Most colorimetric protein assay methods can be divided into two groups based on the type of chemistry involved: those involving protein-copper chelation with secondary detection of the reduced copper and those based on protein-dye binding with direct detection of the color change associated with the bound dye.
Most commercial protein assay reagents are well-characterized, robust products that provide consistent, reliable results. Nevertheless, each assay reagent has its limitations; having a basic understanding of the chemistries involved with each type of assay is essential for selecting an appropriate method for a given sample and for correctly evaluating results.
Copper-based protein assays, including the BCA and Lowry methods, depend on the well-known "biuret reaction", whereby peptides containing three or more amino acid residues form a colored chelate complex with cupric ions (Cu2+) in an alkaline environment containing sodium potassium tartrate. This became known as the biuret reaction because it is chemically similar to a complex that forms with the organic compound biuret (NH2-CO-NH-CO-NH2) and the cupric ion. Biuret, a product of excess urea and heat, reacts with copper to form a light blue tetradentate complex.
Figure 1. Diagram of the biuret reaction. By reducing the copper ion from cupric to cuprous form, the reaction produces a faint blue-violet color.
Figure 2. Structures of urea, biuret and peptide. Because polypeptides have a structure similar to biuret, they are able to complex with copper by the biuret reaction.
Single amino acids and dipeptides do not give the biuret reaction, but tripeptides and larger polypeptides or proteins will react to produce a light blue to violet complex that absorbs light at 540 nm. One cupric ion forms a colored coordination complex with four to six nearby peptides bonds. The intensity of the color produced is proportional to the number of peptide bonds participating in the reaction. Thus, the biuret reaction is the basis for a simple and rapid colorimetric reagent of the same name for quantitatively determining total protein concentration. The working range for the biuret assay is 5-160 mg/mL, which is adequate for some types of industrial applications but not nearly sensitive enough for most protein research needs.
The BCA Protein Assay was introduced by Smith, et al., in 1985. Since then it has become the most popular method for colorimetric detection and quantitation of total protein. One particular benefit is that, unlike other methods available at that time (e.g., Bradford and Lowry assays), the BCA Protein Assay is compatible with samples that contain up to 5% surfactants (detergents). In addition, the BCA Assay responds more uniformly to different proteins than the Bradford method.
The BCA Protein Assay combines the protein-induced biuret reaction (see above) with the highly sensitive and selective colorimetric detection of the resulting cuprous cation (Cu1+) by bicinchoninic acid (BCA). Thus, two steps are involved. First is the biuret reaction, whose faint blue color results from the reduction of cupric ion to cuprous ion. Second is the chelation of BCA with the cuprous ion, resulting in an intense purple color. The purple colored reaction product is formed by the chelation of two molecules of BCA with one cuprous ion. The BCA/copper complex is water-soluble and exhibits a strong linear absorbance at 562 nm with increasing protein concentrations. The purple color can be measured at any wavelength between 550 nm and 570 nm with minimal (less than 10%) loss of signal. The BCA reagent is approximately 100 times more sensitive (lower limit of detection) than the biuret reagent.
Figure 3. The reaction of BCA with cuprous ion. Two molecules of BCA bind to each molecule of copper that had been reduced by a peptide-mediated biuret reaction.
The reaction that leads to BCA color formation as a result of the reduction of Cu2+ is especially influenced by the presence of three particular amino acid residues in proteins: cysteine/cystine, tyrosine and tryptophan. Apparently these amino acids enhance copper reduction independently in the biuret reaction, thereby causing formation of a colored BCA-Cu1+ chelate. However, studies performed with di- and tripeptides indicate that these produce more color than can be accounted for by the four individual BCA-reactive amino acids. In other words, the peptide backbone (and thus the total amount of protein) is the major contributor to the reduction of copper in the biuret reaction and color development in the BCA assay. Slight protein-to-protein variation in the BCA protein assay results from differences among proteins in composition with respect to these three amino acids.
The binding of BCA to cuprous ion effectively removes the weakly chelated peptides of the biuret reaction. Those peptide groups are then free to bind another molecule of cupric ion. Therefore, if bicinchoninic acid and copper are present in large excess (as they always are in BCA protein assay reagents), the protein assay does not reach an end-point. In addition, the rate of BCA color formation is dependent on the incubation temperature. Consequently, the key to obtaining accurate results with the BCA assay method is to assay standards and unknown samples simultaneously so that they both receive identical incubation time and temperature. Assuming that the assay is performed in this way, the assay characteristic enables one to speed development or wait longer for desired colored development as needed.
Substances that reduce copper will also produce color in the BCA assay, thus interfering with the accuracy of the protein quantitation. Reagents that chelate the copper also interfere by reducing the amount of BCA color produced with protein. Certain single amino acids (cysteine or cystine, tyrosine and tryptophan) will also produce color and interfere in BCA assays. Tech Tips and specialized versions of BCA protein assay products address one or another of these sample- incompatibility issues.
