urea cycle, and in part acts as a nitrogen donor in many biosynthetic pathways.
Pyruvate enters gluconeogenesis and is used for glucose synthesis.
The newly formed glucose diffuses into the bloodstream and reaches the peripheral tissues where, due to glycolysis, is converted into pyruvate that can accept amino groups from the free amino acids, thus closing the cycle.
Therefore, the glucose-alanine cycle provides a link between the metabolism of carbohydrates and amino acids, as schematically described below.[6]
Glucose → Pyruvate → Alanine → Pyruvate → Glucose
Glucose-Alanine Cycle
The glucose-alanine cycle occurs not only between the skeletal muscle, the first tissue in which it was observed, and the liver, but involves other cells and extrahepatic tissues including cells of the immune system, such as lymphoid organs.
The analysis of the steps of the glucose-alanine cycle is made considering the cycle between skeletal muscle and the liver.
Both intracellular and extracellular proteins are continuously hydrolyzed to the constituent amino acids and resynthesized, and the rate at which these processes occur is balanced precisely, thereby preventing loss of fat free mass.[3]
However, under catabolic conditions, such as intense and prolonged exercise or fasting, the rate of muscle protein breakdown exceeds synthesis. This leads to the liberation of amino acids, some of which are used for energy and others for gluconeogenesis. And the oxidation of the carbon skeletons of amino acids, in particular branched chain amino acids or BCAA, leucine, isoleucine and valine, may be a significant source of energy for the muscle. For example, after about 90 minutes of strenuous exercise, amino acid oxidation in muscle provides 10-15 percent of the energy needed for contraction.
The utilization of the carbon skeletons of amino acids for energy involves the removal of the amino group, and then the excretion of amino nitrogen in a non-toxic form.[5][6]
The removal of the alpha-amino group occurs by transamination, that can be summarized as follows:
alpha-Keto acid + Amino acid ⇄ New amino acid + New alpha-keto acid
Such reactions, catalyzed by enzymes called aminotransferases or transaminases (EC 2.6.1-) are freely reversible.
Branched chain amino acids, for example, transfer the amino group to alpha-ketoglutarate or 2-oxoglutaric acid, to form glutamate and the α-keto acid derived from the original amino acid, in a reaction catalyzed by branched chain aminotransferase or BCAT (EC 2.6.1.42).
The glucose-alanine cycle in skeletal muscle
In skeletal muscle, the newly formed glutamate may react with ammonia to form glutamine, for many tissues and organs, such as the brain, the major vehicle for interorgan transport of nitrogen.[7] The reaction is catalyzed by the cytosolic enzyme glutamine synthetase (EC 6.3.1.2), and consumes an ATP.
Glutamate + NH4+ + ATP → Glutamine + ADP + Pi
In this case, glutamate leaves the Cahill cycle.
Alternatively, and in contrast to what happens in most of the other tissues, the newly formed glutamate may transfer the amino group to pyruvate, derived from glycolysis, to form alanine and alpha-ketoglutarate. This transamination is catalyzed by alanine aminotransferase or ALT (EC 2.6.1.2), an enzyme found in most animal and plant tissues.
The alanine produced and that derived directly from protein breakdown, and muscle proteins are rich in alanine, can leave the cell and be carried by the bloodstream to the liver; in this way the amino group reaches the liver. And the rate at which alanine formed by transamination of pyruvate is transferred into the circulation is proportional to the intracellular pyruvate production.
Note: Alanine and glutamine are the major sources of nitrogen and carbon in interorgan amino acid metabolism.
The glucose-alanine cycle in the liver
Once in the liver, a hepatic alanine aminotransferase catalyzes a transamination in which alanine, the major gluconeogenic amino acid, acts as an amino group donor and alpha-ketoglutarate as an alpha-keto acid acceptor.[6] The products of the reaction are pyruvate, i.e. the carbon skeleton of alanine, and glutamate.
Glutamate, in the reaction catalyzed by glutamate dehydrogenase (EC 1.4.1.2), an enzyme present in the mitochondrial matrix, forms ammonium ion, which enters the urea cycle, and alpha-ketoglutarate, which can enter the Krebs cycle. This reaction is an anaplerotic reaction that links amino acid metabolism with the citric acid cycle.[6]
However, glutamate can also react with oxaloacetate to form aspartate and alpha-ketoglutarate, in a reaction catalyzed by aspartate aminotransferase (EC 2.6.1.1). Aspartate is involved in the formation of urea as well as in the synthesis of purines and pyrimidines.
Also the pyruvate produced may have different metabolic fates: it can be oxidized for ATP production, and then leave the glucose-alanine cycle, or enter the gluconeogenesis pathway, and thus continue in the cycle.
The glucose produced is released from the liver into the bloodstream and delivered to various tissues that require it, as the skeletal muscle, in which it is used for pyruvate synthesis. In turn, the newly formed pyruvate may react with glutamate, thus closing the cycle.
Transaminases
As previously mentioned, the removal of the amino group from amino acids occurs through transamination. These reactions are catalyzed by enzymes called aminotransferases or transaminases.[6]
They are cytosolic enzymes, present in all cells and particularly abundant in the liver, kidney, intestine and muscle; they require pyridoxal phosphate or PLP, the active form of vitamin B6 or pyridoxine, as a coenzyme, which is tightly bound to the active site.
In transamination reactions, the amino group of free amino acids, except of threonine and lysine, is channeled towards a small number of keto acids, notably pyruvate, oxaloacetate and alpha-ketoglutarate.
Cells contain different types of aminotransferases: many are specific for alpha-ketoglutarate as alpha-keto acid acceptor, but differ in specificity for the amino acid, from which they are named. Examples are the aforementioned alanine aminotransferase, also called alanine transaminase and glutamic pyruvic transferase or GPT, and aspartate aminotransferase or AST, also called glutamic-oxaloacetic transaminase or GOT.
It should be underlined that there is no net deamination in these reactions, no loss of amino groups, as the alpha-keto acid acceptor is aminated and the amino acid deaminated.
It transports nitrogen in a non-toxic form from peripheral tissues to the liver.
It transports pyruvate, a gluconeogenic substrate, to the liver.
It removes pyruvate from peripheral tissues. This leads to a higher production of ATP from glucose in these tissues. In fact, the NADH produced during glycolysis can enter the mitochondria and be oxidized through oxidative phosphorylation.
It allows to maintain a relatively high concentration of alanine in hepatocytes, sufficient to inhibit protein degradation.
It may play a role in host defense against infectious diseases.
Finally, it is important to underline that there is no net synthesis of glucose in the glucose-alanine cycle.
Energy cost
Like the Cori cycle, also the glucose-alanine cycle has an energy cost, equal to 3-5 ATP.
The part of the cycle that takes place in peripheral tissues involves the production of 5-7 ATP per molecule of glucose:
2 ATP are produced by glycolysis;
3-5 ATP derive from NADH/FADH2.
Instead in the liver, gluconeogenesis and the urea cycle cost 10 ATP:
6 ATP are consumed in the during gluconeogenesis per molecule of glucose synthesized;
4 ATP are consumed in the urea cycle per molecule of urea synthesized.
The glucose-alanine cycle, like the Cori cycle, shifts part of the metabolic burden from extrahepatic tissues to the liver.[6] However, the energy cost paid by the liver is justified by the advantages that the cycle brings to the whole body, as it allows, in particular conditions, an efficient breakdown of proteins in extrahepatic tissues, especially skeletal muscle, which in turn allows to obtain gluconeogenic substrates as well as the use of amino acids for energy in extrahepatic tissues.
Similarities and differences with the Cori cycle
There are some analogies between the two cycles, which are listed below.[6][7]
The Cahill cycle partially overlaps the Cori cycle when pyruvate is converted to glucose and the monosaccharide is transported to extrahepatic tissues, in which it is converted again to pyruvate via the glycolytic pathway.
The entry into gluconeogenesis pathway is similar for the two cycles: both alanine and lactate are converted to pyruvate.
Like the Cori cycle, the glucose-alanine cycle occurs between different cell types, unlike metabolic pathways such as glycolysis, citric acid cycle or gluconeogenesis that occur within individual cells
Cori cycle vs Glucose-Alanine Cycle
Below, some differences between the two cycles.
