Keep things as simple as possible when starting out. I have worked and am working with difficult proteins and here's what I do. Here is what I do if I KNOW the protein is going to be hard to isolate. I would know this because I would have tried doing things the easy way by expressing at 37C.

1.) Pick a low copy vector to clone into ( If there are codon issues then use Rosetta II cells otherwise use BL21).

2.) Transform

3.) Pick 20 or so colonies and grow an overnight culture (if you pick one colony it may not be a good overproduced so this gaurantees you are always getting the average)

4.) Inoculate 50 mL culture with 1% of overnight culture. (here you may want to try different type of media like TB and NZCYM or autoinducing media)

5.) Induce at ~0.6 O.D with 0.1 mM IPTG (make sure you cool down the bacteria to the desired temperature before inducing)

6.) Grow at 16 C for 24 hours

7.) Isolate bacteria and sonicate a couple of times in different buffers contain different amount of salt ( 0 - 500 mM NaCL). If protein pI is 7.4 then got AT LEAST one unit above or below in your buffers. So pH to 8.4 or 6.4. Also include 10 - 20% glycerol. Also, always use reducing agents. I use 10 mM BME.

8.) Spin down and run a gel of supernatant.

9.) If you get nothing then you are probably getting inclusion bodies and there's little you can do except go into a different expression system of use a different vector. For example, use pSUMO vector or express in yeast.

All my proteins are annoying but I got all of them expressing by following this sort of workflow.

I use TB for media because it is pH'ed at 7 so your cells are good and happy for 24 hours of expression. LB is not pH'ed and will become acidic from acetate that e. coli produce.

use poly buffers and polybuffer exchangers in between the range of near pH to your enzyme.

Dear Morey,

We have some point to consider:

1) The pH of the medium should be not modified, because it is the optimum for the bacterial multiplication. Moreover, it isn´t the same of the interior of bacterial cell.

2) The more time, the better the folding. Do not try to grow the bacteria fastly.

3) As you know, the bacterias do not have organelle, so that they never can fold well complex proteins. 16 cys it is really a lot of! I recommend to express in some yeast, like Pichia pastoris. You can try to fold it in vitro, but it is a big challenge.

Good Luck!

Att.,

Angela.

Thanks a lot for the answers!

@Angela: How do you know if the disulphide bonds would be formed between the correct pair of Cys residues? For e.g. the crystal structure of this protein shows only ONE disulphide bond while the rest of the Cys residues are free! Would a reducing environment (bacterial cytoplasm) be more suitable for such proteins than an oxidizing environment (periplasm)?

Thank you for your fast reply. You have luck it haven't 8 disulfide bonds. In any case, a reducing environment is more adequate. Nevertheless, the cytoplasm is not so recommendable. Normally, the proteins form inclusion body in the cytoplasm, but can fold well in periplasm. The best way is express it in secreted form, in this case you need a signal peptide. (Other advantage is that you have very low contamination with bacterial proteins in secreted medium).

Another way is to treat the aggregate with urea and DTT or beta-mercaptoethanol, in order to bring it into solution, and after it you have to fold your protein. In the literature you can find same conditions to fold. You have to do a screen: Take only same mL of your sample and try various solutions, time and temperatures.

The protein I'm working on is a large 66KDa protein, and I wouldn't prefer trying refolding experiments with it. I am thinking more on the lines of using a solubility tag, targeting it to a periplasm or even easier, using competent cells that support disulphide bond formation.

Have you used the SHuffle T7 competent cells from New England Biolabs for expressing disulphide protein?

For the induction you do not need to use1mM IPTG. Following induction for 1 -3 hours, collect the bacteria, resuspend in 1/10 volume of 140 mM NaCl, 0.5mM EDTA, 20 mM Tris-HCl pH 7.8, 0.05-0.5 per cent Tween 20 or Nonidet NP-40 and freeze at -20C. Next day, thaw the bacteria and sonicate at 30 sec burst to clarity. Keep the bacteria suspension at all times on ice. Centrifuge to collect the clear supernatant and use your purification on Affinity column of choice. If the volume of your culture is large, use 45-50 percent ammonium sulphate precipitation followed by dialysis to remove the salt. You may need another centrifugation to remove any precipitate formed during dialysis. If needed, you could use 1mM DTT in your buffer which is better than mercaptoethanol. Do not forget to check the induction of your protein on SDS using appropriate controls. The induction has to be titrated using various molars of IPTG for optimization.

best of luck

Have you check if your protein is expressed in inclusion bodies? Is your protein His-tagged?

Keep things as simple as possible when starting out. I have worked and am working with difficult proteins and here's what I do. Here is what I do if I KNOW the protein is going to be hard to isolate. I would know this because I would have tried doing things the easy way by expressing at 37C.