The latest advance in colorimetric protein assays is the Pierce Rapid Gold BCA Protein Assay which retains the high sensitivity and linearity of the traditional BCA assay, but provides ready-to-read results within 5 minutes with room temperature (RT) incubation. In contrast, with traditional BCA assays—depending on the protocol—incubation times range from 30 minutes to 2 hours with temperatures ranging from 37₀ C to 60 ₀ C.
Like the conventional BCA assay, the Pierce Rapid Gold BCA Protein assay involves the reduction of copper by proteins in an alkaline medium (biuret reaction) to produce sensitive and selective colorimetric detection by a new copper chelator. The amount of reduced copper is proportional to the amount of protein present in the solution. The selective copper chelator forms an orange-gold–colored complex that strongly absorbs light at 480 nm. This representative data compares the performance of the conventional and newly adapted BCA protein assays.
Figure 4. Protein concentration determination in lysates using the standard Pierce BCA Protein Assay and Pierce Rapid Gold BCA Protein Assay. Both assays were conducted according to the manufacturer’s protocols, in a microplate format. For the standard BCA assay, 25 μL of sample was added to 200 μL of BCA working reagent and incubated for 30 minutes at 37°C. For the Pierce Rapid Gold BCA Protein Assay, 20 μL of sample was added to 200 μL of the Pierce Rapid Gold BCA working reagent and incubated at room temperature for 5 minutes.
The Lowry protein assay is named after Oliver H. Lowry, who developed and introduced it (Lowry, et al., 1951). It offered a significant improvement over previous protein assays and his paper became one of the most cited references in life science literature for many years. The Modified Lowry Protein Assay uses a stable reagent that replaces two unstable reagents described by Lowry. Essentially, the assay is an enhanced biuret assay involving copper chelation chemistry.
Although the mechanism of color formation for the Lowry assay is similar to that of the BCA protein assay, there are several significant differences between the two. The exact mechanism of color formation in the Lowry assay remains poorly understood. The assay is performed in two distinct steps. First, protein is reacted with alkaline cupric sulfate in the presence of tartrate for 10 minutes at room temperature. During this incubation, a tetradentate copper complex forms from four peptide bonds and one atom of copper (this is the "biuret reaction"). Second, a phosphomolybdic-phosphotungstic acid solution is added. This compound (called Folin-phenol reagent) becomes reduced, producing an intense blue color. It is believed that the color enhancement occurs when the tetradentate copper complex transfers electrons to the phosphomolybdic-phosphotungstic acid complex. The blue color continues to intensify during a 30 minute room temperature incubation. It has been suggested that during the 30 minute incubation, a rearrangement of the initial unstable blue complex leads to the stable final blue colored complex which has higher absorbance (Lowry, et al. 1951; Legler, et al. 1985).
The final blue color is optimally measured at 750nm, but it can be measured at any wavelength between 650 nm and 750 nm with little loss of color intensity. It is best to measure the color at 750 nm since few other substances absorb light at that wavelength.
Figure 5. Protocol summary for the Modified Lowry Protein Assay.
For small peptides, the amount of color increases with the size of the peptide. The presence of any of five amino acid residues (tyrosine, tryptophan, cysteine, histidine and asparagine) in the peptide or protein backbone further enhances the amount of color produced because they contribute additional reducing equivalents to further reduce the phosphomolybdic/phosphotungstic acid complex. With the exception of tyrosine and tryptophan, free amino acids will not produce a colored product with the Lowry reagent; however, most dipeptides can be detected. In the absence of any of the five amino acids listed above in the peptide backbone, proteins containing proline residues have a lower color response with the Lowry reagent due to the amino acid interfering with complex formation.
Unlike in the BCA assay, the secondary binding step in the Lowry method does not involve detachment of the peptide-copper chelate. Therefore, the Lowry method is effectively an end-point assay. Although it is always best to include internal standards in any protein assay, it is possible to obtain reasonable protein estimations with this assay method by comparing to a previously-plotted standard curve.
The protocol requires that the Folin phenol reagent be added to each tube precisely at the end of the ten minute incubation. At the alkaline pH of the Lowry reagent, the Folin phenol reagent is almost immediately inactivated. Therefore, it is best to add the Folin phenol reagent at the precise time while simultaneously mixing each tube. Because this is somewhat cumbersome, some practice is required to obtain consistent results. This also limits the total number of samples that can be assayed in a single run. If a ten second interval between tubes is used, the maximum number of tubes that can be assayed within ten minutes is sixty (10 seconds/tube x 60 tubes = 600 seconds or 10 minutes).
