Protein Assay Essay

Bradford protein assay

Considerations for use

The Bradford assay is very fast and uses about the same amount of protein as the Lowry assay. It is fairly accurate and samples that are out of range can be retested within minutes. The Bradford is recommended for general use, especially for determining protein content of cell fractions and assesing protein concentrations for gel electrophoresis.

Assay materials including color reagent, protein standard, and instruction booklet are available from Bio-Rad Corporation. The method described below is for a 100 µl sample volume using 5 ml color reagent. It is sensitive to about 5 to 200 micrograms protein, depending on the dye quality. In assays using 5 ml color reagent prepared in lab, the sensitive range is closer to 5 to 100 µg protein. Scale down the volume for the "microassay procedure," which uses 1 ml cuvettes. Protocols, including use of microtiter plates are described in the flyer that comes with the Bio-Rad kit.


The assay is based on the observation that the absorbance maximum for an acidic solution of Coomassie Brilliant Blue G-250 shifts from 465 nm to 595 nm when binding to protein occurs. Both hydrophobic and ionic interactions stabilize the anionic form of the dye, causing a visible color change. The assay is useful since the extinction coefficient of a dye-albumin complex solution is constant over a 10-fold concentration range.


In addition to standard liquid handling supplies a visible light spectrophotometer is needed, with maximum transmission in the region of 595 nm, on the border of the visible spectrum (no special lamp or filter usually needed). Glass or polystyrene (cheap) cuvettes may be used, however the color reagent stains both. Disposable cuvettes are recommended.



  1. Bradford reagent: Dissolve 100 mg Coomassie Brilliant Blue G-250 in 50 ml 95% ethanol, add 100 ml 85% (w/v) phosphoric acid. Dilute to 1 liter when the dye has completely dissolved, and filter through Whatman #1 paper just before use.
  2. (Optional) 1 M NaOH (to be used if samples are not readily soluble in the color reagent).
The Bradford reagent should be a light brown in color. Filtration may have to be repeated to rid the reagent of blue components. The Bio-Rad concentrate is expensive, but the lots of dye used have apparently been screened for maximum effectiveness. "Homemade" reagent works quite well but is usually not as sensitive as the Bio-Rad product.


  1. Warm up the spectrophotometer before use.
  2. Dilute unknowns if necessary to obtain between 5 and 100 µg protein in at least one assay tube containing 100 µl sample
  3. If desirred, add an equal volume of 1 M NaOH to each sample and vortex (see Comments below). Add NaOH to standards as well if this option is used.
  4. Prepare standards containing a range of 5 to 100 micrograms protein (albumin or gamma globulin are recommended) in 100 µl volume. See how to set up an assay for suggestions as to setting up the standards.
  5. Add 5 ml dye reagent and incubate 5 min.
  6. Measure the absorbance at 595 nm.


Prepare a standard curve of absorbance versus micrograms protein and determine amounts from the curve. Determine concentrations of original samples from the amount protein, volume/sample, and dilution factor, if any.


The dye reagent reacts primarily with arginine residues and less so with histidine, lysine, tyrosine, tryptophan, and phenylalanine residues. Obviously, the assay is less accurate for basic or acidic proteins. The Bradford assay is rather sensitive to bovine serum albumin, more so than "average" proteins, by about a factor of two. Immunoglogin G (IgG - gamma globulin) is the preferred protein standard. The addition of 1 M NaOH was suggested by Stoscheck (1990) to allow the solubilization of membrane proteins and reduce the protein-to-protein variation in color yield.


  • Bradford, MM. A rapid and sensitive for the quantitation of microgram quantitites of protein utilizing the principle of protein-dye binding. Analytical Biochemistry 72: 248-254. 1976.
  • Stoscheck, CM. Quantitation of Protein. Methods in Enzymology 182: 50-69 (1990).

This article is about the scientific procedure. For other uses, see Bradford (disambiguation).

The Bradford protein assay was developed by Marion M. Bradford in 1976.[1] It is a quick and accurate[2]spectroscopic analytical procedure used to measure the concentration of protein in a solution. It is subjective, i.e., dependent on the amino acid composition of the measured protein.