1.) Pick a low copy vector to clone into ( If there are codon issues then use Rosetta II cells otherwise use BL21).

2.) Transform

3.) Pick 20 or so colonies and grow an overnight culture (if you pick one colony it may not be a good overproduced so this gaurantees you are always getting the average)

4.) Inoculate 50 mL culture with 1% of overnight culture. (here you may want to try different type of media like TB and NZCYM or autoinducing media)

5.) Induce at ~0.6 O.D with 0.1 mM IPTG (make sure you cool down the bacteria to the desired temperature before inducing)

6.) Grow at 16 C for 24 hours

7.) Isolate bacteria and sonicate a couple of times in different buffers contain different amount of salt ( 0 - 500 mM NaCL). If protein pI is 7.4 then got AT LEAST one unit above or below in your buffers. So pH to 8.4 or 6.4. Also include 10 - 20% glycerol. Also, always use reducing agents. I use 10 mM BME.

8.) Spin down and run a gel of supernatant.

9.) If you get nothing then you are probably getting inclusion bodies and there's little you can do except go into a different expression system of use a different vector. For example, use pSUMO vector or express in yeast.

All my proteins are annoying but I got all of them expressing by following this sort of workflow.

I use TB for media because it is pH'ed at 7 so your cells are good and happy for 24 hours of expression. LB is not pH'ed and will become acidic from acetate that e. coli produce.

I should have said TB is buffered at pH 7.

expression of soluble protein or IBs depends on protein nature and some fermentation conditions, the higher temperature and IPTG concentration the higher percantage of IBs formation therefore if you want your protein in soluble form then you should try lower temperature and lower IPTG concentration, other words you slow down expression and cells have time to fold your protein but if you have over-expression you will get IBs because the cells do not have time and resources to fold your over-expressed amount of protein.

E.coli does not produce acetate on LB or TB mediums because it is a protein medium without sugar, acetate is product of over-oxidation of sugar therefore no sugar no acetate, but if you add glucose then you will get acetate and lower pH that even might stop growth of bacteria.

The TB medium has higher concentration of protein nutrients and results in higher cell yield that proportionally should give you higher yield of your protein escpecially if it is in form of IBs, if it is a soluble one then the highest yield is function of time especially if it is enzyme and for soluble protein you should find the optimum time if you want to do it, typically 4 hrs is a standard time, for enzyme the time might be even lower, around 2 hrs.

Thank you all for your replies and detailed procedures!

I tried changing the lysis buffer pH in the range - 6.0 to 7.5, but this did not seem to help. The existing literature shows that the protein was purified AT pH 8.0!! (although they produce it in insect cells). So, I guess pH shouldn't make a big difference to protein solubility. Also, I tried reducing the induction temperature to 16˚C and IPTG concentration to 0.1mM but it didn't show any detectable increase in protein solubility.

I am now using a HIs-tag staining kit for gels, so that I can visualize the protein on the gel directly. With this I see that some amount of the protein is soluble though majority is in the pellet. I am now trying to purify it with Ni beads to see if I can separate the protein from the impurities.

Here are some more questions:

Would adding sorbitol or glycerol/ ethanol/ betaine HCl in te growth media help solubilize the protein?

I think that could be informative for you to know if your protein is in the inclusion bodies or if it is insoluble for his pI, because you should follow two different strategies. http://kirschner.med.harvard.edu/files/protocols/QIAGEN_QIAexpressionist_EN.pdf

Hi,

from my experience, induction at lower temperature (16-18°C) and addition of sorbitol and betain keep the protein solubilized.

However, if you have soch troubles, have you considered other system, where you could use other pH?

Or add larger tag, so that it keep your protein soluble.

Hi everyone,

Thanks a lot for all your replies.

I am finally able to purify my protein of interest from the soluble fraction using Ni beads although I use no imidazole in the binding buffer (as low as 8mM Imidazole causes its elution). Even though I get a lot of non-specific binding I am able to further purify the protein using ion exchange and gel filtration.

I am following the purification using a His-tag in-gel stain that stains His-tagged proteins. However, I am concerned about the low affinity of my protein for Ni beads. Especially since I am counting on the His-tag in-gel stain and the Ni affinity, both of which use the same procedure for detection. Is it possible that a bacterial protein with a chain of HIs residues might also give the same result?

I have attached a gel of the Ni affinity purification with the following samples in the lanes. Binding buffer has no imidazole and elution buffer has 200mM Imidazole.

