SDS-Polyacrylamide Gel Electrophoresis (SDS-PAGE):

    SDS-Polyacrylamide Gel Electrophoresis (SDS-PAGE) is probably the worlds most widely used biochemical method. In the early 60's scientists first appreciated the utility of polyacrylamide gels as a convenient and versatile alternative to starch gels Ornstein 1964, Davis 1964, thus developing polyacrylamide gel electrophoresis or PAGE. The inclusion of ionic detergent Sodium Dodecyl Sulphate (SDS) to the gel and the sample was an important addition to this work. Shapiro et al. were one of the first to make use of this approach (Shapiro et al. 1967). Laemmli showed that proteins could be reliably fractionated by SDS-PAGE, which he described in a figure legend in a Nature paper (Laemmli, 1970). This paper used the method to study specifically the proteins of bacteriophage T4, and made use of Tris based buffers, as used by most researchers today. At about the same time Weber and Osborn showed that the method had general applicability and also showed, in a very systematic study, that the relative mobility of proteins in SDS-PAGE gels correlated quite well with their molecular weights (Weber and Osborn 1969). However these gels were based on phosphate buffers, which subsequently have been less widely used. After these papers appeared SDS-PAGE gels became wildly popular, first in tubes, then as slabs and then as minigels. Ultimately 2-dimensional gel techniques, using isoelectric focusing of proteins in one direction followed by regular SDS-PAGE in the other direction were developed. The first influential paper in this area was published by O'Farrell (O'Farrell 1975). People also began to appreciate that it was possible to transfer proteins out of SDS-PAGE gels onto agarose, or much more conveniently, onto nitrocellulose membranes and there stain them with antibodies. The most influential early paper making use of nitrocellulose membranes was that of Towbin et al. (Towbin et al 1979). Later studies used other kinds of membranes, notably the nylon like material PVDF, which allowed proteins transferred from SDS-PAGE gels to be probed with antibodies but also subjected to direct peptide sequencing (Matsudaira, 1975).

    The reason that SDS-PAGE works is in overview, this; SDS is a powerful detergent, which has a very hydrophobic end (the lipid like dodecyl part) and highly charged part (the sulfate group). The dodecyl part interacts with hydrophobic amino acids in proteins. Since the 3D structure of most proteins depends on interactions between hydrophobic amino acids in their core, the detergent destroys 3D structures, transforming what were globular proteins into linear molecules now coated with negatively charged SDS groups. After boiling in SDS proteins therefore become elongated with negative charges arrayed down them, so they will move towards a positive electrode. The reason why beta-mercaptoethanol is usually included in the sample buffer is to cleave disulfide bonds within or between molecules, allowing molecules to adopt an extended monomeric form. It is not surprising that the largest extended molecules are generally retarded the most by polyacrylamide gels, and the smallest ones the least. Since some proteins have few or no hydrophobic residues it is also not surprising that such molecules don't run on SDS page in a fashion which accurately reflects their molecular weight. Modifications such as phosphorylation and especially glycosylation can also cause proteins to run more or much more slowly than expected. Finally cross-linked proteins don't run as their molecular weight would predict, generally running slower particularly on higher percentage gels. However a particular protein runs on a particular position on a particular percentage acrylamide gel in a reliable manner.

    You'll need some sort of apparatus to run the gel in. Biorad and several other companies make these. You'll also need an apparatus to pour the gels in, unless you are buying premade gels, also available from many companies. Below are formulae we have been using for years, the exact origins of which are lost in recent prehistory. They work about as well as any other methods that are around. There are generally two gels, the resolving gel and the stacking gel. The stacking gel is of very low acrylamide concentration and is used to form the wells into which the protein is loaded. The low acrylamide concentration also allows most proteins to be concentrated at the dye front, so that dilute protein samples can be compared to concentrated samples on the same gel. The higher the acrylamide concentration the resolving gel the slower the proteins go through the gel. On lower percentage gels proteins go faster, so you should fix on a percentage that puts the proteins you are interested in somewhere in the resolving region of the gel.

    The solutions are made up as below; All except the ammonium persulfate can be stored at room temperature for several months. Acrylamide/bisacrylamide solutions may be light sensitive, so many people store these in dark colored or aluminium foil covered bottles. The ammonium persulfate should be made up each week, and stored at 4°C. Ammonium persulfate is rather unstable and decays to produce free radical SO₄^- ions, which react with the acrylamide molecules and initiates their polymerization. The formula for acrylamide is CH₂=CHCONH₂, and polymerization occurs by opening the double bond. So an acrylamide can react with another acrylamide to produce a linear polyacrylamide molecule, and the incorporation every now and then of a bisacrylamide can generate cross links between such linear molecules. Bisacrylamide is basically two acrylamides bound together, the formula being (CH₂=CHCONH)₂CH₂; you can vary the amount of this, the usual range being 1 part in 20 to 1 part in 50. The TEMED is (CH₃)₂NCH₂CH₂N(CH₃)₂ and acts as a catalyst, speeding up the decay of the ammonium persulfate. Polymerization is quicker and more uniform if you degas the first three solutions for a few minutes in an Ehrlenmeyer flask on a house vacuum prior to addition of the last three reagents. Molecular oxygen inhibits polymerization by reacting with the free radical SO₄^- ions, which is actually the reason why PAGE gels are poured in tubes or between plates and not in open top horizontal apparatuses, as can be done with agarose. Also it's a good idea to layer some isopropanol on top of the gel as this prevents oxygen getting in and inhibiting polymerization. If your gel doesn't polymerize it's most likely because the ammonium persulfate has gone off; this is only stable for a few days at 4 Degrees Centigrade. So just add more newly made solution and things should be fine. Alternatively you may have missed out one of the other components, easy enough to do...

Mixes for 20mls running gel solution, enough for two minigels, measure out the other components and make up to a final volume of 20 mls with distilled water. Note "ul" means microliter, many browsers have trouble displaying Greek characters correctly. For different volumes or for other acrylamide concentrations you can use our on-line SDS-PAGE solution calculator or our on-line SDS-PAGE solution calculator for use with Biorad 40% acrylamide solution.

Acrylamide range 5-12%

Acrylamide range 13.5-27%

Stacking Gel Solution, good for 2 minigels, 10 mls total volume, so measure out other components and make up to 10 mls final volume with distilled water is fine. Final concentration of acrylamide is 4.44%. To make other stacking gel concentrations you can use our on-line SDS-PAGE solution calculator, which can determine how much of each solution you need for stacking gels, and also allow you to select acrylamide concentrations other than 4.44%.

Solutions needed:

22.2 % Acrylamide/Bisacrylamide mix 22.2 g acrylamide, 0.6 g bis-acrylamide (37:1 cross-linker ratio) to 100 ml water, filtered.

44.4 % Acrylamide/Bisacrylamide mix 44.4 g acrylamide, 1.2 g bis-acrylamide (37:1 cross-linker ratio) to 100 ml water, filtered.

Reservoir/Running buffer = 57.6 g Glycine, 12 g Tris base, 4 g SDS, water to 4 litres

Stain solution = 2.5 g Coomassie Brilliant Blue R-250, 450 mls methanol, 100 mls glacial acetic acid, water to 1 liter.

Destain solution = 300 mls methanol, 400 mls acetic acid, water to 4 liters.

Sample buffer 5X = make up 100 mls and store away 5-10 mls aliquots.

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