Hi all,
I have a protein which, once purified (e.g., in its elution buffer), is very easy to aggregate, as manifested by the formation of flocculent mass floating in the solution, and a reduction of the soluble protein level. And its aggregation can be caused by even mild agitation/gentle shaking of it with resin, not to mention rolling with resin. Since rolling/regular agitation helps its capture by the resin (second step of a tandem affinity purification), I guess I cannot get around this?
Another fact is that this protein contains a very long (500 amino acid), continuous intrinsically disordered region, which may be related to this aggregation propensity?
So, I wondered, if there is any additives I can add to the solution to stabilise the protein? I have heard of high salt concentrations as a solution, but since I want to cleave the protein's tag and high salt can inhibit the cleavage, I try to avoid this approach. I have also heard of using mixture of glutamate & arginine (20mM?), and using Tween20, to stabilise protein. Have anyone tried similar approaches, especially on disordered proteins? But novel suggestions are alway welcome! Thanks very much!!!
I think it depends upon what protease you are using to cleave your tag. If you are using Sumo protease to cleave a Sumo tag, the protease still possesses cleavage activity in 20% (v/v) glycerol or 2 M Urea or 400 mM NaCl or 300 mM Imidizole. It is pretty hardy (see table 2).
I can get near complete Sumo tag cleavage at 4°C cold room for over 24 hours with Sumo protease with 2 mL of 10 mg/mL of my recombinant Sumo tagged protein in a buffer of 20 mM Tris-HCl (pH = 8), 400 mM NaCl, 100 mM Arginine, 10% (v/v) glycerol ... using 0.1 mg/mL Sumo protease.
Some other tag cleaving proteases might be more fragile. But sumo protease is hardy. I think 20% (v/v) glycerol might make a big difference in solubility. Tag specific proteases have specific pH requirements which I think is about 8. Make sure the pH of your buffers is correct.
If you have a hardy protease like sumo protease, cleavage will just take longer under suboptimal conditions. Do something else or take a nap in the meantime.
Adding 20% glycerol to increase the viscosity of the solution may help. Another thought is to add 0.5 M trehalose, which can help to stabilize protein structure.
Interesting question. Have you tested if a larger versus smaller air-water interface or higher protein concentration promote or reduce aggregation? To avoid aggregation, I like to try agents that stabilize folded states (Trimethylamine N-oxide (TMAO) an osmolyte that enhances protein stability), destabilize the unfolded state (sarcosine), or break the structure of water (potassium thiocyanate (KSCN)).
Adam B Shapiro Hi Adam, thanks for the information! I was thinking of similar approaches. I know that some people add 1M sorbitol (or high glycerol concentrations) to but only to their lysis buffers and remove this crowding reagent immediately afterwards. I was curious - if glycerol, sorbitol (any many other similar reagents) really help stabilise proteins then why don't we have them around all along?
I think your advice is worth trying. Thanks again!
John Tainer Hi John, thanks for the suggestions! No I haven't tested the area of air-water interface although it is likely to contribute to my observations. Primarily because this protein is too precious...
These reagents do sound new to me. Are they known to interfere with common enzymatic assays (e.g., protein phosphorylation, pulldown, ...)? Thanks!!
High concentrations of solutes like glycerol increase the viscosity of solutions, making chromatography for purification difficult due to low flow rate and high column pressure.
One common method of storing enzymes is in 50% glycerol at -20oC. The glycerol keeps the protein solution from freezing, which is convenient when you want to withdraw a portion repeatedly, and stabilizes the protein. The glycerol can be added after the purification is completed.
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