The Lowry assay reagent forms precipitates in the presence of detergents or potassium ions. When potassium ions are the cause, the problem can sometimes be overcome by centrifuging the tube and measuring the color in the supernatant. Most surfactants cause precipitation of the reagent even at very low concentrations. One exception is sodium dodecyl sulfate (SDS), which is compatible with the reagent at concentrations up to 1% in the sample. Chelating agents interfere by binding copper and preventing formation of the copper peptide bond complex. Reducing agents and free thiols also interfere by reducing the phosphotungstate-phosphomolybdate complex, immediately forming an intensely blue colored product upon their addition.
The Modified Lowry Protein Assay Reagent must be refrigerated for long-term storage, but it must be warmed to room temperature before use. Using cold Modified Lowry Protein Assay Reagent will result in low absorbance values.
Use of Coomassie G-250 dye as a colorimetric reagent for the detection and quantitation of total protein was first described by Dr. Marion Bradford in 1976 (Bradford, 1976). Thermo Scientific Pierce Coomassie (Bradford) Assays are variants of the reagent first reported by Bradford.
Figure 6. Chemical structure of Coomassie dye. Formulated in a low-pH phosphoric acid buffer, this colloidal form of Coomassie dye is the basis for Bradford protein assay reagents.
In the acidic environment of the reagent, protein binds to the Coomassie dye. This results in a spectral shift from the reddish/brown form of the dye (absorbance maximum at 465 nm) to the blue form of the dye (absorbance maximum at 610 nm). The difference between the two forms of the dye is greatest at 595 nm, so that is the optimal wavelength to measure the blue color from the Coomassie dye-protein complex. If desired, the blue color can be measured at any wavelength between 575 nm and 615 nm. At the two extremes (575 nm and 615 nm) there is a loss of about 10% in the measured amount of color (absorbance) compared to that obtained at 595 nm.
Development of color in Bradford protein assays is associated with the presence of certain basic amino acids (primarily arginine, lysine and histidine) in the protein. Van der Waals forces and hydrophobic interactions also participate in the binding of the dye by protein. The number of Coomassie dye ligands bound to each protein molecule is approximately proportional to the number of positive charges found on the protein. Free amino acids, peptides and low molecular weight proteins do not produce color with Coomassie dye reagents. In general, the mass of a peptide or protein must be at least 3000 daltons to be detectable with this reagent. In some applications this can be an advantage. For example, the Coomassie Protein Assay has been used to measure "high molecular weight proteins" during fermentation in the beer brewing industry.
Coomassie dye binding assays are the fastest and easiest to perform of all protein assays. The assay is performed at room temperature and no special equipment is required. Standard and unknown samples are added to tubes containing preformulated Coomassie assay reagent and the resultant blue color is measured at 595 nm following a short room temperature incubation. The Coomassie dye-containing protein assays are compatible with most salts, solvents, buffers, thiols, reducing substances and metal chelating agents encountered in protein samples.
The main disadvantage of Coomassie based protein assays is their incompatibility with surfactants at concentrations routinely used to solubilize membrane proteins. In general, the presence of a surfactant in the sample, even at low concentrations, causes precipitation of the reagent. In addition, the Coomassie dye reagent is highly acidic, so proteins with poor acid-solubility cannot be assayed with this reagent. Finally, Coomassie reagents result in about twice as much protein-to-protein variation as copper chelation-based assay reagents.
Figure 7. Absorbance spectra for protein standards in the Thermo Scientific Pierce Coomassie Plus (Bradford) Protein Assay. Protein standards are 0, 125, 250, 500, 750, 1000, 1500 and 2000 µg/mL of bovine serum albumin, respectively. The 2000 µg/mL line is drawn thicker than the others to orient the sequence. Notice that an inverse relationship between protein concentration and absorbance occurs below 525 nm (maximum at 465 nm).
The ready-to-use liquid Coomassie dye reagents should be mixed gently by inversion just before use. The dye in these liquid reagents forms loose aggregates within 60 minutes in undisturbed solutions. Gentle mixing of the reagent by inversion of the bottle will uniformly disperse the dye and ensure that aliquots are homogeneous. After binding to protein, the dye also forms protein-dye aggregates. Fortunately, these protein-dye aggregates can be dispersed easily by mixing the reaction tube. This is common to all liquid Coomassie dye reagents. Because these aggregates form relatively quickly, it is also best to routinely mix (vortex for 2-3 seconds) each tube or plate just before measuring the color.
Introduced in 2008, the Thermo Scientific Pierce 660 nm Protein Assay is a dye-based reagent that offers the same convenience as Coomassie-based assays while overcoming several of their disadvantages. In particular, the Pierce 660 nm Assay is compatible with most detergents and produces a more linear response curve.