The Bradford assay, a colorimetric protein assay, is based on an absorbance shift of the dye Coomassie Brilliant Blue G-250. The Coomassie Brilliant Blue G-250 dye exists in three forms: anionic (blue), neutral (green), and cationic (red).[3] Under acidic conditions, the red form of the dye is converted into its blue form, binding to the protein being assayed. If there's no protein to bind then, the solution will remain brown. The dye forms a strong, noncovalent complex with the protein's carboxyl group by Van der Waals force and amino group through electrostatic interactions.[1] During the formation of this complex, the red form of Coomassie dye first donates its free electron to the ionizable groups on the protein, which causes a disruption of the protein's native state, consequently exposing its hydrophobic pockets. These pockets in the protein's tertiary structure bind non-covalently to the non-polar region of the dye via the first bond interaction (van der Waals forces) which position the positive amine groups in proximity with the negative charge of the dye. The bond is further strengthened by the second bond interaction between the two, the ionic interaction. The binding of the protein stabilizes the blue form of the Coomassie dye; thus the amount of the complex present in solution is a measure for the protein concentration, and can be estimated by use of an absorbance reading.[citation needed]

The cationic (unbound) form is green / red and has an absorption spectrum maximum historically held to be at 465 nm. The anionic bound form of the dye which is held together by hydrophobic and ionic interactions, has an absorption spectrum maximum historically held to be at 595 nm.[4] The increase of absorbance at 595 nm is proportional to the amount of bound dye, and thus to the amount (concentration) of protein present in the sample.[citation needed]

Unlike other protein assays, the Bradford protein assay is less susceptible to interference by various chemical compounds such as sodium, potassium or even carbohydrates like sucrose, that may be present in protein samples.[2] An exception of note is elevated concentrations of detergent. Sodium dodecyl sulfate (SDS), a common detergent, may be found in protein extracts because it is used to lyse cells by disrupting the membrane lipid bilayer and to denature proteins for SDS-PAGE. While other detergents interfere with the assay at high concentration, the interference caused by SDS is of two different modes, and each occurs at a different concentration. When SDS concentrations are below critical micelle concentration (known as CMC, 0.00333%W/V to 0.0667%) in a Coomassie dye solution, the detergent tends to bind strongly with the protein, inhibiting the protein binding sites for the dye reagent. This can cause underestimations of protein concentration in solution. When SDS concentrations are above CMC, the detergent associates strongly with the green form of the Coomassie dye, causing the equilibrium to shift, thereby producing more of the blue form. This causes an increase in the absorbance at 595 nm independent of protein presence.[citation needed]

Other interference may come from the buffer used when preparing the protein sample. A high concentration of buffer will cause an overestimated protein concentration due to depletion of free protons from the solution by conjugate base from the buffer. This will not be a problem if a low concentration of protein (subsequently the buffer) is used.[citation needed]

In order to measure the absorbance of a colorless compound a Bradford assay must be performed. Some colorless compounds such as proteins can be quantified at an Optical Density of 280 nm due to the presence of aromatic rings such as Tryptophan, Tyrosine and Phenylalanine but if none of these amino acids are present then the absorption cannot be measured at 280 nm.[5]


Many protein-containing solutions have the highest absorption at 280 nm in the spectrophotometer, the UV range. This requires spectrophotometers capable of measuring in the UV range, which many cannot. Additionally, the absorption maxima at 280 nm requires that proteins contain aromatic amino acids such as tyrosine (Y), phenylalanine (F) and/or tryptophan (W). Not all proteins contain these amino acids which will skew the concentration measurements. If nucleic acids are present in the sample, they would also absorb light at 280 nm, skewing the results further. By using the Bradford protein assay, one can avoid all of these complications by simply mixing the protein samples with the Coomassie Brilliant Blue G-250 dye (Bradford reagent) and measuring their absorbances at 595 nm, which is in the Vis range.[6]

The procedure for Bradford protein assay is very easy and simple to follow. It is done in one step where the Bradford reagent is added to a test tube along with the sample. After mixing well, the mixture almost immediately changes to a blue color. When the dye binds to the proteins through a process that takes about 2 minutes, a change in the absorption maximum of the dye from 495 nm to 595 nm in acidic solutions occurs.[2] This dye creates strong noncovalent bonds with the proteins, via electrostatic interactions with the amino and carboxyl groups, as well as Van Der Waals interactions. Only the molecules that bind to the proteins in solution exhibit this change in absorption, which eliminates the concern that unbound molecules of the dye might contribute to the experimentally obtained absorption reading. This process is more beneficial since it is less pricey than other methods, easy to use, and has high sensitivity of the dye for protein.[7]

After 5 minutes of incubation, the absorbance can be read at 595 nm using a spectrophotometer; an easily accessible machine.

