Fischer-Rosanoff convention

Martin André Rosanoff the Fischer-Rosanoff ConventionIn 1906, the Russian-American chemist Martin André Rosanoff, working at that time at New York University, chose glyceraldehyde, a monosaccharide, as the standards for denoting the stereochemistry of carbohydrates and other chiral molecules, a nomenclature system called the Fischer-Rosanoff convention, or Rosanoff convention, or D-L system.
Because Rosanoff didn’t know the absolute configuration of glyceraldehyde, he assigned in a completely arbitrary manner:

  • the D prefix, from the Latin dexter, meaning “right”, to (+)-glyceraldehyde, the dextrorotatory enantiomer, thus assuming that the configuration, in Fischer projections, was that with the hydroxyl group (–OH)  attached to the chiral center on the right side of the molecule;
  • the L prefix, from the Latin laevus, meaning “left”, to (-)-glyceraldehyde, the levorotatory enantiomer, thus assuming that the configuration, in Fischer projections, was that with the hydroxyl group attached to the chiral center on the left  side of the molecule.
    Fischer-Rosanoff convention: D-glyceraldehyde and L-glyceraldehyde as standard for the stereochemistry of chiral molecules

Although Fischer rejected this nomenclature system, it was universally accepted and used to obtain the relative configurations of the chiral molecules. How? The configuration about a chiral center is related to that of glyceraldehyde by converting its groups to those of the monosaccharide through reactions that occur with retention of configuration, namely, reactions that do not break any of the bonds to the chiral center. This means that the spatial arrangement of the groups around the chiral center in the reagents and products is same. The Fischer convention allows to divide the chiral molecules, such as amino acids and monosaccharides, into two classes, known as the D series and the L series, depending on whether the configuration of the groups around the chiral center is related to that of D-glyceraldehyde or L-glyceraldehyde.

Note: there is no correlation between retention of configuration and sign of the rotatory power: the D-L system does not specify the sign of the rotation of plane-polarized light caused by the chiral molecule, but simply correlates the configuration of the molecule with that of the glyceraldehyde.

CONTENTS

Fischer-Rosanoff convention and carbohydrates

Monosaccharides can be aldoses or ketoses. Aldoses, and ketoses with more than three carbon atoms have at least one chiral center, and, by convention, they belong to the D series or to the L series if the configuration of the chiral carbon farthest from the carbonyl carbon, the carbon with the highest oxidation state, is same as that of D-glyceraldehyde or L-glyceraldehyde, respectively.
In Fischer projections the longest chain of carbon atoms is oriented vertically, and the atoms are numbered so that the carbonyl carbon has the lowest possible number, then, C-1 in aldoses and C-2 in ketoses.
Aldoses, ketoses, carbonyl carbon, and asymmetric center taken as the reference centerNote: in Nature, D-sugars are much more abundant than L-sugars.

If the sign of the rotation of plane-polarized light must be specified in the name, the prefixes (+) or (-) can be employed in addition to the D and L prefixes. For example, fructose, which is levorotatory, can be named D-(-)-fructose, whereas glucose, which is dextrorotatory, can be named D-(+)-glucose.

Fischer-Rosanoff convention and α-amino acids

Amino acids, depending on the position of the amino group (–NH2) with respect to the carboxyl group (–COOH) can be classified as:

  • α-amino acids, in which the amino group is attached to the α-carbon;
  • β-amino acids, in which the amino group is attached to the β-carbon;
  • γ-amino acids, in which the amino group is attached to the γ-carbon;
  • δ -amino acids, in which the amino group is attached to the δ-carbon.

α-Amino acids belong to the D series or to the L series if the configuration of the –NH2, –COOH, –R, and H groups attached to the α-carbon, the chiral center, is the same of the hydroxyl, aldehyde (–CHO), and hydroxymethyl (–CH2OH), and H groups of D-glyceraldehyde or L-glyceraldehyde, respectively.
In Fischer projections the molecules are arranged so that the carboxylic group, namely, the carbon with the highest oxidation state, is at the top, and the R group at the bottom.
Fischer-Rosanoff convention and alpha-amino acids of the D-series and L-series Among α-amino acids, proteinogenic amino acids, with the exception of glycine whose α-carbon is not chiral, have the L configuration, hence, they are L-α-amino acids.

Note: in Nature, L-α-amino acids are much more abundant than all the other amino acids, which do not participate in protein synthesis.

Relative and absolute configurations

When Rosanoff arbitrarily assigned the D prefix to (+)-glyceraldehyde and the L prefix to (-)-glyceraldehyde, he had 50/50 chance of being correct.
In the early 1950s, a new technique, the x-ray diffraction analysis, made possible to establish the absolute configuration of chiral molecules.Johannes Martin Bijvoet and the absolute configurations of chiral molecules In 1951 a Dutch chemist, Johannes Martin Bijvoet established the absolute configuration of sodium rubidium (+)‐tartrate tetrahydrate and, comparing it with glyceraldehyde, demonstrated that Rosanoff’s guess was right. Consequently, the configurations of the chiral compounds obtained by relating them to that of glyceraldehyde were the same as their absolute configurations: hence, the relative configurations became absolute configurations.

Ambiguities of the Fischer-Rosanoff convention

The Fischer-Rosanoff convention gives rise to uncertainties with molecules with more than one chiral center. For example, considering D-(+)-glucose, the D-L system gives information about the configuration of C-2, but no information about the other asymmetric centers, namely, C-3, C-4, and C-5.
Asymmetric centers of D-(+)-GlucoseIn these cases, the RS system, developed in 1956 by Robert Sidney Cahn, Christopher Ingold, and Vladimir Prelog, labeling each chiral center, allows to describe accurately the stereochemistry of the molecule. In the case of D-(+)-glucose, the molecule has the (2R,3S,4R,5R)-configuration.
It should also be noted that depending on the chiral center taken as the reference center, the same molecule can belong to both the D and L series.

References

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

IUPAC. Compendium of Chemical Terminology, 2nd ed. (the “Gold Book”). Compiled by A. D. McNaught and A. Wilkinson. Blackwell Scientific Publications, Oxford (1997). Online version (2019-) created by S. J. Chalk. ISBN 0-9678550-9-8. doi:10.1351/goldbook

Mason S.F. Molecular optical activity and the chiral discriminations. Cambridge University Press, 2009 [Google eBooks]

Moran L.A., Horton H.R., Scrimgeour K.G., Perry M.D. Principles of Biochemistry. 5th Edition. Pearson, 2012

A. Rosanoff. On Fischer’s classification of stereo-isomers. J Am Chem Soc 1906:28(1);114-121. doi:10.1021/ja01967a014

Fischer projections

Hermann Emil FischerIn 1891, Hermann Emil Fischer, a German chemist, Nobel Laureate in chemistry in 1902, developed a systematic method for the two-dimensional representation of chiral molecules, the so-called Fischer projections or Fischer projection formulas.
Despite they are two-dimensional structures, Fischer projections  preserve information about the stereochemistry of the molecules and, although not being a representation of how molecules might look in solution, are still widely used by biochemists to define the stereochemistry of amino acids, carbohydrates, nucleic acids, terpenes, steroids, and other molecules of biological interest.

CONTENTS

How to draw Fischer projections

By considering a molecule with a single chiral center, e.g. a carbon atom, for drawing the Fischer projections, the tetrahedral structure is rotated so that two groups point downward, whereas two groups point upwards. Then, you draw a cross, place the chiral center at the center of the cross, and arrange the molecule so that the groups pointing downward, that is, behind the plane of the paper, are attached to the ends of the vertical line, and the groups pointing upwards, that is, out front from the plane of the paper, are attached to the ends of the horizontal line.
How to draw Fischer projections of molecules with one chiral center
For compounds  with more than one chiral center, the same procedure is applied to each asimmetric center.
It is also possible to convert a Fischer projection into a three-dimensional representation, for example using the wedges and dashes of perspective formulas, where the two horizontal bonds are represented by solid wedges, whereas the vertical bonds are represented by dashed lines.

How to manipulate Fischer projection formulas?

  • Since Fischer projections represent three-dimensional molecules on a two-dimensional sheet of paper, some rules must be respected to avoid changing the configuration.
    • The projections must not be lifted out the plane of the paper, because this causes enantiomer is converted into the other enantiomer.
    • If you rotate the projections in the plane of the paper, you obtain the same enantiomer if you rotate the structures by 180° in either direction, because the vertical groups must lie below  the plane of the paper, whereas the horizontal groups above. Conversely, the rotation by 90° or 270° in either direction causes an enantiomer is converted into the other enantiomer.

    Rules for manipulating Fischer projections

  • An odd number of exchanges of two groups leads to the other enantiomer.

References

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

IUPAC. Compendium of Chemical Terminology, 2nd ed. (the “Gold Book”). Compiled by A. D. McNaught and A. Wilkinson. Blackwell Scientific Publications, Oxford (1997). Online version (2019-) created by S. J. Chalk. ISBN 0-9678550-9-8. doi:10.1351/goldbook

Moran L.A., Horton H.R., Scrimgeour K.G., Perry M.D. Principles of Biochemistry. 5th Edition. Pearson, 2012

Voet D. and Voet J.D. Biochemistry. 4th Edition. John Wiley J. & Sons, Inc. 2011

Energy yield of glycogen under aerobic and anaerobic conditions

Glycogen, together with lipids, is a reserve of energy for use when needed. Although quantitatively lower than lipids, glycogen has some metabolic advantages.

  • It is mobilized faster than lipids.
  • It is a storage of glucose used to maintain the blood glucose level during fasting, physical activity, and between meals.
  • It is an energy source used both in aerobic and anaerobic conditions.

By considering the last point, the energy yield of glucose released from glycogen is different under aerobic and anaerobic conditions. The metabolic bases of these differences are analyzed below.

CONTENTS

Energy yield of glycogen stores under anaerobic conditions

Under anaerobic conditions, the oxidation of glucose to lactate via anaerobic glycolysis yields two molecules of ATP.
Below, the yield of ATP from anaerobic oxidation of glucose released during glycogenolysis by the action of glycogen phosphorylase (EC 2.4.1.1), and debranching enzyme (EC 3.2.1.33) is considered.
Note: glycogen phosphorylase releases about 90% of stored glucose in the form of glucose-1-phoshate (G-1-P), and debranching enzyme the remaining 10% in the form of glucose.


Glycogen phosphorylase and the oxidation of G-1-P under anaerobic conditions

Glycogen synthesis from glucose requires 2 ATP for each molecule of glucose.
The release of glucose-1-phosphate by the action of glycogen phosphorylase allows the recovery of one of the 2 molecules of ATP used in the preparatory phase of glycolysis. The anaerobic oxidation of glucose-6-phosphate, produced from glucose-1-phosphate (G-1-P) by the action of phosphoglucomutase (EC 5.4.2.2), yields therefore three molecules of ATP and not two, because:

  • one molecule of ATP, instead of two, is used in the preparatory phase of glycolysis, because hexokinase reaction (EC 2.7.1.1) is bypassed;
  • four molecules of ATP are produced in the payoff phase of glycolysis.

The cost-gain rate is 1/3, namely, there is an energy yield of about 66,7%.
The overall reaction is:

Glycogen(n glucose residues) + 3 ADP + 3 Pi → Glycogen(n-1 glucose residues) + 2 Lactate + 3 ATP

By considering the two molecules of ATP used in the synthesis of glycogen and the anaerobic oxidation of glucose-1-phosphate to lactate, there is a yield of one molecule of ATP for each molecule of glucose stored. The overall reaction is:

Glucose + ADP + Pi → 2 Lactate + ATP

Debranching enzyme and the oxidation of glucose under anaerobic conditions

By considering the glucose released by the action of debranching enzyme, the yield of ATP is zero because:

  • two molecules of ATP are used in the synthesis of glycogen from glucose;
  • debranching enzyme releases glucose, then, two molecules of ATP will be used in the preparatory phase of glycolysis;
  • four molecules of ATP are produced in the payoff phase of glycolysis.

If we now consider the oxidation to lactate of all glucose released from glycogen, there is an energy yield equal to:

1-{[(1/3)*0,9]+[(2/2)*0,1]}=0,60

Then, under anaerobic conditions, there is an energy yield of 60%, hence, glycogen is a good storage form of energy.

Energy yield of glycogen stores under aerobic conditions

Under aerobic conditions, the oxidation of glucose to CO2 and H2O via glycolysis, pyruvate dehydrogenase complex, Krebs cycle, mitochondrial electron transport chain, and oxidative phosphorylation yields about 30 molecules of ATP.
Below, the yield of ATP from aerobic oxidation of glucose released from glycogen by the action of glycogen phosphorylase and debranching enzyme is considered.

Glycogen phosphorylase and the oxidation of G-1-P under aerobic condition

The oxidation of glucose-6-phosphate, produced from glucose-1-phosphate by the action of phosphoglucomutase, to CO2 and H2O yields 31 molecules of ATP, and not 30, because only one molecule of ATP is used in the preparatory phase of glycolysis. The cost-gain rate is 1/31, namely, there is an energy yield of about 97%.
The overall reaction is:

Glycogen(n glucose residues) + 31 ADP + 31 Pi → Glycogen(n-1 glucose residues) + 31 ATP + 6 CO2 + 6 H2O

By considering the two molecules of ATP used in the synthesis of glycogen and the aerobic oxidation of glucose-1-phosphate to CO2 and H2O, there is a yield of 29 molecules of ATP for each molecule of glucose stored.
The overall reaction is:

Glucose + 29 ADP + 30 Pi → 29 ATP + 6 CO2 + 6 H2O

Debranching enzyme and the oxidation of glucose under aerobic condition

By considering the glucose released by the action of debranching enzyme, there is a yield of 30 molecules of ATP, because two molecules of ATP are used in the preparatory phase of glycolysis. The cost-gain rate is 2/30, namely, there is an energy yield of about 93,3%.

If we now consider the oxidation to CO2 and H2O of all glucose released from glycogen, there is an energy yield equal to:

1-{[(1/31)*0,9]+[(2/30)*0,1]}=0,96

Energy yield of glycogen aerobic anaerobic conditionsThen, under aerobic conditions, there is an energy yield of 96%, hence, glycogen is a extremely efficient storage form of energy, with a gain of 36% compared to anaerobic conditions.

