Bradford Assay: Difference between revisions

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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.  
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'''


'''Protocol'''
'''Protocol'''


''Some things to know'':
* This protocol assumes the use of bovine serum albumin (BSA) for the standard curve, which has a linear range of 200 μg/mL to 1000 μg/mL within Bio-Rad's protein assay dye.
* If your protein is within a supernatant containing an acid indicator, such as phenol red, this may interfere with the assay. If your protein is within a solution containing phenol red, then ensure that all samples, standards, and blanks are diluted with that media (e.g. Serum free DMEM).
* Each tube will have extra volume to ensure you have enough for the last replicate and to avoid air bubbles.
* Reverse pipetting technique is highly recommended to avoid air bubbles and to ensure accuracy.
''For a 96 well plate''
''For a 96 well plate''


# Using a 0.2 μm filter and 15 mL syringe, filter Bio-Rad Protein Assay Dye Reagent Concentrate (Bradford Dye) into a 1.5 mL Eppendorf tube or 5 mL tube. Let warm to room temperature.  
# 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.  
# Collect lysate or supernatant containing protein. Keep on ice.
# Collect lysate or supernatant containing protein. Keep on ice.
# In separate 1.5 mL tubes, dilute the sample using ddH2O (or whatever buffer/media is appropriate) into 1:2 and 1:10 dilutions.  
# 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)
# Prepare a 10 mg/mL stock solution of BSA by dissolving BSA in ddH2O. Prepare a dilution series using ddH2O (or whatever buffer/media is appropriate) in 1.5 mL tubes: 1 mg/mL, 0.8 mg/mL, 0.6 mg/mL, 0.4 mg/mL, and 0.2 mg/mL.  
#*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.
# See '''Figure 1''' for a general plate setup. 80 μL of sample and standard will be used per well.
#*Mix by pipetting up and down. Avoid creating bubbles in the tube
# Spin down the plate at 300 rpm for 1 min to remove bubbles and any liquid on the side of the well.
#Set up 500 μL dilutions of your sample of interest. Make dilutions of 1:5, 1:10, and 1:25.  
# Pour filtered Bradford Dye into a multichannel pipette reservoir. Using a multichannel pipette, pipet dye into each well such that the dye is diluted 1/5 (for example, if 80 μL of sample was loaded per well, then 20 μL of dye will be added in each well) If a multichannel pipette is not available, use the micropipette and pipet into each well '''as quickly as possible'''.  
#*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!)
#Add 150 μL of sample or standard to each well following '''Figure 1'''. Avoid bubbles by only pipetting to the first stop.
# 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.
# Incubate for at least 5 minutes at room temperature. Do not exceed 1 hr.  
# Incubate for at least 5 minutes at room temperature. Do not exceed 1 hr.  
# Take off the lid of the plate, and place it into a plate reader/spectrometer.
# Take off the lid of the plate, and place it into a plate reader/spectrometer.
# Measure absorbance at 595 nm.
# Measure absorbance at 595 nm.
[[File:Bradford plate layout.png|thumb|'''Figure 1.''' General plate layout |border|center]]
{| class="wikitable"
|+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
|}
[[File:96 well plate for wiki (1).png|left|thumb|500x500px|'''Figure 1''']]
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 
 


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# Export results from spectrometer to Excel.
# Export results from spectrometer to Excel.
# Subtract the average absorbance of each well by the average absorbance of the blank.
# 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.
# Plot the known standard concentrations (X-axis; 0.2, 0.4, 0.6, 0.8, and 1.0 mg/mL) with the average absorbance at each concentration (Y-axis). '''Add a line of best fit and display the equation with R<sup>2</sup> value.''' Ideally, the R<sup>2</sup> value should be ≥ 0.99. R<sup>2</sup> < 0.94 curves should not be used.  
# 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 R<sup>2</sup> value.''' Ideally, the R<sup>2</sup> value should be ≥ 0.99. R<sup>2</sup> < 0.94 curves should not be used.  
# 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.
# 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.
# If the calculated concentration falls within the linear range of the standard curve (0.2 - 1.0 mg/mL), then it is accurate. If necessary, determine the concentration of the undiluted sample by multiplying by the dilution factor.  
# 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.


#  
#  


#
#

Latest revision as of 14:06, 24 May 2022

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.
    • 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.
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