DiaPharma Group, Inc.

Basic Principles

What is a chromogenic substrate?

Enzymes are proteins that catalyze most of the chemical reactions that take place in the body. They make it possible for chemical reactions to occur at neutral pH and body temperature. The chemical compound upon which the enzyme exerts its catalytic activity is called a substrate. Proteolytic enzymes act on their natural substrates, proteins and peptides by hydrolyzing one or more peptide bond(s). This process is usually highly specific in the sense that only peptide bonds adjacent to certain amino acids are cleaved.

Chromogenic substrates are peptides that react with proteolytic enzymes under the formation of color. They are made synthetically and are designed to possess a selectivity similar to that of the natural substrate for the enzyme. Attached to the peptide part of the chromogenic substrate is a chemical group which when released after the enzyme cleavage gives rise to color. The color change can be followed spectrophotometrically and is proportional to the proteolytic activity.

The chromogenic substrate technology was developed in the early 1970s, and has since then become a tool of substantial importance in basic research. The majority of chromogenic substrate applications are found in various clinical fields. In particular they have been used to generate fundamental knowledge of the mechanisms regulating blood coagulation and fibrinolysis. Furthermore, products based on chromogenic substrate technology have brought a new generation of diagnostics into the clinical laboratory.

Proteolytic enzymes

In the living organism, proteolytic enzymes (proteases) are produced to degrade and modify proteins. A main task for proteolytic enzymes is to degrade proteins into peptides or amino acids to be used either as an energy source or as building blocks for resynthesis of proteins. Furthermore, proteolytic enzymes modify cellular environments and facilitate cell migration in connection with wound repair and cancer, ovulation and implantation of the fertilized egg, embryonic morphogenesis, and involution of mammary glands after lactation.

Another important function of the proteases is their role as regulators in processes such as inflammation, infection and blood clotting. Most proteolytic enzymes are highly specific for their substrates. The classification of proteases, however, is not based on their choice of substrate but on their mechanism of action.

Four different groups of proteolytic enzymes, named after the active site amino acid residue responsible for the catalytic activity, are generally distinguished: the aspartic proteases (e.g. pepsin), the cystein proteases (e.g. cathepsin B and cathepsin H), the serine proteases (e.g. trypsin, thrombin and plasmin) and metalloproteases (e.g. collagenases and gelatinases).Although the members of each group of proteolytic enzymes may have very diverse biological functions, amino acid analysis often shows a high degree of structural similarity between them. Detailed knowledge of the structure and mechanism of action of one enzyme can in many cases reveal an understanding of the structure and functions of other enzymes within the same group.

Classes of proteases
Name	active site
serine proteases	Ser His Asp*
cystein proteases	Cys His Asp*
aspartic proteases	Asp Asp
metallo proteases	His His Zn2+

*Asp not always present

Serine proteases

The most extensively studied group of proteolytic enzymes comprises the serine proteases. As indicated by the name each member of this group have a reactive seryl amino acid residue in its active site. The serine proteases are divided into two families: the trypsins and the subtilisins.

The trypsin family is the largest and contains, among others, trypsin and chymotrypsin, elastase, mast cell tryptase, and many of the factors regulating blood coagulation and fibrinolysis. The trypsin type of enzymes have a highly similar amino acid content. They are found in vertebrates and other animals, as well as in fungi and procaryotic cells. In contrast, the subtilisins are only found in bacteria. Members of the trypsin family are classified according to the type of amino acid that occurs at the preferred cleavage site.

Elastase and chymotrypsin cleave after hydrophobic and aromatic amino acids, while other trypsin-like proteases cleave only at the C-terminal side of the basic amino acids arginine or lysine. The amino acid sequence and thus also the three-dimensional structure differ completely between the trypsins and the subtilisins. The catalytically active domains of trypsin and subtilisin have therefore most probably evolved independently, converging from two different genes.

However, since the three amino acids of functional importance at the active sites, serine (Ser), aspartic acid (Asp) and histidine (His), are arranged in the same geometrical relationship in all members of the two families the proteolytic mechanisms are very similar.

