Finding the right buffer to keep a protein in solution and obtain optimal activity is definitely a trial-and-process. First, I would not assume that you are far enough away from the pI is ensure solubility. Second, you should try a number of standard buffers such as sodium phosphate, HEPES, and many others that can be found in the literature and in classical works such as Methods in Enzymology. Third, don't forget that ionic strength can be critical. Many proteins like a "salty" buffer, such as 100-500 mM NaCl. Fourth, your buffer exchange may have removed a critical metal ion that is essential to maintain structure. You are fortunate in being able to measure activity. Use that to determine which combination of buffer component and salts give the best activity and optimal solubility. Good Luck!

Finding the right buffer to keep a protein in solution and obtain optimal activity is definitely a trial-and-process. First, I would not assume that you are far enough away from the pI is ensure solubility. Second, you should try a number of standard buffers such as sodium phosphate, HEPES, and many others that can be found in the literature and in classical works such as Methods in Enzymology. Third, don't forget that ionic strength can be critical. Many proteins like a "salty" buffer, such as 100-500 mM NaCl. Fourth, your buffer exchange may have removed a critical metal ion that is essential to maintain structure. You are fortunate in being able to measure activity. Use that to determine which combination of buffer component and salts give the best activity and optimal solubility. Good Luck!

Inorder to be better informed it would be helpful to address the following issues. You have not indicated the purity of the preperation. Even though the activity of the preperation is appropriate then does the precipatate represent an impurity? What does gel electrophoresis indicate regarding your preperation before and after precipation? What is the protein concentration after concentratiing? The pH appears OK, increasing salt concentration may be advantageous.

It would be a trial & error method. HEPES may be a better choice. You are only one pH unit away from the PI (where the solubility is minimum). Try somewhere around pH 6.0 if your protein shows activity. What is the best pH for your protein activity? Is it a cytosolic protein? As Ben & James have pointed out, try increasing the ionic strength of the buffer.

Thanks for the suggestions, very insightful. My elution buffer contains 0.5M NaCl, but as I mentioned I'm exchanging into PBS (~140mM NaCl) so perhaps the suggestions to increase ionic strength would improve the solubility. After buffer exchange, the fractions are clear with no precipitate. It's only after concentrating that I notice the cloudiness and particulates forming.

The purity of the Ni-NTA purified product is very high (see attached gel, lanes 2-5. lanes 3 and 5 are the reduced version of the protein). The concentration when the solution is resuspended is ~1.4 mg/mL, when it's spun down it's about 0.7 mg/mL.

What would your recommendations be on pH, how far away from the pI should I go, and which direction (acidic or basic)? Obviously the protein has to be active in whichever direction I choose, but is there a general method to selecting these parameters? I would like to avoid wasting a lot of protein trying to determine which buffers and pHs to try, since this protein is low-expressing. The protein is an antibody fragment, which is secreted into the supernatant, harvested and purified.

You always want to stay consistently on one side of the PI to avoid even briefly going through the PI. So if you are PI is 8.4 and you've been working at 7.3, if you change the change the pH, you need to go lower.

However, I don't think the pH is the problem. You are already one unit away and the protein is stable in the His elution buffer (I am assuming 50 mM phosphate, pH 7, 500 mM NaCl, 150 mM Imidazole).

therefore either:

1. Your protein likes high salt (probable)

2. You need to lower the pH to 7 or below (less probable)

3. Your protein is stabilized by high concentrations of imidazole (possible but unlikely)

If your protein is performing well even under turbid conditions, spinning the undissolved part is not a bad idea. Changing the composition of buffer may affect the activity of your protein hence the entire protocol setup. Just give a spin at 3500g and measure the activity. If it loses your protein activity then surely you had to go for hit-trial buffer compositions. Good luck!

I've already met the problem when switching to phosphate based buffers, which isn't so much surprising as phosphate is a rather chaotropic buffer. Why don't you try Tris, which is less chaotropic than phosphate, or even Hepes, which is rather kosmotropic, and all got about the same buffering range...

Second, if you have acces to thermofluor technology (Differential Scanning Fluorimetry), this would be a good way to test in which buffer+salt condition your proteins is the most stable.

