Dear Hyeontae Seo ,

There are several methods that can be used to de-oligomerize a protein. The method chosen will depend on the properties of the protein and the oligomer, and the desired outcome.

One method is to use a denaturant, such as urea or guanidine hydrochloride, to disrupt the interactions between the subunits of the oligomer. This can be done by adding the denaturant to the protein sample and then dialyzing it to remove the denaturant.

Another method is to use a detergent, such as SDS or Triton X-100, to solubilize the oligomer and disrupt the interactions between the subunits. This can be done by adding the detergent to the protein sample and then centrifuging it to remove any insoluble material.

A third method is to use a chaotropic agent such as thiourea, which can disrupt the non-covalent interactions between the subunits of the oligomer. This can be done by adding the chaotropic agent to the protein sample and then centrifuging it to remove any insoluble material.

Finally, you can use a method called size exclusion chromatography (SEC) which separates molecules based on their size by passing them through a column packed with beads of a specific size. The beads act as a sieve allowing molecules of a certain size to pass through, while larger molecules are excluded. This will allow you to separate the oligomers from the monomeric proteins.

Keep in mind that these methods are not guaranteed to work for all proteins and oligomers, and may require optimization for your specific protein.

Are you certain that this is an oligomer of your protein? Could it instead be a chaperone such as HSP70 or similar that is binding?

Can you describe your expression system, and do you have a gel to show us the molecular weight of a) your protein, and b) the contaminant band?

I guess you would like to keep the protein in its folded form and not denature it, which would happen if you followed most of the suggestions above.

First of all you need to know if it's really homo-oligomers or e.g. chaperones that migrate with your protein.

Urea and Guanidine are generally used in high concentrations for denaturing the protein. With urea, you can try to stay below 2 M, most proteins would still be folded but oligomers could dissociate, but as soon as you remove the urea it could form oligomers again.

You can't suggest SDS and Triton in the same sentence as if they were alternatives to each other. SDS is denaturing most proteins, Triton X-100 usually doesn't! Plus, centrifuging will definitely not remove one of the proteins because the detergents usually solubilize proteins. It's unlikely that you could remove one of them by centrifugation.

So, Triton or some milder detergents like Tween 20 or 80 could help dissociating oligomers.

Selective precipitation could work if the properties of the single protein, the contaminant and the complex are different enough, but again this should not be done with a harsh reagent like thiourea, e.g. ammonium sulfate could be useful.

Higher concentrations of salt (up to 1-2 M NaCl) could help dissociating oligomers and subsequent separation by size-exclusion chromatography could work.

What made you think that junk is oligomerized? Did you observe doubled or higher state folded MW of the target protein? Oligomers seemed as multiplied MW of the POI (if homo-oligomers as indicated previously) but this would be not typical since you are performing reducing PAGE...High contraction of SDS in the sample buffer and running buffer should de-oligomerize it already. Do you able to confirm by native page or MS analysis? In gel digested SDS-PAGE band of the junk can also indicate the oligomerization of the same protein.

This may be a stubborn interfering one also...Affinity purification can improve the purity but also prove whether it is oligo or a contaminant...

Otherwise, for efficient deoligomerization, the suggested ones above completely fulfilled the request...

If you perform a Western blot against your protein of interest, does the antibody also show a signal where these higher MW bands show up? That is probably the easiest way to confirm if they are oligomers.

For some proteins (especially membrane proteins), aggregation can be promoted by the high temperature typically used to enhance denaturation. Try denaturing at 37 degrees C or room temperature for 20 minutes or so and load alongside a sample that you've prepared using your normal method to see if there is any difference.

Are you lysing directly in SDS-PAGE loading buffer? Perhaps try lysing your cells mechanically and/or with a non-ionic buffer like Triton-X-100 or Tween 80, remove unlysed cells by centrifugation and then prepare SDS-PAGE samples from the supernatant.