Preparation of Polyacrylamide Gels

Role of Reagents Involved
Reagents. Acrylamide and N, N’ -Methylene bisacrylamide, a stock solution containing 29% (w/v) acrylamide and 1% (w/v) N, N’ Methylene-bisacrylamide, should be prepared in deionized, warm water (to assist the dissolution of the bisacrylamide. Check that the pH of the solution is 7.0 or less, and store the solution in dark bottles at room temperature. Fresh solutions should be prepared every few months.

A 10% stock solution of sodium dodecyl sulfate (SDS) should be prepared in deionized water and stored at room temperature. Tris buffers for the preparations of resolving and stacking gels—it is essential that these buffers be prepared with Tris base. After the Tris base has been dissolved in deionized water, the pH of the solution should be adjusted with HCl

TEMED (N, N, N’, N’-tetramethylethylenediamine)—TEMED accelerates the polymerization of acrylamide and bisacrylamide by catalyzing the formulation of free radicals from ammonium persulfate.

Ammonium persulfate—Ammonium persulfate provides the free radicals that drive polymerization of acrylamide and bisacrylamide. A small amount of a 10% (w/v) stock solution should be prepared in deionized water and stored at 4°C. Ammonium persulfate decomposes slowly, and fresh solutions should be prepared weekly.

Tris-glycine electrophoresis buffer—This buffer contains 25 mM Tris base, 250 mM glycine (electrophoresis grade) (pH 8.3), 0.1% 50S. A 5X stock can be made by dissolving 15.1 g of Tris base and 94 g of glycine in 900 mL of deionized water. Then 50 mL of a 10% (w/v) stock solution of electrophoresisgarlic 50S is adjusted to 1000 mL with water.

Casting of 50S-Polyacrylamide Gels
Assemble the glass plates according to the apparatus manufacturer’s instructions. Determine the volume of the gel mold (this information is usually provided by the manufacturer). In an Erlenmeyer flask, prepare the appropriate volume of solution containing the desired concentration of acrylamide for the resolving gel. Polymerization will begin as soon as the TEMED has been added. Without delay, swirl the mixture rapidly.

Pour the acrylamide solution into the gap between the glass plates. Leave sufficient space for the stacking gel (the length to the teeth of the comb plus 1 cm). Using a Pasteur pipette, carefully overlay the acrylamide solution with 0.1% 50S (for gels containing 8% acrylamide) or isobutanol (for gels containing ≈ 10% acrylamide). Place the gel in a vertical position at room temperature.

For the large-scale isolation method, after polymerization is complete (30 min), pour off the overlay, and wash the top of the gel several times with deionized water to remove any un-polymerized acrylamide. Drain as much fluid as possible from the top of the gel, and then remove any remaining water with the edge of the paper towel.

Prepare the stacking gel as follows: in disposable plastic tubes, prepare the appropriate volume of solution, containing the desired concentration of acrylamide. Polymerization will begin as soon as the TEMEO has been added. Without delay, swirl the mixture rapidly and proceed to the next step. Pour the stacking gel solution directly onto the surface of the polymerized resolving gel. Immediately insert a clean Teflon comb into the stacking gel solution, being careful to avoid trapping air bubbles. Add more stacking gel solution to fill the spaces between the combs completely. Place the gel in a vertical position at room temperature.

While the stacking gel is polymerizing, prepare the samples by heating them to 100°C for 3 minutes in 1 × 50S gel loading buffer to denature the proteins.

1 × 50S gel loading buffer
50 mM Tris-CI (pH 6.8)		1.2 mL
100 mM dithiothreitol/p mercaptoethanol		0.95 mL
2% 50S (electrophoresis grade)		2 mL
0.1 % bormophenol blue		0.5 mL
10% glycerol		1 mL

1 X SDS gel-loading buffer lacking dithiothreitol/p mercaptoethanol can be stored at room temperature. Dithiothreitol/p-mercaptoethanol should then be added, just before the buffer is used, from a 1-M stock.

After polymerization is complete (30 minutes), remove the Teflon comb carefully. Wash the wells immediately with deionized water to remove any unpolymerized acrylamide. Mount the gel in the electrophoresis apparatus. Add Tris-glycine electrophoresis buffer to the top and bottom reservoirs. Remove the bubbles that become trapped at the bottom of the gel between the glass plates. This is best done with a bent hypodermic needle attached to a syringe.

Load up to 15 mL of each of the samples in the predetermined order into the bottom of the wells. Load an equal volume of l X SDS gel-loading buffer into any wells that are unused. Attach the electrophoresis apparatus to an electric power supply (the positive electrode should be connected to the bottom buffer reservoir). Apply a voltage of 8 V/cm to the gel. After the dye front has moved into the resolving gel, increase the voltage to 50 V/cm and run the gel until the bromophenol blue reaches the bottom of the resolving gel (about 4 hours). Then turn off the power supply.

Remove the glass plates from the electrophoresis apparatus and place them on a paper towel. Using a spatula, dry the plates apart. Mark the orientation of the gel by cutting a comer from the bottom of the gel that is closest to the leftmost well (slot 1). Important: do not cut the comer from gels that are to be used for Western blotting.

The gel can now be fixed, stained with Coomassie Brilliant Blue, fluorographed or autoradiographed, or used to establish a Western blot.

Staining of SDS-Polyacrylamide Gels
Polypeptides separated by SDS-polyacrylamide gels can be simultaneously fixed with methanol: glacial acetic acid and stained with Coomassie Brilliant Blue R250, a triphenylmethane textile dye also known as Acid Blue 83. The gel is immersed for several hours in a concentrated methanol/acetic acid solution of the dye, and excess dye is then allowed to diffuse from the gel during a prolonged period of destaining.

Dissolve 0.15 g of Coomassie Brilliant Blue R250 in 90 mL of methanol: water (1:1 v/v) and 10 mL of glacial acetic acid. Filter the solution through a Whatman No.1 filter to remove any particulate matter.

Immerse the gel in at least 5 volumes of staining solution and place on a slowly rotating platform for a maximum of 4 hours at room temperature. Remove the stain and save it for future use. Destain the gel by soaking it in the methanol/acetic acid solution (step 1) without the dye on a slowly rocking platform for 4–8 hours, changing the destaining solution 3 or 4 times. The more thoroughly the gel is destained, the smaller the amount of protein that can be detected by staining with Coomassie Brilliant Blue. After destaining, gels may be stored indefinitely in water containing 20% glycerol.