References

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

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

Voet D. and Voet J.D. Biochemistry. 4th Edition. John Wiley J. & Sons, Inc. 2011

RS system: the priority rules

In 1956, Robert Sidney Cahn, Christopher Ingold and Vladimir Prelog developed a nomenclature system that, based on a few simple rules, allows to assign the absolute configuration of each chirality center in a molecule.
This nomenclature system, called the RS system, or the Cahn-Ingold-Prelog (CIP) system, when added to the IUPAC system of nomenclature, allow to name accurately and unambiguously the chiral molecules, even when there is more than one asymmetric center.
Chiral molecules are, in most cases, able to rotate plane-polarized light when light passes through a solution containing them. In this regard, it should be emphasized that the sign of the rotation of plane-polarized light caused by a chiral compound provides no information concerning RS configuration of its chiral centers.
The Fischer-Rosanoff convention is another way to describe the configuration of chiral molecules. However, compared to RS system, it labels the whole molecule and not each chirality center, and is often ambiguous for molecules with two or more chirality centers.

CONTENTS

The priority rules of the RS system

The RS system assigns a priority sequence to the groups attached to the chirality center and, tracing a curved arrow from the highest priority group to the lowest, labels each chiral center R or S.

First rule

A priority sequence is assigned to the groups based on the atomic number of the atoms directly attached to the chiral center.

  • The atom with the highest atomic number is assigned the highest priority.
  • The atom with the lowest atomic number is assigned the lowest priority.

For example, if an oxygen atom, O, atomic number 8, a carbon atom, C, atomic number 6, a chlorine atom, Cl, atomic number 17, and a bromine atom, Br, atomic number 35 are attached to the chiral center, the order of priority is: Br > Cl > O > C.
For isotopes, the atom with the highest atomic mass is assigned the highest priority.

Second rule

When different groups are attached to the chiral center through identical atoms, the priority sequence is assigned based on the atomic number of the next atoms bound, then moving outward from the chirality center until the first point of difference is reached.
If, for example, –CH3, –CH2CH3 and –CH2OH groups are attached to a chiral center, there are three identical atoms directly attached to the chiral center. Analyzing the next atoms bound, we have:

  • for the methyl group –CH3
H, H, H
  • for the ethyl group –CH2CH3
H, H, C
  • for the hydroxymethyl group –CH2OH
H, H, O

RS system priority sequence first rule

Because the atomic number of oxygen is higher than that of carbon, that, in turn, is higher than that of hydrogen, the order of priority is –CH2OH > –CH2CH3 > –CH3
The order of priority of some groups is:

–I > –Br > –Cl > –SH > –OR > –OH > –NHR > –NH2 > –COOR > –COOH > –CHO > –CH2OH > –C6H5 > –CH3 > –2H > –1H

Note that the groups attached to a chirality center must not have identical priority ranking, because, in that case, the center cannot be chiral.

Once the priority sequence has been established, the molecule is oriented in space so that the group with the lowest priority is pointed away from the viewer, then behind the chiral center. Now, trace a curved arrow, a circle, from the highest priority group to the lowest.

  • If you move in a clockwise direction, the configuration of chiral center is R, from the Latin rectus, meaning “right”.
  • If you move in a counterclockwise direction, the configuration of chiral center is S, from the Latin sinister, meaning “left”.

R configuration chiral center

Third rule

This is the third rule of the RS system, by which we can determine the configuration of a chirality center when there are double or triple bonds in the groups attached to the chirality center.
To assign priorities, the atoms engaged in double or triple bonds are considered duplicated and tripled, respectively.
RS system priority sequence third ruleIn the case of a C=Y double bond, one Y atom is attached to the carbon atom, and one carbon atom is attached to the Y atom.
In the case of a C≡Y triple bond, two Y atoms are attached to the carbon atom, and two carbon atoms are attached to the Y atom.

RS system and multiple chiral centers

When two or more chirality centers are present in a molecule, each center is analyzed separately using the rules previously described.
Consider 2,3-butanediol. The molecule has two chiral centers, carbon 2 and carbon 3, and exists as three stereoisomers: two enantiomers and a meso compound. What is the RS configuration of the chiral centers of the enantiomer shown in figure?RS configuration chiral centers (2R,3R)-2,3-ButanediolConsider carbon 2. The order of priority of the groups is –OH > –CH2OHCH3 > –CH3 > –H. Rotate the molecule so that the hydrogen, the lowest priority group, is pointed away from the viewer. Tracing a path from –OH, the highest priority group, to –CH3, the lowest priority group, we move in a clockwise direction: the configuration of the carbon 2 is, therefore, R. Applying the same procedure to carbon 3, its configuration is R. Then, the enantiomer shown in figure is (2R,3R)-2,3-butanediol.

Amino acids and gliceraldeide

In the Fischer-Rosanoff convention, all the proteinogenic amino acids are L-amino acids. In the RS system, with the exception of glycine, that is not chiral, and cysteine ​​that, due to the presence of the thiol group, is (R)-cysteine, all the other proteinogenic amino acids are (S)-amino acids.
Threonine and isoleucine have two chirality centers, the α-carbon and a carbon atom on the side chain, and exist as three stereoisomers: two enantiomers and a meso compound. The forms of the two amino acids isolated from proteins are (2S,3R)-threonine and (2S,3S)-isoleucine, in Fischer-Rosanoff convention, L-threonine and L-isoleucine.
In the RS system, L-glyceraldehyde is (S)-glyceraldehyde, and, obviously, D-glyceraldehyde is (R)-glyceraldehyde.

References

Cahn R.S., Ingold C., Prelog V. Specification of molecular chirality. Angew Chem 1966:5(4); 385-415. doi:10.1002/anie.196603851

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

IUPAC. Compendium of Chemical Terminology, 2nd ed. (the “Gold Book”). Compiled by A. D. McNaught and A. Wilkinson. Blackwell Scientific Publications, Oxford (1997). Online version (2019-) created by S. J. Chalk. ISBN 0-9678550-9-8. doi:10.1351/goldbook

Moran L.A., Horton H.R., Scrimgeour K.G., Perry M.D. Principles of Biochemistry. 5th Edition. Pearson, 2012

Prelog V. and Helmchen G. Basic principles of the CIP‐system and proposals for a revision. Angew Chem 1982:21(8);567-83. doi:10.1002/anie.198205671

Rosanoff M.A. On Fischer’s classification of stereo-isomers. J Am Chem Soc 1906:28(1);114-121. doi:10.1021/ja01967a014

Voet D. and Voet J.D. Biochemistry. 4th Edition. John Wiley J. & Sons, Inc. 2011

Chirality

The definition of chirality, from the Greek cheir meaning “hand”, is due to Lord Kelvin who enunciated it during the “Baltimore Lectures”, a series of lectures held at Johns Hopkins University in Baltimore, starting from October 1st 1884, and published twenty years later, in 1904, in which the English scientist, among other things, stated: “I call any geometrical figure, or groups of points, chiral, and say it has chirality, if its image in a plane mirror, ideally realized, cannot be brought to coincide with itself“.
Lord Kelvin and The Definition of ChiralityChirality is, therefore, the geometric property of a group of points or atoms in space, or of a solid object, of not being superimposable on its mirror image. These structures, defined as chiral, have the peculiar property of being devoid of symmetry elements of the second kind, namely, a mirror plane, an center of inversion, or a rotation-reflection.
The environment is rich in chiral objects: your hands are the example par excellence, but there are many others, from the shell of a snail to a spiral galaxy. In chemistry, and especially in organic chemistry, chirality is a property of primary importance, because molecules such as carbohydrates, many amino acids, as well as many drugs, are chiral.
Chiral molecules can exist in two forms, mirror images of each other and non-superimposable, namely, there is no combination of rotations or translations on the plane of the sheet that allows their superposition. Such molecules are called enantiomers, from the Greek enántios, meaning “opposite” and meros,meaning “part”.
The most common cause of chirality in a molecule is the presence of a chirality center or chiral center, also called asymmetric center, namely, an atom that bears a set of atoms or functional groups in a spatial arrangement so that the resulting molecule can exist as two enantiomers.
Enantiomers are a kind of stereoisomers, that, in turn, can be defined as isomers having the same number and kind of atoms and bonds, but differing in the spatial orientation of the atoms.

CONTENTS

Enantiomers

Two enantiomers of a chiral molecule, being non-superimposable, are different compounds. How do they differ?
Each pair of enantiomers has identical physical and chemical properties towards achiral properties, such as melting point, boiling point, refractive index, infrared spectrum, the solubility in the same solvent, or the same reaction rate with achiral reagents.
The differences emerge when they interacts with chemical and physical phenomena that have chiral properties.

  • From the chemical point of view, two enantiomers can be distinguished by the way they interact with chiral structures, such as the binding site of a chiral receptor or the active site of a chiral enzyme.
  • From the physical point of view, they differ in their interaction with plane-polarized light, that has chiral properties, namely, they have optical activity.

Chirality and optical activity

The optical activity of materials such as quartz and, more importantly, of organic compounds such as sugars or tartaric acid, was discovered in 1815 by the French scientist Jean-Baptiste Biot.
Chiral molecules can be classified based on the direction in which plane-polarized light is rotated when it passes through a solution containing them.

  • If a solution containing one enantiomer rotates plane-polarized light in a clockwise direction from the observer’s point of view, the molecule is called dextrorotatory or dextrorotary, from the Latin dexter, meaning “right”, and is designated by the prefixes (+)-, or d from dextro-.
  • If a solution containing one enantiomer rotates plane-polarized light in a counterclockwise direction from the observer’s point of view, the molecule is called levorotatory or levorotary, from the Latin laevus, meaning “left”, and is designated by the prefixes (-), or l from laevo-.

Obviously, if we consider a pair of enantiomers, one is dextrorotatory and the other levorotatory.
At present it is not possible to reliably predict the magnitude, direction, or sign of the rotation of plane-polarized light caused by an enantiomer. On the other hand, the optical activity of a molecule provides no information on the spatial arrangement of the chemical groups attached to the chirality center.
Note: a system containing molecules that having the same chirality sense is called enantiomerically pure or enantiopure.

Pasteur and the discovery of enantiomers

Louis Pasteur and the discovery of enantiomersIn 1848, thirty three years after Biot’s work, studies on the optical activity of molecules led Louis Pasteur, who had been a student of Biot, to note that, following the recrystallization of a concentrated aqueous solution of sodium ammonium tartrate, optically inactive, two kinds of crystals precipitated, that were non-superimposable mirror images of each other. After separating them with tweezers, Pasteur discovered that the solutions obtained by dissolving equimolar amounts of the two kind of crystals were optically active and, interestingly, the rotation angle of plane-polarized light was equal in magnitude but opposite in sign. Because the differences in optical activity were due to the dissolved sodium ammonium tartrate crystals, Pasteur hypothesized that the molecules themselves should be non-superimposable mirror images of each other, like their crystals: they were what we now call enantiomers. And it is Pasteur who first used the term asymmetry to describe this property, then called chirality by Lord Kelvin.

Racemic mixtures

A solution containing an equal amount of each member of a pair of enantiomers is called racemic mixture or racemate. These solutions are optically inactive: there is no net rotation of plane-polarized light since the amount of dextrorotatory and levorotatory molecules is exactly the same.
Unlike what happens in biochemical processes, the chemical synthesis of chiral molecules that does not involve chiral reactants, or that is not followed by methods of separation of enantiomers, inevitably leads to the production of a racemic mixture.
The pharmaceutical chemistry is among the sectors most affected by this. As previously mentioned, two enantiomers are different compounds. Many chiral drugs are synthesized as racemic mixtures, but most often the desired pharmacological activity resides in one enantiomer, called eutomer; the other, called distomer, is less active or inactive. An example is ibuprofen, an arylpropionic acid derivative, and anti-inflammatory drug: only the S enantiomer has the pharmacological activity.

Enantiomers of IbuprofenArylpropionic derivatives are sold as racemic mixtures: a racemase converts the distomer to the eutomer in the liver.
However, it is also possible that the distomer causes harmful effects and must be eliminated from the racemic mixture. A tragic example is thalidomide, a sedative and anti-nausea drug sold as a racemic mixture from the 1950s until 1961, and taken also during pregnancy.

Enantiomers of ThalidomideThe distomer, the S enantiomer, could cause serious birth defects, particularly phocomelia. This is probably the most striking example of the importance of the chiral properties of molecules, which prompted health care organizations to promote the synthesis of drugs, including thalidomide, containing a single enantiomer by the pharmaceutical industry.

Chirality centers

Any tetrahedral atom that bears four different substituents can be a chirality center.
Carbon atom is the classic example, but also other atoms from group IVA of the periodic table, such as the semimetals silicon (Si) and germanium (Ge), have a tetrahedral arrangement and can be chiral centers. Another example is the phosphorus atom in organic phosphate esters that has a tetrahedral arrangement, then, when it binds four different substituents it is a chiral center.
The nitrogen atom of a tertiary amine, an amine in which the nitrogen is bounded to three different groups, is a chiral center. In these compounds, nitrogen is located at the center of a tetrahedron and its four sp3 hybrid orbitals point to the vertices, three of which are occupied by the three substituents, whereas the nonbonding electron pair points towards the fourth.
Nitrogen inversion and chiralityAt room temperature, nitrogen rapidly inverts its configuration. The phenomenon is known as nitrogen inversion, namely, a rapid oscillation of the atom and its ligands, during which nitrogen passes through a planar sp2-hybridized transition state. As a consequence, if the nitrogen atom is the only chiral center of the molecule, there is no optical activity because a racemic mixture exists. The inversion of configuration does not occur only in some cases in which nitrogen is part of a cyclic structure that prevents it. Therefore, the presence of a chiral center could be not sufficient to allow the separation of the respective enantiomers.

Note: in 1874, Jacobus Henricus van ‘t Hoff and Joseph Achille Le Bel, based on the work of Pasteur, first formulated the theory of the tetrahedral carbon atom. For this work van ‘t Hoff received the first Nobel Prize in chemistry in 1901.

Chirality in the absence of a chiral center

Chirality can also occur in the absence of a chiral center, due to the lack of free rotation around a double or a single bond, as in the case of:

  • allene derivatives, organic compounds in which there are two cumulative double bonds, namely, two double bonds localized on the same carbon atom;
  • biphenyl derivatives.

Chirality is due to the presence of an axis of chiralityIn this case, chirality is due to the presence of an axis of chirality.

