The virtual laboratory: Enzyme purification
copyright © 1982 - 2006 David A Bender
Virtual Laboratory main menu
Click here to run the program
Each time you run this program you will see a different enzyme, with a randomly selected molecular mass and subunit composition.
There are various reasons why you might want to purify an enzyme, including:
to characterise it - determine its kinetics (including its kcat, the activity per mol of enzyme) and inhibitor sensitivity without interference from other enzymes that may compete for the substrate, remove the product or act on the inhibitor. From such studies you can begin to determine its mechanism of action;
to raise antibodies against it, in order to:
determine amounts of enzyme protein in tissues under various conditions by immunoassay;
localise the enzyme in tissues and sub-cellular compartments by immunohistological techniques;
to crystallise it, in order to determine its structure by X-ray crystallography and other techniques;
to produce an enzyme preparation that can be used industrially to catalyse just one reaction with no undesired by-products. Even when an industrially useful enzyme has been prepared by genetic modification of a micro-organism so that it is over-expressed, it is normally necessary to undertake at least a partial purification to obtain a useful product.
Enzyme purification is an art rather than a science, in that there is no way of knowing when you begin which techniques are most likely to be useful. Every protein is different, and successful purification depends on the application of different techniques, and assessing each step in the procedure for its effectiveness in removing other proteins and its efficiency of recovery of your enzyme.
In this simulation you will purify an enzyme using a variety of techniques.
At each stage of your purification you will see the volume of your sample, the enzyme activity /mL and the protein concentration; you should calculate the recovery of enzyme through each stage, and the degree of purification you have achieved to date.
To calculate the recovery of enzyme you need to note the initial activity of the enzyme (and the volume of the initial sample) - this is shown on the results screen when you perform the ammonium sulphate fractionation - and the activity of the enzyme (and sample volume) at the present stage of the purification. Simply multiply the activity /mL by the volume of the sample to obtain the total amount of enzyme, then express this as a percentage of the total activity in your original sample.
To calculate the purification of the enzyme at each stage you need to calculate its specific activity - i.e. the activity per mg protein, and express this as a multiple of the activity /mg protein in your original sample.
By the final stage you should have achieved purification to homogeneity (as determined by gel filtration chromatography). At this stage, having estimated its molecular mass by gel filtration, and knowing the amount of protein present in the sample, you can calculate the kcat (turnover number) of your enzyme. This is the activity (µmol of substrate converted / minute) divided by the amount of enzyme present in your sample (in mol, calculated from the amount of protein present divided by the molecular mass of the enzyme).
The techniques you will use are:
Ammonium sulphate fractionation
Ion exchange chromatography
Affinity chromatography
Gel filtration - to allow you to estimate the molecular mass of the enzyme
SDS polyacrylamide gel electrophoresis - this shows the individual subunits of the enzyme (if there are multiple subunits in your enzyme)
Each technique may only be used when the appropriate previous steps have been performed, but you can always go back and repeat one of the earlier steps, then carry the purification through using this new set of results.
Ammonium sulphate fractionation
Proteins are soluble in aqueous media because they have hydrophilic amino acid side-chains facing outwards that can interact with water. These are provided by the basic amino acids (arginine, histidine, arginine and lysine), the acidic amino acids (aspartate and glutamate) and the neutral hydrophilic amino acids (asparagine, glutamine, serine, threonine, tyrosine and cysteine).
Any compound that interferes with these interactions between amino acid side-chains and water, by reducing the available water, will reduce the solubility of the protein. As interactions with water become less marked, so protein-protein interactions become more important, and the protein will aggregate and come out of solution. Provided that the temperature is maintained low enough (around 4C), the protein is not irreversibly denatured, but the precipitate can be redissolved in buffer.
A number of different methods can be used to reduce hydrophilic interactions and precipitate out proteins reasonably selectively, including:
Salting out with ammonium sulphate
Selective precipitation with an organic solvent that both reduces available water and also decreases the dielectric constant of the solution. Ethanol fractionation is commonly used to separate protein fractions from blood plasma. Historically, acetone precipitation was an important way of obtaining a dry powder that could be stored for some time without losing activity;
Selective precipitation using non-ionic polymers such as polyethylene glycol, which both reduce the available water and may also interact directly with some of the proteins in the mixture, in much the same way as do the polymers used in gel filtration.
Ammonium sulphate is highly hydrated, and a concentrated ammonium sulphate
solution reduces the available water very considerably.
The diagram on the right shows two proteins, with their hydrophilic regions coloured blue. The protein on the left has relatively few hydrophilic regions, and hence will aggregate and precipitate at a relatively low concentration of ammonium sulphate - perhaps around 20 - 30% saturation. By contrast, the protein on the right has considerably more hydrophilic regions, and hence will remain in solution until the concentration of ammonium sulphate is considerably higher - perhaps around 50 - 60% saturation.
This means that it is possible to separate proteins from a mixture on the basis of their relative hydrophilicity by gradually increasing the concentration of ammonium sulphate.
