SDS-PAGE is widely used to analyze the proteins in complex extracts. The most commonly used methods are derived from the discontinuous SDS-PAGE system first described by Laemmli (1970). The system actually consists of two gels - a resolving (aka running) gel in which proteins are resolved on the basis of their molecular weights (MWs) and a stacking gel in which proteins are concentrated prior to entering the resolving gel. Differences in the compositions of the stacking gel, resolving gel and electrophoresis buffer produce a system that is capable of finely resolving proteins according to their MWs.
Gel electrophoresis of macromolecules
In gel electrophoresis, an electric field is used to move charged molecules through
a matrix of a polymerized substance such as agarose or polyacrylamide. The rates at which individual molecules move through the gel depend on the properties of both the separation system and the molecules themselves. Gel matrices are permeated with networks of pores through which the molecules move. The amount of resistance that the matrix presents to the movement of a molecule depends on the diameter of the pore as well as the size and geometry of the molecule. Researchers can control the size of the pore by adjusting the concentration of gel monomer within a certain range. In general, smaller, more highly charged molecules migrate more rapidly through gels than larger or less charged molecules. The mobility of a molecule is also affected by the buffer system and the strength of the electrophoretic field used for the separation.
You have already used agarose gel electrophoresis to separate DNA molecules. Recall that the size of a linear DNA molecule can be estimated from the rate at which it moves through an agarose gel, because DNA molecules have a uniform charge to mass ratio. Protein electrophoresis is somewhat more complicated than DNA electrophoresis. Proteins are much smaller than DNA molecules, so polyacrylamide gels are used for their separation. In addition, proteins are much more structurally diverse than DNA, so chemical treatments (see below) are used to impart a uniform geometry and charge/mass ratio to the proteins.
Chemistry of acrylamide polymerization
The polyacrylamide gels used to separate proteins are formed by the chemical polymerization of acrylamide and a cross-linking reagent, N,N’methylenebisacrylamide (opposite page). Investigators are able to control the size of the pores in the gel by adjusting the concentration of acrylamide, as well as the ratio of acrylamide to bisacrylamide. Raising either the concentration of acrylamide or bisacrylamide, while holding the other concentration constant, will decrease the pore size of the gel. Polymerization occurs because of free oxygen radicals that react with the vinyl groups in acrylamide and bisacrylamide, as shown in the figure below. The oxygen radicals are generated from the catalyst, ammonium persulfate (APS), when it reacts with a second catalyst, N,N,N’,N’-tetramethylethylenediamine (TEMED).
Acrylamide gel polymerization.
Ammonium persulfate and TEMED catalyze the polymerization of acrylamide and bis-acrylamide monomers into a crosslinked network.
Proteins are denatured prior to electrophoresis
Compared to DNA molecules, proteins are structurally very diverse. Proteins show tremendous variation in their amino acid compositions and in the distribution of amino acids
in their folded structures, features with important implications for electrophoresis. Recall that proteins are mixtures of hydrophobic and hydrophilic amino acids and that the primary sequence of the protein determines its final folded form. Because of the hydrophobic effect, the surfaces
of proteins proteins have a higher frequency of polar and charged amino acids than the interiors, where hydrophobic residues predominate. Folded proteins assume many different geometries and their surfaces are mosaics with respect to the distribution of R groups with different chemistries. Because proteins are so diverse with respect to their surface charges and geometries, the molecular weights of folded proteins cannot be simply determined by their migration rate in an electric field. Postively and negatively charged proteins would migrate in different directions!
To resolve the proteins in a sample according to their size, investigators must convert the proteins to a uniform geometry and impart a uniform charge/mass ratio to the proteins. In SDS- PAGE, the solution is to denature the proteins by boiling them with the anionic detergent, sodium dodecyl sulfate (SDS) and 2-mercaptoethanol. The combination of heat and detergent is sufficient to break the many noncovalent bonds that stabilize protein folds, and 2-mercaptoethanol breaks any covalent bonds between cysteine residues. Like other detergents, SDS is an amphipathic molecule, consisting of a hydrophobic 12-carbon chain and a hydrophilic sulfate group. The SDS hydrocarbon chain permeates the protein interior and binds to hydrophobic groups, reducing the protein to a random coil, coated with negatively charged detergent molecules all along its length. Denatured proteins bind quite a lot of SDS, amounting to ~1.4 g SDS/g protein, or ~one SDS mol- ecule for every two amino acids.
Discontinuities between the stacking and running gels underlie the resolving power of the SDS-PAGE gels
The Laemmli (1970) SDS-PAGE system can be considered a 3-component system. The stacking and running (resolving) gels have different pore sizes, ionic strengths and pHs. The third component is the electrophoresis buffer (25 mM Tris, 192 mM glycine,, 0.1% SDS, pH ~8.3), which contains large amounts of glycine. The ionization state of the glycine is critical to the separation. At neutral pH, glycine is a zwitterion, with a negatively charged carboxyl group and a positively charged amino group. The pKa of the amino group is 9.6, considerably higher than the pH of the chamber buffer. Consequently, very little glycine has a negative charge in the chamber buffer or stacking gel, and significant ionization does not occur until the glycine enters the more alkaline pH 8.8 environment of the running gel. Let’s follow the progress of protein samples during SDS-PAGE to see how differences in the composi- tion of these three components generate the high resolving power of SDS-PAGE gels.
