Hi Saranpal! In principle both ways to measure protein concentration should be fine. However, there are some things to consider. At very low protein concentrations, I found that any protein concentration determination techniques tends to have a higher error. Also, some specifics about your protein can influence the results. For example if you have reducing agents or detergents in your sample this can significantly change the read-out. For Bradford, you also have to take into account that the colored complex only has a limited life time (I think it's on the order of 30min to 1hr). And you always have to be careful about your calibration standards.

In principle, I personally like measuring OD280 (when possible) as it is a very direct read-out (unless nucleotides or detergents or a very low extinction coefficient chane this). But having said that, I found that sometimes errors of up to 2x difference between techniques exist... My preferred workaround is to stick to one technique religously, so that I can compare prep to prep. And finally, if you run a gel with different dilutions of your protein (and a comparison gel with different, known concentrations of a standard protein), the band intensity can help you decide which assay to trust more in your specific case. I hope this helps some! Good luck!

Hi Saranpal! In principle both ways to measure protein concentration should be fine. However, there are some things to consider. At very low protein concentrations, I found that any protein concentration determination techniques tends to have a higher error. Also, some specifics about your protein can influence the results. For example if you have reducing agents or detergents in your sample this can significantly change the read-out. For Bradford, you also have to take into account that the colored complex only has a limited life time (I think it's on the order of 30min to 1hr). And you always have to be careful about your calibration standards.

In principle, I personally like measuring OD280 (when possible) as it is a very direct read-out (unless nucleotides or detergents or a very low extinction coefficient chane this). But having said that, I found that sometimes errors of up to 2x difference between techniques exist... My preferred workaround is to stick to one technique religously, so that I can compare prep to prep. And finally, if you run a gel with different dilutions of your protein (and a comparison gel with different, known concentrations of a standard protein), the band intensity can help you decide which assay to trust more in your specific case. I hope this helps some! Good luck!

Bradford assay is the most common assay for protein quantitation. Depending on expected concentration of your sample, you need to choose between a Micro Assay or a Macro Assay. I would make sure the r2 value of the standard is no less than 0.98 as values less than this makes big differences in accuracy.

I suggest you to read the paper from Zor & Selinger (Linearization of the Bradford Protein Assay Increases ItsSensitivity: Theoretical and Experimental Studies.

Tsaffrir Zor and Zvi Selinger, ANALYTICAL BIOCHEMISTRY 236, 302–308, 1996). The difference is using the ratio of the absorbances 590 nm over 450 nm, linear with protein concentration.

I use Bradford or BCA if I have enough sample volume. I will use NanoDrop only I have a very small amount of lysate. Actually, using exactly the same lysates to measure twice the results will be different. But, as long as it is a systemic error the usually the end results should not be too off.

The UV280 (NanoDrop) should be used only for purified enzyme. If you have anything like crude extract, *everything* in your sample will absorb.

Hi Saranpal,

unfortunately you didn't provide informations about the buffer your protein is solved in, if you have a purified protein or a mixture and which reference protein you used for the Bradford assay.

These points would be crucial to give you good advice.

Just some general considerations: As Paul Lesbates already pointed out the composition of your protein is very important in respect of Tryptophan and Tyrosin, but since you used the specific. Ext. Coefficient this should be fine.

Have you tried to dillute your protein (1:2, 1:5, 1:10) to see if you are in the linear range of absorption detection? If you have detergents or other interfering components in your sample, these might result in seemingly higher protein concentrations, but this should be sorted out be the use of a correct blank for both UV280 and Bradford.

For the Bradford I would also recommend to use dillutions of your sample to check for measurements within the linear range of your assay and detection system. Also make sure to measure the Bradford as soon as possible, since your protein/Dye complex will precipitate within a quarter of an hour.

If your protein is unusual in respect of aminoacid composition (very hydrophobic, many/view Trp, etc.) the right reference protein might also be an issue, since BSA not allways is the right choice.

But as said before, more informations about your sample would help to give the best advice.

Best,

Kai

Thank you everyone for your insightful thoughts and detailed explanation pertaining my query. It was indeed helpful!!

Paul, thanks for the link to Bitesizebio, I must have missed out this article though I am a frequent visitor/reader of bitesize bio.

To answer question raised by Kai:

Yes, I have purified my protein and my protein is dissolved in 50mM Tris-HCl supplemented with 100mM NaCl (Imidazole has been removed/dialyzed) calibrated to pH7.5 and I have used the same buffer to dilute the Bradford Dye in 4:1 ration (Buffer to dye ratio). Yes, my protein is a hydrophobic protein as it is a rubber membrane protein. For this reason, I have actually fused my protein to a larger tag system (namely the Nus A tag) to obtain a highly soluble protein. Therefore, the ext. coefficient value that I computed into the NanoDrop is the value of my protein fused with Nus A tag.

@ Saranpal

If you can dilute your protein and still be in the linear range, I recommend diluting it with Guanidinium hydrochloride prior to A280 readings. The experimental extinction coefficients are closer to the theoretical values if your protein is unfolded and the aromatic groups can rotate freely in solution. Just blank with the same ratio buffer/Guanidinium hydrochloride.

Hi Stefan,

Thank you for your suggestion. Could you also kindly suggest what should be the molarity of the denaturant to dilute my protein? Also, is it possible if I were to use urea instead of guanidium hydrochloride?

Thank you.

8M Urea or 6M guanidinium are usually cited. I never used urea so I can't tell but I see no reason why you couldn't.

I have the same problem of having two different measurements of the same recombination protein sample before and after Triton X-100 treatment. 

You are advised to proceed with caution when measuring the concentration of protein in Triton X-100, as this non-ionic detergent is capable of absorbing UV at 280nm. 

what if the recombinant protein contains several disulfide bonds.  The chaotrophic reagent can also helps to unfold the protein in this case? so that we can still use predicted extinction coefficient from Expassy ProtParam to calculation of concentration?

The UV-Vis-based method of protein quantitation relies on absorbance at 280 nm by a protein’s aromatic amino acids, predominantly tryptophan with minor contributions from tyrosine.Therefore, protein extinction coefficients can vary widely (greater than two-fold difference between extinction coefficients of albumin and immunoglobulin G). Those proteins that do not contain tryptophan, such as Protein A, are very difficult to quantify accurately based on 280 nm absorbance. Also, secondary, tertiary, and quaternary structure all affect absorbance, therefore factors such as pH, ionic strength, etc. can alter the absorbance spectrum.

I prefer the FTIR quantitation (Direct DetectTM ) detecting amide bonds, which are present in all proteins. Although protein structure influences the exact wavenumber of the Amide bond maximum, the intensity of this band can be used to accurately assess concentration.