Enzyme assay

From Wikipedia, the free encyclopedia

https://en.wikipedia.org/wiki/Enzyme_assay 

 

Beckman DU640 UV/Vis spectrophotometer

Enzyme assays are laboratory methods for measuring enzymatic activity. They are vital for the study of enzyme kinetics and enzyme inhibition.

Enzyme units

The quantity or concentration of an enzyme can be expressed in molar amounts, as with any other chemical, or in terms of activity in enzyme units.

Enzyme activity

Enzyme activity is a measure of the quantity of active enzyme present and is thus dependent on various physical conditions, which should be specified.

It is calculated using the following formula:

${\displaystyle \mathrm{a}=\mathrm{n}=\mathrm{r}\times \mathrm{V}}$

where

${\displaystyle \mathrm{a}=}$ Enzyme activity

${\displaystyle \mathrm{n}=}$ Moles of substrate converted per unit time

${\displaystyle \mathrm{r}=}$ Rate of the reaction

${\displaystyle \mathrm{V}=}$ Reaction volume

The SI unit is the katal, 1 katal = 1 mol s⁻¹ (mole per second), but this is an excessively large unit. A more practical and commonly used value is enzyme unit (U) = 1 μmol min⁻¹ (micromole per minute). 1 U corresponds to 16.67 nanokatals.

Enzyme activity as given in katal generally refers to that of the assumed natural target substrate of the enzyme. Enzyme activity can also be given as that of certain standardized substrates, such as gelatin, then measured in gelatin digesting units (GDU), or milk proteins, then measured in milk clotting units (MCU). The units GDU and MCU are based on how fast one gram of the enzyme will digest gelatin or milk proteins, respectively. 1 GDU approximately equals 1.5 MCU.

An increased amount of substrate will increase the rate of reaction with enzymes, however once past a certain point, the rate of reaction will level out because the amount of active sites available has stayed constant.

Specific activity

The specific activity of an enzyme is another common unit. This is the activity of an enzyme per milligram of total protein (expressed in μmol min⁻¹ mg⁻¹). Specific activity gives a measurement of enzyme purity in the mixture. It is the micro moles of product formed by an enzyme in a given amount of time (minutes) under given conditions per milligram of total proteins. Specific activity is equal to the rate of reaction multiplied by the volume of reaction divided by the mass of total protein. The SI unit is katal/kg, but a more practical unit is μmol/mgmin.

Specific activity is a measure of enzyme processivity (the capability of enzyme to be processed), at a specific (usually saturating) substrate concentration, and is usually constant for a pure enzyme.

An active site titration process can be done for the elimination of errors arising from differences in cultivation batches and/or misfolded enzyme and similar issues. This is a measure of the amount of active enzyme, calculated by e.g. titrating the amount of active sites present by employing an irreversible inhibitor. The specific activity should then be expressed as μmol min⁻¹ mg⁻¹ active enzyme. If the molecular weight of the enzyme is known, the turnover number, or μmol product per second per μmol of active enzyme, can be calculated from the specific activity. The turnover number can be visualized as the number of times each enzyme molecule carries out its catalytic cycle per second.

Related terminology

The rate of a reaction is the concentration of substrate disappearing (or product produced) per unit time (mol L⁻¹ s⁻¹).

The % purity is 100% × (specific activity of enzyme sample / specific activity of pure enzyme). The impure sample has lower specific activity because some of the mass is not actually enzyme. If the specific activity of 100% pure enzyme is known, then an impure sample will have a lower specific activity, allowing purity to be calculated and then getting a clear result.

Types of assays

All enzyme assays measure either the consumption of substrate or production of product over time. A large number of different methods of measuring the concentrations of substrates and products exist and many enzymes can be assayed in several different ways. Biochemists usually study enzyme-catalysed reactions using four types of experiments:

• Initial rate experiments. When an enzyme is mixed with a large excess of the substrate, the enzyme-substrate intermediate builds up in a fast initial transient. Then the reaction achieves a steady-state kinetics in which enzyme substrate intermediates remains approximately constant over time and the reaction rate changes relatively slowly. Rates are measured for a short period after the attainment of the quasi-steady state, typically by monitoring the accumulation of product with time. Because the measurements are carried out for a very short period and because of the large excess of substrate, the approximation that the amount of free substrate is approximately equal to the amount of the initial substrate can be made. The initial rate experiment is the simplest to perform and analyze, being relatively free from complications such as back-reaction and enzyme degradation. It is therefore by far the most commonly used type of experiment in enzyme kinetics.
• Progress curve experiments. In these experiments, the kinetic parameters are determined from expressions for the species concentrations as a function of time. The concentration of the substrate or product is recorded in time after the initial fast transient and for a sufficiently long period to allow the reaction to approach equilibrium. Progress curve experiments were widely used in the early period of enzyme kinetics, but are less common now.
• Transient kinetics experiments. In these experiments, reaction behaviour is tracked during the initial fast transient as the intermediate reaches the steady-state kinetics period. These experiments are more difficult to perform than either of the above two classes because they require specialist techniques (such as flash photolysis of caged compounds) or rapid mixing (such as stopped-flow, quenched flow or continuous flow).
• Relaxation experiments. In these experiments, an equilibrium mixture of enzyme, substrate and product is perturbed, for instance by a temperature, pressure or pH jump, and the return to equilibrium is monitored. The analysis of these experiments requires consideration of the fully reversible reaction. Moreover, relaxation experiments are relatively insensitive to mechanistic details and are thus not typically used for mechanism identification, although they can be under appropriate conditions.

