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Bis-Tris Gels: Sharpen Up Your Protein Bands

Previously, Nick talked about SDS-PAGE. Today I am going to tell you about a tweak that will improve your SDS-PAGE protein gels.

Add Some Bis

It involves using Bis-Tris gel buffers. Although Bis-Tris adds a considerable cost to the technique, it has several advantages:

1. Bis-Tris gels are acidic, in contrast to the alkaline conditions found in conventional SDS-PAGE gels. This supresses cysteine reoxidation, which prevents proteins from cross-linking via di-sulphide bonds in the gel.

2. Sodium bissulphate, a reducing agent, is present throughout the buffer system. Unlike conventional PAGE, this means that the reducing environment is maintained all the way through the gel, which also helps to prevent disulphide bond formation.

3. Low molecular weight proteins do not run faster towards the end of the gel.

4. Conventional PAGE protein gels degrade after a month or two as the acrylamide breaks down to acryclic acid. This does not happen with Bis-Tris gels, which means they have a much longer shelf life.

Together, these factors mean that the resolution obtained on Bis-Tris SDS-PAGE gels is significantly greater than with conventional SDS-PAGE gels.

How to Make Your Own Bis-Tris Gels

Here’s how to make a Bis-Tris gel.  The method is not very different from the conventional methods of casting protein gels, just replace the Tris buffer that you use in the stacking and resolving gels with the Bis-tris buffer and omit SDS from the gel. Run at at a constant voltage of 150V.

Use this buffer to separate small proteins (2-50 kDa):

5X Low MW Running Buffer

250 mM MES
250 mM Tris
5 mM EDTA
0.5% SDS
Add sodium bisulfite (running buffer reducing agent) to 5 mM (add fresh before run) from a 1M stock

Alternatively, use this one for higher molecular weight proteins (>20 kDa).

5X High MW Running Buffer

250 mM MOPS
250 mM Tris
5 mM EDTA
0.5% SDS

Add sodium bisulfite (running buffer reducing agent) to 5 mM (add fresh before run) from a 1M stock

200X Running Buffer Reducing Agent

1 M sodium bisulfite

Add to running buffer at 5mM final concentration

3.5X Gel Buffer

1.25 M bis-Tris (pH 6.5-6.8 with HCl)

Bis-Tris gels were developed by Tim Updyke and Sheldon Engelhorn for Invitrogen. Similar gels are marketed by Invitrogen under the NuPAGE label.

Check out this page for a full description on running Bis-Tris protein gels.

If you try this, or already use them, let us know how these gels work for you.

Originally published on September 12, 2008.  Updated and revised on July 21, 2016.

25 Comments

  1. wes on August 16, 2016 at 9:47 pm

    Nice article, Bala. One quibble, I think you meant cysteine rather than cytosine in your point #1.

    • Dr Amanda Welch on August 17, 2016 at 6:31 pm

      Thank you for catching that!

  2. Bill Gillis on February 13, 2015 at 3:53 pm

    Do you have any idea about the sterilzation (autoclave or filter), storage (cold, out of light), and shelf life of these solutions?

    I found that sigma says bis-tris solutions can be filtered or autoclaved, and are stable at 2-8*C (no comment about room temperature)
    https://www.sigmaaldrich.com/content/dam/sigma-aldrich/docs/Sigma/Product_Information_Sheet/b7535pis.pdf

    Sigma provided no data regarding sodium bisulfite, but found reference from dow chemicals saying that a 10% solution (equal to 1 M of sodium bisulfite) is readily oxidized and is only stable for one week, whereas a 30% solution can last up to 6 months. Still no idea if it can be autoclaved/filtered
    http://dowac.custhelp.com/app/answers/detail/a_id/2261

    also, i’m assuming the 5X high-MW running buffer can be sterilized or autoclaved, but may turn yellow (not sure if that would affect the gel/transfer). Most mops solutions need to be protected from light.

    250 mM MOPS 250 mM Tris 5 mM EDTA 0.5% SDS No Need to pH.
    http://openwetware.org/wiki/MOPS

  3. ynokelpe on December 12, 2012 at 9:54 pm

    I have a quick question I ran into while using this procedure. The 8% gel I was making polymerized without the addition of TEMED. I know for sure I did not add TEMED. TEMED is just the initiator for the ammonium persulfate, but why shouldn’t it be necessary in these gels?

  4. TijsClaessens on July 23, 2012 at 11:14 am

    Hi Bala,

    Nice article.

    I was just wondering, can you use sodium bisulfite in regular SDS PAGE as well, or is it also pH dependent?

    Thanks in advance,

    Tijs Claessens

  5. Claus Jensen on October 27, 2010 at 2:06 pm

    Hi

    I had the same trouble with a distorted yellow band in the bottom of the gel. I think I have solved this problem. After you have mounted the gel in the running unit and added running buffer (I use Hoefer) just wash the bottom of the gel several times with a pipette in order to equilibrate the gel with the running buffer. I think this also explains why you never see this phenomenon with the precast gels from Novex, as these have a “vertical” gel connection to the running buffer thus giving a much better equilibration (in the Hoefer gels there are often bubbles in the bottom of the gel).

