we have tested several precipitation methods (methanol/chroroforme, ethanol, ...).

It seems, that ethanol precipitation works best for our "test protein". Simply add 9 volumes ice-cold ethanol (100%) to one volume of buffer containing the protein in 8M urea. Incubate at least 1h at -20°C and then spin down the precipitate. Wash the pellet with 90% ice-cold ethanol, remove the supernatant as good as possible and resuspend the pellet in a suitable buffer. We used PBS with 0,1% SDS - it worked perfect!

What about using purification columns? More expensive, but you can somehow control which chemicals you want to get rid off. Solid-phase extraction technologies could be adapted to your proteins. We use "Oasis columns" from Waters to extract proteins from urine before LC-MS analyses. Perhaps it could be adapted to your process?

GFC columns can be used for this case....gel filtration column- sephadex packed or Dextran packed either ready made columns(prepacked in case if you have instrument ) or ordinary chromatography column in gravity flow

Size exclusion chromatography is for sure a good option, but i guess that if you have to choose between this and dialysis the latter is easier and more economic..but why not dialysis??

thank you for the answers - but GFC columns or other columns will not work since the protein usually precipitate after removal of urea - then it will be lost in the column.

At the end of the process, the protein has to be in a physiological buffer which limits the use of solvents.

You can try to add octylglucoside, 1% before dialysation

thank you for the comment, Marco.

Dialysis works usually very well (protein precipitates) - but since we have only small amounts of protein we lose a lot sticking at the dialysis membrane or even at tips / tube walls. We have tested the addition of Triton X-100 without big success.

I have a rather unorthodox idea. How about you precipitate them both in a glass vessel (to limit loss on plastic) using lyophilization (for example). Then extract urea with ethanol (your protein should still be precipitated). Then, lyophilize again to evaporate the ethanol and redissolve in a buffer of choice?

The easiest way is to dilute the protein to bring down the urea concentration 1000 fold lower and while you dilute it just make sure you keep on stirring the solution if you plan on refolding the protein. The protein can be then concentrated down by buffer exchanging using amicon filters

thank you for all the suggestions.

since we are handling only small amounts of protein (1 - 2 mg), dialysis results sometimes in a very dramatic loss of protein due to precipitated protein sticking everywhere (membrane, tube, tips). the same is true for refolding by dilution followed by concentration using amicon filters.

I like the precipitation idea from Iva and we will try it out. But we have to do it in small scale and I think the poor solubility of urea in ethanol could be a problem.

Thank you Reinhold! I thought of that too. The solubility should be about 20g per liter of ethanol, and about 170g/L in methanol. Either should be fine, and of course, many rounds of small volume shots + some heat (maybe). Good luck and let us know if it worked!

If you are working with volumes less than 1mL, you could either cut or melt a hole in the cap of a 1.6mL or 2mL Eppendorf tube (~1/2 the diameter), cut a small square of dialysis membrane larger than the diameter of the tube's cap, add your sample to the Eppendorf tube, then close the cap carefully (to avoid cutting/breaking the membrane) with the membrane in between, securing the membrane between the tube walls and the cap. I have used this method at least a couple times to dialyze a protein solution to precipitation, and then centrifuge the tube after dialysis to collect all of the precipitated protein before removing the residual supernatant.

we have tested several precipitation methods (methanol/chroroforme, ethanol, ...).

It seems, that ethanol precipitation works best for our "test protein". Simply add 9 volumes ice-cold ethanol (100%) to one volume of buffer containing the protein in 8M urea. Incubate at least 1h at -20°C and then spin down the precipitate. Wash the pellet with 90% ice-cold ethanol, remove the supernatant as good as possible and resuspend the pellet in a suitable buffer. We used PBS with 0,1% SDS - it worked perfect!

Thank you for the update Reinhold. I think this is an important lesson to remember. Sometimes the "old fashioned approach" is forgotten. I love how you simplified the protocol. Congratulations on your success.

@Reinhold Horlacher thank you very much for so simple protocol. Yet it would be lot helpful if we can site a reference for that in our papers.

Hi @Reinhold Horlacher, I like your simple method to remove 8 M urea. May I know how do you determine the 8 M urea is being removed completely? I use 8 M urea too to isolate spore coat proteins from Bacillus licheniformis. What would you do after precipitate your protein sample?

Let say if I precipitate my spore coat proteins samples in 8 M urea, can I use size exclusion chromatography (SEC) to obtain a single pure protein rather than a mixture of spore coat proteins?

I tried using 8M urea to equilibrate SEC and run my sample under 8 M urea denatured condition. When I ran for gel electrophoresis, a single collected fraction from SEC did have few protein bands appeared on the gel.

Can someone give me some recommendations on it? Many thanks.

