Denaturing PAGE using sodium dodecyl sulfate (SDS) is the most common mode of electrophoresis used in assessing the pharmaceutical quality of protein products. Typically, analytical electrophoresis of proteins is carried out under conditions that ensure dissociation of the proteins into their individual polypeptide subunits and that minimize aggregation of these subunits. 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 SDS, become negatively charged, and exhibit a consistent charge-to-weight ratio regardless of protein type. Because the amount of SDS bound is almost always proportional to the molecular weight of the polypeptide and is typically independent of its sequence, SDSpolypeptide complexes migrate through polyacrylamide gels in reasonable accordance with the size of the polypeptide.
Modifications to the polypeptide backbone, such as N- or O-linked glycosylation, however, have a significant impact on the apparent molecular weight of a protein. This is due to the fact that SDS does not bind to a carbohydrate moiety in a manner similar to that of the polypeptide. Thus, a consistent charge-to-weight ratio is not maintained. The apparent molecular weight of proteins having undergone post-translational modifications is not a true reflection of the weight of the polypeptide chain.
For some analyses, complete dissociation of protein to peptide subunits is not desirable. In the absence of treatment with reducing agents, disulfide covalent bonds remain intact, preserving the oligomeric form of the protein. Oligomeric SDSprotein complexes migrate more slowly than their SDSpolypeptide subunits. In addition, nonreduced proteins may not be completely saturated with SDS and hence may not bind the detergent in a constant weight ratio. This makes molecular weight determinations of these molecules less straightforward than analyses of fully denatured polypeptides, because, for valid comparisons, it is necessary that both standards and unknown proteins be in similar configurations. However, the staining of a single band in such a gel is a criterion of purity.
Characteristics of a Discontinuous Buffer System
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 SDSprotein complexes to form into a thin zone (stack) and migrate between the chloride and glycinate phases. Within a broad limit, 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 SDSpolypeptide complexes to separate on the basis of their molecular weights.
Preparation of Gels
In a discontinuous buffer SDSpolyacrylamide gel, it is important to pour the resolving gel, let the gel set, and then pour the stacking gel, because the composition of acrylamidebisacrylamide, buffer, and pH are different.
Gel Stock Solutions
30% AcrylamideBisacrylamide Solution
Prepare a solution containing 290 g of acrylamide and 10 g of methylene bisacrylamide per L of warm water, and filter. [noteAcrylamide and methylene bisacrylamide are slowly converted during storage to acrylic acid and bisacrylic acid, respectively. This deamidation reaction is catalyzed by light and alkali. The pH of the solution must be 7.0 or lower. Store the solution in dark bottles at room temperature. Fresh solutions are prepared every month.]
Ammonium Persulfate Solution
Prepare a small quantity of solution having a concentration of 100 g of ammonium persulfate per L, and store at 4
Ammonium persulfate provides the free radicals that drive polymerization of acrylamide and bisacrylamide. Ammonium persulfate decomposes slowly; therefore, prepare fresh solutions weekly.]
Use an electrophoresis-grade reagent. [noteTEMED accelerates the polymerization of acrylamide and bisacrylamide by catalyzing the formation of free radicals from ammonium persulfate. Because TEMED works only as a free base, polymerization is inhibited at low pH.]
Use an electrophoresis-grade reagent. Prepare a solution having a concentration of about 100 g of SDS per L, and store at room temperature.
1.5 M Buffer Solution
Transfer about 90.8 g of Tris to a 500-mL flask, dissolve in 400 mL of water, adjust with hydrochloric acid to a pH of 8.8, dilute with water to volume, and mix.
1 M Buffer Solution
Transfer about 60.6 g of Tris to a 500-mL flask, add 400 mL of water, adjust with hydrochloric acid to a pH of 6.8, dilute with water to volume, and mix.
Clean two glass plates (10 cm × 8 cm), the polytef comb, the two spacers, and the silicone rubber tubing (0.6 mm × 35 cm) with mild detergent, rinse thoroughly with water, and blot dry.
Lubricate the spacers and the tubing with nonsilicone grease. Apply the spacers along each of the two short sides of the glass plate 2 mm away from the edges and 2 mm away from the long side corresponding to the bottom of the gel.
Begin to lay the tubing on the glass plate by using one spacer as a guide. Carefully twist the tubing at the bottom of the spacer, and follow the long side of the glass plate. While holding the tubing with one finger along the long side, twist the tubing again, and lay it on the second short side of the glass plate, using the spacer as a guide.
Place the second glass plate in perfect alignment with the first, and hold the gel mold together by hand pressure. Apply two clamps on each of the two short sides of the mold. Carefully apply four clamps on the longer side of the mold, thus forming the bottom of the mold. Verify that the tubing is running along the edge of the glass plates and has not been extruded while placing the clamps. The mold is now ready for the pouring of the gel.