Meso compounds

Meso compounds or meso isomers are stereoisomers with two or more chiral centers that are superimposable on their mirror image, then achiral and, as such, optically inactive. Moreover, they have an internal mirror plane that bisects the molecule, with each half a mirror image of the other. Then, meso compounds can be classified as diastereomers, namely, stereoisomers which are not enantiomers.
For a molecule with n chirality centers, the maximum number of possible stereoisomers is 2n.
Consider 2,3-butanediol. The molecule has two chirality centers, the carbons 2 and 3, so there are 22 = 4 possible stereoisomers, whose structures are depicted in the figure, in the Fischer projections, indicated as A, B, C, D.
Stereoisomers, chirality centers, and meso compoundsStructures A and B are mirror images of each other and non-superimposable, then they are a pair of enantiomers.
Structures C and D are mirror images of each other, but are superimposable. In fact, if we rotate structure C or D of 180 degrees, the two structures are superimposable. Then, they are not a pair of enantiomers: they are the same molecule with opposite orientation. Moreover, they have an internal mirror plane, that bisects the molecule, giving two halves, each a mirror image of the other. Structure C, or D, is therefore a meso compound because it has chiral centers, is superimposable on its mirror image, and has internal mirror plane that divides the molecule into two mirror‐image halves.

References

Capozziello S. and Lattanzi A. Geometrical approach to central molecular chirality: a chirality selection rule. Chirality 2003;15:227-230. doi:10.1002/chir.10191

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

IUPAC. Compendium of Chemical Terminology, 2nd ed. (the “Gold Book”). Compiled by A. D. McNaught and A. Wilkinson. Blackwell Scientific Publications, Oxford (1997). Online version (2019-) created by S. J. Chalk. ISBN 0-9678550-9-8. doi:10.1351/goldbook

Kelvin WT. Baltimore lectures on molecular dynamics and the wave theory of light. Clay C. J., London: 1904:619. doi:https://archive.org/details/baltimorelecture00kelviala/mode/2

Voet D. and Voet J.D. Biochemistry. 4th Edition. John Wiley J. & Sons, Inc. 2011

Isomerism

The phenomenon that two or more different chemical compounds have the same molecular formula is called isomerism, from the Greek isos meaning “equal”, and meros meaning “part”, a concept and term introduced by the Swedish scientist Jacob Berzelius in 1830.
Isomerism is a consequence of the fact that the atoms of a molecular formula can be arranged in different ways to give compounds, called isomers, that differ in physical and chemical properties.
There are two types of isomerism: structural isomerism and stereoisomerism, which can be divided into further subtypes.

Tree Diagram for Types of Isomerism

CONTENTS

Structural isomerism

In structural isomerism, also called constitutional isomerism, isomers differ from each other in that the constituent atoms are linked in different ways and sequences.
There are several subtypes of structural isomerism: positional, functional group and chain isomerism.

Positional isomers

In positional isomerism, also called position isomerism, isomers have the same functional groups but in different positions on the same carbon chain.
An example is the compound with molecular formula C6H4Br2, of which there are three isomers: 1,2-dibromobenzene, 1,3-dibromobenzene and 1,4-dibromobenzene. These isomers differ in the position of the bromine atoms on the cyclic structure.

Positional Isomerism

Another example is the compound with molecular formula C3H8O, of which there are two isomers: 1-propanol or n-propyl alcohol, and 2-propanol or isopropyl alcohol. These isomers differ in the position of the hydroxyl group on the carbon chain.

Position Isomers

Functional group isomers

Functional group isomerism, also called functional isomerism, occurs when the atoms form different functional groups.
An example the compound with molecular formula C2H6O, of which there are two isomers: dimethyl ether and ethanol or ethyl alcohol, that have different functional groups, an ether group, –O–, and a hydroxyl group, –OH, respectively.

Functional Group Isomerism

Chain isomers

In chain isomerism, isomers differ in the arrangement of the carbon chains, that may be branched or straight.
An example is the compound with the molecular formula C5H12, of which there are three isomers: n-pentane, 2-methylbutane or isopentane and 2,2-dimethylpropane or neopentane.

Chain Isomers

Stereoisomerism

In stereoisomerism, isomers have the same number and kind of atoms and bonds, but differ in the orientation of the atoms in space. Such isomers are called stereoisomers, from the Greek stereos, meaning “solid”.
There are two subtypes of stereoisomerism, conformational isomerism and configurational isomerism; the latter can be further subdivided into optical isomerism and geometrical isomerism.

Conformational isomerism

In conformational isomerism, the stereoisomers can be interconverted by rotation around one or more single bonds, the σ bonds. These rotations produce different arrangements of atoms in space that are non-superimposable. And the number of possible conformations a molecule can adopted is theoretically unlimited, ranging from the lowest energy structure, the most stable, to the highest energy structure, the less stable. Such isomers are called conformer.
For example, if we consider ethane, C2H4, looking at the molecule from one end down the carbon-carbon bond, using the Newman projection, hydrogen atoms of a methyl group can be, with respect to the hydrogen atoms of the other methyl group, in one of the following conformations.

  • The eclipsed conformation, in which hydrogen atoms of a methyl group are hidden behind those of the other methyl group, then, the angle between carbon-hydrogen bonds on the front and rear carbons, called a dihedral angle, could be 0, 120, 240, 360 degrees. This is the highest energy conformation, then is the less stable.
  • The staggered conformation, in which hydrogen atoms of a methyl group are completely offset from those of the other methyl group, namely, dihedral angles could be 60, 180 or 360 degrees. This is the lowest energy conformation, then the most stable.
  • The skew conformation, corresponding to one of the intermediate conformations between the previous ones.

Ethane Conformers: Newman Projections
The stability of ethane conformers is due to how the electron pairs of the carbon-hydrogen bonds of the two methyl groups are overlapped:

  • in the staggered conformations they are as far away from each other as possible;
  • in the eclipsed conformations they are as close as possible to each other.

The potential energy barrier between these two conformations is small, about 2.8 kcal/mole (11.7 kJ/mole). At room temperature, the kinetic energy of the molecules is 15-20 kcal/mole (62.7-83.6 kJ/mole), more than enough to allow free rotation around the carbon-carbon bond. As a consequence, it is not possible to isolate any particular conformation of ethane.
Note: the potential energy barrier to rotation around double carbon-carbon bonds is about 63 kcal/mole (264 kJ/mole), corresponding to the energy required to break the π bond. (See geometric isomerism). This value is about three times the kinetic energy of the molecules at room temperature at which, then, free rotation is precluded. Only at temperatures above 300 °C molecules acquire enough thermal energy to break the π bond, allowing free rotation around the remaining σ bond. This allows the trans-isomer to be rearranged to the cis-isomer or vice versa.

Configurational isomerism

In configurational isomerism, the interconversion between the stereoisomers does not occur as a result of rotations around single bonds but involves bond breaking and new bond forming, then it doesn’t occur spontaneously at room temperature.
There are two subtypes of configurational isomerism: optical isomerism and geometrical isomerism.

Optical isomers

Optical isomerism occurs in molecules that have one or more chirality centers or chiral centers, namely, tetrahedral atoms that bear four different ligands. The chiral center can be a carbon, phosphorus, sulfur or nitrogen atom.

Chiral Center or Chirality Center
Note: the word chirality derives from the Greek cheiros, meaning “hand”.
Optical isomers lack of a center of symmetry or a plane of symmetry, are mirror image of each other, and cannot be superimposed on one another. Such stereoisomers are called enantiomers, from the Greek enántios, meaning “opposite”, and meros, meaning “part”.
Unlike the other isomers, two enantiomers have identical physical and chemical properties with two exceptions.

  • The direction of rotation of the plane of polarized light, hence the name of optical isomerism.
    If a solution of one enantiomer rotates the plane of polarized light in a clockwise direction, the enantiomer is labeled (+). Conversely, a solution of the other enantiomer rotates the plane of polarized light in a counterclockwise direction by the same angle, and the enantiomer is labeled (-).
  • Although indistinguishable by most techniques, two enantiomers can be distinguished in a chiral environment like the active site of chiral enzymes.

Note: for a molecule with n chiral centers, the maximum number of stereoisomers is equal to 2n.

Geometric isomers

Geometric isomerism, also called cis-trans isomerism, occurs when atoms cannot freely rotate due to a rigid structure such as in:

  • compounds with carbon-carbon, carbon-nitrogen or nitrogen-nitrogen double bonds, where the rigidity is due to the double bond;
  • cyclic compounds, where the rigidity is due to the ring structure.

An example of geometrical isomerism due to the presence of a carbon-carbon double bond is stilbene, C14H12, of which there are two isomers. In one isomer, called cis isomer, the same groups are on the same side of the double bond, whereas in the other, called trans isomer, the same groups are on opposite sides.

Geometric Isomerism

Note: the terms trans and cis are from the Latin trans, meaning “across”, and cis, meaning “on this side of”.
Among the cyclic compounds of carbon, cis-trans isomerism not complicated by the presence of chiral centers occurs in structures with an even number of carbon atoms and substituted in opposite positions, namely, para-substituted. An example is 1,4-dimethylcyclohexane, a cycloalkane, compounds of general formula CnH2n, of which there are two stereoisomers, cis-1,4-dimethylcyclohexane and trans-1,4- dimethylcyclohexane.

cis-trans Isomers

This kind of stereoisomerism cannot exist if one of the atoms that cannot freely rotate carries two groups the same. Why? For the switching between the trans and cis isomers the groups attached to atoms that cannot freely rotate have to be swapped. If there are two groups the same, the switch leads to the formation of the same molecule.
Note: geometric isomers are a special case of diastereomers or diastereoisomers, that, in turn, are stereoisomers that are not mirror image of each other. The other diastereomers are the meso compounds and non-enantiomeric optical isomers.

References

Graham Solomons, T. W., Fryhle C.B.,  Snyder S.A. Solomons’ organic chemistry. 12th Edition. John Wiley & Sons Incorporated, 2017

Morris D.G. Stereochemistry. Royal Society of Chemistry, 2001 [GoogleBook]

North M. Principles and applications of stereochemistry. 1th Edition. CRC Press, 1998

Voet D. and Voet J.D. Biochemistry. 4th Edition. John Wiley J. & Sons, Inc. 2011

Osmotic pressure

In solution, solvent molecules tend to move from a region of higher concentration to one of lower concentration. When two different solutions are separated by a semipermeable membrane, namely, a membrane that allows certain ions or molecules to pass, in this case the solvent molecules, a net flow of solvent molecules from the side with higher concentration to the side with lower concentration will occur. This net flow through the semipermeable membrane produces a pressure called osmotic pressure, indicated as Π, that can be defined as the force that must be applied to prevent the movement of the solvent molecules through a semipermeable membrane.
Osmotic pressure: two different solutions are separated by a semipermeable membrane
Osmotic pressure, together with boiling point elevation, freezing point depression, and vapor pressure lowering, is one of the four colligative properties of solutions, properties that do not dependent of the chemical properties of the solute particles, namely ions, molecules or supramolecular structures, but depend only on the number of solute particles present in solution.
For  a solutions of n solutes, the equation that describes osmotic pressure is the sum of the contributions of each solute:

Π = RT(i1c1 + i2c2 + … + incn)

The equation is known as the van ’t Hoff equation, where:

  • T is the absolute temperature in Kelvin;
  • R is the ideal gas constant = 8.314 J/mol K;
  • c is the molar concentration of the solute;
  • i is the van ’t Hoff factor.

CONTENTS

van ’t Hoff factor

The van ‘t Hoff factor is a measure of the degree of dissociation of solutes in solution, and is described by the equation:

i = 1 + α(n-1)

where:

  • α is the degree of dissociation of the solute molecules, equal to the ratio between the moles of the solute molecules that have dissociated and the number of the original moles, and is comprised between 0, for substances that do not ionize or dissociate in solution, and 1, for substances that completely dissociate or ionize;
  • n is the number of ions formed from the dissociation of the solute molecule.

For non ionizable compounds, such as glucose, glycogen or starch, n = 1, and i = 1.
For compounds that completely dissociate, such as strong acids and strong bases or salts, the van ‘t Hoff factor is a whole number greater than one, as α = 1 and n is equal to at least 2. For example, if we consider sodium chloride, NaCl, potassium chloride, KCl, or calcium chloride, CaCl2, in dilute solution:

NaCl → Na+ + Cl
KCl → K+ + Cl
CaCl2 → Ca2+ + 2 Cl

So in the first two cases i = 2, whereas with calcium chloride, i = 3.
Finally, for substances that do not completely ionize, such as weak acids and weak bases, i is not an integer.

The product of the van ’t Hoff factor and the molar concentration of the solute particles, ic, is the osmolarity of the solution, namely, the concentration of the solute particles osmotically active per liter of solution.

Osmotic pressure, osmosis, and plasma membranes

Osmosis can be defined as the net movement or flow of solvent molecules through a semipermeable membrane driven by osmotic pressure differences across the membrane, to try to equal the concentration of the solute on the two sides of the membrane itself.
In biological systems, water is the solvent and plasma membranes are the semipermeable membranes. Plasma membranes allow water molecules to pass, due to protein channels, known as aquaporins, as well as small non-polar molecules that diffuse rapidly across them, whereas they are impermeable to ions and macromolecules. Inside the cell there are macromolecules, such as nucleic acids, proteins, glycogen, and supramolecular aggregates, for example multienzyme complexes, but also ions in a higher concentration than that of the extracellular environment. This causes osmotic pressure to drive water from outside to inside the cell. If this net flow of water toward the inside of the cell is not counterbalanced, cell swells, and plasma membrane is distended until the cell bursts, that is, an osmotic lysis occurs. Under physiological conditions, this does not happen because during evolution several mechanisms have been developed to oppose, and in some cases even exploit, these osmotic forces. Two of these are energy-dependent ion pumps and, in plants, bacteria and fungi, the cell wall.

Energy dependent ion pumps

Ion pumps reduce, at the expense of ATP, the intracellular concentrations of specific ions with respect to their concentrations in the extracellular environment, thereby creating an unequal distribution of the ions across the plasma membrane, namely, an ion gradient. In this way the cell counterbalances the osmotic forces due to the ions and macromolecules trapped inside it. An example of energy-dependent ion pump is Na+/K+ ATPase, which reduces the concentration of Na+ inside the cell relative to the outside.

