Audited —·Last updated 29 Jul 2026·5 citations·Tier 2·0 uses

Protein Concentration Calculator — A280, E1% and Standard Curve

Turn an A280 reading or a Bradford/BCA standard curve into mg/mL and µM. Predicts ε from Trp, Tyr and cystine using Pace 1995 or Gill & von Hippel 1989.

Protein Concentration Calculator

Method
The blanked reading: A280 for the first four methods, or the assay wavelength (595 nm for Bradford, 562 nm for BCA) for the standard curve. Bench spectrophotometers are linear over roughly 0.1–1.5 AU.
AU
1 cm for a standard cuvette, 0.1 cm for a 1 mm cell. Microvolume instruments usually report an absorbance already normalised to 1 cm — if yours does, leave this at 1. Not used by the standard-curve method, which already embeds the path length.
cm
Get this from ExPASy ProtParam for your sequence, or measure it by the Edelhoch method as Pace et al. recommend. The default of 42925 is what Pace's coefficients give for 2 Trp, 20 Tyr and 17 cystines (mature bovine serum albumin).
M⁻¹cm⁻¹
Number of Trp residues in the mature chain. Trp dominates ε at 280 nm, which is why Pace et al. call the prediction less reliable for proteins with none.
Number of Tyr residues in the mature chain.
Number of disulfide BONDS, not free cysteines — a protein with 35 cysteines forming 17 bridges and one free thiol has 17 cystines. Enter 0 for a fully reduced protein.
Sometimes written A(1%, 280) or E(0.1%) × 10. If your vendor quotes 'A280 of 1 mg/mL = 0.66', that is an E1% of 6.6. The generic '1 AU = 1 mg/mL' rule of thumb corresponds to E1% = 10 and is only a rough average.
Needed for the µM output in every method. The default of 66433 is mature bovine serum albumin (583 residues). Take yours from ExPASy ProtParam or a mass spectrum.
g/mol
From the linear fit of your Bradford or BCA standards. Must be positive — more protein must give more signal. If your standards were in µg/mL, divide the slope by 1000 to convert it to AU per mg/mL.
AU per mg/mL
The y-intercept of the same fit — typically the blank's residual absorbance. A reading below this implies a negative concentration, which the calculator refuses rather than reports.
AU
How many fold you diluted before reading. 1 means neat; 10 means a 1:10 dilution (one part sample, nine parts buffer). The result is scaled back up to the original sample.
×
Volume of the ORIGINAL, undiluted sample you want the total protein mass for.
µL
Concentration
1.3155
Concentration of the original, undiluted sample. Equal to the cuvette figure multiplied by the dilution factor.
Molar concentration
19.802 µM
In the cuvette
1.3155 mg/mL
Total protein
0.6578 mg
Method and caveats
A280 with a molar absorption coefficient you supplied (42925 M-1 cm-1). Beer-Lambert assumes no nucleic-acid contamination and no light scattering; check the A260/A280 ratio and the A320 baseline before trusting it. The reading of 0.85 AU is inside the roughly 0.1-1.5 AU window over which bench spectrophotometers are linear (instrument convention, not a published standard). This is a measured, method-dependent value, not a clinical or population figure.

Background.

This protein concentration calculator turns an absorbance reading into mg/mL, µM and total milligrams by five different routes, and it tells you on the page which paper's constants each route uses. That last part matters more than it sounds. The two standard methods for predicting a protein's molar absorption coefficient from its sequence disagree by about nine percent, and a concentration is only as good as the coefficient behind it.

The four A280 routes all rest on the Beer–Lambert law, A = ε·c·l. Give the calculator an absorbance, a path length and an ε, and it returns the molar concentration; multiply by the molecular weight and you have mg/mL. Where the routes differ is how you get ε. You can supply it directly, from ExPASy ProtParam or from an Edelhoch measurement. You can have it predicted from your tryptophan, tyrosine and cystine counts using Pace and colleagues' 1995 coefficients — 5500 per Trp, 1490 per Tyr, 125 per cystine — which are calibrated for a native protein in water. Or you can use Gill and von Hippel's 1989 coefficients — 5690, 1280 and 120 — which were measured in 6 M guanidine hydrochloride on denatured protein. Both papers are legitimate, widely cited and still in use; applied to the same 2-Trp, 20-Tyr, 17-cystine protein they give 42,925 and 39,020 M⁻¹cm⁻¹ respectively, a nine percent gap that lands directly in your answer. This page ships both as separate labelled modes rather than picking one quietly.

