Tips for Successful SDS-PAGE

Running SDS-PAGE or sodium dodecyl sulfate-polyacrylamide gel electrophoresis is a basic biotechnology technique that is used frequently, but often can be tricky to set up and achieve good results. This post will give a general overview of SDS-PAGE electrophoresis, as well as, give some tips and troubleshooting ideas to help you have a successful SDS-PAGE run!

Quick Overview

SDS-PAGE gels are used to analyze proteins by separating them according to their molecular weight. In their native form, proteins have a complex structure and often come in varying shapes and sizes making electrophoresis difficult. SDS is an anionic detergent that complexes with proteins, and alongside a reducing agent, like 2-mercaptoethanol, is used to unfold a protein’s secondary structure by breaking the bonds that link subunits together. This process is called denaturation, where protein structure is broken down and unfolded from its native state. After the proteins are denatured, they can be loaded into the wells on the polyacrylamide gels. An electric current is used to move the SDS treated proteins towards the positive electrode, in other words, it causes the proteins to migrate from the top of the gel to the bottom. The polyacrylamide gel is porous, where small proteins are able to migrate quickly through the small pockets of the gel. The larger proteins cannot move as quickly through these small pockets, meaning these proteins will migrate slower. This results in protein bands forming, which can be later analyzed by staining to visualize the different molecular weights of each band. 

Tips for Successful Gels

  1. Remove Comb Cleanly
  • Keep your gel straight and sturdy in one hand, while removing the comb with your other hand. When removing the comb, try pulling it gently upwards to prevent damage to wells.
  1. Prep Gel for Chamber
  • Some gels will have tape near the bottom that needs to be removed before putting it in the chamber, if it is not removed samples will not run properly. Not all gels come with tape, so if there is none, you can continue with the next steps in the procedure. Make sure your gel is in the proper orientation before placing it in the chamber. Chambers vary, so check to make sure you place the gel correctly according to your chamber’s directions. If using a vertical protein electrophoresis chamber from Edvotek, the short plate side of the gel should be facing the interior of the chamber. Lastly, when placing the gel in its place, make sure it is straight and is completely secure before adding the electrophoresis buffer and rinsing wells. 
  1. Rinse Wells
  • Rinsing wells will ensure that samples have a clear path to be loaded into wells, so the samples can settle evenly at the bottom of each well. Using a transfer pipet, rinse wells with the electrophoresis buffer until it seems clean enough for the samples to be loaded (rinsing each well 2 or 3 times will usually be enough). 

Troubleshooting SDS-PAGE 

  1. If your gels are not running properly, here are some things you can do.
  • Make sure that you are using the right buffer, and that it was prepared correctly. Remake the buffer if necessary.
  • Check the buffer volume in the chamber. Ensure that the buffer is covering the wells completely. If there’s not enough, just add some more buffer!
  • Make sure that the tape on the gel was removed (if there is tape).
  • Check that the leads are completely connected to the power supply and the chamber.
  1. If your gel resolution is not good or band separation is lacking, here are some things you can do.
  • Check the quality of your polyacrylamide gels. These gels tend to last around 3 months when stored at 4° C. If you have kept them for longer than this time period, visually check that gels don’t look cracked or dried out, as this could cause issues with your resolution and sample migration. 
  • Check that the concentration of the gel and the buffer is correct.
  • Make sure that the wells weren’t overloaded with the samples. 
  1. If your gel has band distortion or bands are faint, here are some things you can do.
  • Check that the gel is running at the recommended voltage indicated in the protocol. Running gels at a higher voltage than recommended can cause overheating, leading to migrated bands being distorted in a smiling manner, instead of bands migrating even and straight. Lower the voltage, if necessary, to achieve better band migration.
  • Check that you didn’t overload samples. Load the volume indicated in the protocol for optimal results. 
  • If your bands look like they migrated well, but are fainter than desired, try staining your gel for a longer period of time.
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