Enzyme assays can be split into two groups according to their sampling method: continuous assays, where the assay gives a continuous reading of activity, and discontinuous assays, where samples are taken, the reaction stopped and then the concentration of substrates/products determined.

Temperature-controlled cuvette holder in a spectrophotometer.

Continuous assays

Continuous assays are most convenient, with one assay giving the rate of reaction with no further work necessary. There are many different types of continuous assays.

Spectrophotometric

In spectrophotometric assays, you follow the course of the reaction by measuring a change in how much light the assay solution absorbs. If this light is in the visible region you can actually see a change in the color of the assay, and these are called colorimetric assays. The MTT assay, a redox assay using a tetrazolium dye as substrate is an example of a colorimetric assay.

UV light is often used, since the common coenzymes NADH and NADPH absorb UV light in their reduced forms, but do not in their oxidized forms. An oxidoreductase using NADH as a substrate could therefore be assayed by following the decrease in UV absorbance at a wavelength of 340 nm as it consumes the coenzyme.

Direct versus coupled assays

Coupled assay for hexokinase using glucose-6-phosphate dehydrogenase.

Even when the enzyme reaction does not result in a change in the absorbance of light, it can still be possible to use a spectrophotometric assay for the enzyme by using a coupled assay. Here, the product of one reaction is used as the substrate of another, easily detectable reaction. For example, figure 1 shows the coupled assay for the enzyme hexokinase, which can be assayed by coupling its production of glucose-6-phosphate to NADPH production, using glucose-6-phosphate dehydrogenase.

Fluorometric

Fluorescence is when a molecule emits light of one wavelength after absorbing light of a different wavelength. Fluorometric assays use a difference in the fluorescence of substrate from product to measure the enzyme reaction. These assays are in general much more sensitive than spectrophotometric assays, but can suffer from interference caused by impurities and the instability of many fluorescent compounds when exposed to light.

An example of these assays is again the use of the nucleotide coenzymes NADH and NADPH. Here, the reduced forms are fluorescent and the oxidised forms non-fluorescent. Oxidation reactions can therefore be followed by a decrease in fluorescence and reduction reactions by an increase. Synthetic substrates that release a fluorescent dye in an enzyme-catalyzed reaction are also available, such as 4-methylumbelliferyl-β-D-galactoside for assaying β-galactosidase or 4-methylumbelliferyl-butyrate for assaying Candida rugosa lipase.

Calorimetric

Chemiluminescence of luminol

Calorimetry is the measurement of the heat released or absorbed by chemical reactions. These assays are very general, since many reactions involve some change in heat and with use of a microcalorimeter, not much enzyme or substrate is required. These assays can be used to measure reactions that are impossible to assay in any other way.

Chemiluminescent

Chemiluminescence is the emission of light by a chemical reaction. Some enzyme reactions produce light and this can be measured to detect product formation. These types of assay can be extremely sensitive, since the light produced can be captured by photographic film over days or weeks, but can be hard to quantify, because not all the light released by a reaction will be detected.

The detection of horseradish peroxidase by enzymatic chemiluminescence (ECL) is a common method of detecting antibodies in western blotting. Another example is the enzyme luciferase, this is found in fireflies and naturally produces light from its substrate luciferin.

Light scattering

Static light scattering measures the product of weight-averaged molar mass and concentration of macromolecules in solution. Given a fixed total concentration of one or more species over the measurement time, the scattering signal is a direct measure of the weight-averaged molar mass of the solution, which will vary as complexes form or dissociate. Hence the measurement quantifies the stoichiometry of the complexes as well as kinetics. Light scattering assays of protein kinetics is a very general technique that does not require an enzyme.

Microscale thermophoresis

Microscale thermophoresis (MST) measures the size, charge and hydration entropy of molecules/substrates at equilibrium. The thermophoretic movement of a fluorescently labeled substrate changes significantly as it is modified by an enzyme. This enzymatic activity can be measured with high time resolution in real time. The material consumption of the all optical MST method is very low, only 5 μl sample volume and 10nM enzyme concentration are needed to measure the enzymatic rate constants for activity and inhibition. MST allows analysts to measure the modification of two different substrates at once (multiplexing) if both substrates are labeled with different fluorophores. Thus substrate competition experiments can be performed.

