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Best protocol to solubilize bacterial fusion proteins?


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#1 radioactivestar88

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Posted 08 April 2016 - 02:48 AM

Hello everyone I need your advice for the solubilisation of a His tagged protein produced in E.coli.
Before performing the Ni-NTA column I would like to figure out how to extract most of the protein from the cells but I'm struggling because on each step of the purification I extract only small quantities.
Here's what I already tested: I usually begin with a sonication of my bacterial cell pellet in Tris 20mM. On the resulting pellet I tried to add Urea 8M or Gu-HCl followed by 4 different mild detergents at 0,5% (Nonidet P40, Tween20, Deoxycholate, or CHAPS).
I put each step of this protocol on SDS-PAGE but the band corresponding to my protein remains always thin line of the same size and intensity. I also added some SDS 0,1% on the remaining pellet of all the samples to see if there was still some protein: here I saw that some protocols worked slightly better than others but there are not great differences.

Do you have any suggestions to help me?
Thank you!



#2 Missle

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Posted 08 April 2016 - 05:04 AM

When you process the total cell protein pellet, do you see strong expression?



#3 radioactivestar88

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Posted 08 April 2016 - 05:50 AM

It is not strong, but I think it can be a bias related to the problem of extraction. I followed the production over the 4 hours, and I can see a band showing up after 2h and getting a bit more intense in 3h and 4h, but it is not a huge band it can easily be mistaken for a "basal" protein of the bacteria (I don't know if it is the right word, sorry).



#4 labtastic

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Posted 13 April 2016 - 12:37 PM

There isn't any sense in troubleshooting anything until you know for certain your protein is expressing.

 

Run an expression culture and a non-expressing culture side by side. Remove some cells, normalize to OD, sonicate briefly, add SDS to a final concentration of 2% from a 10% stock. Add a few microliters to SDS sample buffer, boil and run on gel. Stain with coomassie to see band with correct molecular weight in expression culture that is not present in non-expressing culture. If you see nothing, consider a western-blot probing for his6 tag. 

 

Assuming you have expression, then determine where your protein is fractionating upon lysis. It will be either (a) soluble, ( b ) in membranes, or ( c ) in inclusion bodies. Soluble protein should remain in the supernatant after cell lysis by sonication and centrifugation. Membrane and inclusion body proteins will be in pellet. To distinguish between membrane and inclusion bodies, solubilize in a non-denaturing detergent e.g. triton. Membrane protein will solubilize while inclusion body protein will not. Inclusion body protein will solubilize in 8M urea or 6M guanidine, as well as denaturing detergents (i.e. SDS).

 

Also lyse your protein in buffer with salt (e.g. 200-500 mM NaCl). It is possible your protein is precipitating upon lysis because you have no salt in there.


Edited by labtastic, 13 April 2016 - 12:39 PM.


#5 radioactivestar88

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Posted 18 April 2016 - 03:44 AM

That's what I did. I see a band corresponding to my protein, it is expressed but I'm solubilising a small amount on each of the steps you described (actually I tried this sequence: 1) sonication in lisis buffer 2) urea + non denaturing detergent on the sonication pellet). The quantity I obtain both in the soluble fraction and the membrane+inclusion bodies fraction is comparable. So my question is: how can I solubilise "at once" a good amount of protein?



#6 mdfenko

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Posted 18 April 2016 - 06:46 AM

there are handbooks available at the ge lifesciences website which can give you information on purifying recombinant proteins (amongst other, useful, handbooks):

 

 

http://www.gelifesci...port/handbooks/


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