Western Blot Protocol:

Lysis and Sample Preparation
  1. Add Laemmli buffer to your samples so that the final protein concentration is approximately 1 ug / ul. The protein mass will vary between cell lines and tissues, so it is best if this concentration is determined empirically by a total protein assay (e.g. Bradford Assay, BCA assay).
  2. If you are lysing cells from cell-culture plates, use the tip of your pipette to mix the Laemmli buffer around the well and scrape the bottom of the well to solublize all cells and proteins.
  3. Draw up the Laemmli buffer with your pipette and place into a clean, autoclaved eppendorf tube.
  4. Boil the protein samples for 5 minutes in a water bath. Make sure that you use an object to put downward pressure on the lids of the eppendorf tubes. If not, the heat will increase the pressure inside the tubes to such an extent that the lids will pop off and water will get inside.
  5. After boiling, spin the samples in a table-top centrifuge at approximately 15,000xg for 3 minutes.
  6. Let the samples cool to room temperature before loading the gel.
Gel Casting (Polyacrylamide Gel Recipes , Bis Tris Gel Recipes)
  1. First things first: the best gels are poured and polymerized the evening before the actual run is supposed to take place. Pour the gels the evening of the day before you plan to do the electrophoresis. They can be stored at 4°C overnight as long as they are covered tightly in Saran wrap.
  2. We recommend using a 1.o mm gel because it should give you the most efficient transfer.
  3. Assemble the plates according to your individual manufacturer’s instructions. Normally, there is a glass frame and a cover plate which fit together to form a 1.0 mm space into which the gel is poured.
  4. Label two conical tubes: one with the “stacking” gel, and the other with the specific percentage you’ll be using (e.g. 10%).
  5. Fill each tube with the indicated amounts of water, SDS (standard western protocon only), acrylamide, and buffer, according to the gel recipes.
  6. We recommend making a fresh stock of 10% APS because APS is unstable in water. It’s easy enough to make a 10% APS solution (W/V) in water in an eppendorf tube.
  7. To the resolving gel solution only, add the indicated amounts of APS and TEMED and quickly swirl the conical tube to mix the solution.
  8. Pour the solution into the glass gel frame straight from the conical tube, with the mouth of the tube against the lip of the cover plate.
  9. Fill the frame about 75% of the way so that there will be enough room for the stacking gel.
  10. Overlay n-butanol on top of the solution in the frame while it polymerizes.
  11. The gel should polymerize within an hour. If there is any excess solution in the conical tube, you can use that as a guide for when the resolving gel is done polymerizing.
  12. Once the resolving gel is fully polymerized, dump off the n-butanol and wash away any residual butanol with ddH20 three times.
  13. Wick away any residual water with a clean, dry paper towel.
  14. Set the frame back down and secure it to get ready to pour the stacking gel. Get a clean, 1.0 mm comb ready to insert into the stacking buffer.
  15. Add 10% APS and TEMED to the stacking gel solution according to the gel recipe.
  16. Pour the stacking gel on top of the resolving gel in the frame. Fill it all the way to the top.
  17. Insert the comb, being careful that no air bubbles are trapped between teeth of the comb.
  18. Again, the polymerization will take approximately one hour. Use excess solution in the conical tube as a guide for when the polymerization process is complete.
  19. The best gels are ones that sit overnight at 4°C while wrapped in Saran wrap. This is the recommended method. If the gel must be used that day, allow about two hours to pass from the time the stacking gel is poured.

Gel Running
  1. Make 1x tris-glycine-SDS running buffer. (For a bis-tris gel, use MOPS-SDS running buffer).
  2. Load the gels into the gel holders being sure to keep a tight seal between the gel- cast and the gasket.
  3. Pour the running buffer into the middle of the gels and check for leaks.
  4. Pour the rest of the running buffer into the bottom of the running container.
  5. Remove combs and use a pipette to clean away any unpolymerized acrylamide.
  6. Make a marker for each gel (10 microliters marker + 10 ul blue SDS-buffer)
  7. Load all 20 ul of the marker into the first well.
  8. Load 25 ul of the lysate (1 ug/ul) into the rest of the wells.
  9. Fill any empty wells with Laemmli SDS-buffer.
  10. Set gel to 90v and run for approximately 2 hours.
  1. Remove gel from the gel-cast and place into Transfer buffer.
  2. Cut filter paper in half and soak in transfer buffer (You need six halves per gel.)
  3. Cut a piece of PVDF membrane for transfer and soak in methanol for at least one minute.
  4. Place three pieces transfer-buffer soaked filter paper on the transfer apparatus (roll out any air bubbles every time).
  5. Take the membrane from out of the methanol and rinse it in transfer buffer.
  6. Place the membrane onto the three pieces of filter paper and gently roll out any air bubbles.
  7. Place the gel on top of the membrane in the correct orientation. The marker should be on the same side that it was when you loaded the gel.
  8. Gently roll out any air bubbles.
  9. Place three more pieces of filter paper on top of the gel.
  10. Set the transfer apparatus to 15V for 45 minutes and hit start.
  1. When the transfer is complete, remove the PVDF membrane and place directly in TBST (Be careful not to let the membrane dry at this point.) This step removes any excess methanol from the blot.
  2. Place the PVDF membrane into a fresh tray with your choice of blocking buffer. We recommend 5% BSA in TBST for most applications.
  3. Incubate the membrane in blocking buffer for at least two hours with gentle agitation on a shaker.

  1. Dilute the primary antibody 1:1000 in 5% BSA in TBST.
  2. Remove the western blot membrane from the tray with blocking buffer and place into a fresh, clean tray with the primary antibody solution.
  3. Incubate 2 hours to overnight under gentle agitation on a shaker. If incubating overnight, place the membrane at 4°C.
  4. After primary antibody incubation, wash the western blot membrane 3 time for 5 minutes each with TBST. Be sure to use at least 25 ml of TBST for each wash.
  5. Incubate the washed PVDF memrbane with secondary antibody at a 1:2000 dilution in 2.5% BSA in TBST. The secondary antibody can be diluted further if the signal is too strong. However, for most applications, a dilution of at 1:1000-1:2000 will give the best signal to background ratio for your western blots.
  6. Leave the secondary antibody on for 30 minutes to 1 hour.
  7. Wash the membrane 4 times with TBST with gentle agitation on a shaker. Use at least 25 ml for each wash. After the fourth wash, place your western blot membrane into a fresh, clean tray of TBST.
  1. The ECL should be kept at 4°C. Remove the ECL from the fridge and mix an equal amount of substrate and oxidizing agent in a fresh, clean tray.
  2. Remove the membrane from its tray of TBST and dab the excess buffer off of a corner onto a clean paper towel. Do not let the membrane dry out. Just remove the excess buffer before placing the membrane into the ECL.
  3. Place the membrane into the ECL for 45 seconds.
  4. Remove the membrane from the ECL tray and dab off excess ECL onto a clean paper towel. The membrane should stay wet.
  5. Place the ECL-soaked membrane into a clean cassette between transparent page protectors.
  6. Roll out any air bubbles that may obstruct the film development.
  7. Close the cassette, grab a time and film, and go to the nearest darkroom to develop.
  8. Generally, a good starting exposure time is 30 seconds. This time can be adjusted to get an exposure in the linear range

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