Cell wall

Plant cells are surrounded by an extracellular matrix, the cell wall, that, being non expandable and positioned next to the plasma membrane, allows cell to resist osmotic forces that would cause its swelling and finally the lysis. How? Inside mature plant cells, the vacuoles are the largest organelles, occupying about 80% of the total cell volume. Large quantities of solutes, for the most part organic and inorganic acids, are accumulated within them and osmotically draw water, causing their swelling. In turn, this causes the tonoplast, the membrane that surrounds the vacuole, to press the plasma membrane against the cell wall, that mechanically opposes to these forces and avoids the osmotic lysis. This osmotic pressure is called turgor pressure, and can reach up 2 MPa, that is, 20 atmospheres, a value about 10 times higher than the air pressure in tires. It is responsible for the rigidity of non woody parts of plants, is involved in plant growth, as well as in:

  • wilting of vegetables, due to its reduction;
  • plants movements, such as:
    • the circadian movements of the leaves;
    • the movements of the leaves of Dionaea muscipula, the Venus flytrap, or of the leaves of the sensitive plants such as Mimosa pudica.

Even in bacteria and fungi, the plasma membrane is surrounded by a cell wall that stably withholds the internal pressure, then preventing osmotic lysis of the cell.

Isotonic, hypotonic, and hypertonic solutions

By comparing the osmotic pressure of two solutions separated by a semipermeable membrane, it is possible to define three types of solutions, briefly described below.

  • The solutions are isotonic when they have the same osmotic pressure.
  • If the solutions have different osmotic pressures, that with the higher osmotic pressure is defined hypertonic with respect to the other.
  • If the solutions have different osmotic pressures, that with the lower osmotic pressure is defined hypotonic with respect to the other.

In biological systems, the cytosol is the reference solution; then, if we place a cell in a:

  • isotonic solution, no net flow of water occurs into or out of the plasma membrane;
  • hypertonic solution, there is a net flow of water out of the cell, therefore the cell loses water and shrinks;
  • hypotonic solution, there is a net flow of water into the cell, the cell swells and can burst, i.e., an osmotic lysis can occur.

In addition to ion pumps and the cell wall, in the course of evolution multicellular organisms have developed another mechanism to oppose the osmotic forces: to surround the cells with an isotonic solution or close to isotonicity that prevents, or at least limits, a net inflow or outflow of water. An example is plasma, that is, the liquid component of blood, which, due to the presence of salts and proteins, primarily albumin in humans, has an osmolarity similar to that of the cytosol.

Osmotic pressure, starch and glycogen

Living organisms store glucose in the form of polymers, glycogen in animals, fungi, bacteria, and starch in photoautotrophs, but not in the free form. In this way they avoid that the osmotic pressure exerted by the carbohydrate stores becomes too high. Indeed, since osmotic pressure, like the other colligative properties, depends only on the number of solute molecules, storing millions of glucose units in the form of a significantly lower number of polysaccharides allows to avoid an excessive pressure. Here are some examples.

  • A gram of polysaccharide, e.g. glycogen or starch, composed of 1000 glucose units has an effect on osmotic pressure lower than that of a milligram of free glucose.
  • In hepatocyte, if the glucose stored in the form of glycogen was present in the free form, its concentration would be about 0.4 M, whereas the polysaccharide concentration of about 0.04 μM, and this would cause a net flow of water inside the cell such as to lead to osmotic lysis.
    Furthermore, even if osmotic lysis could be avoided, there would be problems with the transport of glucose into the cell. In humans, under physiological conditions, the blood glucose concentration ranges from 3.33 to 5.56 mmol/L (60-100 mg/dL); if glucose was present in the free form, its intracellular concentration would be 120 to 72 times greater than that of the blood, and its transport into the hepatocyte would entail a large energy expenditure.

References

Beauzamy L., Nakayama N., and Boudaoud A. Flowers under pressure: ins and outs of turgor regulation in development. Ann Bot 2014;114(7):1517-33. doi:10.1093/aob/mcu187

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

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

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

Michal G., Schomburg D. Biochemical pathways. An atlas of biochemistry and molecular biology. 2nd Edition. John Wiley J. & Sons, Inc. 2012

Moran L.A., Horton H.R., Scrimgeour K.G., Perry M.D. Principles of Biochemistry. 5th Edition. Pearson, 2012

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

Multifunctional enzymes

Multifunctional enzymes are proteins in which two or more enzymatic activities, that catalyze consecutive steps of a metabolic pathway, are located on the same polypeptide chain. It seems likely they have arisen by gene fusion events, and represent, like the multienzyme complexes, a product of evolution to maximize the catalytic efficiency, providing advantages that you wouldn’t have if such enzymatic activities were present on distinct proteins dissolved in the cytosol.

CONTENTS

What advantages do multifunctional enzymes provide?

Living organisms fight against the natural processes of decay that, if not counteracted, leads to an increasing disorder, until death. At the molecular level, the maintenance of life is made possible by the extraordinary effectiveness achieved by enzymes in accelerating chemical reactions and avoiding side reactions. The rate of ATP turnover in a mammalian cell gives us an idea of the rate at which cellular metabolism proceeds: every 1-2 minutes, the entire ATP pool is turned over, namely, hydrolyzed and restored by phosphorylation. This corresponds to the turnover of about 107 molecules of ATP per second, and, for the human body, to the turnover of  about 1 gram of ATP every minute. Some enzymes have even achieved catalytic perfection, that is, they are so efficient that nearly every collision with their substrates results in catalysis.
Multifunctional enzymesAnd one of the factors limiting the rate of an enzymatic reaction is just the frequency with which substrates and enzymes collide. The simplest way to increase the frequency of collisions would be to increase the concentration of substrates and enzymes. However, due to the large number of different reactions that take place in the cell, this route is not feasible. In other words, there is a limit to the concentration that substrates and enzymes can reach, concentrations that are in the micromolar range for substrates, and even lower for enzymes. Exceptions are glycolytic enzymes in muscle cells and erythrocytes, present in concentrations of the order of 0.1 mM and even higher.
One of the routes taken by evolution to increase the rate at which enzymatic reactions proceed is to select molecular structures, such as multifunctional enzymes and multienzyme complexes, that allow, through the optimization of the spatial organization of the enzymes of a metabolic pathway, to minimize the distance that the product of reaction A must travel to reach the active site that catalyzes the reaction B in the sequence, and so on, thus obtaining the substrate or metabolic channeling of the pathway itself. For some multifunctional enzymes and multienzyme complexes the channeling is obtained through intramolecular channels.
Metabolic channeling  increases the catalytic efficiency, and then the reaction rate, in various ways, briefly described below.

  • It minimizes the diffusion of substrates in the bulk solvent, then their dilution; this allows to obtain high local concentrations, even when their concentration in the cell is low, thus increasing the frequency of enzyme-substrate collisions.
  • It minimizes the time required by substrates to diffuse from one active site to the next.
  • It minimizes the probability of side reactions.
  • It minimizes the probability that labile intermediates are degraded.

Multifunctional enzymes offer advantages with regard to the regulation of their synthesis, too: being encoded by a single gene, it is possible to coordinate the synthesis of all the enzymatic activities.
Finally, like multienzyme complexes, multifunctional enzymes allow the coordinated control of their catalytic activities. And, because the enzyme that catalyzes the committed step of the sequence often catalyzes the first reaction, this prevents the synthesis of unneeded molecules, which would be produced if the control point were downstream of the first reaction, as well as a waste of energy and the removal of metabolites from other metabolic pathways.

Examples of multifunctional enzymes

Like multienzyme complexes, multifunctional enzymes, too, are very common and involved in many metabolic pathways, both anabolic and catabolic.
Here are some examples.

Acetyl-CoA carboxylase

Acetyl-CoA carboxylase or ACC (EC 6.4.1.2), a biotin-dependent carboxylase, is composed of two enzymes, a biotin carboxylase (EC 6.3.4.14) and a carboxyltransferase, plus a biotin carboxyl-carrier protein or BCCP. ACC catalyzes the synthesis of malonyl-CoA by the carboxylation of acetyl-CoA. The reaction, which is the committed step of fatty acid synthesis, proceeds in two steps. In the first step, biotin carboxylase catalyzes, at the expense of ATP, the carboxylation of a nitrogen atom of biotin, that acts as a carbon dioxide (CO2) carrier, while the source of CO2 is bicarbonate ion. In the second step, carboxyltransferase catalyzes the transfer of the carboxyl group from carboxybiotin to acetyl-CoA to form malonyl-CoA. Malonyl-CoA is the donor of an activated two carbon unit to fatty acid synthase (EC 2.3.1.85) during fatty acid elongation.
In mammals and birds, acetyl-CoA carboxylase is a multifunctional enzyme, as biotin carboxylase activity and carboxyltransferase activity, plus BCCP, are located on the same polypeptide chain.
Conversely, in bacteria it is a multienzyme complex made up of three distinct polypeptide chains, namely, the two enzymes plus BCCP.
Both forms are present in higher plants.

Type I fatty acid synthase

Fatty acid synthase or FAS catalyzes the synthesis of palmitic acid using malonyl-CoA, the product of the reaction catalyzed by acetyl-CoA carboxylase, as a donor of two-carbon units.
There are two types of FAS.
In animals and fungi, it is a multifunctional enzyme, and is called type I. In animals it is an homodimer, and each polypeptide chain contains all seven enzymatic activities plus acyl carrier protein or ACP. In yeast and fungi FAS consists of two multifunctional subunits, called α and β, arranged in an α6β6 heterododecameric structure.
In most prokaryotes and in plants, fatty acid synthase, called type II, it is not a multifunctional enzyme but a multienzyme complex, being composed of distinct enzymes plus ACP.

PRA-isomerase:IGP synthase

The synthesis of the amino acid tryptophan from chorismate involves several steps, briefly described below.
In the first step, glutamine donates a nitrogen to the indole ring of chorismate, that is converted to anthranilate, and glutamine to glutamate; the reaction is catalyzed by anthranilate synthase (EC 4.1.3.27). Anthranilate is phosphoribosylated to form N-(5’-phosphoribosyl)-anthranilate or PRA, in a reaction catalyzed by anthranilate phosphoribosyltransferase (EC 2.4.2.18); in the reaction 5-phosphoribosyl-1-pyrophosphate or PRPP acts as a donor of a 5-phosphoribosyl group. In the next step, catalyzed by PRA isomerase (EC 5.3.1.24), PRA is isomerized to enol-1-o-carboxyphenylamino-1-deoxyribulose phosphate or CdRP. CdRP is converted to indole-3-glycerol phosphate or IGP, in a reaction catalyzed by indole-3-glycerol phosphate synthase or IGP synthase (EC 4.1.1.48). Finally, tryptophan synthase (EC 4.2.1.20) catalyzes the last two steps of the pathway: the conversion of IGP to indole, a hydrolysis, and the reaction of indole with a serine to form tryptophan.
In E. coli, PRA isomerase and IGP synthase are located on a single polypeptide chain, which is therefore a bifunctional enzyme. In other microorganisms, such as Bacillus subtilis, Salmonella typhimurium and Pseudomonas putida, the two catalytic activities located on distinct polypeptide chains.
Conversely, tryptophan synthase is a classic example of a multienzyme complex, and one of the best characterized examples of metabolic channeling.

Glutamine-PRPP amidotransferase

Glutamine-PRPP amidotransferase or GPATase (EC 2.4.2.14) catalyzes the first of ten steps leading to de novo synthesis of purine nucleotides, namely, the formation of 5-phosphoribosylamine through the transfer of the glutamine amide nitrogen to PRPP. Note that glutamine acts as a nitrogen donor.
The reaction proceeds in two steps, which take place on different active sites, an N-terminal active site and a C-terminal active site. In the first step, the N-terminal active site catalyzes the hydrolysis of glutamine amide nitrogen to form glutamate and ammonia. In the second step, catalyzed by the C-terminal active site, which has phosphoribosyltransferase activity, the released ammonia is attached at the C-1 of PRPP to form 5-phosphoribosylamine. In this step the inversion of configuration about the C-1 position of the ribose, from α to β, occurs, then establishing the anomeric form of the future nucleotide.
There are three control points that cooperate in the regulation of de novo synthesis of purine nucleotides, and the reaction catalyzed by glutamine-PRPP amidotransferase, the first committed step of the pathway, is the first control point.
Like in bacterial carbamoyl phosphate synthetase complex (EC 6.3.4.16), the active sites of this multifunctional enzyme are connected through an intramolecular channel. However, this channel is shorter, being about 20 Å long, and lined by conserved nonpolar amino acids, then, it is highly hydrophobic. Lacking hydrogen-bonding groups, it does not impede the diffusion of the ammonia to the other active site.

CAD

The de novo synthesis of pyrimidine nucleotides occurs through a series of enzymatic reactions that, unlike de novo synthesis of purine nucleotides, begins with the formation of the pyrimidine ring, which is then bound to ribose 5-phosphate. The first three steps of the pathway are catalyzed sequentially by carbamoyl phosphate synthetase, aspartate transcarbamoylase (EC 2.1.3.2), and dihydroorotase (EC 3.5.2.3), and are common to all species.
In the first step, carbamoyl phosphate synthetase, which has two enzymatic activities, namely, a glutamine-dependent amidotransferase and a synthase, catalyzes the synthesis of carbamoyl phosphate from glutamine, bicarbonate ion and ATP. In the second step, which is the committed step of the metabolic pathway and is catalyzed by aspartate transcarbamoylase, carbamoyl phosphate reacts with aspartate to form N-carbamoyl aspartate. Finally, dihydroorotase, catalyzing the removal of H2O from N-carbamoyl aspartate, leads to the closure of the pyrimidine ring to form of L-dihydroorotate.
In eukaryotes, particularly in mammals, in Drosophila and Dictyostelium, a genus of amoebae, the three enzymatic activities are located on a single polypeptide chain, encoded by a gene derived from a gene fusion event occurred at least 100 million years ago. The multifunctional enzyme, known by the acronym CAD, is a homomultimer of three subunits or more.
Conversely, in prokaryotes, the three enzymes are distinct, and carbamoyl phosphate synthase is an example of a multienzyme complex.
In yeasts the dihydroorotase is present on a distinct protein.
Studies on enzyme activity have revealed the existence of a substrate channeling, more effective in yeast protein, with respect to the first two steps, than in that of mammals.