The main difference concerns the three carbon intermediate that from peripheral tissues reach the liver: lactate in the Cori cycle, and alanine in the glucose-alanine cycle.
Another difference concerns the fate of the NADH produced by glycolysis in peripheral tissues.
In the Cori cycle, the coenzyme acts as reducing agent to reduce pyruvate to lactate, in the reaction catalyzed by lactate dehydrogenase (EC 1.1.1.27).
In the glucose-alanine cycle, this reduction does not occur and the electrons of NADH can be transported into the mitochondria via the malate-aspartate and glycerol 3-phosphate shuttles, generating NADH, the first shuttle, and FADH2, the other shuttle. And the yield of ATP from NADH and FADH2 is 2.5 and 1.5, respectively.
Finally, from the previous point, it is clear that, unlike the Cori cycle, the Cahill cycle requires the presence of oxygen and mitochondria in the peripheral tissues.
References
^ Felig P., Pozefsk T., Marlis E., Cahill G.F. Alanine: key role in gluconeogenesis. Science 1970;167(3920):1003-1004. doi:10.1126/science.167.3920.1003
^ Gropper S.S., Smith J.L., Groff J.L. Advanced nutrition and human metabolism. Cengage Learning, 2009
^ Lecker S.H., Goldberg A.L. and Mitch W.E. Protein degradation by the ubiquitin–proteasome pathway in normal and disease states. J Am Soc Nephrol 2006;17(7):1807-1819.doi:10.1681/ASN.2006010083
^ Mallette L. E., Exton J.H., and Park C.R. Control of gluconeogenesis from amino acids in the perfused rat liver. J Biol Chem 1969;244(20):5713-5723. doi:10.1016/S0021-9258(18)63618-X
^ abc Moran L.A., Horton H.R., Scrimgeour K.G., Perry M.D. Principles of Biochemistry. 5th Edition. Pearson, 2012
^ abcdefgh Nelson D.L., Cox M.M. Lehninger. Principles of biochemistry. 6th Edition. W.H. Freeman and Company, 2012
^ abc Wu G. Amino acids: biochemistry and nutrition. CRC Press, 2010
The Cori cycle was discovered by Carl Ferdinand Cori and Gerty Theresa Radnitz, a husband-and-wife team, in the ‘30s and ‘40s of the last century. They demonstrated the existence of a metabolic cooperation between the skeletal muscle working under hypoxic conditions and the liver.[1]
The cycle allows the conversion of lactate, the conjugated base of lactic acid and its dominant form at physiological pH, into glucose, thus ensuring a continuous supply of the monosaccharide to peripheral tissues. The importance of this cycle is also demonstrated by the fact that it is responsible for about 40 percent of plasma glucose turnover.[7][8]
From a biochemical point of view, the Cori cycle links gluconeogenesis with anaerobic glycolysis, using different tissues to compartmentalize opposing metabolic pathways.[10]
This metabolic cooperation was demonstrated to exist also between other extrahepatic tissues and liver. Indeed, like the glucose-alanine cycle, the Cori cycle is active between the liver and all those tissues that do not completely oxidize glucose to CO2 and H2O.[11]
The analysis of the steps of the Cori cycle is made considering the lactate produced by red blood cells and muscle fibers.[12]
The cycle begins with the conversion of glucose to lactate, through anaerobic glycolysis and the subsequent action of lactate dehydrogenase (EC 1.1.1.27), which catalyzes the reduction of pyruvate, the conjugate base of pyruvic acid.
This is followed by the diffusion of lactate from the cell into the bloodstream, by which it is transported to the liver, its main user, and the renal cortex, in particular the proximal tubules, as these are another site where gluconeogenesis occurs.
In the liver and renal cortex, lactate is oxidized to pyruvate, in a reaction catalyzed by lactate dehydrogenase. Pyruvate is then converted to glucose by the gluconeogenic pathway.[8]
Finally, glucose diffuses into the bloodstream and reaches red blood cells or muscle fibers, closing the cycle.[10][11]
Red blood cells
Mature red blood cells, being devoid of a nucleus, ribosomes, and mitochondria, are smaller than most other cells. The small size allows them to pass through tiny capillaries, but the lack of mitochondria makes them completely dependent on anaerobic glycolysis for ATP production. This means that these cells continuously produce lactic acid.[8][10]
The availability of NAD+ is essential for glycolysis to proceed as well as for its rate. Indeed, the oxidized form of the coenzyme is required for the oxidation of glyceraldehyde 3-phosphate to 1,3-bisphosphoglycerate in the reaction catalyzed by glyceraldehyde 3-phosphate dehydrogenase (EC 1.2.1.12).
The accumulation of NADH is avoided by the reduction of pyruvate to lactate, in a reaction catalyzed by lactate dehydrogenase, where NADH acts as reducing agent, oxidizing to NAD+.[4]
Muscle fibers
Fast-twitch muscle fibers contain a reduced number of mitochondria and, under hypoxic conditions, such as during intense exercise, produce significant amounts of lactic acid.[8] In fact, in such conditions:
the rate of pyruvate production by glycolysis exceeds the rate of its oxidation by the citric acid cycle, so that less than 10 percent of the pyruvate enters the citric acid cycle;
the rate at which oxygen is taken up by the cells is not sufficient to allow aerobic oxidation of all the NADH produced.[3]
In such conditions, anaerobic glycolysis leads to the production of 2 ATP per molecule of glucose, 3 if the glucose comes from muscle glycogen, therefore, much lower than the 29-30 ATP produced by the complete oxidation of the monosaccharide.[9] However, the rate of ATP production by anaerobic glycolysis is greater than that produced by the complete oxidation of glucose.[10]
Finally, as in red blood cells, the reaction catalyzed by lactate dehydrogenase, regenerating NAD+, allows glycolysis to proceed, but produces lactate.[4]
Lactate
Lactic acid is an end product of metabolism that must be converted back into pyruvate to be used.[3]
The plasma membrane of most cells is freely permeable to both pyruvate and lactate, that can thus reach the bloodstream.[3] And, regarding for example the skeletal muscle, the amount of lactate that leaves the cell is greater than that of pyruvate due to the high NADH/NAD+ ratio in the cytosol and to the catalytic properties of LDH isozyme present in the muscle fibers.[5]
Once into the bloodstream, lactate reaches, among others, the liver and the renal cortex, where it is oxidized to pyruvate, in the reaction catalyzed by tissue-specific isozymes of lactate dehydrogenase.
In the hepatocyte, this oxidation is favored by the low NADH/NAD+ ratio in the cytosol. Then, pyruvate can enter gluconeogenesis, the next step of the Cori cycle, to be converted to glucose.[3]
Glucose enters into the bloodstream and is delivered to the muscle and red blood cells, thus closing the cycle. Obviously the monosaccharide also reaches all other tissues and cells that require it.
Energy cost
The Cori cycle results in a net consumption of 4 ATP.
The gluconeogenic leg of the cycle consumes 2 GTP and 4 ATP per molecule of glucose synthesized, namely, 6 ATP.
ATP-consuming reactions are catalyzed by:
pyruvate carboxylase (EC 6.4.1.1): one ATP;
phosphoenolpyruvate carboxykinase (EC 4.1.1.32): one GTP;
glyceraldehyde 3-phosphate dehydrogenase (EC 1.2.1.12): one ATP.
Since two molecules of lactate are required for the synthesis of one molecule of glucose, the net cost is 2 x 3 = 6 high energy bonds per molecule of glucose.[2]
Conversely, the glycolytic leg of the cycle produces only 2 ATP per molecule of glucose.
Therefore, more energy is required to produce glucose from lactate than that obtained by anaerobic glycolysis in extrahepatic tissues. This explains why the Cori cycle cannot be sustained indefinitely.
Is the Cori cycle a futile cycle?
The continuous breakdown and resynthesis of glucose feature of the Cori cycle might seem like a waste of energy. In reality, this cycle allows the effective functioning of many extrahepatic cells at the expense of the liver and the renal cortex. Therefore, it is more correct to define it as a substrate cycle rather than a futile cycle.[10][ROSENTHAL] Here below, some examples.
