SDS Polyacrylamide Gel Electrophoresis

CHP - updated: Oct. 29, 1998


1. Set up gel plates.

Square back plate l-15 cm X w-16 cm and 1 notched plate.

0.75 mm spacers: 2 shorter spacers ~14 cm long, and 1 longer spacer ~18 cm long.

Place two short spacers on the sides and the long spacer along the bottom of one glass plate.

(These spacers were cut for a larger piece of glass, so ensure the flat (uncut) end is aligned against the other spacer.

• Clamp using either 1- or 2-inch binder clamps.

Use two clamps at the bottom (on the spacer, not on the gel area)

Use one of the 2 inch clamps or two of the 1 inch clamps on each side; ensure the joint of the bottom and side spacers is clamped.


2. Seal the gel with 2 % agar (bactoagar not expensive agarose).

Melt in microwave 1 min. on medium (or till melted).

Drip the agar (using Pasteur pipet) along the seams of the two glass plates (except the top).

The "joint" of the side and bottom spacer is most likely to leak; it needs to be sealed well.

• Let one side sit a few sec. to harden; seal other side, let it harden a few sec.

• Seal bottom adding extra drop of agar at the side bottom spacer interface. One can prop the sealed gel upside down in the white racks.


3. Mix the Lower or Separating gel; see recipes (note they will make exactly two lower gels).

• Mix the buffer, water, and acrylamide (in the best of all worlds, degas, since acrylamide polymerase is inhibited by O2).

• Add 10 % ammonium persulfate (APS) [found in the desiccator in the weight room] (made within one week) and TEMED. Swirl.

• Pour so the level is about 1 cm below the comb. Use a 10 ml pipet.

Try not to trap any bubbles.

• Prop flask containing excess gel solution at an angle; this helps to determine when the gel has polymerized.


4. Gently layer top with water or n-butanol on top.

Acrylamide polymerization is inhibited by O2; this layering ensures a flat interface.

• Polymerization should occur within 30 min.

This can be observed by the "gel-ing" of the gel solution in the flask or when a defined line is observed between the water and lower gel.


5. Prepare to pour the upper / stacking gel.

• Pour off water layer: wipe the area between the glass plates with filter paper.

Try not to touch the gel.


6. Mix Upper /Stacking gel.

• Mix the water, buffer and acrylamide, then add 10% APS and TEMED, swirl.

• Either place comb in, then tip gel and pour gel in slowly to avoid trapping bubbles. (Cammie's choice),

• Or pour the stacking gel and put in the comb.

• Prop flask at an angle to help determine when the gel has polymerized

• Polymerization should occur in 30 min,

Look for defined line around the wells.


7. Setting up the Gel

• Take the clamps off .

• Take out the bottom spacer, get out as much agar as you can, as it may cause problems with trapped bubbles.

• Before taking out the comb, you may consider marking the well positions.

• Take out comb by gently pulling straight out. To get out any unpolymerized acrylamide, wash wells with running buffer, using a syringe with a 38-gauge needle.

• Clamp gel to the gel boxes using large binder clamps.

At the "ears" of the glass plates so it's clamped tightly to the gaskets.


8. Pour running buffer in.

• To get bubbles out from the bottom area between the glass plates, use a 10 ml syringe with a bent 16 or 18 gauge needle. Fill syringe with buffer, push along the bottom of the gel with the point of the needle between the plates while expelling buffer. This will "chase" the bubbles out.

These bubble interfere with the electrical current conduction

Some people have sucess with filling the bottom reservoir and putting the gel in at an angle to decrease the amount of trapped bubbles.


9. Loading gels

• We use a 100 or 50 ul Hamilton syringe, rinsing well between samples (6x in the bottom reservoir). The 20 well combs hold between 30-50 ul depend how deep you put them in. Gel loading tips can be used.

• Insert point of the needle near the bottom of the well, slowly load sample, as you pull the needle out.


10. Running gels

• Attach leads: the SDS-coated (negatively charged) protein will run to the positive red lead

- red on the bottom black on the top.


11. Settings

• SDS gels are run at 20 mAmps/gel (these 15x16 cm gels)

• The heat generated by the combination of current and voltage is less during stacking; if your protein is sensitive to this, use these settings. We use this with tailspike, it decreases the thermal unfolding of the N -terminus in the presence of SDS, decreasing the band below native tailspike. (Bao-Lu's species)

• For other applications one can use 150 volt total, not matter the number of gels (the current will adjust accordingly).


12. Run till dye front comes off.


13. Stain -- King Lab Method (from Myoung Hee Yu).

30 min in stain #2:

10% isopropanol

10% acetic acid

0.003 % Coomassie Blue


1 hr or longer in stain #3:

10% acetic acid

0.003 % Coomassie Blue


Destain as necessary:

10% acetic acid

1 kimwipe to absorb the stain