The solution of proteins to be analyzed is first mixed with SDS, an anionic detergent which denatures secondary and non–disulfide–linked tertiary structures, and applies a negative charge to each protein in proportion to its mass. Without SDS, different proteins with similar molecular weights would migrate differently due to differences in folding, as differences in folding patterns would cause some proteins to better fit through the gel matrix than others. Adding SDS solves this problem, as it linearizes the proteins so that they may be separated strictly by molecular weight (primary structure, or number (and size) of amino acids). The SDS binds to the protein in a ratio of approximately 1.4 g SDS per 1.0 g protein (although binding ratios can vary from 1.1-2.2 g SDS/g protein), giving an approximately uniform mass:charge ratio for most proteins, so that the distance of migration through the gel can be assumed to be directly related to only the size of the protein. A tracking dye may be added to the protein solution to allow the experimenter to track the progress of the protein solution through the gel during the electrophoretic run.
Chemical ingredients and its roles
''Polyacrylamide gel (PAG)'' had been known as a potential embedding medium for sectioning tissues as early as 1954. Two independent groups Davis and Raymond employed PAG in electrophoresis in 1959. It possesses several electrophoretically desirable features that made it a versatile medium. Polyacrylamide gel separates protein molecules according to both size and charge. It is a synthetic gel, thermo-stable, transparent, strong, relatively chemically inert, can be prepared with a wide range of average pore sizes, can withstand high voltage gradients, feasible to various staining and destaining procedures and can be digested to extract separated fractions or dried for autoradiography and permanent recording. DISC electrophoresis utilizes gels of different pore sizes. The name DISC was derived from the discontinuities in the electrophoretic matrix and coincidentally from the discoid shape of the separated zones of ions (Anbalagan, 1999). There are two layers of gel, namely stacking or spacer gel, and resolving or separating gel.
The stacking gel is a large pore polyacrylamide gel (4%). This gel is prepared with Tris buffer pH 6.8 of about 2 pH units lower than that of electrophoresis buffer. These conditions provide an environment for Kohlrausch reactions, as a result, proteins are concentrated to several fold and a thin starting zone of the order of 19 ?m is achieved in a few minutes. This gel is cast over the resolving gel. The height of the stacking gel region was always maintained more than double the height and the volume of the sample to be applied.
The resolving gel is a small pore polyacrylamide gel (3 30%). The Tris buffer used is of pH 8.8. In this gel, macro molecules separate according to their size. In the present experiment, 8%, 10% and 12% Resolving gel were used for separating different range of proteins. 8% gel for 24 â€“ 205 kD proteins, 10% gel for 14-205 kD proteins and 12% gel for 14-66 kD proteins
Tris (tris (hydroxy methyl) aminomethane) (C4H11NO3; mw: 121.14). It has been used as a buffer because it is an innocuous substance to most proteins. Its pKa is 8.3 at 20 Â°C and reasonably a very satisfactory buffer in the pH range 7.0 â€“ 9.0.
Glycine (Amino Acetic Acid) (C2H5NO2; mw: 75.07). Glycine has been used as the source of trailing ion or slow ion because its pKa is 9.69 and mobility of glycinate are such that the effective mobility can be set at a value below that of the slowest known proteins of net negative charge in the pH range. The minimum pH of this range is somewhere around 8.0.
Acrylamide (C3H5NO; mw: 71.08). It is a white crystalline powder. While dissolving in water, autopolymerisation of acrylamide takes place. It is a slow spontaneous process by which acrylamide molecules join together by head on tail fashion. But in presence of free radicals generating system, acrylamide monomers are activated into a free-radical state. These activated monomers polymerise quickly and form long chain polymers. This kind of reaction is known as Vinyladdition polymerisation. A solution of these polymer chains becomes viscous but does not form a gel, because the chains simply slide over one another. Gel formation requires hooking various chains together. Acrylamide is a neurotoxin. It is also essential to store acrylamide in a cool dark and dry place to reduce autopolymerisation and hydrolysis.