The Cori cycle allows to dispose of part of the lactic acid that red blood cells produce.[8]
Under intense exercise conditions, anaerobic glycolysis represents a primary means of ATP production for muscle fibers. But this could lead to an intracellular accumulation of lactate, and a consequent reduction in intracellular pH. Obviously, such accumulation does not occur, also thanks to the Cori cycle, in which the gluconeogenic tissues pay the cost of the disposal of a large part of the muscle lactate.[3] And the oxygen debt which occurs after an intense exercise is largely due to the increased oxygen demand of the hepatocytes, in which the oxidation of fatty acids, their main fuel, provides the ATP required for gluconeogenesis.[6][12]
During trauma, sepsis, burns, or after major surgery, an intense cell proliferation occurs in the wound, that is a hypoxic tissue, and in bone marrow. This in turn results in higher production of lactic acid, an increase in the flux through the Cori cycle, and an increase in ATP consumption in the liver, which, as mentioned, is supported by an increase in the oxidation of fatty acids. A similar condition seems to occur also in cancer patients with progressive weight loss.[2]
The Cori cycle is also important during overnight fasting and starvation.[6]
Cori cycle and glucose-alanine cycle
There are similarities and differences between the two cycles.
The glucose-alanine cycle and the Cori cycle are metabolic pathways that, through the intermediacy of the bloodstream, extend across different organs and help ensure a continuous supply of glucose to tissues.[12] In both cycles entry into gluconeogenesis involves the conversion of lactate and alanine into pyruvate. Finally, in both cycle, the glucose produced is then transported to peripheral tissues where the glycolytic pathway regenerates pyruvate.[13]
Their main difference lies instead in the three-carbon intermediate that is recycled: in the Cori cycle the carbon is returned to the liver as pyruvate, whereas in the glucose-alanine cycle it is returned to the liver as alanine.[8] Additionally, the metabolic fate of NADH also differs: in the Cori cycle it acts as a reducing agent in the reaction catalyzed by lactate dehydrogenase, whereas in the glucose-alanine cycle the electrons of NADH are used for the synthesis of ATP in the mitochondrion. Therefore, another difference is that the glucose-alanine cycle requires the presence of oxygen, while the Cori cycle does not.[8]
^ Gleeson T.T. Post-exercise lactate metabolism: a comparative review of sites, pathways, and regulation. Annu Rev Physiol 1996;58:565-81. doi:10.1146/annurev.ph.58.030196.003025
^ ab Moran L.A., Horton H.R., Scrimgeour K.G., Perry M.D. Principles of Biochemistry. 5th Edition. Pearson, 2012
Bile salts and bile acids are polar cholesterol derivatives, and represent the major route for the elimination of the steroid from the body.
They are molecules with similar but not identical structures, and diverse physical and biological characteristics.
They are synthesized in the liver, stored in the gallbladder, secreted into the duodenum, and finally, for the most part, reabsorbed in the ileum.
Because at physiological pH these molecules are present as anions, the terms bile acid and bile salts are used herein as synonyms.
Bile salts have similarities and differences with cholesterol molecule.
Like the steroid, they have a nucleus composed of four fused rings: three cyclohexane rings, labeled A, B and C, and a cyclopentane ring, labeled D. This structure is the perhydrocyclopentanophenanthrene, more commonly known as steroid nucleus.
Bile Acids and Their Conjugates
In higher vertebrates, they have 24 carbon atoms, as the side chain is three carbons shorter than the original. In lower vertebrates, bile acids have 25, 26, or 27 carbon atoms. The side chain ends with a carboxyl group, ionized at pH 7, that can be linked to the amino acid glycine or taurine.
In addition to the hydroxyl group at position 3, they have hydroxyl groups at positions 7 and/or 12.
All this makes them much more polar than cholesterol.
Since A and B rings are fused in cis configuration, the planar structure of the steroid nucleus is curved, and it is possible to identify:
a concave side, which is hydrophilic because the hydroxyl groups and the carboxyl group of the side chain, with or without the linked amino acid, are oriented towards it;
a convex side, which is hydrophobic because the methyl groups present at position 18 and 19 are orientated towards it.
Cholic Acid Structure
Therefore, having both polar and nonpolar groups, they are amphiphilic molecules and excellent surfactants. However, their chemical structure makes them different from many other surfactants, often composed of a polar head region and a nonpolar tail.
Primary, conjugated and secondary bile salts
Primary bile acids are those synthesized directly from cholesterol in the hepatocytes. In humans, the most important are cholic acid and chenodeoxycholic acid, which make up 80 percent of all bile acids. Before being secreted into the biliary tree, they are almost completely conjugated, up to 98 percent, with the glycine or taurine, to form glycoconjugates and tauroconjugates, respectively. In particular, approximately 75 percent of cholic acid and chenodeoxycholic acid are conjugated with glycine, to form glycocholic acid and glycochenodeoxycholic acid, the remaining 25 percent with taurine, to form taurocholic acid and taurochenodeoxycholic.
Synthesis of Conjugated Bile Acids
Conjugated bile acids are molecules with more hydrophilic groups than unconjugated bile acids, therefore they are much more effective emulsifiers. In fact, conjugation decreases the pKa of bile acids, from about 6, a value typical of non-conjugated molecules, to about 4 for glycocholic acid, and about 2 for taurocholic acid. This makes that conjugated bile acids are ionized in a broader range of pH to form the corresponding salts.
The hydrophilicity of the common acid and bile salts decreases in the following order: glycine-conjugated < taurine-conjugated < lithocholic acid < deoxycholic acid < chenodeoxycholic acid < cholic acid < ursodeoxycholic acid.
Finally, conjugation also decreases the cytotoxicity of primary bile acids.
Secondary bile acids are formed from primary bile acids which have not been reabsorbed from the small intestine. Once they reach the colon, they can undergo several modifications by bacteria of the gut microbiota, which is part of the larger human microbiota, to form secondary bile acids, which, along with short-chain fatty acids, are two major type of bacterial metabolites produced in the gut. They make up the remaining 20 percent of the body’s bile acid pool.
Another way of categorizing bile salts is based on their conjugation with glycine and taurine and their degree of hydroxylation. On this basis, three categories are identified.
Trihydroxy conjugates, such as taurocholic acid and glycocholic acid.
Dihydroxy conjugates, such as glycodeoxycholic acid, glycochenodeoxycholic acid, taurochenodeoxycholic acid, and taurodeoxycholic acid. They account for about 60 percent of bile salts present in the bile.
Unconjugated forms, such as cholic acid, deoxycholic acid, chenodeoxycholic acid, and lithocholic acid.
Function of bile acids
All their physiological functions are performed in the conjugated form.
They are the major route for the elimination of cholesterol from human body.
Indeed, humans do not have the enzymes to break open the cyclohexane rings or the cyclopentane ring of the steroid nucleus, nor to oxidize cholesterol to CO2 and water.
The other mechanism to eliminate the steroid from the body is as cholesterol per se in the bile.
Bile salts are strong surfactants. And in particular, di- and trihydroxy conjugates are the best surfactants among bile acids, much more effective than unconjugated counterparts, since they have more polar groups.
Once in contact with apolar lipids in the lumen of the small intestine, the convex apolar surface interacts with the apolar lipids, such as triglycerides, cholesterol esters, and ester of fat-soluble vitamins, whereas the concave polar surface interacts with the surrounding aqueous medium. This increases the dispersion of apolar lipids in the aqueous medium, as it allows the formation of tiny lipid droplets, increasing the surface area for:
lipase activity, mainly pancreatic lipase, (bile salts also play a direct role in the activation of this enzyme);
intestinal esterase activity.
Subsequently, they facilitate lipid absorption, as well as absorption of fat soluble vitamins by the intestinal mucosa, thanks to the formation of mixed micelles.
Bile acids perform a similar function in the gallbladder where, forming mixed micelles with phospholipids, they prevent the precipitation of cholesterol.
Note: as a consequence of the arrangement of polar and nonpolar groups, bile acids form micelles in aqueous solution, usually made up of less than 10 monomers, as long as their concentration is above the so-called critical micellar concentration or CMC.
At the intestinal level, they modulate the secretion of pancreatic enzymes and cholecystokinin.
In the small and large intestine, they have a potent antimicrobial activity, mainly deoxycholic acid, in particular against Gram-positive bacteria. This activity may be due to oxidative DNA damage, and/or to the damage of the cell membrane. Therefore, they play an important role in the prevention of bacterial overgrowth, but also in the regulation of gut microbiota composition.
