This month's question from the Molecular Biology Forums (online at molecularbiology. forums.biotechniques.com) comes from the “Protein Methods” section. Entries have been edited for concision and clarity. Mentions of specific products and manufacturers have been retained from the original posts, but they do not represent endorsements by, or the opinions of, BioTechniques.

Molecular Biology Techniques Q&A

How can I improve the migration of my proteins? (Thread 21786)

Q I am trying to run a native gel. I used the EZ stain protein marker that I usually use for SDS-PAGE. The red band and the higher blue band seemed to merge and the dimer monomers that I am trying to detect by Western blot ended up very close to each other.

The second time I ran the gel, the marker got stuck in the stacking gel. The separating and stacking gel buffer is 1.5 M Tris-HCl, pH 8.8. The polymerization around my wells is not very uniform and looks uneven. Is that common for native gels? I am using freshly made aminopropyl triethoxysilane (APS), and a new bottle of acrylamide and TEMED. I use 1% detergent deoxycholate (DOC) as the cathode buffer. Does anyone have any advice on what else I can change?

A1 The stacking gel buffer should be pH 6.8.

The prestained standard you are using is denatured and not suitable for native gel electrophoresis. Your best indicator during the run is the tracking dye. You can use more than one if you wish to see separation of colors during the run. If you don't have a native protein standard mix, then you can make your own with any purified native proteins you have on hand. Charge is an important factor in the separation, often even more important than size.

A2 Running a native gel after you establish the SDS gel should be no problem as long as you leave out all of the denaturing and reducing reagents and steps such as DTT, βME, SDS, boiling, etc. If your buffer works well for SDS-PAGE, it should work in a native gel if you leave out the SDS.

A3 What is the acrylamide concentration of your stacking gel? I use a 3.5% stack when I run a native gel. This is a higher bisacrylamide percentage than my running gel. It is cloudy after polymerization, which I catalyze with riboflavin and light. It takes a while to get complete polymerization and I have to flush the wells with electrode buffer prior to loading the gel.

A4 What is the p1 of your protein? I guess it has low pI since your buffer's pH is 8.8. Is the marker you used suitable for running at this pH?

Q I am trying to detect a monomer and dimer but there seems to be an additional band. I think I put the lysate through multiple rounds of freeze and thaw. Could some unwanted degradation explain the appearance of this additional band?

A Yes. Is the additional band higher or lower on the gel? The protein could either have aggregated or degraded. Your lysate will contain proteases and some inhibitors have a short lifespan. Have you performed this analysis before? Your additional band might be the same protein with a different post-translational modification.

Q The additional band is higher up. I prepared the lysates last week and will use a new sample that I have not thawed yet to see if that makes a difference.

I wait for almost an hour for the gel to polymerize. When polymerized, it does not look like an SDS-PAGE gel and has some wrinkling. Is that common?

A1 An hour to complete polymerization is not unusual. Polymerization should become visible within 20 min in a native gel. Giving the gel longer to polymerize is actually necessary when running gels for peptide sequencing since you need to ensure that there are no acrylamide monomers remaining.

A2 The problem may occur while making the gel solution. Are you mixing by inversion in a large enough tube prior to casting your gels? I got clumpy gels when I mixed a 15-µL solution in a 15-µL tube so I switched to a 50-µL tube and that solved the problem. Since you don't have SDS in the mixture, you could probably even shake the container forcefully or vortex it without risking bubble creation.

A3 Do not shake the solution vigorously! That will allow for the incorporation of atmospheric gasses into the solution. One of those gasses, oxygen, will inhibit polymerization. Swirling or mixing by inversion will be sufficient if thorough.

A4 If the wrinkles in your gel don't clear up after letting the gel sit for a long time then you may be using too much APS.

Does your gel warm up during polymerization? Heating could cause convection currents in the solution that show up after polymerization. Do you see the waves throughout the gel or just at the top of the wells? If they are only at the top of the wells, then let the gel polymerize for a longer period of time.

A5 If your gel looks okay but the samples still don't run correctly, you will need to look at your sample. Do you see streaks in the sample? Did you clarify the sample prior to loading? Do you use the DOC in your electrode buffer to ensure that the sample remains soluble and is there also DOC in your sample and gel? If not, your sample may precipitate.

Q In my assay, DOC is meant to help disassociate another protein from my protein of interest. It is in my sample buffer but not the gel.

A I would recommend adding DOC to the gel to avoid shocking the protein with a transition.

If that doesn't solve the problem, you can remove large aggregates and other particles by filtering or centrifuging your sample.

Some recommend degassing the gel solution, but this is not necessary if you don't mix too vigorously. You could try that if you are still getting poor polymerization.