I have a major problem that I have been facing for last few months. I have a protein which is His-tagged at the C-terminus. Its a fusion of 5 monomers and its weight is around 42 KDa. The experimental setup is like this
- Protein expression:
- pET21b plasmid
- BL21 (DE3) bacterial strain
- 1 mM IPTG induction of protein production at OD600 = 0.6
- 4 hrs induction time
Protein purification:
- Cell lysis by sonication
- Ni-NTA affinity chromatogrphy
- Size Exclusion Chromatography (SEC)
The binding buffer that I'm using is 50 mM, 150 mM, 250 mM and 500 mM Imidazole + 50 mM Tris (pH 7.8) + 150 mM NaCl. It binds to the column in very very little amount and on observing on SDS PAGE it is been observed but with lot of impurities. I have tried it more than 6 times but no success I am getting. Anyone has any idea what might be the problem
Can we see the SDS-PAGE?
It would be particularly useful to see the pre-induction and post-induction, clarified lysate fractions to determine relative yield.
First off I would say that 50mM imidazole is pretty high for the binding buffer.
I would be using 10-20 mM imidazole in my binidng buffer, I generally avoid tris buffers with IMAC and use HEPES pH 7.5 or another suitable buffer if the pI of the protein is too close that pH.
Thanks for your comments. I have attached the SDS Page Images
Thanks
Mohit
Dear Mohit,
I don't know what volume you load on your SDS-PAGE analysis. Considering this volume is not high and considering that the band we observe around 45 kDa is the protein of interest, the protein expression doesn't seems bad to me. Since I don't know how many ml you have after SEC, the yield of 7 mg/ml doesn't mean anything to me.
As Stanley Ng suggested, the major part of the protein is eluted in washes. A revision of imidazole concentration will be a nice starting point.
Your SEC is quite promising as it suggests that contaminants can be separated (there aren't binding your protein of interest strongly). Because I'm sure you will improve the IMAC yield, an additional purification step using an ion exchange can be an option. Alternatively, you can play on salt concentration during IMAC to abrogate non-specific interactions (be careful to don't precipitate your protein of interest with high ionic strength).
Thank you for the SDS-PAGE, your protein is flowing through, I would hazard a guess of 80-90% loss.
Try again with less imidazole, ~20mM.
In fact if you have your flow through and wash dilute 3 fold and reapply to your regenerated imac resin, wash with 20mM imidazole.
I would use as little IMAC resin as possible, don't underload. I would allow overloading of target as IMAC responds well to your target protein competing off non-specific proteins. You may get some loss through this, but your prep will be much purer as a result.
If you haven't already done it then it would be worth while carrying out an anti His tag western blot to make sure the His tag isn't being clipped off. If this looks okay then your His tag may be buried and poorly accessible to the nickel resin. Ive had success by adding a low amount of urea (2-4M) or detergent to the lysis buffer before binding. It's another cloning step but swoping the tag to the n terminus is worth trying ( I had recent success doing this with a 220kDa protein expressed in insect cells). I often add a short linker region before my His tag to try and minimise the tag being buried. Another option you have of course is to denature (guanidine or urea) and then refold. . Again another cloning step is to use larger fusion tags e.g. his-sumo, His-thioredoxin are very good solubility/affinity tags. Best of luck! Eddie
Using 50 mM Imidizole in your binding buffer should result in all your protein going into the flow-through along with all the other impurities.
I recommend 5 mM Imidizole in your binding buffer.
The binding affinity of His-tagged proteins to Ni-NTA is usually never as great in practice as in theory.
I am looking at your images. I don't see the supernatant in the SDS PAGE. Did you apply the supernatant to see if any of your protein is in the supernatant. Are you applying the lysate or supernatant onto the column? This isn't clear to me. If your protein is in the supernatant then you could get rid of that big contaminant with near 95 kDa by applying the supernatant to the column, dodging that impurity mostly entirely.
I suppose it is possible that that big contaminant band is actually your protein of interest as a dimer at double the molecular weight. I wouldn't think a dimer would survive SDS, but do you boil your fractions in SDS at 100 centigrade in a hot-block for at least 10 minutes before loading the gel?
If that really is a dimer what if it is a cystine dimer that has a disulfide bond? SDS wouldn't break a disulfide bond, but boiling with Dithiothreitol (DTT) as well would reduce the disulfide bond and you shouldn't see that band anymore if I am right.
Dear Adron
I am applying lysate onto the column. I am not putting supernatent onto the column. Yes I do boil my fractions at 100 degree for atleast 10 minutes. My gene is without a cysteine.
That I am also trying to modify my sequence and put a point mutation to change Lysine to cysteine in my gene and change of histag to N teminal instead of C terminal
Another idea: Does your lysate contain any traces of a chelator like EDTA?
These tend to strip the nickel from the Ni-NTA beads.
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