First I would put a drop of the precipitate solution on a glass slide and look at it under a microscope – is it crystalline or amorphous? If it is crystalline you may be able to spin out a highly purified sample of your protein and re-solubilize it. If it is not crystals, you can adjust the bottom lighting you can easily see precipitate as a cloudy drop and try pushing the pH, salt or other additives up or down by pulling a liquid thread connecting with drops of different solutions. In this way you can test with a drop if any changes will re-solubilize your sample. Once you know what may help, you could switch buffers quickly by running your sample down a de-salting column instead of dialyzing, which is slow.

What was your buffers composition and protein concentration? To help you it is essential. Could be you concentrated your protein during dialysis, thus you have agregation...

Hello Piwowarski,

20mM Tris,pH 7.5, 150mM KCl, 5%glycerol and protein conc. was around 15microgram/microlitre.

This is probable glycerol sucks the water of your protein concentrating it. What was the protein concentration before dialysis?

Sorry didn't noticed, my fault;) Congratulations! You have obtained terrific amount of your protein I can only dream in case of mine:(

I know the proteins starts precipitating at the conc. 5 microgram/microlitre. If your protein volume decreased twice you had 30ug/ul final most of the proteins tends to crystallize or aggregate. If you don't need such a high concentration, dilute your prep prior the dialysis, or exclude glycerol, or both... Keep fingers cross!

We occasionally get precipitation during dialysis, I think some proteins do not like being in the presence of imidazole for long periods. One solution to avoid this problem is to use a desalting column which rapidly buffer exchanges away the imidazole.

Also 15 mg/ml is a pretty high concentration, you could try diluting it and performing the dialysis at a lower concentration, then concentrate it back down afterwards.

Hello Piwowarski and Goult,

I didn't get completely that what is the effect of high or low conc. of glycerol in dialysis?

Your precipitation is probably because of the high concentration.

During dialysis all the components tends to have equilibrium on both sides of the membrane. Since water is migrating fasten than glycerol, goes outside the bag to the glycerol rich buffer ("glycerol sucks the water "). This phenomena is widely used to concentrate proteins, the volume of your sample can decrease few times after ON process. Of course glycerol goes into the sample as well, thus after ON you have equal conc. in and out the bag. To avoid the volume changes you need to have equal glycerol conc. in the sample and buffer.

I recommend for the first try:

Dialyze in 250mM NaCl, 5%Glycerol, 100mM EDTA (pH7.5), 25mM Tris-HcL pH 7.5. with 2-3 buffer exchanges all 3-4 hours. Then dialyze into 150mM NaCl, 5%Glycerol, 15mM Tris-HcL pH 7.5. Your protein should be soluble.

Use a desalting column and you get the same result in 3 hours instead of 24h.

Spin column with suitable MW cutoff can be used to remove imidazole in several steps taking smaller amount of samples.

Dear Seema and Stefan,

Your techniques can lead to increased protein concentration in the sample. This should be avoided, since high protein concentration is up to now the main suspected causing the disaster happen here…

Dear Jan,

There are several types of spin-column of varying capacity (from 0.2ul,4ml.15ml. etc) which can be used accordingly. To avoid local high conc., the sample can be divided in smaller fractions and then concentrate each fraction individually and keep them in separate tubes. It may take time but precipitation can be avoided.

First I would put a drop of the precipitate solution on a glass slide and look at it under a microscope – is it crystalline or amorphous? If it is crystalline you may be able to spin out a highly purified sample of your protein and re-solubilize it. If it is not crystals, you can adjust the bottom lighting you can easily see precipitate as a cloudy drop and try pushing the pH, salt or other additives up or down by pulling a liquid thread connecting with drops of different solutions. In this way you can test with a drop if any changes will re-solubilize your sample. Once you know what may help, you could switch buffers quickly by running your sample down a de-salting column instead of dialyzing, which is slow.

The protein concentration is definitely high and can cause precipitation especially when the buffer composition is changed. This aspect is critical in this situation.

If one keeps in mind that 15-20 mg/ml solutions are routinely used getting crystals for X-ray crystallography, and crystals are initiated by buffer changes, this phenomenon is easily accepted.