In the last few years, it becomes apparent their regulatory role in the control of energy metabolism, and in particular for the hepatic glucose handling.
Enterohepatic circulation
After fat intake, enteroendocrine cells of the duodenum secrete cholecystokinin into the blood stream. Hormone binding to receptors on smooth muscle cells of the gallbladder promotes their contraction; the hormone also causes the relaxation of the sphincter of Oddi. All this results in the secretion of the bile, and therefore of bile acids into the duodenum.
Under physiological conditions, human bile salt pool is constant, and equal to about 3-5 g. This is made possible by two processes:
their intestinal reabsorption;
their de novo synthesis.
Up to 95 percent of the secreted bile salts is reabsorbed from the gut, not together with the products of lipid digestion, but through a process called enterohepatic circulation.
It is an extremely efficient recycling system, which seems to occur at least two times for each meal, and includes the liver, the biliary tree, the small intestine, the colon, and the portal circulation through which reabsorbed molecules return to the liver. Such recirculation is necessary since liver’s capacity to synthesize bile acids is limited and insufficient to satisfy intestinal needs if the bile salts were excreted in the feces in high amounts.
Most of the bile salts are reabsorbed into the distal ileum, the lower part of the small intestine, by a sodium-dependent transporter within the brush border of the enterocytes, called sodium-dependent bile acid transporter or ASBT, which carries out the cotransport of a molecule of bile acid and two sodium ions.
Within the enterocyte, it is thought that bile acids are transported across the cytosol to the basolateral membrane by the ileal bile acid-binding protein or IBABP. They cross the basolateral membrane by the organic solute transporter alpha-beta or OSTalpha/OSTbeta, pass into the portal circulation, and, bound to albumin, reach the liver.
It should be noted that a small percentage of bile acids reach the liver through the hepatic artery.
A hepatic level, their extraction is very efficient, with a first-pass extraction fraction ranging from 50 to 90 percent, a percentage that depends on bile acid structure. The uptake of conjugated bile acids is mainly mediated by a Na+-dependent active transport system, that is, the sodium-dependent taurocholate cotransporting polypeptide or NTCP. However, a sodium-independent uptake can also occur, carried out by proteins of the family of organic anion transporting polypeptides or OATP, mainly OATP1B1 and OATP1B3.
The rate limiting step in the enterohepatic circulation is their canalicular secretion, largely mediated by the bile salt export pump or BSEP, in an ATP-dependent process. This pump carries monoanionic bile salts, which are the most abundant. Bile acids conjugated with glucuronic acid or sulfate, which are dianionic, are transported by different carriers, such as MRP2 and BCRP.
Note: Serum levels of bile acids vary on the basis of the rate of their reabsorption, and therefore they are higher during meals, when the enterohepatic circulation is more active.
Intestinal metabolism
Bile acids which escape ileal absorption pass into the colon where they partly undergo modifications by intestinal microbiota and are converted to secondary bile acids.
The main reactions are listed below.
Deconjugation
On the side chain, hydrolysis of the C24 N-acyl amide bond can occur, with release of unconjugated bile acids and glycine or taurine. This reaction is catalyzed by bacterial hydrolases present both in the small intestine and in the colon.
7alpha-Dehydroxylation
Quantitatively, it is the most important reaction, carried out by colonic bacterial dehydratases that remove the hydroxyl group at position 7 to form 7-deoxy bile acids. In particular, deoxycholic acid is formed from cholic acid, and lithocholic acid, a toxic secondary bile acid, from chenodeoxycholic acid.
It should be noted that 7alpha-dehydroxylation, unlike oxidation and epimerization, can only occur on unconjugated bile acids, and therefore, deconjugation is an essential prerequisite.
Oxidation and epimerization
They are reactions involving the hydroxyl groups at positions 3, 7 and 12, catalyzed by bacterial hydroxysteroid dehydrogenases. For example, ursodeoxycholic acid derives from the epimerization of chenodeoxycholic acid.
Intestinal Metabolism of Bile Acids
Some of the secondary bile acids are then reabsorbed from the colon and return to the liver. In the hepatocytes, they are reconjugated, if necessary, and resecreted. Those that are not reabsorbed, are excreted in the feces.
Whereas oxidations and deconjugations are carried out by a broad spectrum of anaerobic bacteria, 7alpha-dehydroxylations is carried out by a limited number of colonic anaerobes.
7alpha-Dehydroxylations and deconjugations increase the pKa of the bile acids, and therefore their hydrophobicity, allowing a certain degree of passive absorption across the colonic wall.
The increase of hydrophobicity is also associated with an increased toxicity of these molecules. And a high concentration of secondary bile acids in the bile, blood, and feces has been associated to the pathogenesis of colon cancer.
Soluble fibers and reabsorption
The reabsorption of bile salts can be reduced by chelating action of soluble fibers, such as those found in fresh fruits, legumes, oats and oat bran, which bind them, decreasing their uptake. In turn, this increases bile acid de novo synthesis, up-regulating the expression of the 7alpha-hydroxylase and sterol 12alpha-hydroxylase, and thereby reduces hepatocyte cholesterol concentration.
The depletion of hepatic cholesterol increases the expression of the LDL receptor, and thus reduces plasma concentration of LDL cholesterol. On the other hand, it also stimulates the synthesis of HMG-CoA reductase, the key enzyme in cholesterol biosynthesis.
Note: Some anti-cholesterol drugs act by binding bile acids in the intestine, thereby preventing their reabsorption.
Synthesis
Quantitatively, bile acids are the major product of cholesterol metabolism.
As previously said, enterohepatic circulation and their de novo synthesis maintain a constant bile acid pool size. In particular, de novo synthesis allows the replacement of bile salts excreted in the faces, about 5-10 percent of the body pool, namely ~ 0.5 g/day.
Below, the synthesis of cholic acid and chenodeoxycholic acid, and their conjugation with the amino acids taurine and glycine, is described.
There are two main pathways for bile acid synthesis: the classical pathway and the alternative pathway. In addition, some other minor pathways will also be described.
De Novo Synthesis of Primary Bile Acids and Their Conjugates
The classical or neutral pathway
In humans, up to 90 percent of bile salts are produced via the classical pathway, also referred to as “neutral” pathway since intermediates are neutral molecules.
It is a metabolic pathway present only in the liver, that consists of reactions catalyzed by enzymes localized in the cytosol, endoplasmic reticulum, peroxisomes, and mitochondria, and whose end products are the conjugates of cholic acid and chenodeoxycholic acid.
The first reaction is the hydroxylation at position 7 of cholesterol, to form 7alpha-hydroxycholesterol. The reaction is catalyzed by cholesterol 7alpha-hydroxylase or CYP7A1 (E.C. 1.14.14.23). It is an enzyme localized in the endoplasmic reticulum, and catalyzes the rate-limiting step of the pathway.
7alpha-Hydroxycholesterol undergoes oxidation of the 3beta-hydroxyl group and the shift of the double bond from the 5,6 position to the 4,5 position, to form 7alpha-hydroxy-4-cholesten-3-one. The reaction is catalyzed by 3beta-hydroxy-Δ5-C27-steroid oxidoreductase or HSD3B7 (E.C. 1.1.1.181), an enzyme localized in the endoplasmic reticulum.
7alpha-Hydroxy-4-cholesten-3-one can follow two routes:
to enter the pathway that leads to the synthesis of cholic acid, through the reaction catalyzed by 7alpha-hydroxy-4-cholesten-3-one12alpha-monooxygenase or sterol 12alpha-hydroxylase or CYP8B1 (E.C. 1.14.18.8), an enzyme localized in the endoplasmic reticulum;
to enter the pathway that leads the synthesis of chenodeoxycholic acid, through the reaction catalyzed by 3-oxo-Δ4-steroid 5beta-reductase or AKR1D1 (E.C. 1.3.1.3), a cytosolic enzyme.
It should be underlined that the activity of sterol 12alpha-hydroxylase determines the ratio of cholic acid to chenodeoxycholic acid, and, ultimately, the detergent capacity of bile acid pool. And in fact, the regulation of sterol 12alpha-hydroxylase gene transcription is one of the main regulatory step of the classical pathway.
Therefore, if 7alpha-hydroxy-4-cholesten-3-one proceeds via the reaction catalyzed by sterol 12alpha-hydroxylase, the following reactions will occur.