The detailed assay chemistry is proprietary, but the essential mechanism can be summarized as follows. The reagent contains a proprietary dye-metal complex in an acidic buffer. The dye-metal complex binds to protein in the acidic condition, causing a shift in the dye's absorption maximum, which is measured at 660nm. The reagent is reddish-brown and changes to green upon protein binding.
Figure 8. Absorption maximum of the 660 nm Assay Reagent-metal complex shifts proportionally upon binding to BSA. The absorption spectra were recorded for the Pierce 660 nm Protein Assay Reagent from 340 to 800 nm using a spectrophotometer. Protein in the presence of the reagent-metal complex produces a significant absorbance shift at a wavelength of 660 nm.
The color produced in the assay is stable and increases in proportion to a broad range of increasing protein concentrations. The color change is produced by the deprotonation of the dye at low pH facilitated by protein-binding interactions through positively charged amino acid groups and the negatively charged deprotonated dye-metal complex.
The assay binds to proteins in a manner similar to Coomassie dye. Thus, it has similar protein-to-protein variability to Coomassie (Bradford) assay methods. However, unlike Coomassie-based assays, the Pierce 660 nm Protein Assay is fully compatible with nonionic detergents typically used in protein samples. In fact, when used with the Ionic Detergent Compatibility Reagent (IDCR), the Pierce 660 nm Assay is also compatible with sample containing Laemmli SDS sample buffer with bromophenol blue and other buffers containing common ionic detergents.
Fluorescence-based protein quantification detection methods provide superior sensitivity, which means that less protein sample is used for quantitation, leaving more samples available for experiments. For the assays described below, few steps are required and timing is not critical, as signal duration is typically hours, so the assays can be adapted for automated handling in high-throughput applications. The fluorescence signal can be detected using a fluorometer or microplate reader.
Fluorescent protein assays typically provide researchers with greater sensitivity than what can be measured with colorimetric protein assays. The Thermo Scientific Quanti-iT, Qubit and NanoOrange protein assays are based on the dye molecule binding to detergent coating on proteins and hydrophobic regions of proteins, and resulting in fluorescence while unbound dye is non-fluorescent. The detection wavelength for all three of these assays is 470/570 nm. The first two produce a quasi-linear standard curve from 0.5 – 4 µg in a 200 µl sample, with a sample volume of 1 – 20 µl, while NanoOrange has a lower sensitivity range of 10 ng/ml, ranging up to 10 µg/ml. For detection of lipoproteins or proteins in a complex lipid environment, the CBQCA Protein Quantitation Kit may be utilized. These representative data show a typical standard curve produced using a fluorescent protein assay kit.
Figure 9. Low protein-to-protein variation in the Qubit Protein Assay.
The NanoOrange Protein Quantitation Kit contains a very sensitive and easy assay for protein quantitation, with detection as low as 10 ng/mL of protein in solution. This fluorescent dye is suitable for use with spectrofluorometers and microplate readers. For detection of lipoproteins or proteins in a complex lipid environment, check out our CBQCA Protein Quantitation Kit.
The CBQCA Protein Quantitation Kit is a very sensitive assay for quantitating proteins in solution, capable of detection as low as 10 ng of protein per mL. Similar in sensitivity to our NanoOrange protein quantitation reagent (N-6666), CBQCA is better suited for accurate quantitation of proteins in the presence of lipids, membrane fractions or detergents, and for lipoproteins and small peptides. This assay is based on the reaction of the dye with primary amine groups in the presence of cyanide or thiols, causing it to become fluorescent. Unreacted dye remains non-fluorescent. This protein assay can detect proteins in the range of 10 ng/ml to 150 µg/ml.
Finally, the EZQ Protein Quantitation Kit provides a fluorescence-based protein assay that facilitates fast quantitation of protein samples prepared for gel electrophoresis. The assay is based on the dye binding electrostatically to basic amino acids, supplemented by additional hydrophobic interactions, resulting in fluorescence that can be read at 280 nm and 450/618 nm.
Fluorometers are instruments that measure the intensity of the fluorescent signal from dyes attached to biological molecules as well as naturally fluorescent molecules based on signature excitation (Ex) and emission (Em) wavelengths. Fluorometers were designed to quantify, detect and monitor analytes and their reactions with a high degree of sensitivity and specificity.
The Invitrogen Qubit Fluorometer is a benchtop device that accurately measures DNA, RNA, and protein using the highly sensitive Qubit quantitation assays. In conjunction with optimized algorithms, the Qubit 3 Fluorometer employs fluorescent dyes that only produce signal when bound to the target of interest, thereby minimizing the effects of contaminants— including degraded DNA or RNA—on experimental results.
For Research Use Only. Not for use in diagnostic procedures.