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What does ensemble-based model of enzyme mean?

What does ensemble-based model of enzyme mean?



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I am reading Pan et al. (2000), a paper about dihydrofolate reductase (DHFR). They claim using a ensemble-based model of DHFR.

What is a ensemble-based model?


An 'ensemble' is a set of related models of a structure.

For example, conformational ensembles describe the structure of flexible proteins. There is a database of such structures here : http://pedb.vib.be/.

When NMR is used to determine a structure, the result is an ensemble of structures (see this paper for a comparison with crystallography). However, it is possible to see multiple structures for crystallography as well. This paper on "Accessing protein conformational ensembles using room-temperature X-ray crystallography" describes limitations to the idea of crystal structures having one unique structure.


Metabolic pathways consist of chains and cycles of enzyme-catalysed reactions.

Initially the substrate does not fit perfectly into the active site of the enzyme. When the substrate binds to the active site, this changes the shape of the active site and only then does it perfectly fit the substrate. As the substrate binds it changes the shape of the active site and this weakens the bonds in the substrate and therefore reduces the activation energy. This model is a more precise version of the lock and key one. The reason for this is that it explains why some enzymes can bind to many different substrates. If the shape of the active site changes when a substrate binds, this allows many different but similar substrates to bind to the one enzyme.

Figure 7.6.1 - The induced-fit model


Biology toolbox: Enzyme-substrate interactions and inhibition

Figure 1: A diagram showing the Lock and Key model of enzyme-substrate interaction. Model 2: Induced Fit In the Induced Fit model, the enzyme active site forms in response to substrate binding. In the diagram, sites a, b and c move in response to the binding substrate. So initially the active site is not perfect, but upon binding, it is able to move, which puts the active site under strain. This strain is then able to elicit the energy that’s required for the reaction to occur by stabilizing the transition state and not just binding of the substrate. The enzyme carries out its work by inducing the substrate to take up a transition state on the path to the required product. Figure 2: A diagram showing the Induced Fit model of enzyme substrate interaction

IUB System of Classification and Nomenclature of Enzymes

Ø Enzymes are classified based on the reaction they catalyze.

Ø Usually, the enzymes are named by adding the suffix “-ase” to the name of their substrate or to a word describing their activity.

$. Urease:- Catalyzes the hydrolysis of urea.

$. DNA Polymerase:– Catalyzes the polymerization of DNA.

Ø Many enzymes have common names that provide little information about the reactions that they catalyze.

Ø Example: Trypsin – a proteolytic enzyme secreted by the pancreas.

Rules for the Nomenclature and Classification of Enzymes

Ø Biochemists follow systematic rules for the Nomenclature (naming) and Classification of enzymes.

Ø Rules for the Nomenclature and Classification of enzymes were prepared by the International Union of Biochemistry (IUB*) in 1964.

Ø *IUB is now IUBMB (International Union of Biochemistry and Molecular Biology).

Ø In the IUB system, each enzyme has a name and a unique identification number.

Ø The systematic name of each enzyme consists of TWO parts.

$. Name of the substrate(s)

$. Followed by a word ending in ‘-ase’ specifying the type of reaction

Ø Example: Alcohol Dehydrogenase

Ø The recommended name of the enzyme for everyday uses is often an enzyme’s previously used name (common name)

Ø The unique identification number of the enzyme is called Enzyme Commission Number (E.C. Number)

IUB System of Enzyme Classification

Ø All enzymes were categorized into SEVEN major Classes (*previously six) based on the type of reaction they catalyze.

Ø These groups were subdivided and further subdivided so many categories.

Ø Based on this classification, a ‘four-digit unique number’ (called EC number) is assigned to each enzyme as an identification code (The concept of EC Number is described below).

Six Classes of Enzymes (IUB System)

(1). Oxidoreductases

(2). Transferases

(3). Hydrolases

(5). Isomerases

(7). Translocases

(1). Oxidoreductases

Ø Oxidoreductases catalyze oxidation and reduction reactions (redox reactions).

Ø This reaction involves the transfer of protons or electrons between substrates.

Ø Example: Alcohol Dehydrogenase, Odidase

(2). Transferases

Ø Transferases catalyze the transfer of a functional group from one substance to another.

Ø The groups usually transferred by these enzymes are methyl group, ethyl group, amino group, phosphate group etc.)

Ø Example: Transaminase, Nucleoside Monophosphate Kinase (NMP Kinase)

(3). Hydrolases

Ø Hydrolases catalyze the hydrolysis (breakdown) reaction with water (transfer of functional groups to water)

Ø Example: Lipase, Amylase, Peptidase

Ø Lyases catalyze the addition of groups to double bonds and formation of double bonds by the removal of groups.

Ø Example: Fumarase, Decarboxylase

(5). Isomerase:

Ø Isomerases catalyze the transfer of groups within molecules to yield isomeric forms (isomeric reactions).

Ø It catalyzes the intra-molecular (within the molecule) group transfer.

Ø Example: Triose phosphate isomerase, Phosphoglucomutase

Ø Ligases catalyze the condensation (joining) of two molecules with the expense of energy from ATP hydrolysis.

Ø It catalyzes the formation of C – C, C – S, C – O and C – N bonds by the condensation reaction coupled with ATP cleavage.

Ø Example: Aminoacyl-tRNA synthetase, DNA ligase

(7*). Translocases

Ø *This class is a newly added major class of IUBMB enzyme classification.

Ø Translocases catalyse the movement of ions or molecules across membranes or their separation within membranes.

ATP Synthase (Source: Wikipedia)

Features of IUB System of Classification of Enzymes

Ø All enzymes have been classified into 7 major classes.

Ø Each major class is divided into sub-classes.

Ø Each sub-class is further divided into sub-sub-classes.

Ø Each enzyme has been assigned with a specific code number.

Ø This code number is called Enzyme Commission Number (EC number).

Ø EC number consists of 4 digits, separated by periods.

Ø Each digit in the EC number indicates a specific category in the classification.

$. Example: Alcohol Dehydrogenase (EC No. 1.1.1.1)

$. First digit indicate the major class

$. Second digit indicates sub-class

$. Third digit indicates the sub-sub-class

$. Forth digit indicates the systematic specific name of the enzyme.

Ø The systematic specific name of the enzyme consists of TWO parts.

$. The first part indicates the name of the substrate.

$. The second part indicates the nature of the reaction.

$. Example: Alcohol Dehydrogenases (ADH)

E.C. Number Example:

Ø Consider the following reaction in Glycolysis.

ATP + D-glucose → ADP + D-glucose 6-phosphate

Ø Enzyme responsible for this reaction is Hexokinase.

Ø New name of hexokinase according to IUMB is ATP-Glucose phospho-transferase.

