Protein Concentration Calculator (Bradford · BCA · A280)

How calculations work

• Bradford/BCA: Use a standard curve A = m·C + b. Concentration: C = (A_sample − b)/m. You may paste standards to fit m and b (least squares), with optional zero-intercept.
• A280: Beer–Lambert law, A = ε·l·c. • If using molar ε: c (mg/mL) = (A·MW) / (ε·l). • If using mass ε: c (mg/mL) = A / (ε·l).
• Dilution factor multiplies the calculated concentration.

Understanding Bradford, BCA, and A280

Protein concentration can be estimated in three common ways: colorimetric dye-binding assays (Bradford and BCA) and direct UV absorbance at 280 nm (A280). While all three aim to report the same thing—how much protein is present—they differ in chemistry, dynamic range, sensitivity to reagents, and what assumptions they make. This section explains how each approach works, why a standard curve or extinction coefficient is needed, and how to interpret your results with good laboratory practice.

Bradford & BCA assays

Bradford and BCA are colorimetric assays that convert protein amount into a measurable absorbance. In practice you measure a series of standards (e.g., BSA) and fit a line of the form A = m·C + b, where A is absorbance and C is concentration in mg/mL (or μg/mL). Your unknown is then computed as C = (A_sample − b)/m. In the linear range these assays behave well, but at high concentrations the curve can deviate from linearity; some labs use a quadratic fit for BCA in the upper range. Always include a blank and several standards spaced across the range that covers your samples.

Matrix effects matter. Detergents, chaotropes, and reducing agents can interact with either dye chemistry (e.g., Bradford is more sensitive to detergents; BCA can be affected by strong reducers unless “compatible” protocols are used). If your sample buffer differs from the standard buffer, consider preparing matrix-matched standards (same buffer composition) or diluting samples and standards into a common diluent. Inspect fit quality using R² and residuals; a non-zero intercept can flag background or plate artifacts. If you force the fit through the origin (common when blanks are excellent), verify that residuals stay random and small.

Direct A280 (Beer–Lambert)

Many proteins absorb strongly at 280 nm due to tryptophan and tyrosine residues. The Beer–Lambert law relates absorbance to concentration via A = ε·l·c, with path length l (cm) and extinction coefficient ε. If you know the molar ε (M⁻¹·cm⁻¹) and the molecular weight, you can compute mass concentration as c(mg/mL) = (A·MW)/(ε·l). If you instead have a mass ε (mL·mg⁻¹·cm⁻¹), use c(mg/mL) = A/(ε·l). Microvolume spectrophotometers may auto-correct path length; confirm whether your instrument reports path-corrected A280 or raw absorbance.

A280 assumes your protein’s ε is known and accurate for the exact sequence and redox state (e.g., disulfide formation slightly changes ε). Nucleic acids, heme, or scattering from particulates can inflate A280; if contamination is likely, consider parallel A260/A280 checks or a dye-based assay as an orthogonal method. For best accuracy, measure within 0.1–1.0 absorbance units and apply the correct dilution factor explicitly.

Good reporting practices

• State the method (Bradford, BCA, or A280), wavelength, path length, and dilution factor.
• For assays, report slope, intercept, number of standards, and fit model (linear vs. forced-zero).
• For A280, report ε (molar or mass) and the molecular weight if applicable.
• Flag out-of-range readings and repeat within the validated dynamic range.
• When precision matters, verify with a second method or an orthogonal standard.

Tip: This calculator lets you paste standards to auto-fit the curve, toggle a zero-intercept, and switch between molar or mass extinction coefficients for A280, keeping units and dilution explicit for transparency.