SDS-PAGE, sodium dodecyl sulfate polyacrylamide gel electrophoresis, describes a collection of related techniques widely used in biochemistry, forensics, genetics and molecular biology to separate proteins according to their electrophoretic mobility (a function of the length of a polypeptide chain and its charge). In most proteins, the binding of SDS to the polypeptide chain imparts an even distribution of charge per unit mass, thereby resulting in a fractionation by approximate size during electrophoresis.
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Samples may be any material containing proteins, for example prokaryotic or eukaryotic cells, tissues, viruses, environmental samples, or purified proteins. In the case of solid tissues, these are often first broken down mechanically using a blender (for larger sample volumes), using a homogenizer (smaller volumes), by sonicator or by using cycling of high pressure. Cells may also be broken open by one of the above mechanical methods.
In the case of tissues or cells, a combination of biochemical and mechanical techniques – including various types of filtration and centrifugation – may be used to separate different cell compartments and organelles prior to electrophoresis.
The sample to be analyzed is 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. Heating the samples to at least 60°C further promotes protein denaturation, helping SDS to bind.[1] [2] [3] [4]
A tracking dye may be added to the protein solution. This typically has a higher electrophoretic mobility than the proteins to allow the experimenter to track the progress of the protein solution through the gel during the electrophoretic run.
The gels typically consist of acrylamide, bisacrylamide, SDS, and a buffer with an adjusted pH. The solution may be degassed under a vacuum to prevent the formation of air bubbles during polymerization. [5] A source of free radicals and a stabilizer such as ammonium persulfate and TEMED are added to initiate polymerization.[6] The polymerization reaction results in a gel because of the added bisacrylamide, generally about 1 part in 35 relative to acrylamide, 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 is possible.
Gels are usually polymerized between two glass plates in a gel caster, with a comb inserted at the top to create the sample wells. After the gel is polymerized the comb can be removed and the gel is ready for electrophoresis.
Various buffer systems are used in SDS-PAGE depending on the nature of the sample and the experimental objective. The buffers used at the anode and cathode may be the same or different.[7][8] [9]
An electric field is applied across the gel, causing the negatively-charged proteins to migrate across the gel towards the positive (+) electrode (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 hours- though 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. Proteins may therefore be separated roughly according to size (and thus molecular weight); however certain glycoproteins behave anomalously on SDS gels.
Following electrophoresis, the gel may be stained (most commonly with Coomassie Brilliant Blue R-250 or silver stain), allowing visualization 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 molecular weight size markers of known molecular weight in a separate lane in the gel, in order to calibrate the gel and determine the approximate molecular mass of unknown proteins by comparing the distance travelled relative to the marker.
SDS-PAGE 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 approximately based on size, but aberrant migration of some proteins may occur. Different proteins may also stain differently, which interferes with quantification by staining. SDS-PAGE may also be used as a preparative technique for the purification of proteins. For example, quantitative preparative native continuous polyacrylamide gel electrophoresis (QPNC-PAGE) is a method for separating native metalloproteins in complex biological matrices.
Polyacrylamide gel (PAG) had been known as a potential embedding medium for sectioning tissues as early as 1964, and two independent groups employed PAG in electrophoresis in 1959.[10][11] It possesses several electrophoretically desirable features that make it a versatile medium. It is a synthetic, thermo-stable, transparent, strong, chemically relatively inert gel, and can be prepared with a wide range of average pore sizes.[12] The pore size of a gel is determined by two factors, the total amount of acrylamide present (%T) (T = Total concentration of acrylamide and bisacrylamide monomer) and the amount of cross-linker (%C) (C = bisacrylamide concentration). Pore size decreases with increasing %T; with cross-linking, 5%C gives the smallest pore size. Any increase or decrease in %C from 5% increases the pore size, as pore size with respect to %C is a parabolic function with vertex as 5%C. This appears to be because of non-homogeneous bundling of polymer strands within the gel. This gel material can also withstand high voltage gradients, is amenable to various staining and destaining procedures, and can be digested to extract separated fractions or dried for autoradiography and permanent recording.
The following chemicals are used for processing of the gel and the protein samples visualized in it:
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.
In the 14th century the silver staining technique was developed for colouring the surface of glass. It has been used extensively for this purpose since the 16th century. The colour produced by the early silver stains ranged between light yellow and an orange-red. Camillo Golgi perfected the silver staining for the study of the nervous system. Golgi's method stains a limited number of cells at random in their entirety.[13] The exact chemical mechanism by which this happens is still largely unknown.[14] Silver staining was introduced by Kerenyi and Gallyas as a sensitive procedure to detect trace amounts of proteins in gels.[15] The technique has been extended to the study of other biological macromolecules that have been separated in a variety of supports.[16] Classical Coomassie Brilliant Blue staining can usually detect a 50 ng protein band, Silver staining increases the sensitivity typically 50 times. Many variables can influence the colour intensity and every protein has its own staining characteristics; clean glassware, pure reagents and water of highest purity are the key points to successful staining.[17]
Most protein separations are performed using a "discontinuous" (or DISC) 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. The formation of the ion gradient is achieved by choosing a pH value at which the ions of the buffer are only moderately charged compared to the SDS-coated proteins. These conditions provide an environment in which Kohlrausch reactions determine the molar conductivity. As a result, SDS-coated proteins are concentrated to several fold in a thin zone of the order of 19 μm within a few minutes. At this stage all proteins migrate at the same migration speed by isotachophoresis. 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. [18][19] Separation of the proteins by size is achieved in the lower, "resolving" region of the gel. The resolving gel typically has a much smaller pore size, which leads to a sieving effect that now determines the electrophoretic mobility of the proteins. At the same time, the separating part of the gel also has an pH value in which the buffer ions on average carry a greater charge, causing them to "outrun" the SDS-covered proteins and eliminate the ion gradient and thereby the stacking effect.
A very widespread discontinuous buffer system is the tris-glycine or "Laemmli" system that stacks at a pH of 6.8 and resolves at a pH of ~8.3-9.0. A drawback of this system is that these pH values may promote disulfide bond formation between cysteine residues in the proteins 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 pH values is that the acrylamide gel is more stable lower pH values, so the gels can be stored for long periods of time before use.[20][21]
As voltage is applied, the anions (and negatively charged sample molecules) migrate toward the positive electrode (anode) in the lower chamber, the leading ion is Cl¯ ( high mobility and high concentration); glycinate is the trailing ion (low mobility and low concentration). SDS-protein particles do not migrate freely at the border between the Cl¯ of the gel buffer and the Gly¯ of the cathode buffer. Friedrich Kohlrausch found that Ohm's law also applies to dissolved electrolytes. Because of the voltage drop between the Cl- and Glycine-buffers, proteins are compressed (stacked) into micrometer thin layers.[22] The boundary moves through a pore gradient and the protein stack gradually disperses due to a frictional resistance increase of the gel matrix. Stacking and unstacking occurs continuously in the gradient gel, for every protein at a different position. For a complete protein unstacking the polyacrylamide-gel concentration must exceed 16% T. The two-gel system of "Laemmli" is a simple gradient gel. The pH discontinuity of the buffers is of no significance for the separation quality, and a "stacking-gel" with a different pH is not needed.