Gel filtration chromatography is a convenient method of resolving proteins since separation is usually good and the samples can be easily collect. It works by having two phases. The mobile liquid phase flows down the column and the stationary phase is composed of porous gel beads. The beads are often made of cross-linked dextran (sephadexTM) and their pores can be of different sizes so there is a certain amount of control over which macromolecules can flow down the column quickly and which are hindered. Different size molecules can enter the beads to different extents and therefore they are slowed to different rates. In this way, large molecules are excluded from most of the column and therefore move rapidly down. The molecular mass of the smallest molecule unable to enter the pores of the gel beads is called the exclusion limit of that gel. A typical elution might look like as shown below:
The left-hand peak is due to the largest molecules and hence the right-hand peak is due to the smallest molecules. Therefore each substance has its own elution volume, Ve, and therefore the ratio Ve/Vo (where Vo is the void volume, the elution volume molecules above the exclusion limit emerge) is the relative elution volume and is specific for a solute. You can also estimate the molecular mass of an unknown substance by finding its position on a graph of Ve/Vo against log(molecular mass) for various known substances.
Ion exchange chromatography separates substances on the basis of their charge and therefore is useful when resolving proteins since they often contain charged amino acid side chains. In this process, anion and cation exchangers are used which can be strong or weak acids or bases. For example, DEAE-cellulose (diethylaminoethyl) is a strong base whereas CM-cellulose (carboxymethyl) is a weak acid. These anion/cation exchangers are attached to the side of the chromatography column and initially are bound by electrostatic forces to counterions. When proteins are added, they bind to the ion exchanger and displace the counterion. DEAE-cellulose has Cl- counterions and therefore it is a protein that under the conditions has a single negative charge that will displace the counterion, hence exchange will occur. The chloride ions in this instance will be eluted away with the buffer, which itself is very important in the process, as pH fluctuations should be eliminated as they can influence the charge on the protein. However the buffer should not be so strong that the ionic strength is too high and the protein precipitates, therefore losing adsorbent capacity. Common buffers are phosphate (for cation exchangers) and tris (for anion exchangers). The choice of ion exchanger depends on the properties of the protein required. The figure shows that if a protein is stable in the pH range 6 to 8 then an anion exchanger has to be used since the protein has an overall negative charge as the pH is more basic than the protein’s pI. The separation can be made even more effective if a gradient elution is carried out where the pH of the buffer is continuously varied so that proteins are released at intervals. When they are eluted depends on the distance in pH between the buffer and the pI of that protein since the protein’s charge is more stable the further from its isoelectric point.
Affinity chromatography depends on the protein either being or having a natural ligand to which it binds. This ligand is immobilised and attached to a column, hence as the protein passes through it is tightly bound to the side. The bond is non-covalent yet usually very strong and one of the main advantages of this technique is that it can be extremely specific indeed since the process relies on the biochemical properties of the ligand rather than any physical ones. However the chromatographic column has to be carefully prepared in the first place and ligands must be first covalently bound to an inert matrix which lines the column. One of the best materials for this purpose is agarose, which possesses many free hydroxyl groups to bond with the ligand. Otherwise, if the ligand has a free primary amino group then this can be attached to the agarose by ‘activating’ the agarose with cyanogen bromide. When the ligand is attached to the agarose then the protein soluton can be run through the column. It is essential that the ligand can bind to the protein with sufficient affinity but also release it to allow it to continue down the tube. After elution of the mixture, any protein remaining bound to the column can be obtained by perhaps eluting the column with a substance that has a higher affinity for the protein than the agarose-bound ligand. Alternatively the conditions such as pH, ionic strength and temperature can be changed thus forcing the protein to be released. This technique was essential in the discovery of many proteins and significantly the opiate receptor protein whose presence was confirmed by Pert and Snyder. They used radioactively labelled naloxone (a very potent opiate) and having extracted the cell membrane proteins they ran them through an affinity column and found that the opiate receptor proteins attached themselves to the naloxone. This was a key step leading to the discovery of the bodies own opiates, endorphins, and more specifically, enkephalin. A further development of affinity chromatography are immunoaffinity techniques which cross-link monoclonal antibodies to the column material. This can provide startling levels of specificity although there are several disadvantages. One is that the binding is so strong that the protein can sometimes be denatured by the harsh conditions required to remove it from the column. Also it is usually very difficult to make the antibodies specific to the protein that needs to be purified.