7alpha-Hydroxy-4-cholesten-3-one is hydroxylated at position 12 by sterol 12alpha-hydroxylase, to form 7alpha,12alpha-dihydroxy-4-cholesten-3-one.
7alpha,12alpha-Dihydroxy-4-cholesten-3-one undergoes reduction of the double bond at 4,5 position, in the reaction catalyzed by 3-oxo-Δ4-steroid5beta-reductase, to form 5beta-cholestan-7alpha,12alpha-diol-3-one.
5beta-Cholestan-7alpha,12alpha-diol-3-one undergoes reduction of the hydroxyl group at position 4, in the reaction catalyzed by 3 alpha-hydroxysteroid dehydrogenase or AKR1C4 (EC 1.1.1.213), a cytosolic enzyme, to form 5beta-cholestan-3 alpha,7alpha,12alpha-triol.
5beta-Cholestan-3alpha,7alpha,12alpha-triol undergoes oxidation of the side chain via three reactions catalyzed by sterol 27-hydroxylase or CYP27A1 (EC 1.14.15.15). It is a mitochondrial enzyme also present in extrahepatic tissues and macrophages, which introduces a hydroxyl group at position 27. The hydroxyl group is oxidized to aldehyde, and then to carboxylic acid, to form 3 alpha,7 alpha,12 alpha-trihydroxy-5 beta-cholestanoic acid.
3alpha,7alpha,12alpha-Trihydroxy-5beta-cholestanoic acid is activated to its coenzyme A ester, 3alpha,7alpha,12alpha-trihydroxy-5beta-cholestanoyl-CoA, in the reaction catalyzed by either very long chain acyl-CoA synthetase or VLCS (EC 6.2.1.-), or bile acid CoA synthetase or BACS (EC 6.2.1.7), both localized in the endoplasmic reticulum.
3alpha,7alpha,12alpha-Trihydroxy-5beta-cholestanoyl-CoA is transported to peroxisomes where it undergoes five successive reactions, each catalyzed by a different enzyme. In the last two reactions, the side chain is shortened to four carbon atoms, and finally cholylCoA is formed.
In the last step, the conjugation, via amide bond, of the carboxylic acid group of the side chain with the amino acid glycine or taurine occurs. The reaction is catalyzed by bile acid-CoA:amino acid N-acyltransferase or the BAAT (EC 2.3.1.65), which is predominantly localized in peroxisomes.
The reaction products are thus the conjugated bile acids: glycocholic acid and taurocholic acid.
If 7alpha-hydroxy-4-cholesten-3-one does not proceed via the reaction catalyzed by sterol 12alpha-hydroxylase, it enters the pathway that leads to the synthesis of chenodeoxycholic acid conjugates, through the reactions described below.
7alpha-Hydroxy-4-cholesten-3-one is converted to 7alpha-hydroxy-5beta-cholestan-3-one in the reaction catalyzed by 3-oxo-Δ4-steroid 5 beta-reductase.
7alpha-Hydroxy-5beta-cholestan-3-one is converted to 5beta-cholestan-3alpha,7alpha-diol in the reaction catalyzed by 3alpha-hydroxysteroid dehydrogenase.
Then, the conjugated bile acids glycochenodeoxycholic acid and taurochenodeoxycholic acid are formed by modifications similar to those seen for the conjugation of cholic acid, and catalyzed mostly by the same enzymes.
Note: Unconjugated bile acids formed in the intestine must reach the liver to be reconjugated.
The alternative or acidic pathway
It is prevalent in the fetus and neonate, whereas in adults it leads to the synthesis of less than 10 percent of the bile salts.
This pathway differs from the classical pathway in that:
the intermediate products are acidic molecules, from which the alternative name “acidic pathway”;
the oxidation of the side chain is followed by modifications of the steroid nucleus, and not vice versa;
the final products are conjugates of chenodeoxycholic acid.
The first step involves the conversion of cholesterol into 27-hydroxycholesterol in the reaction catalyzed by sterol 27-hydroxylase.
27-Hydroxycholesterol can follow two routes.
Route A
27-hydroxycholesterol is converted to 3beta-hydroxy-5-cholestenoic acid in a reaction catalyzed by sterol 27-hydroxylase.
3beta-Hydroxy-5-cholestenoic acid is hydroxylated at position 7 in the reaction catalyzed by oxysterol 7alpha-hydroxylase or CYP7B1 (EC 1.14.13.100), an enzyme localized in the endoplasmic reticulum, to form 3beta-7alpha-dihydroxy-5-colestenoic acid.
3beta-7alpha-Dihydroxy-5-cholestenoic acid is converted to 3-oxo-7alpha-hydroxy-4-cholestenoic acid, in the reaction catalyzed by 3beta-hydroxy-Δ5-C27-steroid oxidoreductase.
3-Oxo-7alpha-hydroxy-4-cholestenoic acid, as a result of side chain modifications, forms chenodeoxycholic acid, and then its conjugates.
Route B
27-Hydroxycholesterol is converted to 7alpha,27-dihydroxycholesterol in the reaction catalyzed by oxysterol 7alpha-hydroxylase and cholesterol 7alpha-hydroxylase.
7alpha,27-Dihydroxycholesterol is converted to 7alpha,26-dihydroxy-4-cholesten-3-one in the reaction catalyzed by 3beta-hydroxy-Δ5-C27-steroid oxidoreductase;
7alpha, 26-Dihydroxy-4-cholesten-3-one can be transformed directly to conjugates of chenodeoxycholic acid, or can be converted to 3-oxo-7alpha-hydroxy-4-colestenoic acid, and then undergo side chain modifications and other reactions that lead to the synthesis of the conjugates of chenodeoxycholic acid.
Minor pathways
There are also minor pathways that contribute to bile salt synthesis, although to a lesser extent than classical and alternative pathways.
For example:
A cholesterol 25-hydroxylase (EC 1.14.99.38) is expressed in the liver.
A cholesterol 24-hydroxylase or CYP46A1 (EC 1.14.14.25) is expressed in the brain, and therefore, although the organ cannot export cholesterol, it exports oxysterols.
A nonspecific 7alpha-hydroxylase has also been discovered. It is expressed in all tissues and appears to be involved in the generation of oxysterols, which may be transported to hepatocytes to be converted to chenodeoxycholic acid.
Additionally, sterol 27-hydroxylase is expressed in various tissues, and therefore its reaction products must be transported to the liver to be converted to bile salts.
Bile salts: regulation of synthesis
Regulation of bile acid synthesis occurs via a negative feedback mechanism, particularly on the expression of cholesterol 7alpha-hydroxylase and sterol 12alpha-hydroxylase.
When an excess of bile acids, both free and conjugated, occurs, these molecules bind to the nuclear receptor farnesoid X receptor or FRX, activating it: the most efficacious bile acid is chenodeoxycholic acid, while others, such as ursodeoxycholic acid, do not activate it.
FRX induces the expression of the transcriptional repressor small heterodimer partner or SHP, which in turn interacts with other transcription factors, such as liver receptor homolog-1 or LRH-1, and hepatocyte nuclear factor-4alpha or HNF-4alpha. These transcription factors bind to a sequence in the promoter region of 7alpha-hydroxylase and 12alpha-hydroxylase genes, region called bile acid response elements or BAREs, inhibiting their transcription.
One of the reasons why bile salt synthesis is tightly regulated is because many of their metabolites are toxic.
