Introduction to SDS-PAGE
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Introduction to SDS-PAGEThe separation of macromolecules in an electric field is called electrophoresis. In an early form of electrophoresis, disolved protein mixtures were placed in a U-shaped buffer-filled channel and subjected to an electric field. Resolution was poor and any disturbance of the apparatus compromised the separation. Gels were developed to serve as solid supports for electrophoresis, so that the separated products remain separated and can be easily stained and handled. The development of the stacking gel, which compresses the sample into bands a few micrometers thick, added a major improvement to the resolution of gels (Ornstein, 1964; Davis, 1964). Other landmark improvements to protein electrophoresis were the use of polyacrylamide for control of separation by molecular size, and the use of sodium dodecyl sulfate (SDS; lauryl sulfate) to denature proteins in order to ensure reproducibility of the technique (Weber and Osborn, 1969; U.K. Laemmli, 1970).
SDS is an anionic detergent, meaning its molecules have a net negative charge. It binds to most soluble protein molecules in aqueous solutions over a wide pH range. Polypeptide chains bind amounts of SDS that are proportional to the size of the molecules. The negative charges on SDS destroy most of the complex (secondary and tertiary) structure of proteins, and are strongly attracted toward an anode (positively-charged electrode) in an electric field.
A polyacrylamide gel with acrylamide content above a critical density restrains larger molecules from migrating as fast as smaller molecules. Because the charge-to-mass ratio is nearly the same among SDS-denatured polypeptides, the final separation of proteins is dependent almost entirely on the differences in molecular weight (MW) of polypeptides. In a gel of uniform density the relative migration distance of a protein (Rf) is negatively proportional to the log of its MW. If proteins of known MW are run simultaneously with the unknowns, the relationship between Rf and MW can be plotted, and the MWs of unknown proteins determined. Protein separation by SDS-PAGE is used to determine the relative abundance of major proteins in a sample, their approximate molecular weights, and in what fractions they can be found. The purity of protein samples can be assessed. Different staining methods can be used to detect rare proteins and to learn something about their biochemical properties. Specialized techniques such as Western blotting, two-dimensional electrophoresis, and peptide mapping can be used to detect extremely scarce gene products, to find similarities among them, and to detect and separate isoenzymes of proteins.
Many systems for protein electrophoresis have been developed, with features such as built in gel casting apparatus and water-cooled jackets. SDS-PAGE can be conducted on pre-cast gels, saving the trouble and hazard of working with acrylamide. The following description applies to shop-made casting and running apparatus that are much cheaper than commercially available equipment. In addition to cost effectiveness, an advantage of making one's own gels the first time is a deeper understanding of the process.
Preparations will involve casting of two different layers of acrylamide between glass plates. The lower layer (separating, or resolving, gel) will be responsible for actually separating polypeptides by size. The upper layer (stacking gel) will include the sample wells, and will be of a composition that causes the samples to become compressed (stacked) in order to have thin bands and correspondingly better resolution among bands than if we did not stack them.
We use casting stands to prepare the mini-slab gels. Two clean plates with two teflon spacers make a single cassette. We stack the cassettes upright in the stand with the bottoms of the cassettes tight to the bottom of the stand, using modeling clay to seal a thick acrylic cover in place against the last cassette to make a water-tight chamber. Using a well-former (comb) as a template, we mark a fill line about a centimeter below the bottom of the comb for the height of the first (separating) gel solution.
In the Laemmli system for SDS-PAGE, proteins are pulled into an upper (stacking) gel, then begin to separate when they reach a lower (separating) gel of different compositiion. A discontinuous gel must therefore be poured in two stages. Acrylamide polymerizes spontaneously in the absence of oxygen, so the polymerization process involves complete removal of oxygen from the solution.
The total volume between the plates of our gel cassettes is ten ml, so if we prepare 10 ml separating gel mix per cassette we have more than enough. We typically prepare three cassettes per stand and use the best one of the three. From 30% acrylamide stock (see notes below) we prepare gels of compostion 7 to 15% acrylamide, depending on the range of proteins that we wish to separate. Our separating gel buffer stock (4x concentrated) consists of 0.4% SDS, 1.5 M Tris-Cl, pH 8.8. Per cassette, we mix 2.5 ml buffer stock and sufficient acrylamide stock so that when the mix is brought to final volume with distilled water we have the desired percent acrylamide monomer.