Third, apparently you've got the chance to be able to perform activity assays on your protein, so the best thing to do would likely be to test several conditions with this assay (buffer, salts, even pH...).

Fourth, addition of additives can lower your problem, like up to 10 % glycerol or up to 0,5% Triton or Tween.

A last point is the protein concentration : even if 2mg/mL isn't such a high concentration, remember that some proteins really don't like to be concentrated...

One thing to note on pI is the the one we get from the software is theoretical pI and not the experimental determined so its safe to move more than 1 unit away from pI that just 1 unit.

Some protein can not not be concentrated more due to lack of ideal buffer constituents which could be anything from EDTA, DTT, detergent, or glycerol in addition to buffer and salt. So, don't force the protein to concentrate and loose it.

I would avoid doing experiments with cloudy protein though you have got results because I dont know what is that cloudy material?

DSF is good method to see what is the best buffer based on the melting point.

Thank you all for the valuable suggestions. Seems I have a stubborn protein on my hands.

@Jeffrey, the elution buffer is 50mM tris, 500mM NaCl, and 250mM imidazole. It might be a good idea to just buffer exchange the protein into the same buffer used for elution, but without imidazole present. And maybe lower the pH slightly to closer to 7 or 6.5.

@M. Irfan, the protein is used at very low concentrations for assays such as competitive ELISA and flow cytometry so there is no cloudiness in the samples because they are diluted. I did try centrifuging the samples, but the concentration of the soluble portion is much lower. Do you have any ideas on how to resolubilize the precipitated pellet? I tried several buffers, (PBS, 0.1M phosphate + 0.5M NaCl, and also 0.1M phosphate + 0.5M NaCl + 0.25M imidazole) but didn't have much luck with these.

@Yann, good idea to leave the protein in tris buffer (which is what it is eluted in). I'd like to remove the imidazole though. The protein is stable after elution in tris, but I haven't tried to concentrate it in Tris yet.

@S D the assays were done with low concentrations, so it wasn't cloudy. Adding glycerol to the buffer could be useful as well, especially for long term storage.

I will try to produce more protein and attempt several of the suggestions. Perhaps I can't concentrate the protein at all, which would not be the most favorable way forward.

Not sure if you have figured out a good assay or not yet, but one fairly helpful method for me was to get a hold of some hanging drop crystallography plates. You can go through a whole bunch of buffers in one go by spotting your protein on a cover slip with the test buffer (ex. 1-to-1 ratio) and then place that cover slip over a vacuum greased well and reservoir buffer. Its pretty much the opposite concept of screening for crystals.... I know some NMR guys do it for buffer optimization too. Some times close calls can be sorted out by stressing the proteins in a higher temp incubator.

I would advise you to split your sample and exchange "aliquots" with various buffers. Wait some time under controlled temperature (4 degree) and centrifuge using a table top centrifuge at max speed for some 10-30 min. Do a protein quantification on the supernatant. Things like this can guide you where to go.

Clearly imidazole is something that often (not always) makes protein aggregate. Get rid of it asap. Do you have a membrane protein? We often also use 5-10% glycerol in our buffers (membrane proteins).

Good luck

To add further complications to this issue of buffer exchange, I need to perform a labeling experiment in 50mM sodium borate buffer, pH 8.5-8.7. Considering my protein has a pI of 8.4, is it possible that I could dialyze the protein directly from the ni-nta elution buffer (50mM tris, 0.5M NaCl, 400mM imidazole) into the borate buffer without precipitation? I plan to keep the salt-content high in the borate buffer (0.5M NaCl) to help with solubility, but are my expectations of keeping the protein from precipitating unreasonable?

Susana, your suggestions to work with small aliquots are a good idea, especially in this case.

Just wanted to give everyone an update to this, as the more I've worked with it, the more I've learned. May also be helpful to anyone having the same struggles I was.

Adding 0.1M EDTA to the dialysis buffer really helped to prevent precipitation during buffer exchange for me. It creates an extra dialysis step, but it's worth it. Also I think the EDTA works in some manner to chelate some of the leached nickel that has coordinated to the 6His tag. However it works, it did the job for me. I still get some precipitation during concentrating with a centrifugal filter, but I'm not losing nearly as much protein as when I was having problems with precipitation during dialysis.