At each stage you calculate the volume of saturated ammonium sulphate solution that will be required to achieve a given percentage saturation of your enzyme preparation, which is typically a crude tissue homogenate or perhaps a high-speed supernatant of a tissue homogenate. The ice-cold saturated solution of ammonium sulphate is added slowly to the protein solution, in an ice bath, and stirred continually. When the required amount has been added, the solution is centrifuged, and the precipitate collected, and redissolved in buffer. A higher degree of saturation with ammonium sulphate is then achieved by adding more saturated ammonium sulphate solution in the same way.
Initially you would probably use a number of wide ranges of ammonium sulphate saturation, say 0 - 50% and see whether or not your enzyme is precipitated. If it is, then you can refine the range until you achieve maximum recovery of the enzyme and maximum removal of interfering proteins.
Removing the ammonium sulphate by dialysis
Having precipitated a protein fraction that contains most of your enzyme, and redissolved it in buffer, it is necessary to remove the ammonium sulphate before you can proceed to subsequent steps in the purification process. The simplest way to achieve this is to dialyse the solution.
As shown in the diagram, the enzyme solution is placed in a bag of selectively permeable membrane (e.g. cellophane), immersed in a large volume of buffer that is stirred and maintained at about 4C.The membrane has pores that will permit small molecules such as ammonium and sulphate ions to cross, and hence equilibrate in the larger volume of buffer outside, while not permitting large protein molecules to cross. If the buffer is changed several times, allowing several hours each time for the ammonium sulphate to equilibrate, more or less all of it will be removed from the protein solution.
Dialysis will increase the volume of the enzyme solution, because of the initial osmotic effect of the ammonium sulphate; (this is why it is important to leave an air gap at the top of the membrane tube, to prevent it bursting).
Removing the ammonium sulphate by gel filtration
An alternative way of removing the ammonium sulphate is by gel filtration, using e.g. Sephadex G25, which has small pores that will retard
small molecules such as ammonium and sulphate ions, but will exclude large protein
molecules, so that they are eluted in the void volume of the column. This means
that the early eluate will contain the proteins, more or less free from ammonium
sulphate.
Top of page
Techniques available
Ion exchange chromatography
Different proteins have different numbers of acidic (aspartate and glutamate) and basic (arginine, histidine and lysine) amino acid side chains exposed on the surface, and therefore at a given pH they will have different nett charges. This can be exploited as a means of separating proteins by use of chromatography on a column of an ion exchange resin. More highly charged proteins will bind to the resin more strongly, and hence be retarded in their passage through the column.
Commonly, proteins are eluted from the column by either:
changing the pH of the elution buffer, so changing the charge on the proteins;
gradually increasing the ionic strength of the buffer solution (e.g. with a sodium chloride gradient), so weakening the ionic interactions between the proteins and the ion exchange resin.
The resin used may be either a cation or an anion exchanger; those commonly used in protein purification include:
| functional group | support medium | |
| weak anion exchangers | ||
| DEAE-Sephacel | diethylethylaminoethyl | Sephacel |
| DEAE-Sephadex | diethylethylaminoethyl | Sephadex |
| PEI-cellulose | polyethyleneimine | cellulose |
| weak cation exchangers | ||
| CM-Sephacel | carboxymethyl | Sephacel |
| CM-Sephadex | carboxymethyl | Sephadex |
| Bio-Rex 70 | carboxylic acid | acrylic polymer |
In this simulation you will attempt to separate the proteins by anion exchange chromatography on DEAE-Sephadex, using a sodium chloride gradient for elution. Your first task is to decide an appropriate pH for the initial buffer. This is determined by experiment - if you choose a pH at which the enzyme elutes in the void volume of the column (i.e. it has not bound to the resin) then you should choose a higher pH, in order to ensure that it has a sufficient positive charge to bind to the reactive groups on the resin.
Top of page
Techniques available
Affinity chromatography
The principle of affinity chromatography is that the stationary phase consists of a support medium (e.g. cellulose beads) on which the substrate (or sometimes a coenzyme) has been bound covalently, in such a way that the reactive groups that are essential for enzyme binding are exposed. As the mixture of proteins is passed through the chromatography column, those proteins that have a binding site for the immobilised substrate will bind to the stationary phase, while all otter proteins will be eluted in the void volume of the column.
Once the other proteins have all been eluted, the bound enzyme(s) can be eluted in various ways:
by increasing the ionic strength of the buffer, e.g. with a sodium chloride gradient, so weakening interactions between the enzyme and the immobilised substrate
by changing the pH of the buffer, again weakening interactions between the enzyme and the immobilised substrate
by adding a high concentration of substrate (or a substrate analogue) to the elution buffer, so that there is competition between the free and immobilised substrate for the enzyme protein
Linking the substrate to the support medium
There
are several different activated agarose gels that can be used to attach ligands;
CNBr-agarose is easy to use for the attachment of amines, but does not have
a long spacer between the gel beads and the bound ligand, so that protein binding
may be sterically hindered.
