Gluconeogenesis

Gluconeogenesis: contents in brief

What is gluconeogenesis?

Gluconeogenesis is a metabolic pathway that leads to the synthesis of glucose from pyruvate and other non-carbohydrate precursors, even in non-photosynthetic organisms.
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.

Gluconeogenesis
Fig. 1 – Gluconeogenesis and Glycolysis

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.

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Why is gluconeogenesis important?

Gluconeogenesis is an essential metabolic pathway for at least two reasons.

  • It ensures the maintenance of appropriate blood glucose levels when the liver glycogen is almost depleted and no carbohydrates are ingested.
    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% of the total body stores of the monosaccharide, about 210 g, of which 190 g are stored as muscle and liver glycogen, and 20 g are found in free form in body fluids;
about 75% of the daily glucose requirement, about 160 g.

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.

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Where does gluconeogenesis occurs?

In higher animals, gluconeogenesis occurs in the liver, kidney cortex and epithelial cells of the small intestine, that is, the enterocytes.
Quantitatively, the liver is the major site of gluconeogenesis, accounting for about 90% of the synthesized glucose, followed by kidney cortex, with about 10%. 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 (see below).
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 (see above) within the endoplasmic reticulum cisternae.

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Irreversible steps of gluconeogenesis

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.
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: this ensures the irreversibility of the metabolic pathway.
Below, such reactions are analyzed.

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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.
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

Gluconeogenesis
Fig. 2 – 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.
It is noteworthy that the metabolic pathway for the formation of phosphoenolpyruvate from pyruvate depends on the precursor: pyruvate or alanine, or lactate.

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Phosphoenolpyruvate precursor: pyruvate or alanine

Gluconeogenesis
Fig. 3 – Conversion of Pyruvate to PEP

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.
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).
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. This transfer is needed for gluconeogenesis to proceed, as in the cytosolic the NADH, oxidized in the  reaction catalyzed by glyceraldehydes 3-phosphate dehydrogenase, 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.
Finally, the oxaloacetate is converted to phosphoenolpyruvate in the reaction catalyzed by PEP carboxykinase.

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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.

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 (see above).
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.

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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.

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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 (EC 2.7.1.1) or glucokinase (EC 2.7.1.2), 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. 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.

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Gluconeogenesis: energetically expensive

Like glycolysis, much of the energy consumed is used in the irreversible steps of the process.
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.

Glucose + 2 ADP + 2 Pi + 2 NAD+ → 2 Pyruvate + 2 ATP + 2 NADH + 2 H+ + 2 H2O

Below, the overall equation for gluconeogenesis:

2 Pyruvate + 4 ATP + 2 GTP + 2 NADH+ + 2 H+ + 4 H2O → Glucose + 4 ADP + 2 GDP + 6 Pi + 2 NAD+

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”.

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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 . These apparently uneconomical cycles allow to regulate opposite metabolic pathways. In fact, a substrate cycle involves different enzymes, at least two, whose activity can be regulated separately. A such regulation would not be possible if a single enzyme would operate in both directions. 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.

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Regulation of gluconeogenesis

The regulation of gluconeogenesis and glycolysis involves the enzymes unique to each pathway, and not the common ones.
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.

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Pyruvate carboxylase

Gluconeogenesis
Fig. 4 – Two Alternative Fates for Pyruvate

In the mitochondrion, pyruvate can be converted to:

        • acetyl-CoA, in the reaction catalyzed by pyruvate dehydrogenase complex, reaction that connects glycolysis to the Krebs cycle;
        • oxaloacetate, in the reaction catalyzed by private 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 β-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. Acetyl-CoA is a positive allosteric effector of pyruvate carboxylase, and a negative allosteric effector of pyruvate kinase. Moreover, it inhibits pyruvate dehydrogenase both through end-product inhibition and phosphorylation through the activation of a specific kinase.
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 pyruvate dehydrogenase 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.

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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.

Gluconeogenesis
Fig. 5 – Regulation of FBPase-1 and PFK-1

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.

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PFK-1, FBPase-1 and fructose 2,6-bisphosphate
Gluconeogenesis
Fig. 6 – 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. They act intracellularly through fructose 2,6-bisphosphate or F26BP, 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. 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.
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.

Gluconeogenesis
Fig. 7 – Role of F26BP in the Regulation of Gluconeogenesis and Glycolysis

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.

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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. 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.

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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.

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Xylulose 5-phosphate

Gluconeogenesis
Fig. 8 – Xylulose 5-phosphate

Xylulose 5-phosphate, a product of the hexose monophosphate shunt, 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 lipid synthesis. 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.

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Precursors of gluconeogenesis

Besides the aforementioned pyruvate, the major gluconeogenic precursors are lactate (see above), glycerol, the majority of the amino acids, and, more generally, any compound that can be converted to pyruvate or oxaloacetate.

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Glycerol

Glycerol is released by hydrolysis of triglycerides in adipose tissue, and of glycerophospholipids. With the exception of propionyl-CoA (see below), 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.

Gluconeogenesis
Fig. 9 – Conversion of Glycerol to DHAP

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. 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% of glucose production.

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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.
Gluconeogenesis
Fig. 10 – 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.

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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.

Pyruvate + NAD+ + CoASH → Acetyl-CoA + NADH + H+ + C02

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.

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Propionate

Propionate, a three carbon fatty acid, 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 β-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.
    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 (see above).

The oxidation of propionyl-CoA to succinyl-CoA involves three reactions that occur in the liver and other tissues.

Gluconeogenesis
Fig. 11 – Conversion of Propionyl-CoA to Succinyl-CoA

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. In the subsequent reaction, catalyzed by methylmalonyl-CoA epimerase (EC 5.1.99.1), D-methylmalonyl-CoA is epimerized to its L-stereoisomer. 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.

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References

Bender D.A. Introduction to nutrition and metabolism. 3rd Edition. Taylor & Francis, 2004

Garrett R.H., Grisham C.M. Biochemistry. 4th Edition. Brooks/Cole, Cengage Learning, 2010

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;100:5107-12. 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):2915-21. doi: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):443-54. doi:10.1042/BJ20141171

Nelson D.L., M. M. Cox M.M. Lehninger. Principles of biochemistry. 6th Edition. W.H. Freeman and Company, 2012

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):1342-49. doi:10.1021/acs.jproteome.6b00110

Stipanuk M.H., Caudill M.A. Biochemical, physiological, and molecular aspects of human nutrition. 3rd Edition. Elsevier health sciences, 2013 [Google eBooks]

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):2861-63 [PDF]

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):3483-86 [PDF]

Glucose-alanine cycle

Glucose-alanine cycle: contents in brief

What is the glucose-alanine cycle?

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 and amino groups as alanine to the liver, and  receive glucose from the liver via the bloodstream.
The main steps of the glucose-alanine cycle are summarized below.

  • When in extrahepatic tissues amino acids are used for energy, pyruvate, derived from the glycolytic pathway, 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 α-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 the gluconeogenesis pathway 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 carbohydrate and amino acid metabolism, as schematically described below.

Glucose → Pyruvate → Alanine → Pyruvate → Glucose

Glucose-Alanine Cycle
Fig. 1 – 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.

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The steps of the glucose-alanine cycle

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.
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% 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.
The removal of the α-amino group occurs by transamination, that can be summarized as follows:

α-Keto acid + Amino acid ⇄ New amino acid + New α-keto acid

Such reactions, catalyzed by enzymes called aminotransferases or transaminases (EC 2.6.1) are freely reversible (see below).
Branched chain amino acids, for example, transfer the amino group to α-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).

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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. 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 α-ketoglutarate. This transamination is catalyzed by alanine aminotransferase or ALT (EC 2.6.1.2), an enzyme found in most animal and plant tissues.

Glutamate + Pyruvate ⇄ Alanine + α-Ketoglutarate

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.

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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 α-ketoglutarate as an α-keto acid acceptor. The products of the reaction are pyruvate, i.e. the carbon skeleton of alanine, and glutamate.

Alanine + α-Ketoglutarate ⇄ Glutamate + Pyruvate

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 α-ketoglutarate, which can enter the Krebs cycle. This reaction is an anaplerotic reaction that links amino acid metabolism with the Krebs cycle.

Glucose-Alanine Cycle

However, glutamate can also react  with oxaloacetate to form aspartate and α-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.

Glutamate + Oxaloacetate ⇄ Aspartate + α-Ketoglutarate

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.

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Transaminases

As previously mentioned, the removal of the amino group from amino acids occurs through transamination (see above for the general reaction). These reactions are catalyzed by enzymes called aminotransferases or transaminases.
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 α-keto acids, notably pyruvate, oxaloacetate and α-ketoglutarate.
Cells contain different types of aminotransferases: many are specific for α-ketoglutarate as α-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 α-keto acid acceptor is aminated and the amino acid deaminated.

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Functions of the glucose-alanine cycle

This cycle has various functions.

  • 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.

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Energy cost of the glucose-alanine cycle

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 (see below).

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. 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.

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Similarities and differences between glucose-alanine cycle and Cori cycle

There are some analogies between the two cycles, which are listed below.

  • 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, Krebs cycle or gluconeogenesis that occur within individual cells
Glucose-Alanine Cycle
Fig. 2 – Glucose-Alanine Cycle and Cori 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.

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References

Berg J.M., Tymoczko J.L., and Stryer L. Biochemistry. 5th Edition. W. H. Freeman and Company, 2002

Felig P., Pozefsk T., Marlis E., Cahill G.F. Alanine: key role in gluconeogenesis. Science 1970;167(3920):1003-4. doi:10.1126/science.167.3920.1003

Gropper S.S., Smith J.L., Groff J.L. Advanced nutrition and human metabolism. Cengage Learning, 2009 [Google eBooks]

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):1807-19. 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):5713-23 [PDF]

Nelson D.L., M. M. Cox M.M. Lehninger. Principles of biochemistry. 6th Edition. W.H. Freeman and Company, 2012

Raju S.M., Madala B. Illustrated medical biochemistry. Jaypee Brothers Publishers, 2005 [Google eBooks]

Wu G. Amino acids: biochemistry and nutrition. CRC Press, 2013 [Google eBooks]

Cori cycle: definition, function, biochemistry, involved tissues

Cori cycle: contents in brief

What is the Cori cycle?

The Cori cycle, or glucose-lactate 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 low oxygen conditions and the liver. This cycle can be summarized as follows:

  • the conversion of glucose to lactic acid, or lactate, by anaerobic glycolysis in skeletal muscle cells;
  • the diffusion of lactate from muscle cells into the bloodstream, by which it is transported to the liver;
  • the conversion of lactate to glucose by hepatic gluconeogenesis;
  • the diffusion of glucose from the hepatocytes into the bloodstream, by which it is transported back to the skeletal muscle cells, thereby closing the cycle.

Summarizing, we have: part of the lactate produced in skeletal muscle is converted to glucose in the liver, and transported back to skeletal muscle, thus closing the cycle.

Glucose → Lactate →Glucose

The importance of this cycle is demonstrated by the fact that it may account for about 40% of plasma glucose turnover.

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Where does the Cori cycle occur?

In addition to skeletal muscle, this metabolic cooperation was also demonstrated between other extrahepatic tissues and liver.  Indeed, like the glucose-alanine cycle, the glucose-lactate cycle is active between the liver and all those tissues that do not completely oxidize glucose to CO2 and H2O, in which case pyruvate for conversion to lactate or, by transamination, to alanine would lack (see below).
In addition to skeletal muscle cells, examples of cells that continually produce lactic acid are red blood cells, immune cells in the lymph nodules,  proliferating cells in the bone marrow, and epithelial cells in the skin.
Note: skeletal muscle produces lactate even at rest, although at low rate.

Cori Cycle
Fig. 1 – The Cori Cycle

From a biochemical point of view, the Cori cycle links gluconeogenesis with anaerobic glycolysis, using different tissues to compartmentalize opposing metabolic pathways. In fact, in the same cell, regardless of the cell type, these metabolic pathways are not very active simultaneously. Glycolysis is more active when the cell requires ATP; by contrast, when the demand for ATP is low, gluconeogenesis, in those cells where it occurs, is more active.
And it is noteworthy that, although traditionally the metabolic pathways, such as glycolysis, citric acid cycle, or gluconeogenesis, are considered to be confined within individual cells, the Cori cycle, as well as the glucose-alanine cycle, occurs between different cell types.
Finally, it should be underscored that the Cori cycle also involves the renal cortex, particularly the proximal tubules, another site where gluconeogenesis occurs.

