Biuret test
The Biuret test is a simple, colorimetric chemical assay employed in biochemistry and analytical chemistry to detect the presence of proteins and peptides in a sample through the identification of peptide bonds.[1][2] It involves adding a biuret reagent—typically consisting of copper(II) sulfate (CuSO₄), sodium hydroxide (NaOH), and sodium potassium tartrate—to the sample under alkaline conditions, which results in the formation of a violet or purple-colored complex if peptide bonds are present, with the intensity of the color proportional to the protein concentration.[3][4] The test is named after biuret (a condensation product of urea), which produces the same color reaction, and its underlying principle was first described in 1833 by the chemist Ferdinand Rose during studies of urea derivatives; in Poland, it is also known as Piotrowski's test after Gustaw Piotrowski who applied it to physiological samples.[5] The principle of the Biuret test hinges on the coordination of Cu²⁺ ions with the unprotonated amide nitrogens in peptide bonds (–CO–NH–), forming a tetrahedral chelate complex that absorbs light at approximately 540 nm, allowing for both qualitative observation and quantitative measurement via spectrophotometry.[1][2] This reaction requires at least two peptide bonds (as in tripeptides or larger polypeptides) for detection and is relatively non-specific to individual amino acids, making it suitable for total protein estimation rather than distinguishing protein types; however, it can yield false positives with certain compounds like free histidine due to similar coordination sites.[4][2] The reagent's alkaline environment deprotonates the peptide bonds, enhancing their reactivity with copper, and the tartrate acts as a stabilizer to prevent copper hydroxide precipitation.[3]Overview
Definition and Purpose
The Biuret test is a chemical assay that utilizes copper(II) ions in an alkaline solution to detect the presence of peptide bonds in proteins, resulting in the formation of a violet-colored complex.[6] This colorimetric reaction specifically targets the amide linkages formed between amino acids, distinguishing proteins from free amino acids or other non-peptidic compounds.[7] The primary purpose of the Biuret test is to qualitatively confirm the presence of proteins in biological samples, such as tissues, fluids, or extracts, making it a fundamental tool in biochemical analysis for identifying proteinaceous materials.[6] It also serves a secondary quantitative role, where the intensity of the violet color can be measured spectrophotometrically to estimate protein concentrations, providing a reliable method for routine laboratory assessments in clinical and research settings.[8] Peptide bonds, which are the covalent amide bonds linking the carboxyl group of one amino acid to the amino group of another, are essential for the test's specificity; the reaction requires at least two such bonds (as found in tripeptides or larger polypeptides) for detection, with optimal sensitivity for proteins containing three or more.[6] The standard Biuret test typically detects proteins in the concentration range of 5–160 mg/mL, depending on the protocol and sample volume.[9]Historical Development
The biuret reaction was first observed in 1833 by German chemist Heinrich Rose during studies of metallic salt solutions with protein-containing materials such as egg albumen and bovine blood serum, noting the formation of a violet color upon addition of copper sulfate and potassium hydroxide.[5] The compound biuret (NH₂CONHCONH₂), an impurity arising from urea condensation, was isolated in 1848 by Gustav Heinrich Wiedemann through pyrolysis of urea; its red-violet reaction with alkaline cupric solutions paralleled that of proteins, leading to the test's naming in recognition of this structural and reactivity similarity.[5] The reaction was independently rediscovered in 1857 by Polish physiologist Gustaw Piotrowski, who applied it to physiological samples; consequently, in Poland, the test is known as Piotrowski's test.[10] Initial applications to protein analysis emerged in 1873 when Ritthausen and Pott utilized the reaction to investigate plant proteins, followed by Hofmeister's 1879 adaptation for quantitative estimation of proteins and peptones.[5] By the early 20th century, the term "biuret test" gained widespread recognition in biochemical literature for detecting peptide bonds.[5] The test's adoption accelerated in protein assays during the 1940s, with Kingsley's 1942 formulation of a stable, single-reagent system enabling reliable serum protein measurements via colorimetry.[5] Further refinement came in 1949 through Gornall et al.'s comprehensive evaluation, which standardized the reagent composition without iodide for improved accuracy and stability in clinical settings.[11] Following the 1950s, integration with spectrophotometry at 540 nm absorbance allowed for precise quantitative analysis, solidifying the biuret test's role in routine biochemical protocols.[5]Chemical Principle
Reaction Mechanism
