Western Blot Protocol

Preparing the lysate - From cells

Check that the confluency of the cells is to expectation by viewing under a microscope.
Place the cell culture dish on ice and wash the cells with ice-cold Phosphate Buffer Saline (PBS).
Remove the PBS and add ice-cold lysis buffer. (1 ml per 10⁷ cells/100mm dish or 150cm² flask, 0.5ml per 5x10⁶ cells/60mm dish or 75cm² flask)
Scrape any adherent cells off the dish using a cold plastic cell scraper before gently transferring the cell suspension into a pre-cooled microfuge tube.
Maintain constant agitation for 30 minutes at 4°C.
In a cool microcentrifuge, centrifuge the lysate. For this step you should adjust the centrifugation force and time depending on the cell type - as a guideline use 20 minutes at 12,000 rpm, but this should be determined by the person carrying out the procedure (e.g. leukocytes need a very light centrifugation).
Gently remove the lysate from the centrifuge and place on ice.
Aspirate the supernatant and transfer to a fresh tube kept on ice, and discard the pellet.
Denature the lysate by heating the lysate at 95°C for 5 minutes.

Preparing the lysate - From tissues

Note: For this process we recommend keeping the tissue cool throughout the process to avoid degradation by proteases.

Dissect the tissue of interest with clean tools.
Place the tissue in a round-bottom microfuge tube or Eppendorf tube and immerse in liquid nitrogen to “snap freeze”.
The sample may be stored at -80°C for later use, or keep on ice for immediate homogenisation.
For a ~5 mg piece of tissue, add ~300μl lysis buffer to the tube and homogenise with an electric homogeniser.
Rinse the blade twice with another 2 x 300μl lysis buffer before agitating for 2 hours at 4°C (e.g place on an orbital shaker in the fridge).
Ensure a minimum concentration of 0.1 mg/ml, and an optimal concentration ~1-5 mg/ml.

Note: The volume of lysis buffer should be determined according to the amount of tissue being used. Protein extract should not be so diluted that protein is lost or large volumes are needed to load onto the gel.

Centrifuge for the lysate for 20 min at 12000 rpm at 4°C in a microcentrifuge.
Gently remove the tubes from the centrifuge and place on ice, aspirate and transfer the supernatant to a fresh microtube and keep on ice.
Discard the pellet.
Denature the lysate by heating the lysate at 95°C for 5 minutes.

SDS-PAGE (Sodium Dodecyl Sulphate-PolyAcrylamide Gel Electrophoresis)

Prepare the separating gels according to the protein size you are looking to identify. The following can be used as a guideline:

4-40 kDa - 20%
12-45 kDa - 15%
10-70 kDa - 10%
15-100 kDa - 12.5%
25-100 kDa - 10%
Once the separating gel is set prepare the stacking gel. (Ensure to use a plastic comb to form the wells for lysate application.)
When both gels have set and the samples are ready to load use a pipette with a gel-loading tip to gently load equal amounts of between 20-40μg of each lysate per well.

Note: On loading, take care not to poke through the bottom of the wells with the tip as this will distort the band at the development stage. Do not overfill wells as this will lead to spillage into neighbouring wells and corrupt the results.
Add the buffers to the relevant chambers and ensure that there is a good seal on the gel cassette chamber.
We recommend adding frozen gel ice packs to the outer chamber, which will prevent the buffer from overheating and altering the pore size of the gel, which typically results in a protein "smile" efffect at development.

Run the gel according to the following guidance:

Small peptides: High voltage 150 –300V, offers good resolution so long as the process is quick. The initial high current will align small molecules quickly in the stacking gel to get a sharp band.
Larger peptides: Low constant voltage (80-100V) is a good choice for resolution for 70-80 kDa proteins. This will reduce heat from resistance build up in the gel, but a longer timelapse is recommended.

Note: By keeping the voltage the same, the current is reduced, because resistance builds inside the gel over the running time. For some proteins which do not completely meet reducing conditions, their shape will inevitably lead to increased resistance within more concentrated gels. The heat generated will mean the gels need to be cooled to prevent restructuring the conformation of the proteins.