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The steps of the Cori cycle

The analysis of the steps of the Cori cycle is made considering the lactate produced by red blood cells and skeletal muscle cells.
Mature red blood cells are devoid of mitochondria, nucleus and ribosomes, and obtain the necessary energy only by glycolysis. The availability of NAD+ is essential for glycolysis to proceed as well as for its rate: 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).

Glyceraldehyde 3-phosphate + NAD+ → 1,3-Bisphosphoglycerate + NADH + H+

The accumulation of NADH is avoided by the reduction of pyruvate to lactate, in the reaction catalyzed by lactate dehydrogenase (EC 1.1.1.27), where NADH acts as reducing agent.

Pyruvate + NADH + H+ → Lactate + NAD+

The skeletal muscle, particularly fast-twitch fibers which contain a reduced number of mitochondria, under low oxygen condition, such as during intense exercise, produces significant amounts of lactate. 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% 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.

And, like in red blood cells, the reaction catalyzed by lactate dehydrogenase, regenerating NAD+, allows glycolysis to proceed.
However, lactate is an end product of metabolism that must be converted back into pyruvate to be used.
The plasma membrane of most cells is freely permeable to both pyruvate and lactate that can thus reach the bloodstream. 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 the skeletal muscle isoenzyme of LDH.
Once into the bloodstream, lactate reaches the liver, which is its major user, where it is oxidized to pyruvate in the reaction catalyzed by the liver isoenzyme of lactate dehydrogenase (see below).

Lactate + NAD+ → Pyruvate + NADH + H+

In the hepatocyte, this oxidation is favored by the low NADH/NAD+ ratio in the cytosol.
Then, pyruvate enters the gluconeogenesis pathway to be converted into glucose.
Glucose leaves the liver, enters into the bloodstream and is delivered to the muscle, as well as to other tissues and cells that require it, such as red blood cells and neurons, thus closing the cycle.

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Lactate dehydrogenase

The enzyme is a tetramer composed of two different types of subunits, designed as:

  • H subunit (heart) or B chain;
  • M subunit (muscle) or A chain.

The H subunit predominates in the heart, whereas the M subunit predominates in the  skeletal muscle and liver. Typically, tissues in which a predominantly or exclusively aerobic metabolism occurs, such as the heart, synthesize H subunits to a greater extent than M subunits, whereas tissues in which anaerobic metabolism is important, such as skeletal muscle, synthesize M subunits to a greater extent than H subunits.
The two subunits associate in 5 different ways to form homopolymers, that is, macromolecules formed by repeated, identical subunits, or heteropolymers, that is, macromolecules formed by different subunits. Different LDH  isoenzymes have different catalytic properties, as well as different distribution in various tissues, as indicated below:

  • H4, also called type 1, LDH1, or A4, a homopolymer of H subunits, is found in cardiac muscle, kidney, and red blood cells;
  • H3M1, also called type 2, LDH2, or A3B, has a tissue distribution similar to that of LDH1;
  • H2M2, also called type 3, LDH3, or A2B2, is found in the spleen, brain, white cells, kidney, and lung;
  • H1M3, also called type 4, LDH4, or AB3, is found in the spleen, lung, skeletal muscle, lung, red blood cells, and kidney;
  • M4, also called type 5, LDH5, or B4, a homopolymer of M subunits, is found in the liver, skeletal muscle, and spleen.

The H4 isoenzyme has a higher substrate affinity than the M4 isoenzyme.
The H4 isoenzyme is allosterically inhibited by high levels of pyruvate (its product), whereas the M4 isoenzyme is not.
The other LDH isoenzymes have intermediate properties, depending on the ratio between the two types of subunits.
It is thought that the H4 isoenzyme is the most suitable for catalyzing the oxidation of lactate to pyruvate that, in the heart, due to its exclusively aerobic metabolism, is then completely oxidized to CO2 and H2O. Instead, the M4 isoenzyme is the main isoenzyme found in skeletal muscle, most suitable for catalyzing the reduction of pyruvate to lactate, thus allowing glycolysis to proceed in anaerobic conditions.

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Other metabolic fates of lactate

From the above, it is clear that lactate is not a metabolic dead end, a waste product of glucose metabolism.
And it may have a different fate from that entering the Cori cycle.
For example, in skeletal muscle during recovery following an exhaustive exercise, that is, when oxygen is again available, or if the exercise is of low intensity, lactate is re-oxidized to pyruvate, due to NAD+ availability, and then completely oxidized to CO2 and H20, with a greater production of ATP than in anaerobic condition. In such conditions, the energy stored in NADH will be released, yielding on average 2.5 ATP per molecule of NADH.
In addition, lactate can be taken up by exclusively aerobic tissues, such as heart, to be oxidized to CO2 and H20.

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Energy cost of the Cori cycle

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, that is, 6 ATP.
The ATP-consuming reactions are catalyzed by:

  • pyruvate carboxylase (EC 6.4.1.1): an ATP;
  • phosphoenolpyruvate carboxykinase (EC 4.1.1.32): a GTP;
  • glyceraldehyde 3-phosphate dehydrogenase (EC 1.2.1.12): an ATP.

Since two molecules of lactate are required for the synthesis of one molecule of glucose, the net cost is 2×3=6 high energy bonds per molecule of glucose.
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.

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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. Indeed, this cycle allows the effective functioning of many extrahepatic cells at the expense of the liver and partly of the renal cortex. Below, two examples.

  • Red blood cells
    These cells, lacking a nucleus, ribosomes, and mitochondria, are smaller than most other cells. Their small size allows them to pass through tiny capillaries. However, the lack of mitochondria makes them completely dependent on anaerobic glycolysis for ATP production. Then, the lactate is partly disposed of by the liver and renal cortex.
  • Skeletal muscle
    Its cells, and particularly fast-twitch fibers contracting under low oxygen conditions, such as during intense exercise, produce much lactate.
    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. However, the rate of ATP production by anaerobic glycolysis is greater than that produced by the complete oxidation of glucose. Therefore, to meet the energy requirements of contracting muscle, anaerobic glycolysis is an effective means of ATP production. But this could lead to an intracellular accumulation of lactate, and a consequent reduction in intracellular pH. Obviously, such accumulation does not occur, due also to the Cori cycle, in which the liver pays the cost of the disposal of a large part of the muscle lactate, thereby allowing the muscle to use ATP for the contraction.
    And the oxygen debt, which always occurs after a strenuous 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 from lactate.
  • 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 greater production of lactate, an increase in the flux through the Cori cycle and an increase in ATP consumption in the liver, which, as previously said, is supported by an increase in fatty acid oxidation. Hence, the nutrition plan provided to these patients must be taken into account this increase in energy consumption.
  • A similar condition seems to occur also in cancer patients with progressive weight loss.
  • The Cori cycle is also important during overnight fasting and starvation.

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The Cori cycle and glucose-alanine cycle

These cycles are metabolic pathways that contribute to ensure a continuous delivery of glucose to tissues for which the monosaccharide is  the primary source of energy.
The main difference between the two cycles consists in the three carbon intermediate which is recycled: in the Cori cycle, carbon returns to the liver in the form of pyruvate, whereas in the glucose-alanine cycle in the form of alanine.
For more information, see: glucose-alanine cycle.

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References

Bender D.A. Introduction to nutrition and metabolism. 3rd Edition. Taylor & Francis, 2004

Berg J.M., Tymoczko J.L., and Stryer L. Biochemistry. 5th Edition. W. H. Freeman and Company, 2002

Iqbal S.A., Mido Y. Biochemistry. Discovery Publishing House, 2005 [Google eBook]

Nelson D.L., M. M. Cox M.M. Lehninger. Principles of biochemistry. 6th Edition. W.H. Freeman and Company, 2012

Newsholme E.A., Leech T.R. Functional biochemistry in health and disease. John Wiley J. & Sons, Inc., Publication, 2010 [Google eBook]

Rawn J.D. Biochimica. Mc Graw-Hill, Neil Patterson Publishers, 1990

Rosenthal M.D., Glew R.H. Medical biochemistry – Human metabolism in health and disease. John Wiley J. & Sons, Inc., Publication, 2009

Shils M.E., Olson J.A., Shike M., Ross A.C. Modern nutrition in health and disease. 9th Ed., by Lippincott, Williams & Wilkins, 1999

Stipanuk M.H., Caudill M.A. Biochemical, physiological, and molecular aspects of human nutrition. 3rd Edition. Elsevier health sciences, 2013 [Google eBooks]

Bile salts: definition, functions, enterohepatic circulation, synthesis

Bile salts: contents in brief

What are bile salts

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.

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Chemical structure of bile salts

Bile Salts
Fig. 1 – Chemical Structures of the Most Abundant Bile Acids

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.
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 (see below).
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.

Bile Salts
Fig. 2 – Cholic Acid Structure

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.

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.

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Primary, conjugated and secondary bile salts

Bile Salts
Fig. 3 – Conjugated Bile Acids

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% of all bile acids. Before being secreted into the biliary tree, they are almost completely conjugated, up to 98%, with the glycine or taurine, to form glycoconjugates and tauroconjugates, respectively. In particular, approximately 75% of cholic acid and chenodeoxycholic acid are conjugated with glycine, to form glycocholic acid  and glycochenodeoxycholic acid, the remaining 25% with taurine, to form taurocholic acid and taurochenodeoxycholic.
Conjugated bile acids are molecules with more hydrophilic groups than unconjugated bile acids, therefore with a increased emulsifying capacity. 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 colonic microbiota to form secondary bile acids (see below). They make up the remaining 20% 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% of bile salts present in the bile.
  • Unconjugated forms, such as cholic acid, deoxycholic acid, chenodeoxycholic acid, and lithocholic acid.

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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 the absorption of lipid digestion products, as well as 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.

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Enterohepatic circulation of bile salts

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 (see below).

Up to 95% 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 OSTα/OSTβ, 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%, 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.

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Intestinal metabolism of bile acids

Bile Salts
Fig. 4 – Intestinal Bile Acid 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.
  • 7α-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 7α-dehydroxylation, unlike oxidation and epimerization (see below), 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.

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, 7α-dehydroxylations is carried out by a limited number of colonic anaerobes.
7α-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.

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Soluble fibers and reabsorption of bile salts

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 7α-hydroxylase and sterol 12α-hydroxylase (see below), 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.

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Synthesis of primary bile acids

Bile Salts
Fig. 5 – Primary Bile Acid Biosynthesis

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% 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.

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The classical or neutral pathway

In humans, up to 90% of bile salts are produced via the classical pathway (see fig. 5), 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 7α-hydroxycholesterol. The reaction is catalyzed by cholesterol 7α-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.

Cholesterol + NADPH + H+ + O2 → 7α-Hydroxycholesterol + NADP+ + H2O

  • 7α-Hydroxycholesterol undergoes oxidation of the 3β-hydroxyl group and the shift of the double bond from the 5,6 position to the 4,5 position, to form 7α-hydroxy-4-cholesten-3-one. The reaction is catalyzed by 3β-hydroxy-Δ5-C27-steroid oxidoreductase or HSD3B7 (E.C. 1.1.1.181), an enzyme localized in the endoplasmic reticulum.
  • 7α-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 7α-hydroxy-4-cholesten-3-one 12α-monooxygenase or sterol 12α-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 5β-reductase or AKR1D1 (E.C. 1.3.1.3), a cytosolic enzyme.

It should be underlined that the activity of sterol 12α-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 12α-hydroxylase gene transcription is one of the main regulatory step of the classical pathway.

Therefore, if 7α-hydroxy-4-cholesten-3-one proceeds via the reaction catalyzed by sterol 12α-hydroxylase, the following reactions will occur.

  • 7α-Hydroxy-4-cholesten-3-one is hydroxylated at position 12 by sterol 12α-hydroxylase, to form 7α,12α-dihydroxy-4-cholesten-3-one.
  • 7α,12α-Dihydroxy-4-cholesten-3-one undergoes reduction of the double bond at 4,5 position, in the reaction catalyzed by 3-oxo-Δ4-steroid 5β-reductase, to form 5β-cholestan-7α,12α-diol-3-one.
  • 5β-Cholestan-7α,12α-diol-3-one undergoes reduction of the hydroxyl group at position 4, in the reaction catalyzed by 3α-hydroxysteroid dehydrogenase or AKR1C4 (EC 1.1.1.213), a cytosolic enzyme, to form 5β-cholestan-3α,7α,12α-triol.
  • 5β-Cholestan-3α,7α,12α-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 3α,7α,12α-trihydroxy-5β-cholestanoic acid.
  • 3α,7α,12α-Trihydroxy-5β-cholestanoic  acid is activated to its coenzyme A ester, 3α,7α,12α-trihydroxy-5β-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.
  • 3α,7α,12α-Trihydroxy-5β-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 7α-hydroxy-4-cholesten-3-one does not proceed via the reaction catalyzed by sterol 12α-hydroxylase, it enters the pathway that leads to the synthesis of chenodeoxycholic acid conjugates, through the reactions described below.

