What, precisely, is your aim? Do you just want to see the protein in the gel (e.g., for quantification), or do you want to purify the active enzyme (e.g., for kinetic measurements or industrial use)?

In the former case, I'd start with 7 M urea, 2 M thiourea, 10 mM DTT and 1%SDS. 

In the latter case detergents with fat-like structure (lyso-lipids, fatty acids, FOS-choline, LysoFos, neopentenyl glycols) may be able to to solubilise functional protein. In any case, I'd start with a careful literature search. A day in the library can save you months in the lab!

Hi Rahul, try putting the sample (with detergents) into a sonicating water bath for ~ 1 min before loading on a gel.

Dear Rahul, here few suggestions you can give a try.

1 Incubate your protein with low concentration of urea or guanidine Hcl for overnight.

2 Run SDS PAGE

Hi Rahul,

I think the main issue is not how to visualize the protein on SDS-PAGE, but rather how to reduce or prevent aggregation in first place. Considering all you already tried it seems you are getting very large and stable aggregates, suggesting the protein in unfolded or badly misfolded; I wouldn't bother any further to try and rescue this material. First, you should check that the aggregates you are seeing are actually formed by your target protein, e.g. by Western or dot blot. Following, you should try to improve expression of the protein and try to move it to the soluble fraction. If you provide information on how you are currently obtaining you enzyme I or others can be more specific.

Good luck!

Carlo

Thanks Dr. Carlo and everyone.

The aggregates which are forming includes my protein of interest as well as other proteins in the supernatant. I think, the protein it self is being produced in the production medium in aggregated form which i confirmed by directly loading crude supernatant on gel filtration chromatography. The active fraction along with other protein is coming out in void volume. Same is happening with ammonium sulfate or acetone precipitate may be due to high molecular weight of aggregated proteins.

As I know from the literature, lipases are mostly hydrophobic (ammonium sulfate ppt at 40 % saturation only in my case) in nature and that is why forming aggregates with other protein.

Currently, I am producing the enzyme from a wild Pseudomonas strain in lipase production medium with 1% olive oil as lipase inducer followed by precipitation of protein. 

What, precisely, is your aim? Do you just want to see the protein in the gel (e.g., for quantification), or do you want to purify the active enzyme (e.g., for kinetic measurements or industrial use)?

In the former case, I'd start with 7 M urea, 2 M thiourea, 10 mM DTT and 1%SDS. 

In the latter case detergents with fat-like structure (lyso-lipids, fatty acids, FOS-choline, LysoFos, neopentenyl glycols) may be able to to solubilise functional protein. In any case, I'd start with a careful literature search. A day in the library can save you months in the lab!

Dear Sir,

My final aim is to purify the protein (obviously visualization as single purified band in SDS PAGE for molecular weight determination). In first case as you explained, I have tried SDS up to .5 % and urea up to 4M which caused smear formation in gel might be due to denaturation, and also activity is lost (hence activity stain and protein band of interest could not be detected). The problem what I am facing is that, during native PAGE analysis (w/o any detergent) i am getting two active bands one at gel interface and one around 30KDa. While running the same protein sample in size exclusion (sephadex G100 and G200) for purification, all the protein is being collected in void volume. I have tried the same with SDS as well as CHAPS added in the equilibrium buffer and sample, but did not get success. The problem is to keep the protein in free state for purification to achieve.

I will try surely with detergents what you have suggested.

When you did the electrophoresis of the urea denatured sample, did you add urea to the gel matrix as well (you should). Zymograms will usually not work after urea because urea is a denaturing agent.

When you did the gel filtration of the detergent solubilized enzyme, did you have detergent in the elution buffer? Did you equilibrate the column with the detergent-containing elution buffer for at least 24 h before injecting the sample? Because of micelle formation it does take that long!

Apart from these technical issues: Have you considered the possibility that your protein may form oligomers in its native state? If your monomeric mass is 30 kDa, then, say, a pentamer would be 150 kDa and come out with the void on G100 and close to or with the void (depending on Stokes-radius) on G200. On native PAGE some oligomers may stick together, while others break apart into monomers, which later, when you perform the zymogram, may reassemble. 

By the way, Sephadex is a very old chromatographic matrix, you may wish to consider for example Sephacryl for better resolution. Given the possibility of oligomer formation, I'd try Sephacryl 400 or even 500 and see what happens.

Till now I did not consider this possibility but I think this could be a reason behind the whole problem which til now I did not understand.

I did SDS-PAGE (non-reducing) with my sample followed by development of zymogram after removing sds from gel (using Triton-X). I got very sharp bands of my enzyme which were not aggregated. So formation of oligomers in native PAGE might cause trouble during separation (and also during gel filtration) while SDS cleaved the oligomer and I got a clear band.

So, If this is the case, can you please suggest me in detail how do I proceed with my experiments to purify my enzyme?

As I mentioned before, you may be looking at an association-dissociation equilibrium of your protein. The first question then is whether or not you need dissociating agents like urea or detergent in your buffer. If the oligomer can be purified without them, do so and use, say, Sephacryl 500 for SEC. This way, you do not have to worry about reconstitution and loss of activity incuced by the dissociating agent(s).

If you need dissociating agents to separate your proteins from contaminants, then just deal with the added complication. In this case, your running buffer for SEC needs to contain your detergent at the cmc, and you have to equilibrate the column o/n at moderate flow rate (you can pump the buffer in a closed loop). If you also use urea for solubilization, it also needs to be present in the running buffer. In this case you'd go for the monomer, and hence use, say, Sephacryl 100. Note that Sephacryl is a tradename of Pharmacia LKB (now GE), other companies use different names for similar matrices. Once you have purified your monomer, you need to remove the denaturant(s) to allow re-association. Urea can be removed by dialysis or desalting, detergents by dialysis (for detergents with high cmc) or Biobeads SM-2 (for all detergents).

If you need to read up on detergent use in protein science or on chromatography, you could for example read the relevant chapters of Buxbaum 2011. Btw, SDS does not chemically cleave proteins (as, say, proteases would do), it merely dissociates protein-protein interactions.