The correct answer to this question is to do a full scale stability experiment, with different protein concentrations with different buffers, pH's, adjuvants, freezing conditions etc. Take samples at various times and assay for activity.

Most of us, of course, don't have the time to do such studies for each protein we work on, so a more limited set utilizing whatever is available to you, would be the best solution. Also consider where you need to ship it to, and how you will have to send it. I have shipped proteins from the US to Australia for example, where dry ice is a no-no, so the protein has had to be stable at 4C for a week on wet ice. This required finding buffers that the protein was stable in under lab conditions, and assaying at various times points > 1 week.

One of the simplest things is to simply put your protein into a "neutral" buffer such as PBS (or even simply leave it in what it is in), add an equal volume of glycerol and store at -20C. However, glycerol might interfere with downstream applications, especially if structure determination is going to be involved.

Flash freezing small (single use) aliquots in liquid nitrogen and then storing at -80 seems to be a good method for most proteins. This is usually my first method of choice with an unknown protein. But this requires you can ship on dry ice. Your buffers should be fine for this, although consider the effect of 1 M salt on downstream uses too. Would it be better to dialyze (or dilute) it now, before storing, or let the end user do that? Avoid repeated freeze/thaw cycles as much as possible, until you can verify your protein is OK with that. Aggregation can still occur.

If you have access to a lyophilizer, then this can be a good method. You don't have to dialyze into water - you can use a volatile buffer. However, many proteins will tend to aggregate under many lyophilization conditions, so you will need to reconstitute a sample and ensure it remains active.

For shipping on wet ice, an ammonium sulfate pellet can also work. Precipitate your protein with sufficient ammonium sulfate, pellet, and store at 4C. This might need to be dialyzed by the end user to get it into a buffer appropriate for their assay, of course.

Remember that cofactors can greatly stabilize a protein, so add any of these before storing. Think about adding DTT too to prevent oxidation, and EDTA to chelate metal ions (inhibit metaloproteases)... Filter sterilize your final solution, and consider adding sodium azide for long term storage, esp at 4C.

So, in summary - try all 3 of your own suggestions and see which works best!

The correct answer to this question is to do a full scale stability experiment, with different protein concentrations with different buffers, pH's, adjuvants, freezing conditions etc. Take samples at various times and assay for activity.

Most of us, of course, don't have the time to do such studies for each protein we work on, so a more limited set utilizing whatever is available to you, would be the best solution. Also consider where you need to ship it to, and how you will have to send it. I have shipped proteins from the US to Australia for example, where dry ice is a no-no, so the protein has had to be stable at 4C for a week on wet ice. This required finding buffers that the protein was stable in under lab conditions, and assaying at various times points > 1 week.

One of the simplest things is to simply put your protein into a "neutral" buffer such as PBS (or even simply leave it in what it is in), add an equal volume of glycerol and store at -20C. However, glycerol might interfere with downstream applications, especially if structure determination is going to be involved.

Flash freezing small (single use) aliquots in liquid nitrogen and then storing at -80 seems to be a good method for most proteins. This is usually my first method of choice with an unknown protein. But this requires you can ship on dry ice. Your buffers should be fine for this, although consider the effect of 1 M salt on downstream uses too. Would it be better to dialyze (or dilute) it now, before storing, or let the end user do that? Avoid repeated freeze/thaw cycles as much as possible, until you can verify your protein is OK with that. Aggregation can still occur.

If you have access to a lyophilizer, then this can be a good method. You don't have to dialyze into water - you can use a volatile buffer. However, many proteins will tend to aggregate under many lyophilization conditions, so you will need to reconstitute a sample and ensure it remains active.

For shipping on wet ice, an ammonium sulfate pellet can also work. Precipitate your protein with sufficient ammonium sulfate, pellet, and store at 4C. This might need to be dialyzed by the end user to get it into a buffer appropriate for their assay, of course.

Remember that cofactors can greatly stabilize a protein, so add any of these before storing. Think about adding DTT too to prevent oxidation, and EDTA to chelate metal ions (inhibit metaloproteases)... Filter sterilize your final solution, and consider adding sodium azide for long term storage, esp at 4C.

So, in summary - try all 3 of your own suggestions and see which works best!

