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  Section: Biotechnology Methods » Tools and Techniques in Biological Studies
 
 
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Electrophoresis

 
     
 
Electrophoresis is the migration of charged molecules in response to an electric field. Their rate of migration depends on the strength of the field; on the net charge, size and shape of the molecules, and also on the ionic strength, viscosity, and temperature of the medium in which the molecules are moving. As an analytical tool, electrophoresis is simple, rapid, and highly sensitive. It is used analytically to study the properties of a single charged species, and as a separation technique.

There are a variety of electrophoretic techniques, which yield different information and have different uses. Generally, the samples are run in a support matrix, the most commonly used being agarose and polyacrylamide. These are porous gels, and under appropriate conditions, they provide a means of separating molecules by size. We will focus on those methods used for proteins. These can be denaturing or nondenaturing. Nondenaturing methods allow recovery of active proteins and can be used to analyze enzyme activity or any other analysis that requires a native protein structure. Two commonly used techniques in biochemistry are sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDS-PAGE) and isoelectric focusing (IEF). SDS-PAGE separates proteins according to molecular weight and IEF separates according to isoelectric point. This laboratory exercise will introduce you to SDS-PAGE.
 

 
SDS-PAGE
The gel matrix used is a crosslinked acrylamide polymer. This electrophoretic method separates the proteins according to size (and not charge) due to the presence of SDS. The dodecyl sulfate ions bind to the peptide backbone, both denaturing the proteins and giving them a uniform negative charge.

The gels we will be running use a discontinuous system, meaning that they have 2 parts. One is the separating gel, which has a high concentration of acrylamide and acts as a molecular sieve to separate the proteins according to size. Before reaching this gel, the proteins migrate through a stacking gel, which serves to compress the proteins into a narrow band so they all enter the separating gel at about the same time. The narrow starting band increases the resolution. This part of the gel has a lower concentration of acrylamide to avoid a sieving effect.

The stacking effect is due to the glycine in the buffer, the low pH in the stacking gel, and the higher pH in the running buffer. At the low pH, the glycine has little negative charge, and thus moves slowly. The chloride ions move quickly and a localized voltage gradient develops between the 2. As the gel runs, the low pH of the stacking gel buffer is replaced by the higher pH in the running buffer. This maintains a discontinuity in the pH and keeps the glycine moving forward (any glycine molecules behind would acquire a higher charge and speed up). Since there is no real sieving going on, the proteins (which have intermediate mobility) form a tight band, in order of size, between the slower glycine and the faster chloride ions. The separating gel buffer has a higher pH, so the glycine molecules become more negatively charged and move past the proteins, and the voltage gradient becomes uniform. The proteins slow down in the smaller pore size of the separating gel and separate according to size.

Exercise: You will be given protein molecular weight standards, several different solutions containing individual proteins, and a sample of the same serum you used in the protein quantitation lab. Your job is to determine the molecular weights of the individual proteins and the major components in the serum sample. You will run each sample on 2 gels, one you prepare yourself and a commercial precast gel, and compare the results.

Before doing electrophoresis, you must know the amount of protein in each sample. Determine the protein concentrations of each of your samples using a protein assay before coming to the lab to do any electrophoresis. For this exercise, the only sample of unknown protein concentration is the serum that you used for one of your unknowns last week. The amount of protein to be loaded depends on the thickness and length of the gel, and the staining system to be used. Using the Coomassie Blue staining system, as little as 0.1 mg can be detected, but more will be easier to see. As a guide, use 0.5–5 mg for pure samples (one or very few proteins) and 20–60 mg for complex mixtures where the protein will be distributed amongst many protein bands. Overloading will decrease the resolution.

Protocol: The apparatuses used in gel casting or running electrophoresis vary; make sure you look over the appropriate manuals before you operate.

Caution: Unpolymerized Acrylamide is a Neurotoxin. Be Careful! Do not pour unpolymerized acrylamide down the sink, wait for it to polymerize and dispose of it in the trash.

TEMED (N, N, N', N'-tetramethylethelenediamine) is also not very good for you and is very smelly; avoid breathing it. Open the bottle only as long as necessary, or use it in the hood.

