Western Blot

Western blotting is one of the most commonly performed laboratory techniques used for protein detection. Combining the protein separation ability of gel electrophoresis, with the specific protein recognition of antibodies. It is the go-to technique for any scientist working on protein expression or purification. Its ability to identify a single protein from a complex mixture of proteins has led to its popularity, with a great number of commercially available antibodies, buffers, reagents, and specialist equipment designed to streamline and optimise this procedure.

The standard western blot method takes a mixture of proteins from a sample such as a cell lysate and separates out the proteins according to size by gel electrophoresis, usually SDS-PAGE. The separated proteins are then transferred by electroblotting onto a nitrocellulose or PVDF membrane. The membrane is blocked to prevent non-specific binding and the bound proteins are probed by a primary antibody, which binds specifically to the protein of interest. To enable detection of this membrane-bound protein-antibody complex, a labelled secondary antibody specific for the primary antibody is utilised. The secondary antibody label, commonly horseradish peroxidase (HRP) allows protein visualisation by enhanced chemiluminescence (ECL) on photographic film or in an imaging device.

Below are some basic considerations to take into account when setting up or troubleshooting a standard western blot protocol:

Choice of gel

Choose the correct percentage gel according to the size of protein being detected. For best transfer of large proteins use a low percentage gel. Gels can be hand poured in the lab or precast gels can be purchased to save time and avoid possible errors.


If the source of the protein to be blotted is a cell line, ensure that the protein of interest is expressed in large enough amounts for detection. Cells may require a form of stimulation to increase protein production, or samples may need to be concentrated. Never overload samples.


Samples are typically denatured and reduced during SDS-PAGE. This allows the antibody to bind epitopes that are only accessible when the protein is unfolded. If the antibody recognises an epitope present on the surface of a folded protein, a non-denatured or native gel is required, where SDS is not used in any of the buffers and the sample is not heated. If a non-reduced sample is required for antibody binding, reducing agent (eg. DTT or β-mercaptoethanol) is left out of the buffers. Follow the antibody datasheet instructions regarding reduction/denaturation and titrate the amount of antibody used within your own system to get the best results.


Run the experiment without the primary antibody if high background is an issue. This will show if the primary or the secondary antibody is non-specifically binding. A positive control lane containing the protein of interest will not only show if the assay itself has worked but will indicate the size the experimental samples should be running on the gel. Also useful are loading control antibodies, which detect a selected constitutively expressed (housekeeping) protein within your cell lysate. This helps confirm consistent loading between lanes. Choose a control protein that is not the same molecular weight as your protein of interest.


This is an important step as it prevents the antibody binding non-specifically. The standard blocking buffer contains 5% non-fat dry milk or 5% BSA. Milk-based blocking agents are not recommended for use with antibodies specific to phosphorylated proteins. Commercially available blocking agents can help optimise the sensitivity of the assay or deal with problems related to high background staining.

Any reagent used in an assay must be within its expiry date and stored as recommended by the manufacturer. Follow the manufacturer’s instructions for use, but be aware that optimisation of experimental conditions, such as incubation times and antibody concentrations, may be required for the best results in your laboratory setting.

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