As protein concentrations increases, packing of protein molecules will increase by the increase in the number of protein:protein interactions. In this case, in an aqueous environment, hydrophobic surfaces will pack and water will be excluded, and macromolecular structures form. The reduction in surface area - a big rock has a smaller surface area than several small pieces - changes the number of interactions with the medium, as the number of surfaces available for the surrounding medium to interact is reduced. This causes loss in energy from bonding, which needs to be compensated. Thus, critically depending on the medium composition this loss may be unfavorable, causing the protein to leave the solution as aggregates or crystals. Whether the protein will leave the solution now depends critically on the whether the medium is able to compensate for its lost interactions, with new stabler interactions - which then again depends on how the protein packs and what regions are exposed. This is where glycerol, salt and agents like this come in.

As John Tainer says above, you may even have crystals !!!!!

In many cases, free -SH groups are notorious for rapidly inducing aggregation by -S-S- bond formation. This can increase packing, and hence, precipitation. These aggregates can be prevented by using 20 mM DTT or more (yes a high concentration), to prevent -S-S- bond formation. Aggregates - sometimes- in this case can be broken by incubating the aggregates in the ORIGINAL buffer with DTT at 37C or even room temp. Then dialysis is carried out in presence of high DTT. Don't forget now DTT will be in all solutions you use the protein at high concentrations.

Unless you want to get crystals, keep the concentration below 5 mg/ml at least. This should be more than enough to store and use the protein in most experiments.

Good luck!!

In my experience all proteins will precipitate with increasing concentration at the pivot point concentration depending upon protein charge, salt concentration, buffer, and solvent. DNA binding proteins tend to prefer higher salt to stay in solution from those I've worked on. You are evidently using macromolecular crowding to stabilize your proteins and the precipitation is driven by denaturation I suspect rather than saturation. Crowding can stabilize the folded state as the folded protein uses less space than an unfolded polypeptide chain. You may want to find other conditions besides crowding to stabilize your proteins however,

Dear Stefan,

Actually Prem Subramaniam already explained all the issues concerning protein aggregation/crystalization. His impressive post taught me the the bases of my experience... John Tainer also agrees "all proteins will precipitate with increasing concentration...." My experience is that yeast THO complex is precipitating at concentration of approx. 5mg/ml (PMID:22314234). It seems you have big luck working with friendly proteins love neighbourhood...

I've never had the opportunity to work with proteins that can be concentrated to 15 mg/mL without precipitation, even by adding glycerol and high salt concentration to the protein samples (which indeed usually improves protein solubility). So congratulations Stefan if "all of your proteins behave better when concentrated", but I do not believe this is a general tendency for proteins.

From my experience, I've noticed that dialysis is the worst method for buffer exchange, in particular when the protein has an intrinsic propensity to precipitate. One could believe that a slow process like dialysis may avoid precipitation when changing the buffer in comparison with a fast process like dilution with the new buffer. This is actually not what I've observed with the proteins I've worked with. That's why I prefer to use centrifugal filters (with a suitable MW cutoff) for desalting or buffer exchange when working with "difficult" proteins. I agree Jan that such concentration systems can lead to precipitation if you concentrate too much the protein, but this can be easily controlled by choosing a suitable centrifugal filter and measuring protein concentration with a Nanodrop system (for instance) during the process. Even if unexpected, I've noticed that the dilution/concentration process lead to less precipitation than dialysis.

Evaluate how different protein concentrations influences the precipitation of the sample after storage. Generally it is better to store proteins more concentrated than dilute. Losses due to adsorption will be relatively less when the sample is at 10 mg/ml than at 0.5 mg/ml. However, if storing the protein too concentrated leads to precipitation, store the protein at a dilute concentration and then concentrate the sample immediately after thawing and right before crystallization experiments.

In some instances a protein will crystallize during a freeze thaw so be sure to check the precipitate under magnification for the presence of microcrystals.

Try warming a precipitate protein by holding or rubbing the tube in your warm hands or holding the tube under a stream of warm (approximately 37 degrees Celsius) tap water with gentle swirling. Be gentle. Do not shake or vortex. Avoid foaming

https://www.researchgate.net/publication/309041540_Important_Factors_Influencing_Protein_Crystallization

Article Important Factors Influencing Protein Crystallization

 Hi,  Ajit...

 I would like to know that have you succeeded in preventing protein aggregation during dialysis...

This is an old thread I'm replying to, but since Anamika seems to be interested I feel I should share my experiences with this...Sometimes during the protein purification stage, nickel can elute off the column into the elution buffer containing your protein sample.  Once you dialyze or desalt the imidazole, which was occupying the His-tag, residual nickel will bind to His-tagged protein and aggregate it out of solution.  This problem has been solved in two different ways:

Good luck!