  • 7α-Hydroxy-4-cholesten-3-one is converted to 7α-hydroxy-5β-cholestan-3-one in the reaction catalyzed by 3-oxo-Δ4-steroid 5β-reductase.
  • 7α-Hydroxy-5β-cholestan-3-one is converted to 5β-cholestan-3α,7α-diol in the reaction catalyzed by 3α-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.

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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% of the bile salts.
This pathway  (see fig. 5) 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 3β-hydroxy-5-cholestenoic acid in a reaction catalyzed by sterol 27-hydroxylase.
  • 3β-Hydroxy-5-cholestenoic acid is hydroxylated at position 7 in the reaction catalyzed by oxysterol 7α-hydroxylase or CYP7B1 (EC 1.14.13.100), an enzyme localized in the endoplasmic reticulum, to form 3β-7α-dihydroxy-5-colestenoic acid.
  • 3β-7α-Dihydroxy-5-cholestenoic acid is converted to 3-oxo-7α-hydroxy-4-cholestenoic acid, in the reaction catalyzed by 3β-hydroxy-Δ5-C27-steroid oxidoreductase.
  • 3-Oxo-7α-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 7α,27-dihydroxycholesterol in the reaction catalyzed by oxysterol 7α-hydroxylase and cholesterol 7α-hydroxylase.
  • 7α,27-Dihydroxycholesterol is converted to 7α,26-dihydroxy-4-cholesten-3-one in the reaction catalyzed by 3β-hydroxy-Δ5-C27-steroid oxidoreductase;

7α, 26-Dihydroxy-4-cholesten-3-one can be transformed directly to conjugates of chenodeoxycholic acid, or can be converted to 3-oxo-7α-hydroxy-4-colestenoic acid,  and then undergo side chain modifications and other reactions that lead to the synthesis of the conjugates of chenodeoxycholic acid.

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Minor pathways

There are also minor pathways (see fig. 5) 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 7α-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.

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Bile salts: regulation of synthesis

Regulation of bile acid synthesis occurs via a negative feedback mechanism, particularly on the expression of cholesterol 7α-hydroxylase and sterol 12α-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-4α or HNF-4α. These transcription factors bind to a sequence in the promoter region of 7α-hydroxylase and 12α-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.

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References

Chiang J.Y.L. Bile acids: regulation of synthesis. J Lipid Res 2009;50(10):1955-66. doi:10.1194/jlr.R900010-JLR200

Gropper S.S., Smith J.L. Advanced nutrition and human metabolism. 6h Edition. Cengage Learning, 2012 [Google eBook]

Moghimipour E., Ameri A., and Handali S. Absorption-enhancing effects of bile salts. Molecules 2015;20(8); 14451-73. 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):804-16. doi:10.3748/wjg.15.804

Rawn J.D. Biochimica. Mc Graw-Hill, Neil Patterson Publishers, 1990

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):456-68. doi:10.1038/ncpgasthep1179

Human gut microbiota: definition, composition, and the effect of diet

Human gut microbiota: contents in brief

Definition and composition of the human gut microbiota

Gut Microbiota
Fig. 1 – Lactobacillus acidophilus

The human gastrointestinal tract is one of the most fierce and competitive ecological niches. It harbors viruses, eukaryotes, bacteria, and one member of Archaebacteria, Methanobrevibacter smithii.
Bacteria vary in proportion and amount all along the gastrointestinal tract; the greatest amount is found in the colon, which contains over 400 different species belonging to 9 phyla or divisions (of the 30 recognized phyla), and hereafter you refer to them as gut microbiota.
These are the phyla and some of their most represented genera.

  • Actinobacteria (Gram-positive bacteria); Bifidobacterium, Collinsella, Eggerthella, and Propionibacterium.
  • Bacteroidetes (Gram-negative bacteria); more than 20 genera including Bacteroides, Prevotella and Corynebacterium.
  • Cyanobacteria (Gram-negative bacteria).
  • Firmicutes (Gram-positive bacteria); at least 250 genera, including Mycoplasma, Bacillus, Clostridium, Dorea, Faecalibacterium, Ruminococcus, Eubacterium, Staphylococcus, Streptococcus, Lactobacillus, Lactococcus, Enterococcus, Sporobacter, and Roseburia.
  • Fusobacteria (Gram-negative bacteria);
  • Lentisphaerae (Gram-negative bacteria).
  • Proteobacteria (Gram-negative bacteria); Escherichia, Klebsiella, Shigella, Salmonella, Citrobacter, Helicobacter, and Serratia.
  • Spirochaeates (Gram-negative bacteria).
  • Verrucomicrobia (Gram-negative bacteria).

The presence of a small subset of the bacterial world in the colon is the result of a strong selective pressure which acted, during evolution, on both the microbial colonizers, selecting organisms very well adapted to this environment, and the intestinal niche. And nevertheless, each individual harbors an unique bacterial community in his gut.
Despite the high variability existing both with regard to taxa and between individuals, it has been proposed, but not accepted by all researchers, that in most adults the bacterial gut microbiota can be classified into variants or “enterotypes”, on the basis of the ratio of the abundance of the genera Bacteroides and Prevotella. This seems to indicate that there is a limited number of well balanced symbiotic states, which could respond differently to factors such as diet, age, genetics, and drug intake (see below).

Adult’s gut harbors a large and diverse community of DNA and RNA viruses made up of about 2,000 different genotypes, none of which is dominant. Indeed, the most abundant virus accounts for only about 6% of the community, whereas in infants the most abundant virus accounts over 40% of the community. The majority of DNA viruses are bacteriophages or phages, that is, viruses that infect bacteria (they are the most abundant biological entity on earth, with an estimated population of about 1031 units), whereas the majority of RNA viruses are plant viruses.

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Factors affecting gut microbiota composition and development

The intestinal bacterial community is regulated by several factors, most of which are listed below.

  • The diet of the host.
    It seems to be the most important factor.
    Traditionally considered sterile, mother’s milk harbors a rich microbiota consisting of more than 700 species, dominated by staphylococci, streptococci, bifidobacteria and lactic acid bacteria. Therefore, it is a major source for the colonization of the breastfed infant gut, and it was suggested that this mode of colonization is closely correlated with infant’s health status, because, among other functions, it could protect against infections and contribute to the maturation of the immune system. Breast milk affects intestinal microbiota also indirectly, through the presence of oligosaccharides with prebiotic activity that stimulate the growth of specific bacterial groups including staphylococci and bifidobacteria.
    A recent study has compared the intestinal microbiota of European and African children (respectively from Florence and a rural village in Burkina Faso) between the ages of 1 and 6 years old. It has highlighted the dominant role of diet over variables such as climate, geography, hygiene and health services (it was also observed the absence of significant differences in the expression of key genes regulating the immune function, which suggests a functional similarity between the two groups). Indeed infants, as long as they are breastfed, have a very similar gut microbiota, rich in Actinobacteria, mainly Bifidobacterium (see below). The subsequent introduction of solid foods in the two groups, a Western diet rich in animal fat and protein in European children, and low in animal protein but rich in complex carbohydrates in African children, leads to a differentiation in the Firmicutes/Bacteroidetes ratio between the two groups. Gram-positive bacteria, mainly Firmicutes, were more abundant than Gram-negative bacteria in European children, whereas Gram-negative bacteria, mainly Bacteroidetes, prevailed over Gram-positive bacteria in African children.
    And the long-term diets are strongly associated to the enterotype partitioning. Indeed, it has been observed that:

a diet high in animal fat and protein, i.e. a Western-type diet, leads to a gut microbiota dominated by the Bacteroides enterotype;
a diet high in complex carbohydrates, typical of agrarian societies, leads to the prevalence of the Prevotella enterotype.

Similar results emerged from the aforementioned study on children. In the Europeans, gut microbiota was dominated by taxa typical of Bacteroides enterotype, whereas in the Burkina Faso children, Prevotella enterotype dominates.
With short-term changes in the diet (10 days), such as the switch from a low-fat and high-fiber diet to a high-fat and low-fiber diet and vice versa, changes were observed in the composition of the microbiome (within 24 hours), but no stable change in the enterotype partitioning. And this underlines as a long-term diet is needed for a change in the enterotypes of the gut microbiota.
Dietary interventions can also result in changes in the gut virome, which moves to a new state, that is, changes occur in the proportions of the pre-existing viral populations, towards which subjects on the same diet converge.

  • pH, bile salts and digestive enzymes.
    The stomach, due to its low pH, is a hostile environment for bacteria, which are not present in high numbers, about 102-103 bacterial cells/gram of tissue. In addition to Helicobacter pylori, able to cause gastritis and gastric ulcers, microorganisms of the genus Lactobacillus are also present.
    Reached the duodenum, an increase in bacterial cell number occurs, 104-105 bacterial cells/gram of tissue; and similar bacterial concentrations are present in the jejunum and proximal ileum. The low number of microorganisms present in the small intestine is due to the inhospitable environment, consequent to the fact that there is the opening of the ampulla of Vater in the descending part of the duodenum, which pours pancreatic juice and bile into the duodenum, that is, pancreatic enzymes and bile salts, which damage microorganisms.
    In the terminal portion of the ileum, where the activities of pancreatic enzymes and bile salts are lower, there are about 107 bacterial cells/gram of tissue, and up to 1012-1014 bacterial cells/gram of tissue in the colon, so that bacteria represent a large proportion, about 40%, of the fecal mass.
    The distribution of bacteria along the intestine is strategic. In the duodenum and jejunum, the amount of available nutrients is much higher than that found in the terminal portion of the ileum, where just water, fiber, and electrolytes remain. Therefore, the presence of large number of bacteria in the terminal portion of the ileum, and even more in the colon, is not a problem. The problem would be to find a high bacterial concentration in the duodenum, jejunum, and proximal parts of the ileum; and there is a disease condition, called small intestinal bacterial overgrowth or SIBO, in which the number of bacteria in the small intestine increases by about 10-15 times. This puts them in a position to compete with the host for nutrients and give rise to gastrointestinal disturbances such as diarrhea.
  • The geographical position and the resulting differences in lifestyle, diet, religion etc.
    For example, a kind of geographical gradient occurs in the microbiota of European infants, with a higher number of Bifidobacterium species and some of Clostridium in Northern infants, whereas Southern infants have higher levels of Bacteroides, Lactobacillus and Eubacterium.
  • The mode of delivery (see below).
  • The genetics of the host.
  • The health status of the infant and mother.
    For example, in mothers with inflammatory bowel disease or IBD, Faecalibacterium prausnitzii, a bacterium that produces butyrate (an important source of energy for intestinal cells), and with anti-inflammatory activity is depleted, whereas there is an increase in the number of adherent Escherichia coli.
  • The treatment with antibiotics.
  • Bacterial infections and predators.
    Bacteriocins, i.e. proteins with antibacterial activity, and bacteriophages.
    Phages play an important role in controlling the abundance and composition of the gut microbiota. In particular, they could play a major role in the colonization of the newborn, infecting the dominant bacteria thus allowing to another bacterial strain to become abundant.
    This model of predator-prey dynamics, called “kill the winner”, suggests that the blooms of a specific bacterial species would lead to blooms of their corresponding bacteriophages, followed by a decline in their abundance. Therefore, the most abundant bacteriophage genotype will not be the same at different times. And although some the gene sequences present in the infant gut virome are stable over the first three months of life, dramatic changes occur in the overall composition of the viral community between the first and second week of life. During this time period also the bacterial community is extremely dynamic (see below).
  • The competition for space and nutrients.

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The composition of the gut microbiota throughout life

Gut Microbiota
Fig. 2 – Development of Intestinal Microflora

The development of the intestinal microbial ecosystem is a complex and crucial event in human life, highly variable from individual to individual, and influenced by the factors outlined above.
In utero, the gut is considered sterile, but is rapidly colonized by microbes at birth, as the infant is born with an immunological tolerance instructed by the mother.
However, recent studies show the presence of bacteria in the placental tissue, umbilical cord blood, fetal membranes and amniotic fluid from healthy newborns without signs of infection or inflammation. And for example, the meconium of premature infants, born to healthy mothers, contains a specific microbiota, with Firmicutes as the main phylum, and predominance of staphylococci, whereas Proteobacteria, in particular species such as Escherichia coli, Klebsiella pneumoniae, Serratia marcescens, but also enterococci are more abundant in the faeces.
Note: the meconium is free of detectable viruses.
It seems that both vaginal and gut bacteria may gain access to the fetus, although via different route of entry: by ascending entry the vaginal ones, by dendritic cells of the immune system the gut ones. Therefore, there could exist a fetal microbiota.

