Bradford Assay

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Introduction

The Bradford assay should be used to quantify total protein within a sample relative to a standard curve. This is important when running proteins on SDS-PAGE/Western blot, where each lane must contain an identical amount of total protein. This way, the intensity of the bands can be compared between different lanes, as standardizing the amount of protein loaded per lane will account for any variation in cell count from which the proteins were derived.

Protocol

For a 96 well plate

  1. Using a 0.2 or 0.45 μm filter and 15 mL syringe, filter Bio-Rad Protein Assay Dye Reagent Concentrate (Bradford Dye) into a 15 mL falcon tube. Dilute 1:5 with ddH2O (1 part concentrate, 4 parts ddH2O). Invert to mix. Let warm to room temperature.
  2. Collect lysate or supernatant containing protein. Keep on ice.
  3. Label eight 1.5 mL tubes as: 1, 2, 3, 4, 5, 6, 7, 8. Set up the BSA standard by following Table 1. (Use the Bio-Rad 2 mg/mL stock. Do not make BSA yourself)
    • Make sure the diluent is the same as buffer that the protein of interest is in. For example, if in PBS, use PBS to dilute the standard. If in RIPA buffer, use RIPA, etc.
    • Mix by pipetting up and down. Avoid creating bubbles in the tube
  4. Set up 500 μL dilutions of your sample of interest. Make dilutions of 1:5, 1:10, and 1:25.
    • If you think your sample is very concentrated, instead make 1:10, 1:25, and 1:25 dilutions
    • If you think your sample is very dilute, use undiluted, 1:2, and 1:5 dilutions. (This uses a lot of sample however!)
  5. Add 150 μL of sample or standard to each well following Figure 1. Avoid bubbles by only pipetting to the first stop.
  6. Pour the dye into a multichannel pipette reservoir. Using a multichannel pipette, pipet 150 μL of dye into each well. Mix by pipetting carefully up and down.
  7. Incubate for at least 5 minutes at room temperature. Do not exceed 1 hr.
  8. Take off the lid of the plate, and place it into a plate reader/spectrometer.
  9. Measure absorbance at 595 nm.
Table 1.
Tube # Standard volume (μL) Source of standard Diluent volume (μL) Final concentration (μg/mL)
1 10 2 mg/mL stock 790 25
2 10 2 mg/mL stock 990 20
3 6 2 mg/mL stock 794 15
4 500 Tube 2 500 10
5 500 Tube 4 500 5
6 500 Tube 5 500 2.5
7 500 Tube 6 500 1.25
8 - - 500 0
Figure 1









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Analysis using the standard curve

  1. Export results from spectrometer to Excel.
  2. Subtract the average absorbance of each standard/sample by the average absorbance of the blank. Essentially, you are setting the blank absorption to 0 and making all readings relative to that.
  3. Plot the known standard concentrations (X-axis: 0, 1.25, 2.5, 5, 10, 15, 20, and 25 μg/mL) with the average absorbance at each concentration (Y-axis). Add a line of best fit and display the equation with R2 value. Ideally, the R2 value should be ≥ 0.99. R2 < 0.94 curves should not be used.
  4. Calculate the average absorbance for the samples of interest by using the equation of the line graph y = mx+b (Absorbance = Slope*Concentration + b) and solve for x.
  5. If the calculated concentration falls within the linear range of the standard curve (1.25 - 25 μg/mL), then it is accurate. Determine the concentration of the undiluted sample by multiplying by the dilution factor.