Discontinuous assays

Discontinuous assays are when samples are taken from an enzyme reaction at intervals and the amount of product production or substrate consumption is measured in these samples.

Radiometric

Radiometric assays measure the incorporation of radioactivity into substrates or its release from substrates. The radioactive isotopes most frequently used in these assays are ¹⁴C, ³²P, ³⁵S and ¹²⁵I. Since radioactive isotopes can allow the specific labelling of a single atom of a substrate, these assays are both extremely sensitive and specific. They are frequently used in biochemistry and are often the only way of measuring a specific reaction in crude extracts (the complex mixtures of enzymes produced when you lyse cells).

Radioactivity is usually measured in these procedures using a scintillation counter.

Chromatographic

Chromatographic assays measure product formation by separating the reaction mixture into its components by chromatography. This is usually done by high-performance liquid chromatography (HPLC), but can also use the simpler technique of thin layer chromatography. Although this approach can need a lot of material, its sensitivity can be increased by labelling the substrates/products with a radioactive or fluorescent tag. Assay sensitivity has also been increased by switching protocols to improved chromatographic instruments (e.g. ultra-high pressure liquid chromatography) that operate at pump pressure a few-fold higher than HPLC instruments (see High-performance liquid chromatography#Pump pressure).

Factors affecting assays

A pressure chamber for measuring enzyme activity at high pressure.

Several factors effect the assay outcome and a recent review summarizes the various parameters that needs to be monitored to keep an assay up and running.

Salt Concentration

Most enzymes cannot tolerate extremely high salt concentrations. The ions interfere with the weak ionic bonds of proteins. Typical enzymes are active in salt concentrations of 1-500 mM. As usual there are exceptions such as the halophilic algae and bacteria.

Effects of Temperature

All enzymes work within a range of temperature specific to the organism. Increases in temperature generally lead to increases in reaction rates. There is a limit to the increase because higher temperatures lead to a sharp decrease in reaction rates. This is due to the denaturating (alteration) of protein structure resulting from the breakdown of the weak ionic and hydrogen bonding that stabilize the three-dimensional structure of the enzyme active site. The "optimum" temperature for human enzymes is usually between 35 and 40 °C. The average temperature for humans is 37 °C. Human enzymes start to denature quickly at temperatures above 40 °C. Enzymes from thermophilic archaea found in the hot springs are stable up to 100 °C. However, the idea of an "optimum" rate of an enzyme reaction is misleading, as the rate observed at any temperature is the product of two rates, the reaction rate and the denaturation rate. If you were to use an assay measuring activity for one second, it would give high activity at high temperatures, however if you were to use an assay measuring product formation over an hour, it would give you low activity at these temperatures.

Effects of pH

Most enzymes are sensitive to pH and have specific ranges of activity. All have an optimum pH. The pH can stop enzyme activity by denaturating (altering) the three-dimensional shape of the enzyme by breaking ionic, and hydrogen bonds. Most enzymes function between a pH of 6 and 8; however pepsin in the stomach works best at a pH of 2 and trypsin at a pH of 8.

Substrate Saturation

Increasing the substrate concentration increases the rate of reaction (enzyme activity). However, enzyme saturation limits reaction rates. An enzyme is saturated when the active sites of all the molecules are occupied most of the time. At the saturation point, the reaction will not speed up, no matter how much additional substrate is added. The graph of the reaction rate will plateau.

Level of crowding

Large amounts of macromolecules in a solution will alter the rates and equilibrium constants of enzyme reactions, through an effect called macromolecular crowding.

List of enzyme assays

• MTT assay
• Fluorescein diacetate hydrolysis
• para-Nitrophenylphosphate

Nicotinamide adenine dinucleotide

From Wikipedia, the free encyclopedia

https://en.wikipedia.org/wiki/Nicotinamide_adenine_dinucleotide 

Nicotinamide adenine dinucleotide

Names
Other names Diphosphopyridine nucleotide (DPN+), Coenzyme I
Identifiers
CAS Number	53-84-9  53-59-8 (phosphate)  58-68-4 (NADH)
3D model (JSmol)	NAD+: Interactive image NADH: Interactive image
ChEBI	CHEBI:16908
ChEMBL	ChEMBL1628272
ChemSpider	5681
DrugBank	DB00157
ECHA InfoCard	100.000.169
IUPHAR/BPS	2451
KEGG	C00003
PubChem CID	925
RTECS number	UU3450000
UNII	0U46U6E8UK  BY8P107XEP (phosphate)  4J24DQ0916 (NADH)

InChI

SMILES
Properties
Chemical formula	C21H28N7O14P2
Molar mass	663.43 g/mol
Appearance	White powder
Melting point	160 °C (320 °F; 433 K)
Hazards
Occupational safety and health (OHS/OSH):
Main hazards	Not hazardous
NFPA 704 (fire diamond)	1 1 0
Except where otherwise noted, data are given for materials in their standard state (at 25 °C [77 °F], 100 kPa).

