Running Protein Gels

Solutions

10X Running Buffer (0.25 M Tris, 1.92 M glycine, 1% SDS)
121 g Tris
577 g glycine
40 g SDS
ddh20 to 4 L (check pH at 1:10 dilution (pH=~8.8)).

5X Sample Buffer (0.3125 M Tris pH 6.8, 10% SDS, 50% glycerol, 0.005% bromophenol blue, 25% 2-mercaptoethanol)
1.56 ml 2M Tris pH 6.8
1 g SDS
5 ml glycerol
0.2 ml 0.25% solution of bromophenol blue
2.5 ml 2-mercaptoethanol
ddH20 to 10ml.

30:0.8 Acrylamide:bis

0.5 M Tris pH 6.8

10% SDS

10% APS

TEMED

3 M Tris pH 8.8

Pouring Gels

Staining Protocols

Modified from Hoefer Protein Electrophoresis Applications Guide

Standard Staining Solutions

Staining Solution (0.025% Coomassie Brilliant blue R 250, 40% methanol,7% acetic acid)
0.5 g Coomassie Brilliant blue R
800 ml methanol
Stir until dissolved. Then add:
140 ml acetic acid
ddH20 to 2 L
Store at room temperature for up to 6 months.

Destaining Solution I (40% methanol, 7% acetic acid)
400 ml methanol
70 ml acetic acid
ddH20 to 1 L
Store at room temperature ad infinitum.

Destaining Solution II (7% acetic acid, 5% methanol)
700 ml acetic acid
500 ml methanol
ddH20 to 10 L
Store at room temperature ad infinitum.

Standard Coomassie Blue Protocol

1. Place gel in Staining Solution. Shake slowly for 1 hr to overnight.
   

2. Replace the Staining Solution with Destaining Solution I. Shake slowly for 30 minutes.
   

3. Remove Destaining Solution I and replace with Destaining Solution II. Addition of Kimwipes to one corner of the staining tray will help remove Coomassie blue from the gel without changing the destaining solution. Replace tissues when they are saturated with Coomassie blue.
   

4. To minimize cracking, add 1% glycerol to the last destain before drying the gel.

Really Rapid Coomassie Staining Protocol

1. Place gel in staining solution in a small box with a lid. (An empty pipet tip box works well.)

2. Microwave on high for 1 min, then shake 10-20 min.

3. Replace staining solution with Destain I; Add some Kimwipes or a folded-up paper towel to help absorb the stain.

4. Microwave on high for 1 min and shake until the bands emerge clearly from the background.

5. Pour out Destain I, replace with water. The gel will continue to destain a little bit.

6. Dry down the gel when you get around to it.

Really Rapid Silver Staining Protocol

1. Fix: 50% methanol, 10% acetic acid (100 ml).

2. Microwave on high for 1 min, then shake 15 min.

3. Wash: dH₂0 (100 ml).

4. Microwave on high for 1 min, then shake 10 min (or until the gel is rehydrated).

5. Reduce: 5 µg/ml DTT in dH₂0 (100 mls) (i.e. 32.5 µl of 0.1M DTT into 100 mls).

6. Microwave on high for 1 min, then shake 15 min (or until the gel has cooled).

7. Stain: 0.1% AgNO₃ in dH₂0 (100 ml) (Dilute a 10% AgNO₃ stock).

8. DO NOT MICROWAVE. Shake 15 min.

9. Develop: Make fresh developer solution for each gel (200 ml: 40 ml 15% Na₂CO₃ + 160 ml dH₂0).

10. Wash quickly with dH₂0 -- 2 X. Shake while washing.

11. Wash quickly with developer -- 2 X 50 mls.

12. Add 100 µl of 37% formaldehyde to the remaining 100 mls of developer.

13. Pour developer on gel and shake until bands are seen.

14. STOP by adding 5 mls of 2.3 M citric acid (should be bubbling).

15. Shake in STOP for 10 min, then wash out with dH₂0 several times (otherwise the background turns yellow).

16. Soak in 5% glycerol for 15 min or longer before drying gel.

If preparing a sample for mass spectrometry, you may wish to use the protocol found here.

Westerns

Transfer

1. Optional: soak the gel in semi dry transfer buffer (48mM Tris, 39mM glycine, 0.037% SDS, 20% methanol) 10-20’.

2. Cut the membrane (Immobilon-P/PVDF) and 6 pieces of blotting paper (e.g. Schleicher and Schuell GB002) to the same size as the gel (~8.5 x 5.7 cm). Wet the membrane for 15 seconds in 100% methanol, then soak for 2’ in water, then equilibrate the membrane for 5 minutes in transfer buffer.

3. Assemble the transfer stack:

1. Three pieces of blot paper soaked in transfer buffer

2. Gel

3. Pre-soaked membrane

4. Three pieces of blot paper soaked in transfer buffer

4. Place stack upside down (gel side up) on blotter.

5. Blot at 0.8 mA/cm² (~38 mA per gel) for 1-2 hrs.

6. Place the membrane on a piece of blot paper and dry for 2 hrs (or longer) at room temp or soak the membrane in 100% methanol for 10 sec. and dry 15 min.

Antibodies and Chemiluminescence:

1. Wet membrane 5 sec. in 100% methanol, then incubate briefly in TBST (25 mM Tris, 140 mM NaCl, 3 mM KCl, 0.05% Tween-20, pH 8.0).

2. Stain with India Ink (Pelikan, Fount India Black) 0.1% in TBST with 0.02% azide (save and store at 4° C).

3. Wash 2 times with TBST.

4. Block for 1hr at RT or O/N at 4° C with 5% nonfat dry milk in TBST and 0.02% azide (save and store at 4° C).

5. Wash briefly with TBST.

6. Incubate with first antibody in TBST, 2% milk, 0.02% azide (save and store at 4° C) for 1-2 hr at RT (or O/N at 4° C).

7. Quick wash with TBST, then wash 3x 10’ with TBST.

8. Incubate with secondary antibody (1:5000 dilution) in TBST, 2% milk (no azide!) for 20-60’.

9. Quick wash with TBST, then wash 3x 10’ with TBST.

10. Prepare ECL reagents: mix the two reagents 1:1 (~1ml per blot) and pipette ~1ml on a piece of Saran wrap taped to the bench.

11. Drag the blot along the edge of the tray, drain excess TBST, then place the blot with protein side down on the ECL solution.

12. Incubate for 1’ then drain excess reagent and transfer the blot to a plastic report cover.

13. Expose immediately (few seconds — 1 hr).

Stripping

1. After detection wash the membrane 2x 10’ in TBST.

2. Incubate 30’ at 50 deg C in a closed container in stripping buffer (65 mM Tris-HCl pH 6.7, 100 mM beta-mercaptoethanol, 2% SDS). If stripping is incomplete increase temperature to 60 or 70 deg C.

3. Wash 3x 10’ in a large volume of TBST at RT.

4. The membrane is now ready to be blocked.