How can you optimize electrophoresis in protein analysis?

I highly recommend using the Protein SImple automated western blot platforms like the 'Jess' and 'Peggy Sue'. We use the Jess in our lab and get great results. Sample and buffer prep takes about 20 mins and the actual run only takes 3 hours.

Proteins and protein complexes can be also separated based on their m/z (mass over charge) in a non denaturating buffer (Tris-Glycine without any SDS added) and SDS free gels. This also requires a specific ladder for comparison and non denaturating conditions for sample preparation

SDS Tris-glycine buffer is commonly used to separate proteins based on their size. SDS imparts a negative charge to the proteins and maintains them in a linear, denatured form. You can either use commercially available highly concentrated buffers or prepare your own. For silver staining gels, ultrapure water should be used to eliminate backgrounds and increase clarity of bands when preparing solutions, rinsing gels, and washing containers.

If your bands appear as smiling or uneven, you should adjust the voltage. Smiling bands and faint bands on western blots are primarily caused by higher voltage. When you are running your gels, make sure the buffer does not overheat. Overused buffers can create resistance, leading to heating, and consequently, the appearance of faint or abnormal bands.

Most commercially-available gels can be purchased in 4-20% cassettes. This allows users to evaluate protein samples that consist of a variety of sizes without having to analyze each protein component separately.

Proteins will be generally separated using polyacrylamide gels and not agarose. Agarose will be more commonly used for nucleic acid electrophoresis.

Sample preparation for proteins separations in denaturating conditions usually include a boiling step (e.g. 1 min at 100 Celsius) to facilitate protein denaturation in addition to the contribution of SDS and DTT. It is also helpful for to add Glycerol (or similar sugars) to the sample buffer to increase its density and facilitate sample loading in the gel wells.

Preparing samples for protein electrophoresis often requires sample buffers with detergents like SDS and reducing agents such as DTT. Ensuring appropriate concentrations of both is key to achieving optimal denaturing. Less soluble hydrophobic proteins may require the addition of small concentrations of chaotropic agents like urea to be added to the sample buffer to ensure a proper charge to mass ratio and solubility. Additionally, the temperature and length of time heating the sample prior to loading needs to be fitting for the sample buffer used to ensure full denaturing without degrading the proteins in the sample.

There are cases where it’s not possible to quantify a protein sample by spectrophotometry or colorimetry. This can occur if your sample contains a compound that absorbs at a similar wavelength as proteins or interacts with colorimetry reagents. Most commercial protein ladders specify the amount of each standard used per volume, so when they are ran with your protein samples, you can quantify your samples based on band size and intensity. This quantification can be optimized by loading one microgram of protein ladder with a serial dilution of your samples, then performing imaging analysis to roughly determine protein quantity.

Coosmassie staining is most common for protein staining, so make sure to destain your Coomassie gels so you can properly visualize your protein.

Use a loading control. It is advisable to incorporate a loading control, such as a protein ladder or molecular weight markers, to help estimate the size of protein bands accurately. Handle gels carefully. Select the appropriate percentage and type of polyacrylamide gel (e.g., gradient gels for SDS-PAGE) based on the size range of proteins you are analyzing. We usually use a lower acrylamide gel in stacking layer compared to the loading gel. Consider running the gel in a cold room or use ice-bath around the equipment.

After starting your gel, always check for electrolysis bubbles. That will determine if your gel is running. If your gel isn't running, the most likely mistake is forgetting to remove to sticker across the bottom of the gel.