Bisacrylamide (N,Nâ€™-Methylenebisacrylamide) (C7H10N2O2; mw: 154.17). Bisacrylamide is the most frequently used cross linking agent for poly acrylamide gels. Chemically it is thought of having two-acrylamide molecules coupled head to head at their non-reactive ends. Bisacrylamide was preserved at 4 Â°C.
Sodium Dodecyl Sulfate (SDS) (C12H25NaO4S; mw: 288.38). SDS is the most common dissociating agent used to denature native proteins to individual polypeptides. When a protein mixture is heated to 100 Â°C in presence of SDS, the detergent wraps around the polypeptide backbone. It binds to polypeptides in a constant weight ratio of 1.4 g/g of polypeptide. In this process, the intrinsic charges of polypeptides becomes negligible when compared to the negative charges contributed by SDS. Thus polypeptides after treatment becomes a rod like structure possessing a uniform charge density, that is same net negative charge per unit length. Mobilities of these proteins will be a linear function of the logarithms of their molecular weights.
TEMED (N, N, Nâ€™, Nâ€™-tetramethylethylenediamine) (C6H16N2; mw: 116.21). Chemical polymerisation of acrylamide gel is used for SDS-PAGE. It can be initiated by ammonium persulfate and the quaternary amine, N,N,Nâ€™,Nâ€™-tetramethylethylenediamine (TEMED). The rate of polymerisation and the properties of the resulting gel depends on the concentration of APS and TEMED. Increasing the amount of APS and TEMED results in a decrease in the average polymer chain length, an increase in gel turbidity and a decrease in gel elasticity. Decreasing the amount of initiators shows the reverse effect. The lowest catalysts concentrations that will allow polymerisation in the optimal period of time should be used. APS and TEMED are used, approximately in equimolar concentrations in the range of 1 to 10 mM.
Chemicals for processing and visualization
The following chemicals are used for processing of the gel and the protein samples visualized in it:
Bromophenol Blue (BPB) (3â€™, 3â€™â€™, 5â€™, 5â€™â€™-Tetrabromophenolsulphonephthalein) (C19H10Br4O5S; mw: 669.99). BPB is the universal marker dye. Proteins and nucleic acids are mostly colourless. When they are subjected to electrophoresis, it is important to stop the run before they run off the gel. BPB is the most commonly employed tracking dye, because it is viable in alkali and neutral pH, it is a small molecule, it is ionisable and it is negatively charged above pH 4.6 and hence moves towards cathode. Being a small molecule it moves ahead of most proteins and nucleic acids. As it reaches the anodic end of the electrophoresis medium electrophoresis is stopped. It can bind with proteins weakly and give blue colour.
Glycerol (C3H8O3; mw: 92.09). It is a preservative and a weighing agent. Addition of glycerol (20-30 or 50%) is often recommended for the storage of enzymes. Glycerol maintains the protein solution at very low temperature, without freezing. It also helps to weigh down the sample into the wells without being spread while loading.
Coomassie Brilliant Blue (CBB) (C45H44N3NaO7S2; mw: 825.97). CBB is the most popular protein stain. It is an anionic dye, which binds with proteins non-specifically. The structure of CBB is predominantly non-polar. So is usually used (0.025%) in methanolic solution (40%) and Acetic Acid (7%). Proteins in the gel are fixed by acetic acid and simultaneously stained. The excess dye incorporated in the gel can be removed by destaining with the same solution but without the dye. The proteins are detected as blue bands on a clear background. As SDS is also anionic, it may interfere with staining process. Therefore, large volume of staining solution is recommended, approximately ten times the volume of the gel.
Butanol (C4H10O; mw: 74.12). Water saturated butanol is used as an overlay solution on the resolving gel.