1. Pellet

2. supernatent

3. Flow through

4. Eluate - 1

5. Eluate -2

6. M.W Marker

I read that you are using Triton X-100 in your buffer. Even though, it is non-ionic it is not the best detergent in buffers for the purification of proteins. I suggest that you should try 0.5-1% of Nonidet NP-40 or Tween 20 as detergent in your column buffers. You can also use KCl instead of NaCl and increase the salt concentration to 140mM (physiological). The other change I suggest is to use Hepes buffer which has a higher pH stability than Tris/HCl.

If you would prefer to stick to the conditions you are working on, dialyze the eluates to remove imidazole and reapply on to the regenerated column to perform a second purification to eliminate the contaminants.

As seen from your gel, you have good starting material. Did you calculate how much of it is in eluate 1 and 2 (% of the total)?

If the disulfide bonds play a role in aggregation, you should observe higher mwt products even on your SDS-PAGE.

Finally, the purification can be optimized by using a linear gradient of Imidazole than a step wise elution. This will eliminate additional unspecific factors enriched on the matrix.

Is all your protein eluted with 8 mM imidazole? In such case, you can elute it with only say 10 mM imidazole and the rest of bound proteins elute independently in other fraction.

Hi everyone,

Thanks a lot for your advice and comments.

I was finally able to express and purify the protein, so I thought I'd share my experience for others who might be facing similar problems.

I checked the literature for purification of this protein and found that pH was not the problem. The problem was probably the solubility - most of the protein was going into the pellet, and yield - the little protein that was soluble was too low to be distinguishable from the bacterial proteins.

My first goal was to have maximum possible expression while keeping majority of the protein soluble. I reduced the induction temperature to 20˚C and inducer concentration to 1mM IPTG in order to get most of the protein soluble. After Ni affinity purification, I could see a band at the desired M.W., however, there were still a lot of protein contaminations and I wasn't sure about the presence of my protein. I had to do a 2nd step of purification. Since my protein was prone to aggreagation, reducing the NaCl conc for ion exchange resulted in precipitation and low final yield. Using gel filtration wasn't able to reduce the contaminants and led to further dilution of the protein.

In order to remove the unspecific binding, I used lower amount of beads and increased the imidazole conc in the wash buffer to 40mM. Also, I did a batch elution with multiple bead volume elutions. After this, my protein was enriched and the contaminants were considerably reduced. After that I used gel filtration to further purify the protein.

I was using LB throughout, and then tested differernt media and conditions to improve the yield further. Using TB turned out to be a better alternative giving twice the yield and the same purity in the end.

Cheers!

Just a comment about pI - the calculated pI from the sequence does not actually have to correspond to the actual pI, which can only be determined by iso-electric focussing.

I have worked on a protein where the calculated pI differed by 2 pH units from the actual one. So don't draw too many conclusions from the calculated pI, just use it as a rough guess of the actual one.

Yes, that is sort of true for my protein as well. Besides, even at pH = pI, there could be localized charges leading to certain parts of the protein having a positive/negative charge.

Inaki, no its not any nucleic acid binding protein. That publication was from an earlier lab I worked at. :P

Thanks for your overview. I may find it useful for my purification also. Can you clarify the following for me?

1. At what imidazole conc did the protein elute?

2. What is a batch elution with multiple bead volume elutions?(can you specify the conc imidazole also?)

Thanks

Also:

3. Did you do anything about the aggregation? What pH buffer are you storing it in?

4. Do you still think you had purified and artifact or was that the protein itself?

Hi Nandita,

1. I used a batch purification and my protein eluted with 200mM Imidazole. The exact concentration of elution might be between 100-200mM Imidazole. However, this depends on the protein you are studying, and doesn't have much correlation with protein pI.

2. I used 2x bead volume of elution buffer or more, and repeated this 3-4 times.

3. I'm using pH 8 buffer, and my protein is fine in it. I used this pH because it was mentioned in the literature. Later, I also found that HEPES buffer was better suited for my protein.

4. I am sure it is my protein because it interacts with another protein known to be its binding partner. If its an enzyme you are purying, its easier to detect the activity.

Message me if you have any other doubts or want to discuss about your purification in detail.

Thanks! will do if I am stuck

Hi everyone!

I know I am late in this topic, but I just want to share my experience in proteins purification. In our lab we used to purify a lot of different enzymes from a crude extract, so to perform the process in a easy way we start to use isoelectric focusing, to do that we bought the rotofor system from BioRad

Try to reduce the IPTG concentration (0.1mM enough) , and reduce the temperature after induction it may reduce the inclusionbodies formation. your buffer is good nothing to worry about that. if you still get the problem you can go with detergent washes like sodium deoxy cholate and solubilize with urea or guanidine hydrochloride. then refold it with Argenine.