Laboratory 8 SDS-PAGE (Polyacrylamide Gel Electrophoresis) of HRP Objectives: The purpose of this method is to separate proteins according to their size and in reference to standard proteins of known size. The method is often used to determine the relative size and purity of a protein sample. Students should be able to: Prepare samples for SDS-PAGE Successfully run a SDS-PAGE Stain and destain a SDS-PAGE Analyze SDS-PAGE data. Introduction: Since the purpose is to separate many different protein molecules of varying shapes and sizes, the first goal is to get them to be linear so that the proteins no longer have any secondary, tertiary or quaternary structure (i.e. they should have the same linear shape). Consider two proteins that are each 500 amino acids long but one is shaped like a closed umbrella while the other one looks like an open umbrella. If you tried to run down the street with both of these molecules under your arms, which one would be more likely to slow you down, even though they weigh exactly the same? This analogy helps point out that not only the mass but also the shape of an object will determine how well it can move through an environment. So what is needed is a way to convert all proteins to the same shape by boiling them in SDS. The cartoon in Figure 1 depicts what happens to a protein when it is boiled with the denaturing detergent SDS. The top portion of the figure shows a protein with negative and positive charges due to the charged R-groups of the particular amino acids in the protein. The large H represents hydrophobic domains where nonpolar R-groups have collected in an attempt to get away from the polar water that surrounds the protein. The bottom portion shows that SDS can break up hydrophobic areas and coat proteins with many negative charges, which overwhelm any positive charge in the protein due to positively, charged R-groups. The resulting protein has been denatured by SDS (reduced to its primary structure) and as a result has been linearized. R-groups have collected in an attempt to get away from the polar water that surrounds the protein. The bottom portion shows that SDS can break up hydrophobic areas and coat proteins with many negative charges, which overwhelm any positive charge in the protein due to positively, charged R-groups. The resulting protein has been denatured by SDS (reduced to its primary structure) and as a result has been linearized. Figure 1: The effect of SDS and mercaptoethanol on protein structure. SDS (sodium dodecyl sulfate) is an anionic detergent which denatures proteins by associating non-covalently the polypeptide backbone binding specifically in a mass ratio of 1.4:1. In so doing, SDS confers a negative charge to the polypeptide in proportion to its length - ie: the denatured polypeptides become "rods" of negatively charged clouds with equal charge or charge densities per unit length. It is usually necessary to reduce disulfide bridges in proteins. This is done with 2- mercaptoethanol or -ME and DTT respectively). So a protein that started out like the one shown in the top part of Figure 1 will be converted into the one shown in the bottom part of Figure 1. The end result has two important features: 1) all proteins contain only primary structure and 2) all proteins have a large negative charge which means they will all migrate towards the positive pole when placed in an electric field. If these denatured are put into an electric field, they will all move towards the positive pole at the same rate, with no separation by size. So what is needed is a matrix crosslinked 8-1
polyacrylamide gels so proteins move at different rates based on their size. A polyacrylamide gel is not solid but is made of a labyrinth of tunnels through a meshwork of fibers. Figure 2 shows a slab of polyacrylamide (dark gray) with tunnels (different sized red rings with shading to depict depth) exposed on the edge. Notice that there are many different sizes of tunnels scattered randomly throughout the gel. Figure 4: Movement of SDS denatured proteins through a PAGE gel. (red plus) at the far end and the negative pole (black minus) at the closer end. Since all the proteins have strong negative charges, they will all move in the direction the arrow is pointing (run to red). Figure 2: Cross-section of PAGE gel. Figure 3 shows a cross-section of two selected tunnels (only two are shown for clarity of the diagram). These