Hello Christiane- Yes, it is possible because mobility depends upon charge/size ratio, and in SDS PAGE, charge is contributed by SDS binding so, for a single protein due to slight difference in SDS binding due to shape or incomplete denaturation of protein in question. We observed multiple bands of a thermophilic protein on SDS PAGE but several proteins classified as kinetically stable protein display his abnormal SDS binding. see following reference for a more general view.

Structural basis of protein kinetic stability: resistance to sodium dodecyl sulfate suggests a central role for rigidity and a bias toward β-sheet structure

M Manning, W Colón - Biochemistry, 2004 .

Besides this, to confirm the observation, you should do peptide mass fingerprinting by MALDI-TOF.

in Summary , it seems to possible for a protein!

Hello Christiane- Yes, it is possible because mobility depends upon charge/size ratio, and in SDS PAGE, charge is contributed by SDS binding so, for a single protein due to slight difference in SDS binding due to shape or incomplete denaturation of protein in question. We observed multiple bands of a thermophilic protein on SDS PAGE but several proteins classified as kinetically stable protein display his abnormal SDS binding. see following reference for a more general view.

Structural basis of protein kinetic stability: resistance to sodium dodecyl sulfate suggests a central role for rigidity and a bias toward β-sheet structure

M Manning, W Colón - Biochemistry, 2004 .

Besides this, to confirm the observation, you should do peptide mass fingerprinting by MALDI-TOF.

in Summary , it seems to possible for a protein!

Did you check the same with immunoblot? Just confirm it to avoid any further confusion(that one of the protein may be a contaminant, in least chance!).

Yes, Ananda, my next step is to perform a immunoblot, but I find very difficult not to be the same protein, because it went throught three consecutive chromatographies in specific conditions. Anyway, I know the blot is indispensable.

Thank you both.

Yes I agree Christiana, In my case and others, chaperones do elutue with the protein of interest even after multiple purifications. For your blot please use a specific monoclonal antibody not anti His antibody for example (it gave a positive signal with my negative protein as the contaminant "SlyD" has a long Histidine stretch on it).

Good luck!

Hi Christiane,

It is possible that proteolysis shortens your enzyme during expression by a few amino acids and this makes it insoluble. If so, the proportion of i.f.:s.f. would increase as a function of expression time (I don't know if you did a time-course experiment to optimize the expression time). I also suggest to use mass spectrometry to check that the different bands are the same protein form.

SDS and reducing agent (usually beta-mercaptoethanol or dtt) should fully denature your proteins and there should be no difference between the two samples. But note that you might not have enough reducing agent for very concentrated samples (e.g. inclusion bodies) resulting in an unstructured, but knotted, polypeptide (due to remaining cysteine-cysteine crossbridges) that has a smaller dynamic radius (and size) than the fully reduced form. Adding additional beta-mercaptoethanol when preparing sample would rule this case out.

Is the loading (size of bands) roughly the same for the two different protein samples? I've observed that this can sometimes affect the protein mobility.

Sanjeev's answer is certainly valid for proteins originating from thermophylic species. However, if your protein originates from a normal temperature organism, then it should be fully denatured on the PAGE. Otherwise, this problem would crop up much more often.

Hi Patrick, thank you for your answer. I think it make sense what you said about the reducing agent.

In relation to what you said about the loading, if you mean the amount of each sample that was loaded into the gel, it wasn't the same, even because the i.f. was a purified sample and the s.f. was partial, so the amount It was loaded comprised other proteins. Do you think it may be an explanation?

Regarding loading, I'm considering the total amount of protein in the lane. If it is very high then some distortions can occur. You may not have enough SDS to initially coat the protein in order for it to migrate as quickly as it should. Eventually, you will recruit enough SDS from the buffer in the stacking gel to overcome this problem.

We generally run 10 microliters of protein solution from each 1ml fraction of our FPLC runs. In the cases where we have high, pure protein concentrations of 5 mg/ml, we see some distortion of the bands to higher molecular weight.

I would expect retardation for i.b. (1 microliter of dense i.b. suspension would be 10-30% protein or 100-300 micrograms protein). Looking through the description of your problem again, I see the result is the opposite.

If you are running the two samples and a standard, you might want to consider running a few different protein concentrations to see if the migration changes as a function of concentration. Note, such an observation would not be due to dimerization at higher concentrations.

For protein purification, low concentrations look better for publications as the bands will be sharper and more true. For personal use, a higher concentration gives you a better idea of what other proteins might still be present in very low concentrations. So sometimes it is a good idea to run a couple of gels of good purification runs so that you have a good figure for your monthly report and any publication.

Another potential problem is salty samples. If one of the samples contains very high salt and the other very low salt then you could get a distortion. The PAGE buffer contains a decent salt concentration so high means more than 1 molar.

Dear Christiane, could be you have the mysterious dimer of your protein fully discussed here:

https://www.researchgate.net/post/Dimer_at_sds-page

Could you please show your gel picture, maybe the solution is there and you don't see it;)

Here it is Jan.

The C+ is the purified inclusion bodies

Right next to it, is the Capto S unbound, where you can see the upper band which should correspond to the soluble protein (Before the capto S I did an affinity)

Then I took this fraction and did a second affinity purification (arrows)

Sorry for this examination, I’ not from FBI;)

The C+ is the purified inclusion bodies

_________Native buffer or in urea/guanidine ?

Right next to it, is the Capto S unbound, where you can see the upper band which should correspond to the soluble protein (Before the capto S I did an affinity)

_________You mean the weak 80kDa or strong 75kDa?

Then I took this fraction and did a second affinity purification (arrows)

________Which of them you expected?

Keep in touch-:)))

Well, non-FBI Jan ..... lol

1 - The inclusion bodies were purified in denaturing conditions (8M urea)

2- I mean the strong one

3- The second affinity sample was the Capto S unbound.

Got it?

My primary diagnosis is you have the sample in C+ somehow pushed by urea, thus is migrating fasten. CaptoSUB and fr.12, 13 the band could be the same size, the difference could be due to the gel smiling. Wash the slots as I’ve described here:

https://www.researchgate.net/post/After_transferring_on_to_a_membrane_step_in_western_blot_my_marker_and_protein_bands_are_smeared_Why_does_this_happen2

Load C+near 12, 13 it will exclude gel smiling.

Keep fingers cross :)