When the dye reaches the bottom of the gel, turn the power off.
Proteins will slowly elute from the gel at this point, and we recommend not storing the gel - proceed immediately to transfer.

Transfer of proteins - Western Blot transfer

Nitrocellulose or PVDF are the most common types of membrane available, and the transfer can be carried out under wet or semi-dry conditions. In both cases the membrane should be fully wet prior to use, and care should be taken to ensure the gel and the membrane are bound together without any bubbles forming between them. Gently rolling over the layers with a roller can ensure an even covering.

Wet transfer

The gel and membrane are sandwiched between sponge and paper (sponge/filter paper/gel/membrane/filter paper/sponge). (PVDF membranes must be activated with methanol prior to this stage due to its hydrophobic surface.)
Clamp the sandwich tightly together and lock into the gel cassette, ensuring no air bubbles have formed between the gel and membrane.
Submerge the cassette into transfer buffer prior to transfer initiation. The buffer in the western blot tank should fully cover the gel cassette.
It is essential that the direction of the membrane and gel are positioned with the gel on the cathode side of the cassette and the membrane on the anode side, thus ensuring the travel of proteins from the negative electric potential towards the positive electrode. The proteins will migrate from the gel and bind to the membrane.

Semi-dry transfer

This process is more simplistic than the wet transfer in that no tank is used for the transfer. Instead, a sandwich of filter paper/gel/membrane/filter paper is wetted in transfer buffer and placed directly between positive and negative electrodes (similarly to a close-top grill).
Like the wet transfer, it is important that the membrane is closest to the positive electrode and the gel closest to the negative electrode.
Note: The proportion of Tris and Glycine in the transfer buffer may be altered for this type of transfer.

How to block the membrane

It is best to block with 5% serum of a species from the same host species as your secondary antibody. The IgG from the serum will then prevent the secondary antibody binding non-specific sequences. However, as BSA is a cheap resource and generally not used in human research, BSA and milk are commonly used.
Depending on the type of protein you are looking to identify, traditionally there are 2 types of blocking solutions which are used. Bovine Serum Albumin (BSA) is found in the serum of calves and binds to non-specific sequences, which help to prevent non-specific antibody binding.
Non-fat milk or milk powder may also be used. These can be tried interchangeably, but there are circumstances in which it may be preferable to use one over the other. Some antibodies give a stronger signal on membranes blocked with BSA as opposed to milk - BSA is recommended for phospho-proteins.

For the blocking, prepare a 5% milk or BSA solution to a solution of 5g per 100 ml Tris Buffer Saline Tween20 (TBST).
Mix the solution well until solids have fully dissolved. Filtering may also be applied to remove any impurities.
After removing the membrane from the transfer machine, incubate with the blocking solution for 1 hour at room temperature, or overnight at 4°C, with agitation. Ensure the solution covers the whole membrane during agitation to prevent any background occuring later in development.
Wash for 5 seconds in TBST after the incubation.

Incubation with the primary antibody

Dilute primary antibody in TBST at the recommended dilution. You may also include some milk powder or BSA in the solution as this shouldn't interfere with antibody binding.
Incubate the membrane with the diluted primary antibody solution for 1 hour at 37°C, 2 hours at room temperature or overnight at 4°C with agitation.
Remove the antibody solution.
Wash the membrane 3 times for 5-10 minutes each time at room temperature in TBST (50mM Tris, 100mM NaCl, 0.05% Tween-20, pH 7.6), with agitation.
Note: Increasing the concentration of Tween20 to 0.1% reduces the background and increases the specificity, but it will also reduce the sensitivity.

Incubation with secondary antibody

Incubate the membrane with the diluted secondary antibody-conjugated solution (according to recommended guidance) in TBST for 1 hour at room temperature with agitation.
Upon completion remove the antibody solution and repeat with 3 wash cycles.

Note: Unlike the primary antibody solution, the secondary antibody solution should not contain any blocking components like BSA or milk powder.

Chemiluminescent Reaction

Prepare and use the chemiluminescent substrate according to the manufacturer’s instructions. Following substrate exposure to the membrane wrap the membrane and expose to X-ray films for between 10 seconds to 1-hour (depending on recommended guidance). Exposure times could vary significantly according to the amount of antibody and antigen applied.