DMSO absorbs strongly in the UV, it will not work well with CD. Arginine, although it frequently works well for protein folding, both absorbs and has its own CD signal. You can either try adjusting the salt concentration as suggested, try switching chloride for another more kosmotropic anion, and/or try a non-detergent sulfobetaine (some should be relatively UV transparent)

Although kosmotropic anions usually stabilize proteins over more chaotropic ones (i.e sulfate is usually more stabilizing then chloride) many exceptions exist.

Also note glycerol can be purified by passage through a column of activated carbon to remove some of the UV absorbtion. Chloride is relatively safe in comparison in terms of absorption compared to the others

http://www.sciencedirect.com/science/article/pii/S1359027896000089#bib13

If you know your protein is a little hydrophobic, then expecting it soluble in PBS is not wise. You might try to have it in a variety of salt concn and see what works best. Salt ions modulate H-bonding capacity of the proteins and changing the concn will alter the conformation of the protein--making it more soluble or more precipitable. You might also want to alter the solvent pH. However, the worry will be this: how good an idea will it be to do CD on a protein with altered conformation?

Another alternative will be to have your protein in a variety of aqueous solutions of differing DMSO concn. Providing both hydrophilic and hydrophobic moieties in the solvent will keep your protein in a native conformation, since it is a little hydrophobic; but I may be wrong! DMSO, if it keeps it soluble, will be ideal since its max absorbancy is at 268nm, well beyond your unacceptable range of 190-230.

To try the above, have your protein diluted in the respective salt or DMSO solutions. Do not go for dialysis; this is for two reasons: (i) dialysis invariably leads to precipitation of all proteins, and (ii) you can monitor dissolution in small volume very effectively. For example, dilute 10ul of your concentrated protein prep (much of which you may expect as precipitate) into 90ul of various salt/DMSO solutions, let is gently rock at various temperature. Then, centriguge really hard. Save the nearly-100ul supernatant in a separate tube and add 100ul of a suitable buffer to the residual pellet--even if you do not see any. Now, compare the supernatant with the respective pellet to ascertain how much was solubilized in what condition. One simple way to do is by WB, if you have an antibody for the protein. If not, you will have to search for alternative detection techniques; it is difficult to think about it without knowing what it is.

Once you figure out what solution is best, then you can remove urea/glycerol or whatever by dialysing your protein sample with the selected solvent at selected temperature. You do not worry now for precipitation, since you are already providing the solubilizing condition! Good luck.

But in case you find a better method, do come back here and tell us what it is.

I think one can use at least 0.6m urea without it un folding. If you have an activity assay, I would do this, or try putting your protein in different concentrations of urea, and possibly arginine ( to prevent aggregation). And then just use phosphate buffer without the salt, as this could make things worse perhaps. So I would take your refolded protein, and set up several different buffer conditions, and dilute it into the different buffers, and see which one gives the best yield in terms of soluble protein.

Or if sucrose doesn't absorb at a 220-260nm, I would try to exchange the sucrose for the glycerol. But I don't know if PBS is such a good idea either. Maybe try phosphate buffer first, ie about 20mM.

You can also buy refold kit buffers already made, and see if your protein likes anything else besides the buffer you already use.

Can detergent be used when takins a CD spectra. That would certainly help as Sven has suggested, if a Reliable CD spectra is obtained with detergent present.

I looked up the spectra of sucrose and it looks like it does absorb pretty well in the 200-300 regions. Many detergents absorb well in the 200-300nm region. Any suggestions Sven?

If your protein aggregates due to its hydrophobicity it might be an option to dialyse against Phosphate buffer with low concentrated detergent like SDS or CTAB. This might stabilize your protein, too. However, one should check their absorbance in advance.

If this attempt fails due to high absorbance of the detergents it may worth having a look at the Hofmeister series and replacing NaCl and/or KCl by a low concentration of Calcium iodide.

Marcia, urea will have, I guess, max absorbancy at a little earlier than 222nm, not sure of the exact value. This is owing to the carbonyl bouble bond. Even as 0.6M is not strong enough to denature the protein, it is pretty high a concn for absorption spectrum. Also not sure why you suggest phosphate buffer (PBS is based on phosphate buffer, which Marisol is already using); unless you are suggesting a variety of phosphate-based buffer derivatives (varying phosphate concn, varying salt type/concn). MOPS is not an option, as are chloride or citrate ions. If using Tris-based buffers, the pH adjustment must be done with sulfuric or phosphoric acid. By far, phosphate buffer is the safe bet, but since that is causing the protein to aggregate, I felt spiking with DMSO might help. Most detergents will absorb in the unacceptable range. Compounds with double bonds or cyclic moieties should be avoided. The more inorganic the solvent, the better; but there has to be some aliphatic component in the solvent so as to keep this troublesome protein soluble.

DMSO absorbs strongly in the UV, it will not work well with CD. Arginine, although it frequently works well for protein folding, both absorbs and has its own CD signal. You can either try adjusting the salt concentration as suggested, try switching chloride for another more kosmotropic anion, and/or try a non-detergent sulfobetaine (some should be relatively UV transparent)

Although kosmotropic anions usually stabilize proteins over more chaotropic ones (i.e sulfate is usually more stabilizing then chloride) many exceptions exist.

Also note glycerol can be purified by passage through a column of activated carbon to remove some of the UV absorbtion. Chloride is relatively safe in comparison in terms of absorption compared to the others

http://www.sciencedirect.com/science/article/pii/S1359027896000089#bib13

Anil, I also do not think detergent will work, as you and others have pointed out. I just thought maybe someone had tried a certain detergent that would be good with CD. I had unfortunately never heard of one. In terms of DMSO, I was concerned not only of its strong absorbance close to 230nm but also that most proteins don't tolerate concentrations above 1 %. At any rate, I like Jeff's suggestion to use the purified glycerol as a first try, and then maybe to solubilize with the non detergent betaines and possibly arginine additives. In terms of the phosphate vs. PBS, it is likely that the salt in the PBS is what is the problem, but I like your idea also about switching to something else that is completely different. The proteins I worked with seemed to always be more soluble in tris based buffers rather than phosphate.

Thank you very much to all for your suggestions. I will definitely come back with the outcome. To answer Sven's question, my protein is a cystein protease found in the lysosomes of a parasite.

The refolding buffer I use is composed of: 0.1 M Tris-Hcl, 0.001 edta, .25 arginine, and 20 % glycerol.