Western blots can be ugly. I mean down-right, horrifically, wall-of-shame ugly. Not only can they be embarrassing to show to your colleagues, but the ugliness can obscure your results, making it impossible to interpret your data.
Blotting consists of many experimental steps, which makes the technique naturally error-prone. Although standardized protocols exist, many fail to point out the small details that could turn a really ugly Western blot into a thing of beauty.
To save you the humiliation of presenting an ugly blot at lab meeting, I am sharing my 10 tips to good-looking Westerns.
Buy commercial solutions to pour your gels
We were preparing our own solutions to pour gels in the lab until we realized that pH imbalances and contamination of the solutions were happening more often that we would have liked (or imagined!). Once we started using commercial reagents, we never had problems with our gels again. They are not expensive (especially when compared to repeating experiments over and over again to get a pretty blot) and it eliminates one source of error easily.
Shake it! But at the right speed
There are many steps that involve the incubation of the membrane in a shaker. Shaking is more critical than people think, especially when incubating with the primary or secondary antibodies. The correct speed of the shaker (not too fast/too slow) will determine if the antibody binds uniformly to the membrane. Too fast won’t allow the antibody to bind and too slow will impede the uniform binding of the antibody. Be sure that you test the correct speed and always set it up accordingly when you perform your experiments.
Dissolve the milk (or blocking buffer) properly to avoid black dots
Preparing the blocking buffer is often done in a hurry, but this can lead to black dots that make your blot ugly. Make sure you prepare your blocking buffer ahead of time so everything can dissolve properly. If you still have problems with black dots, filter your blocking buffer before use to remove any particulates.
Always use a positive control
When you test a new antibody make sure you use a positive control. Many companies (for example Cell Signaling) offer a list of positive controls for many of their antibodies. These can either be purchased or prepared in the lab. Often it’s just a cell lysate of a particular cell type. Having a positive control rules out the possibility that the antibody simply does not work if you have to perform troubleshooting.
Cut your membrane to incubate different antibodies
When performing blots for many samples, optimize the process to avoid running multiple gels. If the proteins you are interested in are different sizes, cut your membrane after the blocking step, so that you can incubate one half of the blot with one antibody and the other half of the blot with another antibody. Just make sure you are confident where your protein runs – you don’t want to accidentally cut your protein in half!
Be careful when re-using antibodies
Re-using antibodies is generally not recommended. However, they are expensive and some of them can be used two or three times (in general it works better when using antibodies for loading control proteins like actin or GAPDH). Make sure that you mark your antibody-containing tube properly so that you know how many times you have used it.
Don’t abandon your antibody at the bench
Antibodies are very sensitive to temperature changes! If they are kept at -20º, carry them in a mini-cooler so that they are protected against temperature changes. Forgetting your antibody for a while at the bench can make it useless for your next Western blot.
Avoid buying cheap antibodies
There are many companies that produce antibodies of low quality that have not even been tested. They might appear to be cheaper but these antibodies will drive you crazy when you have to test the antibody many times. Sometimes, cheaper options end up being very expensive!
Use an internal standard to ease quantification between different membranes
When you have to run an experiment with many samples that do not all fit on one gel, you need to have the same sample on each of the membranes to use as a “control” for quantification. It will save you a lot of time if you produce a big amount of your internal standard and have it always ready for all your membranes.
Carefully choose your loading control
To quantify your blots you always need to use a separate protein as a loading control. Normally, actin or GAPDH are used. However, these proteins are not always constant under all conditions. Be sure that you investigate in the literature and in your own experiments that these proteins do not change expression under your experimental conditions.
Following these guidelines will help take some of the ugliness out of your Western blots.
Do you have any tips of your own?