Nicotinamide adenine dinucleotide (NAD) is a coenzyme central to metabolism. Found in all living cells, NAD is called a dinucleotide because it consists of two nucleotides joined through their phosphate groups. One nucleotide contains an adenine nucleobase and the other nicotinamide. NAD exists in two forms: an oxidized and reduced form, abbreviated as NAD⁺ and NADH (H for hydrogen), respectively.

In metabolism, nicotinamide adenine dinucleotide is involved in redox reactions, carrying electrons from one reaction to another. The cofactor is, therefore, found in two forms in cells: NAD⁺ is an oxidizing agent – it accepts electrons from other molecules and becomes reduced. This reaction, also with H⁺, forms NADH, which can then be used as a reducing agent to donate electrons. These electron transfer reactions are the main function of NAD. However, it is also used in other cellular processes, most notably as a substrate of enzymes in adding or removing chemical groups to or from, respectively, proteins, in posttranslational modifications. Because of the importance of these functions, the enzymes involved in NAD metabolism are targets for drug discovery.

In organisms, NAD can be synthesized from simple building-blocks (de novo) from either tryptophan or aspartic acid, each a case of an amino acid; alternatively, more complex components of the coenzymes are taken up from nutritive compounds such as niacin; similar compounds are produced by reactions that break down the structure of NAD, providing a salvage pathway that “recycles” them back into their respective active form.

Some NAD is converted into the coenzyme nicotinamide adenine dinucleotide phosphate (NADP); its chemistry largely parallels that of NAD, though predominantly its role is as a cofactor in anabolic metabolism.

The NAD⁺ chemical species’ superscripted addition sign reflects the formal charge on one of its nitrogen atoms; this species is actually a singly charged anion – carrying a (negative) ionic charge of 1 – under conditions of physiological pH.

Physical and chemical properties

Further information: Redox

Nicotinamide adenine dinucleotide consists of two nucleosides joined by pyrophosphate. The nucleosides each contain a ribose ring, one with adenine attached to the first carbon atom (the 1' position) (adenosine diphosphate ribose) and the other with nicotinamide at this position.^[3][4]

The redox reactions of nicotinamide adenine dinucleotide.

The compound accepts or donates the equivalent of H⁻. Such reactions (summarized in formula below) involve the removal of two hydrogen atoms from the reactant (R), in the form of a hydride ion (H⁻), and a proton (H⁺). The proton is released into solution, while the reductant RH₂ is oxidized and NAD⁺ reduced to NADH by transfer of the hydride to the nicotinamide ring.

RH₂ + NAD⁺ → NADH + H⁺ + R;

From the hydride electron pair, one electron is transferred to the positively charged nitrogen atom of the nicotinamide ring of NAD⁺, and the second hydrogen atom transferred to the C4 carbon atom opposite this N atom. The midpoint potential of the NAD⁺/NADH redox pair is −0.32 volts, which makes NADH a moderately strong reducing agent. The reaction is easily reversible, when NADH reduces another molecule and is re-oxidized to NAD⁺. This means the coenzyme can continuously cycle between the NAD⁺ and NADH forms without being consumed.

In appearance, all forms of this coenzyme are white amorphous powders that are hygroscopic and highly water-soluble. The solids are stable if stored dry and in the dark. Solutions of NAD⁺ are colorless and stable for about a week at 4 °C and neutral pH, but decompose rapidly in acidic or alkaline solutions. Upon decomposition, they form products that are enzyme inhibitors.

UV absorption spectra of NAD⁺ and NADH.

Both NAD⁺ and NADH strongly absorb ultraviolet light because of the adenine. For example, peak absorption of NAD⁺ is at a wavelength of 259 nanometers (nm), with an extinction coefficient of 16,900 M⁻¹cm⁻¹. NADH also absorbs at higher wavelengths, with a second peak in UV absorption at 339 nm with an extinction coefficient of 6,220 M⁻¹cm⁻¹. This difference in the ultraviolet absorption spectra between the oxidized and reduced forms of the coenzymes at higher wavelengths makes it simple to measure the conversion of one to another in enzyme assays – by measuring the amount of UV absorption at 340 nm using a spectrophotometer.

NAD⁺ and NADH also differ in their fluorescence. Freely diffusing NADH in aqueous solution, when excited at the nicotinamide absorbance of ~335 nm (near UV), fluoresces at 445–460 nm (violet to blue) with a fluorescence lifetime of 0.4 nanoseconds, while NAD⁺ does not fluoresce. The properties of the fluorescence signal changes when NADH binds to proteins, so these changes can be used to measure dissociation constants, which are useful in the study of enzyme kinetics. These changes in fluorescence are also used to measure changes in the redox state of living cells, through fluorescence microscopy.

Concentration and state in cells

In rat liver, the total amount of NAD⁺ and NADH is approximately 1 μmole per gram of wet weight, about 10 times the concentration of NADP⁺ and NADPH in the same cells. The actual concentration of NAD⁺ in cell cytosol is harder to measure, with recent estimates in animal cells ranging around 0.3 mM, and approximately 1.0 to 2.0 mM in yeast. However, more than 80% of NADH fluorescence in mitochondria is from bound form, so the concentration in solution is much lower.