    Best Regards
    Claus

    • Ankit Patel on November 9, 2016 at 10:03 pm

      Hi Claus,
      I have the same problem with my Bis tris gels. Up to 3/4th of the gel runs nicely but after that the yellow front dye distorts in the middle. I submerged bottom of the gel in MOPS running buffer for few before loading the sample and starting the run but it didn’t help. Any advise on this will be much appreciated.
      Thanks
      Ankit

      • Aleks on June 20, 2018 at 4:41 pm

        Hi Ankit,
        I wanted to ask whether you could find a solution to your problem. I am experiencing the same with my gels and would be grateful for any advice.
        Kind regards,
        Aleks

  6. bala on February 11, 2010 at 7:50 am

    @Foncy
    If I’m right, the invitrogen MOPS buffer doesn’t have sodium bisulfite but they do recommend a special loading buffer which contains the reducing agent!

  7. Foncy on February 10, 2010 at 10:39 pm

    I am looking to try these gels out. I notice the invitrogen MOPS buffer recipe does not include the Sodium Bisulfite. Is this necessary?

  8. yc on January 9, 2010 at 12:05 pm

    I have no problem with these gels. Run great at 180V constant voltage.

    • Ankit on November 13, 2016 at 1:35 pm

      Hi YC,
      I have problem with my Bis tris gels. Up to 3/4th of the gel runs nicely but after that the yellow front dye distorts in the middle. The bands are sharp except this problem. Any advise on this.
      Thanks
      Ankit

  9. Dan on August 28, 2009 at 3:33 pm

    i’m pretty sure that’s the same buffer composition (just 20X, instead of 10X). My gels are great, nice sharp bands, etc, except for the problem at the bottom of the gel.

  10. Rex on August 27, 2009 at 4:08 pm

    Hi Bala,
    Thanks for giving the buffer composition. Any other input from you will be greatly appreciated.
    Dan,
    I also use the Hoefer system. If u find out any remedy, let me know aswell. Did u try the buffer mentioned by Bala? How was it? Are you seeing sharper bands? My bands were not as good as what I see in Tris-Gycine system.

  11. Dan on August 26, 2009 at 6:47 pm

    I’ve tried making my own buffer or using commercial buffer; doesn’t make much if any difference. Rex – are you also using a Hoefer system?

  12. bala on August 25, 2009 at 12:01 pm

    Guys, I’m still trying to get this system right! invitrogen suggests the following running buffer composition.Give it a try..

    MOPS SDS buffer 20x (500ml)

    MOPS 104.6g
    Tris 60.6g
    SDS 10.0g
    EDTA 3.0g

    Add water to makeup 500ml

  13. Rex on August 24, 2009 at 5:06 pm

    Hi Bala,

    I recently tried the BIS-TRIS system as described by u. I faced the same problem as described by others here. The dye front turned acidic and distorted at the end. I changed the whole running buffer and tried to run it again. But no luck.

    Whether some one sorted out this problem? Kindly let me know.

    Thanks in advance.

  14. Dan on July 23, 2009 at 7:38 pm

    I have the same problem with the mini-Hoefer system gels with the dye front getting wavy and acidic; I still have the problem using invitrogen’s LDS loading buffer. If you use a LOT of buffer in the bottom chamber, seems to be a bit better (and sometimes runs fine). I haven’t tried changing the buffer halfway. Anyone else have a solution to this problem?

  15. bala on July 21, 2009 at 10:21 pm

    The normal transfer buffer with methanol works well, however I’ll post an additional buffer composition that Invitrogen recommends, in a day or two.

  16. Francesco on July 21, 2009 at 9:47 am

    I´m wondering which type of transfer buffer I should use for a bis-tris gel.
    Would it be necessary to use the running buffer composition with the addition of methanol?
    Do you have any hint?

  17. Venkatramana on January 10, 2009 at 12:19 pm

    When I tried this method I found that the protein bands are wavy. What could be the reason

  18. Liam on September 29, 2008 at 2:39 pm

    Bala

    I’ll give it a try and report back.

    Thanks
    Liam

  19. Bala on September 23, 2008 at 9:22 am

    @Liam, I just had confused your problem with another one that i faced, at times the gel might run pretty slowly! which is because of the buffer exchange.Regarding your query,you could try adding some sodium bisulphite in the loading buffer (lamelli) – i did not have this problem when i used the NuPAGE loading buffer!

  20. bala on September 23, 2008 at 9:04 am

    Liam, Infact i did face this problem – i was advised to run the gel at constant current (and i used a maximum voltage of 200V).The rationale being that the reservoir buffer completely displaces the gel buffer.The trick did work! 😉

  21. Liam on September 23, 2008 at 7:07 am

    Hi Bala

    I was wondering if anyone else had tried to run this gel system on the smaller gel tanks, like the mini-Hoefer system (8×5.5cm). The gels run beautifully until about 1.5cm from the end, when things go all wavy, the dye front goes acidic, and basically the gel distorts (although after staining it returns back to normal). I’ve measured pH, run at low mA (as opposed to the patent which describes to run at 150V), and have remade the buffers very carefully. Only thing left to try is to replace the buffer halfway through the run. Any ideas ?

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