References
Chiang J.Y.L. Bile acids: regulation of synthesis. J Lipid Res 2009;50(10):1955-1966. doi:10.1194/jlr.R900010-JLR200
Gropper S.S., Smith J.L. Advanced nutrition and human metabolism. 6th Edition. Cengage Learning, 2012
Moghimipour E., Ameri A., and Handali S. Absorption-enhancing effects of bile salts. Molecules 2015;20(8); 14451-14473. doi:10.3390/molecules200814451
Monte M.J., Marin J.J.G., Antelo A., Vazquez-Tato J. Bile acids: Chemistry, physiology, and pathophysiology. World J Gastroenterol 2009;15(7):804-816. doi:10.3748/wjg.15.804
Rosenthal M.D., Glew R.H. Medical biochemistry – Human metabolism in health and disease. John Wiley J. & Sons, Inc., Publication, 2009
Sundaram S.S., Bove K.E., Lovell M.A. and Sokol R.J. Mechanisms of Disease: inborn errors of bile acid synthesis. Nat Clin Pract Gastroenterol Hepatol 2008;5(8):456-468. doi:10.1038/ncpgasthep1179
Alpha-amylase (EC 3.2.1.1) is an enzyme that catalyzes the hydrolysis of α-(1→4) glycosidic bonds present on the straight chains of amylose and amylopectin, the two major components of starch granules.[2]
It is widely distributed in animals, plants, fungi, in the Ascomycota and Basidiomycota phyla, bacteria, in the genus Bacillus, and Archaea. Its wide distribution suggests a central role in carbohydrate metabolism.[15]
In humans, alpha-amylase is encoded by two different genes, and is synthesized mainly in the parotid glands and exocrine pancreas.[9] In plants, the alpha-amylase gene family consists of six different members.[11]
In animals, it is involved in the first step of starch digestion, which leads to the release of the disaccharide maltose, the oligosaccharides maltotriose and alpha-limit dextrins, and small amounts of glucose.[5]
Since starch is the main dietary source of carbohydrates, and therefore of glucose and energy, the activity of alpha-amylase may influence postprandial blood glucose levels.[8]
Alpha-amylase is used in different industrial sectors, as well as in clinical diagnostics.[7]
In plants, and particularly in grasses such as wheat, barley and rice, it is crucial for seed germination and grain maturation.[11][17]
The first detection of an amylase activity dates back to 1811, by the Russian chemist Konstantin Sigizmundovich Kirchhoff, who, working with wheat, discovered an enzymatic activity able to broke down starch.[16]
In 1831, the German chemist Erhard Friedrich Leuchs observed the degradation of starch when mixed with human saliva. Leuchs named ptyalin the agent responsible for such degradation. A few years later, in 1833, Anselme Payen and Jean-François Persoz, French chemists who worked in a sugar factory, isolated from barley an enzyme able to degrade starch, and named it diastase.[5]
The enzyme was named alpha-amylase by Richard Kuhn, in 1925, since he discovered that the hydrolysis products were in the alpha configuration.[13]
In 1894, alpha-amylase was the first enzyme to be produced industrially, extracted from a fungal source, and used to treat digestive disorders. Later, starting from 1917, it was also extracted from Bacillus subtilis and Bacillus mesentericus. Currently, it is mainly produced by genetically modified organisms.[15]
Alpha-amylase, and more generally amylases, of which two other types are known, beta-amylase (EC 3.2.1.2) e la gamma-amylase (EC 3.2.1.3), are widely utilized in different industrial sectors, such as textiles, detergent, feed, food, the production of beer and whiskey, as well as cosmetics, pharmaceuticals, and clinical diagnostics.[7]
Alpha-amylases can be classified, based on the degree of hydrolysis of the substrate, as liquifying, if they cleave 30-40 percent of starch glycosidic bonds, and saccharifying, if they cleave 50-60 percent of the bonds.[15]
Human alpha-amylase genes
In many mammals, including humans, alpha-amylase is encoded by two distinct but closely related genes, present on the short arm of chromosome 1, and named:
AMY1, which encodes for the protein present in saliva, in mammary gland and in some tumors, such as lung tumor;[5]
AMY2, which encodes for the enzyme produced by the pancreas. AMY2 is present in two forms, AMY2A and AMY2B, whose copy number differs in different populations, although two copies per gene seems to be a frequent pattern. It also appears that copy numbers of AMY2 may be associated with those of AMY1.[9]
Activity of alpha-amylase
Alpha-amylase catalyze the first step in starch digestion, namely, the hydrolysis of α-(1→4) glycosidic bonds within the linear chains of amylose and amylopectin, acting at random location along the α-glucans. The enzyme is therefore an endoglycosidases, has an optimal pH of about 7, requires the presence of calcium ions, and leads to the release of disaccharides and oligosaccharides in alpha configuration, in particular, maltose and maltotriose from amylose, and maltose, alpha-limiting dextrins and, in small amount, glucose from amylopectin.[2]
Alpha-amylase
Alpha-amylase, like other enzymes involved in starch metabolism such as pullulanase (EC 3.2.1.41), isoamylase (EC 3.2.1.68), starch branching enzymes (EC 2.4.1.18), and starch debranching enzymes, belongs to the glycoside hydrolase families 13 or GH-13 families, enzymes capable of acting on substrates containing alpha-glycosidic bonds.[10] Two enzymes involved in trehalose metabolism also belong to this family, trehalose synthase (EC 5.4.99.16), which catalyzes the interconversion of maltose and trehalose, and trehalose 6-phosphate hydrolase (EC 3.2.1.93), which hydrolyses trehalose 6-phosphate to yield a molecule of glucose and a molecule of glucose 6-phosphate.
Salivary alpha-amylase
Salivary alpha-amylase or ptyalin is mostly synthesized by the parotid glands, and to a lesser extent by submandibular glands. It is present in a glycosylated- and non-glycosylated form, and is involved in the first, limited, stage of starch digestion in the oral cavity. The enzyme is one of the most abundant salivary proteins, accounting for 40-50 percent of the total salivary proteins, and, with lingual lipase (EC 3.1.1.3), which is involved in lipid digestion, such as digestion of triglycerides and cholesterol esters, is one of the most important salivary enzymes.[3]
Pancreatic alpha-amylase
Pancreatic alpha-amylase is synthesized by the acinar cells of the pancreas.[12] It is one of the major proteins of the pancreatic juice, and, once in the duodenum, it attacks amylose and amylopectin, completing the first stage of the amylolysis. The amylolytic products are then attacked by the alpha-glucosidases of enterocyte brush border, namely, sucrase-isomaltase (EC 3.2.1.48) and maltase-glucoamylase (EC 3.2.1.20), which are also able to hydrolyze the α-(1→2) glycosidic bond of sucrose.[14]
Mammalian pancreatic alpha-amylase is able to bind to N-linked glycans of brush border membrane glycoproteins.[1] Such bond allows the cooperation with sucrase-isomaltase, cooperation that leads to a high increase in glucose production, up to 240 percent. However, it has been observed that glucose uptake by SGLT1, an intestinal transporter involved in the absorption of monosaccharides, is inhibited at high alpha-amylase concentrations in the intestinal lumen. It has been hypothesized that this inhibition may represent an innate regulatory mechanism to avoid sudden increases in glucose absorption and thus facilitate the regulation of postprandial blood glucose levels.[6]
Blood sugar control
Postprandial glycemic control is considered an effective strategy in preventing the development of type II diabetes. Among non-pharmacological methods there is the reduction of the ability of alpha-amylase to attack starch through a food processing leading to starch retrogradation, or by means of molecules present in foods which, with different modes of action, are able to reduce its activity.[8]
Food processing
Starch rich foods are subjected to hydrothermal treatments, such as cooking in boiling water, that cause starch granule gelatinization, namely, they absorb water, swell and lose most of their crystalline structures, which is replaced by amorphous structures, more easily attacked by alpha-amylases.[4] If gelatinized starch is allowed to cool, recrystallization can occur leading to give what is known as retrograded starch, a type of starch resistant to the action of alpha-amylase, which reaches the colon practically intact where it is fermented by the bacteria of the gut microbiota, like soluble fiber.
Role of polyphenols
The digestion of starch by alpha-amylase is affected by molecules present in foods, such as non-starch cell wall polysaccharides, proteins, lipids, and, among phytochemicals, various types of polyphenols.[4] These compounds can limit the action of alpha-amylase in at least four ways.[8]
Proteins and non-starch polysaccharides are able to limit the loss of the ordered structure of native starch during the hydrothermal processes to which the food is subjected during processing;
Lipids, polyphenols, proteins and non-starch polysaccharides are able to form ordered structures with starch.
Proteins, non-starch polysaccharides and polyphenols are able to form physical barriers that reduce the accessibility of hydrolytic enzymes to alpha-glucan chains.
Polyphenols and non-starch polysaccharides are able to reduce the activity of alpha-amylases and alpha-glucosidases.
Among polyphenols, flavonoids such as proanthocyanidins or condensed tannins seem to be important.[8] In order to be effective, the intake of foods rich in polyphenols, such as black tea or green tea, must be associated with a meal rich in carbohydrates. Gluten seems able to hinder the interaction between polyphenols and starch granules, and therefore reduce their inhibitory effect.
