Knowing exactly how much protein is in a sample — before running a gel, setting up an enzyme assay, or normalizing a Western blot — is one of the most routine measurements in a biochemistry or molecular biology lab. UV spectrophotometry at 280 nm makes this fast and non-destructive, but getting an accurate number depends on choosing the right extinction coefficient and understanding what the Beer-Lambert law is actually measuring.
Why A280 measures protein concentration
Proteins absorb ultraviolet light most strongly around 280 nanometers because of the aromatic side chains of tryptophan and tyrosine (and, more weakly, phenylalanine and disulfide bonds). Under the Beer-Lambert law, absorbance is directly proportional to concentration for a fixed path length, so measuring A280 lets you back-calculate concentration once you know the sample's extinction coefficient — the proportionality constant that links absorbance to how much protein is actually present.
Molar vs. mass extinction coefficients
Extinction coefficients for proteins are published in two different conventions. A molar extinction coefficient (ε_M, in M⁻¹cm⁻¹) describes absorbance per mole of protein and requires the protein's molecular weight to convert the result into a mass concentration (mg/mL). A mass extinction coefficient (often labeled ε(0.1%) or E1%₂₈₀, in (mg/mL)⁻¹cm⁻¹) describes absorbance per unit mass concentration directly, skipping the molecular-weight step entirely. Mixing up the two conventions — plugging a molar value into a mass-based formula or vice versa — is one of the most common sources of an order-of-magnitude error in protein quantification.
Why a theoretical extinction coefficient can be wrong
Molar extinction coefficients are usually calculated theoretically from a protein's amino-acid sequence — counting tryptophan, tyrosine, and cystine (disulfide-bonded cysteine) residues and summing their known per-residue contributions. This works well for most well-folded, soluble proteins, but it can under- or over-estimate absorbance for proteins with unusual folding, bound cofactors, or few aromatic residues. Fusion tags, denaturants, and pH can also shift the true extinction coefficient away from the theoretical value.
When to use a standard curve instead
Colorimetric assays like Bradford (Coomassie dye binding) and BCA (bicinchoninic acid, copper reduction) don't rely on a protein's own UV absorbance at all — they measure a color change proportional to total protein mass, calibrated against a standard curve built from known concentrations of a reference protein, usually bovine serum albumin (BSA). These methods are less sensitive to a specific protein's amino-acid composition than A280, making them a good cross-check — or the primary method — for proteins with few tryptophan/tyrosine residues, or for crude lysates containing many different proteins at once.
Limitations of A280-based estimates
A280 readings are fast but assume the sample is free of other UV-absorbing contaminants — nucleic acids in particular absorb strongly at 260 nm and can bleed into the 280 nm reading, inflating the apparent protein concentration. Turbid or particulate samples scatter light and also inflate the reading. For the most rigorous quantification, cross-check an A280-based estimate against a colorimetric assay (Bradford/BCA) or, where absolute accuracy matters most, an amino-acid analysis.