It is important to know about the protein first. Also give the information about the source of protein, eukaryotic or proteolytic, the size of the protein.

By any chance is the protein a membrane protein? Check for the domains it has, using CDD or Pfam or any other.

I was wondering if the protein has Transmembrane domain or signal peptide in it. Check it here TMHMM http://www.cbs.dtu.dk/services/TMHMM/ and SignalP http://www.cbs.dtu.dk/services/SignalP/

Membrane proteins are bit tough to get in soluble form till you remove your transmembrane or signal peptide part.

I had the similar problem with a protein I was working on. Removed the signal peptide and transmembrane domain and voila it solved everything, got into soluble fraction and got purified as charm, got it crystallized it also.

If the protein does not have transmembrane domain or the signal peptide, then there are various good suggestions given about to try. I have some more on those lines that I have desperately tried and saw them working for my case.

I am not suggesting that now as I know its a lot of work in wet lab even if there's a small change you need to do. Been there seen that :).

Good luck,

Hi,

I suggest you to take a look at QIAexpress handbook. It may help you.

Good Luck.

I experienced a similar problem: insoluble protein whatever IPTG conc. and induction time used.

We solved the problem by building biomasse @ 37°C 'til OD 0.6-0.8, then iducing with IPTG 0.1mM at 12°C for 24 hours; I worked fine. Good luck!!

@thanks Jafar and Patrick....I am surely going to use your troubleshooting suggestions regarding protein solubility.

If proper folding of your protein depends on the formation of disulfide bonds, you can try the Origami or SHuffle strains (From Novagen and NEB, respectively). You can also try co-expressing chaperones (TaKaRa sells a chaperone plasmid set that is ideal for this purpose). Beware that soluble expression does not necessarily imply that your protein is properly folded, as soluble aggregates are not unheard of, especially when using chaperone coexpression. Also, ask yourself if it is reasonable to expect your protein to fold spontaneously or to not aggregate. It might require a folding partner, or be a truncated variant (or a member of a multiprotein complex) that exposes hydrophobic patches to the solven, even if correctly folded.

I have a similar problem in my protein expression, with different strategies for making it soluble; still it is in pellet! My target gene is in a PGEX vector and expressed protein has a GST tag!! Inevitably, I use of sarkosyl at high concentration (10%) for solubilizing the pellet and carry out a folding refolding processing. Take a look at this attachment.

Good luck

You can be do it by:

1)lowering the growth temperature. This decreases the rate of protein synthesis and usualy more soluble protein is obtained.

2) using a weaker promoter (e.g. trc instead of T7).

3)using a lower copy number plasmid.

4)lowering the inducer concentration.

I had the same problem for one of my recombinant proteins. Some times the proteins will end-up in inclusion bodies no matter what the incubation time or IPTG concentration is. For my protein, after bioinformatic analysis I have found out that the vector Tag (GST) have changed the recombinant proteins PI to 7.2. Surprisingly, the protein charged remained around 0 from pH of 5 to 10. Finally I managed to purify the protein from the soluble phase by raising the lysis buffer pH to 11. I suggest that you check your protein characteristics for similar issues.

Good luck!

You can also use the cell free expression in vitro system to produce small amount of proteins(less than 1mg), ref. Trends Biotechnol. 2005 Mar;23(3):150-6.

I had this problem before I also recommend you try to and refold it from the inclusion body with urea or guanidine first...

It is important to know about the protein first. Also give the information about the source of protein, eukaryotic or proteolytic, the size of the protein.

By any chance is the protein a membrane protein? Check for the domains it has, using CDD or Pfam or any other.

I was wondering if the protein has Transmembrane domain or signal peptide in it. Check it here TMHMM http://www.cbs.dtu.dk/services/TMHMM/ and SignalP http://www.cbs.dtu.dk/services/SignalP/

Membrane proteins are bit tough to get in soluble form till you remove your transmembrane or signal peptide part.

I had the similar problem with a protein I was working on. Removed the signal peptide and transmembrane domain and voila it solved everything, got into soluble fraction and got purified as charm, got it crystallized it also.

If the protein does not have transmembrane domain or the signal peptide, then there are various good suggestions given about to try. I have some more on those lines that I have desperately tried and saw them working for my case.

I am not suggesting that now as I know its a lot of work in wet lab even if there's a small change you need to do. Been there seen that :).

Good luck,

@ sanjiv kumar

My protein is bacterial protein and doesn't have any trans-membrane or signal peptide. I have tried with temperature, IPTG conc. and different bacterial strain but still it is coming in pellet.

please suggest me some more troubleshooting method for solving solubility problem.

Hi,

You can try to resuspend your pellet in 6M Guanidinium Chloride, with Beta mercaptoethanol. Then, after 1 hour of solubilisation, centrifuge at maximal speed for 30-45 minutes, and your protein will be in the supernatant.

Then dialyse your protein to renature it.

It is a good purification step

Perhaps ur protrein is using codons that are rarely used in e.coli.u can check rare codon usage using softwares. In my case i managed to improve the solubility by changing the rare codons to e.coli preffered codons (synthesize the gene)

Noor has a good point-- it is particularly important that at approximately the first 10-12 codons are those commonly found in your bacteria.

Another possibility would be trying auto-induction. The basic idea is that instead of using an inducing agent like IPTG you use the native function of the T7 promoter. So you use media containing glucose and lactose and grow the cells. As the glucose is depleted, the cells will slowly start activating their T7 promoters to start to use lactose in place of glucose. This will also slowly induce the promoters on your expression vector and lead to a much more gradual expression than you get from using IPTG.

