E63 Western Blot
updated 03/15/1997 - Martina
(For the PDF version, click here.)
SDS-PAGE
1. Clean the gel plates with EtOH.
2. Pour the separating gel. Leave about 1 cm space below the comb for stacking gel. You’ll need about 4 ml of separating gel/ minigel. Overlay carefully with H2O, and let polymerize for about 20 minutes.
3. Pour off the H2O, insert the comb, and pour the stacking gel. Less than 1 ml necessary/minigel. Let polymerize for about 20 minutes before runing the gel. Alternatively, wrap in the wet paper towel and Saran Wrap and store at 4°C (after 2 days it is still OK, haven’t tried longer storage).
4. If you are running two gels at the same time, start at 36 mA. After samples enter the separating gel, increase to 46 mA or more, depending on how fast you want to be done (with 36mA --> 46 mA current, you will be done in about 1 hour).
Recipes:
| 12% gel, 10 mL | |
| H2O | 3.3 mL |
| 30% acrylamide mix | 4.0 mL |
| 1.5 M Tris, pH 8.8 | 2.5 mL |
| 10% SDS | 0.1 mL |
| 10% APS | 0.1 mL |
| TEMED | 0.004 mL |
| 4 mL stack gel | |
| H2O | 2.7 mL |
| 30% acrylamide mix | 0.67 mL |
| 1 M Tris, pH 6.8 | 0.5 mL |
| 10% SDS | 0.04 mL |
| 10% APS | 0.04 mL |
| TEMED | 0.004 mL |
Solutions:
30% acrylamide
purchased from BioRad, stored in 4°C fridge.
1.5 M Tris
pH 8.8
1 M Tris
pH 6.8
10% SDS
dissolve 10 g of SDS in 90 mL of H2O.
bring the volume to 100 mL.
store @ room temperature.
10% APS
dissolve 10 g of APS in 100 mL of H2O.
bring the volume up to 100 mL.
aliquot in 1 mL aliquots.
store in -20°C.
TEMED
purchased from GIBCO, stored at 4°C.
10x Running Buffer (1 L)
| H20 | 800 mL |
| Tris-base | 30.3 g |
| Glycine | 144.2 g |
| SDS | 10 g |
adjust volume to 1 L.
store @ room temp.
Blotting
1. Take down the gel, cut off the stacking part, and soak in blotting buffer for about 15-30 minutes. Too long might be bad, because proteins can start to leech out (at least that’s what I’ve heard).
2. While the gel is soaking, cut out a piece of membrane exactly the size of a gel (for our minigel, its about 5x8 cm, depending on the size of your stacking gel). If working with nitrocellulose, soak briefly in dH2O, and then in blotting buffer till ready to assemble the sandwich. If using the PVDF membrane (Immobilon P), soak in methanol/ 5 seconds, then dH2O/ 5 minutes, then blotting buffer/15 minutes or till ready to assemble sandwich.
3. Cut out two pieces of Whatman 3MM paper slightly larger then gel (it really doesn’t matter, as long as they fit into the blotting unit). Soak the sponges in blotting buffer.
4. Sandwich. Remember to keep track of the position of gel and membrane (gel must be closer to the - electrode, while membrane to the +.) Wet one piece of Whatman 3MM paper and slide the gel onto it. Place on the sponge. Place the membrane over the gel, roll over with a glass pipette to get rid of air bubbles. Wet the second piece of a Whatman 3MM paper and place on top. Roll over with a pipette. Top with second sponge and place the assembled sandwich into the blotting unit.
5. Blot at 50V/1 hr. For E63-1, this time ensures complete transfer of the protein from the gel to the membrane. 30 minutes is sufficient to get enough E63-1 for immunodetection, but at least half of the protein is left behind in gel (you can check by staining the gel with Commassie blue). For CaM, 30 minutes is sufficient for complete transfer. You don’t have to worry about using cold room under these conditions, the unit will not overheat if you start with cold buffer.
| Western Blotting Buffer | ||
| 3 L | 1 L | |
| Tris-base | 9.09 g | 3.03 g |
| Glycine | 43.23 g | 14.41 g |
| Methanol | 600 mL | 200 mL |
| H2O | up to 3 L | up to 1 L |
| store @ 4°C. |
Immunodetection
1. Take down the blot. If you are using pre-stained MW standards, you will be able to tell whether protein transfer to the membrane was complete. While the membrane is still laying on top of a gel, use a black “BICK” pen to mark positions of MW standards (the blue color tends to fade during following procedures. This can be used later to determine which is the “protein side” of the membrane.)
2. Block 1 hr/RT in 5% (w/v) dry milk in TPBS [PBS with 0.05% (v/v)Tween 20].
3. Wash 3x5 minutes in TPBS.
4. 1° Ab 1hr/RT. For R4694, use 1:50 000 dilution in 5% dry milk in TPBS.
5. Wash 3x5 minutes in TPBS.
6. 2° Ab 30 min/RT. For GAR-HRP (Jackson), use 1:7 500 dilution in TPBS.
7. Wash 2x5 minutes in TPBS, then 2x5 minutes in PBS. Visualize antibody staining with one of the following techniques.
| 10x PBS (1 L) | |
| NaCl | 80 g |
| KCl | 2 g |
| Na2HPO4 | 14.4 g |
| KH2PO4 | 2.4 g |
adjust pH to 7.2.
adjust volume to 1 L.
divide into 4x 250 mL aliquots (500 mL bottles).
autoclave and store @ room temp.
10x TBPS
add 2.5 mL of Tween 20 to 500 mL of 10x PBS.
store at room temp.
ECL Detection (Amersham ECL Kit)
1. Mix solutions #1 and #2 from the kit in equal amounts (for 2 miniblots, I use total volume of 10 mL).
2. Remove blot from PBS, dry excess PBS on paper towel. Place the blot into a small container (protein side up) and pour over it the prepared solution.
3. Gently move for 1 minute. Make sure the whole surface of the blot is covered.
4. Remove blot, dry excess solution, and wrap the blot using Saran Wrap. Tape a piece of a bench paper into a film cassette (to prevent contamination of cassette in case of leakage of solution) and tape the blot (“protein side” up) to it. Alternatively, tape the blot to piece of cardboard, you don’t really need a cassette.
5. In a darkroom, place a film over the blot. If you can see the blot glowing in the dark, try 10 second exposure (be sure to put film down without moving it around too much, otherwise you’ll get smeared bands). Otherwise start with a 1 minute exposure and adjust time according to what you get. WARNING: Our film is sensitive to the orange light bulb in the darkroom.
Note: Save the blot wrapped at 4°C. As long as it doesn’t dry out, it can be stripped and re-probed with a different antibody.
DAB/NiCl/H2O2 detection
1. Dissolve 10 mg of DAB (1 tablet, Sigma) in 20 mL PBS. Might be necessary to warm up the solution (37°C).
2. Add 400 µL of 4% NiCl.
3. Immediately before adding to blot, add 20 µL of 30% H2O2.
4. Pour over the blot and let the bands develop to the desired intensity. Stop the reaction by rinsing the blot in dH2O. Dry the blot between paper towels.