This assay is one of the fastest assays performed on proteins.[8] The total time it takes to set up and complete the assay is under 30 minutes.[9] The entire experiment is done at room temperature.

The Bradford protein assay can measure protein quantities as little as 1 to 20 μg.[10] It is an extremely sensitive technique.

The dye reagent is a stable ready to use product prepared in phosphoric acid. It can remain at room temperature for up to 2 weeks before it starts to degrade.

Protein samples usually contain salts, solvents, buffers, preservatives, reducing agents and metal chelating agents. These molecules are frequently used for solubilizing and stabilizing proteins. Other protein assay like BCA and Lowry are ineffective because molecules like reducing agents interfere with the assay.[11] Using Bradford can be advantageous against these molecules because they are compatible to each other and will not interfere.[12]

The linear graph acquired from the assay (absorbance versus protein concentration in μg/mL) can be easily extrapolated to determine the concentration of proteins by using the slope of the line.

It is sensitive technique. It is also very simple: measuring the OD at 595 nm after 5 minutes of incubation. This method can also make use of a Vis spectrophotometer.[13]


The Bradford assay is linear over a short range, typically from 0 µg/mL to 2000 µg/mL, often making dilutions of a sample necessary before analysis. In making these dilutions, error in one dilution is compounded in further dilutions resulting in a linear relationship that may not always be accurate.

Basic conditions and detergents, such as SDS, can interfere with the dye's ability to bind to the protein through its side chains.[8] However, there are some detergent-compatible Bradford reagents. The Bradford assay depends on the sequence of the protein. Thus, if the protein does not contain an ideal number of aromatic residues, then the dye will not be able to bind to the protein efficiently. Another disadvantage of the Bradford Protein Assay is that this method depends on comparing the absorbance of the protein to that of a standard protein. So if the protein does not react to the dye in a similar way as the standard protein, it is possible that the concentration is off.

The reagents in this method tend to stain the test tubes. Same test tubes cannot be used since the stain would affect the absorbance reading. This method is also time sensitive. When more than one solution is tested, it is important to make sure every sample is incubated for the same amount of time for accurate comparison.[14]

It is also inhibited by the presence of detergents, although this problem can be alleviated by the addition of cyclodextrins to the assay mixture.[15]

Much of the non-linearity stems from the equilibrium between two different forms of the dye which is perturbed by adding the protein. The Bradford assay linearizes by measuring the ratio of the absorbances, 595 over 450 nm. This modified Bradford assay is approximately 10 times more sensitive than the conventional one.[16]

Sample Bradford procedure[edit]


  • Lyophilized bovine plasma gamma globulin
  • Coomassie Brilliant Blue 1
  • 0.15 M NaCl
  • Spectrophotometer and cuvettes
  • Micropipettes

Procedure (Standard Assay, 20-150 µg protein; 200-1500 µg/mL)[edit]

  1. Prepare a series of standards diluted with 0.15 M NaCl to final concentrations of 0 (blank = No protein), 250, 500, 750 and 1500 µg/mL. Also prepare serial dilutions of the unknown sample to be measured.
  2. Add 100 µL of each of the above to a separate test tube (or spectrophotometer tube if using a Spectronic 20).
  3. Add 5.0 mL of Coomassie Blue to each tube and mix by vortex, or inversion.
  4. Adjust the spectrophotometer to a wavelength of 595 nm, using the tube which contains no protein (blank).
  5. Wait 5 minutes and read each of the standards and each of the samples at 595 nm wavelength.
  6. Plot the absorbance of the standards vs. their concentration. Compute the extinction coefficient and calculate the concentrations of the unknown samples.

