Old but not Outdated

by Vasudharani Devanathan, Labtimes 05/2013

Example from the “Hall of shame” at the Rice University: To get a gel like this, you have to open the electrophoresis chamber before the run is completed – with the power on.

Sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDS-PAGE) ist still one of the most frequently applied methods in life science laboratories. Though it’s a simple and very basic technique, it may sometimes go wrong.

When I started my bachelor of science, my professor told me that SDS-PAGE is one of the key techniques in the lab, which will never be outdated. I still remember trying to understand every gel preparation step described in Molecular cloning by Sambrook and Maniatis. My Professor was right, although many modifications exist today; SDS-PAGE is not extinct. It is still in wide use in almost all protein labs.

These are days, however, when we buy “pre-made” commercial gels. We live in a fast world and everyone prefers quick access to everything. You may wonder why one should know how to make a gel when it can be readily bought. The reason is simple: it is important to understand the principle and to know the constituents of a gel, to be able to analyse a failed electrophoresis or a doubtful result.

Gel electrophoresis is used to separate proteins with an electric field and was first formulated by Laemmli in 1970 (Nature 27:680-685). Since, it has become such a common laboratory technique that we take it for granted and most of us don’t know the ABC of how a discontinuous pH gel system actually functions.

I worked in a lab with about 30 PhD students. Around 15 of them ran gels every day – not just one but four; the minimum was two gels per student per day. We used to come as early as possible just to pick up the best gel systems. Sometimes, students came back around ten at night, just to reserve a gel system. In such conditions it is important to run a precise gel and finish the experiment with a beautiful image. Finally, it is all about uploading a beautiful western blot for publications.

Basically you have to consider three simple rules for proper gel preparation:

  • Make your gel with the correct percentage of acrylamide and bisacrylamide
  • Prepare your sample properly using a suitable sample buffer.
  • Have the patience to separate the samples well.

If you carefully follow the above steps, a good deal of problems that could potentially emerge in subsequent western blotting can be avoided. To make the gel correctly, sound knowledge of the components, their contribution and role in the acrylamide gel is essential.

SDS gel is a discontinuous gel which comprises two parts: the stacking gel and the separating or running gel. They both have the same constituents with a slightly different composition (stacking gel: Acrylamide 3% and separating gel: 5% to 20%). As indicated by the name, the stacking gel tightly stacks the proteins according to their molecular weight and prepares them for slow separation. The separating gel dissolves the proteins according to their molecular weight. Apart from the composition, they differ in their pH values. The stacking gel is pH 6.8 and the running gel is pH 8.8.

Acrylamide and bisacrylamide form a meshwork, in which the proteins can easily migrate and separate. Ammonium persulfate (APS) and TEMED help to polymerise the gel, thus leading to a tight meshwork. Stacking gels have spaces for loading samples to be analysed, which are formed with a pre-cut comb. Depending on the number and amount of samples to be loaded, one can choose the number and size of the wells. Chloride is the mobile anion in the gel, while glycine is the mobile anion in the upper and lower tank buffer. Proteins are negatively charged due to SDS and migrate to the bottom of the gel.

All too often, a three-day western blotting experiment does not work because the homemade gel is not casted properly or the commercial gel is outdated. Trouble shooting an SDS-PAGE gel is like searching for a needle in a haystack. Common problems are due to polymerisation failures. If APS and TEMED are not correctly proportioned or not mixed well after being added, the polymerisation is affected. Another reason why gels do not polymerise can be due to back shelf storage of APS. Although most of us love to freeze APS, a hard core ­biochemist and a gel lover would never recommend this – freshly made APS simply polymerises gels better.

Other problems include wavy gel, brittle gel, soft gel and inconsistent mobilities. Wavy gels are mostly due to impatience and too much TEMED or APS added to polymerise the gel quickly. Researchers running many gels a day prefer to make them on one day and store them. In such a case, it is better to clean the glass plates and the comb very well to prevent sticky left over gel pieces from affecting the new gel.

Yet another common problem is overloading lanes, which lead to fused ends in protein bands. This makes it difficult to quantify the gel. Hence, before starting a gel, one should know the amount of protein that can be detected. If you are dealing with an unknown protein or analysing a specific protein from a tissue lysate, it is advisable to start with a very low quantity and load different concentrations in each lane. This helps find the minimum detectable amount of protein necessary to get a sharp and ­visible band in SDS-PAGE.

Last Changed: 17.09.2013

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