References

Alberts B., Johnson A., Lewis J., Morgan D., Raff M., Roberts K., Walter P. Molecular Biology of the Cell. 6th Edition. Garland Science, Taylor & Francis Group, 2015

Eriksen T.A., Kadziola A., Bentsen A-K., Harlow K.W. & Larsen S. Structural basis for the function of Bacillus subtilis phosphoribosyl-pyrophosphate synthetase. Nat Struct Biol 2000:7;303-8. doi:10.1038/74069

Hyde C.C., Ahmed S.A., Padlan E.A., Miles E.W., and Davies D.R. Three-dimensional structure of the tryptophan synthase multienzyme complex from Salmonella typhimurium. J Biol Chem 1988;263(33):17857-71

Hyde C.C., Miles E.W. The tryptophan synthase multienzyme complex: exploring structure-function relationships with X-ray crystallography and mutagenesis. Nat Biotechnol 1990:8;27-32. doi:10.1038/nbt0190-27

Michal G., Schomburg D. Biochemical pathways. An atlas of biochemistry and molecular biology. 2nd Edition. John Wiley J. & Sons, Inc. 2012

Muchmore C.R.A., Krahn J.M, Smith J.L., Kim J.H., Zalkin H. Crystal structure of glutamine phosphoribosylpyrophosphate amidotransferase from Escherichia coli. Protein Sci 1998:7;39-51. doi:10.1002/pro.5560070104

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

Yon-Kahn J., Hervé G. Molecular and cellular enzymology. Springer, 2009 [Google eBook]

Multienzyme complexes

Multienzyme complexes are discrete and stable structures composed of enzymes associated noncovalently that catalyze two or more sequential steps of a metabolic pathway.
They can be considered a step forward in the evolution of catalytic efficiency as they provide advantages that individual enzymes, even those that have achieved catalytic perfection, would not have alone.

CONTENTS

What advantages do multienzyme complexes provide?

During evolution, some enzymes evolved to reach a virtual catalytic perfection, namely, for such enzymes nearly every collision with their substrate results in catalysis. Examples are:

  • fumarase (EC 4.2.1.2), which catalyzes the seventh reaction of the citric acid cycle, the reversible hydration/dehydration of fumarate’s double bond to form malate;
  • acetylcholinesterase (EC 3.1.1.7), which catalyses the hydrolysis of acetylcholine, a neurotransmitter, to choline and acetic acid, which in turn dissociates to form an hydrogen ion and acetate;
  • superoxide dismutase (EC 1.15.1.1), which catalyzes the conversion, and then the inactivation, of the highly reactive superoxide radical, (O2.-), to hydrogen peroxide (H2O2) and water;
  • catalase (EC 1.11.1.6), which catalyzes the degradation of H2O2 to water and oxygen.

Then, the rate at which an enzymatic reaction proceeds is partly determined by the frequency with which enzymes and their substrates collide. Hence, a simple way to increase it is to increase the concentrations of enzymes and substrates. However, their concentrations cannot be high because of the enormous number of different reactions that occur within the cell. And in fact in the cells most reactants are present in micromolar concentrations (10-6 M), whereas most enzymes are present in much lower concentrations.
So, evolution has taken different routes to increase the reaction rate, one of which has been to optimize the spatial organization of enzymes with the formation of multienzyme complexes and multifunctional enzymes, that is, structures that allow minimizing the distance that the product of a reaction must travel to reach the active site that catalyzes the subsequent step in the sequence, being active sites close to each other. In other words, what happens is the substrate or metabolic channeling, that can also occur through intramolecular channels connecting the active sites, as in the case of, among the multienzyme complexes, tryptophan synthase complex (EC 4.2.1.20), whose tunnel was the first to be discovered, and bacterial carbamoyl phosphate synthase complex (EC 6.3.4.16).
Metabolic channeling can increase the reaction rate, but more generally, the catalytic efficiency, in several ways, briefly described below.

  • The diffusion of substrates and products in the bulk solvent is minimized, then their dilution and decrease of concentration, too. This leads to the production of high local concentrations, even when their intracellular concentration is low. In turn this leads to an increase in the frequency of enzyme-substrate collisions.
  • The time required by substrates to diffuse between successive active sites is minimized.
  • The probability of side reactions is minimized.
  • Chemically labile intermediates are protected from degradation by the solvent.

Another metabolic advantage of multienzyme complexes, similarly to what happens with multifunctional enzymes, is that they allow to control coordinately the catalytic activity of the enzymes that compose them. And taking into account that the enzyme that catalyzes the first reaction of a pathway is often the regulatory enzyme, it is possible to avoid:

  • the synthesis of unneeded intermediates, which would be produced if the sequence of reactions were regulated downstream of the first reaction;
  • the removal of metabolites from other pathways as well as a waste of energy.

Examples of multienzyme complexes

From what was said above, it is not surprising that, especially in eukaryotes, the multienzyme complexes, like multifunctional enzymes, are common and involved in different metabolic pathways, both anabolic and catabolic, whereas there are few enzymes that diffuse freely in solution. Below are some examples.

2-Ketoacid dehydrogenase family

A classic example of multienzyme complexes are the three complexes belonging to the 2-ketoacid dehydrogenase family, also called 2-oxoacid dehydrogenase family, namely:

  • the pyruvate dehydrogenase complex (PDC);
  • the branched-chain α-keto acid dehydrogenase complex (BCKDH);
  • the α-ketoglutarate dehydrogenase complex, also called 2-oxoglutarate dehydrogenase (OGDH).

These complexes are similar both from structural and functional points of view.
For example, PDC is composed of multiple copies of three different enzymes:

  • pyruvate dehydrogenase or E1 (EC 1.2.4.1);
  • dihydrolipoyl transacetylase or E2 ;(EC 2.3.1.12);
  • dihydrolipoyl dehydrogenase or E3 (EC 1.8.1.4).

Then, PDC, both in prokaryotes and eukaryotes, has the basic E1-E2-E3 structure, a structure also found in the other two complexes. Moreover, within a given species:

  • dihydrolipoyl dehydrogenase is identical;
  • pyruvate dehydrogenase and dihydrolipoyl transacetylase are homologous.

And, although these enzymes are specific for their substrates, they use the same cofactors, namely, coenzyme A, NAD, thiamine pyrophosphate, FAD, and lipoamide.
In order to differentiate them, for the pyruvate dehydrogenase complex, the α-ketoglutarate dehydrogenase complex, and the branched-chain α-ketoacid dehydrogenase complex, they are indicated, respectively:

  • E1p, E1o and E1b (EC 1.2.4.4);
  • E3p, E3o, and E3b (EC 1.8.1.4).

Note: the eukaryotic PDC is the largest multienzyme complex known, larger than a ribosome, and can be visualized with the electron microscope.

The pyruvate dehydrogenase complex is the bridge between glycolysis and the citric acid cycle, and catalyzes the irreversible oxidative decarboxylation of pyruvate, an α-keto acid. During the reactions the carboxyl group of pyruvate is released as carbon dioxide (CO2) and the resulting acetyl group is transferred to coenzyme A to form acetyl-coenzyme A. Furthermore, two electrons are released and transferred to NAD+.

Multienzyme Complexes
Fig. 1 – The Five Reactions Catalyzed by the PDC

Even during the reactions catalyzed by the α-ketoglutarate dehydrogenase complex and the branched-chain α-keto acid dehydrogenase complex, respectively, the fourth reaction of the citric acid cycle, the oxidation of α-ketoglutarate to succinyl-CoA, and the oxidation of α-ketoacids deriving from the catabolism of the branched-chain amino acids valine, leucine and isoleucine, it occurs:

  • the release of the carboxyl group of the α-keto acid as CO2;
  • the transfer of the resulting acyl group to coenzyme A to form the acyl-CoA derivatives;
  • the reduction of NAD+ to NADH.

The remarkable similarity between protein structures, required cofactors and reaction mechanisms undoubtedly reflect a common evolutionary origin.

What are keto acids?

Keto acids or oxoacids are organic compounds containing two functional groups: a carboxyl acid group and a ketone group. Depending on the position of the ketone group, alpha-keto acids, beta-keto acids and gamma-keto acids can be identified.

  • Alpha-keto acids or 2-oxoacids have the ketone group at position α (2) from the carboxylic acid group, that is, adjacent to it. These compounds are important in biology, being involved in glycolysis, like pyruvic acid, the simplest α-keto acid, and in the citric acid cycle, like oxalacetic acid and α-ketoglutaric acid.
  • Beta-ketoacids or 3-oxoacids have the ketone group is at position β (3) from the carboxylic acid group. An example is acetoacetic acid, the simplest β-ketoacid, and one of the three ketone bodies, together with acetone and β-hydroxybutyric acid, produced by the hepatocyte in presence of an excess of acetyl-CoA, such as during fasting or low-carbohydrate diets.
  • Gamma-keto acids or 4-oxoacids have the ketone group is in position γ (4) from the carboxylic acid group. An example is levulinic acid, the simplest beta-keto acid, deriving from the catabolism of cellulose.
Keto Acids
Fig. 2 – Keto Acids

Tryptophan synthase complex

The tryptophan synthase complex is one of the best-studied examples of substrate channeling. Present in bacteria and plants, but not in animals, in bacteria it is composed of two α and two β subunits associated as αβ dimers, which are considered the functional unit of the complex, in turn associated to form an αββα tetramer.
The complex catalyzes the final two steps of the synthesis of tryptophan. In the first step, indole-3-glycerol phosphate undergoes an aldol cleavage, catalyzed by a lyase (EC 4.1.2.8) present on the α subunits, to yield indole and a molecule of glyceraldehyde 3-phosphate. Indole then reaches the active site of the β subunit via a about 30 Å long hydrophobic tunnel that, in each αβ dimers, connects the two active sites. In the second step, in the presence of pyridoxal 5-phosphate, a condensation between indole and a serine forms tryptophan.

Acetyl-CoA carboxylase

Acetyl-CoA carboxylase (ACC) (EC 6.4.1.2), a member of the biotin-dependent carboxylase family, catalyzes the committed step of de novo fatty acid synthesis, namely, the carboxylation of acetyl-CoA to malonyl-CoA, which, in turn, serves as a donor of two-carbon units for the elongation process leading to the synthesis of palmitic acid, catalyzed by fatty acid synthase (EC 2.3.1.85).
In bacteria, ACC is an multienzyme complex composed of two enzymes, biotin carboxylase (EC 6.3.4.14) and a carboxytransferase, plus a biotin carboxyl-carrier protein or BCCP.
Conversely, in mammals and birds, it is a multifunctional enzyme, as the two enzymatic activities, and BCCP, are present on the same polypeptide chain.
In higher plants both forms are present.

Carbamoyl phosphate synthetase complex

Another well-characterized example of substrate channeling is the bacterial carbamoyl phosphate synthetase complex, which catalyzes the synthesis of carbamoyl phosphate, needed for pyrimidine and arginine synthesis. The complex has a about 100 Å long tunnel that connects the three active sites.
The first active site catalyzes the release of the amide nitrogen of glutamine as ammonium ion, that enters the tunnel and reaches the second active site where, at the expense of ATP, is combined with bicarbonate to yield carbamate, that, in the last active site, is phosphorylated to carbamoyl phosphate.

References

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Hyde C.C., Miles E.W. The tryptophan synthase multienzyme complex: exploring structure-function relationships with X-ray crystallography and mutagenesis. Nat Biotechnol 1990:8;27-32. doi:10.1038/nbt0190-27

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Pyruvate dehydrogenase complex: location, functions, enzymes, and regulation

The pyruvate dehydrogenase complex (PDC) is a mitochondrial multienzyme complex composed of three different enzymes:

  • pyruvate dehydrogenase or E1 (EC 1.2.4.1);
  • dihydrolipoyl transacetylase or dihydrolipoamide acetyltransferase or E2 (EC 2.3.1.12);
  • dihydrolipoyl dehydrogenase or dihydrolipoamide dehydrogenase or E3 (EC 1.8.1.4).

Each of these enzymes is present in multiple copies whose number, and then the size of the complex itself, varies from species to species, with molecular masses ranging from 4×106 to 1×107 daltons.
The multienzyme complex contains additional subunits:

  • five different coenzymes;
  • in plants, fungi, and among animals, aves and mammals, two enzymes with regulatory properties: a Mg2+-dependent pyruvate dehydrogenase kinase (EC 2.7.1.99) and a Ca2+-activated pyruvate dehydrogenase phosphatase(EC 3.1.3.43);
  • in eukaryotes, a binding protein called E3BP.

The pyruvate dehydrogenase complex catalyzes, through five sequential reactions, the oxidative decarboxylation of pyruvate, an α-keto acid, to form a carbon dioxide molecules (CO2) and the acetyl group of acetyl-coenzyme A or acetyl-CoA, with the release of two electrons, carried by NAD.

Pyruvate Dehydrogenase Complex
Fig. 1 – PDC Reaction

The overall reaction is essentially irreversible, with a ΔG°’ of -8.0 kcal/mol (-33.4 kJ/mol), and requires the intervention of the three enzymes, whose activities are sequentially coordinated. During the reactions, the intermediate products remain bound to the enzymes and, at the end of the reaction sequence, the multienzyme complex is ready for the next cycle;.

Pyruvate Dehydrogenase Complex
Fig. 2 – The Five Reactions Catalyzed by the PDC

Note: the pyruvate dehydrogenase complex catalyzes the same reactions through similar mechanisms in all organisms.

CONTENTS

The coenzymes of the pyruvate dehydrogenase complex

Five coenzymes are used in the pyruvate dehydrogenase complex reactions: thiamine pyrophosphate or TPP, flavin adenine dinucleotide or FAD, coenzyme A or CoA, nicotinamide adenine dinucleotide or NAD, and lipoic acid.

  • Thiamine pyrophosphate is the active form of thiamine or vitamin B1.
    Thiamine Pyrophosphate
    Fig. 3 – TPP

    TPP is the coenzyme of pyruvate dehydrogenase, to which it is strictly bound through noncovalent interactions. It is involved in the transfer of hydroxyethyl or “activated aldehyde” groups.

  • Flavin adenine dinucleotide is one the active forms of riboflavin or vitamin B2; the other is flavin mononucleotide (FMN).
    Flavin Adenine Dinucleotide
    Fig. 4 – FAD

    FAD is the coenzyme of dihydrolipoyl dehydrogenase, to which it is strictly bound. Like NAD, it participates in electron transfer, or hydride ion (:H or H+ + 2e) transfer.

  • Coenzyme A consists of a β-mercaptoethylamine group connected to pantothenic acid or vitamin B5 through an amide linkage, which, in turn, is bonded to 3′-phosphoadenosine moiety, through a pyrophosphate bridge.
    CoA is involved in the reaction catalyzed by dihydrolipoyl transacetylase, and acts as a carrier of acyl groups.

