SDS-PAGE gel with two resolving %
TLDR; If you want to use two different resolving gel percentages of SDS-PAGE in the same gel, it’s possible.
This is obviously not peer-reviewed. Reach out to me with any feedback, questions, or suggestions by using my contact form.
Background
This is like those cooking recipes online with the too-long backstory. Feel free to skip to Results/Conclusions and/or Procedure!
Ok. The title. I don’t know what to call this type of SDS-PAGE gel. It’s not the traditional discontinuous gel that has a stacking gel and single resolving gel (a.k.a. “separating gel”) where each has its own pH. It’s that, but there are two resolving gels, hence Disc-DiscR, where “R” is for resolving. “Double-disc gel”? “Two-R gel”? Any suggestions in English or any other alphabetical language?
I wanted to see molecular weights from 5 to 245+ kDa. I was interested mostly in the 5 to 35 kDa range and the 100+ kDa range because I wanted to see monoubiquitin (and short chain free polyubiquitin) as well as high-MW polyubiquitin. It is not possible to get good separation of both ends of the molecular weight range with lab-made gels unless you use a gradient gel former, which I’ve never used (and our lab does not have one). I had been casting one 8% gel and one 15% gel every time, using 2x the amount of my sample, which is limited. The alternative was to use commercially available gradient gels, which, as of this writing, are listed for about $14 a piece from Bio-Rad for the TGX Mini-PROTEAN size. However, my lab is frugal and we try to reduce plastic waste, so we make our own PAGE gels using the Bio-Rad glass casting plates.
Result/Conclusion
[Disclaimer: this is not supposed to be a rigorous reporting of results. The results are not unexpected when one understands SDS-PAGE and western blot, so I don’t feel a large burden of proof in this case, and I’m not trying to be as thorough as I’d be with regular science. Some details may be missing, and I am not proofreading for grammar or typos. I just want to get it out on the internet so folks can use this method.]
To resolve low- and high-MW ubiquitin species, I decided to try an 8% top half and 15% bottom half first. It worked pretty nicely, as you can see from the images in Figure 1. I also tried 14% bottom two-thirds and 8.5% top one-third, which is used for Figure 2.
The two resolving gel percentages are strongly connected at the interface. Moving the gel from the glass cassette to the transfer sandwich did not result in separation of the 8% and 15% gel sections. It behaved like a normal single-percentage resolving gel does with this motion. After the transfer, I played with my first gel—the 1:1 15:8%—trying to pull the two resolving sections apart. I was not able to separate them. Instead, the 8% gel stretched, and as I increased the pulling force, I ripped the 8% gel, which you can see in Figure 1, left side.
Protein can leak horizontally at the interface, as seen in the Coomassie blue staining on the membrane after blotting (Figure 1, cyan boxes). It is hard to know for sure, but I also think the protein transfers just fine at the interface, based on the Coomassie gel stain and the membrane stain (Figure 1). Nevertheless, I’d advise you to make sure your protein of interest does not lie close to where the gel interface is to avoid odd band shapes.
Bands below the interface are well resolved. The samples used in these SDS-PAGEs were yeast (S. cerevisiae) alkaline lysis lysate. In my second gel, 2:1 14:8.5%, I probed for ubiquitin first, treated with azide to inactivate the secondary antibody HRP, then probed for Pgk1, a housekeeping protein in yeast. You can see that the Pgk1 band is well resolved, as are monoubiquitin (~8 kDa) and ubiquitinated proteins lower than 100 kDa (Figure 2).
This works for my needs. I am still optimizing the percentages and vertical gel heights to use.
Figure 1. Left: Coomassie staining of the 15:8% gel after transfer shows the gel interface near the middle of the gel and a rip where I pulled on the gel to try to separate the two gels but only ripped the 8% gel instead. Right: coomassie blue staining of the membrane after blotting.
Figure 2. Left: image of the 14:8.5% 2:1 gel still on top of the membrane right after wet transfer showing the interface above the 100 kDa marker. Right: anti-mouse secondary HRP exposure showing Pgk1 and residual ubiquitin primaries superimposed on the marker layer (separate channel) by GeneSys software. The marker is the same as in Figure 1.
Procedure
Let me know if you think it can be done better.
These are instructions for casting two (2) 1:1 15:8% gels (5% stacking gel) using the Bio-Rad Mini-PROTEAN system with 1 mm-gap glass plates.
Prepare the glass plates as usual (wipe clean plates with a lint-less paper wipe, e.g. KimWipe, wet with 70% ethanol).
Clamp the plates together and place the cassettes on the casting stand as usual.
I usually check the seal of the gasket by squirting some ddH2O into the cavity and then pour out the water just before I add the polymerization activators (TEMED and APS) to the Tris, SDS, acrylamide-bisacrylamide, and water gel solution.
TIP: use vaseline/petroleum jelly on the gasket to help seal any chips in the bottom of the glass plates and prevent leaking, which can happen even if the bottom edges of the two plates are flush even.
Prepare at least 5.5 mL of 15% polyacrylamide gel solution, and right after adding the polymerization activators and mixing the gel solution, add about 2.5 mL of the mixture to each glass casette. Immediately, gently add ddH2O to the top at least 2 centimeters to prevent exposure of the gel to air.
A lab mate uses 70% ethanol to cover the gel instead of ddH2O. It seems to work for her.
Let the 15% gel polymerize (30 to 60 minutes, depending on your recipe and reagent freshness).
The higher percentage gel usually “shrinks” a little in the vertical direction, in my experience.
Prepare at least 5.5 mL of 8% polyacrylamide gel solution. Before adding the activators, pour off as much water as possible from the 15% gel. Immediately add the activators to the gel solution, mix, and add the 8% mixture to the 15% gels. Fill to the level that your normally fill it to (approximately 2.5 mL per gel). Add ddH2O gently.
Let the 8% gel polymerize.
Prepare stacking gel as you normally prepare it. We use 5% gel. Add the stacking gel onto the 8% gel, and insert the comb immediately.
Let the stacking gel polymerize.
Wrap the gels in their glass casettes in soaking wet paper towel (tap water is okay for wetting the paper towel) and seal the wrap in a plastic resealable bag (e.g. “sandwich” size Ziploc bag… You can reuse this bag and not throw away so much cling wrap!).
Store the gels at 4 degrees. Use within 1 week (the sooner the better).
The wet transfer conditions I used were 105 V for 90 minutes in a cold room in Tris-glycine transfer buffer (no methanol).
