Protein purification using Ni-NTA
There are several methods commonly employed for cell lysis (e.g. French press, sonication, cell disruption, etc…) but in all cases it is best to tweak the lysis/binding buffer to reduce aggregation right from the start rather than after the Ni-NTA step. Things to remember are:
- use at least 50 to 100 mM Ni-NTA buffer (TRIS, or HEPES, or phosphate, etc…) at a minimum pH of 7.6 and a maximum pH of 9 (around pH 8 is best unless your protein has a pI of 8, which means your protein would be unstable at pH = 8). Using less than 50 mM might not actually buffer your lysate, which can often be more acidic than you realise. At a pH below 7.6 the binding of 6His-tagged proteins is reduced, so make sure your buffer is strong enough.
- use at least 500 mM NaCl in your lysis buffer as most proteins are more soluble in high salt and the non-specific binding of non-his-tagged proteins is reduced.
- Add 2 to 5 mM betamercaptoethanol (BME) 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. I prefer not to use DTT at this stage as DTT is a stronger reducing agent than BME and will lead to a brown mess even quicker than BME.
- you can also add 5-10% glycerol to your lysis buffer as this helps protein solubility/stability.
- proteins are generally temperature sensitive and will aggregate if kept too warm or too cold. Most proteins should be kept at 4° C during all purification steps after cell harvest, though proteins from thermophilic bacteria can precipitate if keep on ice or at 4° C for too long.
The presence of contaminating proteins can lead to precipitation after Ni-NTA and your protein might actually become stable if you simply add a second or even third purification step such as ion exchange or size exclusion chromatography. For structural studies, further purification steps are generally required to obtain a homogeneous protein sample that runs as a single band on a denaturing SDS-PAGE gel in the presence of 5 mM DTT. Again, buffer choices are important:
- you can dialyse your protein into a different buffer that’s better for your protein than the buffer needed for Ni-NTA. 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).
- use less salt now, 50-150 mM NaCl is usually OK, though some protein are not “happy” with less than 200-300 mM NaCl, particularly those with a pI over 9 or under 5.
- if your protein has several cysteines that can potentially form disulphide bridges, change the BME to DTT at this stage (1 to 2 mM is generally enough). After your last purification step (e.g. after size exclusion chromatography) you won’t need DTT or any other reducing agent if your protein doesn’t have any cysteines at all and your sample is pure (no other contaminating proteins are present). If you still need a reducing agent, consider TCEP at this final stage, it’s probably the most stable BUT it’s also VERY expensive, so there’s no point in using it in earlier steps.
- unless you need to remove it because it interferes with your assays, keeping 5-10% glycerol in all your buffers can help with your protein’s solubility and will also help if you’re planning on freezing it. However, you will generally need to remove the glycerol with a final dialysis step if you plan to use your protein for crystallography as glycerol is a well-known nucleation inhibitor.
- if you need to freeze your protein for long-term storage, freeze it in small aliquots (50 to 100 micro litres). This way you can thaw as much protein as you need for any given experiment. If you freeze a large volume, then thaw it to remove an aliquot and refreeze the rest of the sample, the sample will often precipitate badly when you thaw it again. So … freeze small aliquots instead.
This guide was authored by:
Dr. Antonio Ariza
ORCID ID: 0000-0003-4364-823X
University of Oxford
Sir William Dunn School of Pathology
South Parks Road