Ø The name indicates the transfer of phosphoryl group from ATP to glucose.

Ø E.C. Number of Hexokinase is 2.7.1.1

$. First number (2) indicates the class name (Transferases).

$. Second number (7): sub-class (Phosphotransferases).

$. Third number (1) a phosphotransferase with a hydroxyl group as acceptor.

$. Fourth number (1) D glucose as the phosphoryl group acceptor.

Ø For any enzyme, a trivial name is more commonly used.

Ø Here the name is Hexokinase.

Lehninger A.B., (2018), Textbook of Biochemistry, Ed. 5, Pearson International, New York

Berg, J.M., Tymoczko, J.L. and Stryer, L., 2012. Biochemistry/Jeremy M. Berg, John L. Tymoczko, Lubert Stryer with Gregory J. Gatto, Jr.

Voet, D., Voet, J.G. and Pratt, C.W., 2013. Fundamentals of biochemistry: life at the molecular level

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Lysozyme: a model of enzyme action

Lysozyme is a globular protein with a deep cleft across part of its surface. Six hexoses of the substrate fit into this cleft.

  • With so many oxygen atoms in sugars, as many as 14 hydrogen bonds form between the six amino sugars and certain amino acid R groups such as Arg-114, Asn-37, Asn-44, Trp-62, Trp-63, and Asp-101.
  • Some hydrogen bonds also form with the C=O groups of several peptide bonds.
  • In addition, hydrophobic interactions may help hold the substrate in position.

X-ray crystallography has shown that as lysozyme and its substrate unite, each is slightly deformed. The fourth hexose in the chain (ring #4) becomes twisted out of its normal position. This imposes a strain on the C-O bond on the ring-4 side of the oxygen bridge between rings 4 and 5. It is just at this point that the polysaccharide is broken. A molecule of water is inserted between these two hexoses, which breaks the chain. Here, then, is a structural view of what it means to lower activation energy. The energy needed to break this covalent bond is lower now that the atoms connected by the bond have been distorted from their normal position.

As for lysozyme itself, binding of the substrate induces a small (

0.75Å) movement of certain amino acid residues so the cleft closes slightly over its substrate. So the "lock" as well as the "key" changes shape as the two are brought together. (This is sometimes called "induced fit".)

The amino acid residues in the vicinity of rings 4 and 5 provide a plausible mechanism for completing the catalytic act. Residue 35, glutamic acid (Glu-35), is about 3Å from the -O- bridge that is to be broken. The free carboxyl group of glutamic acid is a hydrogen ion donor and available to transfer H + to the oxygen atom. This would break the already-strained bond between the oxygen atom and the carbon atom of ring 4.

Now having lost an electron, the carbon atom acquires a positive charge. Ionized carbon is normally very unstable, but the attraction of the negatively-charged carboxyl ion of Asp-52 could stabilize it long enough for an -OH ion (from a spontaneously dissociated water molecule) to unite with the carbon. Even at pH 7, water spontaneously dissociates to produce H + and OH - ions. [Discussion] The hydrogen ion (H + ) left over can replace that lost by Glu-35.

In the 20 August 2001 issue of Nature, Vocadlo, D. J., et al., report evidence that Asp-52 stabilizes ring 4 by forming a transient covalent bond rather than through ionic interactions.

In either case, the chain is broken, the two fragments separate from the enzyme, and the enzyme is free to attach to a new location on the bacterial cell wall and continue its work of digesting it.


Section Overview

Enzymes have an active site that provides a unique chemical environment, made up of certain amino acid R groups (residues) in a particular orientations and distance from one another. This unique environment is well-suited to convert particular chemical reactants for that enzyme, called substrates, into unstable intermediates (transition states). Enzymes and substrates are thought to bind with an "induced fit", which means that enzymes and substrates undergo slight conformational adjustments upon substrate contact, leading to binding. This subtle change in enzyme shape allows the enzyme to rapidly bind potential substrates in an "open" conformation" and then generate a tighter "closed" catalytically active alternative conformation only when the correct substrate is correctly aligned in the active site.

Enzymes bind to substrates and can potentially catalyze reactions in four different ways (which might act together in a single enzyme): bringing substrates together in an optimal orientation, compromising the bond structures of substrates so that bonds can be more easily broken, providing optimal environmental conditions (often local pH) for a reaction to occur, and/or participating directly in their chemical reaction by forming transient covalent bonds with their substrates.

Enzyme action must be regulated so that in a given cell at a given time, the desired reactions are being catalyzed and the undesired reactions are not. Inhibition and activation of enzymes via other molecules are important ways that enzymes are regulated. Inhibitors can act competitively or noncompetitively noncompetitive inhibitors are usually allosteric (allo (other) steric (form). Activators can also enhance the function of enzymes allosterically. A common method by which cells regulate the enzymes in metabolic pathways is through feedback inhibition. During feedback inhibition, the products of a metabolic pathway serve as inhibitors (usually allosteric) of one or more of the enzymes (usually the first committed enzyme of the pathway) involved in the pathway that produces them.

Enzyme Active Site and Substrate Specificity

The chemical reactants to which an enzyme binds are the enzyme&rsquos . There may be one or more substrates, depending on the particular chemical reaction. In some reactions, a single-reactant substrate is broken down into multiple products. In others, two substrates may come together to create one larger molecule. Two reactants might also enter a reaction, both become modified, and leave the reaction as two products. The location within the enzyme where the substrate binds is called the enzyme&rsquos . Since enzymes are proteins, there is a unique combination of amino acid R groups within the active site. Each amino acid side-chain is characterized by different properties. The unique combination of amino acids, their positions, sequences, structures, and properties, creates a very specific chemical environment within the active site. This specific environment is suited to bind, albeit briefly, to a specific chemical substrate (or substrates). Due to this jigsaw puzzle-like match between an enzyme and its substrates, enzymes can be extremely specific in their choice of substrates. The &ldquobest fit&rdquo between an enzyme and its substrates results from their respective shapes and the chemical complementarity of the functional groups on each binding partner.

This is an enzyme with two different substrates bound in the active site (here conveniently squished down to 2 dimensions). The enzymes are represented as blobs, except for the active site which identifies three amino acids located in the active site (and shows the R group for one of them). The R group of R180 is interacting with the substrates through hydrogen bonding (represented as dashed lines), as are some groups in the peptide backbone. Amino acid positions are denoted a single letter code for the amino acid followed immediately by "position of the amino acid vs. the N terminal end". For example "R180" means an R (arginine) is the 180th amino acid from the N terminus. (Minor note- the R is this diagram is drawn incorrectly, though teh business end- where it connects to substrate- is OK.)

