How long do you leave the IB in urea? I usually incubated overnight under agitation, and this led to almost complete solubilization.

In addition, you might need to be sure to have washed IB from membrane using a detergent before doing the solubilization in urea.

Hi Barbara. I prepare my IBs and resuspend them in 1ml deionised water after whcih I add approx 20 ml solubilzation buffer ph=8.0 for 2-3 hrs on end to end rotor at RT. I had tried the washing steps earlier using wash buffer- 50mM TRIS, 1Mm edta AND 1% triton, followed by a NaCl wash and buffer only wash. but I was losing substantial amount of protein in each of the washes so I have discontinued them. Even without washing my protein is relatively pure but it is not completely solubilzing thereby yield is less. 

Also, reducing the temperature to 18*C during the expression can be helpful

@Jessica... I have already tried lower temperatures and various inducer concentrations for protein expression but my protein is always in the inclusion body. 

Why were you losing protein in the washing? Was it going in the supernatant? So was 1% triton sufficient for solubilize it, at least partially? If this is the case, you might try to include a detergent with urea, during solubilization of IBs. I would also increase the incubation time to over night and do the agitation more vigorous.

Another important thing is considering if your protein has cysteines that could make covalent bonds if you are not in reducing conditions. If it is the case, the protein that remains aggregated could have formed disulfide bridges that favor the aggregate. I would include a reducing agent as DTT or TCEP. Good luck!

reduce the temperature to 18 or 16 , use 0.1%-1% of sarcosyl or 1% triton in lysis buffer

Hi Ankita, can you harvest your protein before it is deposited into inclusion bodies? That way you have a chance to isolate your protein in a soluble form  

Inclusion body proteins are usually a mess of denatured proteins that can never be renatured.

Try this combination: 1% sarkosyl, 2% Triton-X 100, 20mM CHAPS and then purify using Pierce GST-Glutathione Spin Columns. if your tag is GST

Your problem is not easy to solve. Do you have a specific assay for the protein? If yes then check your small scale prep for activity. If you are just looking at SDS-Page, be careful. A small fraction of your protein may be soluble, but still an aggregate (check by gel filtration, light scattering). You will have to change to a GST vector or some other tag(SUMO, MBP) to get around the issue. In my experience GST fusions are better behaved in E.coli. But you cannot do the denatured purification now. Somebody in your institute may have GST and His tag in the same plasmid, try that. You could attempt to change the protein boundaries that you are expressing.

I have been through this and it is very frustrating. None of the above suggestions may work. Try changing your project, if it is feasible. Some proteins are very difficult to purify from E.coli. One has to be realistic.

Good Luck

What about your protein? Does it contain disulfide bonds? If so, you need to add a renaturation step in your process...

Hi Ankita,

I would use 6-8 M guanidinium HCl in HEPES or Tris with a detegent (OG or triton 5mM)in your case for solubilization. Over night at room temperature should work out.

You probably will lose up to 5-10% of your protein during solubilization but compared to what you are telling it is a small loss. Also try to get rid of EDTA if you are using it. It may counteract with your Ni-Beads.

As Can says, the best strategy is the guanidinium HCl . The DTT is not compatible with common Ni-NTA resins. During treatment with the chaotropic agent, in many cases it is convenient to leave the inclusion bodies for several minutes (10-60 min) under mild agitation to ensure complete dissolution.

In order to recover soluble proteins from the inclusion bodies,  inclusion bodies are often solubilized in the presence of strong denaturants such as urea or guanidinium hydrochloride, followed by the removal of the denaturants under optimal conditions that favor refolding. However it seems that you got problem with urea and could be the same

Try a simple step. change your pH, rather higher than 8. It may work

Guanidium HCL at 6M (not possible to reach higher concentration considering its solubility) should be good. However, you could try to use ionic detergent like Sarkosyl or Fos-Choline derivatives (FC-12 is the common one) which are probably more efficient than CHAPS or TX-100 to solubilize IB.

If one of your goal is to renature the purified protein, choose detergent with high CMC as you will be able to remove them easily by dialysis simultaneously to decrease Gn-HCl concentration.

Even the fact that IMAC columns are known to tolerate "denaturing conditions", it should be better to decrease the denaturant concentration  to optimize interaction of the His tag with the resin. Remember that to remove GnHCL, it should be better to do it step by step, against decreasing concentration of GnHCL first and Urea after (i.e. 1st dialysis vs GnHCl 3M, 2nd vs Urea 4M, 3rd vs Urea 2M and final step against your prefered buffer).

if your protein is coming in inclusion body totally, why not attempt isolating inclusion body and solubilising, instead of purifying in IMAC. Please go through the papers of Dr Amulya Panda of NII

Hi Ankita,

Did you get the protein solubilized from IB significantly. I am facing the same problem. I tried 8M Urea and 6M GnHCl but the yield is less. Most of the papers discuss about refolding but not on solubilization. Please share the details if you have succeded. Also, would like to know the pI of your protein.

Regards