Besides the addition of SDS, proteins may optionally be briefly heated to near boiling in the presence of a reducing agent, such as dithiothreitol (DTT) or 2-mercaptoethanol (beta-mercaptoethanol/BME, which further denatures the proteins by reducing disulfide linkages, thus overcoming some forms of tertiary protein folding, and breaking up quaternary protein structure (oligomeric subunits). This is known as reducing SDS-PAGE, and is most commonly used. Non-reducing SDS-PAGE (no boiling and no reducing agent) may be used when native structure is important in further analysis (e.g. enzyme activity, shown by the use of zymograms). For example, quantitative preparative native continuous polyacrylamide gel electrophoresis (QPNC-PAGE) is a new method for separating native metalloproteins in complex biological matrices.
The denatured proteins are subsequently applied to one end of a layer of polyacrylamide gel submerged in a suitable buffer. An electric current is applied across the gel, causing the negatively-charged proteins to migrate across the gel towards the anode. Depending on their size, each protein will move differently through the gel matrix: short proteins will more easily fit through the pores in the gel, while larger ones will have more difficulty (they encounter more resistance). After a set amount of time (usually a few hoursthough this depends on the voltage applied across the gel; higher voltages run faster but tend to produce somewhat poorer resolution), the proteins will have differentially migrated based on their size; smaller proteins will have traveled farther down the gel, while larger ones will have remained closer to the point of origin. Thus proteins may be separated roughly according to size (and therefore, molecular weight). Following electrophoresis, the gel may be stained (most commonly with Coomassie Brilliant Blue or silver stain), allowing visualisation of the separated proteins, or processed further (e.g. Western blot). After staining, different proteins will appear as distinct bands within the gel. It is common to run "marker proteins" of known molecular weight in a separate lane in the gel, in order to calibrate the gel and determine the weight of unknown proteins by comparing the distance traveled relative to the marker. The gel is actually formed because the acrylamide solution contains a small amount, generally about 1 part in 35 of bisacrylamide, which can form cross-links between two polyacrylamide molecules. The ratio of acrylamide to bisacrylamide can be varied for special purposes. The acrylamide concentration of the gel can also be varied, generally in the range from 5% to 25%. Lower percentage gels are better for resolving very high molecular weight proteins, while much higher percentages are needed to resolve smaller proteins. Determining how much of the various solutions to mix together to make gels of particular acrylamide concentration can be done on line
Gel electrophoresis is usually the first choice as an assay of protein purity due to its reliability and ease. The presence of SDS and the denaturing step causes proteins to be separated solely based on size. False negatives and positives are possible. A co migrating contaminant can appear as the same band as the desired protein. This comigration could also cause a protein to run at a different position or to not be able to penetrate the gel. This is why it is important to stain the entire gel including the stacking section. Coomassie Brilliant Blue will also bind with less affinity to glycoproteins and fibrous proteins, which interferes with quantification (Deutscher 1990).
Most protein separations are performed using a "discontinuous" buffer system that significantly enhances the sharpness of the bands within the gel. During electrophoresis in a discontinuous gel system, an ion gradient is formed in the early stage of electrophoresis that causes all of the proteins to focus into a single sharp band. This occurs in a region of the gel that has larger pores so that the gel matrix does not retard the migration during the focusing or "stacking" event. Negative ions from the buffer in the tank then "outrun" the SDS-covered protein "stack" and eliminate the ion gradient so that the proteins subsequently separate by the sieving action in the lower, "resolving" region of the gel.
Many people continue to use a tris-glycine or "Laemmli" buffering system that stacks and resolves at a pH of ~8.3-9.0. These pHs promote disulfide bond formation between cysteine residues in the proteins, especially when they are present at high concentrations because the pKa of cysteine ranges from 8-9 and because reducing agent present in the loading buffer doesn't co-migrate with the proteins. Recent advances in buffering technology alleviate this problem by resolving the proteins at a pH well below the pKa of cysteine (e.g., bis-tris, pH 6.5) and include reducing agents (e.g. sodium bisulfite) that move into the gel ahead of the proteins to maintain a reducing environment. An additional benefit of using buffers with lower pHs is that the acrylamide gel is more stable so the gels can be stored for long periods of time before use.