If you have an activity assay for this enzyme, you can expose a high concentration of the enzyme to various concentrations of DMSO for various lengths of time, then dilute the enzyme/DMSO mixtures into the activity assay and measure the initial rate of the reaction. You have to control for the residual DMSO concentration in the final reaction by testing the effect of that concentration in the final assay when there was no DMSO in the original incubation.

@Adam B Shapiro. I have the activity assay and I can see the activity is goes down with increasing concentrations of DMSO. What I want to quantify how much folded protein is still in there. For example, if I have added 0.5 mg protein in the reaction, how much is left? maybe 0.1 mg is still active and 0.4 mg is denatured@.

Hi Malaikah,

I think clarifying the language will help you develop a plan, since you can have a partially unfolded protein that is still active.

Adam B Shapiro's suggestion sounds like a good one for addressing the activity of the protein. If the assay shows 30% activity relative to a control sample, you would nominally interpret it to have 30% active protein, 70% inactive protein.

If you want to ask to what degree the ensemble is in its native fold, circular dichroism can answer that. There are a handful of software packages that will interpret spectra in terms of percent helix, sheet, and random coil. I haven't mitigated DMSO issues with CD, but it looks like this paper could point you in the right direction: https://www.sciencedirect.com/science/article/pii/S2666166724000157

As pointed out by Jeremy Pronchik , it is not necessarily true that a given protein molecule is either fully active and in its native conformation or fully inactive and unfolded. A protein molecule's structure might be mildly perturbed by a low concentration of DMSO and retain partial activity. High solvent concentrations, on the other hand, would probably result in major loss of native structure of most proteins.

If a solution of a protein at a reasonably high protein concentration is substantially unfolded, it will almost certainly result in aggregation of protein molecules due to exposure of the hydrophobic interior. In this extreme case, a simple measurement by dynamic light scattering will reveal the presence of aggregated protein. This requires a special instrument, but if the aggregation is severe enough, it can even be detected with a UV-VIS spectrophotometer, which most labs have.

The effect of a solvent or other chaotrope on the stability of a protein can be monitored by measuring the melting temperature of the protein in solution, using either differential scanning calorimetry, or differential scanning fluorescence. Each measurement requires a special instrument.

Many proteins contain tryptophan residues whose fluorescence is sensitive to their environment, specifically the hydrophobicity of their environment. Using a spectrofluorimeter, you can measure the tryptophan emission spectrum to follow major and minor structural changes in the protein. This can also be done using far UV circular dichroism (CD).

Far UV CD, mentioned by Jeremy Pronchik is used to measure changes in the secondary structure elements of the protein. Loss of secondary structure would be a sign of a major loss of native structure. I think the presence of DMSO would interfere with this measurement, however.

In addition to all the good suggestions above, alternatively, zymography combined with native PAGE or Native PAGE alone with densitometry may also produce relative concentrations of the folded active amount of enzyme in comparison to the denatured inactive counterpart...Try BN-PAGE to separate the scrambled enzyme mixture by its shape and size only.

Thank you all for great suggestions. I think Native PAGE is the easiest way to start with.