At this point in the class you should be familiar with the chemical characteristics (charge, polarity, hydrophobicity) of the functional groups. For example, the R group of R180 in the enzyme depicted above is the amino acid Arginine (arginine's single letter code happens to be R, which is a little confusing in this context) and R180's R group consists of several "amino" functional groups. An amino functional group contains a nitrogen (N) and hydrogen (H) atoms. Nitrogen is more electronegative than hydrogen so the covalent bond between N-H is a polar covalent bond. The hydrogen atoms in this bond will have a partial positive charge, and the nitrogen atom will have a partial negative charge. This allows amino groups to form hydrogen bonds with other polar compounds. Likewise, the backbone carbonyl oxygens of Valine (V81) and Glycine (G121) the backbone amino hydrogen of V81 are depicted engaged in hydrogen bonds with the small molecule substrate.

Exercise

Look to see which atoms in the figure above are involved in the hydrogen bonds between the amino acid R groups and the substrate.
Which substrate (the left or right one) do you think is more stable in the active site? Why? How?

This is a depiction of an enzyme active site. Only the amino acids in the active site are drawn the numbers refer to their positions in the primary sequence of the protein (and aren't really important here). The substrate is sitting directly in the center.
Source: Created by Marc T. Facciotti (original work)

Exercise

First, identify the type of molecule in the center of the figure above. Second, draw in and label the appropriate interactions between the R groups and the substrate.

Structural Instability of Enzymes

The fact that active sites are so well-suited to provide specific environmental conditions also means that they are subject to influences by the local environment. It is true that increasing the environmental temperature generally increases reaction rates, enzyme-catalyzed or otherwise. However, increasing or decreasing the temperature outside of an optimal range can affect chemical bonds within the active site in such a way that they are less well suited to bind substrates. High temperatures will eventually cause enzymes, like some other biological molecules, to , a process that changes the natural properties of a substance. Likewise, the pH of the local environment can also affect enzyme function. Active site amino acid residues have their own acidic or basic properties that are optimal for catalysis. These residues are sensitive to changes in pH that can impair the way substrate molecules bind, because the charges on the R groups, and therefore both ionic and H-bonding interactions can change with pH. Enzymes are suited to function best within a certain pH range, and, as with temperature, extreme pH values (acidic or basic) of the environment can cause enzymes to denature.

Enzymes have an optimal pH. The pH at which the enzyme is most active will be the pH where the active site R groups are protonated/deprotonated such that the substrate can enter the active site and the initial step in the reaction can begin. Some enzymes require a very low pH (acidic) to be completely active. In the human body, these enzymes are most likely located in the stomach, or located in lysosomes (a cellular organelle used to digest large compounds inside the cell).
Source: http://biowiki.ucdavis.edu/Biochemis. _pH_Inhibition

The process where enzymes denature usually starts with the unwinding of the tertiary structure through destabilization of the bonds holding the tertiary structure together. Hydrogen bonds, are ionic bonds easily disrupted by mild changes in temperate and pH, the disruption of covalent bonds would require more extreme conditions. Using the chart of enzyme activity and temperature below, make an energy story for the red enzyme. Explain what might be happening from temperature 37C to 95C.

Enzymes have an optimal temperature. The temperature at which the enzyme is most active will usually be the temperature where the structure of the enzyme is stable or uncompromised. Some enzymes require a specific temperature to remain active and not denature.
Source: academic.brooklyn.cuny.edu/bi. ge/enz_act.htm

Induced Fit and Enzyme Function

For many years, scientists thought that enzyme-substrate binding took place in a simple &ldquolock-and-key&rdquo fashion. This model asserted that the enzyme and substrate fit together perfectly in one instantaneous step. However, current research supports a more refined view called . The induced-fit model expands upon the lock-and-key model by describing a more dynamic interaction between enzyme and substrate. As the enzyme and substrate come together, their interaction causes a mild shift in the enzyme&rsquos structure that confirms an more productive binding arrangement between the enzyme and the transition state of the substrate. This energetically favorable binding maximizes the enzyme&rsquos ability to catalyze its reaction.

When an enzyme binds its substrate, an enzyme-substrate complex is formed. This complex lowers the activation energy of the reaction and promotes its rapid progression in one of many ways. On a basic level, enzymes promote chemical reactions that involve more than one substrate by bringing the substrates together in an optimal orientation. The appropriate region (atoms and bonds) of one molecule is juxtaposed to the appropriate region of the other molecule with which it must react. Another way in which enzymes promote the reaction of their substrates is by creating an energetically favorable environment within the active site for the reaction to occur. Certain chemical reactions might proceed best in a slightly acidic or non-polar environment. The chemical properties that emerge from the particular arrangement of amino acid residues within an active site create the energetically favorable environment for an enzyme&rsquos specific substrates to react.

The activation energy required for many reactions includes the energy involved in slightly contorting chemical bonds so that they can more easily react. Enzymatic action can aid this process. The enzyme-substrate complex can lower the activation energy by contorting substrate molecules in such a way as to facilitate bond-breaking. Finally, enzymes can also lower activation energies by taking part in the chemical reaction itself. The amino acid residues can provide certain ions or chemical groups that actually form covalent bonds with substrate molecules as a necessary step of the reaction process. In this case, the enzyme is providing an alternate, lower-transition state energy path to the overall reaction. In all cases, it is important to remember that the enzyme will always return to its original state at the completion of the reaction. After an enzyme is done catalyzing a reaction, it releases its product(s). Always keep in mind that enzymes can also facilitate the reverse reaction. The net flux will depend on the reaction direction that provides a negative ∆G.

According to the induced-fit model, both enzyme and substrate undergo dynamic conformational changes upon binding. The enzyme contorts the substrate into its transition state, thereby increasing the rate of the reaction.

Creating an Energy story for the reaction above

Using the figure above, answer the questions posed in the energy story.
1. What are the reactants? What are the products?
2. What work was accomplished by the enzyme?
3. What state is the energy in initially? What state is the energy transformed into in the final state? This one might be tricky still, but try to identify where the energy is in the initial state and the final state.

Speaking of energy: If a protein "bends" a substrate such that it approaches the transition state, where does the energy for that bending come from?

Enzyme Regulation

Why regulate enzymes?

Cellular needs and conditions vary from cell to cell, and change within individual cells over time. The required enzymes and energetic demands of stomach cells are different from those of fat storage cells, skin cells, blood cells, and nerve cells. Furthermore, a digestive cell works much harder to process and break down nutrients during the time that closely follows a meal compared with many hours after a meal. As these cellular demands and conditions vary, so do the needed amounts and functionality of different enzymes.