Amylase activity in clinical biochemistry
The assay of serum alpha-amylase activity is used diagnostic laboratory test.[7]
Increased serum levels of amylase activity may result from:
acute pancreatitis or a flare-up of chronic pancreatitis;
inflammation of the parotid glands;
obstruction of the secretory ducts of the parotid glands or the pancreatic duct.
Assays currently in use do not distinguish between salivary and pancreatic isoforms. However, if the increase in amylase activity is associated with an increase in lipase activity, the underlying pathology is probably of pancreatic origin.
Wheat alpha-amylase
In plants, and in particular in wheat, alpha-amylase plays a role in the germination and malting, which is a controlled germination that leads to the conversion of the grain into malt, which is also used for the production of whiskey and beer.[11]
In wheat grains, the enzyme is mainly synthesized in the early stages of germination, which are heterotrophic as they rely heavily on stored resources, and therefore also on the starch present in the endosperm.[17] The synthesis occurs in the aleurone layer, and initiates the mobilization of starch reserves.
Alpha-amylase is also involved in one of the main defects of wheat, pre-harvest germination.
References
^ Asanuma-Date K., Hirano Y., Le N., Sano K., Kawasaki N., Hashii N., Hiruta Y., Nakayama K., Umemura M., Ishikawa K., Sakagami H., and Ogawa H. Functional regulation of sugar assimilation by N-glycan-specific interaction of pancreatic α-amylase with glycoproteins of duodenal brush border membrane. J Biol Chem 2012;287(27):23104-18. doi:10.1074/jbc.m111.314658
^ ab Benjamin S., Smitha R.B., Jisha V.N., Pradeep S., Sajith S., Sreedevi S., Priji P., Unni K.N., Sarath Josh M.K. A monograph on amylases from Bacillus spp. Adv biosci biotechnol 2013;4:No.2. doi:10.4236/abb.2013.42032
^ Boehlke C., Zierau O., Hannig C. Salivary amylase – The enzyme of unspecialized euryphagous animals. Arch Oral Biol 2015;60(8):1162-76. doi:10.1016/j.archoralbio.2015.05.008
^ ab Butterworth P.J., Bajka B.H., Edwards C.H., Warren F.J., Ellis P.R. Enzyme kinetic approach for mechanistic insight and predictions of in vivo starch digestibility and the glycaemic index of foods. Trends Food Sci Technol 2022;120:254-264. doi:10.1016/j.tifs.2021.11.015
^ abc Butterworth P.J., Warren F.J. and Ellis P.R. Human α-amylase and starch digestion: an interesting marriage. Starch/Stärke 2011;63:395-405. doi:10.1002/star.201000150
^ Date K., Satoh A., Lida K., Ogawa H. Pancreatic α-amylase controls glucose assimilation by duodenal retrieval through N-glycan-specific binding, endocytosis, and degradation. J Biol Chem 2015;290(28):17439-50. doi:10.1074/jbc.M114.594937
abc de Souza P.M., de Oliveira Magalhães P. Application of microbial α-amylase in industry – A review. Braz J Microbiol 2010;41(4):850-61. doi:10.1590/S1517-83822010000400004
^ abcd Gong L., Feng D., Wang T., Ren Y., Liu Y., Wang J. Inhibitors of α-amylase and α-glucosidase: potential linkage for whole cereal foods on prevention of hyperglycemia. Food Sci Nutr 2020;8(12):6320-6337. doi:10.1002/fsn3.1987
^ ab Iafrate A.J., Feuk L., Rivera M.N., Listewnik M.L., Donahoe P.K., Qi Y., Scherer S.W, Lee C. Detection of large-scale variation in the human genome. Nature Genetics. 2004;36:949–951. doi:10.1038/ng1416
^ Janeček Š., Svensson B., MacGregor E.A. α-Amylase: an enzyme specificity found in various families of glycoside hydrolases. Cell Mol Life Sci 2014;71(7):1149-70. doi:10.1007/s00018-013-1388-z
^ abc Ju L., Pan Z., Zhang H., Li Q., Liang J., Deng G., Yu M., Long H. New insights into the origin and evolution of α-amylase genes in green plants. Sci Rep 2019;9(1):4929. doi:10.1038/s41598-019-41420-w
^ Keller P.J., Allan B.J. The protein composition of human pancreatic juice. J Biol Chem 196725;242(2):281-7. doi:10.1016/S0021-9258(19)81461-8
^ Kuhn R. Mechanism of the action of amylases. Constitution of starch. Justus Liebigs Annalen der Chemie 1925;443:1-71
^ Nichols B.L., Avery S., Sen P., Swallow D.M., Hahn D., Sterchi E. The maltase-glucoamylase gene: common ancestry to sucrase-isomaltase with complementary starch digestion activities. Proc Natl Acad Sci U S A. 2003 Feb 4;100(3):1432-7. doi:10.1073/pnas.0237170100
^ abc Tiwari S.P., Srivastava R., Singh C.S., Shukla K., Singh R.K., Singh P., Singh R., Singh N.L., and Sharma R. Amylases: an overview with special reference to alpha amylase. Journal of Global Biosciences 2015;4(SI1):1886-1901.
^ Zakowski J.J., Bruns D.E. Biochemistry of human alpha amylase isoenzymes. Crit Rev Clin Lab Sci 1985;21(4):283-322. doi:10.3109/10408368509165786
^ ab Zhang Q., Pritchard J., Mieog J., Byrne K., Colgrave M.L., Wang J.R., Ral J.F. Over-expression of a wheat late maturity alpha-amylase type 1 impact on starch properties during grain development and germination. Front Plant Sci 2022;13:811728. doi:10.3389/fpls.2022.811728
One of the main functions of the liver is to participate in the maintaining of blood glucose levels within well defined range (in the healthy state before meals 60-100 mg/dL or 3.33-5.56 mmol/L). To do it the liver releases glucose into the bloodstream in:
fasting state;
between meals;
during physical activity.
Blood glucose levels and hepatic glucose 6-phosphatase
In the liver, glycogen is the storage form of glucose which is released from the molecule not as such, but in the phosphorylated form i.e. with charge, the glucose 1-phosphate (this process is called glycogenolysis). The phosphorylated molecule can’t freely diffuse from the cell, but in the liver it is present the enzyme glucose-6-phosphatase that hydrolyzes glucose 6-phosphate, produced from glucose 1-phosphate in the reaction catalyzed by phosphoglucomutase, to glucose (an irreversible dephosphorylation).
Then, glucose can diffuse from the hepatocyte, via a transporter into the plasma membrane called GLUT2, into the bloodstream to be delivered to extra-hepatic cells, in primis neurons and red blood cells for which it is the main, and for red blood cells the only energy source (neurons, with the exception of those in some brain areas that can use only glucose as energy source, can derive energy from another source, the ketone bodies, which becomes predominant during periods of prolonged fasting).
Note: the liver obtains most of the energy required from the oxidation of fatty acids, a class of lipids, and not from glucose.
Glucose-6-phosphatase is present also in the kidney and gut but not in the muscle and brain; therefore in these tissues glucose-6-phosphate can’t be released from the cell.
Glucose 6-phosphatase plays an important role also in gluconeogenesis.
Glucose 6-phosphatase is present into the membrane of endoplasmic reticulum and the hydrolysis of glucose-6-phosphate occurs into its lumen (therefore this reaction is separated from the process of glycolysis). The presence of a specific transporter, the glucose 6-phosphate translocase, is required to transport the phosphorylated molecule from citosol into the lumen of endoplasmic reticulum. Although a glucose transporter is present into the membrane of endoplasmic reticulum, most of the released glucose is not transported back into the cytosol of the cell but is secreted into the bloodstream. Finally, an ion transporter transports back into the cytosol the inorganic phosphate released into the endoplasmic reticulum.