Polymerization is more uniform if the mix is de-gassed to remove much of the dissolved oxygen, by placing it under a vacuum for 5 minutes or so before polymerization. We initiatiate polymerization by adding freshly prepared10% ammonium persulfate (AP) to the mix followed by N-, N'-tetramethylenediamine (TEMED). The amounts of each depend on the quality of acrylamide used, and should be determined in advance by trial and error. We usually start with 100 µl AP and 10 µl TEMED per 10 ml gel mix, and see how it goes. Once the catalysts are added, polymerization may occur quickly, thus it is necessary to have the casting stand completely ready and to have the overlay solution ready to go (see below). After swirling to mix, we simply pour the solution into the space occupied by the cassettes. The cassettes will self-level eventually, but leveling can be hurried along by adding solution to selected cassettes with a pasteur pipet. Excess solution can be removed by tipping the apparatus and pulling off the excess with a pipet, so that the final level is at the fill mark.
Immediately after pouring the gel mix, it must be overlaid with water-saturated butanol to an additional height of 0.5 cm or so (butanol is the top layer in the stock container). Adding butanol to a single cassette will drive the acrylamide mix down, raising the level in the others, so care must be taken to distribute the butanol equally among the cassettes. The purpose of butanol is to produce a smooth, completely level surface on top of the separating gel, so that bands are straight and uniform. Butanol holds very little water in solution, forming a neat layer on top, which is why we use it. Water would make an effective overlay but would mix with the acrylamide solution, diluting it. In fact, the butanol we use is saturated with water so that it does not dry out the gel mix.
Polymerization can be confirmed by pulling some of the remaining gel mix into the pipet, allowing it to stand, and checking it after 10 min or so. When the gel mix can no longer be expelled by squeezing the bulb, the separating gel is set. It should not take more than 15 minutes for any of the gel mixes to polymerize. If it hasn't gelled by that time, something is probably wrong. Often, first time "gel makers" are misled into thinking the gel hasn't polymerized because the top 0.5 ml or so of the gel mix does not set (some oxygen reaches it through the overlay).
Ten ml of stacking gel mix is sufficient for three cassettes, however for the sake of accuracy it may be preferable to make 20 or 30 ml. Excess can be rinsed and tossed into a wastebasket after it polymerizes. It isn't necessary to degas a stacking mix, because the stacker is simply designed to perform as a matrix through which samples will pass as they are caught up between moving boundaries. It is not designed for uniform separation of proteins. Our stacking gel buffer stock consists of 0.5 M Tris-Cl, pH 6.8, with 0.4% SDS. Typical stackers are 3 to 4.5% acrylamide. We use 4% in order to permit stacking of very large proteins and still retain sufficient mechanical strength to make good sample wells.
Before adding the final two components, which will start polymerization, the butanol should be poured off the separating gels into a sink with tap water running and excess butanol/acrylamide removed from the surfaces with a pipet. We use AP and TEMED in similar proportions as for the separating gel mix, although we sometimes increase the amount of one or both components since lower percentage acrylamide solutions tend to polymerize more slowly. After adding AP and TEMED we immediately swirl the mix and pour it into the cassettes to the tops of the plates. We insert combs one at a time, taking care not to catch bubbles under the teeth, and adjust to make them even if necessary, scraping excess stacking mix off later.
Water-saturated butanol prevents drying of the edge of the resolving gel, retards the diffusion of oxygen into the acrylamide, and levels the surface. Butanol is preferred because it is not miscible with the acrylamide solution, and is much lighter. It is water-saturated in advance so that it does not extract water from the acrylamide mix.
There is no need to de-aerate a stacking solution. Uniformity of the polyacrylamide is not essential, since the stacking gel doesn't separate proteins.
A separating gel of given acrylamide concentration separates proteins effectively within a characteristic range. The largest polypeptides can enter a low percentage gel readily, and are fairly well separated. However, such a gel has a relatively low cutoff. That is, polypeptides below a particluar size are not restricted at all by the gel, and all move at the same pace, along with the tracking dye, regardless of size. A gel of 7% acrylamide composition typically has a cutoff of 45 kiloDaltons. A gel of very high percentage acrylamide may restrict all of the proteins in a mixture. The smallest protein of any significance among the fractions of mammalian blood is hemoglobin (14 kD). The hemoglobin band is readily resolved in a 15% acrylamide gel, but is buried in the dye front in a 7% gel. The problem with running just a 15% gel is that the heavier proteins are so restricted that they are jammed near the top of the gel and are not easily resolved from one another. In fact, it is a good idea to forget about analyzing the top third of such a gel. To take advantage of the characteristics of both low and high percentage gels, we usually run both.