Regulation of Enzymes by Molecules

Enzymes can be regulated in ways that either promote or reduce their activity. Although this inhibition might be the basis of the action of certain poisons, in many cases an enzyme may have evolved to respond to environmental influences (such as the concentration of relevant metabolites, which are not necessarily substrates or products) by regulating its own activity. There are many different kinds of molecules that inhibit or promote enzyme function, and various mechanisms exist for doing so. In some cases of enzyme inhibition, for example, an inhibitor molecule is similar enough to a substrate that it can bind to the active site and simply block the substrate from binding. When this happens, the enzyme is inhibited through , because an inhibitor molecule competes with the substrate for active site binding. On the other hand, in noncompetitive inhibition, an inhibitor molecule binds to the enzyme in a location other than an active site. That binding alters the overall shape of the enzyme such that it no longer binds its substrate effectively. This type of inhibition is called .

Competitive and noncompetitive inhibition affect the rate of reaction differently. Competitive inhibitors affect the initial rate but do not affect the maximal rate, whereas noncompetitive inhibitors affect the maximal rate.

Discuss: Why are the effects of competitive inhibitors overcome by high concentrations of substrate?

Most allosterically regulated enzymes are made up of more than one polypeptide, meaning that they have more than one protein subunit. When an allosteric inhibitor binds to an enzyme, all active sites on the protein subunits are changed slightly such that they bind their substrates with less efficiency. There are as well as inhibitors. Allosteric activators bind to locations on an enzyme away from the active site, inducing a conformational change that increases the affinity of the enzyme&rsquos active site(s) for its substrate(s).

Allosteric inhibitors modify the active site of the enzyme so that substrate binding is reduced or prevented. In contrast, allosteric activators modify the active site of the enzyme so that the affinity for the substrate increases.

Video Link

Check out this short (1 minute) video on competitive vs. noncompetitive enzymatic inhibition. Also, take a look at this video (1.2 minutes) on feedback inhibition.

Many enzymes don&rsquot work optimally, or even at all, unless bound to other specific non-protein helper molecules, either temporarily, through ionic or hydrogen bonds, or permanently through stronger covalent bonds. These helper molecules are termed . Binding to these molecules promotes optimal conformation and function for their respective enzymes. Cofactors may be inorganic ions such as iron (Fe 2+ ) and magnesium (Mg 2+ ), and these ions maybe be linked to larger nonprotein molecules. The term coenzyme is sometimes used to define a subclass of cofactors that are organic helper molecules, with a molecular structure made up of carbon, nitrogen and hydrogen, which are required for enzyme action (for example, a heme group, as a opposed to a metal ion or iron-sulfur cluster). There are also further specialized terms for subclasses of cofactors. These terms are employed loosely, variously, and irregularly by different scientists, and I suggest you stick with the safe, all-encompassing term "cofactor". The most common sources of organic cofactors are dietary vitamins. Vitamin C is a cofactor for multiple enzymes that take part in building the important connective tissue component, collagen. Hence a lack of vitamin C in our diet results in scurvy, a painful disease of connective tissue. An important step in the breakdown of glucose to yield energy is catalysis of pyruvate to acteyl coA by a multi-enzyme complex called pyruvate dehydrogenase. Pyruvate dehydrogenase is a complex of several enzymes that actually requires one inorganic cofactor (a magnesium ion) and five different organic coenzymes to catalyze its specific chemical reaction.

Succinate dehydrogenase, an enzyme involved in both electron transport and the citric acid cycle, is an example of an enzyme that carries many cofactors, allowing it to transport electrons through the enzyme from the original donor molecule (succinate), through FAD/FADH (flavin), iron sulfur clusters, and heme, finally to Q, in the respiratory ETC. The function of this enzyme made possible only via the presence of various cofactors.

Succinate dehydrogenase (SDH) oxidizes succinate to fumarate, using FAD as the immediate 2e- acceptor. The FAD of subunit SDHA is considered a cofactor as it does not leave the enzyme, but is directly oxidized by nearby iron sulfur clusters, within the B subunit of this enzyme. These are in turn oxidized by iron containing heme group within the SDHC and D subunits. Finally the electrons leave thsi complex via transfer to the membrane-diffusible molecule ubiquinone, also known as, coenzyme Q. The electrons will proceed further down respiratory ETC. All of these transfers, of course, can only occur if there is some electron acceptor at the end of the ETC.

Enzyme Compartmentalization

In eukaryotic cells, molecules such as enzymes are usually compartmentalized into different organelles. This allows for yet another level of regulation of enzyme activity. Enzymes required only for certain cellular processes can be housed separately along with their substrates, allowing for more efficient chemical reactions. Examples of this sort of enzyme regulation based on location and proximity include the enzymes involved in the latter stages of cellular respiration, which take place exclusively in the mitochondria, and the enzymes involved in the digestion of cellular debris and foreign materials, located within lysosomes.

Additional Links

Khan Academy

The following links will take you to a series of videos on kinetics. The first link contains 4 videos on reaction rates and the second link contains 9 videos related to the relationship between reaction rates and concentration. These videos are supplemental and are provided to give you an outside resource to further explore enzyme kenetics.


Contents

The reaction catalysed by an enzyme uses exactly the same reactants and produces exactly the same products as the uncatalysed reaction. Like other catalysts, enzymes do not alter the position of equilibrium between substrates and products. [1] However, unlike uncatalysed chemical reactions, enzyme-catalysed reactions display saturation kinetics. [2] For a given enzyme concentration and for relatively low substrate concentrations, the reaction rate increases linearly with substrate concentration the enzyme molecules are largely free to catalyse the reaction, and increasing substrate concentration means an increasing rate at which the enzyme and substrate molecules encounter one another. However, at relatively high substrate concentrations, the reaction rate asymptotically approaches the theoretical maximum the enzyme active sites are almost all occupied by substrates resulting in saturation, and the reaction rate is determined by the intrinsic turnover rate of the enzyme. [3] The substrate concentration midway between these two limiting cases is denoted by KM. Thus, KM is the substrate concentration at which the reaction velocity is half of the maximum velocity. [3]

The two most important kinetic properties of an enzyme are how easily the enzyme becomes saturated with a particular substrate, and the maximum rate it can achieve. Knowing these properties suggests what an enzyme might do in the cell and can show how the enzyme will respond to changes in these conditions.

Enzyme assays are laboratory procedures that measure the rate of enzyme reactions. Since enzymes are not consumed by the reactions they catalyse, enzyme assays usually follow changes in the concentration of either substrates or products to measure the rate of reaction. There are many methods of measurement. Spectrophotometric assays observe change in the absorbance of light between products and reactants radiometric assays involve the incorporation or release of radioactivity to measure the amount of product made over time. Spectrophotometric assays are most convenient since they allow the rate of the reaction to be measured continuously. Although radiometric assays require the removal and counting of samples (i.e., they are discontinuous assays) they are usually extremely sensitive and can measure very low levels of enzyme activity. [4] An analogous approach is to use mass spectrometry to monitor the incorporation or release of stable isotopes as substrate is converted into product. Occasionally, an assay fails and approaches are essential to resurrect a failed assay.

