Currently, I use icIEF technique to determine pI of a protein. I see several peaks (more than 15) in profile. I want to use the neat protein to determine its pI and do not want to introduce any denaturant or reductant.
Any thoughts, suggestions?
Thank you
The isoelectric point of a native protein depends on which ionizable residues face the solution and which are buried in the interior of the structure, particularly when they are involved in contact ion pairs. Further you have to worry about PTM variability (e.g., partial phosphorylation or glycosylation). Then you have sialic acid variation to deal with on the glycosylation (a sign of aging). For example you get 7 silaloforms of transferrin with the asialo form up-regulated in chronic alcoholics. On BSA or HSA, you often see multiple isoforms due to the binding of other peptides to the core protein. Not to mention the monomer/multimer issue. It is always best to denature a sample to determine whether you are dealing with structural differences or PTMs. If multimeric, you should be able to see that on a native sieving gel.
Perform SEC fractionation to separate multimeric and monomeric variants first (a kind of non-reducing, non-denaturant native mode application). After this stage, deglycosylation is needed (PNGase) to convert the glycoform-presenting monomer to its non-glycosylated counterpart (I am not sure about the need for additional o-glycosidases and/or phosphatases treatment). By doing this you may able to get the pI of your protein utilizing gel-based IEF, icIEF, and chromatofocusing (IEX) techniques.
(Note that, for efficient PTM removal, denaturing and reducing may be needed prior to enzyme applications which improve the chopping of the inner side PTM-related conjugates...Indeed, this must be handled in line with the physical-chemical information of your target protein)...
Thanks Luke V Schneider and İsmail Emir Akyildiz for your inputs and thoughts into it. So far, I have tried native form, heat denatured form, zwittergent treated form, heat+zwittergent treated form using 30, 50 and 100kDa filter columns but the protein still precipitate in capillary and there is no sign of improvement so far. Next, I want to try denature protein in presence of SDS and DTT and deglycosylate and try to remove SDS by buffer exchange and see if it works!! Any suggestions on how to completely negate the effect of SDS in the final prep?
IEF can concentrate a protein beyond its saturation concentration, leading to precipitation. This is a field strength issue. If you lower the field strength and wait longer the bands will be more diffuse, but you should still get separation without the precipitation. SDS doesn't solubilize every protein. If you had problems with a zwitterionic surfactant, you may still have problems with SDS. You might also try running with zwitterionic non-detergent sulfobetanes. These help with more lipid-soluble proteins and are fully compatible with both IEF and PAGE methods even at 0.5 to 1 M concentrations. Glycerol can also be helpful (e.g., 20-50%) in the running buffer as it doesn't add current but helps solubility of some proteins.
SDS is virtually impossible to remove by buffer exchange or by absorption. The NDSBs are readily removed by buffer exchange.
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