NAD⁺ concentrations are highest in the mitochondria, constituting 40% to 70% of the total cellular NAD⁺. NAD⁺ in the cytosol is carried into the mitochondrion by a specific membrane transport protein, since the coenzyme cannot diffuse across membranes. The intracellular half-life of NAD⁺ was claimed to be between 1–2 hours by one review, whereas another review gave varying estimates based on compartment: intracellular 1–4 hours, cytoplasmic 2 hours, and mitochondrial 4–6 hours.

The balance between the oxidized and reduced forms of nicotinamide adenine dinucleotide is called the NAD⁺/NADH ratio. This ratio is an important component of what is called the redox state of a cell, a measurement that reflects both the metabolic activities and the health of cells. The effects of the NAD⁺/NADH ratio are complex, controlling the activity of several key enzymes, including glyceraldehyde 3-phosphate dehydrogenase and pyruvate dehydrogenase. In healthy mammalian tissues, estimates of the ratio of free NAD⁺ to NADH in the cytoplasm typically lie around 700:1; the ratio is thus favorable for oxidative reactions. The ratio of total NAD⁺/NADH is much lower, with estimates ranging from 3–10 in mammals. In contrast, the NADP⁺/NADPH ratio is normally about 0.005, so NADPH is the dominant form of this coenzyme. These different ratios are key to the different metabolic roles of NADH and NADPH.

Biosynthesis

NAD⁺ is synthesized through two metabolic pathways. It is produced either in a de novo pathway from amino acids or in salvage pathways by recycling preformed components such as nicotinamide back to NAD⁺. Although most tissues synthesize NAD⁺ by the salvage pathway in mammals, much more de novo synthesis occurs in the liver from tryptophan, and in the kidney and macrophages from nicotinic acid.

De novo production

Some metabolic pathways that synthesize and consume NAD⁺ in vertebrates.The abbreviations are defined in the text.

Most organisms synthesize NAD⁺ from simple components. The specific set of reactions differs among organisms, but a common feature is the generation of quinolinic acid (QA) from an amino acid – either tryptophan (Trp) in animals and some bacteria, or aspartic acid (Asp) in some bacteria and plants. The quinolinic acid is converted to nicotinic acid mononucleotide (NaMN) by transfer of a phosphoribose moiety. An adenylate moiety is then transferred to form nicotinic acid adenine dinucleotide (NaAD). Finally, the nicotinic acid moiety in NaAD is amidated to a nicotinamide (Nam) moiety, forming nicotinamide adenine dinucleotide.

In a further step, some NAD⁺ is converted into NADP⁺ by NAD⁺ kinase, which phosphorylates NAD⁺. In most organisms, this enzyme uses ATP as the source of the phosphate group, although several bacteria such as Mycobacterium tuberculosis and a hyperthermophilic archaeon Pyrococcus horikoshii, use inorganic polyphosphate as an alternative phosphoryl donor.

Salvage pathways use three precursors for NAD⁺.

Salvage pathways

Despite the presence of the de novo pathway, the salvage reactions are essential in humans; a lack of niacin in the diet causes the vitamin deficiency disease pellagra. This high requirement for NAD⁺ results from the constant consumption of the coenzyme in reactions such as posttranslational modifications, since the cycling of NAD⁺ between oxidized and reduced forms in redox reactions does not change the overall levels of the coenzyme. The major source of NAD⁺ in mammals is the salvage pathway which recycles the nicotinamide produced by enzymes utilizing NAD⁺. The first step, and the rate-limiting enzyme in the salvage pathway is nicotinamide phosphoribosyltransferase (NAMPT), which produces nicotinamide mononucleotide (NMN). NMN is the immediate precursor to NAD+ in the salvage pathway.

Besides assembling NAD⁺ de novo from simple amino acid precursors, cells also salvage preformed compounds containing a pyridine base. The three vitamin precursors used in these salvage metabolic pathways are nicotinic acid (NA), nicotinamide (Nam) and nicotinamide riboside (NR). These compounds can be taken up from the diet and are termed vitamin B₃ or niacin. However, these compounds are also produced within cells and by digestion of cellular NAD⁺. Some of the enzymes involved in these salvage pathways appear to be concentrated in the cell nucleus, which may compensate for the high level of reactions that consume NAD⁺ in this organelle. There are some reports that mammalian cells can take up extracellular NAD⁺ from their surroundings, and both nicotinamide and nicotinamide riboside can be absorbed from the gut.

The salvage pathways used in microorganisms differ from those of mammals. Some pathogens, such as the yeast Candida glabrata and the bacterium Haemophilus influenzae are NAD⁺ auxotrophs – they cannot synthesize NAD⁺ – but possess salvage pathways and thus are dependent on external sources of NAD⁺ or its precursors. Even more surprising is the intracellular pathogen Chlamydia trachomatis, which lacks recognizable candidates for any genes involved in the biosynthesis or salvage of both NAD⁺ and NADP⁺, and must acquire these coenzymes from its host.

