157Vol. 50 | No. 3 | 2011

Troubleshooting ForumMolecular Biology Techniques Q&A

Native PAGE: polymerization

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 do not represent endorsements by, or the opinions of, BioTechniques.

How can I troubleshoot a poorly polymerized native gel that interferes with sample separation? (Thread 21786)

Q I used the EZ stain protein marker in a native gel since our lab doesn’t have a native-specific marker, but the red and blue bands seemed to merge. The second time I ran

the gel, the marker never left the stacking gel. In both cases, I was unable to distinguish the dimer monomers I’m trying to detect by Western blot. My stacking and separating buffer is 1.5 M Tris-HCl, pH 8.8. How can I get the native gel to work?

A If your SDS gel is already established, running the native gel shouldn’t be a problem as long as you leave out all the denaturing/reducing steps such as DTT, beta-mercaptoethanol, SDS, boiling, etc. You can use the same buffers as long as you leave out the SDS.

A Prestained standards are denatured and aren’t suitable for native gel electrophoresis. You can make your own native protein standard mix with any purified native proteins you have on hand. Remember that charge is an important factor in the separation, sometimes even more important than size, especially if the proteins are close in size. Your best indicator of the run is the tracking dye. The stacking gel buffer should be pH 6.8.

Q The polymerization around the wells was not very uniform and the gel looked unusual. After pouring the separating gel, it took nearly an hour to polymerize. I saw a transparent polymer-ization in the bottom portion of each of the wells that was preventing the proteins from migrating into the gel. Is that common for native gels? I use 1% detergent deoxycholate (DOC) as the cathode buffer. Could the DOC be precipitating and blocking progression of the sample?

A What is the acrylamide concentration of your stacking gel? I use a 3.5% stack in my native gels, which is a higher bis-acrylamide percentage than in the running gel. I catalyze polym-erization with riboflavin and light and it polymerizes cloudy. It takes time to get complete polymerization and I have to flush the wells with electrode buffer prior to loading.

A I haven’t used a commercial marker for native gels, but always add my proteins as markers. What is the pI of your protein? I guess it’s low since you used a buffer of pH 8.8. Is the marker you used suitable for this pH?

Q I tried again and flushed the wells really well, which allowed me to see the proteins on a Western blot. I am trying to detect a monomer and dimer, but saw an additional band. Can anyone explain the appearance of this additional band?

A Is the additional band higher or lower on the gel? It could represent aggregated or degraded protein. Some protease inhibitors have a short lifespan, so they might allow protease activity in your lysate. The additional band might also be the same protein with different post-trans-lational modifications.

A An hour to complete polymerization is normal. You shouldn’t see any polymerization for at least 20 min with a native gel. Allowing more time for polymerization is necessary when running proteins for peptide sequencing so that you can be certain there are no acryl-amide monomers remaining.

Q The SDS-PAGE gel looks homogenous when it polymerizes, but this native gel looks uneven and wrinkled in places. The separating gel is made up of 14.84 mL distilled

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159Vol. 50 | No. 3 | 2011

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water, 7.5 mL 1.5 M Tris-Cl, pH 8.8, 7.5 mL 30% acrylamide, 150 µL 10% adenosine 5′-phosphosulfate (APS), and 15 µL TEMED.

A What is the pH of the Tris and water solution before you add anything else? Are you mixing by inversion in a large enough tube prior to casting your gels? A 50-mL tube works best for 15 mL solution. Be sure to mix well. Since you don’t have SDS, you can mix the solution quite vigorously.

A Do not shake the solution vigorously! That will allow for solvation of atmospheric gasses into the solution. Oxygen will inhibit polymerization. Swirl or mix gently by inversion. That will be sufficient if you do it thoroughly.

A If you are seeing what it sounds like you are describing, then it is normal. Your gel just hasn’t completely polymerized. Give it another hour or so, if necessary, and the wrinkling should resolve. Your wells should be fully formed and uniform.

Q The wells seem fully polymerized, but there are still wrinkles just below the comb. I used 150 µL APS (10%) and 15 µL TEMED. The remaining gel polymerized in the tube shows the same wrinkled appearance.

A You may be using too much APS. Does the gel warm up during polymerization? Heating can cause convection currents in the solution that show up after polymerization. How far into the gel do you see the waves? Are they present throughout the gel or only at the top of the wells?

Q The waves start just below the comb and I can see them about halfway through the gel. I tried again with new separating buffer at pH 8.8, mixed by inversion, used 130 µL of APS and 15 µL of TEMED and polymerized for over an hour. I still saw the wavy pattern, but it was less than the last time I tried. I ran the gel anyway. The samples settled nicely into the wells, but the gel still ran very strangely.

A If your gel looked okay but the sample still ran poorly, then we need to look at the sample. Are you still using the pre-stained standards? They won’t tell you anything in native PAGE. Do you see streaking in your samples? Did you clarify the sample prior to loading? How do you prepare your sample? You need something to increase the density so you can load the sample under the buffer and tracking dye. I use bromophenol blue (BpB) in glycerol. You could also use a couple of crystals of sucrose and BpB in water or buffer.

Why do you use DOC in your electrode buffer? Is it to ensure that the sample remains soluble? If so, is there DOC in the sample and in the gel? If not, then your sample may be precipitating.

Q I use DOC to help disassociate another protein from my protein of interest. It is present in the 5× sample buffer along with 1 M Tris-HCl, pH 6.8, 1% BpB, glycerol, and water. It’s not included in the gel, but is in the cathode buffer.

What do you mean by “clarifying the sample?” How important is the degassing step? Others use the same materials for SDS-PAGE. Could that affect my results? During the pre-run, I forgot to turn on the stirrer and saw some precipitate.

A If you have material in the sample that interferes with the way it runs, you can clarify it by filtering or centrifuging it to remove large aggregates and other particles. You should add DOC to your gel to avoid shocking the protein with a transition. There should not have been anything present during the pre-run to precipitate, so that could be the key to solving the problem. Where did the precipitate come from? Could it be your water? Deionized water with organics removed is usually the best to use.

Carefully vortexing the sample should not be a problem. Just don’t allow it to foam. (I’m not referring to SDS or DOC foam, but denatured protein foam.) Degassing the gel solution is not necessary if you don’t mix too vigorously, but in this case, you might try it.

What color is your TEMED? It turns a deeper yellow as it ages. Fresh TEMED is nearly colorless. What does the dry APS look like? If it has clumps, it has been exposed to moisture and may not be as effective.

Selected and edited by Kristie Nybo, Ph.D.

BioTechniques 50: 157-159 (March 2011) doi 10.2144/000113625

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