Hi
I'm trying to purify an over-expressed membrane protein which I suspect is stuck in inclusion bodies. I extract the proteins using 8M Urea in a 50mM Tris-HCl (pH 7) buffer containing 20mM imidazole and 0.5M NaCl. I ultracentrifuge the sample before loading it onto the IMAC column. During loading, the column gets clogged quite soon. When I try to elute the protein at very low flow rates on FPLC (to prevent over pressure), the proteins come off very slowly (during most of the linear gradient from 20mM to 500mM Imidazole), especially my protein of interest.
What should I try to prevent the protein from precipitating/clogging onto the column?
Can something like a phosphate buffer possibly have a different effect vs a Tris buffer? Should I try a higher/lower pH? Less NaCl?
PS: pI of my protein is 8.5, truncated version of the protein (just the hydrophilic part) purifies perfectly as it's completely soluble.
Inclusion bodies apart from having our protein of interest, also houses other bacterial hydrophobic protein and large chunks of lipids, which is some how physically bound to the protein of interest. So, these lipids had to be dissolved and removed before solubilizing in 8 M urea.
First lyse the bacterial cell in normal Tris buffer. Centrifuge it and discard the supernatant. Now, wash the bacterial inclusion pellet with detergents like Triton-x100 (up to 1%), sodium deoxycholate (up to 1%) and sarkosyl (up to 1%). U can also try ading Tween 20. After every wash load the flow through to see if any loff of the target protein. After these washes, then solubilze the protein (pellet) in 8 M urea. After solubilizng, again centrifuge (12,000 rpm, 20 mins, 4 degrees) it and load the supernatant on the column (discard the pellet). Hope it will help u!
did you equilibrate the column with the 8M urea, in the buffer.? I would do this. If you didn't centrifuge at a high enough rate, there may also be precipitate. And if the column is at a different temperature than the solution that might be a problem. I would dilute out the protein as well if the solution is too viscous. But I have never had problems with solubilized inclusion bodies in the buffer you are using.
Are you filtering the sample before loading it into the column?
Also, take into account that if the protein has free cysteines, at that pH they may and will form intermolecular disulfide bonds. Try adding 1 mM TCEP to the buffer to reduce oxidized cisteines; it is compatible with Ni-NTA (it is expensive, though).
You are not omitting urea from the elution buffer, right?
Hope it helps!
Thanks for all the replies!
Bhanu P Jagilinki: I did do a single 10mM CHAPS wash of the pellet before solubilizing in urea. I think I'll add additional washing steps (with Triton and Tween) next time to see if that helps, thanks!
Marcia Moss: I did equilibrate the column and I ultracentrifuged at 100 000 x g for 1.5 hours before loading the sample. The sample was slightly viscous, so I'll try diluting it a bit further next time, thanks :)
Alejandro Martin: My protein has two cysteins, I guess there is a chance that it'll form disulfide bonds then? I don't have any TCEP unfortunately, but my column can handle up to 20mM BME, so I'll give that a try to see if it helps! And yes, my elution buffer contains 8M urea :) Thanks!
Sodium deoxycholate is more effective in dissolving and removing the lipid content. So try to include this one also before solubilizing in 8 M urea.
Thanks, I've managed to get hold of some sodium deoxycholate, will try to purify again on Friday if everything goes according to plan and will report back. I really hope it's just a lipid problem and not something else!
Try passing the sample through a 0.22 um filter. If it clogs the filter, it will definitely clog the column.
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