Functions

Rossmann fold in part of the lactate dehydrogenase of Cryptosporidium parvum, showing NAD⁺ in red, beta sheets in yellow, and alpha helices in purple.

Nicotinamide adenine dinucleotide has several essential roles in metabolism. It acts as a coenzyme in redox reactions, as a donor of ADP-ribose moieties in ADP-ribosylation reactions, as a precursor of the second messenger molecule cyclic ADP-ribose, as well as acting as a substrate for bacterial DNA ligases and a group of enzymes called sirtuins that use NAD⁺ to remove acetyl groups from proteins. In addition to these metabolic functions, NAD⁺ emerges as an adenine nucleotide that can be released from cells spontaneously and by regulated mechanisms, and can therefore have important extracellular roles.

Oxidoreductase binding of NAD

Further information: Protein structure and Oxidoreductase

The main role of NAD⁺ in metabolism is the transfer of electrons from one molecule to another. Reactions of this type are catalyzed by a large group of enzymes called oxidoreductases. The correct names for these enzymes contain the names of both their substrates: for example NADH-ubiquinone oxidoreductase catalyzes the oxidation of NADH by coenzyme Q. However, these enzymes are also referred to as dehydrogenases or reductases, with NADH-ubiquinone oxidoreductase commonly being called NADH dehydrogenase or sometimes coenzyme Q reductase.

There are many different superfamilies of enzymes that bind NAD⁺ / NADH. One of the most common superfamilies includes a structural motif known as the Rossmann fold. The motif is named after Michael Rossmann, who was the first scientist to notice how common this structure is within nucleotide-binding proteins.

An example of a NAD-binding bacterial enzyme involved in amino acid metabolism that does not have the Rossmann fold is found in Pseudomonas syringae pv. tomato (PDB: 2CWH​; InterPro: IPR003767).

In this diagram, the hydride acceptor C4 carbon is shown at the top. When the nicotinamide ring lies in the plane of the page with the carboxy-amide to the right, as shown, the hydride donor lies either "above" or "below" the plane of the page. If "above" hydride transfer is class A, if "below" hydride transfer is class B.

When bound in the active site of an oxidoreductase, the nicotinamide ring of the coenzyme is positioned so that it can accept a hydride from the other substrate. Depending on the enzyme, the hydride donor is positioned either "above" or "below" the plane of the planar C4 carbon, as defined in the figure. Class A oxidoreductases transfer the atom from above; class B enzymes transfer it from below. Since the C4 carbon that accepts the hydrogen is prochiral, this can be exploited in enzyme kinetics to give information about the enzyme's mechanism. This is done by mixing an enzyme with a substrate that has deuterium atoms substituted for the hydrogens, so the enzyme will reduce NAD⁺ by transferring deuterium rather than hydrogen. In this case, an enzyme can produce one of two stereoisomers of NADH.

Despite the similarity in how proteins bind the two coenzymes, enzymes almost always show a high level of specificity for either NAD⁺ or NADP⁺. This specificity reflects the distinct metabolic roles of the respective coenzymes, and is the result of distinct sets of amino acid residues in the two types of coenzyme-binding pocket. For instance, in the active site of NADP-dependent enzymes, an ionic bond is formed between a basic amino acid side-chain and the acidic phosphate group of NADP⁺. On the converse, in NAD-dependent enzymes the charge in this pocket is reversed, preventing NADP⁺ from binding. However, there are a few exceptions to this general rule, and enzymes such as aldose reductase, glucose-6-phosphate dehydrogenase, and methylenetetrahydrofolate reductase can use both coenzymes in some species.

Role in redox metabolism

A simplified outline of redox metabolism, showing how NAD⁺ and NADH link the citric acid cycle and oxidative phosphorylation.

 

Further information: Cellular respiration and Oxidative phosphorylation

The redox reactions catalyzed by oxidoreductases are vital in all parts of metabolism, but one particularly important area where these reactions occur is in the release of energy from nutrients. Here, reduced compounds such as glucose and fatty acids are oxidized, thereby releasing energy. This energy is transferred to NAD⁺ by reduction to NADH, as part of beta oxidation, glycolysis, and the citric acid cycle. In eukaryotes the electrons carried by the NADH that is produced in the cytoplasm are transferred into the mitochondrion (to reduce mitochondrial NAD⁺) by mitochondrial shuttles, such as the malate-aspartate shuttle. The mitochondrial NADH is then oxidized in turn by the electron transport chain, which pumps protons across a membrane and generates ATP through oxidative phosphorylation. These shuttle systems also have the same transport function in chloroplasts.

