I want to quantify a homotetrameric protein (non-covalently bond) and I have tried several ways for breaking up the subunits such as DTT, b-mercaptoethanol, Urea and Urea + thiourea but I get mixtures of mono,-di, tri- and tetramers. Does anybody have some suggestions about stronger treatments to apply?
DTT and b-mercaptoethanol are reducing agents. Urea and thiourea are chaotropic agents and they do not reduce proteins. The reducing action of DTT and b-mercaptoethanol are an equilibrium between reduced and not reduced. It is normal to obtain a mix between mono, di, tri and tetramers. If you would like to increase the proportion of monomer, you need to increase the amount of DTT. You could add some urea (up to 8M) and a detergent as SDS, NP40 or Triton-X100 in order to eliminate the 3D structure (but this do not reduce the proteins, just help to generate monomeric protein).
You could use TBP (Biorad), it is an another reducing agent, stronger. Do not mix TBP and DTT or B-Me.
One consideration is what you are doing wit hthe protein after you denature it. If it is for a Western blot, then standard conditions should work, a combination of heat and reducing agent. If you are trying to be gentle as you are planning to do something else with the protein, you have a more difficult situation, as you need to maintain a reduced material in an oxidzing environment, namely an aqueous solution exposed to air. Also, as the tetramer is not bound by disulfide bridges as you sugest, the interactions will be a mix of ionic and hydrophobic, most likely, depending on where the subunits interact. Full denaturation works, as you disrupt secondary and tertiary structure along with quarternary, but trying to get normally folded subunits fully separated would be difficult, anything that disrupts those bonds may hit internal ones maintaiing lower orders of structure as well.
Hi Stephane,
Thank you for your reply.I already tried 9M of urea and I did include 10% SDS in all treatments (sorry I forgot to mention this). I used 20 mg/ml of DTT, how high do you think I could go? Thanks!
Regards, Els
Hi Michael,
I just want to quantify my monomer via Western Blotting, so the whole thing can be denaturated. The thing is I did a 2-D DIGE and did detect the monomer. But now I have switched to Western Blotting, I seem to detect mostly the tetramer. I also don't see a difference in the amount of tetramer present between the different treatments, so it doesn't seem like one is working better than the others. Thank you for any suggestions you can offer.
Regards, Els
I have used a buffer with 50 mM DTT for Westerns. Of course, that buffer is diluted to 25 mM by the sample. WIth the SDS and heating to 65 C for a few minutes, multimeric proteins generally give single bands of the expected molecular weight on Westerns, though that is limitied to the ones you check. If you are loking at only a few proteins out of thousands that may be in a sample, do you ever know if all of them denature fully?
Classically, I use up to 100mM of DTT (sorry you need to calculate in mg/ml...)
To increase DTT activity you could increase the pH above 8 with a Tris solution, the pK of the thiol (-SH) is around 8.3.
I even put it at 95°C for 10 minutes and it still doesn't break up...Is the 100 mM before or after you dilute it 1:1 with the sample?
Yes it is final in the sample buffer. In addition, you need to maintain this concentration in the loading buffer. Do you check your pH ? Basic ? you could use pH paper.
Do you try urea ?
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