In a conical flask, prepare the appropriate volume of solution, containing the desired concentration of acrylamide, as shown in Table 1
. Mix the components in the order shown. Before adding the Ammonium Persulfate Solution
and the TEMED,
pour the solution into a disposable filtration unit equipped with a nitrocellulose filter having a 0.45-µm porosity, and apply vacuum. Allow the solution to degas by swirling the filtration unit, and disconnect the vacuum when no more bubbles are formed in the solution. Add appropriate amounts of Ammonium Persulfate Solution
as shown in Table 1;
swirl; and pour immediately into the gap between the two glass plates of the mold. Leave sufficient space for the stacking gel (the length of the teeth of the comb plus 1 cm). Using a pipet, carefully overlay the solution with water-saturated isobutyl alcohol. Leave the gel in a vertical position at room temperature for polymerization.
Table 1. Preparation of Resolving Gel
After polymerization is complete (about 30 minutes later), pour off the overlay, and wash the top of the gel several times with water to remove the isobutyl alcohol overlay and any unpolymerized acrylamide. Drain as much fluid as possible from the top of the gel, then remove any remaining water with the edge of a paper towel.
In a conical flask, prepare the appropriate volume of solution containing the desired concentration of acrylamide, as shown in Table 2
Table 2. Preparation of Stacking Gel
||Component Volume (mL) per Gel Mold Volume Below
|1.0 M Buffer Solution
|Ammonium Persulfate Solution
Mix the components in the order shown. Before adding the Ammonium Persulfate Solution
and the TEMED
, pour the solution into a disposable filtration unit equipped with a nitrocellulose filter having a 0.45-µm porosity, and apply vacuum. Allow the solution to degas by swirling the filtration unit, and disconnect the vacuum when no more bubbles are formed in the solution. Add appropriate amounts of Ammonium Persulfate Solution
as shown in Table 2
, swirl, and pour immediately into the gap between the two glass plates of the mold directly onto the surface of the polymerized Resolving Gel.
Immediately insert a clean polytef comb into the stacking gel solution, being careful to avoid trapping air bubbles. Add more stacking gel solution to fill the spaces of the comb completely. Leave the gel in a vertical position, and allow it to polymerize at room temperature. After polymerization is complete (about 30 minutes later), carefully remove the polytef comb, and proceed as directed below.
Sample Buffer 1
Dissolve 1.89 g of Tris, 5.0 g of SDS, 50 mg of bromophenol blue, and 25.0 mL of glycerol in 100 mL of water. Adjust with hydrochloric acid to a pH of 6.8, and dilute with water to 125 mL. Before use, dilute with an equal volume of water or sample, and mix.
Sample Buffer 2 (for reducing conditions)
Prepare as directed in Sample Buffer 1 except to add 12.5 mL of 2-mercaptoethanol before adjusting the pH. Alternatively, prepare as directed for Sample Buffer 1 except to start with about 1.93 g of Tris and add a suitable quantity of DTT to obtain a final 100 µM DTT concentration.
Dissolve 151.4 g of Tris, 721.0 g of aminoacetic acid (glycine), and 50.0 g of SDS in water; dilute with water to 5000 mL; and mix to obtain a stock solution. Immediately before use, dilute this stock solution with water to 10 times its volume, mix, and adjust to a pH between 8.1 and 8.8.
Rinse the wells immediately with water or with the Running Buffer
to remove any unpolymerized acrylamide. (If necessary, straighten the teeth of the Stacking Gel
with a blunt hypodermic needle attached to a syringe.) Remove the clamps on one short side, carefully pull out the tubing, and replace the clamps. Proceed similarly on the other short side. Remove the tubing from the bottom part of the gel.
Mount the completed gel in the electrophoresis apparatus. Add the electrophoresis buffers to the top and bottom reservoirs. Remove any bubbles that become trapped at the bottom of the gel between the glass plates. [noteRemoval is best done with a bent hypodermic needle attached to a syringe. Never prerun the gel before loading the samples, because that will destroy the discontinuity of the buffer systems. Before loading the sample, carefully rinse the slot with Running Buffer.]
Prepare the test and standard solutions in the recommended Sample Buffer, and treat as directed in the individual monograph. Apply the appropriate volume of each solution to the Stacking Gel wells.
Start the electrophoresis under the conditions recommended by the manufacturer of the equipment. Electrophoresis running time and current or voltage may need to be varied in order to achieve optimum separation. Check that the dye front is moving into the Resolving Gel. When the dye has reached the bottom of the gel, stop the electrophoresis. Remove the gel assembly from the apparatus, and separate the glass plates. Remove the spacers, cut off and discard the Stacking Gel, and immediately proceed with staining.
