It will work. But, ammonium sulphate does not precipitate target proteins selectively. It often co-precipitates other highly expressed contaminant proteins. You may need to optimize the concentration of ammonium sulphate for selective salting-out of your target proteins.

It is also worth to try other precipitating agents, such as sodium sulphate, PEG, etc...

In the presence of urea, the solubility of your protein will be high in solution. So you need to consider protein concentration and concentration of precipitating agents for selective salting-out of your target proteins.

Yes everything is possible but the question is: what for? In my mind few things can be done with such preparation.

Hi Vikas,

I have experienced a number of proteins that refuse to bind to affinity resin - sometimes the problem has been solved by changing conditions, and other times by modifying the expression construct to use a tag that is more tolerant of the conditions necessary to extract the protein from cells and isolate from lysate in the first instance.

Can you be more descriptive of what it is you have tried (protein pI, the affinity tags in the sequence, affinity resin used, buffer conditions in your binding attempts etc.)? This will allow those of us reading your post to offer more helpful advice based specifically on the evidence presented. Careful selection of the lysis buffer composition may also give conditions that facilitate isolation of your protein and removal of contaminants.

One idea that comes to mind - 8M Urea dissolves almost everything so this will be a very crude solution with significant contamination. If you firstly take your pelleted cells and lyse with a plain saline buffer to remove the easily soluble proteins, then spin at top speed (ie. 48,000x g for 30 minutes), you can remove the soluble proteins in the supernatant. Repeat this, collecting the supernatant each time. Then, use buffer containing 1M urea to extract proteins soluble in 1M, then 2M, then increasing amounts of urea until you attain 8M urea, each time collecting the newly-solubilised proteins and separating from pelleted material. When you have reached 8M urea, perform SDS-PAGE of the collected supernatants and identify which fraction contains your protein of interest and how clean it is.

This means that if your protein is only soluble in 8M urea, you have a chance to remove many impurities that are soluble in lower levels.

Providing the NaCl concentration is low (or 0mM) , ion exchange can be performed in 8M urea, with elution by addition of NaCl or change of pH, all still with urea present.

This is only a starting point of the discussion - if you can reply with more information about your existing conditions we should be able to work this out. Best wishes, Robin

@ Robin Dickinson,

Sir, Since I am getting my protein in insoluble form I am using denaturing condition to purify the protein of interest.

Affinity Chromatography

My protein has pI of 6.5, having N-terminal His-tag. I have tried binding at pH range- 7-8 with 20mM Imidazole, with additives NaCl upto 500mM, Glyceral upto 5%; and my protein start eluting in 50mM Imidazole. While increasing percentage of additive adversely affect binding of protein to Ni-NTA resin.

Ion-Exchange Chromatography

I have also tried anion exchange chromatography with DEAE spharose. I have tried binding pH range from 8-11. Washing with reduced pH and low concentration of NaCl (25mM) and eluted with 50mM NaCl or at reduce pH 7.0 with 25mM NaCl.

Hi Vikas,

>Since I am getting my protein in insoluble form I am using denaturing condition to purify the protein of interest.

Precisely. If you start by lysing the cells in a buffer that only mobilises the soluble proteins, you can separate these and remove them from your preparation to reduce the number of proteins present in your binding process ; then sonicate the pelleted proteins in buffers of increasing Urea content until your protein is solubilised under denaturing conditions. This would then mean that you may be able to remove the imidazole included in the binding buffer to block non-specific binding which in turn may give you a little more leeway for your own protein to bind.

I note that you observe that your protein isn't binding to the resin so well when you already have 20mM Imidazole present, and that elution occurs at 50mM Imidazole. This suggests that your protein isn't binding this resin effectively - it does happen with some proteins. I'm assuming that you're using Nickel resin - which resin chemistry are you using?

ie - Nickel-NTA, Nickel-IDA etc. - these can affect the way the proteins bind.

Secondly, have you considered trying resin equilibrated with Cobalt or with Zinc instead of Nickel? Bear in mind if you do use these that the binding // washing / elution concentrations of Imidazole are very different to those used for Nickel as these resins generally bind less background proteins and so require much less imidazole at the time of binding and elution is often substantially lower in the Imidazole gradient than it would be for Nickel.

Finally, if you are using Nickel resin, I would reduce the starting Imidazole concentration to 5mM instead of 20mM, and move the pH to 8 or even 8.5 for binding as pH7 and lower are often used as mild washing steps for proteins that bind with normal efficiency.

Please feel free to keep discussing this here.