The sample buffer used for SDS-PAGE contains a tracking dye, bromophenol blue (BPB), which will migrate with the leading edge of the proteins being separated on the gel. The sample buffer also contains glycerol, which allows the protein samples to settle into the bottom of the gel wells. The gel is vertically positioned in the electrophoresis apparatus and covered with chamber buffer containing glycine (right, shaded).
Once a voltage is applied, the chloride ions in the sample buffer and stacking gel move rapidly toward the positive pole, forming the leading edge of a moving ion front. Glycine molecules have very little charge in the stacking gel, so they migrate at the rear of the moving ion front. This difference in chloride and glycine mobility sets up a steep voltage gradient in the stacking gel that sweeps along the negatively charged protein-SDS complexes. The large pores of the stacking gel present very little resistance to the movement of protein-SDS complexes, which then “stack up” into a very concentrated region at the interface between the running and stacking gels (right). Protein-SDS complexes remain concentrated at the interface until the slowly migrating glycine molecules reach the boundary between the two gels.
Dramatic changes occur as the glycine ions enter the running gel. The pH of the running gel is closer to the pKa of the glycine amino groups, so a significant fraction of the glycine molecules assume a negative charge. Negatively charged glycine molecules begin to move at the same rate as the chloride ions, thereby eliminating the voltage difference that controlled protein mobility through the stacking gel. The pores in the running gel are much smaller than those of the stacking gel, so the pores present frictional resistance to the migration of proteins. Proteins begin to migrate at different rates, because of the sieving properties of the gel. Smaller protein-SDS complexes migrate more quickly than larger protein- SDS complexes (right). Within a certain range determined by the porosity of the gel, the migration rate of a protein in the running gel is inversely proportional to the logarithm of its MW.
Proteins are visualized with stains.
With few exceptions, naturally-occurring proteins are invisible on SDS-PAGE gels. Consequently, researchers often use pre-stained protein standards to monitor the approximate positions of proteins during electrophoresis. The pre-stained standards are produced by covalently attaching a large number of chromophores to a protein. The addition of the chromophores increases the MW of the protein and also produces more diffuse bands on the
gel. The diffuseness of the bands reflects variation in the number of dye molecules attached to individual protein molecules. We will use prestained standard protein ladders in our gels, so you will be able to visualize the protein separation that is occurring. Yeast proteins will not be visible, however, because they have not been modified with chromophores.
To visualize the positions of proteins after electrophoresis is complete, investigators stain the gels with various dyes that bind noncovalently and with very little specificity to proteins. During the staining process, proteins are also “fixed” in the gel, meaning that proteins become insoluble and unable to diffuse out of the gel. In our experiments, we will use Simply Blue, a colloidal suspension of Coomassie Brilliant Blue G-250. Brilliant Blue G-250 binds proteins nonspecifically through a large number of ionic and Van der Waals interactions. In this procedure, gels are rinsed with water to remove the buffer salts used for electrophoresis and then treated with the colloidal G-250 suspension. Protein bands appear rapidly, and when necessary, the gels can be destained with deionized water to lower the gel background. Brilliant Blue staining intensity is considered to be a quantitative procedure, because with some exceptions, the intensity of a stained band is directly proportional to the amount of protein in a band.
Protein molecular weights can be calculated from their migration on gels
The sizes of proteins in an extract can be calculated by comparing their migration to a set of standard proteins run on the same gel. Researchers select standard proteins that will be well- resolved on the particular gel that they are running. For example, an investigator using a 7.5% gel will select standards with higher molecular weights (MWs) than an investigator using a 15% gel, which is better suited to the analysis of small proteins. The principles used to estimate MWs are the same used for agarose gel electrophoresis. A plot of the log10MW of the standard proteins against the distance that each protein migrated on the gel will give a straight line in the region where the gel has good resolving power. (Note: MW is not the same as the mass of a protein. MW is a dimensionless term. For example, myoglobin has a mass of 16.7 kDa and a MW of 16,700.) The sizes of unknown proteins can be estimated by interpolating experimental values on a graph of standard proteins. Proteins whose molecular weights fall outside this range will not be well- resolved on the gel.
The figure below illustrates several of the points discussed above. The same sets of un- stained and pre-stained protein standards were separated on either 12% or 15% SDS-PAGE gels. The prestained standards in lanes 1-5 are visible without staining, but they become much more pronounced after staining. The unstained standard in lane 6 requires staining to become visible, but the bands are much more discrete and will give more reliable values when calculating MWs of unknown proteins, because chromophores have not been attached to the proteins. The data in lanes 2-5 also demonstrate that Brilliant Blue staining is a quantitative procedure, because the intensity of bands in each lane increases in direct proportion to the amount of protein in the lane.
When analyzing your experimental data, remember to consider the additional amino acids that have been added during the cloning procedure. The Met proteins that you are working with are fusion proteins with additional amino acids at the C-termini the Met proteins. The BG1805 plasmid encodes HA and His6 epitopes, as well as the ZZ immunoglobin binding domain. Together these sequences add a walloping ~19 kDa to the expected mass of S. cerevisiaeMet proteins (Gelperin et al., 2005). The pYES2.1 plasmid encodes 33 amino acids that are added to cloned ORFs. The additional sequences include a V5 epitope tag and a (His)6 purification tag at the C-termini of overexpressed proteins. Together, these amino acids add ~5 kDa to the size of the protein.
The MW of the LacZ control protein without the V5 epitope is ~120 kDa. Because this is such a large protein, it will be very difficult to get an accurate estimate of its MW.