This fact may lead to the suggestion that the arrangement of the three catalytically active amino acids at the active site is very efficient for hydrolysis of peptide bonds. Mammalian serine proteases are usually synthesized as inactive proenzymes, zymogens, consisting of a single peptide chain. Activation occurs when the zymogen is cleaved at one or several specific sites. Most commonly such cleavage is accomplished by the action of another protease. Most serine proteases contain two functionally distinct parts. The region where the catalytically active amino acids are found is very similar in trypsin and chymotrypsin as well as in the serine proteases involved in blood coagulation. The other region is located in the exterior parts of the enzyme. This region is of considerable size in the serine proteases regulating blood coagulation and fibrinolysis and four main types of structures can be distinguished: kringle domains, growth factor domains, vitamin K dependent carboxylated calcium binding domains, and domains homologous to the finger structure of fibronectin.

All four domain types are not present in all groups of serine proteases.

In the living organism, proteolytic enzymes (proteases) are produced to degrade and modify proteins. A main task for proteolytic enzymes is to degrade proteins into peptides or amino acids to be used either as an energy source or as building blocks for resynthesis of proteins. Furthermore, proteolytic enzymes modify cellular environments and facilitate cell migration in connection with wound repair and cancer, ovulation and implantation of the fertilized egg, embryonic morphogenesis, and involution of mammary glands after lactation.

Another important function of the proteases is their role as regulators in processes such as inflammation, infection and blood clotting. Most proteolytic enzymes are highly specific for their substrates. The classification of proteases, however, is not based on their choice of substrate but on their mechanism of action.

Four different groups of proteolytic enzymes, named after the active site amino acid residue responsible for the catalytic activity, are generally distinguished: the aspartic proteases (e.g. pepsin), the cystein proteases (e.g. cathepsin B and cathepsin H), the serine proteases (e.g. trypsin, thrombin and plasmin) and metalloproteases (e.g. collagenases and gelatinases).Although the members of each group of proteolytic enzymes may have very diverse biological functions, amino acid analysis often shows a high degree of structural similarity between them. Detailed knowledge of the structure and mechanism of action of one enzyme can in many cases reveal an understanding of the structure and functions of other enzymes within the same group.

The proteolytic mechanism of serine proteases

The bond-cleaving reaction exerted by a serine protease on its substrate is the result of an interaction between the substrate and the charge relay network of the enzyme. This network, which is present in the active site of all serine proteases, is known as the catalytic triad. It is built up from the side-chains of three specific amino acids (the hydroxy group of serine, the imidazole group of histidine and the carboxylic acid group of aspartic acid) that interact with each other through an array of hydrogen bonds.

The proteolytic action of a serine protease on its substrate comprises several steps starting with the formation of a non-covalent complex between the enzyme and the substrate. A nucleophilic attack by the serine hydroxyl group on the amide carbonyl carbon atom in the substrate results in cleavage of the amide bond and the formation of an acyl-enzyme intermediate.

The acyl-enzyme ester bond is then hydrolysed in the rate limiting step and the enzyme is now free to catalyze the cleavage of another substrate molecule.

Enzyme specificity and substrate selectivity

Specificity is a property of the enzyme and describes how restrictive the enzyme is in its choice of substrate; a completely specific enzyme would have only one substrate. The specificity of the serine proteases is usually not very high since they have similar active sites and act through the same proteolytic mechanism. Consequently, a single serine protease may act on various substrates although at different rates. How the substrate fits the active site of the enzyme is of crucial importance to the outcome of the enzyme-substrate reaction. The bond to be cleaved must have a specific orientation relative to the amino acid side chains of the catalytic triad. The most important factor governing the fit of a substrate for an enzyme is the amino acid sequence around the bond to be cleaved.

Trypsin cleaves amides and esters of the basic amino acids arginine and lysine. Thrombin has a similar preference, but is more specific for arginine than for lysine. Selectivity is a property of the substrate and indicates the degree to which the substrate is bound to and cleaved by different enzymes. The best measure for selectivity is given by the ratio kcat/Km. Synthetic substrates are considerably smaller than the natural substrates and can usually be cleaved by more than one enzyme, i. e. synthetic substrates are not completely selective. The explanation for this is that large substrates such as fibrinogen not only interact with the active site but also with exterior domains of the enzyme. Such interactions allow substrates to discriminate between different serine proteases and fibrinogen thus becomes highly selective for thrombin.