The most sensitive enzyme assays use lasers focused through a microscope to observe changes in single enzyme molecules as they catalyse their reactions. These measurements either use changes in the fluorescence of cofactors during an enzyme's reaction mechanism, or of fluorescent dyes added onto specific sites of the protein to report movements that occur during catalysis. [5] These studies are providing a new view of the kinetics and dynamics of single enzymes, as opposed to traditional enzyme kinetics, which observes the average behaviour of populations of millions of enzyme molecules. [6] [7]

An example progress curve for an enzyme assay is shown above. The enzyme produces product at an initial rate that is approximately linear for a short period after the start of the reaction. As the reaction proceeds and substrate is consumed, the rate continuously slows (so long as substrate is not still at saturating levels). To measure the initial (and maximal) rate, enzyme assays are typically carried out while the reaction has progressed only a few percent towards total completion. The length of the initial rate period depends on the assay conditions and can range from milliseconds to hours. However, equipment for rapidly mixing liquids allows fast kinetic measurements on initial rates of less than one second. [8] These very rapid assays are essential for measuring pre-steady-state kinetics, which are discussed below.

Most enzyme kinetics studies concentrate on this initial, approximately linear part of enzyme reactions. However, it is also possible to measure the complete reaction curve and fit this data to a non-linear rate equation. This way of measuring enzyme reactions is called progress-curve analysis. [9] This approach is useful as an alternative to rapid kinetics when the initial rate is too fast to measure accurately.

Enzymes with single-substrate mechanisms include isomerases such as triosephosphateisomerase or bisphosphoglycerate mutase, intramolecular lyases such as adenylate cyclase and the hammerhead ribozyme, an RNA lyase. [10] However, some enzymes that only have a single substrate do not fall into this category of mechanisms. Catalase is an example of this, as the enzyme reacts with a first molecule of hydrogen peroxide substrate, becomes oxidised and is then reduced by a second molecule of substrate. Although a single substrate is involved, the existence of a modified enzyme intermediate means that the mechanism of catalase is actually a ping–pong mechanism, a type of mechanism that is discussed in the Multi-substrate reactions section below.

Michaelis–Menten kinetics Edit

As enzyme-catalysed reactions are saturable, their rate of catalysis does not show a linear response to increasing substrate. If the initial rate of the reaction is measured over a range of substrate concentrations (denoted as [S]), the initial reaction rate ( v 0 > ) increases as [S] increases, as shown on the right. However, as [S] gets higher, the enzyme becomes saturated with substrate and the initial rate reaches Vmax, the enzyme's maximum rate.

The Michaelis–Menten equation [11] describes how the (initial) reaction rate v0 depends on the position of the substrate-binding equilibrium and the rate constant k2.

This Michaelis–Menten equation is the basis for most single-substrate enzyme kinetics. Two crucial assumptions underlie this equation (apart from the general assumption about the mechanism only involving no intermediate or product inhibition, and there is no allostericity or cooperativity). The first assumption is the so-called quasi-steady-state assumption (or pseudo-steady-state hypothesis), namely that the concentration of the substrate-bound enzyme (and hence also the unbound enzyme) changes much more slowly than those of the product and substrate and thus the change over time of the complex can be set to zero d [ ES ] / d t = ! 0 >/

<=>>> . The second assumption is that the total enzyme concentration does not change over time, thus [ E ] tot = [ E ] + [ ES ] = ! const >_< ext>=>+><=>>< ext>> . A complete derivation can be found here.

The Michaelis constant KM is experimentally defined as the concentration at which the rate of the enzyme reaction is half Vmax, which can be verified by substituting [S] = KM into the Michaelis–Menten equation and can also be seen graphically. If the rate-determining enzymatic step is slow compared to substrate dissociation ( k 2 ≪ k − 1 ll k_<-1>> ), the Michaelis constant KM is roughly the dissociation constant KD of the ES complex.

Thus the product formation rate depends on the enzyme concentration as well as on the substrate concentration, the equation resembles a bimolecular reaction with a corresponding pseudo-second order rate constant k 2 / K M /K_> . This constant is a measure of catalytic efficiency. The most efficient enzymes reach a k 2 / K M /K_> in the range of 10 8 – 10 10 M −1 s −1 . These enzymes are so efficient they effectively catalyse a reaction each time they encounter a substrate molecule and have thus reached an upper theoretical limit for efficiency (diffusion limit) and are sometimes referred to as kinetically perfect enzymes. [12] But most enzymes are far from perfect: the average values of k 2 / K M /K_< m >> and k 2 > are about 10 5 s − 1 M − 1 < m >^<-1>< m >^<-1>> and 10 s − 1 >^<-1>> , respectively. [13]

Direct use of the Michaelis–Menten equation for time course kinetic analysis Edit

The observed velocities predicted by the Michaelis–Menten equation can be used to directly model the time course disappearance of substrate and the production of product through incorporation of the Michaelis–Menten equation into the equation for first order chemical kinetics. This can only be achieved however if one recognises the problem associated with the use of Euler's number in the description of first order chemical kinetics. i.e. ek is a split constant that introduces a systematic error into calculations and can be rewritten as a single constant which represents the remaining substrate after each time period. [14]

In 1983 Stuart Beal (and also independently Santiago Schnell and Claudio Mendoza in 1997) derived a closed form solution for the time course kinetics analysis of the Michaelis-Menten mechanism. [15] [16] The solution, known as the Schnell-Mendoza equation, has the form:

where W[ ] is the Lambert-W function. [17] [18] and where F(t) is

This equation is encompassed by the equation below, obtained by Berberan-Santos, [19] which is also valid when the initial substrate concentration is close to that of enzyme,

Linear plots of the Michaelis–Menten equation Edit

The plot of v versus [S] above is not linear although initially linear at low [S], it bends over to saturate at high [S]. Before the modern era of nonlinear curve-fitting on computers, this nonlinearity could make it difficult to estimate KM and Vmax accurately. Therefore, several researchers developed linearisations of the Michaelis–Menten equation, such as the Lineweaver–Burk plot, the Eadie–Hofstee diagram and the Hanes–Woolf plot. All of these linear representations can be useful for visualising data, but none should be used to determine kinetic parameters, as computer software is readily available that allows for more accurate determination by nonlinear regression methods. [20]

The Lineweaver–Burk plot or double reciprocal plot is a common way of illustrating kinetic data. This is produced by taking the reciprocal of both sides of the Michaelis–Menten equation. As shown on the right, this is a linear form of the Michaelis–Menten equation and produces a straight line with the equation y = mx + c with a y-intercept equivalent to 1/Vmax and an x-intercept of the graph representing −1/KM.

