Hi,
I'm expressing and purifying a membrane protein which at the C-terminus has a short linker followed by a TEV-mEmerald-TwinStrep tag. I can solubilise and purify the protein with Streptactin-XT resin suspension, with the protein being eluted off the resin with my buffer (25 mM HEPES, 200 mM NaCl, 1 mM EDTA, 5% glycerol, 0.05% DDM pH 8 + Roche Protease inhibitors) plus 50 mM biotin and then concentrated down by ultrafiltration, but I am having some trouble cleaving the mEmerald-TwinStrep with TEV protease.
To surmise what I did. After concentration I added the TEV protease directly to the protein on a 1 ug TEV per 5 ug protein basis as the sample volume was small (~250 ul) and the concentration low (0.16 mg/ml) so dialysis was unfeasible, and I deliberately didn't add any DTT or other reducing agent as my protein has a disulphide bond. I then left it at 4 degrees C with end-over-end rotation overnight to cleave. I ran a sample on a gel about 18 hours in, and I would estimate about 50% had cleaved. I therefore left it for another 12 hours before trying to separate the cleaved and uncleaved protein by size exclusion chromatography.
Unfortunately, it still looked like only about 50-60% of the protein had cleaved. I'm wondering if the low cleavage it might have been due to the 50 mM biotin that was still present. I assumed that it wouldn't make any difference to the protease activity, as TEV is typically quite active in most buffers and none of other components of my buffer are typically associated with inhibiting TEV according to literature. The 50 mM biotin is the only factor that I couldn't find any information on, the closest that I could find was on the elution buffer used for the older Streptactin resin (which used 5-10 mM desthbiotin instead of 50 mM biotin), which TEV appears to work fine in. Next time I will probably have to remove the biotin anyway with a desalting column etc so that I can bind the cleaved mEmerald-TwinStrep to the resin as the SEC was unable to sufficienty separate them. The other possibility I've thought of is that I didn't add any reducing agent, although some sources claim that TEV works fine anyway if added in sufficient quantity like I did.
If anyone has any suggestions on how I can increase the cleavage efficiency then I will welcome them.
Thanks!
I can't imagine it would be the biotin's fault. I have cleaved in the presence of 10 mM biotin with no issues. The TEV is generally very tolerant to all manner of conditions and additives.
I think your result is fairly typical for a large structured substrate. Some substrates cleave much more quickly than others. It depends on structural factors that you have little control over. This can affect both the vmax (cleavage velocity) and Km (degree of ultimate cleavage completion).
I don't even think that 50% cleavage is particularly bad. Just double the size of your production. Be happy that you got what you wanted at all.
You can't usually have "complete" cleavage with TEV, due to its naturally high Km for most substrates. If you must for whatever reason go further towards completion, the main thing to do would be to increase the amount of TEV. I see proteins that require a 1:2 or 1:1 ratio with some regularity. I'll give you 50:50 that it will work well enough.
As suggested by John presence of Biotin doesn't have any impact on the cleaving activity of TEV enzyme and you never get 100% cleavage. I usually do is dialyze my protein in the buffer containing no desthiobiotin/biotin overnight at 4 degree without any reducing agent and carry out the cleavage as well simultaneously with the TEV in the dialysis bag. I usually get more than 80% of cleavage by TEV by doing this. In your case since dialysis is not feasible try to desalt it using filters to get rid of biotin and then perform overnight digestion using TEV (increasing the enzyme concentration).
Thanks Shahid Chaudhary and John Schloendorn for your helpful suggestions. When I next purify this protein I shall use a desalting column to remove the biotin and I will use more TEV to try and improve the cleavage. I was actually going to ask when you thought it would be best to desalt (i.e, before/after cleavage), but I shall do it before based on what you say. Would you recommend though to desalt it before (i.e right after elution) or after concentrating the protein though?
To be honest desalting is not required at all after elution but you can try to filter out biotin after elution, exchange buffer and then perform TEV cleavage overnight with increased enzyme concentration.
DEar Samuel
do you have any information about aggregation state of your proteins?
Im my experience the most cases when TEV fail to cut at 100% is due to presence of aggregates and inability to TEV to reach the digestion site. It is happen often when you try to solubilize membrane proteins with MBP fusion for example.
ciao
Manuele
I analysed the purified protein with mass photometry before I added the protease Manuele Martinelli and it didn't suggest that the protein had aggregated - the majority was in a monomeric state, with a small proportions being dimeric. When I was screening detergents for solubilisation efficiency, fluorescent size exclusion didnt suggest that the protein aggregated under the condtions I later used for purification. I'm therefore pretty confident that my protein is not aggregated.
I've noticed that Thermo sell dedicated colums for removal of dyes and biotin, has anyone used these out of interest and do you think they are worth the slightly greater cost compared to the standard Zeba desalting columns they sell? I'm only really interested in the Biotin removal aspect. https://www.thermofisher.com/uk/en/home/life-science/protein-biology/protein-purification-isolation/protein-dialysis-desalting-concentration/small-molecule-removal.html
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