SDS-PAGE (sodium dodecyl sulfate polyacrylamide gel electrophoresis) is the most widely used technique to separate proteins from complicated samples of mixture. It plays a key role in molecular biology and in a wide range of subfields of biological research. Although it’s commonly used, it’s not always fully understood, so let’s give it a try.
SDS-PAGE basically separates proteins according to their molecular weight, based on their rates of migration through the polyacrylamide gel under the influence of an applied electrical field. The movement of any charged species through an electric field is determined by its net charge, molecular radius, and the magnitude of the applied field. The problem with native proteins is that their net charge and molecular radius are not molecular-weight dependent. Instead, the net charge is determined by amino acid composition and molecular radius by the tertiary structure. So if we want to separate proteins based on their molecular weight only, we need to destroy the tertiary structure and mask the intrinsic net charge of the protein. That’s where SDS comes in handy.
SDS is a detergent present in the SDS-PAGE sample buffer where, along with a bit of heat and a reducing agent (normally DTT), it makes the protein linear and uniformly negative charged, and ensures it stays that way all throughout the procedure. After the separation of proteins by SDS-PAGE, I transfer them from the gel to the nitrocellulose membrane. In other words, I perform Western blotting. The proteins transferred from the gels are immobilized at their respective relative migration positions at the time point when the electric current of the gel run was stopped. Blotting makes it possible to detect the proteins on the membrane using specific antibodies. This article will focus on the electrophoresis.
Step 1: Gel choice
Polyacrylamide gels can be purchased, or you can make them in the lab. I prefer the commercial ones because making your own gels increases the risk of error in an already error-prone procedure. Contamination and pH imbalances happen more often than you could imagine in the “homemade” versions. In both cases, make sure to correctly choose the percentage of your gel, as this defines migration and separation of proteins. The rule is: The smaller the size of the protein of interest, the higher the percentage of acrylamide/bis; the bigger the size of the protein of interest, the lower the percentage of acrylamide/bis. Here’s a guide for choosing gels properly:
Once you’ve made your choice, place the gels in the electrophoresis tank and add migration (running) buffer. The gels should be submerged in it. Standard running buffer for PAGE is 1x Tris-glycine (pH 8.3) consisting of 25 mM Tris base, 190 mM glycine, and 0.1% SDS. Acrylamide is a potent neurotoxin, so be sure to wear gloves!
Step 2: What to add to your run other than the samples themselves?
Protein Ladder is the main thing you need to have running besides your samples. It is a molecular weight ladder that will determine the size of your protein of interest at the end of the run. In my lab we use Chameleon Duo Pre-Stained Protein Ladder.
Positive control is something I advise you to incorporate into your experiments—especially when using new protocols or antibodies. This way you will be certain that your antibody is actually capable of binding the protein of interest, even though it might not be present in the analyzed samples.
Loading controls are a way of confirming that protein loading is equal across the gel, and their expression level should not vary between the different samples. Housekeeping genes are often used as loading controls because those are the proteins that exhibit high-level, constitutive expression in the cell type or sample we are analyzing. It is important to select a loading control that has a different molecular weight than the protein of interest—otherwise, you won’t be able to distinguish the bands.
Step 3: Loading the samples and running the SDS-PAGE
I usually use 12-well systems, which have a limit of 20 µl of sample per well for mini gel loading. Thermo Fisher Scientific has a helpful guide, depending on your needs.
Use special gel-loading tips or a micro-syringe for this procedure. Pipette the samples up and down several times before loading, but be careful not to create bubbles. When loading, don’t touch the bottom of the well with the pipette, as this can cause distorted bands. And don’t push the pipette plunger further than the first stop because this causes bubbles and leaking of your sample from the well.
Now you’re all set to run the gel. Duration depends on the voltage and the manufacturer’s recommendations, and can vary from one hour to overnight.
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