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Protein Concentration from A280 Calculator

Protein concentration from an A280 reading uses the Beer–Lambert law: c = A280 / (ε280 × path length), where ε280 is the molar extinction coefficient at 280 nm. This calculator either derives ε280 from your sequence — ε280 = n(Trp)·5500 + n(Tyr)·1490 + n(Cystine)·125, the Gill & von Hippel formula — or takes a measured ε you enter. It then reports concentration in mg/mL and µM, applying your path length and dilution factor. For example, a protein with ε280 = 43,824 M⁻¹cm⁻¹ and MW 66,430 Da reading A280 = 0.66 in a 1 cm cuvette is 1.00 mg/mL (15.1 µM). If your sequence has no Trp, Tyr or paired Cys, ε280 is zero and A280 cannot quantify it — use Bradford or BCA instead.

Measuring absorbance at 280 nm is the fastest, cheapest and most common way to determine the concentration of a purified protein, because it is non-destructive, needs no reagents and takes seconds. The physics is the Beer–Lambert law: absorbance is proportional to concentration, path length and a per-molecule constant ε280. The whole difficulty is knowing ε280. This page is part of the protein calculator set and removes that hurdle by deriving ε280 directly from the amino-acid sequence — or letting you enter a measured value — then carrying the result all the way to mg/mL and µM with path length and dilution handled explicitly.

Proteins absorb at 280 nm almost entirely because of their aromatic residues. Tryptophan is by far the strongest absorber (ε ≈ 5500 M⁻¹cm⁻¹), tyrosine is next (ε ≈ 1490), and a disulfide-bonded cystine pair contributes a little (ε ≈ 125). The Gill & von Hippel equation adds these per-residue contributions to predict ε280 of a folded protein in water to within about ±5%: ε280 = n(Trp)·5500 + n(Tyr)·1490 + n(Cystine)·125. A protein with no tryptophan and few tyrosines absorbs weakly and is hard to quantify this way — which is exactly why the calculator warns you when ε280 comes out at zero.

Once ε280 is known the concentration follows from c(M) = A280 / (ε280 × path), corrected by the dilution factor if you diluted the sample before reading. To get a mass concentration, multiply by the molecular weight: because 1 g/L equals 1 mg/mL, mg/mL = c(M) × MW. The same relationship is often quoted as A(1%,1cm) = 10 · ε280 / MW — the absorbance of a 1% (10 mg/mL) solution in a 1 cm path — and the calculator reports that too, since many protocols and datasheets are written in those units. The molecular weight comes from the sequence automatically, or you can enter it; the molecular weight calculator gives it on its own.

The method has real pitfalls that a bare formula hides, and this page names them. Nucleic acids absorb strongly at 260 nm and their tail inflates A280, so a contaminated prep reads high — check the A260/A280 ratio (pure protein is ~0.57). Reductants like DTT and β-mercaptoethanol absorb near 280 nm and must be in the blank. Denaturants shift ε280, scattering particles raise the baseline, and readings above ~1.5 leave the linear range of most spectrophotometers. Handle those and A280 is an excellent routine measurement; ignore them and it is a source of silent 2-fold errors. After quantifying, pair the result with the molarity calculator or the dilution calculator to prepare working stocks.

Optional. When given, ε280 is computed from the Trp/Tyr/cystine content and MW from the sequence. Leave blank to enter a known ε280 instead.

Molar extinction coefficient at 280 nm from a datasheet or measurement. Used only when the sequence box is empty.

Needed to report mg/mL and A(1%,1cm) when you supply ε280 directly.

Absorbance at 280 nm minus the buffer blank.

1 cm standard cuvette; micro-volume readers often use a shorter effective path.

Factor by which you diluted before reading (10 for a 1:10 dilution).

A disulfide cystine pair adds 125 M⁻¹cm⁻¹; free reduced cysteines add nothing at 280 nm.

Concentration5.74822 mg/mL
Molarity671.141 µM
ε2801490 M⁻¹cm⁻¹
Molecular weight8564.84 Da
A(1%,1cm)1.73967

How to use the protein concentration calculator

If you have the sequence, paste it into the sequence box and ε280 and MW are both derived for you. Choose the cysteine state: oxidized counts disulfide-bonded cystine pairs (each +125), reduced counts none.

If you instead have a measured or datasheet ε280, leave the sequence blank and enter the ε280 in M⁻¹cm⁻¹, plus the molecular weight in Da if you want mg/mL and A(1%,1cm) reported.

Enter the A280 of your sample, blank-corrected against the exact buffer it is in. Set the path length (1 cm for a standard cuvette; micro-volume instruments often use a shorter effective path).