tunnels extend all the way through the gel, but they meander through the gel and do not go in straight lines. Notice the difference in diameter of the two tunnels. You have to remember that when we work with proteins, we work with many copies of each kind of protein. As a result, the collection of proteins of any given size tends to move through the gel at the same rate, even if they do not take exactly the same tunnels. The electrophoresis is run until the small proteins reach about 0.5 cm from the, at which the current is turned off. The gel is then removed from its cassette and incubate with stain, usually Coomassie Blue (same stain used in the Bradford assay). Denatured proteins are normally colorless and thus invisible. After stain, the gels are incubated with several changes of wash buffer to reduce the background. This leaves stained proteins that appear as bands on the gel (Figure 5) Figure 3: Layer of PAGE gel. After preparation of the gel, the protein samples that have been boiled in SDS and mercaptoethanol are applied to the gel and the system is electrophoresis by turning on the current. If all the proteins enter the gel at the same time and have the same force pulling them towards the other end, which ones will be able to move through the gel faster? Small molecules can maneuver through the polyacrylamide forest faster than big molecules. The cartoon in Figure 4 shows a mixture of denatured proteins (pink lines of different lengths) beginning their journey through a polyacrylamide gel (gray slab with tunnels). An electric field is established with the positive pole Figure 5: A stained PAGE gel showing protein bands in each lane. 8-2
The gel represented in Figure 5 has five lanes. Each lane represents a single sample, with lane 1 being the sized standards used for estimating the relative size of each protein band in the samples (lanes 2-5). For todays lab, the samples are put in order from lane 2 to lane 5 representing: lane 2: crude; lane 3: affinity purified; lane 4: gel filtration purified; lane 5: standard HRP. The photo in Figure 6 shows a variety of different proteins being separated on a gel. This particular image is showing a serial dilution of Figure 6: Photo of an actual gel stained using a silver stain. the same protein sample to indicate how little protein is needed (16 picograms = 16 x 10-12 grams) in order to be detected using a sensitive silver stain. Coomassie stain is less sensitive, and 5-50 ug is typically loaded. Staining the gel: Coomassie Brilliant Blue R 250 is the most commonly used staining procedure for the detection of proteins. It is the method of choice if SDS is used in the electrophoresis of proteins, and is sensitive for a range of 0.5 to 20 micrograms of protein. Staining solution typically contains 50% distilled water, 40% methanol, 10% acetic acid, and 0.1% Comassie Blue (100 mg/100 ml). Destain is the same solution without Comassie Blue. Determination of Molecular Weight This is done by SDS-PAGE of proteins of known molecular weight along with the protein to be characterized. A linear relationship exists between the logarithm of the molecular weight of an SDS-denatured polypeptide and its migration distance. A simple way of determining relative molecular weight by electrophoresis (M R) is to plot a standard curve of distance migrated vs. log MW for known samples, and calculate the log MW of the sample after measuring distance migrated on the same gel. Table 1: Sizes of Molecular Weight Markers (Top to Bottom) There is a caveat to this method to keep in mind. SDS-PAGE separates proteins based on their primary structure size but not amino acid sequence. Therefore, if we had many copies of two different proteins that were both 500 amino acids long, they would travel together through the gel in a mixed band. As a result, we would not be able to use SDS-PAGE to separate these two proteins from each other. 8-3
