without knowing a lot about your protein other than it is animal derived membrane protein, my first thoughts are on why your protein is insoluble:

1. Have you tried different expression strains of bacteria: BL21, Xl1 blue, JM109 etc. If your protein has disulphide bonds bacteria will struggle if your protein is cytosol expressed (reducing environment). Try expressing in the periplasmic space which is a more oxidising environment using a suitable signal.

2. Codon bias: E coli and animals do not necessarily share that same use of codons and hence E.coli may have limited supply of certain tRNA's. There are strains available to overcome this

3. Is you protein glycosylated: does it require this for folding, if so E coli can't provide that sophisticated post translational modification.

4. The pH of your lysis buffer maybe too close to the pI. You could try using a slightly more acidic pH. Try lysing your cells with lysosome/sonication instead

5. your protein is sticking to other E. coli proteins etc that are pulling it down: higher salt, detergent levels (try different detergents), change pH may help.

good luck

without knowing a lot about your protein other than it is animal derived membrane protein, my first thoughts are on why your protein is insoluble:

1. Have you tried different expression strains of bacteria: BL21, Xl1 blue, JM109 etc. If your protein has disulphide bonds bacteria will struggle if your protein is cytosol expressed (reducing environment). Try expressing in the periplasmic space which is a more oxidising environment using a suitable signal.

2. Codon bias: E coli and animals do not necessarily share that same use of codons and hence E.coli may have limited supply of certain tRNA's. There are strains available to overcome this

3. Is you protein glycosylated: does it require this for folding, if so E coli can't provide that sophisticated post translational modification.

4. The pH of your lysis buffer maybe too close to the pI. You could try using a slightly more acidic pH. Try lysing your cells with lysosome/sonication instead

5. your protein is sticking to other E. coli proteins etc that are pulling it down: higher salt, detergent levels (try different detergents), change pH may help.

good luck

Hi, I totally agree with Ian, membrane proteins are usually difficult to treat and solubilize

What percentage did you use? A key to efficient solubilization of a membrane protein is having sufficient detergent to fully solubilize all of the lipids (unless you have a detergent that can selectively extract your protein. The amount of detergent will depend upon the amount of cells. There are a number of reviews out there that give pretty good rules of thumb. I have first lysed my cells (2gm wet weight) in something like bugbuster or your own lysis solution. However, lysis using a french press or high pressure homogenizer is better, you should include benzonase or other nuclease in either method of lysis. The membrane fractions, hopefully containing your protein inserted into the membrane (assuming you have used the french press or disrupter, can be collected by high speed centrifugation 120kxg or so. You may want to try a low speed spin first 21kxg for say 10-30min. If your protein comes down it probably is in the low speed spin its likely to be in inclusion bodies. Once you get the membrane fraction, if it still contains your protein, you can try various detergents and different lipid:detergent ratios to see which solublizes your protein best. My colleagues and I have found that membrane protein solubilzation is far more reproducible with the french press/ or pressure disrupter. Lysis by sonication is less reproducible, although it can work if you ensure that you really break up the cells.

I may try pH 6.5 with detergent. The following was provided to me:

The detergent buffer consisted of 20 mM HEPES, pH 7.5, 2 mM CaCl2. Digitonin (Sigma,D141), n-Dodecyl-β-D-Maltoside (DDM) (Invitrogen, P/N BN20051), Triton-X100 (Sigma, T9284) and a mixture of Triton-X100, 0.5% Sodium Desoxycholate (Fisher Sci, S-285), 0.1% SDS, were used for membrane extraction at 4C during 30 min with gentle vortexing, followed by centrifugation at 17000 x g/30 min.

I work with many proteins which are insoluble in bacteria. 6M urea works great for me. I sonicate the bacteria in a buffer containing 6M urea and then purify the protein in urea containing buffer. All proteins sonicated in urea will mostly be in the supernatant. The urea is then removed slowly by dialysis in a step-wise manner (3M urea, 1M urea, 1M KCl etc.). Some more information about this can be found in the manuscript - Gomes et al., J Biol Chem. 2004 Nov 26;279(48):49579-87.

This should work for most proteins, however a few proteins may be irreversibly denatured by high urea concentrations.

I also work on GST fusion protein using Potassium phasphate buffer pH 7.2, 1 mM EDTA and PMSF and 0.1% TritonX 100. I suggest, Please find first that ur protein is coming in soluble form or not. As, Sometimes GST interfere with the proper protein folding due to its large size. Please check the solubility of your protein in different buffer like HEPES etc.

You can also try different E.Coli strain (Codon plus, pLys) useful for Eukaryotic protein expression at low temperature between 20-24 degree celcius after induction.

I also recommend to use 6M urea to solubilize your protein. Just re-suspend your pallet with protein in buffer containing urea and KCl, keep it in all purification steps. You can remove urea by gel filtration or by changing buffer. This worked for me in most of the in-soluble proteins.

Thank you all for your kind suggestions. I already tried different strains of E.coli and also tried various other parameters also.I will follow your suggestions and try it again. Hope for the best.

There are two potential problems you may have that others are talking about. The first is that you have a membrane protein which would be in the insoluble fraction unless you can use the right detergent to solubilize it. Then there is the issue that the protein may be sggregating and in inclusion bodies. For the first case, I would recommend to try what John Flanagan has written. You maybe can extract your protein relatively easily, rather than change the codons, etc etc. as Ian Wilkinson recommends. But, if your protein is still insoluble, and requires addition of urea, then I do think you would have an uphill battle to purify this protein in EColi and have it be active in the end. Then I would recommend switching to Baculoviruses or mammalian cells for expression. We and others worked for months to try to express our membrane proteins in EColi. It was just an uphill struggle. But it did finally work by others but it took years to get some piece of the protein expressed and active in EColi.

I agree with all the suggestions above. It is a common problem to express foreign proteins in bacterial. Some of the proteins maybe toxic to bacterial, so they just through them into inclusion bodies :-)

If you have tried all the options (low temperature expression, low IPTG concentration for induction, strains, detergents, pH, denature and refolding, different cell lysis methods....), You might either consider to use different expression systems such as Baculorviru or others (they are usually lot more expensive than bacterial system) or try different bacterial expression vectors, or maybe co-purify the protein with small chaperone proteins. You may consider using SUMO/His double tag expression vector, which usually helps protein expression level and protein solubility in E.coli. It worked very well in our lab! The other advantage is that the double-tag can be efficiently removed by ULP1 sumo protease and it is a very efficient protease comparing to Thrombin, TEV, and EK, and you only need 1ul (12U) to remove fusion tags from 1mg purified protein with >95% cleavage at 4*C for 16 hours. It is cost effective as well since you don’t need much protease and the cost of ULP1 is inexpensive! For more information: http://www.enzymax.net/sumo%20protease.htm

By the way, GST loses its ability to bind Glutathione resin when denatured, it might not be a good idea to use strong denaturants such as guanidinium or urea in the purification buffers. Ni-resin should be ok to use with 6M Urea.

Good luck to your research!