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Below is the protocol I am using.
Lysis Buffer: 1mM EDTA, 1mM EGTA, 10mM Tris, pH 7.5, 150mM NaCl, 1% Triton x-100, 0.5% Nonidet P-40, 0.2mM Sodium Orthovanadate,
2.5mM Sodium Pyrophosphate, 1mM Glycerol 2-phosphate Disodium Hyd
At the time of use:
Add 10 µl Halt Phosphatase Inhibitor Cocktail and
Add 4 µl PMSF (1mM) to 1ml Lysis Buffer.
Sample prep and Gel electrophoresis (`14 hours)
- Prepare 7.5 % Substrate separating gel with water that has 10 mg of gelatin per 4.0 mL.
- Add butanol on top of gel to get rid of the bubbles - Polymerize for 2 hours
- 4% stacking Gel (~1 Hour).
- Load 2 ug (medium) or 50 ug (tissue) of protein. Add water to the necessary sample to give a constant volume for each sample. Final volume 30 uL (15 of which is 2xNRB)
- Add an equal volume of non-reducing buffer (do not boil)
10% SDS, 4% Sucrose, 0.1% Bromophenol Blue, 0.25 M Tris-HCl pH 6.85 ml stacking gel buffer qs to 10 ml.
- Run gel at 125 volts in 4° until the samples until the dye reaches the very bottom of the gel (~90 min).
- Shake gel with 2.5% triton (2.5mL/100mL) (with 0.02% NaN3 – 20 mg) 2 X for 30 min each to remove SDS and renature proteins.
- Replace triton with dd water and shake for 2 x 10 min. Remove water and shake with in-gel enzyme buffer. This removes the triton. Rinse with a small amount of incubation buffer
- Place gel at 37⁰C in incubation buffer for 20 hours
Next Day
- Rinse for 5 mins with DI water.
- Take a picture of your gel while the markers are still visible.
- Replace with fresh commassie blue and shake for 30 min at room temp
- Replace with destain for 30 min (or longer) until you can see clear bands
