A280 → concentration (Beer–Lambert)
C (mg/mL) = A280 ÷ (ε × l) × MW × dilution factor. Pick a protein or enter ε + MW by hand.
Filling either box overrides the database choice.
Estimate ε from sequence (Pace et al. 1995)
ε = 5500·#Trp + 1490·#Tyr + 125·#Cys
Your reading
Subtracted from A280 to correct turbidity.
Bradford / BCA standard curve
Enter your standard concentrations and their absorbances — the tool fits a least-squares line, reports R², and reads your unknown off the curve.
Standards
| # | Concentration | Absorbance | Remove |
|---|
Unknown sample
Protein unit converter
Convert between mass and molar concentration. Needs the molecular weight — pick from the database or type it in.
Quick MW presets:
Dilution planner (C1V1 = C2V2)
How much stock to pipette, and how much buffer to add, to reach a working concentration.
Worked examples (hand-checked)
1 · BSA from A280
A280 = 1.00 (blank-corrected), 1 cm cuvette, no dilution. BSA: ε = 43,824 M⁻¹cm⁻¹, MW = 66,463 Da.
2 · Bradford standard curve
Standards (mg/mL → A595): 0→0.05, 0.125→0.18, 0.25→0.32, 0.5→0.58, 1.0→1.12. Least squares: slope = 1.0700, intercept = 0.0488, R² = 0.9999 (≥ 0.98 ✓). Unknown A595 = 0.45:
3 · Dilution: 5 mg/mL stock → 0.5 mg/mL working, 1 mL final
Methodology — the formulas behind the numbers
Beer–Lambert law (A280 mode)
Proteins absorb at 280 nm mainly through tryptophan and tyrosine (plus disulfides). Absorbance follows the Beer–Lambert law:
where ε is the molar extinction coefficient (M⁻¹cm⁻¹), l the pathlength in cm, MW the molecular weight in Da, and DF the dilution factor. Molar concentration is c(M) = A ÷ (ε·l); multiply by 10⁶ for µM. The "mass extinction coefficient" ε ÷ MW is the A280 of a 1 mg/mL solution — 0.66 for BSA, 2.65 for lysozyme, 1.40 for IgG.
Extinction coefficient from sequence
When no literature ε exists, the Pace et al. (1995) estimate used by Expasy ProtParam is:
Cysteine's contribution assumes the reduced form; a fully disulfide-bonded protein differs slightly. Treat sequence-based ε as ±5–10% — fine for planning, not for publishing.
Standard curve (Bradford / BCA / Lowry)
Colorimetric assays are relative: dye response depends on amino-acid composition, so you calibrate against a BSA (or IgG) standard series. The tool fits y = m·x + b by ordinary least squares and reports R² = 1 − SSres/SStot. The unknown's concentration is x = (A − b) ÷ m, times its dilution factor. This is an educational estimate — not professional advice; always follow your lab's validated SOP for regulated work.
Honest limitations — when the number lies
- Nucleic-acid contamination. DNA/RNA absorb ~20× more per mass at 260 nm than protein does at 280 nm, and they still absorb at 280. Even 1–2% nucleic acid inflates an A280 reading noticeably. Check A260/A280: pure protein ≈ 0.57; above ~0.8, suspect contamination and re-purify (or treat with nuclease).
- Light scattering. Aggregates and turbid samples scatter light, which the detector reads as absorbance. The optional A320 correction subtracts this background — but a high A320 means you should centrifuge/filter and re-read, not just correct.
- Outside the linear range. Above ~A280 2.0 most detectors saturate; below ~0.05 you're in the noise. Dilute or concentrate into the 0.1–1.0 window.
- Wrong ε, wrong answer. ε varies with buffer pH, denaturants, and disulfide state. The 10-protein database lists literature values — your supplier's certificate of analysis wins.
- Bradford is protein-dependent. Coomassie binds arginine/lysine-rich proteins more strongly; a BSA curve misreads a lysine-poor protein. Match your standard to your sample when accuracy matters.
- NanoDrop pathlength. NanoDrop software already normalizes to the 10 mm equivalent — pick "1 cm cuvette" for its displayed value, and only use the 1 mm / 0.2 mm options for a raw short-path reading.
🧬 Protein or nucleic acid? Pick the right tool.
This calculator is for proteins (A280). If you measured absorbance at 260 nm, you have DNA or RNA — use our DNA concentration calculator instead (A260 → ng/µL, A260/A280 purity, oligo mode). Mixing the two up is the most common concentration mistake on the bench.
Frequently asked questions
How do you calculate protein concentration from A280?
The Beer-Lambert formula is C (mg/mL) = A280 / (extinction coefficient x pathlength) x molecular weight x dilution factor. Example with BSA: A280 = 1.00, extinction coefficient 43,824 M-1cm-1, MW 66,463 Da, 1 cm cuvette -> 1.00 / 43824 x 66463 = 1.5166 mg/mL. The calculator above does this with a 10-protein extinction-coefficient database so you don't have to look values up.
Where do I find the extinction coefficient for my protein?
Three routes, in order of trust: (1) your supplier's certificate of analysis, (2) the curated 10-protein database built into this calculator (BSA, IgG, lysozyme, ovalbumin, RNase A, trypsin, beta-lactoglobulin, ovotransferrin, GFP, carbonic anhydrase II - literature values from Expasy ProtParam), or (3) estimate it from the amino-acid sequence with the Pace et al. 1995 formula: epsilon = 5500 x #Trp + 1490 x #Tyr + 125 x #Cys. The sequence estimator is built into the A280 tab.