References
Nelson D.L., M. M. Cox M.M. Lehninger. Principles of biochemistry. 6th Edition. W.H. Freeman and Company, 2012
Roach P.J., Depaoli-Roach A.A., Hurley T.D and Tagliabracci V.C. Glycogen and its metabolism: some new developments and old themes. Biochem J 2012;441:763-787. doi:10.1042/BJ20111416
Stipanuk M.H., Caudill M.A. Biochemical, physiological, and molecular aspects of human nutrition. 3rd Edition. Elsevier health sciences, 2012
When excess calories are consumed from carbohydrates or proteins, such surplus is used to synthesize fatty acids and then triacylglycerols, while it doesn’t occur if the excess come from fats.
De novo fatty acid synthesis in plants and animals
De novofatty acid synthesis is largely similar among plants and animals.
It occurs in chloroplasts of photosynthetic cells of higher plants, and in cytosol of animal cells by the concerted action of two enzymes: acetyl-CoA carboxylase (EC 6.4.1.2) and fatty acid synthase (EC 2.3.1.85).
Fatty acid synthase catalyzes a repeating four-step sequence by which the fatty acyl chain is extended by two carbons, at the carboxyl end, every each passage through the cycle; this four-step process is the same in all organisms.
In animals, the primary site for lipid metabolism is liver, not the adipose tissue. However, adipose tissue is a major organ system in which fatty acid synthesis occurs, though in humans it is less active than in many other animal species.
Although myristic acid, lauric acid and a trace of stearic acid may also be produced, in animals and plants the main product of these reactions is palmitic acid.
It should be noted that in certain plants, such as palm and coconut, chain termination occurs earlier than palmitic acid release: up to 90% of the fatty acids produced and then present in the oils of these plants are between 8 (caprylic acid) and 14 (myristic acid) carbons long (palmitic acid: 16 carbon atoms).
Synthesis of long chain saturated and unsaturated fatty acids
Palmitic acid is the commonest saturated fatty acid in plant and animal lipids, but generally it is not present in very large proportions because it may be undergo into several metabolic pathways.
In fact:
it is the precursor of stearic acid;
it may be desaturated (insertion of a double bond into fatty acid chain) to palmitoleic acid, the precursor of all fatty acids of omega-7 or n-7 family, in a reaction catalyzed by Δ9-desaturase (EC 1.14.19.1), an ubiquitous enzyme in both plant and animal kingdoms and the most active lipid enzyme in mammalian tissues, the same enzyme that catalyzes the desaturation of stearic acid to oleic acid.
Note: Δ9- desaturase inserts double bounds in the 9-10 position of the fatty acid carbon chain, position numbered from the carboxyl end of the molecule, and:
if the substrate is palmitic acid, the double bond is inserted between n-7 and n-8 position of the chain (in this case numbered from the methyl end of the molecule), so producing palmitoleic acid, the founder of omega-7 series;
if the substrate is stearic acid, the double bond is inserted between n-9 and n-10 position of the chain and oleic acid will be produced.
It may be esterified into complex lipids.
Of course, in plants and animals there are fatty acids longer and/or more unsaturated than these just seen thanks to modification systems (again desaturation and elongation) that catalyze reactions of fatty acid synthesis that are organism- tissue- and cell- specific.
For example, stearic acid may be:
elongated to arachidic acid, behenic acid and lignoceric acid, all saturated fatty acids, in reactions catalyzed by elongases. Again, chain elongation occurs, both in mitochondria and in the smooth endoplasmic reticulum, by the addition of two carbon atom units at a time at the carboxylic end of the fatty acid through the action of fatty acid elongation systems (particularly long and very long saturated fatty acids, from 18 to 24 carbon atoms, are synthesized only on cytosolic face of the smooth endoplasmic reticulum);
desaturated, as seen, to oleic acid, an omega-9 or n-9 fatty acid, in a reaction catalyzed by Δ9-desaturase.
Several researchers have postulated that the reason for which stearic acid does not concur to hypercholesterolemia is its rapid conversion to oleic acid.
Oleic acid is the start point for the synthesis of many other unsaturated fatty acids by reactions of elongation and/or desaturation.
In fact:
gadoleic acid, erucic acid and nervonic acid, all monounsaturated fatty acids.
Saturated fatty acids, and unsaturated fatty acids of the omega-9 series, usually oleic acid (but also palmitoleic acid and other omega-7 fatty acids) are the only fatty acids produced de novo in mammal systems.
Thanks to the consecutive action of the enzymes Δ12-desaturase (1.14.19.6) and Δ15-desaturase (EC 1.14.19.25), that insert a double bond respectively in the 12-13 and 15-16 position of the carbon chain of the fatty acid, oleic acid is converted first to linoleic acid, founder of all the omega-6 polyunsaturated fatty acids, and then to alpha-linolenic acid, founder of all the omega-3 polyunsaturated fatty acids (omega-3 and omega-6 PUFA that will be produced from these precursors through repeating reactions of elongation and desaturation).
In the case of essential fatty acid (EFA) deficiency, oleic acid may be desaturated and elongated to omega-9 polyunsaturated fatty acids, with accumulation especially of Mead acid.
Omega-3 and omega-6 PUFA synthesis
Animal tissues can desaturate fatty acids in the 9-10 position of the chain, thanks to the presence of Δ9 desaturase; as previously seen, if the substrate of the reaction is palmitic acid, the double bond will appear between n-7 and n-8 position, with stearic acid between n-9 and n-10 position, so leading to formation respectively of palmitoleic acid and oleic acid.
Animals lack Δ12- and Δ15-desaturases, enzymes able to desaturate carbon carbon bonds beyond the 9-10 position of the chain. For these reason, they can’t produce de novo omega-3 and omega-6 PUFA (which have double bonds also beyond the 9-10 position), that are so essential fatty acids.
Δ12- and Δ15-desaturases are present in plants; though many land plants lack Δ15-desaturase, also called omega-3 desaturase, planktons and aquatic plants in colder water possess it and produce abundant amounts of the omega-3 fatty acids.
References
Akoh C.C. and Min D.B. “Food lipids: chemistry, nutrition, and biotechnology”. CRC Press Taylor & Francis Group, 2008 3th ed. 2008
Bender D.A. “Benders’ dictionary of nutrition and food technology”. 2006, 8th Edition. Woodhead Publishing. Oxford
Burr G.O. and Burr M.M. A new deficiency disease produced by the rigid exclusion of fat from the diet. Nutr Rev 1973;31(8):148-149. doi:10.1111/j.1753-4887.1973.tb06008.x
Chow Ching K. “Fatty acids in foods and their health implication”. 3rd Edition. CRC Press Taylor & Francis Group, 2008
Rosenthal M.D., Glew R.H. Mediacal biochemistry. Human metabolism in health and disease. John Wiley & Sons, Inc. 2009
Stipanuk M.H., Caudill M.A. Biochemical, physiological, and molecular aspects of human nutrition. 3rd Edition. Elsevier health sciences, 2012
Biochemistry and metabolism
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The pyruvic acid produced in the cytosol, by means of specific transporters, enters the mitochondrion where it can be used for energy, entering the citric acid cycle, and/or biosynthetic/anaplerotic purposes, depending on the needs of the cell.[15]
The C subunit forms a sixth isoform, a homotetramer, named as LDH-6 or C4. 
Keto acids, and in particular alpha-keto acids, are very important in biochemistry, being involved in many metabolic pathways.
The enzyme, like 
PTST1 appears to associate, in the plastid stoma, with GBSS. The complex binds to the nascent starch granule, the protein dissociates from the enzyme, that begins to catalyze the elongation of the malto-oligosaccharides, while the protein returns to the stroma where it recruits another GBSS.
The few branches are connected to the linear chain by α-(1→6) glycosidic bonds, such as in amylopectin and 
And one of the factors limiting the rate of an enzymatic reaction is just the frequency with which substrates and enzymes collide. The simplest way to increase the frequency of collisions would be to increase the concentration of substrates and enzymes. However, due to the large number of different reactions that take place in the cell, this route is not feasible. In other words, there is a limit to the concentration that substrates and enzymes can reach, concentrations that are in the micromolar range for substrates, and even lower for enzymes. Exceptions are the enzymes of 
Then, glucose can diffuse from the hepatocyte, via a transporter into the plasma membrane called GLUT2, into the bloodstream to be delivered to extra-hepatic cells, in primis neurons and red blood cells for which it is the main, and for red blood cells the only energy source (neurons, with the exception of those in some brain areas that can use only glucose as energy source, can derive energy from another source, the ketone bodies, which becomes predominant during periods of prolonged fasting).