  1. Make sure gel plates are clean and dry. Do not get your fingerprints on them or the acrylamide will not polymerize properly.
  2. Prepare gel solutions (separating and stacking), but do not add polymerizing agents, APS and TEMED (this would start the polymerization).
  3. Lay the comb on the unnotched plate and mark (on the outside, using a Sharpie) about 1 cm below the bottom of the teeth. This will be the level of the separating gel. If available, use an alumina (opaque, white) plate, for the notched plate, as this conducts heat away from the gel more efficiently than glass. Set up the gel plates, spacers, and plastic pouch in the gel casting as described in the manufacturer’s directions. When everything is completely ready, add TEMED to the separating gel solution, mix well, and pour it between the plates, up to the mark. Wear gloves if you pour directly from the beaker. You can also use a disposable pipette. Work quickly or the solution will polymerize too soon. Carefully layer isopropanol (or water-saturated butanol) on top of acrylamide so it will polymerize with a flat top surface (i.e., no meniscus). Do this at the side and avoid large drops, so as not to disturb the gel surface. When the leftover acrylamide in the beaker is polymerized, the acrylamide between the plates will also be ready.
  4. If you are running the gel on the same day, prepare samples while the acrylamide is polymerizing. Otherwise, wait until you are ready to run the gel.
    (i) You will need a sample of each unknown substance, plus the molecular weight standards. Prepare samples in screw-cap microcentrifuge tubes. The protein content should be at 1–50 mg in 20–30 mL sample. The total sample volume that can be loaded depends on the thickness of the gel and the diameter of the comb teeth. For Genei apparatuses, this is ~ 30 mL/well. To prepare the sample, mix 7–10 mL of the sample (depending on protein concentration) +20 mL 2X sample buffer containing 10% b-mercapto-ethanol (BME). Use the BME in the hood - it stinks! For dilute samples, mix 40 mL of the sample and 10 mL 5X sample buffer and add 2 mL of BME. Heat to 90°C for 3 minutes to completely denature proteins. It is important to heat samples immediately after the addition of the sample buffer. Partially denatured proteins are much more susceptible to proteolysis and proteases are not the first proteins to get denatured. (Heat samples to 37°C to redissolve SDS before running the gel if samples have been stored after preparation).
    (ii) If you want the proteins in the sample to retain disulfide bonds, do not add BME. If both reduced and nonreduced samples will be run on the same gel, leave at least 3–4 empty wells between samples, since the BME will diffuse between wells and reduce proteins in adjacent samples.
    (iii) MW Stds: 7 mL of Rainbow stds +10 mL of sample buffer (do not make in advance). Heat to 37°C before use.
  5. After the separating gel has polymerized, drain off the isopropanol. Add TEMED to the stacking gel solution, pour the solution between the plates, and insert the comb to make wells for loading samples. The person putting in the comb should wear gloves. Keep an eye on this while it’s polymerizing and add more gel solution if the level falls (as it usually does), or the wells will be too small.
  6. After polymerization, do not cut the bag; we reuse them. The gel may be stored at this point by taping the bag shut to prevent drying. When ready to run the gel: mark the position of each well, since they are difficult to see when full.
  7. Remove comb and rinse wells with running buffer. See the manual directions for setting up the gels in the buffer chambers. The apparatus can run 2 gels simultaneously. There is a blank plate to use when running only one. Fill the upper chamber with running buffer first and check for leaks. Adjust the plates if necessary. Load the samples using a micropipettor with gel-loading tips (these are longer and thinner than the normal tips). This will be demonstrated. Do not load samples in the end wells. Make sure to write down which sample was loaded in each well.
  8. Electrophoresis (takes 1–2 hours).
    Connect the gel apparatus to the power supply and run at 15 mA/gel until the tracking dye (blue) moves past the end of the stacking gel. Increase the current to 20–25 mA/gel but make sure the voltage does not get above 210 V. Run until the blue tracking dye moves to the bottom of the separating gel. For the BioRad apparatus, do not exceed 30 mA, regardless of the number of gels.
  9. Disassemble the apparatus and carefully separate the gel plates using a flat spatula.
    Cut off the stacking gel and any gel below the blue tracking dye. Note the color of each of the molecular weight standards, as they will all be blue after staining. Wash 3X with distilled water.
    Place the gel in a plastic staining container and add Coomassie Blue staining solution. Keep it in this 1 hour overnight. Wash again with water. You can wrap the gel in plastic wrap and Xerox or scan it to have a copy. The gel may also be dried.

Data Analysis
Measure the length of the gel (since you cut off the bottom, this is the distance traveled by the dye).

Measure the distance traveled by each of the molecular weight standards. Measure the distances of each unknown band.

For samples lanes with many bands (serum in this exercise), measure all bands in those with just a few and the major bands in those that have many.

Prepare a standard curve by plotting log MW versus relative mobility (Rf, distance traveled by protein divided by distance traveled by dye). Use this and the mobility of bands from your fractions to determine the MW of the unknown proteins. (Review standard curves from the protein quantitation lab if necessary.) MW of proteins that do not run very far into the gel or run near the dye front will not be accurate.

If you have reduced and unreduced samples, compare the number of bands and MW of each to determine the number of subunits.

Gel Solutions
  1. Separating gel: (15 mL, enough for two gels) 10% acrylamide. 40% Acrylamide/bisacrylamide mix 3.55 mL.
    1.5 M tris pH 8.8, 3.75 mL, H2O 7.4 mL, 10% SDS 150 mL, 10% ammonium persulfate (APS) 150 mL (prepared fresh), TEMED 6 mL.
  2. Stacking gel: (5 mL) 5% acrylamide. Compresses the protein sample into a narrow band for better resolution. 40% Acrylamide/bisacrylamide mix 0.625 mL. 0.5 M tris pH 6.8, 1.25 mL, H2O 3.0 mL, 10% SDS 50 mL, 10% APS 50 mL, TEMED 5 mL.
  3. 2X sample buffer (10 mL)—store in the freezer for an extended time.
  4. SDS must be at room temperature to dissolve
  5. H2O 1.5 mL, 0.5 M Tris pH 6.8, 2.5 mL, 10% SDS (optional) 4.0 mL, glycerol 2.0 mL, BPB 0.01%, b-mercaptoethanol (optional) 0.1 mL.
  6. Running buffer(5L) 30 g Tris Base, 144 g glycine, dissolve in sufficient H2O to make 1.5 L and put into final container.
    Add 1.5 g SDS (Caution: do not inhale dust).
When adding SDS, avoid making too much foam, which makes measuring and pouring difficult.

Final pH should be around 8.3, but do not adjust it or the ionic strength will be too high and the gel will not run properly. If the pH is way off, it was made incorrectly or is old and has some contamination.

The running buffer can also be made more concentrated (5X or 10X) and diluted as needed to save bottle space.
 
     
 
 
     




     
 
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