Procedure (Micro Assay, 1-10 µg protein/mL)[edit]

  1. Prepare standard concentrations of protein of 1, 5, 7.5 and 10 µg/mL. Prepare a blank of NaCl only. Prepare a series of sample dilutions.
  2. Add 100 µL of each of the above to separate tubes (use microcentrifuge tubes) and add 1.0 mL of Coomassie Blue to each tube.
  3. Turn on and adjust a spectrophotometer to a wavelength of 595 nm, and blank the spectrophotometer using 1.5 mL cuvettes.
  4. Wait 2 minutes and read the absorbance of each standard and sample at 595 nm.
  5. Plot the absorbance of the standards vs. their concentration. Compute the extinction coefficient and calculate the concentrations of the unknown samples.

Using data obtained to find concentration of unknown[edit]

In summary, in order to find a standard curve, one must use varying concentrations of BSA (Bovine Serum Albumin)[2] in order to create a standard curve with concentration plotted on the x-axis and absorbance plotted on the y-axis. Only a narrow concentration of BSA is used (2-10 ug/mL) in order to create an accurate standard curve.[17] Using a broad range of protein concentration will make it harder to determine the concentration of the unknown protein. This standard curve is then used to determine the concentration of the unknown protein. The following elaborates on how one goes from the standard curve to the concentration of the unknown.

First, add a line of best fit, or Linear regression and display the equation on the chart. Ideally, the R2 value will be as close to 1 as possible. R represents the sum of the square values of the fit subtracted from each data point. Therefore, if R2 is much less than one, consider redoing the experiment to get one with more reliable data.[18]

The equation displayed on the chart gives a means for calculating the absorbance and therefore concentration of the unknown samples. In Graph 1, x is concentration and y is absorbance, so one must rearrange the equation to solve for x and enter the absorbance of the measured unknown.[19] It is likely that the unknown will have absorbance numbers outside the range of the standard. These should not be included calculations, as the equation given cannot apply to numbers outside of its limitations. In a large scale, one must compute the extinction coefficient using the Beer-Lambert Law A=εLC in which A is the measured absorbance, ε is the slope of the standard curve, L is the length of the cuvette, and C is the concentration being determined.[20] In a micro scale, a cuvette may not be used and therefore one only has to rearrange to solve for x.

In order to attain a concentration that makes sense with the data, the dilutions, concentrations, and units of the unknown must be normalized (Table 1). To do this, one must divide concentration by volume of protein in order to normalize concentration and multiply by amount diluted to correct for any dilution made in the protein before performing the assay.

Alternative assays[edit]

Alternative protein assays include:


Further reading[edit]

  • Bradford, M.M. (1976), "Rapid and sensitive method for the quantitation of microgram quantities of protein utilizing the principle of protein-dye binding", Anal. Biochem., 72: 248–254, doi:10.1016/0003-2697(76)90527-3, PMID 942051 
  • Zor, T.; Selinger, Z. (1996), "Linearization of the Bradford protein assay increases its sensitivity: theoretical and experimental studies", Anal. Biochem., 236: 302–308, doi:10.1006/abio.1996.0171, PMID 8660509 
  • Noble, J.E.; Bailey, M.J.A. (2009), "Quantitation of Protein", Methods Enzymol., 463: 73–95, doi:10.1016/S0076-6879(09)63008-1 
  • Albright, Brian (2009), Mathematical Modeling with Excel, p. 60, ISBN 978-0763765668 
  • Stephenson, Frank Harold (2003), Calculations for molecular biology and biotechnology: a guide to mathematics in the laboratory, p. 252, ISBN 0126657513 
  • Dennison, Clive (2003), "A guide to protein isolation", Focus on structural biology, 3: 39, ISBN 1402012241 
  • Ibanez, Jorge G. (2007), Environmental chemistry: fundamentals, p. 60, ISBN 0387260617 

External links[edit]

