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Western Blot Protocols

By Rachel Stewart, PhD

Western blotting involves the electrophoretic separation of proteins from a complex mixture based on their mass, the transfer of these proteins to a solid matrix, and the detection of specific proteins of interest on the matrix using antibodies. Below are example protocols for harvesting and preparing various types of tissue for the purpose of protein separation and blotting using chemiluminescent detection.

Lysate Preparation

Lysate from adherent culture cells

  1. Aspirate the culture medium from the cells, place the dish on ice, and wash twice with cold PBS.
  2. Add cold lysis buffer to the cells (1 ml / 107 cells in a 100 mm dish or 500 µl / 5x106 cells in a 60 mm dish).
  3. Scrape the cells from the bottom of the dish with a cold cell scraper and transfer the suspension to a cold microcentrifuge tube on ice.
  4. Continue lysis on ice with frequent agitation for 5 – 30 minutes. Alternatively, perform sonication or use a Dounce homogenizer to lyse the cells.
  5. Centrifuge the lysate at 4°C for 15 minutes at ~14,000 x g; the time and speed may vary depending on the cell type.
  6. Transfer the supernatant to a new tube on ice and discard the pellet.

Lysate from suspension culture cells

  1. Pellet cells in the culture medium at 2,000 x g for 5 minutes at 4°C.
  2. Aspirate the supernatant and wash the cells twice with cold PBS on ice.
  3. Resuspend the pelleted cells using lysis buffer.
  4. Continue with the lysis protocol for adherent cells from Step 4.

Lysate from tissue samples

  1. Dissect the tissue sample and immediately snap freeze in liquid nitrogen; store at -80°C for later use or proceed to the next step.
  2. Grind the sample in the presence of liquid nitrogen in a cooled mortar before transferring 1 g of the powdered tissue to a microcentrifuge tube on ice containing 1 ml cold lysis buffer. Alternatively, homogenize 1 g of tissue in 1 ml cold lysis buffer on ice until sufficiently disrupted.
  3. Continue lysis on ice with frequent agitation for 5 – 30 minutes.
  4. Centrifuge the lysate at 4°C for 30 minutes at ~35,000 x g.
  5. Transfer the supernatant to a new tube on ice and discard the pellet.

  1. Determine the total protein concentration for each lysate using BCA, Bradford, or Lowry assays, as appropriate.
  2. Dilute the lysates 1:1 with 2X Laemmli buffer with freshly added β-mercaptoethanol or DTT, and vortex thoroughly.
  3. Denature and reduce the samples by heating at 95°C for 5 minutes or 70°C for 10 minutes, and vortex thoroughly.

  1. Assemble the electrophoresis apparatus, insert the gel cassette, and fill both inner and outer chambers with running buffer according to the manufacturer’s instructions.
  2. Slowly load a molecular weight marker / ladder and the appropriate amount of sample(s) containing sample buffer to the wells of the gel. Use 20 – 30 µg of protein per well as a starting point.
  3. Connect the electrophoresis apparatus to the power supply and run according to the manufacturer’s recommendations. For the first 10 minutes of the run, use 5 – 10 V/cm of gel as the samples concentrate in the stacking gel. Once the dye front moves into the resolving gel, increase the voltage to ~100 V or as recommended by the manufacturer.
  4. Run the gel until the dye front runs off the bottom of the gel (~1 hour).

  1. Ensure the transfer buffer is pre-chilled at 4°C.
  2. Cut a piece of membrane and two pieces of filter paper to the size of the gel.
  3. If using a PVDF membrane, pre-wet the membrane in methanol for 2 minutes and then wash for 30 seconds in 1X transfer buffer. If using nitrocellulose, pre-treatment is not required. Carefully use tweezers to handle the membrane.
  4. Equilibrate the gel in transfer buffer for 10 minutes. To prevent the gel tearing, it may be easiest to move the gel into the transfer buffer while it is still attached to the glass cast / cassette.
  5. Soak the membrane, two pieces of absorbent filter paper, and two sponges in transfer buffer.
  6. Sequentially assemble the transfer sandwich in a shallow dish with transfer buffer, ensuring all components are wet. Start from the back, which will contact the negatively charged electrode (cathode). Layer a sponge, filter paper, the gel, the membrane, and a second piece of filter paper, building toward the positively charged (anode) electrode at the top.
  7. After sandwiching the gel and membrane between filter paper, use a roller or pipette to gently roll over the sandwich to remove air bubbles between the gel and membrane, which can impede transfer efficiency.
  8. Add the final sponge to the top of the stack and firmly lock the sandwich together.
  9. Fit the sandwich into the transfer cassette, ensuring the membrane side of the sandwich faces the anode and the gel side faces the cathode. Perform a wet or semi-dry transfer according to the manufacturer’s instructions. For wet transfer, run at 100 V for 1 – 2 hours or 30 V overnight at 4°C.