If you diluted the sample before reading, enter the dilution factor so the result refers to the original stock (e.g. 10 for a 1:10 dilution).

Press Calculate. The primary card is the concentration (mg/mL when a molecular weight is available, otherwise µM); the cards below give molarity in M and µM, ε280 used, A(1%,1cm) and the inputs echoed back. Carry the result into the dilution calculator to make a working dilution.

Protein Concentration from A280 formula

ε280 = n(Trp)·5500 + n(Tyr)·1490 + n(Cystine)·125 [M⁻¹cm⁻¹]
n(Cystine) = ⌊n(Cys)/2⌋ (oxidized) or 0 (reduced)
c (M) = (A280 × dilution factor) / (ε280 × path length)
concentration (mg/mL) = c (M) × MW (Da)
A(1%,1cm) = 10 · ε280 / MW; concentration (µM) = c (M) × 10⁶
Per-residue coefficients (Trp 5500, Tyr 1490, cystine 125 M⁻¹cm⁻¹) are for a folded protein in water at neutral pH, per Gill & von Hippel (1989) and Pace et al. (1995). The mg/mL step uses the identity 1 g/L = 1 mg/mL, so mg/mL = c(M) × MW.

Variables and units

SymbolMeaningUnit
A280Blank-corrected absorbance at 280 nm—
ε280Molar extinction coefficient at 280 nmM⁻¹cm⁻¹
pathCuvette path lengthcm
DFDilution factor of the measured sample×
cMolar concentrationM (mol/L)
MWMolecular weight of the proteinDa

How protein concentration is calculated

  1. Obtain ε280: either derive it from the sequence with the Gill & von Hippel sum, or enter a measured value.
  2. Obtain MW: from the sequence automatically, or entered, to report a mass concentration.
  3. Apply Beer–Lambert: c(M) = (A280 × dilution factor) / (ε280 × path length).
  4. Convert to mass concentration: mg/mL = c(M) × MW (using 1 g/L = 1 mg/mL).
  5. Report molarity in µM (×10⁶) and A(1%,1cm) = 10 · ε280 / MW.
  6. Flag the common interferences — nucleic acid, reductants, turbidity, out-of-range absorbance.

Worked laboratory examples

Example 1 — The BSA benchmark

Given: ε280 = 43,824 M⁻¹cm⁻¹ · MW = 66,430 Da · A280 = 0.66, path 1 cm, no dilution

  1. c = 0.66 / (43,824 × 1) = 1.506 × 10⁻⁵ M.
  2. mg/mL = 1.506 × 10⁻⁵ × 66,430 = 1.00 mg/mL.
  3. A(1%,1cm) = 10 × 43,824 / 66,430 = 6.60; equivalently a 1 mg/mL (0.1%) solution reads 0.66 — the classic BSA check.

Result: 1.00 mg/mL (15.1 µM)

Example 2 — ε derived from a sequence (ubiquitin)

Given: ubiquitin, 76 residues · composition: 0 Trp, 1 Tyr, 0 Cys · A280 = 1.0, path 1 cm

  1. ε280 = 0×5500 + 1×1490 + 0×125 = 1490 M⁻¹cm⁻¹.
  2. MW from the same sequence = 8564.84 Da.
  3. c = 1.0 / 1490 = 6.71 × 10⁻⁴ M = 671 µM; mg/mL = 6.71 × 10⁻⁴ × 8564.84 = 5.75.
  4. A(1%,1cm) = 10 × 1490 / 8564.84 = 1.74; a 1 mg/mL solution reads 0.174 — low, because ubiquitin has no tryptophan.

Result: ε280 = 1490; A280 1.0 → 5.75 mg/mL (671 µM)

Example 3 — A diluted Trp-rich sample

Given: ε280 = 48,275 M⁻¹cm⁻¹ (a 6-Trp, 10-Tyr, 3-cystine protein) · MW = 50,000 Da · A280 = 0.75 of a 1:5 dilution, path 1 cm

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

Result: 3.88 mg/mL (77.7 µM) in the undiluted stock

Example 4 — Cysteine pairing changes ε

Given: a peptide CCCC (four cysteines) · no Trp or Tyr

  1. Oxidized: 4 Cys form 2 cystines → ε280 = 2 × 125 = 250 M⁻¹cm⁻¹.
  2. Reduced: free cysteines do not absorb at 280 nm → ε280 = 0, and A280 cannot quantify it.
  3. The cysteine-state selector therefore changes whether this sample is measurable at all.

Result: ε280 = 250 (oxidized) vs 0 (reduced)

Want the reasoning behind the numbers? Read How to Determine Protein Concentration from A280.