The Biuret test involves the formation of a coordination complex between copper(II) ions (Cu²⁺) and the peptide bonds of proteins in an alkaline medium. The hydroxide ions (OH⁻) from the reagent, typically provided by sodium hydroxide (NaOH) or potassium hydroxide (KOH), deprotonate the nitrogen atoms in the peptide linkages (-CO-NH-), freeing the lone pairs on these nitrogens to act as electron donors for coordination with Cu²⁺. This deprotonation is essential, as it enables the otherwise neutral peptide nitrogens to bind effectively to the metal ion.[7][12] The coordination occurs when Cu²⁺ ions bind to four deprotonated nitrogen atoms from adjacent peptide bonds, forming a square planar violet complex. This structure requires at least two peptide bonds (corresponding to tripeptides or larger polypeptides), as shorter chains like single amino acids or dipeptides lack the necessary number of free -CO-NH- groups for stable complexation. The test's specificity stems from this requirement for vicinal peptide linkages; non-peptide amides or unrelated nitrogen-containing compounds do not react unless they structurally mimic the biuret moiety (NH₂-CO-NH-CO-NH₂).[9][12] The characteristic mauve to purple color of the complex results from a charge-transfer interaction between the Cu²⁺ center and the peptide ligands, leading to absorption of visible light at approximately 540 nm. The intensity of this color is proportional to the number of peptide bonds present.[9] A simplified equation representing the reaction mechanism is: \ce{2 Cu^{2+} + Protein (with \geq 2 peptide bonds) + 4 OH^- -> [Cu_2(Protein)]^{2+} + 4 H2O} This notation highlights the involvement of two Cu²⁺ ions coordinating with the protein and the consumption of hydroxide ions in deprotonation, with water as a byproduct.[7][13]Biuret Reagent Composition
The Biuret reagent is an alkaline solution primarily composed of 0.15% (w/v) copper(II) sulfate pentahydrate (CuSO₄·5H₂O), 0.6% (w/v) potassium sodium tartrate (also known as Rochelle salt), and approximately 3% (w/v) sodium hydroxide (NaOH) in distilled water. This formulation is commonly used for qualitative detection in biochemical assays.[14][7] Copper sulfate supplies the Cu²⁺ ions that form the violet-colored complex with peptide bonds during the test. Potassium sodium tartrate functions as a chelating agent, binding to Cu²⁺ to maintain its solubility and prevent precipitation of copper hydroxide in the alkaline medium. The alkali (NaOH or KOH) raises the pH above 12, promoting the necessary deprotonation of peptide groups for coordination with Cu²⁺.[13][15][7] Preparation involves dissolving the potassium sodium tartrate in the alkaline solution first, then slowly adding the CuSO₄·5H₂O while stirring, and diluting to the final volume with distilled water to avoid local precipitation. The resulting blue solution is stored in a dark, tightly sealed bottle at room temperature to inhibit photo-oxidation or reduction of the copper ions.[14][5][16] Variations in the reagent include using anhydrous forms of CuSO₄ or tartrate, with proportional water adjustments to achieve equivalent molar concentrations, or opting for commercial pre-mixed solutions that ensure consistency and eliminate preparation errors. The reagent maintains stability for 1–2 months when stored properly; discard if it develops a green tint, signaling copper ion degradation or contamination.[5][3]Experimental Protocol
Materials and Sample Preparation
The Biuret test requires specific laboratory materials to ensure accurate and reproducible protein detection. Essential equipment includes test tubes (such as 13 x 100 mm culture tubes), pipettes capable of dispensing 0.5–2.0 mL volumes, and a spectrophotometer set to measure absorbance at 540 nm for quantitative analysis.[3][17] The core reagent is the Biuret solution, typically prepared or obtained as a ready-to-use formulation containing copper sulfate, sodium potassium tartrate, and sodium hydroxide. Additional items include distilled or deionized water for dilutions and blanks, as well as optional tools like a centrifuge or filter for handling turbid samples.[3][18] Suitable samples for the Biuret test are primarily aqueous solutions containing proteins, such as biological fluids (e.g., serum or urine), tissue homogenates, or extracts from food and beverage matrices like beer. Samples must be solubilized in water or compatible buffers if not already in liquid form; solid or semi-solid materials should be homogenized prior to testing.[17][18] Interfering substances, including chelating agents like EDTA or EGTA, reductive compounds such as DTT, and amine-containing buffers like Tris, should be avoided, as they can disrupt the copper-protein complex formation.[19][20] Sample preparation begins with assessing concentration and clarity to prevent assay inaccuracies. For samples exceeding 10 g/L protein, dilute 1:10 or as needed using distilled water or a neutral buffer to bring concentrations within the linear range of 0.5–10 g/L. Turbid samples require centrifugation at 10,000 × g for 5–10 minutes or filtration to remove debris and particulates. Acidic samples (pH < 7) should be adjusted to pH 8 using 1 M NaOH and incubated for 15 minutes to ensure compatibility with the alkaline Biuret reagent. Typical volumes are 1–2 mL of prepared sample per test, with pH neutrality confirmed before proceeding.[17][3][18] Controls are essential for validation and calibration. Include a negative control using 1 mL of saline or distilled water to establish baseline absorbance, and a positive control prepared from bovine serum albumin (BSA) standards at concentrations like 2 mg/mL or 10 mg/mL to verify reagent performance and construct a standard curve for quantitative tests. Reagent blanks, consisting of water plus Biuret solution, account for background absorbance.[3][17] For low-protein samples (<0.5 g/L), optional precipitation with 5–50% trichloroacetic acid (TCA) followed by redissolution in a smaller volume of water can concentrate the analyte.[17]Step-by-Step Procedure
