Western blot protocol

  1. Stain desired cell population. Direct the first sort (Yield mode) into staining media (HBSS-free + 2% bovine serum). I use a 1.5 mL epi tube with 0.5 mL of media to collect the cells, and I transfer them to a FACS tube for the second sort.
  2. Place 50 µL of 100% TCA into a 1.5 mL epi tube. Aim the second sort (purity mode) at the bottom of the epindorf tube. With a 100 micron flow cell, 30,000 cells brings the volume to ~300 µL.
  3. Mix the tube and place on ice. Complete all desired sorts, then bring the volume in each tube up to 500 µL with distilled water and mix again. The final TCA concentration is 10%.
  4. Spin at highest speed in a cold microfuge. Remove the supernatant. The precipitate is barely visible as a faint white film on the wall of the tube.
  5. Wash twice with 1 mL of acetone. Let the pellet stand to dry. It is stable and can be left overnight if desired.
  6. Re-suspend in 12 µL of solubilization buffer (9M urea, 2% Triton X-100, 1% DTT). Make sure to coat the walls of the tube with the buffer since this is where the precipitate settles.
  7. Add 4 µL of 4x LDS sample buffer (Invitrogen). Heat 70°C for 10 minutes and then store at -80°C.
  8. Run samples on NuPage gels (Invitrogen). I think that the Bis-Tris gradient gels used with NuPage MES buffer gives the best separations for proteins under 100 kDa. For large proteins (200-300 kDa), I’ve had better success with the MOPS buffer or the Tris-glycine gels. The invitrogen website has a table showing how each gel and buffer combination separates. Unfortunately, these are not cheap – but they work very well.
  9. Transfer to PVDF membrane using the NuPage transfer buffer. Then proceed with blocking and washing as per any Western blot protocol. All of the phosphospecific antibodies that I use are from Cell Signaling. I use a low concentration of HRP secondary antibody (1:40,000) and I wash very thoroughly after the secondary antibody (sometime overnight). You have to be more careful about the signal to noise ratio than is necessary when using cell line extracts.
  10. To develop the blots, I use the SuperSignal Femto kit from Thermo and the Hyperfilm ECL from GE. These are very sensitive, but they can cause some issues with background if the blot is not washed thoroughly.
  11. To strip blots, I use 1% SDS, 25 mM glycine pH 2 and rock vigorously for 30 minutes. This is more gentle and causes less background than 2- mercaptoethanol based strip buffers. After stripping, I was with distilled water briefly and then re-block. I’ve been able to strip and reprobe as many as 10 times with this method.

Note: If you do this, the order that you probe with various antibodies is important because for stronger antibodies (e.g. total Akt and total S6) the stripping is incomplete. Start with the most important phospho-antibodies: in my case P-Akt and P-S6. Get through those and then do the total protein antibodies. It also helps if you don’t do proteins of similar size on back-to-back days. For example, doing P-Akt (Ser473) then P-S6 then P-Akt (Thr308) allows me to confirm that the Ser473 antibody has stripped off well before I probe with Thr308 antibody.

Alternative protocol

  1. Double sort desired cell population into 0.5 mL staining media (20,000 or 30,000 cells) in an epi tube.
  2. Spin the cells down by placing the epi tube in the FACS centrifuge and spinning at 1500 RPM. Remove all but 10-20 µL of supernatant. The cell pellet is small but visible.
  3. Resuspend in 0.5 mL Iscove’s + 2%FBS and incubate at 37°C for 30 min.
  4. Spin and remove supernatant as in step 2. Resuspend in 1 mL of cold PBS.
  5. Spin and remove all but 10-20 µL supernatant. Resupend in 100 µL cold PBS.
  6. Add the resuspended cells to a fresh tube that already contains 50 µL of 100% TCA and 350 µL water (after adding the cells the final volume is 500 µL of 10% TCA). Mix and proceed to step 4 above