Tips for Successful Western Blotting
Western blotting is a powerful tool for studying proteins. As a popular technique, encountered by everyone from first-year students to experienced researchers, western blotting has a number of functions. In general, it is useful for identifying, semi-quantifying (it can show relative protein levels, but not an absolute measure) and determining the size of specific proteins. Western blotting is also sometimes called immunoblotting or protein blotting, because it uses an antibody to detect a specific antigen. The “blotting” part refers to the process of transferring the proteins from a gel to a membrane.
The essence of western blotting is to take a sample containing a mixture of proteins (native or denatured), and separate the proteins using gel electrophoresis. The proteins are then transferred to a membrane. The target protein is marked through washing the membrane in a solution containing the primary antibody, which should bind to the protein of interest. A secondary antibody that can be visualised (e.g. through staining or immunofluorescence) is added and binds to the primary antibody. If all goes to plan, the result should be an image with clear, defined bands representing the target protein.
As with any laboratory process, however, there are a multitude of things that can go wrong! Here are a few key points to be aware of for a successful western blot.
1. How should I prepare the samples?
If you are using a tissue sample, it will first need to be broken down by blending or homogenization. Add a buffer to allow lysis of the cells and solubilize the proteins. Protease and phosphatase inhibitors can be included in the buffer to avoid protein breakdown (as these enzymes are sometimes released by cell lysis).
Western blotting is often (but not always) carried out with denatured proteins. This ensures that proteins will be separated by size during electrophoresis and prevents proteins being broken down by proteases. For most samples, boiling for five minutes is a suitable way to denature the proteins.
2. Which type of gel should I use?
Most western blots use polyacrylamide (PA) gels, together with a sodium dodecyl sulfate (SDS) buffer. Western blots using denatured proteins normally use a type of electrophoresis called SDS-PAGE (SDS-polyacrylamide gel electrophoresis.) SDS causes all the proteins to become negatively charged, allowing them to be separated by molecular weight during electrophoresis.
Gel thickness can influence both the quality and quantity of protein detection. Usually, thinner gels allow more efficient protein transfer, ultimately producing protein bands with better definition. Gels are also available with different concentrations of PA. In general, the higher the molecular weight of the target protein, the lower the concentration of PA that will be needed.
3. Do I need to use a blocking agent?
After electrophoresis, proteins are transferred to a membrane so that they can be detected by antibodies. A blocking agent is then used to prevent nonspecific binding between the primary antibody and the membrane.
Non-fat dry milk or bovine serum albumin, together with a small amount of detergent, are commonly-used blocking agents. The proteins in the blocking solution should bind to the membrane in places where the target protein has not bound. This prevents the primary antibody from binding to the membrane, and should therefore reduce background “noise” to give clearer results.
4. How do I work out how much primary antibody to use?
Aim to use an antibody concentration that gives the best ratio of signal to background. A solution with an antibody concentration of anywhere between 0.5µg/mL and 5µg/mL could be appropriate; however, often a concentration of 1µg/mL is sufficient.
You should also consider the relative abundance of the target protein. If you are trying to detect low abundance proteins, you might need to try increasing both the amount of protein and the concentration of the primary antibody.
5. How can I optimize the incubation time?
The standard procedure is to incubate with the primary antibody for one hour at room temperature in order to visualize the target protein. However, overnight incubation at 4℃ should allow plenty of time for the antibody-antigen reaction, increasing the likelihood of a positive signal.
6. Which controls should I use?
It is important to use both positive and negative controls to validate your western blotting results. A useful negative control is to run the procedure using the secondary antibody only – this will help you to determine how much background is due to the secondary antibody.
When you are sourcing antibodies for western blotting, don’t forget St John’s Antibody Validation Project. As part of the Project, you will be able to try up to five free, trial-size samples of our primary antibodies. With around 12,000 antibodies in the project, we are confident you will find something suitable.