How do I convert mg/mL to micromolar for a protein?
Divide by the molecular weight: uM = (mg/mL x 1,000,000) / MW in Daltons, equivalently uM = 1000 x (mg/mL) / MW in kDa. Example: 1 mg/mL BSA (MW 66,463 Da) = 1,000,000 / 66463 = 15.046 uM. The unit converter tab does mg/mL, ug/uL, uM, nM and % w/v in both directions - 1% w/v equals 10 mg/mL.
How does the Bradford assay calculator work?
Colorimetric assays like Bradford and BCA are relative methods: you measure your standards (typically BSA at 0, 0.125, 0.25, 0.5 and 1.0 mg/mL) and the tool fits a least-squares line y = m*x + b, reporting R-squared. Your unknown's concentration is then (absorbance - b) / m, times its dilution factor. The chart shows your points against the fitted line so a bad point is visually obvious.
Bradford vs BCA: which protein assay should I use?
Bradford (read at 595 nm) is fast - 5 minutes, one reagent - but it is thrown off by detergents like SDS and Triton, and Coomassie dye binds arginine-rich proteins more strongly, so the standard should resemble your sample. BCA (read at 562 nm) tolerates up to 5% SDS and most detergents, takes 30+ minutes at 37 C, and is disturbed by reducing agents like DTT and beta-mercaptoethanol. This calculator's curve-fitting works for either; pick the assay your sample chemistry allows.
What is a good R-squared for a protein standard curve?
Aim for R2 >= 0.98. Below that, suspect a pipetting error, a saturated top standard, or dye reagent past its prime - the calculator flags it explicitly. Also check that your unknown's absorbance falls between your lowest and highest standards; reading a concentration off an extrapolated curve is unreliable, and the tool warns you about that too.
What does the A260/A280 ratio mean for a protein sample?
Pure protein reads about 0.57, because aromatic residues absorb at 280 nm. Nucleic acids absorb roughly 20x more per mass at 260 nm, so even slight DNA/RNA contamination pushes the ratio up: above ~0.8, suspect contamination and consider nuclease treatment or re-purification, because the A280 concentration will be overestimated. Enter your A260 in the calculator for an automatic verdict.
How do I correct protein concentration for turbidity?
Aggregates scatter light, which the detector counts as absorbance. Take a reading at 320 nm - where neither protein nor nucleic acid absorbs - and subtract it from A280. The calculator has an optional A320 field that does this. But treat a large A320 as a symptom: centrifuge or filter the sample and re-read rather than trusting a heavily corrected number.
Which pathlength do I pick for NanoDrop readings?
A NanoDrop measures through a 1 mm (or 0.2 mm) liquid column, but its software already scales the displayed value to the 10 mm equivalent - so choose '1 cm cuvette' for the number on screen. Only use the 1 mm / 0.2 mm options for a raw, un-normalized short-path reading; picking 1 mm for an already-normalized value inflates your result 10-fold.
Why is my A280 reading negative or zero?
Almost always a blanking problem: the instrument was blanked with a different buffer than the sample is in, or the blank drifted. Re-blank with the exact sample buffer and re-read. This calculator refuses to compute from a zero or negative A280 instead of silently producing a nonsense number.
What is the linear range for A280 protein measurements?
Beer-Lambert only holds while the detector responds linearly - roughly A280 0.1 to 1.0 for the diluted sample on most instruments. Above ~2.0 the reading saturates and the computed concentration becomes unreliable; below ~0.05 you are in the noise. The calculator flags readings above 2.0; the correct move is to dilute and re-read.
Can I use this calculator for the Lowry assay?
Yes for the math: Lowry is also a colorimetric assay with a standard curve, so enter your standards and read at 750 nm using the assay selector. Just remember Lowry's chemistry is sensitive to many common buffer components (Tris, EDTA, ammonium sulfate), so keep your standards in the same buffer as your samples.
How do I dilute a protein stock to a working concentration?
Use C1V1 = C2V2: stock volume = (target concentration x final volume) / stock concentration. Example: 5 mg/mL stock to 0.5 mg/mL in 1 mL final -> 0.5 x 1.0 / 5 = 0.10 mL stock plus 0.90 mL buffer (100 uL + 900 uL). The dilution planner tab computes both volumes and the dilution factor, and refuses impossible setups like a target above the stock.
What is the difference between protein and DNA concentration measurement?
Different molecules, different wavelengths, different math. Proteins are quantified at 280 nm via aromatic residues with a protein-specific extinction coefficient (this tool). Nucleic acids are quantified at 260 nm with fixed conversion factors - 50 ng/uL per A260 for dsDNA. For DNA/RNA, use our DNA concentration calculator instead; mixing the two up is the most common bench concentration mistake.
How does this compare to the OmniCalculator protein concentration calculator?
OmniCalculator's page covers the basic Beer-Lambert formula in a single thin calculator. This one goes further: a 10-protein extinction-coefficient database with sources, an estimate-epsilon-from-sequence tool, Bradford/BCA/Lowry standard-curve fitting with R2 quality checks and an SVG chart, a five-unit converter, a C1V1 dilution planner, A260/A280 purity interpretation, A320 scatter correction, three hand-checked worked examples, and honest limitations - all computed 100% in your browser.
Can I rely on this for medical, diagnostic or regulated lab work?
No. This is an educational estimate tool, not professional advice: it computes from the numbers you type and cannot judge sample quality, instrument calibration, or protocol validity. For clinical, diagnostic, or GMP/GLP work, use your lab's validated SOPs and calibrated instruments, and confirm critical results with an orthogonal method.