Calculate the stoichiometry of binding that you measured in moles of ligand bound per mole of protein. If the protein is an oligomer, do the calculation on the basis of protein monomers.

Suppose the protein is a monomer and has a single binding site for the ligand. The stoichiometry at saturation is therefore equal to the fraction of the protein that retains ligand-binding capability.  This is the simplest case.

The first complication to consider is that there could be both high-affinity and low-affinity or nonspecific ligand binding to the monomer. This should be apparent from deviation of  the binding data from a single-binding site model (Langmuir isotherm).

Further complicating this analysis is the possibility of cooperativity of binding sites in an oligomeric protein. The Hill equation can be used to fit data to a cooperative binding model. There could also be multiple non-cooperative binding sites with different affinities, for which there are also suitable equations. Non-specific binding usually looks like a linear binding curve superimposed on a saturable binding curve, so it is possible to extract the saturable part of the binding from the data using a suitable equation.

Please consult Chapter 4 in the book Enzymes: A Practical Introduction to Structure, Mechanism, and Data Analysis by Robert A. Copeland for equations.

Unless you know the precise behavior of protein of your interest in its native state versus detergent solubilized (quite possibly with and without other contaminating cellular components to varying degree), it is unlikely that you will be able to accurately quantify the amount.

If on the other hand, if someone has already established a relationship (even an empirical one), just follow their publication.

Best,

If I understand correctly, your initial estimation of 'receptor' density is calculated for crude protein which means that it is a mixture of proteins so you can not determine what percentage of your target protein is actually binding.  The same would be true for your detergent purified protein.  You would need 'pure' target protein and then determine the receptor density of that and then follow Adam Shapiro's suggestions. 

Hi,

Thanks everyone for response. I think I can easily estimate activity from purified protein. 

e.g. 

Bmax 1000 pmol/mg (calculated from saturation binding curve)

First I need to convert pmol to mass 

Protein Mol.Wt  = 50kDa

(1000 x 10^-12 mol x 50 000 g / mol) = 50 microgram

Total amount used in the assay approx 50 microgram

(50/50)*100 = 100%

I can do similar estimation for unpurified protein if I quantify my target protein using WB. 

Does this make sense? 

In your Bmax of 1000 pmol/mg, I would expect that the numerator (1000 pmol) refers to the amount of radioligand that binds to the protein, and the denominator refers to the mg of protein. If that is true, then it is the mg of protein that should be converted to moles. Hence:

1000 pmol ligand/mg protein = 1 µmole ligand/g protein

(10-6 mole ligand/g protein) x (5 x 104 g protein/mole protein) = 0.05 mole ligand/mole protein.

In other words, only 5% of the protein has ligand bound at Bmax.

Assuming all of this binding is specific to the protein, and the protein is 100% pure, only 5% of the protein is capable of binding ligand. If there is non-specific binding, the % active protein could be lower. If the protein is impure, the % could be higher.

Shahid, you are making too many convenient assumptions without evidence, unless you are not posting additional details I alluded to earlier which support the idea that your assumptions are quite valid.

I agree with Adam B Shapiro answers ...good luck.