You do not mention your cell lines, growth conditions or whether you optimized expression and lysis protocols to minimize insolubility. 6HIS or 9HIS under denaturing conditions works quite well. Your pH may be off. Things to consider, clone your gene in fusion with either maltose binding protein or SUMO to increase solubility. Reduce expression temperature to 16 C and express for 16-20 hours. Lower your IPTG concentration. Co-transform with chaperone expression plasmids from Takara. I always avoid having to pursue refolding of denatured proteins. Reviewers always are skeptical about a refolded protein. Insoluble proteins are not fun. When I encounter one, I start over by fusing it to MBP. This always solves my problem. Good luck.

Do you find the protein in the flow through? If so, you can try to use copper instead of nickel in your IMAC column. Or play with pH, it can also increase the binding. Make sure you don't have EDTA in your sample, sometimes protease inhibitors contain EDTA, and that screws any IMAC.

Thanks for your input. We were working with Ni but Co will be tried soon.... But for an efficient purification, I must do something to improve the efficiency of solubilisation either with urea or sarcosyl....

Co binds proteins much weaker than Ni, so I'm not sure if it helps. You could try once to solubilise a sample with SDS, then you'll know how much protein you have there, run an IMAC, SDS-PAGE and Western and then compare with other, non-denaturing detergents.

Hi,

Is your protein known for being insoluble? As it might not be as simple as being a column issue - have you thought of using a different tag, particularly ones that are good for solubility issues, such as GST or NusA? Or changing the host, using an Origami (DE3) rather than BL21?

When you know the protein is soluble (in the detergent environment), lets think the 6XHis is well exposed to Ni or Co ions. If you are usind chelating columns, try to pass your lysate in a very slow flow rate!, if you are adding imidazole on binding, try to keep it as less as possible, since it may also compete with your protein to be bound with Ni/Co.

If inclusion bodies are the only problem then no way, you have to go for refolding your protein which is painful :(

Good luck

What is the protein you are purifying? The nature of the protein will be cruical to know in order to avoid copying and pasting the entire contents of all handbooks of protein-purification.

Anyway: consider an other expression system (like e.g. yeast).

Depending on the purified protein you could consider pre-purification using Ni-NTA in batch purification. I had bad experience using Talon beads with a particular protein (that is to say I am not down voting Talon beads, they just in a certain procedure gave worse results). Also the use of a His-Trap column did not give as well yields as Ni-NTA batch.

Instead of SDS, I would rather first try Triton X100. SDS is known to be difficult to remove from protein preparations and in fact, it can (which I have seen!) drastically reduce the yield of your protein in IMAC.

You do not mention your cell lines, growth conditions or whether you optimized expression and lysis protocols to minimize insolubility. 6HIS or 9HIS under denaturing conditions works quite well. Your pH may be off. Things to consider, clone your gene in fusion with either maltose binding protein or SUMO to increase solubility. Reduce expression temperature to 16 C and express for 16-20 hours. Lower your IPTG concentration. Co-transform with chaperone expression plasmids from Takara. I always avoid having to pursue refolding of denatured proteins. Reviewers always are skeptical about a refolded protein. Insoluble proteins are not fun. When I encounter one, I start over by fusing it to MBP. This always solves my problem. Good luck.

I agree about the exposition of his tail so I don't think you're not binding the protein (of course check flow rate, EDTA, etc). Regarding the detection, I'm not sure if I understood. When are you detecting your protein? Directly in urea 8M or after refold it? If you have problems with solubilization in urea you can try other solutions, e.g. Guanidinium Chloride 6M and adding DTT (maybe you have something very big and insoluble due to a net of disulphide bonds). If your problem is to refold it, then it's more difficult. You should try other refolding method or other additives. There is no law about it and only a trial and error strategy is possible.

To overcome the solubilization problem you may express the His-tag on the other teminus of the protein. I once expressed a protein with C-terminus His-tag and it was expressed as inclusion bodies but when I expressed the same protein with an N-terminus His-tag I got high yield of soluble protein,

Regarding the purification you may increase the residance time to give the protien more time to bind

Hi,

it is difficult to do the refolding of a protein and even more recover the activity. What you could do is to use the DE3 cells as Meena recommended, use low concentrations of IPTG (0,1mM) and grow the cells for long time (over night) at low temperature (16-20 Celsius). For any protein going to inclusion bodies you mast avoid leaky expression.

Good luck!

I assume you are sure your bacteria is expressing protein. Have you tried sonication in addition to Urea (or guanidine)? It may help to destroy big aggregates allowing denaturant to better access the protein in the inclusion bodies. Have you check the pH to be correct before loading the column? I often used recirculating mode to reload over and over the column (using a peristaltic pump) o/n at 4°C. I agree that non denaturing conditions are preferable and it is very important to know the characteristics of your protein before applying a method or another.

Consider also that even when most of the target protein is in aggregates there may be plenty of soluble material with which to work. In this case try increasing the culture volume and you can use non denaturing conditions (in this case sonication can be of some help to increase the amount of solubilized protein)

See if you can find useful hints in the attached (old) protocol.

You could also try to skip the refolding step by increasing the proportion of soluble protein. Cultivating the cells at 18 ºC over night after IPTG induction can do the trick. If your protein still goes to inclusion bodies and need to refold you can try some of the refolding kits available in the market to find the optimal refolding conditions.

If the protein is small (single domain 10-30kDa) chances are pretty good that you will be able to renature it in vitro. In this case I would use 6M GdmHCl instead of 8M Urea to solubilize the inclusion bodies (but you wont be able to run SDS-PAGE of the sample). If done properly, this solubilized (denatured) protein will be already very pure and you wont need to do a Ni-NTA chromatography (maybe after renaturation).

If the protein is a big, multidomain molecule (>30kDa) or a membrane protein chances are slim that you will be able to renature it in high yield. In this case I would optimize the expression conditions (Temperature as low as 16C o/n and/or lower IPTG concentration) - as mentioned by others. ALternatively, use a different cloning construct or expression host.

I did not read all the answers and probably it was mentioned before. But, you can try to use a construct with (6xHist)-MBP (N-terminal to you protein). This Maltose Binding Protein (MBP) will (generally) make protein more soluble.

Also, use minimal growth and induction conditions such as 30 degree for grow and 24 degree for induction with 0,1mM IPTG.

Based on my experience, it is very difficult to archive a good quantity of protein with all those purification methods, such as detergents and non-detergents solubilizations. Plus, it will make your protocol much longer, expensive and might not be feasible.

Hope it might help.

Hi AncI can see already there are so many good suggestion to inmrove the solubility of your protein and if they work i feel you need not to worry about refolding. If finally you have to refolding then as per my experience you can try following things.

First check the PI of your protein and decide the appropriate pH for your buffer. I would prefer rapid dilution method instead of Ni-NTA binding. So just check the concentration of your protein in 8M urea and then do rapid dilution in refolding buffer containging ~2 M of urea and GSH/GSSG (if your protein have Cys then) at a final concentration of 0.1-0.2 mg/ml. Let refolding reaction go for 18 hours at 4C with very mild stirring. Then dislyze your protein against 0 urea buffer with ~5 mM BME. and filtration. I hope it should work. Good luck.

Hi, for improving affinity try using a Cobalt column insted of a Ni column. If you are willing to make bigger changes, use MBP or NusA as affinity tags.

It´s your protein from bacteria? If not see if its a case of codon bias which produces high insolubility, in that case you can use BL-21 codon plus cells.