Polyacrylamide gel electrophoresis (PAGE) is used for the qualitative characterization of proteins in biological preparations, for control of purity and for quantitative determinations.

Analytical gel electrophoresis is an appropriate method with which to identify and to assess the homogeneity of proteins in drug substances. The method is routinely used for the estimation of protein subunit molecular masses and for the determination of the subunit compositions of purified proteins.

Ready-to-use gels and reagents are widely available on the market and can be used instead of those described in this chapter, provided that they give equivalent results and that they meet the validity requirements given below under Validation of the Test.

This method cited is limited to the analysis of monomeric polypeptides with a mass range of 14,000 to 100,000 Da. It is possible to extend the mass range by various techniques (e.g., gradient gels or particular buffer systems), but those techniques are not discussed in this chapter.

Denaturing PAGE using sodium dodecyl sulfate (SDS) (SDS-PAGE) is the most common mode of electrophoresis used in assessing the pharmaceutical quality of protein products and will be the focus of the example method. Typically, analytical electrophoresis of proteins is carried out in polyacrylamide gels under conditions that ensure dissociation of the proteins into their individual polypeptide subunits and that minimize aggregation. Most commonly, the strongly anionic detergent SDS is used in combination with heat to dissociate the proteins before they are loaded on the gel. The denatured polypeptides bind to SDS, become negatively charged, and exhibit a consistent charge-to-mass ratio regardless of protein type. Because the amount of SDS bound is almost always proportional to the molecular mass of the polypeptide and is independent of its sequence, SDS-polypeptide complexes migrate through polyacrylamide gels with mobilities dependent on the size of the polypeptide.

The electrophoretic mobilities of the resultant detergent–polypeptide complexes all assume the same functional relationship to their molecular masses. Migration of SDS complexes is toward the anode in a predictable manner, with low molecular mass complexes migrating faster than larger ones. The molecular mass of a protein can therefore be estimated from its relative mobility in calibrated SDS-PAGE and the occurrence of a single band in such a gel is a criterion of purity.

Modifications to the polypeptide backbone, such as N- or O-linked glycosylation, however, have a significant impact on the apparent molecular mass of a protein, because SDS does not bind to a carbohydrate moiety in a manner similar to the polypeptide. Thus, a consistent charge-to-mass ratio is not maintained. The apparent molecular mass of proteins having undergone post-translational modifications is not a true reflection of the mass of the polypeptide chain.

Polypeptide subunits and three-dimensional structure is often maintained in proteins by the presence of disulfide bonds. A goal of SDS-PAGE analysis under reducing conditions is to disrupt this structure by reducing disulfide bonds. Complete denaturation and dissociation of proteins by treatment with 2-mercaptoethanol or dithiothreitol (DTT) will result in the unfolding of the polypeptide backbone and subsequent complexation with SDS. In these conditions, the molecular mass of the polypeptide subunits can be calculated by linear regression in the presence of suitable molecular mass standards.

For some analyses, complete dissociation of protein into subunit peptides is not desirable. In the absence of treatment with reducing agents such as 2-mercaptoethanol or DTT, disulfide covalent bonds remain intact, preserving the oligomeric form of the protein. Oligomeric SDS-protein complexes migrate more slowly than their SDS-polypeptide subunits. In addition, nonreduced proteins may not be completely saturated with SDS and hence may not bind the detergent in a constant mass ratio. This makes molecular mass determinations of these molecules less straightforward than analyses of fully denatured polypeptides, because it is necessary that both standards and unknown proteins be in similar configurations for valid comparisons. However, the staining of a single band in such a gel is a criterion of purity.

