Prepared by: Dr. Tarek M. Zaida
SDS-PAGE Protocol
SDS Gel Electrophoresis
You will need the following reagents:
3x Sample Buffer (10 ml)
10% w/v (3ml) SDS
10 mM (1.6ml) Dithiothreitol, or (1.6 ml) beta-mercapto-ethanol
100 %
1M
(3 ml) Glycerol
(2.4ml) Tris-HCl, pH 6.8
0.005 g
Bromophenol blue
Store at 4C (or keep frozen for long term)
10x Running (separating) Buffer 1L
30.3 g
Tris base
144 g
Glycine
10 g
SDS
1x Running Gel Solution
For different applications increase your desired percentage acrylamide, make up 30 ml of
running gel by selecting one of the following percentages and mixing the ingredients shown
below. After adding APS and TEMED your gel will polymerize fairly quickly, so do not add
these until you are sure you are ready to pour.
7%
10%
12%
15%
H2 O
15.3 ml
12.3 ml
10.2 ml
7.2 ml
1.5 M Tris-HCl, pH 8.8
7.5 ml
7.5 ml
7.5 ml
7.5 ml
10% (w/v) SDS
0.15 ml
0.15 ml
0.15 ml
0.15 ml
6.9 ml
9.9 ml
12.0 ml
15.0 ml
10% (w/v) ammonium persulfate (APS) 0.15 ml
0.15 ml
0.15 ml
0.15 ml
0.02 ml
0.02 ml
0.02 ml
0.02 ml
Acrylamide/Bis-acrylamide
(30%/0.8% w/v)
TEMED
Stacking Gel Solution (4% Acrylamide)
H2 O
3.075 ml
0.5 M Tris-HCl, pH 6.8
1.25 ml
20% (w/v) SDS
0.025 ml
Acrylamide/Bis-acrylamide
0.67 ml
(30%/0.8% w/v)
10% (w/v) ammonium persulfate (APS)
0.025 ml
TEMED
0.005 ml
Pouring the Gels
Choose a percentage acrylamide based on the molecular weight range of proteins you wish to
separate:
% Gel
M.W. Range
7
50 kDa - 500 kDa
10
20 kDa - 300 kDa
12
10 kDa - 200 kDa
15
3 kDa - 100 kDa
Preparing your Sample
Mix your protein 6 µl : 3 µl with the sample buffer. Heat your sample by either:
a) Boiling for 5 minutes (Works for most proteins)
b) 65 ºC for 10 minutes (If you have smearing using the above procedure)
c) 37 ºC for 30 minutes (Membrane proteins or others that do not enter the gel otherwise may
benefit from this type of sample preparation)
Pouring the separating gel
1. Clean glass plates with soap and water, then with ethanol. Assemble the glass plates and
spacers.
2. Transfer the separating gel to a beaker and add APS, then TEMED.
3. Mix quickly, then add the separating gel solution (using a Pasteur pipette or a dropper) to the
center of the plates to a height about 4 cm from the top for the large plates.
4. Quickly add isobutanol to the top of this until the level reaches the top of
the plates.
Note: (Isobutanol will prevent oxygen from getting into the gel which could oxidize it and inhibit
polymerization)!!!
5. The separating gel should polymerize in 10 to 30 minutes.
Pouring the stacking gel
6. Pour off isobutanol. Pour water several times into the gel plate space to rinse off all the
isobutanol. Dry the watery interplate surface with a piece of Whatmann paper.
7. To polymerize the stacking gel, add APS, then TEMED, mix, then pour on top of the
polymerized separating gel.
8. Insert the comb straight on down, then pour a little more stacking gel on the sides of the comb
to fully seal it. Remove any bubbles from underneath the comb, if possible, by moving the comb
gently from side to side so the bubbles get into the space in between and float up.
9. The stacking gel should polymerize in 10 to 30 minutes.
Load the gel
10. Attach the large gel plates containing the polymerized gel to the apparatus via the clips
provided.
11. Pour the electrophoresis “running” buffer into the chamber. The large gel takes about 7 liter.
12. Remove bubbles trapped at the bottom of the glass plates in the large gel with a syringe.
13. Flush the wells with a syringe just before loading to get rid of any unpolymerized
polyacrylamide that may seep in.
14. When loading the gel, load something (1X loading buffer in blank lanes) in every lane and
the dye front will migrate more evenly.
15. Run large thin gels at a constant current of 20 mA. After the dye front enters the separating
gel, you can turn the current up to 30 mA. Where you stop the gel will depend on how big the
smallest protein is that you want to visualize. If you wait until the dye front just flows out of the
gel, it should take about 6 - 7 hours for a large, thin gel to run.
Loading your protein samples on gel
1.
Pipet your sample into the gel adjusting the volume according to the amount
of protein in your sample. If you are going to stain using Coomassie, don't use
much more than 5µg of your protein of interest to get a nicely defined band.
2.
Be sure to include a lane with molecular weight standards (marker).
3.
Now attach your power leads and run the gel at 16 mA & 100 volts until the
blue dye front reaches the bottom. First turn off the power supply, then
remove remove the gel from the chamber and process further - Visualize your
proteins using Coomassie Brilliant Blue.
Preparation of Coommassie Staining buffer (Stain for 1-2 hours)
0.25g Coommassie Brilliant Blue G-250
50% MeOH
10% CH3COOH
40% H2Od
Destaining Buffer (destain 4 – 24 hours)
5% MeOH
7.5% CH3COOH
87.5% H2Od