Title: Learn: Bradford Assay
Course: BTEC-2020-301-302-303-F21 > Section: Protein Gels

Module2 Bradford Assay

You have now separated proteins by mass using SDS-PAGE. For that gel, we told you how much to load. But how would you know how much protein to load on a gel if it isn’t stated? If you use too little, the bands may be difficult to see; too much, and blobby bands may overlap each other. The volume of protein sample needed depends on the concentration of the protein in your sample.

How do you determine protein concentration? One option is a direct measure of absorbance at 280 nm via spectrophotometry. This is very useful for pure protein samples, if you know the extinction coefficient of the protein in question. (Recall that A = C*l*E, meaning that absorbance is dependent on concentration, path length, and extinction coefficient.) The extinction coefficient describes how a particular substance absorbs a particular wavelength of light. Proteins vary widely in their extinction coefficients, so it is necessary to know the E value to get a good estimate of concentration. Another limitation of direct measurement of absorbance is that it isn’t very sensitive; it isn’t good for concentrations of less than about 0.1 mg / mL. Finally, absorbance from other compounds at 280 can interfere with getting a good estimate of concentration.

Another common way of measuring protein concentration is a dye-binding assay, such as the Bradford assay. In the Bradford assay, you add a particular form of Coomassie blue to the proteins. (You’ve already encountered Coomassie blue when staining your SDS-PAGE gels; it’s the blue dye that lets you see your bands.) The Coomassie used in the Bradford assay starts out a reddish-brown color, but when it binds to protein, it turns blue. Coomassie binds to most proteins with approximately equal affinities. The bluer the solution turns, the more protein it contains. An advantage of the Bradford assay is that it can detect protein levels that are 10-fold lower than that required for direct absorbance. It also does not require knowledge of the extinction coefficient, and is less sensitive to interference from other compounds.

To perform a Bradford assay, you set up a series of standards - samples of known concentration over a wide range - and compare the dye binding of the known concentrations to that of your unknown sample. In this experiment, you will set up standards that range from 0 - 300 ug/mL of BSA (bovine serum albumin, an inexpensive protein often used as a standard). You will also set up several dilutions of your unknown protein. It’s important to set up several dilutions of the unknown, because you need at least one of the dilutions to fall in the range of the standards. You will add the diluted Bradford reagent to each of the samples - standard and unknown - and allow them to incubate 10 minutes. You will notice a color change occurs in the cuvettes. Your standards should have a smooth gradient of color from reddish-brown to dark blue, as shown in the picture.

Bradford assay standards should demonstrate a clear progression of color from reddish-brown (no protein) to bright blue (lots of protein).
Photo by doroquez.

You will use a spectrophotometer set to 595 nm to measure the amount of blue dye in each sample. The data for your standards will be graphed with BSA concentration on the x-axis, and absorbance at 595 nm on the y-axis. The data should be linear. Once you have the equation of the line that describes the data, you can use that equation to determine the concentrations of your unknowns from their absorbance.

A Bradford assay standard curve should show a linear relationship between protein concentration and absorbance at 595 nm. Only the standards are graphed on the standard curve, not the unknowns.
Image: Jean Bower.

The Bradford assay is a very useful tool for estimating the concentration of proteins, but it does have some drawbacks to note.

- Dye binding is dependent on the amino acid composition of a protein.
- If dye binding is not the same for your standard protein and your unknown protein, your concentration estimate will be off.
- Buffer components (esp. detergents like SDS) may interfere with binding.