Since both the oxidized and reduced forms of nicotinamide adenine dinucleotide are used in these linked sets of reactions, the cell maintains significant concentrations of both NAD⁺ and NADH, with the high NAD⁺/NADH ratio allowing this coenzyme to act as both an oxidizing and a reducing agent. In contrast, the main function of NADPH is as a reducing agent in anabolism, with this coenzyme being involved in pathways such as fatty acid synthesis and photosynthesis. Since NADPH is needed to drive redox reactions as a strong reducing agent, the NADP⁺/NADPH ratio is kept very low.

Although it is important in catabolism, NADH is also used in anabolic reactions, such as gluconeogenesis. This need for NADH in anabolism poses a problem for prokaryotes growing on nutrients that release only a small amount of energy. For example, nitrifying bacteria such as Nitrobacter oxidize nitrite to nitrate, which releases sufficient energy to pump protons and generate ATP, but not enough to produce NADH directly. As NADH is still needed for anabolic reactions, these bacteria use a nitrite oxidoreductase to produce enough proton-motive force to run part of the electron transport chain in reverse, generating NADH.

Non-redox roles

The coenzyme NAD⁺ is also consumed in ADP-ribose transfer reactions. For example, enzymes called ADP-ribosyltransferases add the ADP-ribose moiety of this molecule to proteins, in a posttranslational modification called ADP-ribosylation. ADP-ribosylation involves either the addition of a single ADP-ribose moiety, in mono-ADP-ribosylation, or the transferral of ADP-ribose to proteins in long branched chains, which is called poly(ADP-ribosyl)ation. Mono-ADP-ribosylation was first identified as the mechanism of a group of bacterial toxins, notably cholera toxin, but it is also involved in normal cell signaling. Poly(ADP-ribosyl)ation is carried out by the poly(ADP-ribose) polymerases. The poly(ADP-ribose) structure is involved in the regulation of several cellular events and is most important in the cell nucleus, in processes such as DNA repair and telomere maintenance. In addition to these functions within the cell, a group of extracellular ADP-ribosyltransferases has recently been discovered, but their functions remain obscure. NAD⁺ may also be added onto cellular RNA as a 5'-terminal modification.

The structure of cyclic ADP-ribose.

Another function of this coenzyme in cell signaling is as a precursor of cyclic ADP-ribose, which is produced from NAD⁺ by ADP-ribosyl cyclases, as part of a second messenger system. This molecule acts in calcium signaling by releasing calcium from intracellular stores. It does this by binding to and opening a class of calcium channels called ryanodine receptors, which are located in the membranes of organelles, such as the endoplasmic reticulum, and inducing the activation of the transcription factor NAFC3

NAD⁺ is also consumed by different NAD+-consuming enzymes, such as CD38, CD157, PARPs and the NAD-dependent deacetylases (sirtuins,such as Sir2.). These enzymes act by transferring an acetyl group from their substrate protein to the ADP-ribose moiety of NAD⁺; this cleaves the coenzyme and releases nicotinamide and O-acetyl-ADP-ribose. The sirtuins mainly seem to be involved in regulating transcription through deacetylating histones and altering nucleosome structure. However, non-histone proteins can be deacetylated by sirtuins as well. These activities of sirtuins are particularly interesting because of their importance in the regulation of aging.

Other NAD-dependent enzymes include bacterial DNA ligases, which join two DNA ends by using NAD⁺ as a substrate to donate an adenosine monophosphate (AMP) moiety to the 5' phosphate of one DNA end. This intermediate is then attacked by the 3' hydroxyl group of the other DNA end, forming a new phosphodiester bond. This contrasts with eukaryotic DNA ligases, which use ATP to form the DNA-AMP intermediate.

Li et al. have found that NAD⁺ directly regulates protein-protein interactions. They also show that one of the causes of age-related decline in DNA repair may be increased binding of the protein DBC1 (Deleted in Breast Cancer 1) to PARP1 (poly[ADP–ribose] polymerase 1) as NAD⁺ levels decline during aging. Thus, the modulation of NAD⁺ may protect against cancer, radiation, and aging.

Extracellular actions of NAD⁺

In recent years, NAD⁺ has also been recognized as an extracellular signaling molecule involved in cell-to-cell communication. NAD⁺ is released from neurons in blood vessels, urinary bladder, large intestine, from neurosecretory cells, and from brain synaptosomes, and is proposed to be a novel neurotransmitter that transmits information from nerves to effector cells in smooth muscle organs. In plants, the extracellular nicotinamide adenine dinucleotide induces resistance to pathogen infection and the first extracellular NAD receptor has been identified. Further studies are needed to determine the underlying mechanisms of its extracellular actions and their importance for human health and life processes in other organisms.

Clinical significance

The enzymes that make and use NAD⁺ and NADH are important in both pharmacology and the research into future treatments for disease. Drug design and drug development exploits NAD⁺ in three ways: as a direct target of drugs, by designing enzyme inhibitors or activators based on its structure that change the activity of NAD-dependent enzymes, and by trying to inhibit NAD⁺ biosynthesis.