Colonization occurs during delivery by a maternal inoculum, generally composed of aerobic and facultative bacteria (the newborn’s gut initially contains oxygen), then replaced by obligate anaerobes,  bacteria typically present in adulthood, to which they have created a hospitable environment.
Furthermore, there is a small number of different taxa, with a relative dominance of the phyla Actinobacteria and Proteobacteria, that remains unchanged during the first month of life, but not in the subsequent ones as there is a large increase in variability and new genetic variants. Many studies underline that the initial exposure is important in defining the “trajectories” which will lead to the adult ecosystems. Additionally, these initial communities may act as a source of protective or pathogenic microorganisms.

Mother’s vaginal and fecal microbiotas are the main sources of inoculum in vaginally delivered infants. Indeed, infants harbor microbial communities dominated by species of the genera Lactobacillus (the most abundant genus in the vaginal microbiota and early gut microbiota) Bifidobacterium, Prevotella, or Sneathia. And it seems likely that anaerobes, such as members of the phyla Firmicutes and Bacteroidetes, not growing outside of their host, rely on the close contact between mother and offspring for transmission. Finally, due to the presence of oxygen in infant gut, the transmission of strict anaerobes could occur not directly at birth but at a later stage by means of spores.
The first bacteria encountered by infants born by caesarean section are those of the skin and hospital environment, and gut microbiota is dominated by species of the genera Corynebacterium, Staphylococcus and Propionibacterium, with a lower bacterial count and diversity in first weeks of life than infants born vaginally.
Further evidence supporting the hypothesis of vertical transmission is the similarity between the microbiota of meconium and samples obtained from possible sites of contamination.
These “maternal bacteria” do not persist indefinitely, and are replaced by other populations within the first year of life.
Objects, animals, mouths and skin of relatives, and breast milk are secondary sources of inoculum; and breast milk (see below) seems to have a primary role in determining the microbial succession in the gut.
The variation and diversity among children reflect instead the individuality of these microbial exposures.
Note: the delivery mode seems also to influence the immune system during the first year of life, perhaps via the influence on the development of gut microbiota. Infants born by cesarean section have:

  • a lower bacterial count in stool samples at one month of age, mainly due to the higher number of bifidobacteria in infants born vaginally;
  • a higher number of antibody secreting cells, which could reflect an excessive antigen exposure (the intestinal barrier would be more vulnerable to the passage of antigens).

Within a days after birth, a thriving community is established. This community is less stable over time and more variable in composition than that of adults. Very soon, it will be more numerous than that of the child’s cells, evolving according to a temporal pattern highly variable from individual to individual.
Viruses, absent at birth, reach about 108 units/gram wet weight of faeces by the end of the first week of life, therefore representing a dynamic and abundant component of the developing gut microbiota. However, viral community has an extremely low diversity, like bacteria, and is dominated by phages, which probably influence the abundance and diversity of co-occurring bacteria, as seen above. The initial source of the viruses is unknown; of course, maternal and/or environmental inocula are among the possibilities. Notably, the earliest viruses could be the result of induction of prophages from the “newborn” gut bacterial flora, hypothesis supported by the observation that more than 25% of the phage sequences seem to be very similar to those of phages infecting bacteria such as Lactococcus, Lactobacillus, Enterococcus, and Streptococcus, which are abundant in breast milk.

By the end of the first month of life it is thought that the initial phase of rapid acquisition of microorganism is over.
In 1-month-old-infants, the most abundant bacteria belong to the genera Bacteroides and Escherichia, whereas Bifidobacterium, along with Ruminococcus, appear and grow to become dominant in the gastrointestinal tract of the breastfed infants between 1 and 11 months. Bifidobacteria such as Bifidobacterium longum subspecies infantis:

  • are known to be closely related to breastfeeding;
  • are among the best characterized commensal bacteria;
  • are considered probiotics, that is, microorganisms which can confer health benefits to the host.

Their abundance confers also benefits through competitive exclusion, that is, they are an obstacle to colonization by pathogens. And indeed, Escherichia and Bacteroides can become preponderant if Bifidobacterium is not adequately present in the gut.
In contrast, bacteria of the genera Escherichia (e.g. E. coli), Clostridium (e.g. C. difficile), Bacteroides (e.g. B. fragilis) and Lactobacillus are present in higher levels in formula-fed infants than in breastfed infants.
Although breast-fed infants receive only breast milk until weaning, their microbiota can show a large variability in the abundances of bacterial taxa, with differences between individuals also with regard to the temporal patterns of variation. These variations may be due to diseases, treatments with antibiotics, changes in host lifestyle, random colonization events, as well as differences in immune responses to the gut colonizing microbes. However, it is not yet clear how these factors contribute to shape infant gut microbiota.
It seems that also the virome changes rapidly after birth, as the majority of the viral sequences present in the first week of life are not found after the second week. Moreover, the repertoire expands rapidly in number and diversity during the first three months. This is in contrast with the stability observed in the adult virome, where 95% of the sequences are conserved over time.

In normal condition, towards the end of the first year of life, babies have consumed an adult-like diet for a significant time period and should have developed a microbial community with characteristics similar to those found in the adult gut, such as:

  • a more stable composition, phylogenetically more complex, and progressively more similar among different subjects;
  • a preponderance of Firmicutes and Bacteroidetes, followed by Verrucomicrobia and a very low abundance of Proteobacteria;
  • an increase in short-chain fatty acid (SCFA) levels and bacterial load in the feces;
  • an increase of genes associated with xenobiotic degradation, vitamin biosynthesis, and carbohydrate

Interestingly, the significant turnover of taxa occurring from birth to the end of the first year is accompanied by a remarkable constancy in the overall functional capabilities.
Towards the end of the first year of life also the early viral colonizers were replaced by a community specific to the child.

The gut microbiota reaches maturity at about 2.5 years of age, fully resembling the adult gut microbiota.
The selection of the most adapted bacteria is the result of various factors.

  • The transition to an adult diet.
  • An increased fitness to the intestinal environment of the taxa that typically dominate the adult gut microbiota than the early colonizers.
  • The significant changes in the intestinal environment, result of the developmental changes in the intestinal mucosa.
  • The effects of the microbiota itself.

Therefore, the first 2-3 years of life are the most critical period in which you can intervene to shape the microbiota as best as possible, and so optimize child growth and development.

From a chaotic beginning, all this leads to the establishment of the gut ecosystem typical of the young adult, which is relatively stable over time until old age (viral, archaeal and eukaryotic components included), and dominated, at least in the western population, by members of the phyla Firmicutes, about 60% of the bacterial communities, Bacteroidetes and Actinobacteria (mainly belonging to the Bifidobacterium genus), each comprising about 10% of the bacterial community, followed by Proteobacteria and Verrucomicrobia. The genera Bacteroides, Clostridium, Faecalibacterium, Ruminococcus and Eubacterium make up, together with Methanobrevibacter smithii, the large majority of the adult gut microbial community.
It should be noted that different data were obtained from analysis of populations of African rural areas, as seen above.
And the gut microbiota is sufficiently similar among subjects to allow the identification of a shared core microbiome.
Stability and resilience, however, are subject to numerous variables among which, as previously said, diet seems to be one of the most important. Therefore, in order to maintain the stability of the gut microbiota, the variables have to be kept constant, or in the case of diseases prevented (also through vaccinations). However, the stability and resilience could be harmful if the dominant community is pathogenic.

The gut microbiota undergoes substantial changes in the elderly. In a study conducted in Ireland on 161 healthy people aged 65 years and over, the gut microbiota is distinct from that of younger adults in the majority of subjects, with a composition that seems to be dominated by the phyla Bacteroidetes, the main ones, and Firmicutes, with almost inverted percentages than those found in younger adults (although large variations across subjects were observed). And there are Faecalibacterium, about 6% of the main genera, followed by species of the genera Ruminococcus, Roseburia and Bifidobacterium (the latter about 0.4%) among the most abundant genera.
Also the variability in the composition of the community is greater than in younger adults; this could be due to the increase in morbidities associated with aging and the subsequent increased intake of medications, as well as to changes in the diet.

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References

Breitbart M., Haynes M., Kelley S., Angly F., Edwards R.A., Felts B., Mahaffy J.M., Mueller J., Nulton J., Rayhawk S., Rodriguez-Brito B., Salamon P., Rohwer F. Viral diversity and dynamics in an infant gut. Res Microbiol 2008;159:367-73. doi:10.1016/j.resmic.2008.04.006

Claesson M.J., Cusack S., O’Sullivan O., Greene-Diniz R., de Weerd H., Flannery E., Marchesi J.R., Falush D., Dinan T., Fitzgerald G., et al. Composition, variability, and temporal stability of the intestinal microbiota of the elderly. Proc Natl Acad Sci USA 2011;108(Suppl 1);4586-91. doi:10.1073/pnas.1000097107

Clemente J.C., Ursell L.K., Wegener Parfrey L., and Knight R. The impact of the gut microbiota on human health: an integrative view. Cell 2012;148:1258-70. doi:10.1016/j.cell.2012.01.035

De Filippo c., Cavalieri D., Di Paola M., Ramazzotti M., Poullet J.B., Massart S., Collini S., Pieraccini G., and Lionetti P. Impact of diet in shaping gut microbiota revealed by a comparative study in children from Europe and rural Africa. Proc Natl Acad Sci 2010;107(33):14691-6. doi:10.1073/pnas.1005963107

Dominguez-Bello M.G., Costello E.K., Contreras M., Magris M., Hidalgo G., Fierer N., and Knight R. Delivery mode shapes the acquisition and structure of the initial microbiota across multiple body habitats in newborns. Proc Natl Acad Sci 2010;107:11971-5. doi:10.1073/pnas.1002601107

Fernández L., Langa S., Martín V., Maldonado A., Jiménez E., Martín R., Rodríguez J.M. The human milk microbiota: origin and potential roles in health and disease. Pharmacol Res 2013;69(1):1-10. doi:10.1073/pnas.1002601107

Huurre A., Kalliomäki M., Rautava S., Rinne M., Salminen S., and Isolauri E. Mode of delivery-effects on gut microbiota and humoral immunity. Neonatology 2008;93:236-40. doi:10.1159/000111102

Koenig J.E., Spor A., Scalfone N., Fricker A.D., Stombaugh J., Knight R., Angenent L.T., and Ley R.E. Succession of microbial consortia in the developing infant gut microbiome. Proc Natl Acad Sci 2011;108(1):4578-85. doi:10.1073/pnas.1000081107

Ley R.E., Peterson D.A., and Gordon J.I. Ecological and evolutionary forces shaping microbial diversity in the human intestine. Cell 2006;124(4):837-48. doi:10.1016/j.cell.2006.02.017

Minot S., Sinha R., Chen J., Li H., Keilbaugh S.A., Wu G.D., Lewis J.D., and Bushman F.D. The human gut virome: inter-individual variation and dynamic response to diet. Genome Res 2011;21:1616-1625. doi:10.1101/gr.122705.111

Moreno-Indias I.M., Cardona F., Tinahones F.J. and Queipo-Ortuño M.I. Impact of the gut microbiota on the development of obesity and type 2 diabetes mellitus. Front Microbiol 2014;5(190):1-10 . doi:10.3389/fmicb.2014.00190

Newburg D.S. & Morelli L. Human milk and infant intestinal mucosal glycans guide succession of the neonatal intestinal microbiota. Pediatr Res 2015;77:115-120. doi:10.1038/pr.2014.178

Palmer C., Bik E.M., DiGiulio D.B., Relman D.A., and Brown P.O. Development of the human infant intestinal microbiota. PLoS Biol 2007;5(7):e177. doi:10.1371/journal.pbio.0050177

Rodrıguez J.M., Murphy K., Stanton C., Ross R.P., I. Kober O.I., Juge N., Avershina E., Rudi K., Narbad A., Jenmalm M.C., Marchesi J.R. and Collado M.C. The composition of the gut microbiota throughout life, with an emphasis on early life. Microb Ecol Health Dis 2015;26:26050. doi:10.3402/mehd.v26.26050

Wu G.D., Chen J., Hoffmann C., Bittinger K., Chen Y.Y., Keilbaugh S.A., Bewtra M., Knights D., Walters W.A., Knight R., et al. Linking long-term dietary patterns with gut microbial enterotypes. Science 2011;334:105-8. doi:10.1126/science.1208344

Human microbiota: definition, composition, functions, antibiotics

Human microbiota: contents in brief

What is the human microbiota?