In the teaching lab we recommend that alternate teams prepare low or high percent gels, with each team exchanging samples with a team that prepared the other type gel. Each team, then, would load its set of samples, appropriate standards, and another team's samples on its gel, and have its samples loaded onto another percent gel as well. In addition to expanding the range of resolution of bands, this practice allows comparison between identical fractions prepared by different teams, to control for inconsistencies in fractionation, sample preparation, etc.
When the stacking gel has set, the cassettes should be rinsed free of any excess liquid, leaving the combs in place. If casting stands are used, the clay is scraped off of the front cover and the cover removed. Gel cassettes are separated with the aid of a single edged razor blade if necessary (having beveled plates helps). After scraping off any excess stacking gel, the surfaces of the plates must be rinsed and dried, and the best gels selected. Small air spaces may appear between stacking gel and resolving gel or between the gel and the glass plates. As the outside pressure on the plates is relieved the glass expands, creating some spaces. As long as there is no continuous channel from the top to the bottom of the gel, the spaces will not influence protein migration.
The assembly of a gel running stand varies with the type of apparatus. The top of the cassette must be continuous with an upper buffer chamber and the bottom must be continuous with a lower chamber so that current will run through the gel itself. The cassette must be sealed in place using gaskets or a sealant such as agarose. In a teaching lab the assembly is best described by going through the procedure, using a film, and/or having a demonstration set up. We fill both the upper and lower buffer compartments with an electrode buffer (running buffer) consisting of 25 mM Tris, 192 mM glycine, 0.1% sodium dodecyl sulfate. We do not adjust pH of the electrode buffer. We remove the comb from the gel before filling the upper buffer compartment.
Hamilton syringes work well for loading samples into the wells. Ideally, the glycerol in a sample causes it to sink neatly to the bottom of the well, allowing as much as 20 µl or even more to be loaded. If the combs do not fit well or the plates are not clean the sample often hangs up, and we are limited to 10 µl or so.
The anode (+ electrode) must be connected to the bottom chamber and the cathode to the top chamber. The negatively-charged proteins will move toward the anode, of course. Gels are usually run at a voltage that will run the tracking dye to the bottom as quickly as possible without overheating the gels. Overheating can distort the acrylamide or even crack the plates. The voltage to be used is determined empirically - 150 volts works well for us.
When the dye front is nearly at the bottom of the gel it is time to stop the run. For low percent gels with a tight dye front, the dye should be on the verge of running off the gel. When the percent acrylamide is high the dye front may be diffuse, since the dye is not homogeneous. If you know the approximate position of the lowest protein band you can let the dye run off. Only experience will tell you when it is appropriate to stop the run. Before removing gels the power must be turned off and cables removed (using one hand, to avoid making a circuit).
Removal the gel from the cassette is better demonstrated than described. The plates are separated and the gel is dropped into a staining dish containing deionized water. After a quick rinse, the water is poured off and stain added. Staining usually requires incubation overnight, with agitation.
The dye actually penetrates the entire gel, however it only sticks permanently to the proteins. Excess dye is washed out by 'destaining' with acetic acid/methanol, also with agitation. It is most efficient to destain in two steps, starting with 50% methanol, 10% acetic acid for 1-2 hours, then using 7% methanol, 10% acetic methanol to finish. The first solution shrinks the gel, squeezing out much of the liquid component, and the gel swells and clears in the second solution. Properly stained/destained gels should display a pattern of blue protein bands against a clear background. The gels can be dried down or photographed for later analysis and documentation.
The original dye front, consisting of bromphenol blue dye, disappears during the process. In fact, bromphenol blue is a pH indicator which turns light yellow under acid conditions, prior to being washed out. In low percentage gels, sufficient protein may run with the dye front so that the position of the bromphenol blue front is permanently marked with unresolved proteins, often forming a continuous "front" across the bottom of the gel. In higher % gels, a distinct dye front is usually not obtained.
Coomassie blue may not stain some proteins, especially those with high carbohydrate content. Stains such as periodic acid-Schiff (PAS), fast green, or Kodak 'Stain's all' may detect different patterns. Silver staining is generally used when detection of very faint proteins is necessary.
Routine staining with Coomasie Blue is straightforward - about the only ways to ruin a gel at this point are physical damage (ripping the gel, for example), letting dye pool and precipitate in the gel, forgetting the alcohol at some step, allowing protein to dissolve and diffuse out of the gel. If that happens, the information is lost.
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