Naturally, no experimental values can be taken at negative 1/[S] the lower limiting value 1/[S] = 0 (the y-intercept) corresponds to an infinite substrate concentration, where 1/v=1/Vmax as shown at the right thus, the x-intercept is an extrapolation of the experimental data taken at positive concentrations. More generally, the Lineweaver–Burk plot skews the importance of measurements taken at low substrate concentrations and, thus, can yield inaccurate estimates of Vmax and KM. [21] A more accurate linear plotting method is the Eadie–Hofstee plot. In this case, v is plotted against v/[S]. In the third common linear representation, the Hanes–Woolf plot, [S]/v is plotted against [S]. In general, data normalisation can help diminish the amount of experimental work and can increase the reliability of the output, and is suitable for both graphical and numerical analysis. [22]

Practical significance of kinetic constants Edit

The study of enzyme kinetics is important for two basic reasons. Firstly, it helps explain how enzymes work, and secondly, it helps predict how enzymes behave in living organisms. The kinetic constants defined above, KM and Vmax, are critical to attempts to understand how enzymes work together to control metabolism.

Making these predictions is not trivial, even for simple systems. For example, oxaloacetate is formed by malate dehydrogenase within the mitochondrion. Oxaloacetate can then be consumed by citrate synthase, phosphoenolpyruvate carboxykinase or aspartate aminotransferase, feeding into the citric acid cycle, gluconeogenesis or aspartic acid biosynthesis, respectively. Being able to predict how much oxaloacetate goes into which pathway requires knowledge of the concentration of oxaloacetate as well as the concentration and kinetics of each of these enzymes. This aim of predicting the behaviour of metabolic pathways reaches its most complex expression in the synthesis of huge amounts of kinetic and gene expression data into mathematical models of entire organisms. Alternatively, one useful simplification of the metabolic modelling problem is to ignore the underlying enzyme kinetics and only rely on information about the reaction network's stoichiometry, a technique called flux balance analysis. [23] [24]

Michaelis–Menten kinetics with intermediate Edit

One could also consider the less simple case

where a complex with the enzyme and an intermediate exists and the intermediate is converted into product in a second step. In this case we have a very similar equation [25]

but the constants are different

Multi-substrate reactions follow complex rate equations that describe how the substrates bind and in what sequence. The analysis of these reactions is much simpler if the concentration of substrate A is kept constant and substrate B varied. Under these conditions, the enzyme behaves just like a single-substrate enzyme and a plot of v by [S] gives apparent KM and Vmax constants for substrate B. If a set of these measurements is performed at different fixed concentrations of A, these data can be used to work out what the mechanism of the reaction is. For an enzyme that takes two substrates A and B and turns them into two products P and Q, there are two types of mechanism: ternary complex and ping–pong.

Ternary-complex mechanisms Edit

In these enzymes, both substrates bind to the enzyme at the same time to produce an EAB ternary complex. The order of binding can either be random (in a random mechanism) or substrates have to bind in a particular sequence (in an ordered mechanism). When a set of v by [S] curves (fixed A, varying B) from an enzyme with a ternary-complex mechanism are plotted in a Lineweaver–Burk plot, the set of lines produced will intersect.

Enzymes with ternary-complex mechanisms include glutathione S-transferase, [26] dihydrofolate reductase [27] and DNA polymerase. [28] The following links show short animations of the ternary-complex mechanisms of the enzymes dihydrofolate reductase [β] and DNA polymerase [γ] .

Ping–pong mechanisms Edit

As shown on the right, enzymes with a ping-pong mechanism can exist in two states, E and a chemically modified form of the enzyme E* this modified enzyme is known as an intermediate. In such mechanisms, substrate A binds, changes the enzyme to E* by, for example, transferring a chemical group to the active site, and is then released. Only after the first substrate is released can substrate B bind and react with the modified enzyme, regenerating the unmodified E form. When a set of v by [S] curves (fixed A, varying B) from an enzyme with a ping–pong mechanism are plotted in a Lineweaver–Burk plot, a set of parallel lines will be produced. This is called a secondary plot.

Enzymes with ping–pong mechanisms include some oxidoreductases such as thioredoxin peroxidase, [29] transferases such as acylneuraminate cytidylyltransferase [30] and serine proteases such as trypsin and chymotrypsin. [31] Serine proteases are a very common and diverse family of enzymes, including digestive enzymes (trypsin, chymotrypsin, and elastase), several enzymes of the blood clotting cascade and many others. In these serine proteases, the E* intermediate is an acyl-enzyme species formed by the attack of an active site serine residue on a peptide bond in a protein substrate. A short animation showing the mechanism of chymotrypsin is linked here. [δ]

External factors may limit the ability of an enzyme to catalyse a reaction in both directions (whereas the nature of a catalyst in itself means that it cannot catalyse just one direction, according to the principle of microscopic reversibility). We consider the case of an enzyme that catalyses the reaction in both directions:

Many different enzyme systems follow non Michaelis-Menten behavior. [33] A select few examples include kinetics of self-catalytic enzymes, cooperative and allosteric enzymes, interfacial and intracellular enzymes, processive enzymes and so forth. Some enzymes produce a sigmoid v by [S] plot, which often indicates cooperative binding of substrate to the active site. This means that the binding of one substrate molecule affects the binding of subsequent substrate molecules. This behavior is most common in multimeric enzymes with several interacting active sites. [34] [35] Here, the mechanism of cooperation is similar to that of hemoglobin, with binding of substrate to one active site altering the affinity of the other active sites for substrate molecules. Positive cooperativity occurs when binding of the first substrate molecule increases the affinity of the other active sites for substrate. Negative cooperativity occurs when binding of the first substrate decreases the affinity of the enzyme for other substrate molecules.

Allosteric enzymes include mammalian tyrosyl tRNA-synthetase, which shows negative cooperativity, [36] and bacterial aspartate transcarbamoylase [37] and phosphofructokinase, [38] which show positive cooperativity.

Cooperativity is surprisingly common and can help regulate the responses of enzymes to changes in the concentrations of their substrates. [34] Positive cooperativity makes enzymes much more sensitive to [S] and their activities can show large changes over a narrow range of substrate concentration. Conversely, negative cooperativity makes enzymes insensitive to small changes in [S].

The Hill equation (biochemistry) [39] is often used to describe the degree of cooperativity quantitatively in non-Michaelis–Menten kinetics. The derived Hill coefficient n measures how much the binding of substrate to one active site affects the binding of substrate to the other active sites. A Hill coefficient of <1 indicates negative cooperativity and a coefficient of >1 indicates positive cooperativity.