Human Microbiota
Fig. 1 – Lactobacillus casei

It has been known for almost a century that humans harbor a microbial ecosystem, known as human microbiota, remarkably dense and diverse, made up of a  number of viruses and cells much higher than those of the human body, and that accounts for one to three percent of body weight. All the genes encoded by the human body’s microbial ecosystem, which are about 1,000 times more numerous than those of our genome, make up the human microbiome. Microorganisms colonize all the surfaces of the body that are exposed to the environment. Indeed, distinct microbial communities are found on the skin, in the vagina, in the respiratory tract, and along the whole intestinal tract, from the mouth up to rectum, the last part of the intestine.

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Composition of the human microbiota

Human Microbiota
Fig. 2 – Escherichia coli

It is composed of organisms from all taxa.

  • Bacteria, at least 100 trillion (1014) cells, a number ten times greater than that of the human body. They are found in very high concentration in the intestinal tract, up to 1012-1014/gram of tissue, where they form one of the most densely populated microbial habitats on Earth. In the gut, bacteria mainly belong to the Firmicutes, Bacteroidetes and Actinobacteria phyla. Fusobacteria (oropharynx), Tenericutes, Proteobacteria, and Verrucomicrobia are other phyla present in our body.
    Note: bacterial communities in a given body region resemble themselves much more across individuals than those from different body regions of the same individual; for example, bacterial communities of the upper respiratory tract are much more similar across individuals than those of the skin or intestine of the same individual.
  • Viruses, by far the most numerous organisms, about quadrillion units. The genomes of all the viruses harbored in the human body make up the human virome. In the past, viruses and eukaryotes (see below) have been studied focusing on pathogenic microorganisms, but in recent years the attention has also shifted on many non-pathogenic members of these groups. And many of the viral gene sequences found are new, which suggests that there is still much to learn about the human virome. Finally, just like for bacteria, there is considerable interpersonal variability.
  • Archaebacteria, primarily those belonging to the order Methanobacteriales, with Methanobrevibacter smithii predominant in the human gut (up to 10% of all anaerobes).
  • Eukaryotes, and the parasites of the genera Giardia and Entamoeba have probably been among the first to be identified. But there is also a great abundance and diversity of fungal species, belonging to genera such as Candida, Penicillium, Aspergillus, Hemispora, Fusarium, Geotrichum, Hormodendrum, Cryptococcus, Saccharomyces, and Blastocystis.
Human Microbiota
Fig. 3 – Candida albicans

Based on the relationships with the human host, microorganisms may be classified as commensals or pathogens.

  • Commensals cause no harm to the host, with which they establish a symbiotic relationship that generally brings benefits to both.
  • On the contrary, pathogens are able to cause diseases, but fortunately represent a small percentage of the human microbiota. These microorganisms establish a symbiosis with the human host and benefit from it at the expense of the host. They can cause disease:

if they move from their niche, such as the intestine, into another one where they do not usually reside, such as the vagina or bladder (as in the case of Candida albicans, normally present in the intestine, but in very small quantities);
in patients with impaired immunological defenses, such as after an immunosuppressive therapy.

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Functions of the human microbiota

Human Microbiota
Fig. 4 – Bifidobacterium longum

Sometimes referred to as “the forgotten organ“, human microbiota, mainly with its intestinal bacterial members, plays many important functions that can lead to nutritional, immunological, and developmental benefits, but can also cause diseases. Here are some examples.

  • It is involved in the development of the gastrointestinal system of the newborn, as shown by experiments carried out on germ-free animals in which, for example, the thickness of the intestinal mucosa is thinner than that of colonized animals, therefore more easily subject to rupture.
  • It contributes to energy harvest from nutrients, due to its ability to ferment indigestible carbohydrates, promote the absorption of monosaccharides and the storage of the derived energy. This has probably been a very strong evolutionary force that has played a major role in favor of the fact that these bacteria became our symbionts.
  • It contributes to the maintenance of the acidic pH of the skin and in the colon.
  • It is involved in the metabolism of xenobiotics and several polyphenols.
  • It improves water and mineral absorption in the colon.
  • It increases the speed of intestinal transit, slower in germ-free animals.
  • It has an important role in resistance to colonization by pathogens, primarily in the vagina and gut.
  • It is involved in the biosynthesis of isoprenoids and vitamins through the methylerythritol phosphate pathway.
  • It stimulates angiogenesis.
  • In the intestinal tract, it interacts with the immune system, providing signals for promoting the maturation of immune cells and the normal development of immune functions. And this is perhaps the most important effect of the symbiosis between the human host and microorganisms. Experiments carried out on germ-free animals have shown, for example, that:

macrophages, the cells that engulf pathogens and then present their antigens to the immune system, are found in much smaller amounts than those present in the colonized intestine, and if placed in the presence of bacteria they fail to find and therefore engulf them, unlike macrophages extracted from a colonized intestine;
there is not the chronic non-specific inflammation, present in the normal intestine as a result of the presence of bacteria (and of what we eat).

  • Changes in its composition can contribute to the development of obesity and metabolic syndrome.
  • It protects against the development of type I diabetes.
  • Many diseases, both in children and adults, such as stomach cancer, lymphoma of mucosa-associated lymphoid tissue, necrotizing enterocolitis (an important cause of morbidity and mortality in premature babies) or chronic intestinal diseases, are, and others seem to be, related to the gut microbiota.

In conclusion, it seems very likely that the human body represents a superorganism, result of years of evolution and made up of human cells, and the resulting metabolic and physiological capacities, as well as an additional organ, the microbiota.

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Human Microbiome Project

Human Microbiota
Fig. 5- Human Microbiome Project

The bacterial component of the human microbiota is the subject of most studies including a large-scale project started in 2008 called “Human Microbiome Project“, whose aim is to characterize the microbiome associated with multiple body sites, such as the skin, mouth, nose, vagina and intestine, in 242 healthy adults. These studies have shown a great variability in the composition of the human microbiota; for example, twins share less than 50% of their bacterial taxa at the species level, and an even smaller percentage of viruses. The factors that shape the composition of bacterial communities begin to be understood: for example, the genetic characteristics of the host play an important, although this is not true for the viral community. And metagenomic studies have shown that, despite the great interpersonal variability in microbial community composition, there is a core of shared genes encoding signaling and metabolic pathways. It appears namely that the assembly and the structure of the microbial community does not occur according to the species but the more functional set of genes. Therefore, disease states of these communities might be better identified by atypical distribution of functional classes of genes.

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Effects of antibiotics on the human microbiota

Human Microbiota
Fig. 6 – Clostridium difficile

The microbiota in healthy adult humans is generally stable over time. However, its composition can be altered by factors such as dietary changes, urbanization, travel, and especially the use of broad-spectrum antibiotics. Here are some examples of the effects of antibiotic treatments.

  • There is a long-term reduction in microbial diversity.
  • The taxa affected vary from individual to individual (even up to a third of the taxa).
  • Several taxa do not recover even after 6 months from treatment.
  • Once the bacterial communities have reshaped, a reduced resistance to colonization occurs. This allows foreign and/or pathogen bacteria, able to grow more than the commensals, to cause permanent changes in human microbiota structure, as well as acute diseases, such as the dangerous pseudomembranous colitis, and chronic diseases, as it is suspected for asthma following the use and abuse of antibiotics in childhood. Moreover, their repeated use has been suggested to increase the pool of antibiotic-resistance genes in our microbiome. In support of this hypothesis, a decrease in the number of antibiotic-resistant pathogens has been observed in some European countries following the reduction in the number of antibiotics prescribed.

Finally, you must not underestimate the fact that the intestinal microflora is involved in many chemical transformations, and its alteration could be implicated in the development of cancer and obesity. However, regarding use of antibiotics, you should be underlined that if western population has a life expectancy higher than in the past is also because you do not die of infectious diseases!

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References

Burke C., Steinberg P., Rusch D., Kjelleberg S., and Thomas T. Bacterial community assembly based on functional genes rather than species. Proc Natl Acad Sci USA 2011;108:14288-93. doi:10.1073/pnas.1101591108

Clemente J.C., Ursell L.K., Wegener Parfrey L., and Knight R. The impact of the gut microbiota on human health: an integrative view. Cell 2012;148:1258-70. doi:10.1016/j.cell.2012.01.035

Gill S.R., Pop M., Deboy R.T., Eckburg P.B., Turnbaugh P.J., Samuel B.S., Gordon J.I., Relman D.A., Fraser-Liggett C.M., and Nelson K.E. Metagenomic analysis of the human distal gut microbiome. Science 2006;312:1355-59. doi:10.1126/science.1124234

Palmer C., Bik E.M., DiGiulio D.B., Relman D.A., and Brown P.O. Development of the human infant intestinal microbiota. PLoS Biol 2007;5(7):e177. doi:10.1371/journal.pbio.0050177

The Human Microbiome  Project

Turnbaugh P.J., Gordon J.I. The core gut microbiome, energy balance and obesity. J Physiol 2009;587:4153-58. doi:10.1113/jphysiol.2009.174136

Zhang, T., Breitbart, M., Lee, W., Run, J.-Q., Wei, C., Soh, S., Hibberd, M., Liu, E., Rohwer, F., Ruan, Y. Prevalence of plant viruses in the RNA viral community of human feces. PLoS Biol 2006;4(1):e3. doi:10.1371/journal.pbio.0040003

Flavonoid biosynthesis pathway

Flavonoid biosynthesis pathway: contents in brief

The flavonoid biosynthetic pathway in plants

Flavonoid Biosynthesis
Fig. 1 – Fig. 1 – Biosynthesis of Flavonoids

The biosynthesis of flavonoids, probably the best characterized pathway of plant secondary metabolism, is part of the phenylpropanoid pathway that, in addition to flavonoids, leads to the formation of a wide range of phenolic compounds, such as hydroxycinnamic acids, stilbenes, lignans and lignins.
Flavonoid biosynthesis is linked to primary metabolism through both mitochondria- and plastid-derived molecules. Since it seems that most of the involved enzymes characterized to date operate in protein complexes located in the cell cytosol, these molecules must be exported to the cytoplasm to be used.
The end products are transported to different intracellular or extracellular locations, with flavonoids involved in pigmentation usually transported into the vacuoles.
The biosynthesis of this group of polyphenols requires one p-coumaroyl-CoA and three malonyl-CoA molecules as initial substrates.

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Biosynthesis of p-coumaroyl-CoA

Flavonoid Biosynthesis
Fig. 2 – p-Cumaroyl-CoA

p-Coumaroyl-CoA is the pivotal branch-point metabolite in the phenylpropanoid pathway, being the precursor of a wide variety of phenolic compounds, both flavonoid and non-flavonoid polyphenols.
It is produced from phenylalanine via three reactions catalyzed by cytosolic enzymes collectively called group I or early-acting enzymes, in order of action:

  • phenylalanine ammonia lyase (EC 4.3.1.24);
  • cinnamate 4-hydroxylase or cinnamic acid 4-hydroxylase (EC 1.14.13.11);
  • 4-cumarato: CoA ligase or hydroxycinnamic: CoA ligase (EC 6.2.1.12).

They seems to be associated in a multienzyme complex anchored to the endoplasmic reticulum membrane. The anchoring is probably ensured by cinnamate 4-hydroxylase that inserts its N-terminal domain into the membrane of the endoplasmic reticulum itself. These complexes, referred to as “metabolons”, allow the product of a reaction to be channeled directly as substrate to the active site of the enzyme that catalyzes the consecutive reaction in the metabolic pathway.
With the exception of cinnamate 4-hydroxylase, the enzymes which act downstream of phenylalanine ammonia lyase are encoded by small gene families in all species analyzed so far.
The different isoenzymes show distinct temporal, tissue, and elicitor-induced patterns of expression. It seems, in fact, that each member of each family can be used mainly for the synthesis of a specific compound, thus acting as a control point for carbon flux among the metabolic pathways leading to lignan, lignin, and flavonoid biosynthesis.

Note: phenylalanine is a product of the shikimic acid pathway, which converts simple precursors derived from carbohydrate metabolism, phosphoenolpyruvate and erythrose-4-phosphate, into the aromatic amino acids phenylalanine, tyrosine and tryptophan. Unlike plants and microorganisms, animals do not possess the shikimic acid pathway, and are not able to synthesize the three above-mentioned amino acids, which are therefore essential nutrients.