Because cancer cells utilize increased glycolysis, and because NAD enhances glycolysis, nicotinamide phosphoribosyltransferase (NAD salvage pathway) is often amplified in cancer cells.

It has been studied for its potential use in the therapy of neurodegenerative diseases such as Alzheimer's and Parkinson's disease and Muliple sclerosis. A placebo-controlled clinical trial of NADH (which excluded NADH precursors) in people with Parkinson's failed to show any effect.

NAD⁺ is also a direct target of the drug isoniazid, which is used in the treatment of tuberculosis, an infection caused by Mycobacterium tuberculosis. Isoniazid is a prodrug and once it has entered the bacteria, it is activated by a peroxidase enzyme, which oxidizes the compound into a free radical form. This radical then reacts with NADH, to produce adducts that are very potent inhibitors of the enzymes enoyl-acyl carrier protein reductase, and dihydrofolate reductase.

Since many oxidoreductases use NAD⁺ and NADH as substrates, and bind them using a highly conserved structural motif, the idea that inhibitors based on NAD⁺ could be specific to one enzyme is surprising. However, this can be possible: for example, inhibitors based on the compounds mycophenolic acid and tiazofurin inhibit IMP dehydrogenase at the NAD⁺ binding site. Because of the importance of this enzyme in purine metabolism, these compounds may be useful as anti-cancer, anti-viral, or immunosuppressive drugs. Other drugs are not enzyme inhibitors, but instead activate enzymes involved in NAD⁺ metabolism. Sirtuins are a particularly interesting target for such drugs, since activation of these NAD-dependent deacetylases extends lifespan in some animal models. Compounds such as resveratrol increase the activity of these enzymes, which may be important in their ability to delay aging in both vertebrate, and invertebrate model organisms. In one experiment, mice given NAD for one week had improved nuclear-mitochrondrial communication.

Because of the differences in the metabolic pathways of NAD⁺ biosynthesis between organisms, such as between bacteria and humans, this area of metabolism is a promising area for the development of new antibiotics. For example, the enzyme nicotinamidase, which converts nicotinamide to nicotinic acid, is a target for drug design, as this enzyme is absent in humans but present in yeast and bacteria.

In bacteriology, NAD, sometimes referred to factor V, is used as a supplement to culture media for some fastidious bacteria.

History

Arthur Harden, co-discoverer of NAD

 

Further information: History of biochemistry

The coenzyme NAD⁺ was first discovered by the British biochemists Arthur Harden and William John Young in 1906. They noticed that adding boiled and filtered yeast extract greatly accelerated alcoholic fermentation in unboiled yeast extracts. They called the unidentified factor responsible for this effect a coferment. Through a long and difficult purification from yeast extracts, this heat-stable factor was identified as a nucleotide sugar phosphate by Hans von Euler-Chelpin. In 1936, the German scientist Otto Heinrich Warburg showed the function of the nucleotide coenzyme in hydride transfer and identified the nicotinamide portion as the site of redox reactions.

Vitamin precursors of NAD⁺ were first identified in 1938, when Conrad Elvehjem showed that liver has an "anti-black tongue" activity in the form of nicotinamide. Then, in 1939, he provided the first strong evidence that niacin is used to synthesize NAD⁺. In the early 1940s, Arthur Kornberg was the first to detect an enzyme in the biosynthetic pathway. In 1949, the American biochemists Morris Friedkin and Albert L. Lehninger proved that NADH linked metabolic pathways such as the citric acid cycle with the synthesis of ATP in oxidative phosphorylation. In 1958, Jack Preiss and Philip Handler discovered the intermediates and enzymes involved in the biosynthesis of NAD⁺; salvage synthesis from nicotinic acid is termed the Preiss-Handler pathway. In 2004, Charles Brenner and co-workers uncovered the nicotinamide riboside kinase pathway to NAD⁺.

The non-redox roles of NAD(P) were discovered later. The first to be identified was the use of NAD⁺ as the ADP-ribose donor in ADP-ribosylation reactions, observed in the early 1960s. Studies in the 1980s and 1990s revealed the activities of NAD⁺ and NADP⁺ metabolites in cell signaling – such as the action of cyclic ADP-ribose, which was discovered in 1987.

The metabolism of NAD⁺ remained an area of intense research into the 21st century, with interest heightened after the discovery of the NAD⁺-dependent protein deacetylases called sirtuins in 2000, by Shin-ichiro Imai and coworkers in the laboratory of Leonard P. Guarente. In 2009 Imai proposed the "NAD World" hypothesis that key regulators of aging and longevity in mammals are sirtuin 1 and the primary NAD⁺ synthesizing enzyme nicotinamide phosphoribosyltransferase (NAMPT). In 2016 Imai expanded his hypothesis to "NAD World 2.0", which postulates that extracellular NAMPT from adipose tissue maintains NAD⁺ in the hypothalamus (the control center) in conjunction with myokines from skeletal muscle cells.