In the first moment after an enzyme is mixed with substrate, no product has been formed and no intermediates exist. The study of the next few milliseconds of the reaction is called pre-steady-state kinetics. Pre-steady-state kinetics is therefore concerned with the formation and consumption of enzyme–substrate intermediates (such as ES or E*) until their steady-state concentrations are reached.

This approach was first applied to the hydrolysis reaction catalysed by chymotrypsin. [40] Often, the detection of an intermediate is a vital piece of evidence in investigations of what mechanism an enzyme follows. For example, in the ping–pong mechanisms that are shown above, rapid kinetic measurements can follow the release of product P and measure the formation of the modified enzyme intermediate E*. [41] In the case of chymotrypsin, this intermediate is formed by an attack on the substrate by the nucleophilic serine in the active site and the formation of the acyl-enzyme intermediate.

In the figure to the right, the enzyme produces E* rapidly in the first few seconds of the reaction. The rate then slows as steady state is reached. This rapid burst phase of the reaction measures a single turnover of the enzyme. Consequently, the amount of product released in this burst, shown as the intercept on the y-axis of the graph, also gives the amount of functional enzyme which is present in the assay. [42]

An important goal of measuring enzyme kinetics is to determine the chemical mechanism of an enzyme reaction, i.e., the sequence of chemical steps that transform substrate into product. The kinetic approaches discussed above will show at what rates intermediates are formed and inter-converted, but they cannot identify exactly what these intermediates are.

Kinetic measurements taken under various solution conditions or on slightly modified enzymes or substrates often shed light on this chemical mechanism, as they reveal the rate-determining step or intermediates in the reaction. For example, the breaking of a covalent bond to a hydrogen atom is a common rate-determining step. Which of the possible hydrogen transfers is rate determining can be shown by measuring the kinetic effects of substituting each hydrogen by deuterium, its stable isotope. The rate will change when the critical hydrogen is replaced, due to a primary kinetic isotope effect, which occurs because bonds to deuterium are harder to break than bonds to hydrogen. [43] It is also possible to measure similar effects with other isotope substitutions, such as 13 C/ 12 C and 18 O/ 16 O, but these effects are more subtle. [44]

Isotopes can also be used to reveal the fate of various parts of the substrate molecules in the final products. For example, it is sometimes difficult to discern the origin of an oxygen atom in the final product since it may have come from water or from part of the substrate. This may be determined by systematically substituting oxygen's stable isotope 18 O into the various molecules that participate in the reaction and checking for the isotope in the product. [45] The chemical mechanism can also be elucidated by examining the kinetics and isotope effects under different pH conditions, [46] by altering the metal ions or other bound cofactors, [47] by site-directed mutagenesis of conserved amino acid residues, or by studying the behaviour of the enzyme in the presence of analogues of the substrate(s). [48]

Enzyme inhibitors are molecules that reduce or abolish enzyme activity, while enzyme activators are molecules that increase the catalytic rate of enzymes. These interactions can be either reversible (i.e., removal of the inhibitor restores enzyme activity) or irreversible (i.e., the inhibitor permanently inactivates the enzyme).

Reversible inhibitors Edit

Traditionally reversible enzyme inhibitors have been classified as competitive, uncompetitive, or non-competitive, according to their effects on KM and Vmax. These different effects result from the inhibitor binding to the enzyme E, to the enzyme–substrate complex ES, or to both, respectively. The division of these classes arises from a problem in their derivation and results in the need to use two different binding constants for one binding event. The binding of an inhibitor and its effect on the enzymatic activity are two distinctly different things, another problem the traditional equations fail to acknowledge. In noncompetitive inhibition the binding of the inhibitor results in 100% inhibition of the enzyme only, and fails to consider the possibility of anything in between. [49] In noncompetitive inhibition, the inhibitor will bind to an enzyme at its allosteric site therefore, the binding affinity, or inverse of KM, of the substrate with the enzyme will remain the same. On the other hand, the Vmax will decrease relative to an uninhibited enzyme. On a Lineweaver-Burk plot, the presence of a noncompetitive inhibitor is illustrated by a change in the y-intercept, defined as 1/Vmax. The x-intercept, defined as −1/KM, will remain the same. In competitive inhibition, the inhibitor will bind to an enzyme at the active site, competing with the substrate. As a result, the KM will increase and the Vmax will remain the same. [50] The common form of the inhibitory term also obscures the relationship between the inhibitor binding to the enzyme and its relationship to any other binding term be it the Michaelis–Menten equation or a dose response curve associated with ligand receptor binding. To demonstrate the relationship the following rearrangement can be made:

Adding zero to the bottom ([I]-[I])

This notation demonstrates that similar to the Michaelis–Menten equation, where the rate of reaction depends on the percent of the enzyme population interacting with substrate, the effect of the inhibitor is a result of the percent of the enzyme population interacting with inhibitor. The only problem with this equation in its present form is that it assumes absolute inhibition of the enzyme with inhibitor binding, when in fact there can be a wide range of effects anywhere from 100% inhibition of substrate turn over to just >0%. To account for this the equation can be easily modified to allow for different degrees of inhibition by including a delta Vmax term.

This term can then define the residual enzymatic activity present when the inhibitor is interacting with individual enzymes in the population. However the inclusion of this term has the added value of allowing for the possibility of activation if the secondary Vmax term turns out to be higher than the initial term. To account for the possibly of activation as well the notation can then be rewritten replacing the inhibitor "I" with a modifier term denoted here as "X".

While this terminology results in a simplified way of dealing with kinetic effects relating to the maximum velocity of the Michaelis–Menten equation, it highlights potential problems with the term used to describe effects relating to the KM. The KM relating to the affinity of the enzyme for the substrate should in most cases relate to potential changes in the binding site of the enzyme which would directly result from enzyme inhibitor interactions. As such a term similar to the one proposed above to modulate Vmax should be appropriate in most situations: [51]

A few examples of reversible inhibition belonging to the competitive and uncompetitive models have been discussed in the following papers. [52] [53] [54]

Irreversible inhibitors Edit

Enzyme inhibitors can also irreversibly inactivate enzymes, usually by covalently modifying active site residues. These reactions, which may be called suicide substrates, follow exponential decay functions and are usually saturable. Below saturation, they follow first order kinetics with respect to inhibitor. Irreversible inhibition could be classified into two distinct types. Affinity labelling is a type of irreversible inhibition where a functional group that is highly reactive modifies a catalytically critical residue on the protein of interest to bring about inhibition. Mechanism-based inhibition, on the other hand, involves binding of the inhibitor followed by enzyme mediated alterations that transform the latter into a reactive group that irreversibly modifies the enzyme.