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Phenylalanine ammonia lyase (PAL)

Flavonoid Biosynthesis
Fig. 3 – Phenylalanine Ammonia Lyase

It is one of the most studied and best characterized enzymes of plant secondary metabolism. It requires no cofactors and catalyzes the reaction that links primary and secondary metabolism: the deamination of phenylalanine to trans-cinnamic acid, with the release of nitrogen as ammonia and introduction of a trans double bond between carbon atoms 7 and 8 of the side chain. Therefore, it directs the flow of carbon from the shikimic acid pathway to the different branches of the phenylpropanoid pathway. The released ammonia is probably fixed in the reaction catalyzed by glutamine synthetase.
The enzyme from monocots is also able to act as tyrosine ammonia lyase (EC 4.3.1.25), converting tyrosine to p-coumaric acid directly, (therefore without the 4-hydroxylation step), but with a lower efficiency.
In all plant species investigated,  several copies of phenylalanine ammonia lyase gene are found, copies that probably respond differentially to internal and external stimuli. Indeed, gene transcription, and then enzyme activity, are under the control of both internal developmental and external environmental stimuli. Here are some examples that require increased enzyme activity.

  • The flowering.
  • The  production of lignin to strengthen the secondary wall of xylem cells.
  • The production of flower pigments that attract pollinators.
  • Pathogen infections, that require the production of phenylpropanoid phytoalexins, or exposure to UV rays.

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Cinnamate 4-hydroxylase (C4H)

Flavonoid Biosynthesis
Fig. 4 – Cinnamate 4-Hydroxylase

It belongs to the cytochrome P450 superfamily (EC 1.14.-.-), is a microsomal monooxygenase containing a heme cofactor, and dependent on both O2  and NADPH. It catalyzes the formation of p-coumaric acid  through the introduction of a hydroxyl group in 4-position of trans-cinnamic acid (this hydroxyl group is present in most flavonoids). This reaction is also part of the biosynthesis of hydroxycinnamic acids.
Increases in transcription rates and enzyme activity are observed in correlation with the synthesis of phytoalexins (in response to fungal infections), lignification as well as wounding.

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4-Coumarate:CoA ligase (4CL)

Flavonoid Biosynthesis
Fig. 5 – 4-Coumarate:CoA Ligase

With Mg2+ as a cofactor, it catalyzes the ATP-dependent activation of the carboxyl group of p-coumaric acid and other hydroxycinnamic acids, metabolically rather inert molecules, through the formation of the corresponding CoA-thioester. Generally, p-coumaric acid and caffeic acid are the preferred substrates, followed by ferulic acid and 5-hydroxyferulic acid, low activity against trans-cinnamic acid and none against sinapic acid. These CoA-thioesters are able to enter various reactions such as:

  • reduction to alcohol (monolignols) or aldehydes;
  • stilbene and flavonoid biosynthesis;
  • transfer to acceptor molecules.

It should finally be pointed out that the activation of the carboxyl group can also be obtained through an UDP-glucose-dependent transfer to glucose.

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Biosynthesis of malonyl-CoA

Malonyl-CoA does not derived from the phenylpropanoid pathway, but from the reaction catalyzed by acetyl-CoA carboxylase (EC 6.4.1.2, the cytosolic form, see below). The enzyme, with biotin and Mg2+ as cofactors, catalyzes the ATP-dependent carboxylation of acetyl-CoA, using bicarbonate as a source of carbon dioxide (CO2).

Flavonoid Biosynthesis
Fig. 6 – Acetyl-CoA Carboxylase

It is found both in the plastids, where it participates in the synthesis of fatty acids, and the cytoplasm, and is the latter that catalyzes the formation of malonyl-CoA that is used in the biosynthesis of flavonoids and other compounds. Increases in the transcription rate of the gene and enzyme activity are induced in response to stimuli that increase the biosynthesis of these polyphenols, such as exposure to pathogenic fungi or UV-rays.
In turn, acetyl-CoA is produced in plastids, mitochondria, peroxisomes and cytosol through different metabolic pathways. The molecules used in the biosynthesis of malonyl-CoA, and therefore of the flavonoids, are  the cytosolic ones, produced in the reaction catalyzed by ATP-citrate lyase (EC 2.3.3.8) that cleaves citrate, in the presence of CoA and ATP, to form oxaloacetate and acetyl-CoA, plus ADP and inorganic phosphate.

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The first steps in flavonoid biosynthesis

Flavonoid Biosynthesis
Fig. 7 – Naringenin Chalcone

The first step in flavonoid biosynthesis is catalyzed by chalcone synthase (EC 2.3.1.74), an enzyme anchored to the endoplasmic reticulum and with no known cofactors.
From one p-coumaroyl-CoA and three malonyl-CoA, it catalyzes sequential condensation and decarboxylation reactions in the course of which a polyketide intermediate is formed. The polyketide undergoes cyclizations and aromatizations leading to the formation of the A ring. The product of the reactions is naringenin chalcone (2′,4,4′,6′-tetrahydroxychalcone), a 6′-hydroxychalcone and the first flavonoid to be synthesized by plants.
The reaction, cytosolic, is irreversible due to the release of three CO2 and 4 CoA.

Flavonoid Biosynthesis
Fig. 8 – Basic Flavonoid Skeleton

The B ring and the three-carbon bridge of the molecule originate from p-coumaroyl-CoA (and therefore from phenylalanine), the A ring from the three malonyl-CoA units.
Also 6’-deoxychalcone can be produced; its synthesis is thought to involve an additional reduction step catalyzed by polyketide reductase (EC. 1.1.1.-).
Chalcone synthase from some plant species, such as barley (Hordeum vulgare), accepts as substrates also caffeoil-CoA, feruloil-CoA and cinnamoyl-CoA.
It is the most abundant enzyme of the phenylpropanoid pathway, probably because it has a low catalytic activity, and, in fact, is considered to be the rate-limiting enzyme in flavonoid biosynthesis.
As for phenylalanine ammonia lyase, chalcone synthase gene expression is under the control of both internal and external factors. In some plants, one or two isoenzymes are found, while in others up to 9.
Chalcone synthase belongs to polyketide synthase group, present in bacteria, fungi and plants. These enzymes are able to catalyze the production of polyketide chains through sequential condensations of acetate units provided by malonyl-CoA units. They also includes stilbene synthase (EC 2.3.1.146), which catalyzes the formation of resveratrol, a non flavonoid polyphenol compound that has attracted much interest because of its potential health benefits.

Flavonoid Biosynthesis
Fig. 9 – (2S)-Naringenin

Generally, chalcones do not accumulate in plants because naringenin chalcone is converted to (2S)-naringenin, a flavanone, in the reaction catalyzed by chalcone isomerase (EC 5.5.1.6). The enzyme, the first of the flavonoid biosynthesis to be discovered, catalyzes a stereospecific isomerization and closes the C ring. Two types of chalcone isomerases are known, called type I and II. Type I enzymes use only 6′-hydroxychalcone substrates, like naringenin chalcone, while type II, prevalent in legumes, use both 6′-hydroxy- and 6′-deoxychalcone substrates.
It should be noted that with 6′-hydroxychalcones, isomerization can also occur nonenzymically to form a racemic mixture, both in vitro and in vivo, enough to allow a moderate synthesis of anthocyanins. On the contrary, under physiological conditions 6′-deoxychalcones are stable, and so the activity of type II chalcone isomerases is required to form flavanones.
The enzyme increases the rate of the reaction of 107 fold compared to the spontaneous reaction, but with a higher kinetics for the 6′-hydroxychalcones than 6′-deoxychalcones. Finally, it produces (2S)-flavanones, which are the biosynthetically required enantiomers.
As other enzymes in flavonoid biosynthesis, also chalcone isomerase gene expression is subject to strict control. And, as phenylalanine ammonia lyase and chalcone synthase, it is induced by elicitors.

Flavonoid Biosynthesis
Fig. 10 – Dihydrokaempferol

In the reaction catalysed by flavanone-3β-hydroxylase (EC 1.14.11.9), (2S)-flavanones undergo a stereospecific isomerization that converts them into the respective (2R,3R)-dihydroflavonols. In particular, naringenin is converted into dihydrokaempferol.
The enzyme is a cytosolic non-heme-dependent dioxygenase, dependent on Fe2+ and 2-oxoglutarate, and therefore belonging to the family of 2-oxoglutarate-dependent dioxygenase (which distinguishes them from the other hydroxylases of the flavonoid biosynthetic pathway which are cytochrome P450 enzymes).
Naringenin chalcone, (2S)-naringenin, and dihydrokaempferol are central intermediates in flavonoid biosynthesis, since they act as branch-point compounds from which the synthesis of distinct flavonoid subclasses can occur. For example, directly or indirectly:

Not all of these side metabolic pathways are present in every plant species, or are active within each tissue type of a given plant. Like enzymes previously seen, the activity of those involved in these “side-routes” is subject to strict control, resulting in a tissue-specific profile of flavonoid compounds. For example, grape seeds, flesh and skin have distinct anthocyanin, catechin, flavonol and condensed tannin profiles, whose synthesis and accumulation are strictly and temporally coordinated during the ripening process.

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References

Andersen Ø.M., Markham K.R. Flavonoids: chemistry, biochemistry, and applications. CRC Press Taylor & Francis Group, 2006

de la Rosa L.A., Alvarez-Parrilla E., Gonzàlez-Aguilar G.A. Fruit and vegetable phytochemicals: chemistry, nutritional value, and stability. 1th Edition. Wiley J. & Sons, Inc., Publication, 2010

Heldt H-W. Plant biochemistry – 3th Edition. Elsevier Academic Press, 2005

Vogt T. Phenylpropanoid biosynthesis. Mol Plant 2010;3(1):2-20. doi:http://dx.doi.org/10.1093/mp/ssp106

Wink M. Biochemistry of plant secondary metabolism – 2nd Edition. Annual plant reviews (v. 40), Wiley J. & Sons, Inc., Publication, 2010

Lignans: definition, chemical structure, biosynthesis, metabolism, foods

Lignans: contents in brief

What are lignans?

Lignans are a subgroup of non-flavonoid polyphenols.
They are widely distributed in the plant kingdom, being present in more than 55 plant families, where they act as antioxidants and defense molecules against pathogenic fungi and bacteria.
In humans, epidemiological and physiological studies have shown that they can exert positive effects in the prevention of lifestyle-related diseases, such as type II diabetes and cancer. For example, an increased dietary intake of these polyphenols correlates with a reduction in the occurrence of certain types of estrogen-related tumors, such as breast cancer in postmenopausal women.
In addition, some lignans have also aroused pharmacological interest. Examples are:

  • podophyllotoxin, obtained from plants of the genus Podophyllum (Berberidaceae family); it is a mitotic toxin whose derivatives have been used as chemotherapeutic agents;
  • arctigenin and tracheologin, obtained from tropical climbing plants; they have antiviral properties and have been tested in the search for a drug to treat AIDS .

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Chemical structure of lignans

Lignans
Fig. 1 – Phenylpropane unit

Their basic chemical structure consists of two phenylpropane units linked by a C-C bond between the central atoms of the respective side chains (position 8 or β), also called β-β’ bond. 3-3′, 8-O-4′, or 8-3′ bonds are observed less frequently; in these cases the dimers are called neolignans. Hence, their chemical structure is referred to as (C6-C3)2, and they are included in the phenylpropanoid group, as well as their precursors: the hydroxycinnamic acids (see below).
Based on their carbon skeleton, cyclization pattern, and the way in which oxygen is incorporated in the molecule skeleton, they can be divided into 8 subgroups: furans, furofurans, dibenzylbutanes, dibenzylbutyrolactones, dibenzocyclooctadienes, dibenzylbutyrolactols, aryltetralins and arylnaphthalenes. Furthermore, there is considerable variability regarding the oxidation level of both the propyl side chains and the aromatic rings.
They are not present in the free form in nature, but linked to other molecules, mainly as glycosylated derivatives.
Among the most common lignans, secoisolariciresinol (the most abundant one), lariciresinol, pinoresinol, matairesinol and 7-hydroxymatairesinol are found.

Note: they occur not only as dimers but also as more complex oligomers, such as dilignans and sesquilignans.

