The Bradford assay for protein is widely used because of its sensitivity, speed, convenience, lack of need for a UV-capable spectrophotometer, and adaptability to 96-well plates. The "Bradford Reagent" is an acidic stain which turns blue when it interacts with protein. The resulting absorbance is best determined at 595 nm. The reagent is typically sold as a concentrated stock at 5-fold working strength. The protein standard or unknown must be sufficiently diluted not to interfere with the low pH achieved by the acid in the reagent.

Microplate Bradford Assay

Reagent and protein are mixed in wells of a flat-bottomed, 96-well plate, and the absorbance is read at 595 nm (or the closest available wavelength available on a 96-well plate reader, which may be 570 nm or 620 nm). As with all assays, the amounts of protein placed in the wells must be within the range where the assay result is directly proportional to protein concentration, the "linear range" of the assay. In practice, the Bradford result is often not quite linear, and so a curve rather than a straight line will better fit the results.

According to the manufacturer, this assay is ten-fold more sensitive than absorbance at 280 nm. If we take an absorbance of 0.1 to be the minimum reliably detectable, A₂₈₀ can detect a minimum of about 75 micrograms/ml. The microplate Bradford assay achieves an absorbance of 0.1 at about 6 micrograms/ml. Moreover, the microplate Bradford has the advantage that large numbers of samples can be read quickly without expensive UV-capable cuvettes.

When full to the brim, one well in a 96-well plate holds 300 microliters; 200 microliters is a comfortable volume to use per well. The Bradford reagent (stain in phosphoric acid and a water/methanol mixture) is designed to be diluted 5-fold with protein. Therefore, in each well we'll put 160 microliters of protein, diluted in water; to this, we'll add 40 microliters of Bradford reagent, thereby achieving the recommended 5-fold dilution of reagent.

Standard Curve

The Bradford assay is calibrated by using a pure protein of known concentration, called the "standard protein". Like other protein assays, the result of the Bradford assay differs for different proteins. You are aware that A₂₈₀ is about twice as sensitive to IgG as to BSA. The Bradford is the reverse: about twice as sensitive to BSA as to IgG. It is best to use the same protein for the standard which you wish to estimate in the unknown. However, it is acceptable to use a different protein for the standard provided you know the correction factor for the unknown protein. By convention, proteins are diluted in distilled water for the Bradford assay.

1. Put 160 microliters water in well A-1, which will be taken to be the blank, and subtracted from the absorbances of all other wells.

2. Use standard protein at 0.05 mg/ml (50 micrograms/ml; dilute the standard provided to 0.05 mg/ml if it is more concentrated). Perform triplicate dilutions as follows:

          microliters/well
         ------------------
                 Standard @   Assay
   Row   Water   0.05 mg/ml   mg/ml**
   
   B     144      16*
   C     120      40
   D      80      80
   E       0     160
   
   * Use P20 pipet; P200 is suitable for 30 or more microliters.
   ** Calculate this for the 160 microliter volume, not the final 200
   microliter volume.  This makes it easier to interpret your unknowns.
   

3. Add 40 microliters of Bradford reagent to each well, and pipet in and out eight times to mix well. When pipetting, avoid making bubbles by keeping the pipet tip well below the surface. If you start at the blank and move to the same or higher concentrations, you can use the same yellow pipet tip for all wells. (If you want to blow residual mixture out of the tip, do so on a tissue, not into the Bradford reagent.)

4. Check wells for bubbles. If bubbles are present, use a P20 to add 5 microliters of ethanol to the surface of the well -- this breaks bubbles without affecting the absorbance.

5. Read the absorbances of your wells at the wavelength available on the reader closest to 595 nm. (The reagent manufacturer recommends reading the assay 5 to 60 minutes after mixing.) Write the time when you read the plate on the printout (unless the reader prints the time).

6. Are the absorbances within an acceptable range? Given the absorbance of the blank (about 0.38; some readers report this and some don't), what is the reliable range for the corrected absorbances?