    Coenzyme A
    Fig. 5 – Coenzyme A

    The β-mercaptoethylamine moiety terminates with a sulfhydryl group (–SH), a reactive thiol crucial for the role played by the coenzyme, because the acyl groups bonded to it through a thioester bond have a high standard free energy of hydrolysis. This provides acyl groups with a high transfer potential, equal to -31.5 kcal/mol (-7.5 kJ/mol), slightly more exergonic [1 kJ/mol (0.2 kcal/mol )] than that for the hydrolysis of ATP to ADP and Pi. Thioesters have therefore a high transfer potential of the acyl group and can donate it to a variety of molecules, that is, such acyl group can be considered as an activated group ready for a transfer. It is also possible to state that the formation of the thioester bond allows to conserve a portion of the free energy derived from the oxidation of the metabolic fuel. It should be noted that coenzyme A is also abbreviated as CoA-SH to emphasize the role played by the thiol group.
    Note: in the thioester bond a sulfur atom sits in the position where an oxygen atom is in the ester bond.

 Thioester and Ester Bonds
Fig. 6 – Ester and Thioester Bonds
  • Nicotinamide adenine dinucleotide can be synthesized from tryptophan, an essential amino acid, or from niacin or vitamin B3 or vitamin PP, from Pellagra-Preventing, the source of the nicotinamide moiety.
Nicotinamide Adenine Dinucleotide
Fig. 7 – NAD

NAD is involved in the reaction catalyzed dihydrolipoyl dehydrogenase, and, like FAD, participates in electron transfer, or hydride transfer.

  • Unlike the other coenzymes of the pyruvate dehydrogenase complex, lipoic acid does not derive, directly or indirectly, from vitamins and/or essential amino acids, that is, from building blocks that cannot be synthesized de novo by the organism and must be supplied by the diet.
    It is the coenzyme of dihydrolipoyl transacetylase, to which it is covalently bound, through an amide linkage, to the ɛ-amino group of a lysine residue to form a lipoyl-lysine or lipoamide, the so-called lipoyllysyl arm. It couples electron transfer to acyl group transfer.

    Pyruvate Dehydrogenase Complex
    Fig. 8 – Lipoyl-lysine

    Lipoic acid has two thiol groups that can undergo a reversible intramolecular oxidation to form a disulfide bridge (-S-S-), a reaction analogous to that between two cysteine (Cys) residues of a protein.
    Because the disulfide bridge (note: a cyclic disulfide) is capable of undergoing redox reactions, during the reactions catalyzed by the pyruvate dehydrogenase complex, it is first reduced to dihydrolipoamide, a dithiol or the reduced form of the prosthetic group, and then, reoxidized to the cyclic form.

Note
Many enzymes require small non protein components, called cofactors, for their catalytic activity. Cofactors can be metal ions or small organic or metalloorganic molecules, and are classified as coenzymes and prosthetic groups.
A prosthetic group is cofactor that binds tightly to an enzyme by non-covalent or covalent bond, namely, it is permanently bound to the protein.
A coenzyme is cofactor that is not permanently bound to the enzyme.

Where is the pyruvate dehydrogenase complex located?

In eukaryotes, the pyruvate dehydrogenase complex, like the enzymes for citric acid cycle and oxidation of fatty acids, is located in the mitochondrion, where is associated with the surface of the inner membrane facing the matrix.
In prokaryotes, it is located in the cytosol.

Functions of the pyruvate dehydrogenase complex

The main functions of the pyruvate dehydrogenase complex are to produce acetyl-CoA and NADH.

Acetyl-CoA
Fig. 9 – Acetyl-CoA
  • The acetyl group linked to coenzyme A, an activated acetate, depending on the metabolic conditions within the cell and/or the cell type, can be:

oxidized to two carbon dioxide molecules via the citric acid cycle reactions to harvest a portion of the potential energy stored in the form of ATP or GTP;
utilized for the synthesis of fatty acids, cholesterol, steroids, isoprenoids, ketone bodies and acetylcholine.

It is therefore possible to state that, depending on the metabolic conditions and/or cell type, the pyruvate dehydrogenase complex commits carbon intermediates from amino acid and glucose catabolism to:

citric acid cycle, and then to the production of energy, e.g. in skeletal muscle in aerobic conditions, and, always, in cardiac muscle;
synthesis of lipids and acetylcholine.

  • In aerobic organisms, NADH can be oxidized to NAD+ via hydride ion transfer to the mitochondrial electron transport chain that, in turn, carries the two electrons to molecular oxygen (O2), allowing the production of 2.5 ATP molecules per pair of electrons.
    Pyruvate Dehydrogenase Complex
    Fig. 10 – Nicotinamide Ring: Reduced Form

    Note: in anaerobic organisms there are electron acceptors alternative to oxygen, such as sulfate or nitrate.

Conceptually, the pyruvate dehydrogenase complex is the bridge between glycolysis and the citric acid cycle. However, due to the irreversibility of the overall reaction catalyzed by the multienzyme complex, it acts as an one-way bridge: pyruvate can be decarboxylated, oxidized and the remaining acetyl unit linked to the CoA, but it is not possible to carry out the opposite reaction, namely, to convert acetyl-CoA into pyruvate.
The irreversibility of this reaction, and the absence of alternative pathways, explain why it is not possible to use acetyl-CoA, and therefore fatty acids, as a substrate for gluconeogenesis (see below).

Other sources of acetyl-CoA

Other than pyruvate, the acetyl group of acetyl-CoA can derive from the oxidation of fatty acids and the catabolism of many amino acids. However, regardless of its origin, acetyl-CoA represents an entry compound for new carbon units into the citric acid cycle. And, it is also possible to state that the acetyl group of acetyl-CoA represents the form in which most of the carbon enters the cycle.

Sources of pyruvate

Pyruvate can derive from different cytosolic sources.

  • Under physiological conditions, in most cells it derives mainly from glycolysis: the oxidation of a glucose molecule yields two pyruvate molecules.
  • Lactate can be oxidized to pyruvate in the reaction catalyzed by lactate dehydrogenase (EC 1.1.1.27). In fact, isoenzymes in which the H subunit predominates, such as LDH1 or H4, a homopolymer of H subunits found in cardiac muscle, a tissue with completely aerobic metabolism, preferentially catalyze lactate oxidation.
    Lactate oxidation can occurs in hepatocytes, too, during gluconeogenesis, favored by the low NADH/NAD+ ratio in the cytosol.
    It should be noted that lactate is a metabolite deriving from the catabolism of glucose, and therefore from carbohydrate catabolism.
  • Another source is malate, by the reaction catalyzed by the cytosolic malic enzyme (EC 1.1.1.40), which is implicated in the exchange of intermediates of the citric acid cycle, such as oxaloacetate, malate and citrate, between the cytosol and the mitochondrial matrix.

Malate + NADP+ → Pyruvate + CO2 + NADPH + H+

  • Finally, the carbon skeleton of six amino acids, namely, alanine, cysteine, glycine, serine, threonine and tryptophan, can be partly or completely converted into pyruvate.

In turn, pyruvate is a metabolic intermediate which can be metabolized by different pathways, both anabolic and catabolic. Examples are gluconeogenesis, the synthesis of lipids, and the oxidative metabolism. In addition, it can have an anaplerotic function, playing a role in the maintenance of the metabolic flow through the acid cycle citric.

Mitochondrial pyruvate transport

In eukaryotes, glycolysis occurs in the cytosol whereas all the subsequent steps of the aerobic metabolism, that is, the reactions catalyzed by the pyruvate dehydrogenase complex, the citric acid cycle, the electron transport chain, and the oxidative phosphorylation occur in the mitochondria.
Similarly to most other metabolites and anions, the transport of pyruvate across the outer mitochondrial membrane is probably mediated by a relatively non-specific, voltage-dependent anion channel. Conversely, its transport across the inner mitochondrial membrane occurs through a specific transporter made up of two proteins named MPC1 and MPC2, acronym of mitochondrial pyruvate carrier, which form a hetero-oligomeric complex in the membrane.

Structure of the pyruvate dehydrogenase complex

Although the pyruvate dehydrogenase complex is composed of multiple copies of three different enzymes and catalyzes the same reactions by similar mechanisms in all the organisms in which it is present, it has a very different quaternary structure.
The structure of E. coli multienzyme complex, which has a weight of ∼4,600 kD and a diameter of ∼300 Å, was the first to be characterized, thanks to the work of Lester Reed. In this complex, 24 units of dihydrolipoyl transacetylase form a structure with cubic symmetry, namely, the enzymes, associated as trimers, are placed at the corners of a cube. Dimers of pyruvate dehydrogenase are associated with dihydrolipoyl transacetylase core, at the center of each edge of the cube, for a total of 24 units. Finally, dimers of dihydrolipoyl dehydrogenase are located at the center of each of the six faces of the cube, for a total of 12 units. Note that the entire complex is composed of 60 units. A similar structure with cubic symmetry is also found in most other Gram-negative bacteria.
In some Gram-positive bacteria and in eukaryotes, the pyruvate dehydrogenase complex has a dodecahedral form, namely, that of a regular polyhedron with 20 vertices, 12 pentagonal faces, and 30 edges, with icosahedral symmetry, also called I symmetry. Considering for example the multienzyme complex present in mitochondria, it is the largest known multienzyme complex, with a weight of ∼10,000 kD and a diameter of ∼500 Å, so more than 5 times the size of a ribosome. Noteworthy, it can be visualized with the electron microscope. The complex is composed of a dodecahedral core, with a diameter of about 25 nm, formed, as in Gram-negative bacteria, by dihydrolipoyl transacetylase, but consisting of 20 trimers of the enzyme,  for a total of 60 units, located at the vertices of the structure. The core is surrounded by 30 units of pyruvate dehydrogenase, one centered on each edge, and 12 units of dihydrolipoyl dehydrogenase, one centered on each face. The entire complex is therefore composed of 102 units.

The quaternary structure of the complex is further complicated, as mentioned previously, by the presence of three additional subunits: a pyruvate dehydrogenase kinase, a pyruvate dehydrogenase phosphatase, and the E3-binding protein.
Kinase and phosphatase are bound to the dihydrolipoyl transacetylase core.
E3BP is bound to each of the 12 pentagonal faces, and therefore is present in about 12 copies. It is required to bind dihydrolipoyl dehydrogenase to the core of dihydrolipoyl transacetylase, as demonstrated by the fact that its partial proteolysis decreases the binding ability of the dehydrogenase. In E3-binding protein it is possible to identify a C-terminal domain, that has no catalytic activity, and a lipoamide-containing domain, similar to that of dihydrolipoyl transacetylase, capable of accepting an acetyl group, too. However, the removal of this domain does not cause any reduction of the catalytic activity of the multienzyme complex.

Structure of pyruvate dehydrogenase or E1

Pyruvate dehydrogenase of eukaryotes and some Gram-positive bacteria is composed of two different polypeptide chains, called α and β, associated to form a 2-fold symmetric α2β2 heterotetramer. Conversely, in E. coli and other Gram-negative bacteria the two subunits are fused to form a single polypeptide chain, and the enzyme is a homodimer.
The enzyme has two active sites.
Considering the heterotetrameric structure of Bacillus stearothermophilus pyruvate dehydrogenase, a Gram-positive bacteria, each thiamine pyrophosphate binds between the N-terminal domains of an α and a β subunit, at the end of a ∼21 Å deep funnel-shaped channel leading to the active site, with its reactive group, the thiazole ring (see fig. 3), closest to the channel entrance. At the entrance this channel there are also two conserved loops, essential both for the catalytic activity of the enzyme and for its regulation. The X-ray analysis of B. stearothermophilus enzyme, when it binds both TPP and the peripheral subunit-binding domain (PSBD) of dihydrolipoyl transacetylase, which binds to the C-terminal domain of the β subunits, has revealed that, in addition to a heterotetramer with a very tight structure, the two active sites have a different structure, in particular regarding the arrangement of the two conserved loops. In fact, in one enzyme subunit, in the presence of the activated form of thiamine pyrophosphate, the inner loop is ordered in a way that it blocks the entrance to the active site, whereas the loop at the entrance of the other active site is disordered and does not block the entrance. This explains, from a structural point of view, the observed differences in the rate of substrate binding exhibited by the two active sites. A similar arrangement and asymmetry have been observed in all thiamine pyrophosphate-dependent enzymes of which the structure has been solved.
In addition to TPP and a magnesium ion (Mg2+), located in each of the two active sites, a third Mg2+ is located at the centre of the tetramer, within a ∼20 Å deep solvent-filled tunnel that connects the two active sites. The tunnel is largely lined by 10 conserved amino acid residues from all four subunits, in particular glutamate (Glu) and aspartate (Asp), six and four, respectively, plus other acidic residues around the TPP aminopyrimidine ring. And it should be underlined the absence of basic residues to neutralize them. Similar tunnels have been found in all thiamine pyrophosphate-dependent enzymes with known crystalline structure, with dimeric or tetrameric structure, for example in transketolase, an enzyme of the pentose phosphate pathway.

Note: B. stearothermophilus belongs to the phylum Firmicutes and is recently renamed Geobacillus stearothermophilus.

What is the function of the acidic tunnel?

Through mutagenesis experiments on B. stearothermophilus pyruvate dehydrogenase, the tunnel has been shown to play a role in the catalytic mechanism.
The change of some of the aforementioned acidic residues to neutral amino acids does not alter, compared with the wild-type pyruvate dehydrogenase:

  • the efficiency of incorporation of the modified enzyme into the multienzyme complex;
  • the structure of active sites;
  • the quaternary structure of the enzyme.

However, rate of decarboxylation is reduced by over 70% compared to the wild-type enzyme, as well as, once the multienzyme complex is assembled with the mutant pyruvate dehydrogenase, the PDC activity, which is reduced by over 85% compared to the wild-type complex. But how does this occur?
Because the distance between the substituted amino acids and the active sites is ≥7 Å, that is, these amino acids are remote from the active sites of pyruvate dehydrogenase, they cannot directly influence its catalytic activity. Then, the catalytic mechanism described below was proposed.
Considering the apoenzyme, thiamine pyrophosphate binds fast and strongly to the first active site, is activated, and the active site is closed, thus protecting the zwitterionic thiazolium from the external environment.
Conversely, in the second active site TPP binds, but is not activated, and the active site remains in an open conformation.
In the first active site, pyruvate reacts with the thiazolium C-2, and thiamine pyrophosphate of the second active site, which is a general acid, donates a proton to the first site. The result is a decarboxylation in the first site and the activation of the coenzyme in the second site, which is then closed.
It should be noted that while the activation of the first thiamine pyrophosphate is the result of the binding to the active site, the activation of the second coenzyme, and therefore of the second active site, is coupled to the decarboxylation of pyruvate in the first active site. Or, from another point of view, while an active site requires a general acid, the other requires a general base.
Protons are needed for the catalytic activity, and their transport between the two active sites occurs via the acidic tunnel. They are reversibly shuttled along a chain of donor-acceptor groups provided by glutamate and aspartate residues and the entrained water, that act as a proton wire.
It seems, therefore, that unlike many other enzymes, in which the communication between the active sites occurs through conformational changes and subunit rearrangements, in pyruvate dehydrogenase and in the other TPP-dependent enzymes, the proton wire is the molecular basis of such communication.
At this point, the holoenzyme has been formed and the active sites are in a dynamic equilibrium, each exchanging between the dormant and the activated state. This seems to be the state in which the enzyme is found in vivo at the start of each catalytic cycle.
A consequence of such a mechanism is that, as the catalytic cycles occur, the two active sites are out of phase with each other, namely, when an active site requires a general acid, the other requires a general base, and vice versa.
Finally, it should be noted that this mechanism allows the switching of the loops that close the active sites so as to:

  • coordinate substrate uptake and product release;
  • explain the asymmetry existing between the two active sites.

