How to Determine Protein Concentration from A280

Reading the absorbance at 280 nm is the fastest, cheapest and most common way to determine the concentration of a purified protein: it is non-destructive, needs no reagents or standard curve, and takes seconds. The whole method rests on one equation — the Beer–Lambert law — and one number you have to know or predict: the molar extinction coefficient ε280. This guide derives ε280 straight from an amino-acid sequence, works the absorbance-to-concentration arithmetic through real examples, converts between the molar (ε) and percent (E¹%) conventions, and names the interferences that silently double your error. The protein concentration calculator runs the same calculation and derives ε280 from your sequence so you can check each step.

Why proteins absorb at 280 nm

Proteins absorb ultraviolet light near 280 nm almost entirely because of their aromatic side chains. Tryptophan is by far the strongest absorber, tyrosine is next, and a disulfide-bonded cystine pair contributes a little. The peptide backbone itself absorbs much lower, near 205 nm, so at 280 nm you are reading the aromatics fairly specifically. The practical consequence is important: a protein with no tryptophan and few tyrosines absorbs weakly and is hard to quantify this way, and one with none of the three cannot be measured at 280 nm at all.

The Beer–Lambert law

Absorbance is proportional to concentration, path length and a per-molecule constant:

A = ε · c · l

where A is the (blank-corrected) absorbance, ε is the molar extinction coefficient in M⁻¹cm⁻¹, c is the molar concentration in mol/L, and l is the path length in cm. Rearranged to solve for concentration, with a dilution factor (DF) folded in:

c (M) = (A280 × DF) / (ε280 × l)

To get a mass concentration, multiply by the molecular weight (MW in Da = g/mol). Because 1 g/L equals 1 mg/mL:

concentration (mg/mL) = c (M) × MW

and concentration (µM) = c (M) × 10⁶. The molecular weight comes from the sequence via the protein molecular weight calculator.

Deriving ε280 from the sequence

You usually do not have a measured ε, but you can predict it from the sequence with the Gill & von Hippel equation, accurate to about ±5% for a folded protein in water:

ε280 = n(Trp)·5500 + n(Tyr)·1490 + n(Cystine)·125 [M⁻¹cm⁻¹]

Residue	ε contribution (M⁻¹cm⁻¹)
Tryptophan (W)	5500
Tyrosine (Y)	1490
Cystine (disulfide-bonded Cys pair)	125

Here n(Cystine) = ⌊n(Cys)/2⌋ when the cysteines are oxidized (paired into disulfides) and 0 when they are reduced (free thiols). Count the residues, multiply, add — that is ε280.

Worked example: ε from a sequence, then concentration

Take ubiquitin (76 residues). Its composition is 0 Trp, 1 Tyr, 0 Cys, so:

ε280 = 0×5500 + 1×1490 + 0×125 = 1490 M⁻¹cm⁻¹
MW from the same sequence = 8564.84 Da (see the how to calculate protein molecular weight guide)
Suppose you read A280 = 1.0 in a 1 cm cuvette, undiluted:
- c = 1.0 / (1490 × 1) = 6.71 × 10⁻⁴ M = 671 µM
- mg/mL = 6.71 × 10⁻⁴ × 8564.84 = 5.75 mg/mL

Ubiquitin’s ε is low precisely because it has a single tyrosine and no tryptophan — a 1.0 absorbance already means a fairly concentrated 5.75 mg/mL solution.

The BSA benchmark: using a known ε

When you have a measured or datasheet ε, skip the sequence step. Bovine serum albumin (BSA) is the classic reference: ε280 = 43,824 M⁻¹cm⁻¹, MW = 66,430 Da.

A280 = 0.66, path 1 cm: c = 0.66 / 43,824 = 1.506 × 10⁻⁵ M
mg/mL = 1.506 × 10⁻⁵ × 66,430 = 1.00 mg/mL

That is the origin of the rule of thumb “a 1 mg/mL BSA solution reads A280 ≈ 0.66.” It is a useful sanity check on any spectrophotometer.

ε versus A(1%,1cm): the percent convention

Many datasheets quote the percent extinction coefficient A(1%,1cm) (also written E¹%) instead of the molar ε. It is the absorbance of a 1% w/v solution — which is 1 g/100 mL = 10 g/L = 10 mg/mL — in a 1 cm path. Convert between the two like this:

A 1% solution has molarity c = 10 g/L ÷ MW g/mol = 10/MW.
So A(1%,1cm) = ε · (10/MW) = 10 · ε / MW, and conversely ε = A(1%,1cm) · MW / 10.