  1. Block the membrane in ~10 ml blocking buffer (5% BSA / TBST or 5% non-fat milk / TBST) in a small box for 1 – 2 hours at room temperature with gentle agitation.
  2. Incubate the membrane with the primary antibody diluted in TBST, with or without additional blocking agent, for 2 hours at room temperature or overnight at 4°C. Refer to the antibody’s datasheet for recommended dilution ranges. If using a heat-sealed plastic bag for the incubation, ~2 ml of primary antibody solution may be sufficient.
  3. Wash the membrane three times with 20 ml of TBST for five minutes each with gentle agitation.
  4. Incubate the membrane with the secondary antibody diluted in 10 ml TBST for 1 hour at room temperature. Refer to the antibody’s datasheet for the recommended dilution, but 1:5,000 is typical for HRP-conjugated secondary antibodies.
  5. Wash the membrane three times with ~20 ml of TBST for five minutes each with gentle agitation.
  6. If using an HRP-conjugated secondary antibody, use a chemiluminescent substrate kit for detection. Refer to the manufacturer’s instructions to prepare the substrate. Place the membrane on a plastic sheet, apply the substrate solution, and image the chemiluminescent signal with a CCD camera-based imaging system.

Mild Stripping

  1. Incubate the membrane in mild stripping buffer for 10 minutes with gentle agitation.
  2. Replace with new buffer and incubate for 10 minutes with gentle agitation.
  3. Wash with two changes of PBS for 10 minutes each.
  4. Wash with two changes of TBST for 10 minutes each.
  5. Proceed with blocking and antibody incubation steps.

Harsh Stripping

  1. Warm the harsh stripping buffer to 50°C in a fume hood.
  2. Incubate the membrane with the buffer for 45 minutes at 50°C with gentle agitation in a fume hood. Dispose of β-mercaptoethanol appropriately.
  3. Rinse the membrane under running water for 1 minute.
  4. Wash with three changes of TBST for 5 minutes each.
  5. Proceed with blocking and antibody incubation steps.

Buffers and Reagents

RIPA

  • 50 mM Tris, pH 8.0
  • 150 mM NaCl
  • 1.0% NP-40 or Triton X-100
  • 0.5% Sodium deoxycholate
  • 0.1% SDS (sodium dodecyl sulfate)

NP-40

  • 50 mM Tris, pH 8.0
  • 150 mM NaCl
  • 1.0% NP-40 (or Triton X-100)

Tris-HCl

  • 20 mM Tris-HCl, pH 7.5

Tris-Triton

  • 10 mM Tris, pH 7.4
  • 100 mM NaCl
  • 1 mM EDTA
  • 1 mM EGTA
  • 1% Triton X-100
  • Glycerol
  • 0.1% SDS
  • 0.5% Deoxycholate

2X Laemmli Sample Buffer

  1. 0.125 M Tris-HCl
  2. 4% SDS
  3. 5% β-mercaptoethanol
  4. 20% Glycerol
  5. 0.004% Bromophenol blue
  6. pH to 6.8

Component Stacking Gel 4% Resolving Gel 7.5% Resolving Gel 12%
30% Acrylamide / Bisacrylamide 1.98 ml 3.75 ml 6.0 ml
0.5M Tris-HCl pH 6.8 3.78 ml -- --
1.5M Tris-HCl pH 8.8 -- 3.75 ml 3.75 ml
10% SDS 150 µl 150 µl 150 µl
diH2O 9 ml 9 ml 5.03 ml
TEMED 15 µl 15 µl 7.5 µl
10% APS 75 µl 75 µl 75 µl
Total Volume 15 ml 15 ml 15 ml

  1. Prepare fresh APS and clean the glass plates of the gel cast with ethanol.
  2. Prepare the stacking and resolving gel solutions without TEMED and APS and assemble the gel casting system. Tip: You can check for leakage within your cast by filling the cast with diH2O first.
  3. Add TEMED and APS to the resolving gel solution, mix quickly but thoroughly with a pipette, and add to the cast to the appropriate level.
  4. Immediately overlay the resolving gel with isopropanol to ensure the gel is level, and allow polymerization to proceed for 15 – 30 minutes.
  5. Rinse away the isopropanol with diH2O and gently dry the area above the resolving gel with filter paper.
  6. Add TEMED and APS to the stacking gel solution, mix quickly, and add to the cast above the resolving gel.
  7. Place a comb gently into the solution, avoiding the formation of bubbles.
  8. Allow polymerization to proceed for 30 – 45 minutes.
  9. Gently remove the comb by pulling it straight upwards and rinse the wells with diH2O.

Running buffer (Tris-Glycine / SDS)

  • 25 mM Tris base
  • 190 mM Glycine
  • 0.1% SDS
  • pH to 8.3

Transfer buffer (Wet)

  • 25 mM Tris base
  • 190 mM Glycine
  • 20% Methanol
  • pH to 8.3

Transfer buffer (Semi-dry)

  • 48 mM Tris
  • 39 mM Glycine
  • 20% Methanol
  • 0.04% SDS

  • 24 g Tris base
  • 88 g NaCl
  • pH to 7.6 with HCl
  • H2O to 1 L

  • 100 ml 10X TBS
  • 900 ml H2O
  • 1 ml Tween-20

5% Milk Blocking Buffer

  • 5% w/v non-fat milk
  • 1X TBST

5% BSA Blocking Buffer

  • 5% w/v BSA
  • 1X TBST

Mild Stripping Buffer (1 L)

  • 15 g Glycine
  • 1 g SDS
  • 10 ml Tween-20
  • H2O to 1 L
  • pH to 2.2 with HCl

Harsh Stripping Buffer (100 ml)

  • 12.5 ml 0.5M Tris-HCl pH 6.8
  • 20 ml 10% SDS
  • 0.8 ml β-mercaptoethanol
  • H2O to 100 ml