The most popular electrophoretic method for the characterization of a complex mixture of proteins involves the use of a discontinuous buffer system consisting of two contiguous, but distinct, gels: a resolving or separating (lower) gel and a stacking (upper) gel. The two gels are cast with different porosities, pHs, and ionic strengths. In addition, different mobile ions are used in the gel and electrode buffers. The buffer discontinuity concentrates large volumes of sample in the stacking gel, resulting in improved resolution. When power is applied, a voltage drop develops across the sample solution that drives the proteins into the stacking gel. Glycinate ions from the electrode buffer follow the proteins into the stacking gel. A moving boundary region is rapidly formed with the highly mobile chloride ions in the front and the relatively slow glycinate ions in the rear. A localized high-voltage gradient forms between the leading and trailing ion fronts, causing the SDS-protein complexes to form into a thin zone (stack) and migrate between the chloride and glycinate phases. Within broad limits, regardless of the height of the applied sample, all SDS proteins condense into a very narrow region and enter the resolving gel as a well-defined, thin zone of high protein density. The large-pore stacking gel does not retard the migration of most proteins and serves mainly as an anticonvective medium. At the interface between the stacking and resolving gels, the proteins experience a sharp retardation due to the restrictive pore size of the resolving gel. Once in the resolving gel, proteins continue to be slowed by the sieving of the matrix. The glycinate ions overtake the proteins, which then move in a space of uniform pH formed by tris(hydroxymethyl)aminomethane (Tris) and glycine. Molecular sieving causes the SDS-polypeptide complexes to separate on the basis of their molecular masses.

Coomassie staining is the most common protein staining method, with a detection level on the order of 1 to 10 µg of protein per band. Silver staining is the most sensitive method for staining proteins in gels and a band containing 10 to 100 ng can be detected.

All of the steps in gel staining are done at room temperature with gentle shaking (e.g., on an orbital shaker platform). Gloves must be worn when staining the gels, because fingertips will stain.

Depending on the method used, the gels are treated in a slightly different way. For Coomassie staining, after the destaining step, allow the gel to stand in a 100 g per L glycerol solution for at least 2 hours. For silver staining, add to the final rinsing step a 5-minute incubation in a 20 g per L glycerol solution.

Immerse two sheets of porous cellulose film in water, and incubate for 5 to 10 minutes. Place one of the sheets on a drying frame. Carefully lift the gel, and place it on the cellulose film. Remove any trapped air bubbles, and pour a few mL of water around the edges of the gel. Place the second sheet on top, and remove any trapped air bubbles. Complete the assembly of the drying frame. Place in an oven, and leave at room temperature until dry.

Molecular masses of proteins are determined by comparison of their mobilities with those of several marker proteins of known molecular mass. Mixtures of proteins with precisely known molecular masses blended for uniform staining are available for calibrating gels. They are available in various molecular mass ranges. Concentrated stock solutions of proteins of known molecular mass are diluted in an appropriate sample buffer and loaded on the same gel as the protein sample to be studied.

Immediately after the gel has been run, the position of the bromophenol blue tracking dye is marked to identify the leading edge of the electrophoretic ion front. This can be done by cutting notches in the edges of the gel or by inserting a needle soaked in India ink into the gel at the dye front. After staining, measure the migration distances of each protein band (markers and unknowns) from the top of the Resolving Gel. Divide the migration distance of each protein by the distance traveled by the tracking dye. The normalized migration distances so obtained are called the relative mobilities of the proteins (relative to the dye front) and conventionally denoted as RF. Construct a plot of the logarithm of the molecular masses (MR) of the protein standards as functions of the RF values. Note that the graphs are slightly sigmoid. Unknown molecular masses can be estimated by linear regression analysis of interpolation from the curves of log MR against RF as long as the values obtained for the unknown samples are positioned along the linear part of the graph.

The test is not valid unless the proteins of the molecular mass marker are distributed along 80% of the length of the gel and over the required separation range (e.g., the range covering the product and its dimer or the products and its related impurities); the separation obtained for the relevant protein bands shows a linear relationship between the logarithm of the molecular mass and the RF. Additional validation requirements with respect to the solution under test may be specified in individual monographs.

Where the impurity limit is specified in the individual monograph, a reference solution corresponding to that level of impurity should be prepared by diluting the test solution. For example, where the limit is 5.0%, a reference solution would be a 1:20 dilution of the test solution. No impurity—any band other than the main band—in the electropherogram obtained from the test solution may be more intense than the main band obtained with the reference solution.

Under validated conditions impurities may be quantified by normalization to the main band, using an integrating densitometer. In this case, the responses must be validated for linearity.