Procedure: Pre-Lab: No calculations required for this lab Materials Needed From last two weeks: crude HRP, affinity purified HRP, gel filtration purified and concentratned HRP, HRP standard. 1 precast gel, gel apparatus with top, (Two groups per bench will share a gel box) Molecular weight markers 1X SDS Running buffer, 800 ml and antioxidant/ box Microtubes, 5, 500 ul locking Sample treatment buffer Sample reducing buffer Antioxidant, 500 ul /gel apparatus power supply staining bin Coomassie Blue stain: 0.1 % Coomassie Blue R-250, 10% methanol, 10 % acetic acid, 100 ml Destain: 10% acetic acid, 10% methanol Storage: 10% acetic acid Tinfoil Large spatula/gel knife Sample Preparation and Gel Loading 1. Start a 250 ml boiling water bath by filling the beakers almost to the top. Add a boiling stone. Cover with aluminum foil and make 5 holes (one for each sample) in the top by piercing with a yellow tip. Push down the tinfoil so the top is in contact with the water (do this before it is hot!). 2. Prepare samples according to Table 2 below in 500 ul lock top microtubes. Table 2: Sample preparation Sample Volume Sample (ul) Volume Sample Buffer(uL) Volume Reducing Agent (ul) Standards 10 2.5 1.5 Crude 10 2.5 1.5 Affinity 40 10 4 purified Gel filtration 40 10 4 purified Standard HRP 10 2.5 1.5 3. Centrifuge the samples for several seconds to get all the solution mixed and to the bottom of the tube. 4. Lock all five of the microtubes and put them into the holes in the tinfoil covering the boiling water bath and boil for 5 minutes. Be sure the samples are immersed in the boiling water. 5. While the samples are boiling, assemble the gel box as shown in Figure 7 in the unlocked format. 8-4
6. Remove two gels (one per group pair) from their plastic coverings. Insert the gels into the Xcel Mini gel box with both fronts facing the Buffer Core. If only one gel is being used, then add a plastic plate on the opposite side of the Buffer Core. Add a plastic insert BACK behind the plates (see Figure 7). Figure 7: Assembly of the gel box. 7. Clamp the gel assembly into the box by pushing the lever IN (Figure 8). Figure 8: Gel box in LOCKED configuration. 8. Load the upper chamber with 200 ml of 1X SDS Running buffer (it should cover the wells) containing 500 ul antioxidant. Fill the lower chamber with 600 ml of 1X SDS running buffer 9. After the samples have boiled for 5 min, remove the tinfoil and let samples cool for several seconds. Use the medium micropipette and yellow tip to load each sample. Take up the sample, slide the tip inside the Buffer Core and ALONG THE BACK PANEL OF THE GEL into the well assigned (lanes 1-5 for samples 1-5), and dispense to the first stop only. BE CAREFUL TO NOT BLOW AIR INTO THE WELL, i.e., don t use the 2 nd stop of the pipette. 8-5
10. After all samples are added, snap in the gel box top and run at 150 V until the tracking dye almost reaches the bottom of the gel (approximately 35 minutes). Stain and destain the gel. 1. After electrophoresis remove the gel box top, unlock the gel cassette and pull out the gel. With the front plate up, pry apart the cassette at its edges with your fingers or with the gel knife. Pull the front plate (the smaller plate) of the cassette away from the bottom plate, being sure the gel stays with the bottom plate (if not, use a spatula to push it down). Use the gel knife to transfer to a staining bin, 2. Add enough Coomassie stain to cokmpletely cover the gel, cover with tinfoil and label for ID, and place on the shaking platform in the incubator. 3. Shake for 15 min, empty the stain back into its original container (use a transfer pipette to remove). 4. Add destaining solution to cover the gel, cover with tinfoil, and wash 10 min. 5. If bands aren't clear then repeat wash (discard wash in waste). 6. If bands are still not clear, shake overnight and take pictures the next morning. 7. The wash solutions and electrophoresis buffer should be transferred to waste liquids. 8. Make a standard curve using the molecular weight standards and determine the molecular weight of horseradish peroxidase (HRP). Laboratory Report The nominal MW of the standards are listed in the Table1 and are reported from top to bottom of the gel. Identify each standard protein band and provide a table with MW and distance traveled from bottom of the loading well to the front of the band. Prepare a standard curve for the PAGE gel stained with Coomassie Blue. Identify the HRP bands for each sample using the band in the HRP standard (lane 5) for comparision. From the standard curve calculate the MW of HRP for each of the sample lanes.. List the total number of bands observed for each of the lanes in the PAGE gel. Do the purification steps increase the purity of the HRP? How did you decide? Is the combination of affinity purification followed by gel filtration an appropriate combination of column types for the purification of HRP? How did you decide? 8-6