Note: an apoenzyme is an enzyme that lacks the association of its cofactors. Conversely, an holoenzyme is an apoenzyme together with its cofactors. The apoenzyme is a catalytically inactive enzyme, whereas the holoenzyme is a catalytically active enzyme.

Structure of dihydrolipoyl transacetylase or E2

Three functionally distinct domains can be identify in the structure of dihydrolipoyl transacetylase: an N-terminal lipoyl domain, a peripheral subunit-binding domain, and a C-terminal catalytic domain or acyltransferase domain. These domains are connected by 20- to 40 amino acid residues rich in alanine and proline, hydrophobic amino acids that are interspersed with charged residues. These linkers are highly flexible and largely extended, that allows the three domains to kept away from each other.

Pyruvate Dehydrogenase Complex
Fig. 11 – E2 Domains

Note: flexible linkers are present in E3BP, too.

  • The N-terminal lipoyl domain is composed of ∼80 amino acid residues, and is so called because it binds lipoic acid. The number of these domains depend on the species, ranging from one to three. For example, there is one domain in B. stearothermophilus and in yeasts, two in Streptococcus faecalis and in mammals, and three in Azotobacter vinelandii and E. coli.
    The link between the ɛ-amino group of a lysine residue and lipoic acid leads to the formation of a flexible arm, the lipoyl-lysine, which has a maximum extended length of ∼14 Å. Adding the polypeptide segment which connects the N-terminal domain to the adjacent domain, whose length is greater than 140 Å, the resulting flexible tether is able to swings the lipoyl group between the active sites of pyruvate dehydrogenase and dihydrolipoyl dehydrogenase, as well as to interact with neighboring dihydrolipoyl transacetylases of the core.
    It should be noted that the number of these tethers is 3 x 24 = 72 in E. coli, whereas in mammals 2 x 60 = 120, based on the number of N-terminal domains and the units of dihydrolipoyl transacetylase.
    One pyruvate dehydrogenase can therefore acetylate numerous dihydrolipoyl transacetylases, and one dihydrolipoyl dehydrogenase can reoxidize many dihydrolipoamide groups.
    Moreover, it also occurs:

an interchange of the acetyl groups between the lipoyl groups of the dihydrolipoyl transacetylase core;
the exchange of both acetyl groups and disulfides between the tethered arms.

  • PSBD is composed of ∼35 amino acid residues arranged to form a globular structure that binds to both pyruvate dehydrogenase and dihydrolipoyl dehydrogenase, that is, it holds the multienzyme complex together.
  • The C-terminal catalytic domain, which, of course, contains the active site, is composed of ∼250 amino acid residues arranged to form a hollow cage-like structure containing channels large enough to allow substrates and products to diffuse in and out. For example, CoA ad lipoamide, the two substrate of dihydrolipoyl transacetylase, bind, in their extended conformation, at the opposite ends of a channel located at the interface between each pair of subunits in each trimers.

Structure of dihydrolipoyl dehydrogenase or E3

The structure of dihydrolipoyl dehydrogenase was deduced from studies of the enzyme in several microorganisms. It has a homodimeric structure, with each ∼470 amino acid residue chain folded into four domains, from the N-terminal to the C-terminal end: a FAD-binding domain, a NAD+-binding domain, a central domain, and an interface domain. All domains participate in the formation of the active site.
FAD is almost completely hidden inside the protein because, unlike thiol or NADH, it is easily oxidizable and must therefore be protected from the surrounding solution, namely, from O2. In fact, in the absence of the nicotinamide coenzyme, the phenol side chain of a tyrosine residue (Tyr), for example Tyr181 in the Gram-negative bacteria Pseudomonas putida, covers the NAD+-binding pocket so as to protect FADH2 from the contact with the surrounding solution.

Pyruvate Dehydrogenase Complex
Fig. 12 – FADH2

Conversely, when NAD+ is located in the active site, the phenol side chain of the aforementioned tyrosine residue is interposed between the nicotinamide ring and the flavin ring.
In the active site of the enzyme’s oxidized form is also present a redox-active disulphide bridge. It forms between two cysteine residues located in a highly conserved segment of the polypeptide chain, e.g., Cys43 and Cys48 in P. putida, and is located on opposite side of the flavin ring with respect to the nicotinamide ring. The disulphide bridge links consecutive turns in a segment of a distorted α-helix, and, noteworthy, in the absence of such distortion, Cα atoms of the two cysteine residues would be too distant to allow the disulfide bridge to form.
Dihydrolipoyl dehydrogenase has therefore two electron acceptors: FAD and the redox-active disulphide bridge.
Note: the heterocyclic rings of NAD and FAD are parallel and in contact through van der Waals interactions; S48 is also in contact through van der Waals interactions with the flavin ring, on the opposite side of it from the NAD ring.

Reaction of pyruvate dehydrogenase or E1

In the reaction sequence catalyzed by components of the pyruvate dehydrogenase complex, pyruvate dehydrogenase catalyzes the first two steps, namely:

  • the decarboxylation of pyruvate to form CO2 and the hydroxyethyl-TPP intermediate;
  • the reductive acetylation of the lipoyl group of dihydrolipoyl transacetylase.

The first reaction is essentially identical to pyruvate decarboxylase reaction (EC 4.1.1.1), which carries out a non-oxidative decarboxylation in glucose fermentation to ethanol. What differs is the fate of the hydroxyethyl group bound to thiamine pyrophosphate that, in the reaction catalyzed by pyruvate dehydrogenase is transferred to the next enzyme in the sequence, dihydrolipoyl transacetylase, whereas in the reaction catalyzed by pyruvate decarboxylase is converted into acetaldehyde.

Catalytic mechanism of pyruvate dehydrogenase or E1

In thiamine pyrophosphate-dependent enzymes, the thiazolium ring is the active center, but only as dipolar carbanion or ylid, namely, as a dipolar ion, or zwitterion (German for “hybrid ion”), with positive charge on the N-3 and negative charge on C-2. Conversely, the positively charged thiazolium ring, that is, positive charged nitrogen and no charge on C-2, can be defined as “dormant” or inactive form.
The reaction begins with the nucleophilic attack by C-2 carbanion to the carbonyl carbon of pyruvate, which has the oxidation state of an aldehyde, and leads to the formation of a covalent bond between coenzyme and pyruvate.
Then, the cleavage of C-1–C-2 bond of pyruvate occurs. This leads to the release of the carboxyl group, namely, of the C-1 as CO2, while the remaining carbon atoms, C-2 and C-3, stay bound to the thiamine pyrophosphate as hydroxyethyl group. The cleavage of the C-1–C-2 bond, and therefore the decarboxylation of pyruvate, is favored by the fact that the negative charge on the C-2 carbon, that is unstable, is stabilized by the presence in the thiazolium ring of the positively charged N-3, a imine nitrogen (C=N+), that is, due to the presence of an electrophilic or electron deficient structure that acts as an electron sink or electron trap, in which the carbanion electrons can be delocalized by resonance.
At this point, the intermediate stabilized by resonance can be protonated to form hydroxyethyl-TPP.
Note: this first reaction catalyzed by pyruvate dehydrogenase is that in which the private dehydrogenase complex exercises its substratum specificity; furthermore, it is the slowest of the five reactions, hence limiting the rate of the overall reaction.

Pyruvate Dehydrogenase Complex
Fig. 13 – E1: Catalytic Mechanism

The enzyme then catalyzes the oxidation of the hydroxyethyl group to an acetyl group, and its transfer on lipoyllysyl arm of dihydrolipoyl transacetylase. The reaction begins with the formation of a carbanion on the hydroxylic carbon of the hydroxyethyl-TPP, by the removal of the carbon-linked proton by an enzyme base.
The carbanion carries out a nucleophilic attack on the lipoamide disulfide, with the formation of a high-energy acetyl-thioester bond with one of the two -SH groups. In this reaction, the oxidation of the hydroxyethyl group to an acetyl group occurs with the concomitant reduction of the lipoamide disulfide bond: the two electrons removed from the hydroxyethyl group are used to reduce the disulfide. This reaction is therefore a reductive acetylation accompanied by the regeneration of the active form of pyruvate dehydrogenase, namely, the enzyme with the thiazolium C-2 in the deprotonated form, the ylid or dipolar carbanion form.

Note that the energy derived from the oxidation of the hydroxyethyl group to an acetyl group drives the formation of the thioester bond between the acetyl group and coenzyme A.

Note: as previously said, the lipoyllysyl arm, arranged in an extended conformation in the channel where TPP is also found, allows the transfer of hydroxyethyl from hydroxyethyl-TPP to CoA, that is, it can move from the active site of pyruvate dehydrogenase to the active sites of dihydrolipoyl transacetylase, and then of dihydrolipoyl dehydrogenase.

A deeper look on thiamine pyrophosphate

Thiamine pyrophosphate molecule consists of three chemical moieties, from which its chemistry and enzymology depend: a thiazolium ring, a 4-aminopyrimidine ring, and the diphosphate side chain (see fig. 3).
The diphosphate side chain binds the cofactor to the enzyme via the formation of electrostatic bonds between the negative charges carried by its phosphoryl groups and the positive charges carried by Ca2+ and Mg2+ ions, in turn, bound to highly conserved sequences, GlyAspGly (GDG) and GlyAspGly-X26-AsnAsn (GDG-X26-NN), respectively.
The thiazolium ring plays a central role in catalysis, due to its ability to form the C-2 carbanion, that is, a nucleophilic center on the C-2 atom.
Note: as mentioned previously in this article, once bound to the enzyme, thiamine pyrophosphate locates in the active site so that the thiazolium ring is positioned close to the channel entrance leading to the active site.
The aminopyrimidine ring has a dual function:

  • it anchors the coenzyme holding it in place;
  • it has a specific catalytic role, participating in acid/base catalysis, as evidenced by studies with thiamine pyrophosphate analogs in which each of the three nitrogen atoms of the ring were replaced in turn. These studies demonstrated that the N-1’ atom and the N-4’-amino group are required, whereas the other nitrogen atom of the ring, the N-3’ atom, is required to a lesser extent.

How is the dipolar carbanion of thiamine pyrophosphate formed?

Three tautomeric forms of the aminopyrimidine ring can be identified in the enzyme-bound coenzyme not involved in the reaction:

  • the canonical 4’-aminopyrimidine tautomer;
  • the N-1 protonated form, that is, 4-aminopyrimidinium ion;
  • the 1’,4’-iminopyrimidine tautomer.

It seems that the 1′,4′-imino tautomer is the tautomer that undergoes deprotonation, before the entry of the substrate into the active site. The C-2 of the thiazolium ring is “much more acidic than most =C-H groups found in other molecules” (see also in the article on the pentose phosphate pathway). The higher acidity, i.e., the fact that the C-2 proton is easily dissociable, is due to the presence of the quaternary nitrogen on the thiazolium ring, a positively charged nitrogen atom able to electrostatically stabilize the resulting carbanion. In the deprotonation reaction, the amino group of the aminopyrimidine ring seems to play an essential role: it acts as a base and is suitably positioned to accept the proton. However, in the 4’-aminopyrimidine tautomer one of its protons sterically collides with the C-2 proton; in addition, its pK is too low to carry out the deprotonation efficiently. A mechanism was therefore proposed, in which the side chain of a conserved glutamate residue, for example βGlu59 in B. stearothermophilus, or Glu51 in Saccharomyces uvarum (brewer’s yeast) pyruvate decarboxylase, donates a proton to the aminopyrimidine, converting it to its 1′,4′-iminopyrimidine tautomer that, accepting the C-2 proton, returns to the canonical 4′-aminopyrimidinic form and allows the formation of the carbanion.

Note: carbanion formation on C-2 is a consequence of an intramolecular proton transfer.

Deprotonation of thiamine pyrophosphate and closure of pyruvate dehydrogenase active site

The loss of the C-2 proton of the thiazolium ring leads, from a positively charged ring, to a dipolar ion, or zwitterion. This change in state of charge triggers a conformational change in one of the two conserved loops at the entrance of the active site channel, specifically, the inner of these loops, that, in turn, leads to the closure of the channel to the surrounding water environment. In this closed conformation the thiazolium carbanion is protected against electrophiles.
To sum up: the deprotonation of thiamine pyrophosphate leads to the closure of the active site and the protection of the newly formed dipolar carbanion, that is, TPP-dependent enzymes would be only active in closed conformation.
Conversely, in the other active site, thiamine pyrophosphate is not in the ylid form, the channel is open, and the site is inactive.

Reaction of dihydrolipoyl transacetylase or E2

In the reaction sequence catalyzed by components of the pyruvate dehydrogenase complex, dihydrolipoyl transacetylase catalyzes the third step, namely, the transfer of the acetyl group from acetyl-dihydrolipoamide to CoA to form acetyl-CoA and dihydrolipoamide, the fully reduced form of lipoamide, the dithiol.
It should be noted that the acetyl group, initially bound by ester linkage to one of the –SH group of lipoamide is next bound to the –SH group of coenzyme A, again by ester bond, hence the term transesterification.

Catalytic mechanism of dihydrolipoyl transacetylase or E2

During the reaction, the sulfhydryl group of coenzyme A carries out a nucleophilic attack on the carbonyl carbon of the acetyl group of acetyl dihydrolipoamide-dihydrolipoyl transacetylase to form a transient tetrahedral intermediate, that “decomposes” to dihydrolipoamide-dihydrolipoyl transacetylase and acetyl-CoA.