Philosophical discourse on reversibility and irreversibility of inhibition Edit

Having discussed reversible inhibition and irreversible inhibition in the above two headings, it would have to be pointed out that the concept of reversibility (or irreversibility) is a purely theoretical construct exclusively dependent on the time-frame of the assay, i.e., a reversible assay involving association and dissociation of the inhibitor molecule in the minute timescales would seem irreversible if an assay assess the outcome in the seconds and vice versa. There is a continuum of inhibitor behaviors spanning reversibility and irreversibility at a given non-arbitrary assay time frame. There are inhibitors that show slow-onset behavior [52] and most of these inhibitors, invariably, also show tight-binding to the protein target of interest. [52] [53]

The favoured model for the enzyme–substrate interaction is the induced fit model. [55] This model proposes that the initial interaction between enzyme and substrate is relatively weak, but that these weak interactions rapidly induce conformational changes in the enzyme that strengthen binding. These conformational changes also bring catalytic residues in the active site close to the chemical bonds in the substrate that will be altered in the reaction. [56] Conformational changes can be measured using circular dichroism or dual polarisation interferometry. After binding takes place, one or more mechanisms of catalysis lower the energy of the reaction's transition state by providing an alternative chemical pathway for the reaction. Mechanisms of catalysis include catalysis by bond strain by proximity and orientation by active-site proton donors or acceptors covalent catalysis and quantum tunnelling. [41] [57]

Enzyme kinetics cannot prove which modes of catalysis are used by an enzyme. However, some kinetic data can suggest possibilities to be examined by other techniques. For example, a ping–pong mechanism with burst-phase pre-steady-state kinetics would suggest covalent catalysis might be important in this enzyme's mechanism. Alternatively, the observation of a strong pH effect on Vmax but not KM might indicate that a residue in the active site needs to be in a particular ionisation state for catalysis to occur.

In 1902 Victor Henri proposed a quantitative theory of enzyme kinetics, [58] but at the time the experimental significance of the hydrogen ion concentration was not yet recognized. After Peter Lauritz Sørensen had defined the logarithmic pH-scale and introduced the concept of buffering in 1909 [59] the German chemist Leonor Michaelis and Dr. Maud Leonora Menten (a postdoctoral researcher in Michaelis's lab at the time) repeated Henri's experiments and confirmed his equation, which is now generally referred to as Michaelis-Menten kinetics (sometimes also Henri-Michaelis-Menten kinetics). [60] Their work was further developed by G. E. Briggs and J. B. S. Haldane, who derived kinetic equations that are still widely considered today a starting point in modeling enzymatic activity. [61]

The major contribution of the Henri-Michaelis-Menten approach was to think of enzyme reactions in two stages. In the first, the substrate binds reversibly to the enzyme, forming the enzyme-substrate complex. This is sometimes called the Michaelis complex. The enzyme then catalyzes the chemical step in the reaction and releases the product. The kinetics of many enzymes is adequately described by the simple Michaelis-Menten model, but all enzymes have internal motions that are not accounted for in the model and can have significant contributions to the overall reaction kinetics. This can be modeled by introducing several Michaelis-Menten pathways that are connected with fluctuating rates, [62] [63] [64] which is a mathematical extension of the basic Michaelis Menten mechanism. [65]

ENZO (Enzyme Kinetics) is a graphical interface tool for building kinetic models of enzyme catalyzed reactions. ENZO automatically generates the corresponding differential equations from a stipulated enzyme reaction scheme. These differential equations are processed by a numerical solver and a regression algorithm which fits the coefficients of differential equations to experimentally observed time course curves. ENZO allows rapid evaluation of rival reaction schemes and can be used for routine tests in enzyme kinetics. [66]


Enzymes can catalyze reactions through a variety of mechanisms. Some of these include:

  • Bond strain: enzymes can destabilize bonds within the substrate.
  • Proximity and orientation: conformational changes in the enzyme upon substrate binding can bring reactive groups closer together or orient them so they can react.
  • Proton donors and acceptors: the presence of acidic or basic groups can affect bond polarization and reaction speed.
  • Electrostatic catalysis: electrostatic attractions between the enzyme and the substrate can stabilize the activated complex.
  • Covalent catalysis: covalent bonding to side chains or cofactors can lower the energy of the transition state.

As such, enzymes show that evolutionary biology has produced highly effective catalysts.

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The lock and key hypothesis/ the induced fit model

The lock and key hypothesis explains how enzymes can be so specific with their substrates and the reactions they catalyse. It describes how the enzyme’s active site has a very unique shape that complements the shape of a specific substrate. They can therefore fit exactly together.

The lock and key mechanism often means that an enzyme has specificity for one particular substrate, but it can also be compatible with a family of substrates, such as those possessing particular functional groups or modifications.

However, the lock and key hypothesis is now out of date, and scientists have developed a new model to explain how enzymes and substrates fit together. An important feature of enzymes not covered under the lock and key hypothesis, is that the active site changes shape after the substrate has bound. This ensures an even tighter fit and more precise bonding. This has lead to another hypothesis, the induced fit model, which explains that the contact of the substrate with the active site induces the enzyme to change shape. Once the product is generated, it leaves the surface of the enzyme, which turns back to its original shape.

Because enzymes are very specific in their activity, they are also very sensitive to changes in the environment, and require specific conditions to functions. Factors such as temperature, pH, concentration of the enzyme, and the concentration of the substrate all affect the rate of the reaction.


One gene–one enzyme hypothesis

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One gene–one enzyme hypothesis, idea advanced in the early 1940s that each gene controls the synthesis or activity of a single enzyme. The concept, which united the fields of genetics and biochemistry, was proposed by American geneticist George Wells Beadle and American biochemist Edward L. Tatum, who conducted their studies in the mold Neurospora crassa. Their experiments involved first exposing the mold to mutation-inducing X-rays and then culturing it in a minimal growth medium that contained only the basic nutrients that the wild-type, or nonmutated, strain of mold needed to survive. They found that the mutant strains of mold required the addition of specific amino acids to the minimal medium in order to grow. Using this information, the researchers were able to associate mutations in specific genes to the disruption of individual enzymes in the metabolic pathways that normally produced the missing amino acids. This discovery won Beadle and Tatum the 1958 Nobel Prize for Physiology or Medicine (shared with American geneticist Joshua Lederberg).

Although the hypothesis was amply verified in principle, it has undergone considerable sophistication since the 1940s. Today it is known that not all genes encode an enzyme and that some enzymes are made up of several short polypeptides encoded by two or more genes.

This article was most recently revised and updated by Kara Rogers, Senior Editor.


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