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Biosynthesis of lignans

Lignans
Fig. 2 – Lignan Biosynthesis

In this section, we will examine the synthesis of some of the most common lignans.
The pathway starts from 3 of the 4 most common dietary hydroxycinnamic acids: p-coumaric acid, sinapic acid, and ferulic acid (caffeic acid is not a precursor of this subgroup of polyphenols). Therefore, they arise from the shikimic acid pathway, via phenylalanine.
The first three reactions reduce the carboxylic group of the hydroxycinnamates to alcohol group, with formation of the corresponding alcohols, called monolignols, that is, p-coumaric alcohol, sinapyl alcohol and coniferyl alcohol. These molecules also enter the pathway of lignin biosynthesis.

  • The first step, which leads to the activation of the hydroxycinnamic acids, is catalysed by hydroxycinnamate:CoA ligases, commonly called p-coumarate:CoA ligases (EC 6.2.1.12), with formation of the corresponding hydroxycinnamate-CoAs, namely, feruloil-CoA, p- coumaroyl-CoA and sinapil-CoA.
  • In the second step, a NADPH-dependent cinnamoyl-CoA: oxidoreductase, also called cinnamoyl-CoA reductase (EC1.2.1.44) catalyzes the formation of the corresponding aldehydes, and the release of coenzyme A.
  • In the last step, a NADPH-dependent cinnamyl alcohol dehydrogenase, also called monolignol dehydrogenase (EC 1.1.1.195), catalyzes the reduction of the aldehyde group to an alcohol group, with the formation of the aforementioned monolignols.
Lignans
Fig. 3 – (-)-Matairesinol

The next step, the dimerization of monolignols, involves the intervention of stereoselective mechanisms, or, more precisely, enantioselective mechanisms. In fact, most of the plant lignans exists as (+)- or (-)-enantiomers, whose relative amounts can vary from species to species, but also in different organs on the same plant, depending on the type of reactions involved.
The dimerization can occur through enzymatic reactions involving laccases (EC 1.10.3.2). These enzymes catalyze the formation of radicals that, dimerizing, form a racemic mixture. However, this does not explain how the enantiomeric mixtures found in plants are formed. The most accepted mechanism to explain the stereospecific synthesis involves the action of the laccase and of a protein able to direct the synthesis toward one or the other of the two enantiomeric forms: the dirigent protein. The reaction scheme might be: the enzyme catalyzes the synthesis of phenylpropanoid radicals that are orientated in such a way to obtain the desired stereospecific coupling by the dirigent protein.
For example, pinoresinol synthase, consisting of laccase and dirigent protein, catalyzes the stereospecific synthesis of (+)-pinoresinol from two units of coniferyl alcohol. (+)-Pinoresinol, in two consecutive stereospecific reactions catalyzed by NADPH-dependent pinoresinol/lariciresinol reductase (EC 1.23.1.2), is first reduced to (+)-lariciresinol, and then to (-)-secoisolariciresinol. (-)-Secoisolariciresinol, in the reaction catalyzed by NAD(P)-dependent secoisolariciresinol dehydrogenase (EC 1.1.1.331) is oxidized to (-)-matairesinol.

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Metabolism of lignans by human gut microbiota

Their importance to human health is due largely to their metabolism by colonic microbiota, which carries out deglycosylations, para-dehydroxylations, and meta-demethylations without enantiomeric inversion. Indeed, this metabolization leads to the formation molecules with a modest estrogen-like activity (phytoestrogens), a situation similar to that observed with some isoflavones, such as those of soybean, some coumarins, and some stilbenes. These active metabolites are the so-called “mammalian lignans or enterolignans”, such as the aglycones of enterodiol and enterolactone, formed from secoisolariciresinol and matairesinol, respectively.
Studies conducted on animals fed diets rich in lignans have shown their presence as intact molecules, in low concentrations, in serum, suggesting that they may be absorbed as such from the intestine. These molecules exhibit estrogen-independent actions, both in vivo and in vitro, such as inhibition of angiogenesis, reduction of diabetes, and suppression of tumor growth.
Note: the term “phytoestrogen” refers to molecules with estrogenic or antiandrogenic activity, at least in vitro.

Once absorbed, they enter the enterohepatic circulation, and, in the liver, may undergo phase II reactions and be sulfated or glucuronidated, and finally excreted in the urine.

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Food rich in lignans

Lignans
Fig. 4 – (-)-Secoisolariciresinol

The richest dietary source is flaxseed (linseed), that contains mainly secoisolariciresinol, but also lariciresinol, pinoresinol and matairesinol in good quantity (for a total amount of more than 3.7 mg/100 g dry weight). They are also found in sesame seeds.
Another important source is whole grains.
They are also present in other foods, but in concentrations from one hundred to one thousand times lower than those of flaxseed. Examples are:

  • beverages, generally more abundant in red wine, followed in descending order by black tea, soy milk and coffee;
  • fruits, such as apricots, pears, peaches, strawberries;
  • among vegetables, Brassicaceae, garlic, asparagus and carrots;
  • lentils and beans.

Their presence in grains and, to a lesser extent in red wine and fruit, makes them, at least in individuals who follow a Mediterranean-style eating pattern, the main source of phytoestrogens.

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References

Andersen Ø.M., Markham K.R. Flavonoids: chemistry, biochemistry, and applications. CRC Press Taylor & Francis Group, 2006

de la Rosa L.A., Alvarez-Parrilla E., Gonzàlez-Aguilar G.A. Fruit and vegetable phytochemicals: chemistry, nutritional value, and stability. 1th Edition. Wiley J. & Sons, Inc., Publication, 2010

Heldt H-W. Plant biochemistry – 3th Edition. Elsevier Academic Press, 2005

Manach C., Scalbert A., Morand C., Rémésy C., and Jime´nez L. Polyphenols: food sources and bioavailability. Am J Clin Nutr 2004;79(5):727-47 [Abstract]

Tsao R. Chemistry and biochemistry of dietary polyphenols. Nutrients 2010;2:1231-46. doi:10.3390/nu2121231

Satake H, Koyama T., Bahabadi S.E., Matsumoto E., Ono E. and Murata J. Essences in metabolic engineering of lignan biosynthesis. Metabolites 2015;5:270-90. doi:10.3390/metabo5020270

van Duynhoven J., Vaughan E.E., Jacobs D.M., Kemperman R.A., van Velzen E.J.J, Gross G., Roger L.C., Possemiers S., Smilde A.K., Doré J., Westerhuis J.A.,and Van de Wiele T. Metabolic fate of polyphenols in the human superorganism. PNAS 2011;108(suppl. 1):4531-8. doi:10.1073/pnas.1000098107

Wink M. Biochemistry of plant secondary metabolism – 2nd Edition. Annual plant reviews (v. 40), Wiley J. & Sons, Inc., Publication, 2010

Hydroxycinnamic acids: definition, chemical structure, synthesis, foods

Hydroxycinnamic acids: contents in brief

What are hydroxycinnamic acids?

Hydroxycinnamic acids or hydroxycinnamates are phenolic compounds belonging to non-flavonoid polyphenols.
They are present in all parts of fruits and vegetables although the highest concentrations are found in the outer part of ripe fruits, concentrations that decrease during ripening, while the total amount increases as the size of the fruits increases.

Their dietary intake has been associated with the prevention of the development of chronic diseases such as:

  • cardiovascular disease;
  • cancer;
  • type-2 diabetes.

These effects do seem to be due not only to their high antioxidant activity (that depends upon the hydroxylation pattern of the aromatic ring, see below), but also to other mechanisms of action such as, e.g., the reduction of intestinal absorption of glucose or the modulation of secretion of some gut hormones.

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Chemical structure of hydroxycinnamic acids

Hydroxycinnamic Acids
Fig. 1 – C6-C3 Skeleton

Their basic structure is a benzene ring to which a three carbon chain is attached, structure that is referred to as C6-C3. Therefore they can be included in the phenylpropanoid group.
The main dietary hydroxycinnamates are:

  • caffeic acid or 3,4-dihydroxycinnamic acid;
  • ferulic acid or 4-hydroxy-3-methoxycinnamic acid;
  • sinapic acid or 4-hydroxy-3,5-dimethoxycinnamic acid;
  • p-coumaric acid or 4-coumaric acid or 4-hydroxycinnamic acid.

In nature, they are associated with other molecules to form, e.g., glycosylated derivatives or esters of tartaric acid, quinic acid, or shikimic acid. In addition, several hundreds of anthocyanins acylated with the aforementioned hydroxycinnamates have been identified (in descending order with p-coumaric acid, more than 150, caffeic acid, about 100, ferulic acid, about 60, and sinapic acid, about 25). They are rarely present in the free form, except in processed foods that have undergone fermentation, sterilization or freezing. For example, an overlong storage of blood orange fruits causes a massive hydrolysis of hydroxycinnamic derivatives to free acids, and this in turn could lead to the formation of malodorous compounds such as vinyl phenols, indicators of too advanced senescence of the fruit.

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Biosynthesis of hydroxycinnamic acids

Hydroxycinnamic Acids
Fig. 2 – Biosynthesis of Hydroxycinnamates

Hydroxycinnamate biosynthesis consists of a series of enzymatic reactions subsequent to that catalyzed by phenylalanine ammonia lyase (PAL). This enzyme catalyzes the deamination of phenylalanine to yield trans-cinnamic acid, so linking the aromatic amino acid to the hydroxycinnamic acids and their activated forms.
In the first step, a hydroxyl group is attached at the 4-position of the aromatic ring of trans-cinnamic acid to form p-coumaric acid (reaction catalysed by cinnamic acid 4-hydroxylase). The addition of a second hydroxyl group at the 3-position of the ring of p-coumaric acid leads to the formation of caffeic acid (reaction catalysed by p-coumarate 3-hydroxylase or phenolase), while the O-methylation of the hydroxyl group at the 3-position yields ferulic acid (reaction catalyzed by catechol-O-methyltransferase). In turn, ferulic acid is converted into sinapic acid through two reactions: a hydroxylation at the 5-position to form 5-hydroxy ferulic acid (reaction catalysed by ferulate 5-hydroxylase), and the subsequent O-methylation of the same hydroxyl group (reaction catalyzed by catechol-O-methyltransferase).
Hydroxycinnamic acids are not present in high quantities since they are rapidly converted to glucose esters or coenzyme A (CoA) esters, in reactions catalyzed by O-glucosyltransferases and hydroxycinnamate:CoA ligases, respectively. These activated intermediates are branch points, being able to participate in a wide range of reactions such as condensation with malonyl-CoA to form flavonoids, or the NADPH-dependent reduction to form lignans (precursors of lignin).

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The main hydroxycinnamic acids in foods

Kiwis, blueberries, plums, cherries, apples, pears, chicory, artichokes, carrots, lettuce, eggplant, wheat and coffee are among the richest sources.

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Caffeic acid

Hydroxycinnamic Acids
Fig. 3 – Caffeic Acid

It is generally, both in the free form and bound to other molecules, the most abundant hydroxycinnamic acid in vegetables and most of the fruits, where it represents between 75 and 100% of the hydroxycinnamates.
The richest sources are coffee (drink), carrots, lettuce, potatoes, even sweet ones, and berries such as blueberries, cranberries and blackberries.
Smaller quantities are present in grapes and grape-derived products, orange juice, apples, plums, peaches, and tomatoes.
Caffeic acid and quinic acid bind to form chlorogenic acid, present in many fruit and in high concentration in coffee.

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Ferulic acid

Hydroxycinnamic Acids
Fig. 4 – Ferulic Acid

It is the most abundant hydroxycinnamic acid in cereals, which are also its main dietary source.
In wheat grain, its content is between 0.8 and 2 g/kg dry weight, which represents up to 90% of the total polyphenols. It is found chiefly, up to 98% of the total content, in the aleurone layer and pericarp (that is, the outer parts of the grain), and therefore its content in wheat flours depends upon the degree of refining, while the main source is obviously the bran. The molecule is present mainly in the trans form, and esterified with arabinoxylans and hemicelluloses. And in fact, in wheat bran the soluble free form represents only about 10% of its total amount. Dimers were also found, which form bridge structures between chains of hemicellulose.
In fruits and vegetables, ferulic acid is much less common than caffeic acid. The main sources are asparagus, eggplant and broccoli; lower quantities are found in blackberries, blueberries, cranberries, apples, carrots, potatoes, beets, coffee and orange juice.

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Sinapic acid

Hydroxycinnamic Acids
Fig. 5 – Sinapic Acid

The highest amounts are found in citrus peel and seeds (in orange juice, the amount is much lower); appreciable quantities in Chinese cabbage and in some varieties of cranberries.

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p-Coumaric acid

Hydroxycinnamic Acids
Fig. 6 – p-Coumaric Acid

High amounts are present in eggplant, the richest source, broccoli and asparagus; other sources are sweet cherries, plums, blueberries, cranberries, citrus peel and seeds, and orange juice.

