protein precipitates:HELP !

Shane Albright insomnia at
Sun Oct 16 21:37:01 EST 1994

In article <37j39o$ss6 at>, Clemens Suter-Crazzolara
<un691cs at> wrote:

. . . <STUFF DELETED> . . .

> Inprincipal, I don't think it is the histidine tag that is causing the
> problems. I am also testing the pH, currently at 8.9, will test the
> more acid buffers also. Triton, or tween is apparently not so good:
> CHAPS seems to be the detergent of choice. I added 10 mM to my buffer,
> at different pHs it gets quite turbid: and my protein still precipitates.
> On the other hand, I heard from a colleague that 1-2 M urea will keep
> most proteins in solution, and they remain usually biologically active
> (that is probably what you want too, isn't it ?). So this I will test
> also: to first concentrate the protein, add urea to 6 M and slowly dialyse
> down to 0 urea and check when it percipitates.
> I have found no references in the literature yet about this problem. Perhaps
> some other readers out there could help us ?
> lots of luck,
> clemens, heidelberg


I don't think that there really is a general solution to this problem, and
it may be that your protein simply cannot be made soluble when expressed
in E. coli.  I'm assuming that the protein is in inclusion bodies to begin
with, since you purify everything under denaturing conditions, and one
approach may be to change media pH, induction conditions, etc. to get more
soluble protein, then purify under native conditions.  Renaturation is
tricky, unfortunately, and often just leads to inactive aggregates.

That being said, I can tell you my recent experiences at trying to
renature a His-tagged protein from 6M guandine HCl.  You might find it
applicable to your situation.  My protein is about 50 kd with the His tag,
and is almost entirely in inclusion bodies.  I tried a bunch of approaches
for the renaturation following the Ni column, and the common denominator
was that my protein aggregated, and after a few days fell out of solution
(at 4 deg C).  0.1% NP-40 prevented a large part of the aggregation (but
not all of it) and kept the protein from precipitating.  This protein
binds DNA, and gel shifts indicated that only a very small fraction of my
renatured stuff was active.  I did some research on protein refolding and
discovered two important points:

   1) Speed is important, since the folding is reversible, but aggregation
is not.  You want to make the transition from denatured to folded as quick
as possible.  Otherwise, aggregates are favored.  However, non-denaturing
amts of Gdn HCl (<=1M) or urea (<=2M) can help prevent mis-folding.

   2) Renaturation at >100ug/ml is probably doomed to failure.  Aggregates
form much too easily.  Concentration should be done after you get active,
renatured protein.

Following these guidelines, I designed a protocol that appears to work
well. I bind protein to the Ni-beads (Qiagen's Ni-NTA agarose) in 6M
guanidine, then dilute them (yes, the beads with protein attached)
200-fold into rapidly stirring buffer containing 1M guanidine and allow
stirring to continue at 4 deg for >24 hours.  Then I can recover the beads
by centrifugation, resuspend in a small vol of buffer, pour a column, and
elute with 0.5M imidazole/1M guanidine.  I then dilute to lower the
guanidine to 0.1M and use a DNA-affinity column to concentrate active
protein and to get rid of all the guanidine.  

About 50% of the protein never comes off the beads, and about 20-30% of
what does is active.  So 10-15% final yield with nice specific activity.

Hope this helps.  Feel free to email me if you have any questions, or if
you have any other input into this discussion.

Shane Albright
insomnia at

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