To perform the standard Biuret test, begin by preparing the sample as outlined in the materials section, ensuring it is a clear solution containing potential peptide bonds, such as protein extracts in aqueous buffer. For quantitative analysis, use the same sample-to-reagent ratio for samples and standards.[3] The procedure involves the following steps:- Pipette 1 mL of the prepared sample into a clean test tube (typically 13 x 100 mm).[3]
- Add an equal volume (1 mL) of Biuret reagent to the test tube containing the sample.[7]
- Gently mix the contents by swirling or shaking the tube to ensure homogeneity without introducing excessive foam.[7]
- Incubate the mixture at room temperature (approximately 20–25°C) for 5–10 minutes; for a faster reaction, heat the tube in a water bath at 37°C for 2–5 minutes.[3][18]
- Observe the color development, which typically occurs within 2–30 minutes after mixing; if needed, gently shake again for uniformity.[7][3]
Results and Analysis
Qualitative Interpretation
The qualitative interpretation of the Biuret test relies on observing color changes in the solution following the addition of the reagent and a brief incubation period. A positive result is indicated by the development of a violet or purple (often described as mauve) coloration, which arises from the formation of a complex between cupric ions and the peptide bonds in proteins containing at least two such bonds. This color change confirms the presence of proteins, such as polypeptides or larger macromolecules, and the intensity of the violet hue generally correlates with the protein concentration in the sample, allowing for rough estimation by eye.[7][9] In contrast, a negative result shows no color change, with the solution retaining the blue hue of the Biuret reagent itself. This outcome typically signifies the absence of proteins or peptides with sufficient peptide bonds, as seen in samples containing only free amino acids, monosaccharides like sugars, or other non-proteinaceous compounds that do not form the requisite complex.[7][24] For accurate visual assessment, the reaction mixture should be compared against positive and negative controls—such as a known protein solution for violet and distilled water for blue—to establish baselines. Viewing the tubes against a white background, like a white tile or paper, enhances contrast and clarity in detecting subtle color shifts. The test's visual detection limit is approximately 5 mg/mL of protein, below which changes may not be reliably discernible by naked eye.[24][9]Quantitative Measurement
The quantitative measurement of protein concentration using the Biuret test relies on spectrophotometric analysis of the purple coordination complex formed with proteins. After the reaction, the absorbance is measured at 540 nm using a UV-Vis spectrophotometer, with a blank (reagent without protein) subtracted to correct for any background absorbance from the reagent itself. This wavelength corresponds to the maximum absorption of the complex, providing a reliable signal proportional to protein content.[3][25] The absorbance follows the Beer-Lambert law, given by A = \epsilon l c where A is the absorbance, \epsilon is the specific absorptivity (approximately 0.2 mL/(mg·cm) for many proteins at 540 nm), l is the optical path length (typically 1 cm in standard cuvettes), and c is the protein concentration in mg/mL. This relationship allows direct correlation between the measured absorbance and protein amount, though the exact \epsilon can vary slightly depending on the protein type.[12] For accurate quantification, a calibration curve is essential and is typically prepared using bovine serum albumin (BSA) as the standard protein, with concentrations ranging from 0 to 10 mg/mL. Serial dilutions of a BSA stock solution are reacted with the biuret reagent under identical conditions to the samples, and their absorbances at 540 nm are plotted against known concentrations to yield a linear standard curve (absorbance vs. concentration). The slope of this curve represents the response factor specific to the assay conditions.[26][3] Once calibrated, the unknown sample concentration is determined by dividing the corrected sample absorbance by the slope of the standard curve, yielding the protein concentration in mg/mL: c = \frac{A_{\text{sample}}}{\text{slope}}. This approach accounts for instrumental and reagent variations. The assay demonstrates linearity over a protein concentration range of 5 to 100 mg/mL; samples exceeding this range must be diluted and re-assayed to ensure measurements fall within the linear portion.[3][25]Applications