For BSA: A(1%,1cm) = 10 × 43,824 / 66,430 = 6.60 (the literature E¹% for BSA is ≈ 6.67). Note the factor of ten: a 1% (10 mg/mL) solution reads 6.60, while a 1 mg/mL (0.1%) solution reads 0.66 — ten times less. Getting this factor wrong is a common and expensive mistake. If a protocol says “an A280 of 1.0 corresponds to X mg/mL”, then X = MW/ε = 10 / A(1%,1cm).

Cysteine state matters

The 125 M⁻¹cm⁻¹ cystine term applies only to disulfide-bonded cysteine pairs. If your protein is reduced (free thiols, e.g. in DTT), those cysteines contribute essentially nothing at 280 nm. Consider the peptide CCCC (four cysteines, no Trp or Tyr):

Oxidized: 4 Cys form 2 cystines → ε280 = 2 × 125 = 250 M⁻¹cm⁻¹
Reduced: no cystine → ε280 = 0, and A280 cannot quantify it at all

The calculator has a cysteine-state selector for exactly this reason. Match it to the redox state of your sample.

Path length and dilution factor

Two corrections sit in the denominator and numerator of the Beer–Lambert rearrangement and are easy to drop:

Path length. A standard cuvette is 1 cm, but micro-volume instruments (NanoDrop-style) use a shorter effective path that the software usually corrects for automatically — confirm whether your reported A280 is already path-normalized.
Dilution factor. If you diluted the sample to bring it into range, multiply the result by that factor to recover the stock concentration.

For example, a Trp-rich protein with ε280 = 48,275 M⁻¹cm⁻¹ and MW 50,000 Da, read at A280 = 0.75 after a 1:5 dilution:

c = (0.75 × 5) / (48,275 × 1) = 3.75 / 48,275 = 7.77 × 10⁻⁵ M
mg/mL = 7.77 × 10⁻⁵ × 50,000 = 3.88 mg/mL for the original stock
µM = 7.77 × 10⁻⁵ × 10⁶ = 77.7 µM

Interferences — and when not to use A280

A280 is excellent for a pure, aromatic-containing protein, but several things bias it:

Nucleic-acid contamination. DNA and RNA absorb strongly at 260 nm and their tail inflates A280. Check the A260/A280 ratio: pure protein is ≈ 0.57, pure DNA ≈ 1.8. A ratio near or above 1.0 means contamination and an overestimated concentration — use a two-wavelength correction or clean up the sample. (For nucleic acids themselves, the DNA concentration calculator works from A260 instead.)
Reductants and buffer components. DTT, β-mercaptoethanol, imidazole and some detergents absorb near 280 nm. Blank against the exact sample buffer, not water.
Turbidity and scattering. Aggregated or particulate samples raise the whole baseline.
Out-of-range absorbance. Most spectrophotometers are linear only to about A ≈ 1.5–2.0; dilute into range and multiply back.
Aromatic-poor proteins. If ε280 ≈ 0, use a colorimetric assay (Bradford, BCA, Lowry) or a 205 nm peptide-bond reading instead — these need a standard curve and consume sample, but work where UV cannot.

Common mistakes

Dropping the factor of ten between A(1%,1cm) and ε/MW — the single most common error on this page.
Forgetting the dilution factor, so you report the diluted, not the stock, concentration.
Blanking against water instead of the sample buffer, counting reductant absorbance as protein.
Using oxidized ε for a reduced sample (or vice versa) when the protein is cysteine-rich.
Ignoring nucleic-acid contamination and reading a high A280 as pure protein.
Assuming every protein absorbs at 280 nm — an aromatic-poor protein gives a near-zero ε and a meaningless result.

Where this fits in the wider workflow

A280 quantitation sits at the end of a purification and feeds everything downstream. Get the exact mass first with the protein molecular weight calculator, predict the isoelectric point to choose your chromatography condition (the how to calculate protein isoelectric point guide covers it), then measure A280 to find out how much you recovered. Carry the concentration into the dilution calculator or molarity calculator to make working stocks. All three tools live in the protein calculators hub and share the same sequence input, so ε280, MW and concentration stay consistent end to end.