@saurabh: we have no publication you can cite. What about: personal communication R. Horlacher, Trenzyme GmbH (Germany)? Thank you.

@Kokwei: we do not determine the Urea content after precipitation since trace amounts are no problem for us. You should be able to run the SEC in the presence of urea as you have done, but due to the viscosity of the buffer it will not be easy. But keep in mind, that you will have denatured proteins and the behaviour of the column will change and you could not compare the results you obtain with runs you have done under native conditions. Therefore, you should "calibrate" the SEC also under urea conditions.

@Reinhold Horlacher. I have one question. Removing urea prior to trypsin digestion gives me less recovey when i resuspend protein directly in ABC. I was thinking to remove Urea after trypsin digestion. Is it prossible to repeat same protocol for peptide recovery???

@Reinhold, I calibrated the SEC with 8 M urea. It seemed I coudn't get a single band on SDS-PAGE gel.

@Saurabh

Iprotease will not cleave in 8M urea, you have to bring down the urea to at least 4M (better 1 or 2M).

I think, the precipitation of small fragments will also be a problem - but we never tested this issue.

@Kokwei

do you see some purification? depending on the complexity of your sample you may not be able to get a single band. please check also the parameters of your SEC column (sample volume, flow rate, range of seperation, ...)

You can try desalting or buffer exchanges on a concentrators

Alternative to dialysis is ultrafiltration where your urea buffer can be replaced by your buffer of choice. Methanol/chloroform precipitation is also a good alternative, but if you had urea, this means that your proteins are denatured and it might be challenge to resolubilised the denatured proteins in an acquous solution without urea. Thus the progressive removal of urea by dialysis or ultrafiltration might be a better alternative if you need to get your protein in an acquous buffer.

We use solid phase extraction spin C18 columns to remove urea (and other salts).. it is effective and efficient. I use it regularly for my MS and I am very satisfied with it. There are plenty of them available from Empore, Microspin, Macrospin..

Regarding urea during trypsin digestion, the recommended concentration is close to 1M for Promega trypsin. Diluting the sample will help and then using SPE columns will be a effective to remove urea and recovery of peptides..

A couple of folks have asked the question. For many years I have used size exclusion chromatography using the "Spin Column" method to remove/desalt solutes away from macromolecules. At first, I demonstrated to my satisfaction that I could remove high concentrations of sugars using coloured dyes. However, not being satisfied with what my eyes could see of colour, I tried using a C14-labelled compound and the effluent from the spin column yielded background C14 counts. The trick in your case is going to be making sure that your column has sufficient bed volume to exchange all of the urea out in the top portion of the column.

In my case, I was preparing my own G-50 size exclusion gel columns to meet with my particular application. The nifty part of this process is that once you obtain the appropriate spin conditions, the volume you put on top of the column is recovered 100% in the effluent and there is complete solvent exchange with the bed volume.

The classical method is to prepare your column where the sample volume is, in your case, much smaller than 10% of the bed volume...I'd press toward 1% if you can. Your G-50 is equilibrated with the end buffer of interest. Allow all of the liquid in the column to come to the top of your column (I prepared my spin columns in syringe bodies with a frit or glass wool at the bottom...frit is better if you can get one...I often cut mine to size with a cork bore.) Place your column/syringe into a test tube and spin at 1000 x rpm for 1 minute in a table top swinging bucked centrifuge (Beckman, frequently used for sedimentation of cells). This first spin is meant to spin out just the void volume of your G-50 column. The syringe is then placed in a second tube, your sample applied to the top allowing it to wick into the G-50, and tube spun again at 1000 x rpm for 1 minute. The effluent should have the same volume as that added, would be in your exchange buffer, and should have no urea. It does work nicely and can easily be tested without your protein. I was able to exchange out very high sugar concentrations.

Good luck. It will work for you...you just have to play with the speeds, rpm/g-force/time, etc. so that your applied sample volume is exactly recovered in the effluent of the spin column.

Using small anion exchanger column chromatography to remove urea is fast and easy approach. A gel-filtration column is also OK, but takes more time.

As I stated above and related to Niyaz's comment, a spin column takes all but 2 minutes in the cold and the urea concentration is zero with no change in protein concentration. Traditional gel filtration and ion exchange column approaches will always generate a ~4 to potentially 10 fold decrease in protein concentration to which you would need to add a protein concentration step.

Good luck.

Follow up on Urea dialysis, wanted to ask if it can be done at 4C or should be at room temp. Explaining my situation, my protein is soluble in 8M urea but to further purify it, I cant use 8M urea so I need to remove it and the ppt I will dissolve in some another solvent. Will urea ppt in 4C and will be also be in my protein ppt. I have to do O/N dialysis.