7. If the absorbances are not in the acceptable range, increase or decrease the concentration of the starting protein standard to put the values in the desired range, and run the standard curve again. If you run it again, transfer the old blank to a different well in row A, and make a new one in well A1, so it matches the new standard curve.

8. Optional: Once you have results in an acceptable range, you may wish to do a more detailed standard curve, such as this one in duplicate:

   Row   Water   Standard   mg/ml
   
   A     160       0
   B     144      16
   C     130      30
   D     115      45
   E      90      70
   F      60     100
   G      30     130
   H       0     160
   

9. Graph your results (absorbance on vertical axis/ordinate/Y vs. protein mg/ml on horizontal axis/abcissa/X).

10. Optional: Re-read your plate. Have the absorbances changed significantly? How much time elapsed between readings?

11. When finished with your plate, dump it into the sink, and rinse the wells several times with distilled water (to remove protein), and then ethanol until all blue stain is removed. Rinse again with distilled water, and allow it to dry with the cover off for later re-use.

Determination of Unknowns (Fractions from Sephadex Column)

The absorbance in the Bradford assay varies from run to run depending on the batch of reagent used, the time between mixing and reading, and which reader is used. Therefore it is best to run a standard curve alongside your unknowns, mixing the reagent in the standard wells at the same time as in your unknown wells.

The total amount of protein put into our Sephadex columns is roughly 100 milligrams. This will be distributed in about 40 one-ml fractions. What sensitivity do we need? If a fraction has less than 0.5 mg/ml, we don't need to know the concentration accurately since it will not contribute enough to a peak to be included in a pool. The assay can detect 0.005 mg/ml, so we'll dilute each fraction 100-fold, making 0.5 mg/ml in the undiluted fraction the minimum we can reliably detect. This requires that we consume only a tiny portion of each fraction.

12. Set up a standard curve of protein dilutions in your plate.

13. Plan a plate layout with two wells per unknown fraction. The first well will of each pair will be used only to make a 1/10 dilution; no Bradford reagent will be added to it. The second well will contain the 1/100 dilution, to which Bradford reagent will be added. Into each of the pairs of wells for your unknowns, put 144 microliters water.

14. With a P20, add 16 microliters of unknown to the first well of each pair, and with the pipet touching the bottom corner, pipet in and out ten times to mix. Transfer 16 microliters of the 1/10 dilution to the second well of the pair, and mix. Since the purpose of testing fractions is to decide which fractions to pool, only rough accuracy is required at this point. Therefore, you can use the same tip for many unknown dilutions.

15. Add 40 microliters of Bradford reagent to the second well of each pair (the 1/100 dilution), and pipet in and out 8-10 times to mix well. When pipetting, avoid making bubbles by keeping the pipet tip well below the surface, touching the bottom corner.

16. Check wells for bubbles. If bubbles are present, use a P20 to add 5 microliters of ethanol to the surface of the well -- this breaks bubbles without affecting the absorbance.

17. Read the absorbances of your wells at the wavelength available on the reader closest to 595 nm. (The reagent manufacturer recommends reading the assay 5 to 60 minutes after mixing.) Write the time when you read the plate on the printout.

18. Graph your standard curve (absorbance on the Y axis vs. protein mg/ml on the X axis).

19. Estimate the concentrations of your unknowns from your graph. Graph these (mg/ml on the ordinate vs. fraction number on the abcissa). Since it takes twice the concentration of IgG as BSA to give equal absorbances in the Bradford assay, if you used a BSA standard curve, correct the estimated unknown concentrations: multiply "BSA" mg/ml by two to estimate IgG mg/ml.

20. Decide which fractions to pool. Consult an instructor to verify your plan. Pool the fractions, recording the total volume.

21. Into available wells in your 96-well plate, make a suitable dilution, in triplicate, of each pool. Make a new standard curve series. Read the plate and determine the IgG concentration in your pools. Multiply the concentration times the total volume to get total mg IgG in each pool.

22. When finished with your plate, dump it into the sink, and rinse the wells several times with distilled water (to remove protein), and then ethanol until all blue stain is removed. Rinse again with distilled water, and allow it to dry with the cover off for later re-use.