I purified a lipase by HiTrap FPLC and dialyzed the collected samples overnight in a solution of Na2HPO4 (50mM), NaCl (200mM), EDTA (50mM), DTT (2mM), pH 8. Since I got a high volume of sample (80 mL), I would like to concentrate my lipase to 12 mL using an Amicon 30kDa for my 2nd purification (size exclusion FPLC). When I was concentrating the protein, I saw that the protein start precipitating. Do you know what reagent I could use to prevent the precipitation of protein at high concentration ?
Thank you,
Hang
There are several possible reasons for this observation:
1) The isoelectric point of your protein might be about pH 8.0, therefore when you concentrate your protein at this pH it starts to aggregate and precipitate. All proteins are prone to aggregation and precipitation in the buffers with pH close to their isoelectric points. Try to change the pH of your buffer for dialysis 1.0 pH unit up or down (pH 7.0 or 9.0).
2) Since your dialysis buffer had DTT (2 mM) I guess your protein has some disulphide bonds. DTT is very unstable at pH 8.0 and over the time of dialysis and further steps it might be inactivated. After that, the formation of the disulphide bonds in your protein starts in random and possibly not natural way. That might cause the formation of different disulphide oligomers of your protein which could be insoluble. After dialysis, try to add freshly prepared 2 mM DTT to your protein solution, incubate 1 h, and then make Amicon concentration.
3) 50 mM EDTA is a little excessive concentration for inhibition of possible metal dependent proteases (usually 5 mM is enough). Some proteins are stable (soluble) at high concentrations only when they have their natural metal ligand bound as a part of their structure. When you strip your protein from that metal with EDTA at high protein concentration, they might aggregate and precipitate. Check if your protein is a metal binding protein, and how it might affect its properties.
There are several possible reasons for this observation:
1) The isoelectric point of your protein might be about pH 8.0, therefore when you concentrate your protein at this pH it starts to aggregate and precipitate. All proteins are prone to aggregation and precipitation in the buffers with pH close to their isoelectric points. Try to change the pH of your buffer for dialysis 1.0 pH unit up or down (pH 7.0 or 9.0).
2) Since your dialysis buffer had DTT (2 mM) I guess your protein has some disulphide bonds. DTT is very unstable at pH 8.0 and over the time of dialysis and further steps it might be inactivated. After that, the formation of the disulphide bonds in your protein starts in random and possibly not natural way. That might cause the formation of different disulphide oligomers of your protein which could be insoluble. After dialysis, try to add freshly prepared 2 mM DTT to your protein solution, incubate 1 h, and then make Amicon concentration.
3) 50 mM EDTA is a little excessive concentration for inhibition of possible metal dependent proteases (usually 5 mM is enough). Some proteins are stable (soluble) at high concentrations only when they have their natural metal ligand bound as a part of their structure. When you strip your protein from that metal with EDTA at high protein concentration, they might aggregate and precipitate. Check if your protein is a metal binding protein, and how it might affect its properties.
Additionally, you might find that the solubility of the protein is affected by the ionic strength (higher or lower) and/or detergent. Since the protein is a lipase, it seems reasonable to include a detergent to occupy any exposed hydrophobic surfaces. I would recommend a detergent with small micelles that won't cause trouble with gel filtration chromatography, such as CHAPS.
Hi there,
Tee following paper, about the addition of glutamate and arginine to improve protein solubility and stability, might be interesting:
http://wolfson.huji.ac.il/purification/PDF/Literature/Golovanov2004.pdf
Agree with Adam about detergents. Our proteins are universally less than happy and hate being concentrated. For that reason we try to avoid concentrating at all and just try to purify with sufficiently high concentrations to maintain the same concentrations we need throughout our subsequent treatments. Our protein was really helped by adding a MBP tag for solubility. That was the best tag and the only other tag that made any difference was GST (also large) but MBP was still pretty dramatically better. Tags can always be removed immediately prior to your experiment (we like the precision protease). Detergents definitely help a lot but are incompatible with some of our protocols. You should note that as Adam alluded too, large micelles like NP40 will concentrate just like your protein in the concentrator so your detergent concentration will go up too. Our proteins are called "amyloid-like" so they love to aggregate which we've taken the unorthodox approach of purifying in 1 M urea, which is low enough to not denature the protein to any extent we can measure and makes it much easier to work with at high concentrations. Despite all this, we cannot concentrate to get the high level of concentrations that we've achieved by just optimizing our initial purification. So by starting off with a concentration that is orders of magnitude higher than we need (largely by optimizing expression and solubility during lysis) for any of our experiments, we have no worries that subsequent steps will dilute our samples too much for our needs.
Some time your sample may not like salt at higher concentration. Dialyze against water and then concentrate it. I was lucky to concentrate some antibodies to 200 mg/mL by this way
I agree with Viktor. If you concentrate your protein, a positive second virial coefficient might be pretend protein aggregation/precipitation. You can reach it by lowering the pH down to 6.0 or 5.5(depends on the isoelectric point), or by addition of stabilizing salts like Arginine HCl. A different approach could be the addition of non-ionic surfactants like polysorbat 20/80 in low concentrations to prevent shear stress during the centrifugation.
All fine suggestions above and some more in order of preference:
1. Minimise the protein concentration when possible, split your sample and do 2 column runs.
2. Ensure a strong net repulsive regime (more than 1 pH unit below pI), or positive second virial coefficient (B22) or interaction parameter (kD). (Alexander Klaks suggestion).
3. After point 2. reduce the salt content (ionic strength) to 150 or 100 mM as this shields ionic protein-protein interactions, which can be good in net attractive conditions but reduces repulsion in repulsive conditions.
The bottom line is it is currently difficult to predict the right conditions, so it will require some empirical testing based upon positive past experience if you really have to go to high concentrations. As someone who works with high concentration proteins, try low concentrations if possible!
All the comments above contain very useful suggestion (except dialysis against water...) on how to tweak buffer composition to prevent aggregation during concentration in a spin device. You did not specify what kind of device you are using, but some of the below may be beneficial:
1. spin your device slower than the recommended rpm, stopping the centrifuge frequently and gently mixing the solution inside the device to prevent very high concentrations forming in proximity of the membrane
2. the better solution is to use a stirred cell or a TFF system to prevent any concentration gradient at the membrane.
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