Hi Lane,

I agree to try the above suggestions as well. Is your protein soluble in that buffer to begin with, and did you do any optimization tests? How good was your expression to begin with, and do you know for sure the gunk crashing is your protein of interest?

To begin trouble shooting buffers, sometimes simply varying the salt helps in solubility. In my experience, increasing the salt concentration can help, and reduces non-specific binding. You can also try to add small amounts of detergents or glycerol (but this depends on your down stream applications as well!). You can also vary the buffering solution and strength. I had a his-tagged protein that was soluble in a phosphate buffer, but not in tris or HEPES.

Good luck!

Hi Lane,

I would start with a fresh buffer. Maybe the salt concentration is to high so that your protein precipitates. Did you check, if your protein is indeed precipitating (western blot of the white gunk)?

Really unusual/interesting problem...

Good luck!

Hello Lane,

before purifing the protein, u should load the protein on sds page, to check how much ur protein is actually expressed,if its expression is good ,u can actually then proceed to purification at range of lower temperatures.

Best of luck

Hi Lane,

I agree to try the above suggestions as well. Is your protein soluble in that buffer to begin with, and did you do any optimization tests? How good was your expression to begin with, and do you know for sure the gunk crashing is your protein of interest?

To begin trouble shooting buffers, sometimes simply varying the salt helps in solubility. In my experience, increasing the salt concentration can help, and reduces non-specific binding. You can also try to add small amounts of detergents or glycerol (but this depends on your down stream applications as well!). You can also vary the buffering solution and strength. I had a his-tagged protein that was soluble in a phosphate buffer, but not in tris or HEPES.

Good luck!

Perhaps there is something in your lysate (a substrate or another protein that forms a complex with the one you are purifying) that is essential for your protein's stability.

Is your protein membrane-associated? Maybe a bit of detergent would improve the solubility...

Is that really your buffer - 150 mM Tris sounds like a lot? I would use 20 mM NaP (or Tris, but I prefer NaP), and up the NaCl to 150 or 300 mM.

I would like to ask how you cleared your lysate. At what g-force was the spin? You may want to not only separate unbroken cells but also include a second centrifugation step to remove inclusion bodies.

Protein purification can be annoying! It looks like you already have lots of good suggestions.

- what is the predicted pI of your protein? Make sure that makes sense with the pH you're using.

- Is it a DNA/RNA binding protein and did you treat with DNase or RNase? I've had colleagues whose proteins became unhappy when the binding partner was gone.

- Definately try different monovalent concentrations. Also, I had a protein that became insoluble when a tag was cleaved off, so I did the cleavage step in a much larger volume - you can increase the volume during resin binding (might not help...).

- try adding a little bit of various detergents (NP-40, trition, etc) to see if it helps.

-10-20%glycerol or sucrose to keep your protein happier?

Good luck!

I totally agree with Valerie and Amanda. The first thing you should try changing is the pH of buffers. It's worth to check the pH of your cell lysate it can be different from that one you use for lysis. Notice that pH of Tris buffers is temperature dependent!

I too agree with the above suggestions and also with Mathew. 40 mM imidazol is quite high unless your protein binds very strongly the Nickel. It is never an easy task to find what it is failing. You should try some of the interesting suggestions above such a bit of detergent, glycerol, …play with salt concentration, ….Qiagen has a user manual for NTA-agarose protein purification with a troubleshooting section but those are guidelines. I think that purifying a protein needs that you establish a kind of scientific empathy with it at each step. For this you get to know it.

Also, if it is not a typo error, then 150 mM Tris buffer is a bit strange to me. Normally a buffers like this are only used to re-equilibrate quickly the buffer as a result of previous treatment either with an acid or base. You should avoid Tris when using metal-affinity purification but it can be used if its concentration is not too high.

Dear Lane,

Is your protein binding properly to Ni-NTA with that much imidazole in the binding buffer? If there is no loss in flow through then its fine otherwise try to decrease the imidazole concentration to 5-10 mM only in binding buffer and then optimize the wash conditions from 20-50mM in wash to make sure there is no protein loss in wash. Also are you sure that the protein ppt in wash is definitely your protein of interest? If possible, check whether the buffer you are using for binding is suitable for your protein to begin with? I usually go for 0.05 M Tris and it gives good results. You can also use 1XPBS. It also works well.

While I -as several other poster- use more NaCl and less Imidazol for washing, the main point seems to be that your protein was happy in this buffer and then is not happy anymore. So it seems you removed something that it needed.

Should you know co-factors, add them to your buffer. Same for other salts/metals (maybe something you tested in activity tests?)

What kind of protein is it?

hi...Ni-NTA works best at basic pH , so try to use 50mM Tris pH 8.5 with 300mM NaCl, 20 mM Imidazole, 5 mM B-ME, do not add DTT, add 10% glycerol. Give some wash with high salt buffer - 1M NaCl. keep less NaCl 150mM in elution buffer. But Tris pH should be 8.5. One more thing, protein tends to precipitate when it is tooo much that can be handled at some concentration in the buffer. So when you are dissolving your cell pellet before sonication/lysis, use more buffer- atleast 20 ml for a pellet of 1L. A diluted protein is less likely to precipitate.

Do reply if something works.

Protein estimation and preferably PAGE analysis, pI, etc, knowledge before hand helps to plan your strategy of purification.

Did your protein precipitate once it was eluted? We had this problem and it was found that nickel leaching out of the column was interacting with our protein and causing it to precipitate. We treated the precipitated sample with EDTA and dialyzed against PBS and the protein went into solution. You may also need to bring the protein concentration to < 3 mg/ml while dialyzing to make it soluble. Good Luck.

Although I am not that experienced, I would suggest you to load some of the white stuff on an SDS-PAGE, to see whether it is your protein or a protein complex. Perhaps, your protein is much concentrated under these conditions and it precipitates forming this white stuff. It might also be in a conformation that doesn't allow it being soluble, either because of the buffers you used or the way you handled it during the cells' lysis (a way that may be hard for the protein you work with). If this is the case, try to dilute it by adding some μls (or mls) of an 8M guanidine HCl solution (to reach a final concentration of 6M Gnd HCl). In this way you will succeed to lower the concentration of your protein and also denature it. Keep it in RT for ~1h to denature and then dialyze it against 150mM Tris, 50mM NaCl, to renature (refold). If there is not only your protein but a complex of proteins, you might succeed - in some way- to unbound the one you are interested in from the complex