I'm trying to bind protein to the surface of nanoparticles which have carboxylated previously and activated by EDC-NHS. I use 0.2-3 mg/mL protein concentrations for 2mg/mL of nanoparticles. How can I find out the quantity of protein binded to nanoparticles?
(I have tested Bradford assay, but I think it is not suitable for this concentration values.)
Hi Sayed, do you mean that the proteins bound to your nanoparticles are below the detection limits of the Bradford assay?
Bradford should be good up to 0.1-0.2 mg/ml protein.
If you have less protein, then fluorescence assay using fluorescamine or FITC might be the way to go.
BTW, if you have carboxylated nanoparticles, you can incubate them with EDC at pH 5 and then add protein.
The NHS ester formed through EDC activation of carboxyl groups is very labile and only lasts for minutes at pH 7.
If you can separate the nanoparticles from the solution by centrifugation or ultracentrifugation, you can measure the amount of protein bound by measuring the reduction of the protein concentration in the supernatant using any convenient protein assay, such as UV absorbance, Bradford, BCA, Lowry assay, fluorescence-based method, SDS-PAGE, etc. If the protein is too dilute, you can concentrate it by TCA precipitation.
Thank you very much for your valuable answers, but as I told I have tested UV absorption and Bradford. my protein concentrations is in the range 0.2-3 mg/mL. Bradford is not linear at this range and also sample dilution may be not correct. UV absorption gives me unacceptable results too. I'm trying SDS-PAGE now...
Hi Sayed,
If the volume of your sample is sufficient to take 10 microliters for the Bradford assay (in triplicate, preferably), then 0.2-0.3 mg/ml should be concentrated enough for the Bradford assay. The Bradford assay can be used nicely with 1-5 micrograms of protein in a 200 microliter assay volume in 96-well clear plastic plates.
0.2-3 mg/ mL is more concentrated than the linear range of Bradford. so I must dilute my samples if I want to use this assay. But here is the problem that the dilution does not affect linearly in the Bradford results. I mean if I dilute them 10 times the absorbance of protein-reagent at 595 nm may not decrese 10 times...
I always find with the Bradford assay that the standard curve with BSA is parabolic and fits well to a 2nd order polynomial over a certain range, say 0-5 micrograms, then flattens out. So it may be necessary to test a set of dilutions to make sure that at least one dilution is in the dynamic range of the standard curve. I recommend using 2-fold serial dilutions.
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