It could be a degradation band that either co-eluted with your protein through all the chromatography steps, or the degradation may have occurred after the chromatography. It could be another protein that binds to your protein. It could be residual TEV protease that didn't get removed somehow.

Make sure you use protease inhibitor cocktail throughout the purification, do all the steps at 4oC, and keep all the fractions on ice as much as possible. Perform the entire purification procedure as quickly as possible. These suggestions should help minimize proteolytic degradation.

Make sure you are completely reducing your protein during the purification and with SDS-PAGE sample buffer. Unreduced disulfide bonds could affect the apparent molecular weight on SDS-PAGE.

As it seems from your information your protein of interest seems to be in much higher concentrations than the contaminant.Having said that...

First,if you use a N-terminal His-Tag,the contaminant could also be a truncated version of your protein and thus gets eluted in the affinity.I suppose the contaminant might not be bound to your protein of interest as you seem to get your protein of interest at the right elution volumes in gel filtration.

Secondly,What gel filtration column are you using??,Make sure you do not overload the column as this would compromise the resolution capacity of the run for proteins that are nearby.

And third,I am just curious that did you concentrate your protein after the all purification steps & if yes,did you see this extra band before concentrating your protein or after that.

I think your protein oxidized during purification steps and dimer formed, compare with standards and controls: standard protein of your interest and its oxidized sample (oxidize some of your standard). Then run a gel and compare all of them to exclude oxidation. 

Protein Purification is not proper you can check with the standards to make sure that you get only the desired protein and make sure you collect only fraction of that particular protein or you can purify the fraction again

Thank you all for your answers and suggestions.

Adam; I used PMSF during my purification and kept adding it to my lysate every 30 minutes. All fractions were kept in ice immediately and the purification steps were done as quickly as possible. I also thought it could be residual TEV protease, but firstly; the size of TEV protease is around 27 kDa so it is a bit smaller than the band which I am seeing. secondly, TEV protease would not bind to the anion exchange column since it has a pI of around 9; so that is the reason why I excluded this possibility. 

Himanshu, my protein had a C-terminal His tag so it cannot be a truncated version of my protein. I am using the Superdex 75 10/300 GF column, and I honestly do not think I overloaded the column since I divided my sample into 6 each 500ul (since the loop can only take 500ul sample at a time. As for concentration my protein, yes I do concentrate after each purification step in order to do a buffer exchange, and I can see the extra band more obviously when the protein is concentrated.

Irada; if a dimer formed then I would see a band at 75 kDa. How do you usually oxidize the standard?

Bahja, you can oxidize your protein with 20 to 100uM H2O2 or more. Add to your standard in RT for about 10-20mins and then 20ug/ml catalase  (or sample buffer for gel and run gel) to stop further oxidation

Hi there,

If your protein is eluted as a monomer from SEC, I would say the lower band on gel comes from degradation though the presence of protease inhibitors. The question may be when does this happen as you haven't mentioned any of the pattern obtained after affinity and IEC? If you want to be sure about the content of this 30kD band, go for MS fingerprint to identify the missing sequence.

Hi, Irada,

Maybe there is a dimer which can be analyzed via SDS-PAGE.

Good lucky

Apart from incompletely oxidised disulfide bonds (you can check this reduction with DTT and alkylation with iodoacetamide) you should also consider cleavage during preparation of the sample for SDS-PAGE. Aspartate-proline bonds (among others) are vulnerable to cleavage at higher temperatures. Boiling samples for SDS-PAGE is common but usually unnecessary as denaturation is usually complete at much lower temperatures.

A final thought: have you calibrated your column so you can estimate the solution size of the protein? If your peak on SEC is larger than on SDS-PAGE, some kind of quaternary structure may be present. This may mask the presence of a smaller version of the protein you are interested in. Assessing both bands by MS will tell you if they are related – implying cleavage.

Is there an alternative start codon present in the coding gene? Can you (by calculation) obtain the second band size by translation at this eventual, alternative start codon? If this is true then the theory of a dimer between your full size protein and a truncated version can be true. MS of the additional band or immunodetection somewhere at the middle of the protein should give you the final answer, good luck.

Your protein may not be monomer and each monomer of the protein could be in different MW. When you run SDS-PAGE, the SDS breaks the links between monomers and each polipeptide with different MW may appear in different distance.

As stated by the other people in this discussion your protein could be a (hetero)dimer. I suggest to pick both band and identify the protein(s) by mass spectrometry. If you need help in the identification just contact me and we can do the MS analysis.

SDS-PAGE denatures protein and confers single charge to your protein. Interaction of your protein with electric field causes possible separation of dimers with hydrophobic interactions resulting in more than a single band. Try NMR and compare MW.

Dominique; I see this band from the first purification step (affinity) and it is carried over through all the subsequent purifications. You are right, I definitely need to send the bands for MS for absolute identification.

Kou; thank you for the manuscript. I never add reducing agents when I purify TEV protease. I add DTT to the digest reaction just after setting it up.

Craig; I also thought it could be from sample preparation for SDS-PAGE, so to rule out this possibility I run another gel without heating the samples and the band was still present!! As for column calibration; I haven't calibrated the column. Sending the bands for MS analysis will surely be by next step; thank you!

Ahmed; your suggestion makes sense- I need to check the sequence to see if that was the case; thank you!

Hi Bahja,

Does your protein contain any disulphide bonds?  If so it is possible that there is a proportion that has been proteolytically cleaved yet the protein remains held together by the disulphide bonds.  If this is the case it will behave almost identically to the uncleaved material as there could be as little as a single hydrolysed peptide bond difference between the two species.

As others have said Mass spec and or N-terminal sequencing on the two bands would probably be informative.

either your solvent system does not seperate the two compounds or there is isomerization during gel filtration