Thanks again for all the suggestions!

I have also faced the same problem, you can make purification buffer in PBS by adding 20mM imidazole and other required component like beta ME, Glycerol, etc. One thing you should keep in mind that the pH and Glycerol component of purification and dialysis buffer shouldn't vary too much as it leads to precipitation. And if it has to vary, try dialysis with a slow gradient type of exchange rather than quick steep change.

I also found that with this particular antibody fragment, switching to HEPES buffer instead of PBS really helped with long term storage. I noticed a lot of precipitation after thawing aliquots in PBS that were snap frozen in liquid nitrogen, then stored at -80C long term. When I switched the dialysis/storage buffers to HEPES instead (and sometimes added 10% glycerol), the precipitation was almost completely eliminated when the aliquots were thawed. Food for thought, if anyone else is having this issue.

If your protein binds to membrane, the Ni column purified protein can get cloudy.  You can raise the salt concentration, which will help the to solubilize the hydrophobic proteins. The cloudy stuff may also be due to lipid.  In that case, you can just remove the precipitate by centrifugation.

here is one peculiar problem I am facing. more or less same precipitation problem. I am trying purification of a protein with pI 5.9. I am getting good yield of of it however, when I take it for concentrating it starts forming insoluble precipitation. I am using Amicon filters and even the low concentrated protein solution is forming precipitation. precipitation is not as usual cloudy material, it looks as if a tissue paper is dissolved in water which is insoluble in any buffer. I won't see this precipitation during dialysis or in any other steps. Buffer which I am using is 50mM tris (tried pH 7.0, 7.5, 8.0), 500mM NaCl, 5mM beta ME, 10% glycerol, 0.1mM PMSF. I have tried Hepes also but no improvement. For changing the buffer I am using Desaltng column. I tried adding Triton X 100 during lysis and affinity purification protocol there also no success. I have not tried acidic buffers.

Please suggest what to look for :(

@Manjulu Try lowering the salt concentration. Higher salt concentrations will favor the interaction of hydrophobic surfaces, which may promote the formation of aggregates. Aggregation varies with salt concentration, with some proteins liking high salt and some liking low salt. However, 500 mM NaCl is at the high end so I would experiment with lower NaCl concentrations.

run the secondary structure of your protein then try to make the outer surface of your protein zero charge

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

Article Important Factors Influencing Protein Crystallization

I am using tris buffer(pH 7) for the purification of my protein whose pI is 8.I get a large band at the top of the well which couldnt be separated using AKTA system.I have tried adding 10 percent glycerol.looks like my protein is forming aggregates in solution but i cannot see the precipitates. should i use Hepes buffer for its purification or which additives are to be recommended.

Anam dilute your protein conc, Increase voltage, check with fresh running buffer, denature for 5-10mints, change the gel percentage, one of them should work for SDS, for AKTA (pH < pI or at a pH > pI ), you could use ion-exchange chromatography (cation- or anion- exchange chromatography, respectively) to purify proteins that are charged and bind to cation- (e.g. CM-cellulose) or anion (e.g. Q-speharose) exchange columns.

Buffer: try with high salt conc=200-300mM NaCl or KCl,

or try with HEPES or Phosphate buffer.

best of luck

Najeeb Ullah thankyou for the suggestions.I will try them now

Phosphate buffer is a poor buffer of choice for preservation due to the different rates of crystallization of the components of phosphate buffer causing pH Shift.(which likely contributed to the aggregation) HEPES/Histidine/Citrate buffers are better.

Also, though scientists do use Tween-20/80 for preservation, they are notoriously hard to remove in subsequent filtration. Use of 10% DMSO or sucrose to stabilize is a better approach as they can be far easily buffer exchanged.

Manjula, even thoguh this is quite a few years late in response your high salt conc. could be affecting the precipitation and formation of visibile fibrils. Addition of 100 mM Arginine can help in solublizing the aggregates if the high salt conc. is otherwise required.

Article Effect of Initial Buffer Composition on pH Changes During Fa...