Figure 1. Coomassie Brilliant Blue G-250, the binding dye for the Bradford Method
Graph 1. Actual BSA data attained from a micro scale UV-Vis Spectrophotometer
Table 1. Actual assay data for determine concentration of unknown based on line of best fit of the above standard curve
  1. ^ abNinfa, Alexander J; Ballou, David P; Benore, Marilee (2008). Fundamental Laboratory Approaches for Biochemistry and Biotechnology. Wiley. p. 113. 
  2. ^ abcdBradford, Marion (1976). "A Rapid and Sensitive Method for the Quantification of Microgram Quantities of Protein Utilizing the Principle of Protein-Dye Binding"(PDF). Analytical Biochemistry. 72: 248–254. doi:10.1006/abio.1976.9999 – via Google Scholar. 
  3. ^"Quick Start TM Bradford Protein Assay"(PDF). Archived from the original(PDF) on 2017-12-17. 
  4. ^"Protein determination by the Bradford method". 
  5. ^P., Ballou, David; Marilee., Benore, (2010). Fundamental laboratory approaches for biochemistry and biotechnology. John Wiley. ISBN 9780470087664. OCLC 420027217. 
  6. ^Ninfa, Ballou, Benore, Alexander J., David P., Marilee (2010). Fundamental Laboratory Approaches for Biochemistry and Biotechnology. United States of America: John Wiley & Sons, Inc. pp. 110, 113. ISBN 978-0-470-08766-4. 
  7. ^Ninfa, Alexander J.; Ballou, David P.; Benore, Marilee (2010). Fundamental Laboratory Approaches for Biochemistry and Biotechnology. John Wiley & Sons Inc. p. 113. ISBN 978-0470087664. 
  8. ^ abOkutucu, Burcu; Dınçer, Ayşşe; Habib, Ömer; Zıhnıoglu, Figen (2007-08-01). "Comparison of five methods for determination of total plasma protein concentration". Journal of Biochemical and Biophysical Methods. 70 (5): 709–711. doi:10.1016/j.jbbm.2007.05.009. 
  9. ^"Protein Assay Technical Handbook"(PDF). 
  10. ^"4.5. Determination of protein concentration". Retrieved 2016-05-19. 
  11. ^barbosa, Helder; Slater K.H, Nigel (3 August 2009). "Protein quantification in the presence of poly(ethylene glycol) and dextran using the Bradford method". Analytical Biochemistry: Methods in the Biological Sciences. 395 (1): 108–110. doi:10.1016/j.ab.2009.07.045. 
  12. ^Ninfa, Alexander J. (2010). Fundamental Laboratory Approaches for Biochemistry and Biotechnology. Wiley. pp. 117–118. ISBN 978-0470087664. 
  13. ^Ninfa, Ballou (1998). Fundamental Approaches to Biochemistry and Biotechnology. Fitzgerald Science Press, Bethesda, MD. pp. 114–116. ISBN 978-0470087664. 
  14. ^Ninfa, Alexander J; Ballou, David P; Benore, Marilee (2009). Fundamental Laboratory Approches for Biochemistry and Biotechnology. Wiley. p. 113. 
  15. ^Rabilloud, Thierry. "A single step protein assay that is both detergent and reducer compatible: The cydex blue assay". Electrophoresis. 37 (20): 2595–2601. arXiv:1610.07373. doi:10.1002/elps.201600270. PMID 27445231. 
  16. ^Zor, Tsaffrir; Selinger, Zvi (1996-05-01). "Linearization of the Bradford Protein Assay Increases Its Sensitivity: Theoretical and Experimental Studies". Analytical Biochemistry. 236 (2): 302–308. doi:10.1006/abio.1996.0171. 
  17. ^"Linearization of the Bradford Protein Assay Increases Its Sensitivity: Theoretical and Experimental Studies"(PDF). November 20, 1995. Archived from the original(PDF) on December 17, 2017. 
  18. ^Albright, Brian (2009). Mathematical Modeling with Excel. p. 60. ISBN 978-0763765668. 
  19. ^Stephenson, Frank Harold (2003). Calculations for molecular biology and biotechnology: a guide to mathematics in the laboratory. p. 252. ISBN 0126657513. 
  20. ^Ibanez, Jorge G. (2007). Environmental chemistry: fundamentals. p. 60. ISBN 0387260617. 

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