Enzyme kinetics

An enzyme acts as a catalyst for a certain chemical reaction. This means that the enzyme decreases the activation energy and reaction time for the reaction without changing the equilibrium. The enzyme is neither consumed nor modified in the reaction. An enzyme-catalyzed reaction starts with the formation of an enzyme-substrate complex, ES.

The complex has two possible outcomes. It can dissociate to free enzyme (E) and substrate (S) with a rate constant k2 , or it can proceed to form product (P) with a rate constant k3 . At a fixed concentration of the enzyme the product is formed at a rate linearly proportional to the substrate concentration. However, after saturation of the active site of each enzyme molecule with its substrate the rate of product formation is independent of the substrate concentration, [S]. The equilibrium in (1) can be pushed towards product formation by increasing the substrate concentration.

(1)

The rate of product formation depends on the concentration of enzyme-substrate complex.

(2)

The rates of formation and consumption of ES can be written as:

rate of formation of ES

(3)

rate of consumption of ES

(4)

At steady state, [ES] is constant and the rates in (3) and (4) are equal:

(5)

Rearrangement of (5) gives an expression for [ES] at steady state:

(6)

The Michaelis constant, Km, is defined below:

(7)

The concentration of free enzyme [E] equals the total enzyme concentration minus the concentration of enzyme-substrate complex:

(8)

Substitution of (7) and (8) into (6) gives:

(9)

Substituting this expression for [ES] in (2) gives:

(10)

The reaction reaches its maximal rate, Vmax, when all active sites of the enzyme are occupied, i.e. when [S] is much larger than Km, which means that [S]/[S]+Km approaches 1.

Thus,

(11)

these conditions product molecules are formed at a constant and maximal velocity. Equation (11) can also be written as:

(12)

k3 reflects the turnover number, which is the maximal number of substrate molecules that can be converted to product per time unit, dimension time^-1.

The Michaelis-Menten equation (13) is obtained from (10) and (11) and explains the relationship between reaction rate and substrate concentration as shown in Figure 5.

(13)

At a low substrate concentration ([S] << Km), then

(14)

i.e. the rate is directly proportional to [S].

This means that an increase in substrate concentration will cause an increase in the reaction rate. On the other hand, at a high substrate concentration ([S] >> Km), then

(15)

i.e. the reaction rate is independent of [S].

A special situation can be identified when the substrate concentration equals Km ([S] = Km); then

(16)

which means that the Michaelis constant Km is equal to the substrate concentration at which the reaction rate is exactly half the maximum rate or Vmax/2.

The Michaelis-Menten equation (13) is transformed into an equation giving a straight line plot by taking the reciprocal of both sides of the equation.

(17)

A plot of 1/V versus 1/[S] is called the Lineweaver-Burk plot or double-reciprocal plot, and has the intercept 1/Vmax and the slope Km/Vmax. Thus, the kinetic parameters Km and Vmax are readily derived by measuring the rate of catalysis at different substrate concentrations.

Multiplication of both sides of equation (17) with the substrate concentration [S] gives

(18)

The plot of [S]/V versus [S] is sometimes referred to as the Hanes plot. The advantage of the Hanes plot over the Lineweaver-Burk plot is that the experimental errors in V give in the former a more or less constant contribution over a wide range of [S] values (15). Consequently, more accurate results are obtained using the Hanes procedure for data treatment.

Chromogenic substrates in practice

Measurements made using chromogenic substrates reflect enzyme activity. Often it is more important to have knowledge about the activity of an enzyme than of the amount or mass - the quantity recorded in an immunological assay. Synthetic substrates are very sensitive, i.e. they can detect very low enzyme activities. They are in fact often more sensitive than a corresponding natural substrate.

This ability of chromogenic substrates to detect low enzyme concentrations makes them useful in, for example, the search for the presence of certain enzyme activities either in research or in quality control procedures. Sometimes there is a lack in correspondence between a natural and a chromogenic substrate in their responses to a certain enzyme preparation. For example, thrombin that has been partly degraded through autohydrolysis (ß-thrombin) reacts just as well with its chromogenic substrate as does the native form of thrombin (a-thrombin) while only native thrombin reacts with the natural substrate fibrinogen.

