Protein Concentration Calculator
Compute protein concentration from absorbance at 595 nm using a linear standard curve. The standard back-calculation for Bradford and similar dye-binding assays in molecular biology labs.
Last updated: May 2026
Compare with similar
About this calculator
Protein quantification by colorimetric assay (Bradford, BCA, Lowry) relies on a dye that binds protein and shifts its absorbance spectrum proportionally to protein concentration. A standard curve is generated by measuring absorbance for known concentrations of a reference protein (usually BSA — bovine serum albumin) and fitting a linear regression A = slope × concentration + intercept. To find an unknown sample's concentration, rearrange to concentration = (A − intercept) / slope. This calculator does exactly that. Variables: absorbance is the sample's measured A595; slope is the standard-curve slope (Δabsorbance per Δconcentration, typically in units of A595 per μg/ml); intercept is the y-intercept of the standard curve at zero protein. Edge cases: slope must be non-zero (no curve, no quantification); a sample absorbance below the intercept gives a negative computed concentration, which is meaningless — likely the sample is below the assay's detection limit. Real-world protocol: Bradford assay covers ~1–25 μg/ml in standard format and ~25–1500 μg/ml in micro format; BCA covers ~20–2000 μg/ml; Lowry covers ~5–100 μg/ml. The Bradford response is non-linear above ~25 μg/ml (the curve flattens), so samples beyond this should be diluted before measurement. Each protein binds the dye slightly differently — IgG, lysozyme, and other basic proteins overestimate at given concentrations because they bind more dye per mg than BSA. For accurate quantification of a single specific protein, use that protein as the standard rather than BSA. Detergents (especially SDS) and reducing agents (DTT, β-mercaptoethanol) interfere with Bradford; BCA tolerates SDS but is sensitive to reducing agents. The choice of assay depends on what your sample buffer contains.
How to use
Example 1 — Standard back-calculation. You measure a sample at A595 = 0.45. Your standard curve has slope = 0.002 A595 per (μg/ml) and intercept = 0.05. Enter Absorbance = 0.45, Slope = 0.002, Intercept = 0.05. Concentration = (0.45 − 0.05) / 0.002 = 0.40 / 0.002 = 200 μg/ml. ✓ The sample contains 200 μg/ml of protein — well within the Bradford linear range, so this number is reliable. Example 2 — Low-concentration sample near detection limit. Sample reads A595 = 0.08, standard curve has slope = 0.002 and intercept = 0.06 (a relatively high intercept can indicate buffer interference). Enter 0.08, 0.002, 0.06. Concentration = (0.08 − 0.06) / 0.002 = 10 μg/ml. ✓ The sample is at the very low end of Bradford's detection range — measurement noise dominates here, and the result has high relative uncertainty (likely ±20%). Repeat with a more concentrated sample if possible, or switch to a more sensitive assay (Bradford micro-format, BCA, or fluorescence-based methods like Qubit).
Frequently asked questions
Which protein assay should I use — Bradford, BCA, or Lowry?
Bradford is the most popular for rapid, simple quantification: fast (5 min), cheap, low-volume, and forgiving. Downsides: incompatible with detergents (SDS) and gives different responses to different proteins, especially highly basic ones (IgG, histones). BCA is more compatible with detergents and reducing agents (a key advantage for membrane-protein work), more linear over a wider range, but slower (30 min incubation at 37 °C) and more expensive. Lowry is the historical gold standard, very sensitive but slow (40 min), uses more reagents, and is interfered with by many common chemicals. UV absorbance at 280 nm (no dye, just protein's intrinsic absorbance) is the fastest method but works only for relatively pure samples and depends on the specific protein's tryptophan/tyrosine content. Choose based on sample composition and required throughput; for most general lab work, Bradford is fine if your buffer is compatible.
How do I generate a good standard curve?
Use a high-purity reference protein (BSA from Sigma or similar, at certified concentration). Prepare 5–7 standards spanning the expected sample concentration range — e.g., 0, 5, 10, 25, 50, 100, 200 μg/ml for Bradford macro-format. Run each standard in triplicate to estimate measurement noise. Plot absorbance vs concentration; fit a linear regression and check the R² — you want R² > 0.99 for reliable back-calculation. If the highest standards curve away from linearity (typical for Bradford above 25 μg/ml or BCA above 1500 μg/ml), exclude them and fit only the linear range. The standard curve must be re-prepared every run because the dye reagent's sensitivity drifts. Always include a blank (no protein, just buffer + reagent) and subtract its absorbance — this should equal your intercept value approximately.
Why does my sample give wildly different concentrations between Bradford and BCA?
Different dye-binding chemistries give different responses to different proteins. Bradford's Coomassie dye binds preferentially to basic and aromatic residues; BCA reduction is sensitive to peptide bonds and to specific amino acids (cysteine, tryptophan, tyrosine). A protein rich in basic residues (lysozyme, histones, IgG) will read 30–100% higher in Bradford than in BCA. A protein rich in cysteine will read higher in BCA than in Bradford. If you need absolute concentration of a specific protein, use that protein as your standard rather than BSA; or use amino acid analysis (gold standard but expensive). For relative comparisons of similar protein populations across conditions, the choice matters less because the bias is consistent.
What are the most common mistakes people make with protein quantification?
The first is not making fresh standard curves — Bradford dye and BCA reagents lose activity over weeks of storage, so an old curve under-reports protein. The second is using BSA standards while quantifying a very different protein and reporting the result as accurate; this is fine for "BSA-equivalent" reporting but misleading for absolute amounts. The third is reading at the wrong wavelength — Bradford is 595 nm, BCA is 562 nm, Lowry is 750 nm; mixing them produces nonsense. The fourth is sample buffer interference — DTT, EDTA, SDS, glycerol, and Triton all affect different assays differently; check compatibility tables before assuming the result is valid. The fifth is sample concentration outside the linear range; very low samples have signal-to-noise problems, very high samples saturate the dye and underestimate concentration. Always dilute very concentrated samples and re-measure to confirm linearity.
When should I not use this calculator?
Skip it when the assay's response is non-linear at the sample concentration (typically Bradford above ~25 μg/ml in macro format) — dilute the sample and re-measure in the linear range instead. Don't use it when sample buffer contains components known to interfere with your assay (SDS for Bradford, reducing agents for BCA, copper-chelators for Lowry); the absorbance reading is invalid regardless of the curve. It's the wrong tool for very pure protein samples where direct UV absorbance at 280 nm with the known extinction coefficient is more accurate; the dye-based methods add unnecessary error. Avoid it for samples with absorbing impurities (haemoglobin, cytochromes, NADH) that contribute background at the measurement wavelength. Finally, don't use it when measurement is below the assay's detection limit (typically ~5 μg/ml for Bradford macro); switch to a more sensitive method (Bradford micro, BCA, Qubit) or concentrate the sample.