Biochemical and Research Uses
In biochemical research, the Biuret test is commonly applied to quantify total protein concentrations in cell lysates and enzyme extracts, offering a straightforward method for evaluating protein recovery from complex biological matrices.[27][28] It is particularly valuable for determining protein levels in fractions derived from gel electrophoresis, facilitating precise sample normalization prior to techniques like SDS-PAGE.[29] The test also plays a key role in educational settings within biochemistry laboratories, where it serves as a foundational demonstration for illustrating the detection of peptide bonds through the characteristic violet color formation in protein-positive samples.[30] This hands-on approach helps students understand protein structure and assay principles without requiring advanced equipment. Furthermore, the Biuret test is frequently integrated into research workflows alongside SDS-PAGE for assessing protein purity and during dialysis procedures to monitor protein retention or buffer exchange efficiency.[31] Its primary advantages include operational simplicity, minimal cost, and specificity for peptide bonds, which avoids interference from nucleic acids such as DNA and RNA in nucleic acid-rich samples.[9] Representative examples of its utility encompass measuring protein yields in recombinant expression systems to optimize production protocols and quantifying protein content in food science analyses, such as in gelatin or peanut extracts.[32][33] The assay's sensitivity typically spans 5–160 mg/mL, suitable for many research-scale determinations.[31]Clinical and Industrial Contexts
In clinical settings, the Biuret test is widely employed for the quantitative measurement of total protein concentrations in serum and urine, aiding in the diagnosis of conditions such as hypo- and hyperproteinemia.[21][34] Low serum protein levels (hypoproteinemia) can indicate malnutrition, liver dysfunction, or nephrotic syndrome, while elevated levels (hyperproteinemia) may signal dehydration or chronic inflammation.[21][35] This assay is integrated into automated clinical chemistry analyzers, enabling rapid and precise analysis with detection limits as low as 0.1–0.15 g/dL, and is classified by the FDA as a Class II medical device under the total protein test system category (product code CEK).[4][36][37] For example, serum total protein measurements contribute to assessing liver and kidney function, where deviations from the normal range of 6.4–8.3 g/dL help evaluate hepatic synthetic capacity or renal protein loss.[38][39] High-throughput implementations of the Biuret method in clinical laboratories support processing hundreds of samples per day through automated systems, enhancing efficiency in routine diagnostics.[4] FDA-approved reagent kits, such as those for the Alinity c analyzer, facilitate this scalability while maintaining compliance with regulatory standards for accuracy and precision.[40][41] In industrial applications, the Biuret test serves as a reliable tool for quality control in the dairy sector, where it quantifies milk protein content to ensure product consistency and compliance with nutritional standards.[42][43] For instance, it detects protein levels in fresh milk and powdered formulations, helping identify adulteration by non-protein additives that dilute nutritional value.[44][45] In the brewing industry, the assay assesses soluble protein in wort and beer, informing haze stability and foam quality during production.[46] Pharmaceutical formulations also utilize the Biuret method for verifying protein concentrations in biopharmaceutical products, ensuring batch-to-batch uniformity in therapeutic protein-based drugs.[47] Regulatory frameworks endorse the Biuret test in these contexts; FDA-cleared kits support clinical protein assays, while the method aligns with guidelines for food protein determination in nutritional labeling and analysis.[37][33] The World Health Organization recognizes spectrophotometric approaches like Biuret for evaluating protein in nutritional assessments, particularly in food safety and quality monitoring.[33]Variants and Alternatives
High-Sensitivity Modifications