Aminohexanoic acid-agarose (CH-agarose, for reaction with amines) and diaminohexane-agarose (AH-agarose, for reaction with carboxylic acids)) have relatively long methylene chains that keep the ligand a significant distance from the gel beads.
An alternative reagent for attaching amines is carbonyldiimadazole-agarose, and epoxy-activated agarose is used for alcohols.
Top of page
Techniques available
Gel filtration or gel exclusion chromatography
Proteins (and other macromolecules) can be separated by their size by chromatography on columns of beads of gel that have small pores, so that smaller molecules spend more time within the pores of the support medium, and hence move more slowly, than larger molecules. This is the technique of gel exclusion chromatography, also know as gel filtration or gel permeation chromatography. By adding a series of coloured markers of known molecular mass to the sample undergoing chromatography, it is possible to calibrate the column, and hence estimate the molecular mass of the enzyme, by comparison of its elution volume with those of the standard markers.
Large molecules travel faster, and hence elute earlier from the column.
Media for gel exclusion chromatography
The media used for gel exclusion chromatography include dextran (Sephadex™), polyacrylamide (Bio-Gel P™) and dextran-polyacrylamide (Sephacryl™) and agarose (Sepharose™ and BioGel A™). Each is available with a variety of different ranges of pore size in the beads, permitting separation of macromolecules of different size.
A gel with a smaller range of pore sizes (and hence a smaller range over which it can separate macromolecules on the basis of their size) will give a higher resolution; a gel with a wider range will give lower resolution, but will permit fractionation of a larger range of sizes as an initial step when the approximate molecular mass of the enzyme is not known.
The particle size of the gel beads (the mesh size) also affects resolution; smaller beads permit higher resolution, but a lower flow rate through the column (and hence a slower separation).
The table shows the useful range for the most commonly used gel filtration media - the lower and upper molecular sizes (in kDa) over which they can be used to separate macromolecules. The upper limit is known as the exclusion limit of the gel - the size above which proteins will elute in the void volume of the column.
| lower limit | upper limit | |
| dextran gels | ||
| Sephadex G-50 | 1.5 | 30 |
| Sephadex G-75 | 3 | 80 |
| Sephadex G-100 | 4 | 150 |
| Sephadex G-150 | 5 | 300 |
| Sephadex G-200 | 5 | 600 |
| polyacrylamide gels | ||
| Bio-Gel P-10 | 1.5 | 20 |
| Bio-Gel P-30 | 2.5 | 40 |
| Bio-Gel P-60 | 3 | 60 |
| Bio-Gel P-100 | 5 | 100 |
| Bio-Gel P-150 | 15 | 150 |
| Bio-Gel P-200 | 30 | 200 |
| Bio-Gel P-300 | 60 | 400 |
| dextran-polyacrylamide gels | ||
| Sephacryl S-200 | 5 | 250 |
| Sephacryl S-300 | 10 | 1500 |
| Sephacryl S-400 | 20 | 8000 |
| agarose gels | ||
| Sepharose 6B | 10 | 4000 |
| Sepharose 4B | 60 | 20,000 |
| Sepharose 2B | 70 | 40,000 |
| Bio-Gel A-0.5 | 10 | 500 |
| Bio-Gel A-1.5 | 10 | 1500 |
| Bio-Gel A-5 | 10 | 5000 |
| Bio-Gel A-15 | 40 | 15,000 |
| Bio-Gel A-50 | 100 | 50,000 |
Top of page
Techniques available
SDS-polyacrylamide gel electrophoresis
Electrophoresis
is a technique for separating molecules on the basis of their charge and size.
At an appropriate pH, the distance they move in an electric field depends on
both their overall charge and also their size.
The support medium for electrophoresis may be a strip of paper or cellulose acetate (as, for example when small peptides are to be separated), or a gel, such as polyacrylamide (as is commonly used for electrophoresis of proteins and nucleic acids).
If the protein is denatured with mercaptoethanol (to reduce disulphide bridges to -SH groups) and the anionic detergent sodium dodecyl sulphate (SDS), it becomes coated with a layer of the detergent, and therefore all proteins will have a similar charge. The distance they move on electrophoresis now depends on their size.
Therefore, electrophoresis on polyacrylamide gel after denaturation with SDS (known as SDS-PAGE) together with standards of known molecular mass permits determination of the molecular mass of a protein. Since the protein has been denatured, it has lost all secondary, tertiary and quaternary structure, and if the native protein consists of more than one subunit, these will be separated, and will migrate in the electric field according to their size.
Polyacrylamide gel
A polyacrylamide
gel is prepared by co-polymerising acrylamide and methylene bis-acrylamide;
the reaction is initiated by a source of radicals (typically ammonium persulphate,
but sometimes riboflavin and uv irradiation) and the catalyst tetramethylene
ethylene diamine (TEMED).
Methylene bis-acrylamide forms cross-links between the chains of polyacrylamide.
The percentage of methylene bis-acrylamide in the mixture, and hence the extent of cross-linking, determines the size of the pores in the gel, and hence the extent to which proteins of different size are retarded in the electric field.
Smaller proteins move faster.
Top of page
Techniques available