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References

Andersen Ø.M., Markham K.R. Flavonoids: chemistry, biochemistry, and applications. CRC Press Taylor & Francis Group, 2006

de la Rosa L.A., Alvarez-Parrilla E., Gonzàlez-Aguilar G.A. Fruit and vegetable phytochemicals: chemistry, nutritional value, and stability. 1th Edition. Wiley J. & Sons, Inc., Publication, 2010

Manach C., Scalbert A., Morand C., Rémésy C., and Jime´nez L. Polyphenols: food sources and bioavailability. Am J Clin Nutr 2004;79(5):727-47 [Abstract]

Preedy V.R. Coffee in health and disease prevention. Academic Press, 2014  [Google eBook]

Zhao Z.,  Moghadasian M.H. Bioavailability of hydroxycinnamates: a brief review of in vivo and in vitro studies. Phytochem Rev 2010;9(1):133-145. doi:10.1007/s11101-009-9145-5

Polyphenols in grapes and wine: chemical composition and biological activities

Polyphenols in grapes and wine: contents in brief

Polyphenols in Grapes
Fig. 1 – Red Grapes

The consumption of grapes and grape-derived products, particularly red wine but only at meals, has been associated with numerous health benefits, which include, in addition to the antioxidant/antiradical effect, also anti-inflammatory, cardioprotective, anticancer, antimicrobial, and neuroprotective activities.
Grapes contain many nutrients such as sugars, vitamins, minerals, fiber and phytochemicals. Among the latter, polyphenols are the most important compounds in determining the health effects of the fruit and derived products.
Indeed, grapes are among the fruits with highest content in polyphenols, whose composition is strongly influenced by several factors such as:

  • cultivar;
  • climate;
  • exposure to disease;
  • processing

Nowadays, the main species of grapes cultivated worldwide are: European grapes, Vitis vinifera, North American grapes, Vitis rotundifolia and Vitis labrusca, and French hybrids.
Note: grapes are not a fruit but an infructescence, that is, an ensemble of fruits (berries): the bunch of grapes. In turn, it consists of a peduncle, a rachis, cap stems or pedicels, and berries.

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What are polyphenols in grapes and wine?

Polyphenols in red grapes and wine are significantly higher, both in quantity and variety, than in white ones. This, according to many researchers, would be the basis of the more health benefits related to the consumption of red grapes and wine than white grapes and derived products.
Polyphenols in grapes and wine are a complex mixture of flavonoid compounds, the most abundant group, and non-flavonoid compounds.
Among flavonoids, they are found:

Among non-flavonoid polyphenols:

Most of the flavonoids present in wine derive from the epidermal layer of the berry skin, while 60-70% of the total polyphenols are present in the grape seeds. It should be noted that more than 70% of grape polyphenols are not extracted and remain in the pomace.
The complex chemical interactions that occur between these compounds, and between them and the other compounds of different nature present in grapes and wine, are probably essential in determining both the quality of the grapes and wine and the broad spectrum of therapeutic effects of these foods.
In wine, the mixture of polyphenols play important functions being able to influence:

  • bitterness;
  • astringency;
  • red color, of which they are among the main responsible;
  • sensitivity to oxidation, being molecules easily oxidizable by atmospheric oxygen.

Finally, they act as preservatives and are the basis of long aging.

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Polyphenols in grapes and wine: anthocyanins

They are flavonoids widely distributed in fruits and vegetables.
They are primarily located in the berry skin (in the outer layers of the hypodermal tissue), to which they confer color, having a hue that varies from red to blue. In some varieties, called “teinturier”, they also accumulate in the flesh of the berry.
There is a close relationship between berry development and the biosynthesis of anthocyanins. The synthesis starts at veraison (when the berry stops growing and changes its color), causes a color change of the berry that turns purple, and reaches the maximum levels at complete ripening.
Among wine flavonoids, they are one of the most potent antioxidants.
Each grape species and cultivars has a unique composition of anthocyanins. Moreover, in grapes of Vitis vinifera, due to a mutation in the gene coding for 5-O-glucosyltransferase, mutation that determines the synthesis of an inactive enzyme, only 3-monoglucoside derivatives are synthesized, while in other species  the glycosylation at position 5 also occurs. Interestingly, 3-monoglucoside derivatives are more intensely colored than 3,5-diglucoside derivatives.

Polyphenols in Grapes
Fig. 2 – Malvidin-3-glucoside

In red grapes and wine, the most abundant anthocyanins are the 3-monoglucosides of malvidin (the most abundant one both in grapes and wine), petunidin, delphinidin, peonidin, and cyanidin. In turn, the hydroxyl group at position 6 of the glucose can be acylated with an acetyl, caffeic or coumaric group, acylation that further enhances the stability.
Anthocyanidins, namely the non-conjugated molecules, are not present in grapes and in wine, except as traces.
Anthocyanins are scarcely present in white grapes and wine.
The composition of anthocyanins in wine is highly influenced both by the type of cultivar and by processing techniques, since they are present in wine as a result of extraction by maceration/fermentation processes. For this reason, wines deriving from similar varieties of grapes can have very different anthocyanin compositions.
Together with proanthocyanidins, they are the most important polyphenols in contributing to some organoleptic properties of red wine, as they are primarily responsible for astringency, bitterness, chemical stability against oxidation, as well as of the color of the young wine. In this regard, it should be underscored that with time their concentration decreases, while the color is due more and more to the formation of polymeric pigments produced by condensation of anthocyanins both among themselves and with other molecules.
During wine aging, proanthocyanidins and anthocyanins react to produce more complex molecules that can  partially precipitate.

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Polyphenols in grapes and wine: flavanols or catechins

Polyphenols in Grapes
Fig. 3 – Catechin

They are, together with condensed tannins, the most abundant flavonoids, representing up to 50% of the total polyphenols in white grapes and between 13% and 30% in red ones.
Their levels in wine depend on the type of cultivar.
Typically, the most abundant flavanol in wine is catechin, but epicatechin and epicatechin-3-gallate are also present.

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Polyphenols of grapes and wine: proanthocyanidins or condensed tannins

Polyphenols in Grapes
Fig. 4 – Procyanidin C1

Composed of catechin monomers, they are present in the berry skin, seeds and rachis of the bunch of grapes as:

  • dimers: the most common are procyanidins B1-B4, but also procyanidins B5-B8 can be present;
  • trimers: procyanidin C1 is the most abundant;
  • tetramers;
  • polymers, containing up to 8 monomers.

Their levels in wine depend on the type of grape varieties and wine-making technology, and, like anthocyanins, are much more abundant in red wines, in particular in aged wines, compared to white ones.
In addition, as previously said, together with anthocyanins, condensed tannins are important in determining some organoleptic properties of the wine.

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Polyphenols of grapes and wine: flavonols

They are present in a large variety of fruit and vegetables, even if in low concentrations.
They are the third most abundant group of flavonoids in grapes, after proanthocyanidins and catechins.
They are mainly present in the outer epidermis of the berry skin, where they play a role both in providing protection against UV-A and UV-B radiations and in copigmentation together with anthocyanins.
Flavanol synthesis begins in the sprout; the highest concentration is reached a few weeks after veraison, then it decreases as the berry increases in size.
Their total amount is very variable, with the red varieties often richer than the white ones.
In grapes, they are present as 3-glucosides and their composition depends on the type of grapes and cultivar:

  • the derivatives of quercetin, kaempferol and isorhamnetin are found in white grapes;
  • the derivatives of myricetin, laricitrin and syringetin are found, together with the previous ones, only in red grapes, due to the lack of expression in white grapes of the gene coding for flavonoid-3′,5′-hydroxylase.
Polyphenols in Grapes
Fig. 5 – Quercetin-3-glucoside

In general, the 3-glucosides and 3-glucuronides of quercetin are the major flavonols in most of the grape varieties. Conversely, quercetin-3-rhamnoside and quercetin aglycone are the major flavonols in muscadine grapes.
In wine and grape juice, unlike grapes, they are also found as aglycones, as a result of the acid hydrolysis that occurs during processing and storage. They are present in wine in a variable amount, and the major molecules are the glycosides of quercetin and myricetin, which alone represent 20-50% of the total flavonols in red wine.

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Polyphenols in grapes and wine: hydroxycinnamates

Polyphenols in Grapes
Fig. 6 – Ferulic Acid

Hydroxycinnamic acids are the main class of non-flavonoid polyphenols in grapes and the major polyphenols in white wine.
The most important are p-coumaric, caffeic, sinapic, and ferulic acids, present in wine as esters with tartaric acid.
They have antioxidant activity and in some white varieties of Vitis vinifera, together with flavonols, are the polyphenols mainly responsible for absorbing UV radiation in the berry.

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Polyphenols in grapes and wine: stilbenes

Polyphenols in Grapes
Fig. 7 – trans-Resveratrol

They are phytoalexins which are produced in low concentrations only by a few edible species, including grapevine (on the contrary, flavonoids are present in all higher plants).
Together with the other polyphenols in grapes and wine, also stilbenes, particularly resveratrol, have been associated with health benefits resulting from the consumption of wine.
Their content increases from the veraison to the ripening of the berry, and is influenced by the type of cultivar, climate, wine-making technology, and fungal pressure.
The main stilbenes present in grapes and wine are:

  • cis- and trans-resveratrol (3,5,4′-trihydroxystilbene);
  • piceid or resveratrol-3-glucopyranoside and astringin or 3′-hydroxy-trans-piceid;
  • piceatannol;
  • dimers and oligomers of resveratrol, called viniferins, of which the most important are:

α-viniferin, a trimer;
β-viniferin, a cyclic tetramer;
γ-viniferin, a highly polymerized oligomer;
ε-viniferin, a cyclic dimer.

In grapes, other glycosylated and isomeric forms of resveratrol and piceatannol, such as resveratroloside, hopeaphenol, or resveratrol di- and tri-glucoside derivatives, have been found in trace amounts.
Glycosylation of stilbenes is important for the modulation of antifungal activity, protection from oxidative degradation, and storage of the wine.
The synthesis of dimers and oligomers of resveratrol, both in grapes and wine, represents a defense mechanism against exogenous attacks or, on the contrary, the result of the action of extracellular enzymes released from pathogens in an attempt to eliminate undesirable compounds.

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Polyphenols in grapes and wine: hydroxybenzoates

Polyphenols in Grapes
Fig. 8 – Gallic Acid

The hydroxybenzoic acid derivatives are a minor component in grapes and wine.
In grapes, gentisic, gallic, p-hydroxybenzoic and protocatechuic acids are the main ones.
Unlike hydroxycinnamates, which are present in wine as esters with tartaric acid, they are found in their free form.
Together with flavonols, proanthocyanidins, catechins, and hydroxycinnamates they are among the responsible of astringency of wine.

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References

Andersen Ø.M., Markham K.R. Flavonoids: chemistry, biochemistry, and applications. CRC Press Taylor & Francis Group, 2006

Basli A, Soulet S., Chaher N., Mérillon J.M., Chibane M., Monti J.P.,1 and Richard T. Wine polyphenols: potential agents in neuroprotection. Oxid Med Cell Longev 2012. doi:10.1155/2012/805762

de la Rosa L.A., Alvarez-Parrilla E., Gonzàlez-Aguilar G.A. Fruit and vegetable phytochemicals: chemistry, nutritional value, and stability. 1th Edition. Wiley J. & Sons, Inc., Publication, 2010

Flamini R., Mattivi F.,  De Rosso M., Arapitsas P. and Bavaresco L. Advanced knowledge of three important classes of grape phenolics: anthocyanins, stilbenes and flavonols. Int J Mol Sci 2013;14:19651-19669. doi:10.3390/ijms141019651

Georgiev V., Ananga A. and Tsolova V. Recent advances and uses of grape flavonoids as nutraceuticals. Nutrients 2014;6: 391-415. doi:10.3390/nu6010391

Guilford J.M. and Pezzuto J.M. Wine and health: a review. Am J Enol Vitic 2011;62(4):471-486. doi:10.5344/ajev.2011.11013

He S., Sun C. and Pan Y. Red wine polyphenols for cancer prevention. Int J Mol Sci 2008;9:842-853. doi:10.3390/ijms9050842

Xia E-Q., Deng G-F., Guo Y-J. and Li H-B. Biological activities of polyphenols from grapes. Int J Mol Sci 2010;11-622-646. doi:10.3390/ijms11020622

Waterhouse A.L. Wine phenolics. Ann N Y Acad Sci 2002;957:21-36. doi:10.1111/j.1749-6632.2002.tb02903.x