INSTRUCTOR: SDS-PAGE is a technique developed to separate proteins according to their relative molecular mass. Proteins can have a net positive or negative charge depending on their amino acid composition and the pH of the solution that they are in. In SDS-PAGE, sodium dodecyl sulfate – SDS, a negatively charged detergent – is used to denature and solubilise proteins.
By heating proteins briefly in the presence of SDS, they are unfolded, and the detergent coats the polypeptides. In this way, the protein is denatured. Because SDS is negatively charged, and many SDS molecules bind to a single protein molecule, the detergent effectively masks the protein's intrinsic charge and gives it a net negative charge proportional to its linear size.
Multi-subunit proteins have more than one polypeptide chain, or subunit, and the noncovalent interactions that hold subunits together in a multi-subunit protein are disrupted by SDS. Some proteins have disulfide bridges between or within subunits. These covalent linkages can be reduced by addition of a reducing agent such as beta-mercaptoethanol or dithiothreitol to protein samples.
The result is a mixture of discrete denatured polypeptides which can be analysed by electrophoresis. Analysis of a protein by SDS-PAGE under reducing conditions and under non-reducing conditions – that is, in the absence of a reducing agent – can therefore give us information on its structure and subunit composition.
A polyacrylamide gel is used as an inert matrix through which the denatured proteins move in response to an electric field. The gel is prepared by polymerisation and cross-linking of acrylamide monomers. The polyacrylamide forms a kind of sieve or three-dimensional mesh through which small molecules can move more freely than large molecules.
When the proteins are applied to or loaded onto a polyacrylamide gel, and an electric field applied across the gel, the denatured polypeptides move towards the anode at a rate dependent on their relative molecular mass, with small proteins moving faster than large proteins. Thus, in a given time the small proteins move further from the origin than do the large proteins.
The pore size of the gel can be varied by using different concentrations of acrylamide. Thus, higher concentrations produce smaller pore sizes and denser gels, as shown in the schematic. Gels ranging from 7% to 15% acrylamide are routinely used, and the acrylamide concentration is chosen to maximise separation of the proteins of interest according to their Mr values.
This animation illustrates the effect of changing the acrylamide concentration. The bands are visible here. But remember, in a real experiment the bands are not visible until after electrophoresis is complete and the gel has been stained. At the low acrylamide concentration of 7%, you can see that the larger proteins near the top of the gel are well separated. But the smaller proteins near the bottom of the gel are not well separated.
At the high acrylamide concentration of 15%, the effect is reversed. The larger proteins remain close together at the top of the gel, while the smaller proteins further down the gel are well separated. Thus, large polypeptides can readily enter a low-percentage acrylamide gel, and are fairly well resolved.
However, such a gel will not retard small polypeptides sufficiently to give good resolution. A very high-percentage acrylamide will restrict all proteins, but very large proteins will be concentrated near the top of the gel and will not be well resolved.
The figure shows a schematic of the electrophoresis setup. The polyacrylamide gel is arranged vertically in a gel tank between two separate reservoirs of buffer. The reservoir contacting the top of the gel where the samples are loaded contains the cathode, whilst the bottom of the gel is dipped in a reservoir containing the anode. When the electric field is applied between these two electrodes, the negatively charged SDS protein complexes move through the gel in the direction of the anode.
After electrophoresis, proteins can be visualised by staining with a dye such as Coomassie blue. If a mixture of protein standards of known Mr – sometimes referred to as markers – are run on a gel alongside a protein of unknown Mr, it is possible to estimate the Mr of the unknown.
To do so, it is necessary to first calculate the relative migration distance of the protein – denoted Rf. This value is given by dividing the distance the protein has travelled by the distance that the dye front has travelled. So for the protein X, the Rf is calculated by dividing the distance travelled by the protein – x prime – by the distance travelled by the dye front – d. Similarly, for protein Y, the Rf is given by dividing y prime by d.
The log of the Mr for each standard is plotted against its Rf value. As you can see from the graph, Rf for a protein is inversely proportional to the log of its Mr. Using the standard plot, it is possible to calculate the Mr of the unknown protein from its Rf value.
In practise, SDS-PAGE can only give a rough estimate of the M r of a protein. It is generally used to compare the profile or relative abundance of proteins in, for example, different cell or tissue extracts, or to follow a protein during different stages of its purification.
While SDS-PAGE can tell us something about the number of different proteins in a sample and their relative molecular masses, it does not definitively identify particular proteins. If we have an antibody which specifically recognises the protein of interest, we can perform a procedure known as Western blotting, or immunoblotting.
In the Western blot technique, the proteins in an SDS-PAGE gel are transferred to a nitrocellulose filter by application of an electric field, as shown here. An antibody against a particular protein – indicated as X – is incubated with the filter, and binds specifically to that protein. A secondary antibody which recognises the first antibody and which is conjugated to an enzyme – such as horseradish peroxidase – is incubated with the filter and binds to the primary antibody.
The presence of the enzyme – and hence the presence of protein X – is revealed by incubation of the filter with a substrate for the enzyme. The product of the reaction may, for example, be coloured, or can be luminescent.
The application of this technique can be illustrated as follows. We have five separate mixtures of proteins, and we want to find out which of the mixtures contains a known protein – protein X. We subject the five protein mixtures to SDS-PAGE on two separate identical gels with a mixture of known markers run alongside the unknown proteins.
One of the gels is stained with Coomassie blue dye to visualise all the proteins. The other gel is blotted onto a nitrocellulose filter in a Western blot procedure, and probed with an antibody specific to protein X.
We know the Mr of protein X, and from the stained gel it is evident that a protein with an Mr similar to that of protein X is present in samples one, two, four, and five. From the Western blot, we can see that protein X is only present in samples one, two, and five. And the polypeptide in lane four – though it has the same apparent Mr as X, is not in fact X.