1. Preparation of the Bradford protein assay reagent

1. Dissolve 100 mg of Coomassie Brilliant Blue G in 50 mL 95% (w/v) ethanol. Mix until Coomassie Brilliant Blue G is completely dissolved.
   CAUTION: Ethanol is flammable and causes eye irritation. Avoid flames and use goggles.
2. To the previous solution, add 100 mL of 85% (w/v) phosphoric acid carefully.
   CAUTION: Phosphoric acid is corrosive to metals and causes skin corrosion, serious eye damage, and acute oral toxicity. Wear gloves and goggles.
3. Add the solution containing Coomassie Brilliant Blue G, ethanol, and phosphoric acid slowly to 600 mL of deionized water.
4. Dilute the solution to a final volume of 1,000 mL. Scale up or down according to the number of samples to be analyzed. As described in the original method⁷, final concentrations in the working Bradford reagent should be 0.01% (w/v) Coomassie Brilliant Blue G, 4.7% (w/v) ethanol, and 8.5% (w/v) phosphoric acid.
5. Remove any insoluble material filtering through filter paper (Whatman #1 paper or equivalent).
6. The reagent is stable for several weeks when stored at room temperature (RT) and protected from light. Filter as necessary since precipitates may form over time.
   ​NOTE: Alternatively, ready to dilute and use Bradford reagents are commercially available. Follow the manufacturer's instructions for preparing the reagent and proceed to the next step.

2. Preparation of protein standard solutions

1. Prepare a stock solution (refer to step 2.4) of an isolated protein to be used as standard. An affordable and commonly used protein is bovine serum albumin (BSA). Other options are ovalbumin and bovine gamma globulin.
2. If the molar absorptivity of the protein used as standard is known, check the concentration of the stock solution in a spectrophotometer.
3. For BSA, a commonly used formula is BSA (mg/mL) = (A280/6.6) × 10, where A280 is the absorbance at 280 nm with a path length of 1 cm read against an appropriate blank (i.e., ε₂₈₀^1% = 6.6)⁷. For example, 0.8 mg/mL BSA has an absorbance of 0.528 at 280 nm.
4. To generate the standard curve, prepare several dilutions of BSA within 0.025 mg/mL and 1.0 mg/mL. These will result in 0.25-10 µg of BSA per well after adding a sample volume of 10 µL per well.
   NOTE: Protein standard solutions should be prepared in a medium with the same composition (final concentration) of the medium used to prepare samples.

3. Assay

1. Dilute the samples to achieve a protein concentration within the standard curve range of 0.025-1.0 mg/mL. Have several (at least 3) sample dilutions within the range.
   NOTE: Samples can be diluted with phosphate-buffered saline (PBS) or any other medium/buffer composition compatible with the Bradford reagent. The final concentration of medium components should not differ between standard and samples.
2. Add 10 µL of each protein standard solution to three wells (i.e., in triplicate) of a 96-well microplate. For the 0 (zero) protein point, add 10 µL of the buffer/medium used to prepare the standard solutions and sample dilutions.
3. To another set of wells, add 10 µL of each sample dilution to three wells (i.e., in triplicate) of the same 96-well microplate. One approach is to add different volumes of a sample and complete with a medium of up to 10 µL per well (e.g., 0.5 µL, 1 µL, 2 µL, 4 µL, and 8 µL of the sample plus 9.5 µL, 9 µL, 8 µL, 6 µL, and 2 µL of medium).
4. Add 250 µL of Bradford protein assay reagent to all wells. A typical microplate setup is shown in Figure 2. In this example, a set of blank wells containing 260 µL (the final volume per well) water was included to serve as blank in a microplate reader (step 4.5). This is required only if absorbance data will also be collected.
   NOTE: Prepare a standard curve in the same microplate of samples each time the assay is performed. In other words, if the number of samples requires another plate, prepare another standard curve in the second plate, and so on.
5. Record results (section 4) within 5-15 min.

Figure 2: A typical plate layout for the Bradford protein assay. Blank refers to three wells containing 260 µL of water to be used as blank in a microplate reader. STD refers to protein standards. S1-S6 are six different samples. SX_1-SX_4 are four different sample dilutions for each sample. Please click here to view a larger version of this figure.

