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The Practical Guide to Running a Perfect Homemade SDS-PAGE Gel

SDS gel

As biochemists, we routinely run SDS-PAGE to analyze our proteins. It can be a repetitive and tedious task to get a “presentable” gel for publication (especially when there are Western blots to follow)… but not if you know the tricks!

Here is my guide to the perfect gel:

Clean the glass plates well

Look out for gel debris on your plates, as it is a little devil that causes leakage and imperfections in your gel.

If you are running sensitive samples (e.g. RNA-protein complex), always remember to spray a little 70% ethanol / DEPC water on the plates and wipe them dry.

Stop SDS-PAGE gel leakage (first checkpoint)

I am using BioRad terminology here, but it is technically the same setup for other brands.

Follow these steps to prevent your gel from leaking out before it polymerizes:

  • Insert the glass plates into the (green) casting frame while the frame is placed on a flat surface (such as your lab bench)
  • Open the “doors” to lock the plates into place
  • Feel the bottom of the plates with your finger – when properly aligned the two glass plates should be perfectly parallel
  • Assemble the casting frame on the transparent stand and fill the compartment with a little water to check for leakage
  • Pour the water out (without dissembling the set up) and you are ready to pour in your acrylamide mix

Overlay the separating gel

It is a general practice to overlay a layer of “liquid” on top of the separating gel before it polymerizes to ensure an even interface between the two layers. This is important to ensure your samples are at the same starting point as they begin their race along the separating gel.

This “liquid” can be isopropanol, ethanol, or water. If you are using the organic solvents, make sure you remove any residue drops with filter paper after the separating gel is set. If you are using water, make sure you overlay it slowly or else it will directly mix with the acrylamide (but on the bright side, you don’t have to worry too much about the residual water). And in either case, keep this layer of “liquid” under 500µl per 75mm-thick mini gel.

Assemble the cassette (second checkpoint to stop the leakage)

We are talking about a different type of leakage here – in which your running buffer is leaking from the cathode to anode, or as we know it, from the inner to the outer compartment.

It is important to stop the leakage as resistance builds up when the buffer level is running low in the cathode. This will cause the whole gel system to warm up quite quickly, resulting in:

  • Smiling / frowning gels
  • Smearing lanes
  • Dissociation of SDS-resistant complexes that are otherwise observable on your gels!

It is in fact quite easy to keep your gel system from leaking. Make sure there is no gel debris on your glass plates and that the plates are firmly pressed against the rubber tubing in the gasket. The key is the contact between the plates and the tubing. Keep them in perfect contact to avoid leakage.

If you are slightly paranoid about this (like I am!), fill the cathode with water and see if there is an obvious drop in water level after two minutes. If not, you are good to go!

Know the risks of reusing your buffer

It is economical to reuse your gel running buffer, but the longer you store these buffers and the more times you reuse them, the higher the chance of contamination. In worst-case scenarios, you will see garbage or non-specific bands on your gel, even in the empty lanes. To avoid this, use fresh buffer for the cathode (the inner compartment) so that the gels are always run in clean buffer.

Don’t sacrifice your gel for speed

The higher voltage / current / power you use to run your gels, the speedier your samples migrate – but it is also more likely to boil up your gel.

To prevent gel overheating, consult the manufacturer’s instructions to choose a reasonable voltage. Remember to pay attention to the specific conditions required for gels with different compositions.

After the run, feel the buffer in the cathode with your finger (unplugged and wearing gloves!!) to check if the setup has overheated.

If you want to run your gel quickly, consider putting an ice pack in the gel tank or moving the whole setup to the cold room.

Do not break the gel

It all comes down to this. Your SDS-PAGE is done and you need to take the delicate gel off the glass plates. Use a scraper to gently separate the two pieces of glasses. Do not fight against nature – let your gel pick whichever side it wants to stick with. Again with the help of the scraper, gently slide your gel out to a container. Practice (and patience) makes perfect.

When working with really delicate gels (low percentage / gradient gels), here is a handy trick: after you have separated the glass plates, put the plate with gel under slowly running water and catch the sliding gel with a container.

There you go, a perfectly run gel that is comparable to the pricy precast gels.  Imagine the time and effort you are going to save when you can run every gel to perfection.

Good luck!

SDS gel

2 Comments

  1. Jacob on November 19, 2015 at 10:21 pm

    I’m just starting the fun with SDS-PAGE but had to solve quite a lot of problems on my own because no one around known anything about running gels (yay!) First of all my mini setup leaked so I used parafilm trick… It still leaked so I resorted to using a small (~300 µl) amount of resolving gel, waiting it for solidify and then adding rest. It’s not perfect and I still wonder if it interfered with my assay but it kept me going. Second my gels got smiley so i overkilled it, using refrigerated buffer and cold packs added to the outside. Still I got some smears and wished to have more obvious bands but I’m getting there…

    Also I had no bromophenol blue lying around and was too lazy to order some, so I used SDS-PAGE loading buffer without the dye and added some DNA Loading buffer directly to the sample just prior to running gel.

    • Soham Bose on June 15, 2016 at 12:36 pm

      Hi Jacob…. I used to have some problem with gel leaking (I don’t have a biorad apparatus), here’s how I overcame the problem….
      First, align the glass plates (remember to clean them first with water and then with 70% EtOH followed by complete drying) with the spacers in between them. Completely wrap three sides of the glass plates (i.e, right , left and bottom part) with STEEL GRIP also commonly known as Black Tape. Then prepare your resolving gel mixture just don’t add APS. Take 1ml of that mixture in a separate 1.5ml eppendorf. To it add 3-4 times APS that you usually add in the resolving Gel mixture. Quickly Mix the contents in the eppendorf and add 800µl it in between the glass plates from one side (you can also confirm it from the leftover 200µl in the eppendorf)… It will solidify very quickly, we call it sealant. Then add the resolving gel followed by stacking gel according to your experimental need. You can always cut the sealant before transferring in case of western blot. This sealant part wont interfere with your experiment… Hope this helps

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