Hello.
I am working with a recombinant enzyme from a marine bacterium, attempting to overexpress it in E. coli and purify it using His-tag affinity chromatography. Since I cannot perform all characterization assays in a single day, I tried different storage methods:
- Flash-freezing some enzyme solution with liquid nitrogen and storing it at -80°C, both with and without 10% glycerol.
- Storing some enzyme solution at 4°C.
After three days, I compared protein concentrations using a Bradford assay. Surprisingly, the enzyme stored at 4°C had the highest concentration, while the flash-frozen sample without glycerol showed a very low concentration, likely due to protein degradation. That is why I use the enzyme stored at 4°C, but I am unsure how long can I use the enzyme.
I hope to use the flash-frozen sample with 10% glycerol, but I am concerned about its impact on enzyme concentration and activity.
Here is my questions:
Any insights would be greatly appreciated! Thank you very much.
A protein should not degrade at -80°C. It might lose activity but it should not degrade. And the Bradford assay only assesses protein and not activity. So the fact that you see a low concentration after storing at -80°C suggests you may have a sampling error problem. Perhaps it wasn't mixed well and when you took a bit out to assay your sample had excluded the protein. Hard to say exactly what happened but I can't imagine it was degradation. You might try again and freeze away small aliquots (that were well mixed) and thaw the entire aliquot to measure.
Some proteins are very stable at 4°C, others less so. It will depend upon your protein. But activity measurement is really the best thing to measure since you don't care if you have a lot of protein if it's all dead.
But I think freezing in 10% glycerol is generally good for most proteins (but maybe not for everything).
I never use glycerol for enzymes, it can affect enzymatic activity (but you can always test it first, many enzymes are just fine). I agree with what Michael said. Other than that, I wonder about your purification... Did you dialize imidazole out before freezing? High concentration of imidazole definitely supports protein precipitation, especially in the context of freezing-thawing, so that would be my first guess of what went wrong.
Katarzyna Radziwon Thank you for your answer!
Yes, I dialyze the enzyme twice (1 hour each) to remove the imidazole. However, I did not dialyze against a salt-free buffer. My dialysis buffer contains 0.3 M NaCl, just like the lysis and elution buffers, because when I tried to reduce the NaCl concentration, my enzyme precipitated inside the dialysis bag. Could that be a problem?
What's the volume of dialysis buffer?
Never do salt-free with a protein. I typically use 150 mM NaCl in storage, 300 mM might contribiute to your problem but I don't think it's the main issue.
If the enzyme works just fine stored at 4C, go with that, I worked with some protein that were fine after months at 4C (but it's generally rare).
Katarzyna Radziwon I used 2 L dialysis buffer for 15 ml of enzyme solution.
Thank you for your insights!
Best to make small beads of the concentrated enzyme solution by allowing small drops to freeze in liquid nitrogen. You can then estimate the enzyme concentration and activity by simply weighing a bead or two. Then dilute with the buffer of your choice. Best to store the beads in small plastic bottles under liquid nitrogen. Has worked for me for the last 45 years.
I just read that the pH of phosphate buffer can decrease when frozen. Could this be a potential source of problems? My final buffer for enzyme storage contains 50 mM phosphate buffer, 0.3 M NaCl, 10 mM 2-mercaptoethanol, and 0.5 mM PMSF.
Should I consider using a different buffer?
Phosphate buffer is not very T dependent but is pressure dependent. Trizma buffer is very T dependent. The key is to make sure your enzyme is frozen as a concentrate. After thawing and weighing the beads, you can then add whatever buffer you wish, Trizma or phosphate and you will be OK. Many people use -80°C, which is OK. Just for me, I prefer liquid N2 for long-term storage.
You have BME in your storage buffer... I assume reducing agent is necessary. BME is not stable, for long term storage use TCEP. That's probably not the main issue here because you'd observe an effect mostly with the 4C sample.
I'm wondering if you ran a gel of thawed sample? You're suspecting degradation, I suspect agreggation. Gel would clarify that. Degradation would be for sure faster at 4C, so that's unlikely. If it's aggregation, make sure you're not concentrating your protein too much (I'd say a safe zone is up to 40 uM, but for many proteins it might be way less). I'd also lower salt concentration so it matches typical PBS, that's around 150 mM. PMSF has also an incredibly short half life, if your buffers are filtered and your protein pure, there's no reason to store it with protease inhibitor, especially considering it's not functional any more. Of course, there is a big question, is your protein pure after just Ni? Probably not. His tagged proteins require a polishing step typically.
Michael L. Smith Thank you for your suggestions.
Katarzyna Radziwon I think you're right about the aggregation. I thawed another aliquot of the enzyme, and there were white clumps in the flash-frozen sample without glycerol.
Thank you for your suggestions. I will try to do it.
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