If you want to compare the banding patterns of the standard and the purified enzyme, you have to load equal amounts on SDS-PAGE. In the example you showed above the amount of standard is so low that even if it had the same banding pattern as the new preparation, the additional bands would be under the limit of detection of the staining procedure.

Regarding the appearance of "new" bands under storage (they actually look similar to those showing in your most recent enzyme preparation), one possibility that comes to mind is disulfide oxidation, and another, that there was some unaccounted for variation during sample preparation between the first and the second gel, and therefore the additional bands come from better disaggregation/solubilization of the protein aggregates that show at the top of the gel in time 0.

If you have a proper antibody, why don't you do a Western blot to try to find out more about the identity of the mistery bands?

Hope it helps!

One possibility is that the reducing agent in your sample buffer has become oxidized, such that a disulfide bond in the protein is not getting reduced, resulting in the upper band. Try fresh sample buffer or reducing agent.

Dear Alejandro Martin ,

Thank you very much for your answer and suggestions. I would keep all the conditions and concentrations same for all the samples and see how it goes.

Kind regards

Santosh

Dear Adam B Shapiro ,

Thank you very much for your time in answering the query. In one of the troubleshooting methods, I did try adding some fresh reducing agent to the sample buffer but it did not solve the issue.

Kind regards

Santosh

Santosh Dacha , this might help:

Article Origin of multiple bands of proteins on sodium dodecyl sulph...

Perhaps it might be wise to reduce/alkylate free cysteines before gel electrophoresis. Any decent protocol book on proteomics should give you a hint on how to do it.

Also, there are proteins which aggregate upon heating during sample preparation. Try heating the samples at 50*C for 30 min. instead of 100*C/5 min.

Hi,

I am agree with Adam, I think this is a denaturation problem of your protein samples.

First of all apply heat denaturation 5-10 min in 95-100o C for your samples before adding b-Mercapthoethanol to the sample buffer. Then cool your samples on ice and add b-MEtOH in to your tubes. If you add b-MEtOH before heat denaturation, probably the concentration of b-MEtOH drastically reduced.

If you reuse B-mercaptoethanol (as a denaturation agent) contained sample buffer, after a while the concentration of B-METOH reduces, so its effect. I suggest you use DTT as a denaturation agent (it does smell not as bad as bMEtOH) and every time prepare your sample buffer fresh (95% Sample buffer + 5% bMEtOH or 350 mM DTT final concentration).

I hope this helps.

Good Luck!

The second image protein bands doesn't seems that much crisper while compared to first image. You better check the Tris(pH 8.8 & 6.8) and AB solutions or again prepare them freshly before the run. Also the smear appearing in the second gel image may be due to several conditions like conductivity problem, running buffer concentration and the loading dye. " Coming to your query : Try to standardize your enzyme concentration for the protocol which you are following currently or else change your reference protocol.You said that the second gel was ran after 3 months so there may be loss of protein degradation in between these time. Even though you were storing the samples in -80 degree, the Protease inhibitor's activity will be slowed down or disappear. So go for fresh sample with the above said suggestions.

Did you use the same MW ladder in both gels?

I seem to me you assigned the 63 kDa standard differently in the two gels, and that makes you think the lower band is your protein. It may be exactly the opposite, the upper band is your protein and the lower band the result fro partial proteolysis.

Best wishes,

Rogelio