4. Recording results

1. In a well-illuminated room, hold the microplate parallel to the bench against a uniform white background (e.g., a paper sheet) with one hand. Ensure accurate alignment by placing a bubble level on the plate.
2. With the other hand, hold the smartphone parallelly (some camera applications usefully indicate the inclination of the device) to the bench and the microplate and take one or several pictures of the whole microplate (Figure 3). In iOS devices, turn on the camera level indicator in the camera settings by enabling the Grid option. In Android devices, turn on the camera level indicator in the camera settings by enabling Framing Hints.
3. No special lighting apparatus is required, but take care to avoid shadows and reflections. For example, avoid shading the plate or the background with the smartphone and avoid shading the background with the microplate. Small reflections in the edges of the well area are not a problem; color data can be extracted from a very small area in the center of each well.
4. Briefly check the picture for background uniformity, shadows, and reflections. Also, take a look at the angle of the wells; the center of each well should be directly visible (i.e., not behind well walls).
5. If the comparison between microplate absorbance readings and picture color data is desired, read the microplate at 590 nm and 450 nm in a microplate reader²¹.

Figure 3: Capturing the results of the Bradford protein assay. In a well-illuminated room, the microplate is positioned parallelly to the bench against a uniform background with one hand. With the other hand, the smartphone is hold parallelly to the bench and the microplate. Please click here to view a larger version of this figure.

5. Extracting color data-Automatically

1. Download ImageJ and ReadPlate²², an ImageJ plugin is available at https://imagej.nih.gov/ij/plugins/readplate/index.html (this is a .txt file).
2. Open ImageJ, click on Plugins > Install and select the file downloaded in step 6.1.
3. Set the measurement parameters by clicking on Analyze > Set Measurements and check the following options: Area; Standard deviation; Min & max gray value; Mean gray value; Modal gray value. On the bottom of the window, set Redirect to: None and Decimal places (0-9): 3.
4. Go to File > Open and select the picture of the microplate taken in step 4.
5. Go to Plugins > ReadPlate. Read the instructions and then click OK.
6. Select the number of wells: 96.
7. Using the Rectangular selection tool automatically loaded by the plugin, make a rectangle beginning in the center of the A1 well and ending in the center of the H12 well. Then, click OK.
8. Select the Blue channel and click OK.
9. Confirm the default parameters by clicking on OK.
10. Check if the software delineated an area inside each well and if the selected areas are not covering areas with abnormal shadows or reflection. Click OK.
11. Save the results and repeat steps 5.8-5.10 for the green channel.
12. Calculate the blue-to-green ratio using the mode for each color.
    ​NOTE: The calculation can be done manually or using the software of the reader preferences, such as R, Microsoft Excel, or GraphPad Prism.

6. Extracting color data-Manually

1. Download Inkscape, a free and open-source graphics editor.
   NOTE: Any software with the color picker tool (usually depicted as an eye dropper) can be used to identify the color and extract RGB data.
2. Open Inkscape, go to File > Open. Select the picture of the microplate taken in Step 4.
3. Pick the Select and Transform Objects (S) tool, which is depicted as an arrow on the top-left side, and click on the picture. A dashed line border will indicate the selection.
4. Select the Pick Colors from Image (D) tool, which is depicted as an eye dropper on the left side.
5. Click in the center of a well. The color on the bottom panel ("Fill:") will change accordingly. Click on the color and a Fill and Stroke tab will pop up on the right side.
6. Change the Flat Color dropdown menu to RGB. Record the values shown for the blue and green channels for each well.
7. Calculate the blue-to-green ratio using the recorded values.
   ​NOTE: The calculation can be done manually or using the software of the reader preferences, such as R, Microsoft Excel, or GraphPad Prism.

7. Building standard curves and extrapolating unknowns

1. Plot the average green-to-blue intensity ratio as a function of protein standard concentration.
   NOTE: Plot the data manually or using the software of the reader preferences, such as R, Microsoft Excel, or GraphPad Prism.
2. Calculate the average green-to-blue intensity ratio for each sample and dilution.
3. Check if the signal obtained for the sample lies within the linear range of the protein standard.
4. Ignore any value below or above the minimum or maximum values of the standard curve.
5. Use the linear equation describing the standard curve to extrapolate the amount of protein in the sample. Multiply the calculated values by the dilution factor accordingly.