My suggestions are

1. Try different constructs. N-term and C-term truncations have made all the difference on many crystallography projects. Look for info on domain boundaries, and try sequence alignments with any known homologs/orthologs.

2. Co-expression with any known protein partner(s)

3. Re-fold from the pellet and/or alter your lysis buffer.

Is your protein cytoplasmic or a membrane protein and what is the source? Assuming it to be cytoplasmic - some proteins are very lethal for cell that it can not afford keeping it in cytoplasm at any condition. you can try other strains of E. coli that can synthese lethal proteins (e.g., C41, C43, Plyse, etc). For me Low Salt Sorbitol Betain method just work fine. Even the most rigid ones come to soluble fraction. If you try 5ml induction , mainly you'll see protein in pellet but when you will try a liter prep. you will get protein from soluble fraction.

I am not much up to the bacterial protein but in order to deal with the similar problem in plant proteins, I followed a method where I can precipitate the protein by incubating the extracted protein supernatant in 0.1M Ammonium acetate over night at -20. Then following day I washed the pellet with chilled acetone. Then I found the solubility was really good for 1 and 2D. Also you can do a quality check by taking 1 uL protein solution prior to pelleting and simply blot it on a filter disc. Then stain with coomassie and see whether the solution what you are precipitating is having protein and approximately how much.

1.you check PI of your protein and your buffer should be compatible for protein PI

2. you better clone your insert fused to MBP tag ,this tag some times acts as a chaperon

3.you can tranform in to the ecoli strain containg chaperon plasmids like pkje7

4.while lysing your cells add 0.1-0.2% triton

5. you can add sarkosyl.

http://www.ncbi.nlm.nih.gov/pmc/articles/PMC50983

6. before induction at around 0.5 OD grow your culture at 18 deg for 1 hr and then add IPTG

1. You can try to reduce expression level, for example short after induction incubation, lower IPTG.

2. Different tag can be tried out, as suggested by Lalith

3.. If it is not the problem with buffer compatibility (your protein should have some charge in the buffer you are using), try to extract your protein form the pellet by reuspending in 8 M urea. Then reduce the concentration of urea step by step by diluting. It worked for me.

We had the same problem with some of our GST-constructs - they stay in the so called inclusion bodies. We now use a problem combining the advices of Lalith.

To solve this problem, we freeze the bacteria pellet over night (-80°C) then thaw on ice and resuspend in PBS before adding Xml (0.5%) of 10% sarkosyl, vortex and sonificate. Then we incubate for 15 min on ice before adding Xml (0.5%) of 10% TritonX. Incubate for 1 h on a rotating wheel before spinning down - then your protein should be in the supernatant. If there`s still a very viscid liquid centrifuge at 7200 rcf over night.

Good luck

You can also try to change your bacterial strain. I work with a protein that are expressed just in one kind of E. coli.

Another option is perform a refolding.

Very helpful is an intein tag (which in addition helps during purification by a chitin binding domain), which is completely removed later. See NEB manuals.

Do you Know the pi of your protein? The pH of you lise buffer must be at least one point up or down of the protein's pi. this part is very important, and what kind of buffer to use...

What is the purpose of the protein that you are isolating? When we isolate protein for the purpose of making antibodies, we pray it is insoluble. This makes the clean-up of the protein much easier and cleaner. We simply resolublize the insoluble pellet with denaturing buffer (which contains 6M Guanadine-HCL). We then run this denatured protein through our Talon Column (His-Tag binding). Once we have the purified His-Tagged protein we step wise dialyse the protein back into native buffer so that the protein can slowly refold. (6M, 4M, 3M, 2M, 1M, 500mM, 0M guanadine)...This works very well for the purpose of antibodies, and we almost always end up with soluble protein at the end. Occasionally though certain proteins (like our viral coat protein) will re-precipitate between 500mM and 0mM, as the characteristics of this protein is to form multimers with other coat protein molecules. If the purpose of your experiment is to have active functioning enzyme after isolation, then this method is a bit of a crap shoot, as it will totally depend on whether your protein is able to refold into a conformationally active protein all on it’s own. Hope this helps…good luck!

Are you sure that your protein is found in inclusion bodies and doesn' t aggregate after cell lysis? In several cases protein aggregation during cell lysis can be prevented by adding stabilizing additives (e.g. L-arginine, trehalose) prior lysis.

Good luck

If you tried all the suggestion given by expert here and still not getting the protein soluble, You can try with sarcosyl (Sodium lauroyl sarcosinate ) Just you have to optimize the different percentage of sarcosyl starting from 0.1% - 0.5%.I am sure this will work just you need some patience and optimization.add sarkosyl while lysis.

Good luck.

The best way to make the protein soluble is to play with the PH for example increasing the ph of protein sample to 9 will increase its solubility try to titrate NaoH 0.1M etc

Best wishes

Mohammed

If its possible for you ,clone your gene next to a thioredoxin tag. It helps you to suloblize your protein. I think expression of recombinant proteins in rosetta gami strain of Ecoli influences on solubilizing the expressed protein.

This might be a dumb question, but is the protein denatured?

i think you study the hydrophobic and hydrophilic regions on the protein and charge ions

If you're interested in the natured state of the protein, you might be having problems if the protein is conforming to sequester charges from the solvent. Though that should make it more soluble, atleast in theory. Or there could be protein-protein interactions that are producing michelles which could decrease solubility as the aggregate grows. Have you tried running a western to see what sizes proteins you are getting? Or am I completely off base here? Is there anyway you could resuspend the pellet in a suitable solvent, salt out the peptide and do an A/B extraction? Or maybe even thin layer might be an option..