To achieve lower detection limits than the standard Biuret test, which relies on the formation of a violet complex between Cu²⁺ ions and peptide bonds absorbing at 540 nm, several modifications enhance sensitivity by amplifying the colorimetric signal or improving compatibility with challenging samples.[48] The bicinchoninic acid (BCA) assay represents a key high-sensitivity variant of the Biuret test, where proteins in alkaline medium reduce Cu²⁺ to Cu⁺ via peptide bonds, and bicinchoninic acid then chelates the Cu⁺ to form a highly stable purple complex absorbing at 562 nm.[48] This chelation mechanism extends the Biuret reaction by providing greater color intensity and stability compared to the original tartrate-complexed Cu²⁺, enabling detection in the range of 0.5–20 µg/mL protein.[48] The assay's sensitivity arises from the selective reduction step, which is proportional to protein concentration, and it exhibits low variability across different protein types.[48] Another prominent modification is the Lowry assay, which integrates the Biuret reaction with the Folin-Ciocalteu reagent to further enhance detection. In this method, proteins first react with Cu²⁺ in alkaline conditions to produce Cu⁺, which then reduces the phosphomolybdate-phosphotungstate complex in the Folin-Ciocalteu reagent, yielding a blue color measured at 750 nm. This dual-step amplification allows for protein quantification in the 5–100 µg/mL range, offering approximately 100-fold greater sensitivity than the standard Biuret test alone. Modified versions of the Lowry assay, such as those optimizing reagent stability and incubation times, maintain this range while reducing interferences from common laboratory contaminants. Additional variants include adaptations of the Biuret reaction for solubilizing membrane proteins using surfactants like SDS or Triton X-100, which are compatible at concentrations up to 5% in the BCA format without disrupting the Cu⁺ chelation.[9] Microplate-based implementations of these assays further improve sensitivity and throughput by minimizing sample volumes (e.g., 10–25 µL per well) while preserving the linear detection range.[49] These high-sensitivity modifications provide 10–100 times greater detection limits than the standard Biuret test and excel in detergent compatibility, making them suitable for complex samples like cell lysates or solubilized membrane fractions.[48]Comparison to Other Assays
The Biuret test, which detects peptide bonds in proteins through a colorimetric reaction with copper ions, offers distinct advantages and limitations when compared to other common protein quantification assays such as the Bradford (Coomassie Brilliant Blue), Lowry, and Kjeldahl methods.[50][51] Compared to the Bradford assay, the Biuret test exhibits lower sensitivity, detecting proteins in the range of 0.5–10 mg/mL, whereas Bradford can quantify down to 1–100 µg/mL.[51][50] However, Biuret is more robust against interferences from detergents like SDS or Triton X-100, which can disrupt Bradford's dye-binding to basic and aromatic amino acid residues, leading to variability based on protein composition.[52][50] This makes Biuret preferable for samples containing surfactants, though Bradford remains faster (5 minutes versus Biuret's 10–30 minutes) and suitable for rapid screening of dilute solutions.[52][53] In contrast to the Lowry assay, which combines Biuret's copper reaction with Folin-Ciocalteu reagent for enhanced color development, the standard Biuret test is simpler and quicker, requiring only a single reagent addition and avoiding the multi-step process that takes up to 40 minutes for Lowry.[51][50] Biuret also shows fewer interferences from reducing agents (e.g., DTT, β-mercaptoethanol), EDTA, or cations like K⁺ and Mg²⁺, which can inhibit Lowry's phosphomolybdate reduction step and cause inconsistent results.[51][52] While Lowry achieves higher sensitivity (2–150 µg/mL, approximately 100-fold greater than Biuret), its complexity and susceptibility to sample matrix effects often necessitate careful optimization.[51][53] Relative to the Kjeldahl method, which involves acid digestion to convert nitrogen to ammonium for titration or distillation, the Biuret test provides a direct, non-destructive colorimetric measurement without hazardous chemicals or lengthy digestion (hours versus Biuret's minutes).[51][52] Kjeldahl serves as a reference standard for total nitrogen content, estimating protein via a conversion factor (e.g., 6.25 for many samples), but it lacks specificity by including non-protein nitrogen sources like urea or ammonia, potentially overestimating protein by 10–20% in complex matrices.[51][54] Biuret, while less precise for absolute quantification, excels in quick qualitative or semi-quantitative screening of peptide-containing proteins.[51] Selection of the Biuret test over alternatives depends on sample characteristics: it is ideal for high-concentration proteins (>1 mg/mL) in interference-free matrices, such as purified extracts without detergents or reducing agents, where simplicity and speed are prioritized.[50][53] Bradford or Lowry are better for low-concentration or trace-level detection, while Kjeldahl suits regulatory or total nitrogen assessments despite its labor intensity.[51][54]| Assay | Sensitivity Range | Time Required | Key Interferences | Primary Advantages | Primary Disadvantages |
|---|---|---|---|---|---|
| Biuret | 0.5–10 mg/mL | 10–30 min | Reducing agents, ammonia | Simple, detergent-tolerant | Low sensitivity for dilute samples |
| Bradford | 1–100 µg/mL | 5 min | Detergents, low aromatic residues | Rapid, high sensitivity | Protein composition variability |
| Lowry | 2–150 µg/mL | 30–40 min | Reducing agents, EDTA, cations | High sensitivity, stable endpoint | Complex, multiple interferences |
| Kjeldahl | 0.1–10% N (protein equiv.) | Hours | Non-protein nitrogen (e.g., urea) | Accurate total N reference | Labor-intensive, hazardous |