Detection of Proteins in Gels
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, because a band containing 10 to 100 ng can be detected; but the method is more cumbersome and less rugged. All of the steps in gel staining are performed at room temperature with gentle agitation (e.g., on a rocking platform shaker or equivalent). Gloves must be worn when staining the gels to prevent fingerprint residue staining.
Coomassie Staining Solution
Prepare a solution of Coomassie brilliant blue R-250 having a concentration of 1.25 g per L in a mixture of water, methanol, and glacial acetic acid (5:4:1). Filter, and store at room temperature.
Prepare a mixture of water, methanol, and glacial acetic acid (5:4:1).
Fixing Solution 1
Prepare a mixture of water, methanol, and trichloroacetic acid (5:4:1).
Fixing Solution 2
Transfer 250 mL of methanol to a 500-mL volumetric flask, add 0.27 mL of formaldehyde, dilute with water to volume, and mix.
Silver Nitrate Reagent
To a mixture of 40 mL of 1 M sodium hydroxide and 3 mL of ammonium hydroxide, add, dropwise and with stirring, 8 mL of a 200 g per L solution of silver nitrate; dilute with water to 200 mL, and mix.
Transfer 2.5 mL of a citric acid solution (2 in 100) and 0.27 mL of formaldehyde to a 500.0-mL volumetric flask, dilute with water to volume, and mix.
Prepare a 10% (v/v) solution of acetic acid.
Immerse the gel in an excess of Coomassie Staining Solution, and incubate for at least 1 hour. Remove the Coomassie Staining Solution. Destain the gel with an excess of Destaining Solution. Change the Destaining Solution several times, until the stained protein bands are clearly distinguishable on a clear background. The more thoroughly the gel is destained, the smaller the amount of protein that can be detected. Destaining can be accelerated by including a few g of anion-exchange resin or a small sponge in the Destaining Solution. [noteThe acidalcohol solutions used in this procedure do not completely fix proteins in the gel. This can lead to losses of some low molecular weight proteins during the staining and destaining of thin gels. Permanent fixation is obtainable by incubating the gel in Fixing Solution 1 for 1 hour before it is immersed in the Coomassie Staining Solution.]
Immerse the gel in an excess of Fixing Solution 2, and incubate for 1 hour. Remove Fixing Solution 2, add fresh Fixing Solution 2, and incubate for at least 1 hour, or overnight if convenient. Discard Fixing Solution 2, and wash the gel in an excess of water for 1 hour. Soak the gel for 15 minutes in a 1% solution of glutaraldehyde (v/v). Wash the gel twice, for 15 minutes each time, with an excess of water. Soak the gel in fresh Silver Nitrate Reagent for 15 minutes in darkness. Wash the gel three times, for 5 minutes each time, with an excess of water. Immerse the gel for about 1 minute in Developing Solution until satisfactory staining has been obtained. Stop the development by incubation in the Stopping Solution for 15 minutes, then rinse the gel with water, and proceed with drying as indicated below.
Drying of Gels
For Coomassie staining, after the destaining step, incubate the gel in a glycerol solution (1 in 10) for at least 2 hours. For silver staining, add to the final rinsing step a 5-minute incubation in a glycerol solution (1 in 50).
Immerse two sheets of porous cellophane 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 cellophane sheet. 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 a drying oven, leave at room temperature until dry, or use a commercial gel dryer.
Molecular Weight Determination
Molecular weights of proteins are determined by comparison of their mobilities with those of several marker proteins of known molecular weight. Mixtures of proteins with precisely known molecular weights blended for uniform staining are available for calibrating gels. They are available in various molecular weight ranges. Concentrated stock solutions of proteins of known molecular weight are diluted in a sample buffer and loaded on the same gel as the protein sample to be tested.
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 (semilogarithmic) plot of the logarithm of the molecular weights (MR) of the protein standards as functions of the RF values. [noteThe graphs are slightly sigmoid.] From the graph so obtained, estimate the unknown molecular weights by linear regression analysis or interpolation, as long as unknown samples are positioned along the linear part of the graph.
If the proteins of the molecular weight marker are not distributed along 80% of the length of the gel and over the required separation range (i.e., the range covering the product and its dimer or the products and its related impurities), and the separation obtained for the relevant protein bands does not show a linear relationship between the logarithm of the molecular weight and the RF , the test is not valid.
Quantification of Impurities
Where the impurity limit is specified in the individual monograph, a reference solution corresponding to that level of impurity is prepared by diluting the test solution. For example, where the limit is 5.0%, a reference solution is a 1 in 20 dilution of the test solution. No impurityany band other than the main bandin the electrophoretogram obtained from the test solution is more intense than the main band obtained with the reference solution.
Under validated conditions and when using the Coomassie staining procedure, impurities may be quantified by normalization to the main band, using an integrating densitometer. In this case, the responses must be validated for linearity.