A chromogenic substrate is less selective, i.e. it has less discrimination in its reactivity towards related enzymes compared to the natural substrate. However, this lack of absolute selectivity can be compensated for when setting up chromogenic substrate assays. This is done by the proper selection of type of buffer, pH, relative concentrations of sample and reagents, addition of inhibitors, and/or choice of activator or incubation times. When presented with the opportunity of using one or more chromogenic substrates in a particular experimental setting for which there is no existing method, there are a few considerations that are worthwhile to make.

Substrate
If the specificity of the enzymatic activity to be measure-red is known then comprehensive overviews such as the Selectivity Tables will serve as a guide in selecting a proper substrate. The local distributor of Chromogenix products may also be contacted for advice on the choice of substrate(s). If the specificity of the enzyme is unknown, a screening procedure can be applied. When different substrates are available, such screening of the enzyme specificity can be carried out by comparing the rate of hydrolysis or pNA-generation obtained with the different substrates. Unless certain experience is available to the investigators it is usually advisable to discuss the plan and/or the result with Chromogenix. Advice on the next step can thus be given concerning either continued screening or the selection of a particular substrate that is suitable in the planned investigation.

Contaminating enzymes
If the sample to be tested with a chromogenic substrate contains more than one enzyme that may react with the same substrate, there are a number of measures that can be taken in order to eliminate the interfering/ contaminating activity. A natural or synthetic inhibitor can be introduced, the sample can be further diluted or conditions can be found (different pH and/or buffer) where the relative activities of the present enzymes are optimized. Such considerations can be based on the information below concerning temperature, pH, buffer and ionic strength.

Temperature
The rate by which the chromogenic substrate is cleaved is highly dependent on the temperature. It is therefore important to know at what temperature(s) a particular method is applicable - it may be at room (ambient) temperature, 25, 30, or 37 °C. An increase in temperature of 1 °C causes an increase in the reaction velocity of 2.5-7.5%. The temperature thus must be kept constant during the measurement and if results from different experiments are to be compared they must be performed at the same temperature. It is advisable to run the reactions in thermostated cuvettes and to use preheated stock solutions.

pH
Both Km and kcat are dependent on the pH. This means that kinetic calculations can only be made using results obtained at the same pH. Usually, the enzyme activity is measured at the pH optimum for the proteolytic activity of the enzyme. However, when several proteases are present in the same solution, as, e.g. when the sample is from plasma, it is not always advantageous to search for the pH that gives the maximum reactivity of the enzyme under investigation. Instead it is better to choose a pH where other serine proteases that may compete for the substrate have relatively lower levels of activity.

Buffers
The buffer medium and the concentration of buffer substances must be well defined. Usually Tris-HCl is used since the pKa of Tris buffer is 8.1 (25 °C), which makes it suitable for measurements at pH values between 7.3-9.3, where most of the serine proteases show maximal activities. Furthermore, this buffer is stable - it can even be autoclaved. Tris-imidazole has also been used, but is not to be recommended as imidazole is known to slightly inhibit certain proteases such as trypsin and plasmin.

Ionic strength and other additives
The appropriate ionic strength is usually obtained by adjusting the concentration of NaCl. Further substances that it may be necessary to add are CaCl2 (when Ca-dependent enzymes are studied), NaN3 (or other bactericidal agents) to prevent bacterial growth and polyethylene glycol or Tween 80 to prevent adsorption of the enzymes to the reaction vessel walls.

Substrate handling
The substrate solution is usually prepared by adding sterile water to the dry powder. Substrates with low solubility in water can be dissolved in DMSO (dimethyl sulfoxide) and then diluted in water. The final DMSO concentration should preferably not exceed 10% in the reaction mixture. Substrates dissolved in sterile water are stable for more than 6 months in the refrigerator (2 - 8 °C) and for several weeks at room temperature (25 °C). The stability is considerably reduced in alkaline buffers. Furthermore, contamination by microorganisms and exposure to light for longer periods should be avoided. The substrate concentration should be chosen so that linear kinetics is obtained. A substrate concentration of twice the Km (2 x Km ) is usually appropriate.