The final technique of protein purification is gel electrophoresis. One of the key components is the gel used because it is the size of its pores that determine the speed of movement through the gel and also the efficiency of separation. There are two main gels in use, polyacrylamide (therefore used in PAGE) and agarose. The gel can be cast in a tube shape but often it is a slab in which several samples can be analysed in different lanes of the gel. The buffer used with proteins is usually around pH 9 so the samples have a net negative charge. In this way they will migrate to the anode of the gel. A current of usually 300v is then applied to achieve the separation into discrete bands. Unlike gel filtration, it is the large molecules that are retarded most and therefore these will travel the shortest distance along the gel.
After the separation is complete the proteins can be detected by staining by, for instance, Coomassie brilliant blue. Another method of detection is possible if an antibody to the protein is available and is the Western blot and usually gives very accurate results. Any proteins on the gel can be separated easily by simply cutting the protein band out of the gel after separation. The gel portion can then be easily removed by centrifugation or some other method to leave the pure protein. A slight variation on the standard gel electrophoresis technique is SDS-PAGE, which uses sodium dodecyl sulphate – a detergent. This binds to proteins and because it has a large negative charge itself, masks the effect of the protein’s charge during electrophoresis. In this way, the proteins have similar shapes and charges, so separation is based on molecular masses only. This is a useful way of determining molecular masses by using known marker proteins as well.
How would you devise a strategy for the purification of a protein?
Because there are so many purification techniques available to the scientist, the main factors to consider are the properties of the protein since these will have a major bearing on the effectiveness of the purification. However you first have to obtain the desired protein from a suitable source. In general, you will be looking for a specific protein and it is important to try and find a source that has the highest possible concentration of that protein. For instance, haemoglobin is of course found in erythrocytes and respiratory enzymes in the mitochondria. The method of obtaining at least an impure solution of the protein depends on its source. For animal cell cytosolic proteins, cell lysis is sufficient. For membrane proteins, acetone or toluene extraction is effective. There are more vigorous methods depending on the type of cell, for instance, the French Press forces cells through a small orifice at very high pressure so as to break up plant cells. Ultrasonication disrupts prokaryotic cells very effectively by using high-pressure sound waves. Therefore an intracellular protein firstly requires disruption and solubilisation. These processes need to be high capacity, high yield, low cost and low resolution, since they don’t need to separate the proteins efficiently, just remove the basic background matter. In general assays can be enzymatic, immunological or physical. Extracellular proteins are usually more resistant to harsh conditions and therefore can be more flexible in the methods used to purify them. If many purification steps are used, which is often the case, another very important factor to consider is the order and nature of the steps. For instance, salting out of the proteins cannot then be followed by an ion exchange step because the high ionic strength of the protein solution will interfere and prevent adsorption during the ion exchange step. Also ion exchange should not really be followed directly by gel filtration, as the fraction would need concentrating
Therefore when you devise the strategy of purification, you have to consider not only order of the steps but also capacities of each step. Precipitation methods are often best for the first step since they can handle as large a capacity as you want. Gel filtration capacity depends on the size of the column but can usually handle moderate capacities. Electrophoresis is usually a low capacity operation as is affinity chromatography if your ligands are spread out somewhat. It is important to note that a solution of high salt concentration is not necessarily a disadvantage as this can be altered by dialysis in pure water. With a couple of steps and a bit of time, a solution can easily be ready for ion exchange chromatography. Of course you must also take the properties of the protein into account to prevent its denaturation or inactivation of the active site. This is a subjective matter though and is difficult to discuss further.