How to interpret the result

The primary number is your protein concentration in the units that matter for the next step. When a molecular weight is available it is mg/mL, what you use to load a gel or set up a reaction; the µM value beside it is what a molar binding or kinetics experiment needs.

ε280 is the lever. A sequence-derived ε is accurate to about ±5% for a folded globular protein; a measured ε from a validated standard is better still. If the two disagree by more than that, suspect a modification, a mis-folded or aggregated sample, or contamination.

A(1%,1cm) is the same information in percent-solution units. If a protocol says "an A280 of 1.0 corresponds to X mg/mL", that X is MW / ε280 = 10 / A(1%,1cm), and the calculator gives it so you can match the protocol directly.

When the reading looks wrong, the A260/A280 ratio is the first diagnostic. Pure protein is ~0.57; a ratio near 1.0 or higher signals nucleic-acid contamination that is inflating your A280 and your calculated concentration.

Assumptions

  • The protein is folded and in aqueous buffer at roughly neutral pH; denaturants and extreme pH change ε280.
  • The absorbance is blank-corrected against the exact sample buffer, including any reductant.
  • Beer–Lambert holds — the reading is within the instrument's linear range (ideally A280 < 1.5).
  • In sequence mode, ε280 follows the Gill & von Hippel per-residue coefficients with cysteines paired as chosen.
  • The sample is a single pure protein; contaminants with their own absorbance bias the result.

Limitations

  • A protein lacking Trp, Tyr and paired Cys has ε280 ≈ 0 and cannot be quantified at 280 nm — use Bradford, BCA or a 205 nm peptide-bond reading.
  • Nucleic-acid contamination absorbs at 260 nm and inflates A280; a correction or a clean-up is needed.
  • DTT, β-mercaptoethanol, imidazole and some detergents absorb near 280 nm and must be present in the blank.
  • Sequence-derived ε assumes a native fold; a denatured or aggregated protein absorbs differently.

Common mistakes

  • Forgetting the dilution factor and reporting the diluted, not the stock, concentration.
  • Using the wrong path length for a micro-volume spectrophotometer and being off by its ratio.
  • Blanking against water instead of the actual sample buffer, so reductant absorbance is counted as protein.
  • Ignoring nucleic-acid contamination and reading a high A280 as pure protein.
  • Assuming every protein absorbs at 280 nm — a Trp/Tyr-poor protein gives a near-zero ε and a meaningless result.

FAQ

How do I find ε280 if I do not have the sequence?

Use a published or datasheet value, or measure it against a known concentration. If you have the sequence, this tool derives ε280 from the Trp, Tyr and cystine content using the Gill & von Hippel equation, accurate to about ±5% for a folded protein.

Why is 280 nm used for proteins?

The aromatic side chains tryptophan and tyrosine (and disulfide cystine) absorb UV with a peak near 280 nm, while the peptide backbone absorbs lower, near 205 nm. 280 nm is specific enough to read protein in many buffers without interference from the backbone.

What is A(1%,1cm) and how does it relate to ε280?

A(1%,1cm) is the absorbance of a 1% (10 mg/mL) solution in a 1 cm path. It relates to the molar coefficient by A(1%,1cm) = 10 · ε280 / MW. For BSA, 10 × 43,824 / 66,430 = 6.60; a 1 mg/mL (0.1%) solution is a tenth of that and reads A280 ≈ 0.66.

How do I correct for nucleic-acid contamination?

Measure A260 and A280 and check the ratio (pure protein ≈ 0.57; pure DNA ≈ 1.8). Use a two-wavelength correction such as the Warburg–Christian equation, or better, clean up the sample. A high A260/A280 means your A280-derived concentration is overestimated.

Does the cysteine state really matter?

Yes, when a protein is cysteine-rich. Each disulfide-bonded cystine pair adds 125 M⁻¹cm⁻¹; reduced free cysteines add nothing at 280 nm. The selector lets you match the redox state of your sample so the ε is right.

My A280 is above 1.5 — is that a problem?

Most spectrophotometers are only linear up to about 1.5–2.0 AU. Dilute the sample into the linear range, measure, and multiply by the dilution factor. Reading outside the linear range under-reports the true absorbance and your concentration.

Scientific Sources

External links open in a new tab and carry rel="nofollow". We cite primary references (NIST, NIH/NCBI, peer-reviewed papers) and established manufacturer technical documentation (NEB, Thermo Fisher, Promega, IDT). Calculated results are estimates; your protocol and manufacturer guidance take precedence.

Last reviewed: 2026-09 · Method version: 1.0