I am trying to purify one of my protein of interest having molecular weight around 52 Kda. Upon over expression it comes in inclusion body. So I isolated and washed inclusion body and then dissolved in lysis buffer having 20mM sodium phosphate, 6M gdn-HCL, 500mM NaCl. Then performed Ni-NTA AFFInity chromatography. The binding is performed using similar buffer except used urea and pH was 8.0 washing is done with pH 6.0 and pH 5.0 buffer while elution is done in buffer having pH 4.0. the problem is elution is happening but it consists of a lot of contaminant proteins in form of fine bands as visualized on SDS-PAGE.
ANY SUGGESTION to remove these contaminant bands. I don't wanna another chromatography step. Optimization and modification in protocol is most welcome.
Lanes are marker, inclusion body, fow through, binding buffer, pH6.0 wash pH 5.0 wash pH 4.0 wash.(elute1, 2, 3, 4and so on ).
Hello Vishal,
I would recomend you to perform a gel filtration in a Sephadex 200, for example. It’s very simple and helps you to separate fractions easily. But once you don’t wanna additional chromatography steps, you should try a step by step elution, using a continuous and crescent gradient of imidazole. Are you performing you chromatography in an automated purifier (Akta, for example)? For this protocol you’ll need this equipment. Try the following parameters: Gradient on --> length final volume 20mL --> buffer B (Elution buffer imidazole 500mM) 100% --> fractions collected volume 0,5mL. Thereby, each protein (yours and contaminants) tend to dissociate from the column with different imidazole concentrations. Analyzing the chromatogram and it’s peaks you can monitor fractions elution and get those who have your protein.
Are you perfoming your elution step using urea or gdn-HCl?
I hope it helps!
Bests,
Thanks Dr. De Leo. i am doing it in manual column i don't have access to machines. The elution is performed in presence of Urea.
Hi,
It looks to me like your protein elutes already in your washing steps (pH 6 including). To start with, prolong washing with the binding buffer at pH 8 and eventually add another step with pH7 wash. Then elute at a lower pH if this is preferred over imidazole steps.
My other recommendation is to check the loading buffer and/or electrophoresis buffer in the next SDS-PAGE runs. Your 'contaminants' seem to be present in just about all lanes with about the same concentration, which is weird. This could be a sign of impurities in the running buffer or in the gel loading buffer - eventually prepare fresh and compare with the old one.
Good luck,
Marko
Dear Vishal,
You could try adding 20 mM imidazole to your lysis and initial binding buffers. As an addendum to Thais Canassa De Leo's answer: If you don't have equipment like an Äkta, then you can make a step gradient readily yourself. I used to wash my columns with 20 mM --> 60 mM --> and 100 mM imidazole. You could then elute with between 200 and 500 mM imidazole.
I don't use this method anymore because I do multiple purification steps, including cutting the His tag off of the protein. Using the above method, you will likely lose some of your target protein in the 60--100 mM imidazole washes.
Also, as Marko noted, your protein already seems to elute at pH 6. Given that the band is much more intense than in your actual elution, it is possible that the protein at pH 6 is more pure. If you ran a dilution series of the pH 6 wash, and found a loading volume that has the same intensity as the elution, then you could directly compare the intensities of the contaminating bands. Often, we try to eliminate contaminating bands and in the end if our purification schemes just lower the concentration of all proteins present, we can make it look as if the protein is more pure, even though little has changed.
Best regards,
Jordan
@marco thanks. at present i am following the same and will update the results as we will get. We would also consider to change the loading and running buffer as per your suggestion.
@jordan surely if pH things does not work i would try to go for imidazole based washing and elution as per your suggestion.
You can also use salting-out precipitation to purify your proteins from the cell lysates. 0.8 M- 1.0 M sodium sulphate is good for purification of over-expressed proteins via selective salting-out precipitation.
Good luck!
@Alemu protein is coming in inclusion bodies is salting out works with urea?
Selective salting-out precipitation may not work in the presence of urea as the solubility of the protein will be very high in the presence of urea. Any ways, it is good to try in small scale. I did not try selective salting-out precipitation in the presence of urea and detergents.
Nickel affinity chromatography exploits the fact that protein motifs with several histidine residues are bound to each other (the so-called His6 tag) on materials with nickel chelates.
For the isolation of these fusion proteins, e.g. Sepharose1) containing Ni2 + bound to nitrilotriacetic acid (NTA) was used. Ni2 + can interact with two His residues of the protein in exchange for water. In most cases, this process is eluted with an imidazole gradient. The protein can also be eluted from the column by means of a change in the pH value, but this can sometimes lead to denaturation of the protein in the case of pH-sensitive proteins.
I am always recommending the use of RP-HPLC with gradient elution (please see file; Lysozyme by RP-HPLC).
I have experienced many miserable results using the Ni-NTA affinity chromatography during an Arbeit period at The University of Tokyo (Yayoi-machi, Bunkyo-ku, Tokyo, Japan) from 2009 to 2013. Ni-NTA affinity gel seems to be an unexpectedly hydrophobic material.
Further, the polyacrylamide gel (PAG) is also a very hydrophobic material (please see file; IEF for hydrophobic protein).
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