I hate TRIS buffers. I would recommend using MOPS or another Goode buffer. The main keys to protein stability are purity, concentration and temperature. Storage for long periods between steps can lead to denaturation and loss of activity. You may just have to go entirely through the purification, from start to finish (spending 48 hours in the cold room is not uncommon).

If you concentrate after anion exchange, you can also perform a buffer exchange at the same time to reduce the salt concentration (this is also much faster than dialysis).

Freezing the protein can lead to loss of activity. Try adjusting the pool to a final 30% glycerol concentration and storage at -20.

I think Richard said it all in his first response. All proteins are not created equal so a condition that works for one protein may not work for yours.

I concur, try all three of your suggestions mentioned above. Good luck!

Forgot to add a reference!

http://www.ncbi.nlm.nih.gov/pubmed/20439424?dopt=Abstract

If you purified under native condition without urea solubilisation, have to remove the salts by dialysis or using cut-off membrane then lyophilise the protein samples and take an aliquots to check the stability of protein by assay or SDS-PAGE (to check the protein degradation).

Thank you for all your constructive answer!

@ Richard Heath

After the precipitation by the ammonium sulfate, isn't there a risk that the protein aggregates when you dialyze the salt because of a too high concentration?

Unfortunately I don't have access to the publication in my lab ...

@ Arghavan Golbaz ·

Do you mean that imidazole can have an effect on the storage or just that it is more comfortable to have your protein in a "clean" buffer after thawing?

@Kumar Subramaniam ·

This is an idea but I just can check the stability by SDS PAGE. So we will see!!

You all talk about PBS buffer. Is it because you have had problems with other buffers? And so, can we say that PBS is THE buffer of reference when you begin a storage assay?

I think PBS most often resembles the internal environment of the cell better than other buffers, but all proteins react differently to different buffers, you just need to stumble onto one buffer system/ salt concentration/ protein concentration / storage solution that works for your specific protein.

"you just need to stumble onto one buffer system/ salt concentration/ protein concentration / storage solution that works for your specific protein. "

This is actually what I try to do!

But instead of doing random tests that cost both time and money, I try and find a light in this fog.

Dear Alexander GAY, with my little experience, if the protein ended up in a buffer having traces of imidazole it aggregates ultimately, hence i suggest possibly exchange the buffer to remove the traces of imidazole from the protein and can preserve with glycerol at -80 C. Besides, Dr.Richard Heath has provided extensive and satisfactory solution. You can try out. Good luck..Cheers..:)

Choice of buffer: I always prefer phosphate buffer over any other. PBS pH is around neutral. But the phosphate buffers (Gomori buffers) of different pH can be prepared easily [Please see attached file taken from one of the appendices of Molecular Cloning, Sambrook and Russel]. Some proteins get aggregated when at certain pH conditions. We have to take this also into consideration. So, during imidazole removal, I suggest to exchange protein solution against any of the phosphate buffers of your pH choice. You can even have required NaCl in this buffer. This buffer might sequester any divalent cations in solution (that's the only problem).

Stability of protein: Temperature is one important factor here. Freeze-thaw cycles and sometimes even lyophilization affects protein structure and stability! If you find your protein can be retained in solution form and also active at up to say 8-10 degrees centigrade for long duration (36-48 hrs), it will be great to transport as solution in ice pack with dry ice in good insulator boxes. This should reach destination within a day or two!

All the best!!!

Your protein is soluble in Imidazole, and if you stored your protein along with imidazole, it will precipitate out your protein and will create problems in  downstream applications. You need to dialysis the protein against PBS and which will not effect your protein quality and can be used for further study. The other option is to use a Lyophilizer machine to get the protein in powdered form without any moisture. You can then store your protein at -20C for long time without any confusion and can prepare the concentration whatever you need. It is also helpful to get the same quality protein for your all experiments and you can get the good result with the same sample.

well wisher

is it ok to collect cells and spin to get the pellet, freeze the pellet at -80 and use it in a month for w.blot and other work such as RNA extraction?

Will storing the protein at -80 oC not affect the protein??

I have to use my protein for cell culture studies. So if I store it at -20 with 10% glycerol, will that not interfere with my experiments further? Do I have to dialyse it again and then use for the experiments?