Pyruvate Dehydrogenase Complex
Fig. 14 – E2: Catalytic Mechanism

As previously said, the mobility of the lipoyllysyl arm plays a central role in the reaction mechanism.

Reaction of dihydrolipoyl dehydrogenase or E3

In the reaction sequence catalyzed by components of the pyruvate dehydrogenase complex, dihydrolipoyl dehydrogenase catalyzes the fourth and fifth steps.
The enzyme catalyzes electron transfers needed to regenerate the disulfide bridge of the lipoyl group of dihydrolipoyl transacetylase, that is, to regenerate the oxidized form of the prosthetic group, and thus completing the catalytic cycle of the transacetylase.
The reaction has a ping-pong catalytic mechanism: it occurs in two successive half-reaction, in which each of the two substrates, NAD+ and dihydrolipoamide, reacts in the absence of the other. Moreover during the first half-reaction, the release of the first product and the formation of an enzyme intermediate complex occur before the second substrate binds, while the enzyme underogoes a structural change, whereas in the second half-reaction the release of the second product and the return of the enzyme to its starting state, again, via a structural change, occur.
Considering the ping-pong kinetic mechanism of dihydrolipoyl dehydrogenase:

  • in the first half-reaction the oxidation of dihydrolipoamide to lipoamide occurs;
  • in the second half-reaction the reduction of NAD+ to NADH occurs.

Catalytic mechanism of dihydrolipoyl dehydrogenase or E3

Below, the reaction mechanism of P. putida dihydrolipoyl dehydrogenase is described.
In the first half-reaction, the oxidized dihydrolipoyl dehydrogenase (E), i.e., the enzyme with the disulfide bridge between Cys43 and Cys48, binds dihydrolipoamide (LH2) to form the enzyme-dihydrolipoamide complex (E●LH2). At this point, a sulfur atom of dihydrolipoamide carries out a nucleophilic attack on the sulfur of Cys43, to form the disulfide bridge lipoamide-Cys43 (E-S-S-L), while the sulfur of Cys48 is released as a thiolate ion (S48).
The proton on the second thiol group of lipoamide is then abstracted by histidine (Hys) 451, that acts as a general acid-base catalyst, leading to the formation of a second thiolate ion, this time on the lipoamide (E-S-S-L ●S), that, through a nucleophilic attack, displaces the sulfur of Cys43, S43, aided in this by general acid catalysis by Hys451 which donates a proton to S43. The catalytic action of Hys451 is essential, as demonstrated by mutagenesis studies in which its substitution with a glutamine residue causes the enzyme to retain ∼ 0.4% of the wild-type catalytic activity.
The thiolate anion S48 then contacts, through non-covalent interactions, the flavin ring near 4a position, i.e., an electron pair of S48, which acts as electron donor, is partially transferred to the oxidized flavin ring, which, in turn, is the electron acceptor. The resulting structure is called charge-transfer complex.
Meanwhile, the phenolic side chain of the Tyr181 continues to hinder access to the flavin ring, thus protecting it from oxidation by O2.

Pyruvate Dehydrogenase Complex
Fig. 15 – Dihydrolipoamide Oxidation via E3

To sum up, what occurs is an interchange reaction of disulfide bridges leading to the formation of the oxidized form of lipoamide, the first product, which is released, and the reduced form of the dihydrolipoyl dehydrogenase.

The second half-reaction involves the reduction of NAD+ to NADH + H+ by electron transfer from the reactive disulfide of the enzyme via FAD.

Pyruvate dehydrogenase complex
Fig. 16 – Oxidation of Reduced E3 by NAD+

It begins with the entry of NAD+ into the active site and its binding to form the EH2●NAD+ complex. It should be noted that the entry of the coenzyme causes the phenolic side chain of the Tyr181 to be pushed aside by the nicotinamide ring.
Following the collapse of the charge-transfer complex, a covalent bond is formed between the flavin atom C-4a and S48, to which the extraction of a proton from S43 by the flavin atom N-5 is accompanied, with the formation of the corresponding thiolate anion, S43.
S43carries out a nucleophilic attack on S48, leading to the formation of the redox-active disulphide bridge between Cys43 and Cys48, followed by the breakdown of the covalent bond between S48 and the flavin atom C-4a to form reduced FADH anion, FADH, with negative charge on atom N-1. It should be noted that dihydrolipoyl dehydrogenase is in the oxidized form (E).

Pyruvate Dehydrogenase Complex
Fig. 17 – FADH-

FADH has a transient existence because the proton bound to its N-5 is instantly transferred, as hydride ion, to the nicotinamide atom C-4, that is juxtaposed to flavin atom N-5. This leads to the formation of FAD and of the second product of the reaction, NADH, which is released.
To sum up, what occurs is that the electrons removed from the hydroxyethyl group, which derives from pyruvate, pass, via FAD, to NAD+. The catalytic cycle of dihydrolipoyl dehydrogenase is therefore completed, being the enzyme and its coenzymes in their oxidized form. At this point, the catalytic cycle of the entire pyruvate dehydrogenase complex is completed, too, and the complex is ready for a new reaction cycle.

Note: unlike the thiazolium ring of thiamine pyrophosphate, FAD does not acts as electron trap or electron sink, but rather as an electron conduit between the redox-active disulphide, in its reduced form, and NAD+.

Note: the catalytic mechanism of dihydrolipoyl dehydrogenase has been determined in analogy with that of glutathione reductase (EC 1.8.1.7), at 33% identical and whose structure is more extensively characterized. It should, however, be noted that although the two enzymes catalyze similar reactions, these usually occur in opposite direction:

  • dihydrolipoyl dehydrogenase uses NAD+ to oxidize two –SH groups to a disulfide (–S–S–);
  • glutathione reductase uses NADPH to reduce a –S–S– to two thiol groups.

Nevertheless, their active sites are closely superimposable.

Regulation of the pyruvate dehydrogenase complex

In mammals, the regulation of the activity of the pyruvate dehydrogenase complex is essential, both in the fed and fasted states. In fact, the multienzyme complex plays a central role in metabolism because, catalyzing the irreversible oxidative decarboxylation of pyruvate, represents the entry point of the carbon flux from all carbohydrate sources as well as from ∼50% of carbon skeletons of glucogenic amino acids, that, as a whole, correspond to ∼60% of the daily calorie intake, into:

  • the citric acid cycle, and therefore to the full oxidation to CO2;
  • the synthesis of lipids (fed state) and acetylcholine (see above).

The importance of the regulation of the conversion of pyruvate into acetyl-CoA is also underlined by the fact that mammals, although able to produce glucose from pyruvate, cannot synthesize it from acetyl-CoA, because of the irreversibility of pyruvate dehydrogenase reaction and the absence of alternative pathways. Then, the inhibition of the activity of the complex allows to spare glucose and the amino acids that can be converted into pyruvate, such as alanine, when other fuels, for example acetyl-CoA from fatty acid oxidation, are available.
This explains why the activity of the complex is carefully regulated by:

  • feedback inhibition;
  • nucleotides;
  • covalent modifications, namely, phosphorylation and dephosphorylation of specific target proteins.

Regulation of the pyruvate dehydrogenase complex by feedback inhibition and energy status of the cell

The activity of the dephosphorylated form of the pyruvate dehydrogenase complex is regulated by feedback inhibition.
Acetyl-CoA and NADH allosterically inhibit the enzymes that catalyze their formation, dihydrolipoyl transacetylase and dihydrolipoyl dehydrogenase, respectively.
In addition, CoA and acetyl-CoA, as well as NAD+ and NADH, compete for binding sites on E2 and E3, respectively, that catalyze reversible reactions. This means that, in the presence of high ratios of [Acetyl-CoA]/[CoA] and [NADH]/ [NAD+], the reactions of transacetylation and dehydrogenation work in reverse; therefore, dihydrolipoyl transacetylase cannot accept the hydroxyethyl group from TPP because it is maintained in the acetylated form. This cause thiamine pyrophosphate to remain bound to pyruvate dehydrogenase in its hydroxyethyl form, which, in turn, decreases the rate of pyruvate decarboxylation. Hence, high ratios of [Acetyl-CoA]/[CoA] and [NADH]/[NAD+] indirectly influence pyruvate dehydrogenase activity.

Pyruvate Dehydrogenase Complex
Fig. 18 – PDC Activiy: Feedback Inhibition

Acetyl-CoA and NADH are also produced by fatty acid oxidation, which takes place, like the reactions of the pyruvate dehydrogenase complex, within the mitochondrion. This means that the cell, by regulating the activity of the multienzyme complex, preserves carbohydrate stores when fatty acids are available for energy. For example, during the fasted state, liver, skeletal muscle and many other organs and tissues rely primarily on fatty acid oxidation for energy. Conversely, the activity of the multienzyme complex is increased in the fed state, when many different types of cells and tissues mainly use glucose as a fuel.
More generally, when the production of NADH and/or acetil-CoA exceeds the capacity of the cell to use them for ATP production, the activity of the pyruvate dehydrogenase complex is inhibited. The same is true when there is no need for additional ATP to be produced. Infact, the activity of multienzyme complex is also sensitive to the energy charge of the cell. Through allosteric mechanisms, high ATP levels inhibit the activity of the pyruvate dehydrogenase component of the complex, whereas high ADP levels, that signs that the energy charge of the cell may become low, activate it, thus committing the carbon skeleton of carbohydrates and some amino acids to energy production.

Note: in the skeletal muscle, the activity of the pyruvate dehydrogenase complex increases with increased aerobic activity, resulting in a in greater dependence on glucose as a fuel source.

Regulation of the pyruvate dehydrogenase complex by phosphorylation/dephosphorylation

Unlike prokaryotes, in mammals the activity of the pyruvate dehydrogenase complex is also regulated by covalent modifications, i.e., phosphorylation and dephosphorylation of three specific serine residues of the α subunit of pyruvate dehydrogenase, the enzyme that catalyzes the first, irreversible step of the overall reaction sequence.
Note: as mammalian pyruvate dehydrogenase is an heterotetramer, there are six potential phosphorylation sites.

Pyruvate Dehydrogenase Complex
Fig. 19 – PDC Activiy: Covalent Modifications

Phosphorylation, which inactivates pyruvate dehydrogenase, and then blocks the overall reaction sequence, is catalyzed by pyruvate dehydrogenase kinase. Two of the aforementioned serine residues are located on the more C-terminal loop, at the entrance of the substrate channel leading to the respective active site, and the phosphorylation of only one of them inactivates the pyruvate dehydrogenase, hence demonstrating the out of phase coupling between its active sites.
Conversely, in the dephosphorylated state the complex is active. Dephosphorylation is catalyzed by a specific protein phosphatase, the pyruvate dehydrogenase phosphatase.
The activities of pyruvate dehydrogenase kinase and pyruvate dehydrogenase phosphatase are in turn subject to allosteric regulation by several modulators.

Regulation of pyruvate dehydrogenase kinase

The activity of pyruvate dehydrogenase kinase depends on the ratios of [NADH]/[NAD+], [acetyl-CoA]/[CoA], and [ATP]/[ADP] , as well as on the pyruvate concentration, in the mitochondrial matrix (see fig. 19).

  • High ratios of [NADH]/[NAD+] and [acetyl-CoA]/[CoA], as during the oxidation of fatty acids and ketone bodies, activate the kinase, pyruvate dehydrogenase is phosphorylated, and the pyruvate dehydrogenase complex is inhibited. This allows tissues, such as cardiac muscle, to preserve glucose when fatty acids and/or ketone bodies are utilized for energy, because acetyl-CoA synthesis from pyruvate, and hence from carbohydrates (and some amino acids) is turned off.
    Conversely, when the concentrations of NAD+ and coenzyme A are high the activity of the kinase is inhibited and the multienzyme complex is active.
    Therefore, acetyl-CoA and NADH, two of the three end products of the reactions catalyzed by the pyruvate dehydrogenase complex, allosterically control their synthesis by regulating directly and indirectly, by regulating the activity of pyruvate dehydrogenase kinase, the activity of the complex.
  • A high ratio of [ATP]/[ADP] activates the kinase, and then inhibits the pyruvate dehydrogenase complex.
    Note: unlike many other kinases, such as those involved in the control of glycogen metabolism, pyruvate dehydrogenase kinase is not regulated by cAMP levels, but by molecules that signal changes in energy status of the cell and in the availability of biosynthetic intermediates: ATP and NADH, and acetyl-CoA, respectively.
  • Pyruvate allosterically inhibits pyruvate dehydrogenase kinase.
    When its levels are high, it binds to kinase and inactivates it, pyruvate dehydrogenase is not phosphorylated, and the pyruvate dehydrogenase complex remains active.
  • Pyruvate dehydrogenase kinase is also activated by interaction with dihydrolipoyl transacetylase in its acetylated form, i.e. when acetyl-dihydrolipoamide is present.

Other activators of the kinase is potassium and magnesium ions.

Regulation of pyruvate dehydrogenase phosphatase

The activity of pyruvate dehydrogenase phosphatase depends on the ratios of [NADH]/[NAD+] and [acetyl-CoA]/[CoA], as well as on [Ca2+], in the mitochondrial matrix (see fig. 19).

  • Low ratios of [NADH]/[NAD+] and [acetyl-CoA]/[CoA] activate the phosphatase, pyruvate dehydrogenase is dephosphorylated, and the pyruvate dehydrogenase complex is activated.
    Conversely, when the aforesaid ratios are high, phosphatase activity is reduced, kinase activity is increased, and the multienzyme complex is inhibited.
  • Calcium ion activates pyruvate dehydrogenase phosphatase.
    Ca2+ is an important second messenger that signals the cell requires more energy. Therefore, when its levels are high, as in cardiac muscle cells after epinephrine stimulation or in skeletal muscle cells during the muscular contraction, the phosphatase is active, the complex is dephosphorylated, and then active.
  • Insulin, too, is involved in the control of the activity of the pyruvate dehydrogenase complex through the activation of pyruvate dehydrogenase phosphatase. The hormone, in response to increases in blood glucose, stimulates glycogen synthesis and the synthesis of acetyl-CoA, a precursor in the synthesis of lipids.

Fasting and subsequent refeeding, too, affect the activity of the multienzyme complex.
In tissues such as skeletal muscle, cardiac muscle or kidney, fasting significantly decreases the activity of the complex, whereas refeeding reverses the inhibition of fasting.
In the brain, however, these variations are not observed because the activity of pyruvate dehydrogenase complex is essential for ATP production.

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