I'm trying to purify a protein using HIS-select Ni-affinity column (washing with 25mM Tris pH7, 0.3M NaCl, 10mM imidazole and elution with the same buffer + 250mM imidazole). The protein (pI= 8.3) becomes cloudy/precipitated within 4 h of dialysis into Tris buffer + 0.1M NaCl. (I dialyze immediately after elution, and ptn conc reaches 4 mg/ml) The amount of ppt is less and I remove the supernatant, but the protein still precipitates on storage and gradually loses activity. The protein forms a little ppt at any pH (5-7.5), so it possibly could be the imidazole that the protein is not happy with. Can anyone suggest practical ways to elute the protein without imidazole? (I've read about using histidine/low pH etc, but couldnt find actual protocols where these methods have been used).
Many of the proteins I work with start to precipitate quickly after nickel affinity chromatography. Buffers for nickel affinity chromatography generally have 300-500 mM NaCl in them, but afterwards people dialyse into buffers with 100-200 mM NaCl while removing the imidazole. I have established that in my case it is not the loss of imidazole that has a negative effect but rather that my proteins dislike buffers with low concentrations of NaCl. Give it a try, maybe keeping the NaCl concentration higher than 100 mM might help you. Also, freezing your protein straight after purification will stop it from precipitating and help keep its activity for years ( ... if your protein can be frozen).
His-tagged proteins can aggregate via their tags if there are nickel or other divalent metal ions in the buffer ... sometimes traces of nickel my co-elute if you just recharged your column and it hasn't been extensively washed afterwards. I always add 1 mM EDTA to the dialysis buffer I use to remove the imidazole.
I try to keep the NaCl concentration as close to 500 mM as possible throughout all purification steps. For some of my protein constructs I have to lower it during tag removal because some of the proteases needed to cleave the His-tags are not active at NaCl concentrations over 250 mM, but after the cleaving protocol I raise it again. I always include a gel filtration (size exclusion chromatography) step at the end of purification and after this the proteins seem less affected by lower NaCl concentration than after the initial nickel affinity step.
When I'm done with purification, I concentrate my proteins to the desired concentration (in my case 6 to 20 mg/ml are ok for structural biology studies) and I then dilute the protein (still in the concentrator) with buffer that contains no NaCl until the NaCl concentration of the sample inside the concentrator is around 100-200 mM NaCl. I then concentrate the sample again to the desired protein concentration and freeze the protein at -80 degrees C in small aliquots (50 - 100 microliters). I found that thawing and refreezing the same aliquot more than 3 or 4 times leads to precipitation, so rather than freezing large aliquots I prefer to freeze small aliquots so that I don't have to thaw more protein than I need. Some proteins may need the addition of 5-10% glycerol for freezing properly but I have had no problems freezing them without glycerol so far.
Many of the proteins I work with start to precipitate quickly after nickel affinity chromatography. Buffers for nickel affinity chromatography generally have 300-500 mM NaCl in them, but afterwards people dialyse into buffers with 100-200 mM NaCl while removing the imidazole. I have established that in my case it is not the loss of imidazole that has a negative effect but rather that my proteins dislike buffers with low concentrations of NaCl. Give it a try, maybe keeping the NaCl concentration higher than 100 mM might help you. Also, freezing your protein straight after purification will stop it from precipitating and help keep its activity for years ( ... if your protein can be frozen).
His-tagged proteins can aggregate via their tags if there are nickel or other divalent metal ions in the buffer ... sometimes traces of nickel my co-elute if you just recharged your column and it hasn't been extensively washed afterwards. I always add 1 mM EDTA to the dialysis buffer I use to remove the imidazole.
I try to keep the NaCl concentration as close to 500 mM as possible throughout all purification steps. For some of my protein constructs I have to lower it during tag removal because some of the proteases needed to cleave the His-tags are not active at NaCl concentrations over 250 mM, but after the cleaving protocol I raise it again. I always include a gel filtration (size exclusion chromatography) step at the end of purification and after this the proteins seem less affected by lower NaCl concentration than after the initial nickel affinity step.
When I'm done with purification, I concentrate my proteins to the desired concentration (in my case 6 to 20 mg/ml are ok for structural biology studies) and I then dilute the protein (still in the concentrator) with buffer that contains no NaCl until the NaCl concentration of the sample inside the concentrator is around 100-200 mM NaCl. I then concentrate the sample again to the desired protein concentration and freeze the protein at -80 degrees C in small aliquots (50 - 100 microliters). I found that thawing and refreezing the same aliquot more than 3 or 4 times leads to precipitation, so rather than freezing large aliquots I prefer to freeze small aliquots so that I don't have to thaw more protein than I need. Some proteins may need the addition of 5-10% glycerol for freezing properly but I have had no problems freezing them without glycerol so far.
The protein also comes off the column more concentrated, so this may be an issue. Salt concentration does not have to be high. It just prevents some non specific sticking. I would find out what buffer your protein is soluble in, ie take the protein right after Elution, and dilute it into some different buffers to see what it likes and doesn't like.
one trick that might help is cleaving the tag off immediately after elution - this has worked for some proteins that I have worked on which had this problem. Adding EDTA to the eluted protein immediately ((ie while still in the fraction tubes) has also helped for one protein.
I strongly agree with above suggestions. Here I want to put my two cents. The buffer you are using may not be the optimal condition for your protein. Whether you change the protocol or not, if your protein does not like this buffer condition, it will behave strange. So, its better to optimize your buffer condition first. Dynamic light scattering (DLS) is the good option to see how your protein behaves in hundreds of buffer condition. Then, you can choose or optimize further the best buffer condition to get good quality protein.
Have a look on this paper, http://www.ncbi.nlm.nih.gov/pmc/articles/PMC2954336/
about optimization of buffer condition by DLS.
his-tagged proteins sometimes aggregate after etching Ni from the column. in thise case Ni can act as metal bridges between the proteins. give another chance to your protein by using EDTA to remove the Ni ions. good luck!
ps: imidazole help keeping the his-tagged protein in solution by "masking" the etched Ni ions avoiding the formation of the metal bridges
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