I am working with a protein of approx 35kDa, in SDS PAGE the band is coming at approx 72 kDA. If it is in Dimeric form then why it is not coming as monomeric form. Laoding buffer contains 5% BME, samples were boiled for 10 min.
First of all, are you certain that the 72 K protein is your protein of interest? Does it light up with a specific antibody on W-blot or have you tried mass spec on it to identify it?
If it is your protein of interest then it is unlikely to be dimerising solely through disulphide bridging otherwise BME would resolve the dimer. However, it's worth trying other reducing agents such as DTT and/or TCEP before concluding this.
Boiling in SDS should resolve dimerisation through hydrophobic interactions.
Try adding sodium chloride to 0.3-0.5M final in the sample before boiling in SDS/BME loading dye to resolve the dimer.
Changing the pH to 2 points above or below the pI of the protein might help.
Finally try denaturing with chemicals (8M urea, 6M Guanidine hydrochloride) or using 0.4M arginine to disrupt the dimer.
Hope this helps!
Sometimes, the protein of interest gets glycosylated after post-translation at several positions in its sequence. In that case, the protein appears as a smeared band at a position higher than its actual size. To resolve this, the protein first needs to be deglycosylated using enzymes such as PNGase F. Just check if your protein is a glycosylated one.
First of all, are you certain that the 72 K protein is your protein of interest? Does it light up with a specific antibody on W-blot or have you tried mass spec on it to identify it?
If it is your protein of interest then it is unlikely to be dimerising solely through disulphide bridging otherwise BME would resolve the dimer. However, it's worth trying other reducing agents such as DTT and/or TCEP before concluding this.
Boiling in SDS should resolve dimerisation through hydrophobic interactions.
Try adding sodium chloride to 0.3-0.5M final in the sample before boiling in SDS/BME loading dye to resolve the dimer.
Changing the pH to 2 points above or below the pI of the protein might help.
Finally try denaturing with chemicals (8M urea, 6M Guanidine hydrochloride) or using 0.4M arginine to disrupt the dimer.
Hope this helps!
Dear Rakesh:
The molecular mass you see in SDS-PAGE could be due to two different reasons:
1.- Is it an halophilic protein? If so, you have to keep in mind that molecular mass of halophilic proteins is overstimated by SDS-PAGE due to the high amino acidic residues content these proteins have.
2.- If it is not an halophilic protein, then you could have a dimer which is not completely denaturated under the conditions you have used. As mentioned above, try to increase temperature and time of protein boling (using a samples buffer containing high mercaptoetanol concentration and increasing SDS concentration too). Other reagents such as DTT may also help.
Good luck
Rosa María Martínez Espinosa
Dear Rakesh,
sometime happen that SDS stabilize the protein. You may denature protein at higher temperature without having SDS then you can incubate with SDS for PGE. For reference you may a biochemistry paper by Javed Masood Khan. In this paper author tried to show the role of SDS in protein stabilization.
Hello, I think your protein may be annealing back after you reduce it. So you need to alkylate the reduced sulfite bonds. Several companies "carry reduction-alkylation kits" . Example Bio-Rad Cat #163-2090 or you can do it yourself. It is a good practice to alkylate your samples every time you reduce them. Good luck.
- Badges
- Science method