Polyacrylamide Gels

Setting the Gel

Take a glass plate and a white plate from the draining try in the sink, get the white spacers (black if you want thicker gel) and put in a grey clamp. Push the sandwich down a little bit from being flush with the clamp so there is a greater seal on the base. Mark ~1 cm below the bottom of the well to denote the size of the stacking layer. Place the clamped "sandwich" on a stand to stabilize.

Reagent	Volume (Separation Layer)	Volume (Stacking Layer)	Notes
MilliQ	2.088 ml	1.854 ml
1.5 M Tris-Cl, pH 8.8	1.5 ml	n/a	Don't use this in the stacking layer; kept in cabinets above gel boxes
0.5 M Tris-Cl, pH 6.8	n/a	0.75 ml	Don't use this in the separation layer;kept in cabinets above gel boxes
10% SDS	60 μl	30 μl	kept in cabinets above gel boxes
40% Bis:Acrylamide	2.25 ml	0.3 ml	kept in fridge in adjacent bay
TEMED	9 μl	6 μl	Add this reagent last, then pour gel immediately after;kept in cabinets above gel boxes
10% APS	90 μl	60 μl	Add this reagent last, then pour gel immediately after; Keep an aliquot in the -20, 0.1 g in 1 ml H₂0
Total	5.997 ml	3 ml

Pour the separation mix first up to the line you've marked. After filling to the appropriate volume, take 1 ml of water and squirt over the top of the gel to get rid of bubbles and prevent diffusion of the monomer out of the gel.

About 10 minutes later, you can pour off the water and use a piece of paper to remove droplets. Now add the TEMED and APS to the stacking mix and pour the mix on top of the separation layer. Place the comb in the top.

To run the gel, release the clamps and remove the comb. Clean off the area above the wells to remove stray chunks of gel. If you haven't already, mark the bottom of the wells with a pen so they are easier to find when you are ready to load samples.

Remove the "sandwich" and place in the gel dock using clamps. Put SDS buffer in the upper and lower reservoirs (the buffer should reach the top of the wells in the top chamber and the indentation in the bottom chamber), then use a pipette to flush out the wells with SDS buffer.

If not added already, add 66 μl βME to 1 ml of 6x loading buffer. Add the buffer to samples so that the final dilution is 1/6. Mix well, spin down, and heat on heat block at 95^oC for 5 minutes. Spin down and then add to wells. Use 5 μl of marker (SM0431) (kept in the antibiotic box).

Run the gel at 16 mA until dye has reached separation layer, then run at 32 mA until the dye has reached the bottom of the gel. It's permissible to run the gel at 16 mA for the duration of the gel.

After the gel has finished running, disconnect the apparatus and remove the "sandwich" from the clamps. Separate the sandwich by first pushing one of the spacers to expose, and then torquing it to separate the glass from the white plastic revealing the gel.

Use a spacer to separate the separation layer from the stacking layer and place the former in a plastic container with 0.3% Coomassie Blue. Place the gel on a shaker. Staining time varies, but as a rule of thumb, the gel should be completely blue (no longer translucent) when fully stained.

To destain, dump out the coomassie blue solution, rinse the gel in water, then add destaining solution:

Destaining time varies as well, but when done, the gel should be translucent once more while protein bands should show up clearly in blue. You can add paper towel to better soak up the dye.

The scanner is in wing A. Turn on the scanner first (all three lights should be solid), then the computer. Place the gel on the scanner on the side nearest you with the top of the gel nearest you.

Select the GS-???? instrument from the menu on the right of the screen, then select the dye used to stain the gel and the "translucent" setting.

Run notes will generally be recorded with the particular protocol for which the gel is needed, but independent gels will be recorded here.