My His-tagged purified protein contains 10mM trisCl, 300mM NaCl and 200mM imidazole at which the protein was eluted at pH 8.0. What should be ideal dialysis buffer composition to remove imidazole?
I am using 10mM tris and 300mM NaCl at pH 8.0.
Does NaCl affect protein function? Do I need to remove it also?
As Dominique already mentioned, the choice of buffer composition depends on many factors that vary from protein to protein.
- 10-20 mM buffer (TRIS, HEPES, etc...) is generally sufficient to buffer the protein solution. Choose a pH that's better for your protein than the pH needed for Ni-NTA. The buffers for Ni-NTA require pH > 7.6 or the His-tags won't bind efficiently to the column/beads. Your protein, however, might be happier at a different pH, so after Ni-NTA choose a pH that is at least 1 point away from your protein's pI (though 1.5 to 2 points is preferable) to make sure your protein is properly charged. Choose to go either up or down the pH scale towards the region closest to pH = 7 to 8 rather than towards the extreme ends of the pH scale (e.g. with pI = 8 go towards pH 6 to 7 rather than pH 9 to 10).
- 100-150 mM NaCl is usually OK, though some protein are not "happy" with less than 200-300 mM NaCl, particularly those with pI's over 9.
- If your protein has several cysteins and can potentially form disulphide bridges by itself, then change the BME from the Ni-NTA buffer to DTT at this stage (1 mM is generally enough). BME is a weak reducing agent and will stop working after a day or two, whereas DTT will continue to work at least twice as long. DTT can't be used during Ni-NTA as it will reduce the nickel ions much more quickly than BME. It's a good idea to use BME in Ni-NTA to stop contaminating proteins from aggregating or binding to your protein of interest via disulphide bridges, even if your protein doesn't have any disulphide bridges. However, you won't need BME or DTT after Ni-NTA if your protein doesn't have any cysteins.
Hi there,
You have to define the composition of the buffer so that your protein is happy! There is no general rule : the pH must be in accordance with protein activity and stability and nature and salt concentration adapted too... By default for a "normal" soluble protein, I use 20mM Tris pH 7.5 150mM NaCl and 1mM BME...
Thank you Dominique. I will remove the extra NaCl salts too by using 75mM NaCl strength buffer at pH 8.0.
As Dominique already mentioned, the choice of buffer composition depends on many factors that vary from protein to protein.
- 10-20 mM buffer (TRIS, HEPES, etc...) is generally sufficient to buffer the protein solution. Choose a pH that's better for your protein than the pH needed for Ni-NTA. The buffers for Ni-NTA require pH > 7.6 or the His-tags won't bind efficiently to the column/beads. Your protein, however, might be happier at a different pH, so after Ni-NTA choose a pH that is at least 1 point away from your protein's pI (though 1.5 to 2 points is preferable) to make sure your protein is properly charged. Choose to go either up or down the pH scale towards the region closest to pH = 7 to 8 rather than towards the extreme ends of the pH scale (e.g. with pI = 8 go towards pH 6 to 7 rather than pH 9 to 10).
- 100-150 mM NaCl is usually OK, though some protein are not "happy" with less than 200-300 mM NaCl, particularly those with pI's over 9.
- If your protein has several cysteins and can potentially form disulphide bridges by itself, then change the BME from the Ni-NTA buffer to DTT at this stage (1 mM is generally enough). BME is a weak reducing agent and will stop working after a day or two, whereas DTT will continue to work at least twice as long. DTT can't be used during Ni-NTA as it will reduce the nickel ions much more quickly than BME. It's a good idea to use BME in Ni-NTA to stop contaminating proteins from aggregating or binding to your protein of interest via disulphide bridges, even if your protein doesn't have any disulphide bridges. However, you won't need BME or DTT after Ni-NTA if your protein doesn't have any cysteins.
Hi Gaurav,
Your buffer should be something with minimal additives that keeps your protein functional in solution. Some very good suggestions have been made above.
Your final conditions will depend on what studies you want to do with your protein. If you want to do some structural studies by x-ray crystallography or solution nmr spectroscopy, then you would need high concentration of your protein in ~0.5 mM range. Some other studies such as binding or functional assays do not need such high concentration. So, the studies you aim to do will also dictate your buffer composition.
Thank you Antonio and Soumya for your kind attention on the question.
I am getting slightly white solution of my protein after dialysis. Although, It's showing good activity against the substrate. Why I am getting white solution? My protein sample was quite clear before dialysis.
The white, snowy particles are microscopic aggregates of denatured protein. This means your protein is less "happy" in the new buffer than in the old one. Many proteins stay in solution more easily under high ionic conditions (high salt), particularly those that are highly charged (pI >9 or
Thanks Antonio. I really appreciate your answer. Yes, I did centrifugation and I have lost some of my protein in the pellet. I wish I could put them back in solution!
Hi I have 1 cysteine in my protein; do I need to add BME? How do I check for PI value
You can check your protein isoelectric point value here
https://web.expasy.org/protparam/
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