The fifth route handles colourimetric assays — Bradford, BCA and Lowry — where you read a dye or a copper complex against a standard curve. Feed in the slope and intercept of your fit and the calculator converts a reading into concentration. Read the caveat that comes with it: a colourimetric number is assay-dependent in a way an A280 number is not. Coomassie dye binding in the Bradford assay responds mainly to basic and aromatic residues, while the BCA assay depends on Cu²⁺ reduction by a different residue set, so the two assays legitimately disagree on the same tube. On top of that, the number depends on which protein calibrated the curve: the same sample quantified against BSA standards and against IgG standards will not give the same answer. Always report the assay and the standard with the value.

A few things this calculator deliberately does not do. It does not correct A280 for nucleic-acid contamination or for light scattering, both of which inflate the reading — check your A260/A280 ratio and your A320 baseline instead. It does not fit your standard curve; paste the slope and intercept from your own regression. And it treats the standard curve as linear, which for Bradford is only a local approximation because the dye response is genuinely non-linear over a wide range.

Every number this page returns is a measured, method-dependent value, not a population statistic and not a clinical result. Two honest labs measuring the same tube by two honest methods will get two different numbers, and the useful discipline is to report the method, the constants and the instrument alongside the figure rather than to pretend a single true value exists.

What is protein concentration calculator?

Protein concentration is normally measured in one of two families of ways. The direct route reads the sample's own ultraviolet absorbance at 280 nm, where tryptophan and tyrosine side chains — and, more weakly, disulfide-bonded cystines — absorb. Because absorbance is proportional to concentration through the Beer–Lambert law A = ε·c·l, dividing the reading by the molar absorption coefficient ε and the path length l gives the molar concentration directly. The method is fast, uses no reagents, and does not consume the sample, but it needs an ε for your specific protein and it is confounded by anything else in the tube that absorbs at 280 nm, most notably nucleic acid.

The indirect route uses a colourimetric assay, in which a reagent changes colour in proportion to protein present. Bradford's 1976 Coomassie dye-binding assay and the bicinchoninic acid assay of Smith and colleagues from 1985 are the two most common. These do not need an ε or a molecular weight, work at lower concentrations, and are less sensitive to nucleic acid — but they need a standard curve, they consume the sample, and the answer depends on both the chemistry and the standard protein used to calibrate it.

The molar absorption coefficient itself can be predicted from sequence. Pace, Vajdos, Fee, Grimsley and Gray established the modern coefficients in Protein Science in 1995, from 116 measured values across 80 proteins, and their own conclusion is worth quoting: the prediction is 'quite reliable for proteins containing Trp residues, and less reliable for proteins that do not', and 'the best approach is to measure rather than predict epsilon'. Gill and von Hippel's earlier 1989 paper in Analytical Biochemistry gives a different coefficient set measured under denaturing conditions and claims about five percent accuracy. Both are implemented here, separately labelled, because a reader following one paper and a reader following the other will legitimately get different answers.

How to use this calculator.

  1. Pick the method that matches your data. If you already know ε for your protein, use the first mode. If you know the sequence but not ε, use one of the two composition modes. If your vendor quotes a 1% extinction value, use the fourth. If you ran a Bradford or BCA assay, use the standard curve.
  2. Enter the blanked absorbance. For A280 that is the reading against a buffer blank; for a colourimetric assay it is the reading against the assay blank at 595 nm (Bradford) or 562 nm (BCA).
  3. Set the path length: 1 cm for a standard cuvette, 0.1 cm for a 1 mm cell. If your microvolume instrument already normalises the reading to a 1 cm path, leave it at 1. The standard-curve mode ignores this field, because the curve already embeds the path length.
  4. For the composition modes, count the residues in the mature chain. Cystines are disulfide BONDS, not free cysteines — a protein with 35 cysteines forming 17 bridges plus one free thiol has 17 cystines. Enter 0 for a fully reduced protein.
  5. For the standard-curve mode, paste the slope and intercept from your own linear fit. If your standards were in µg/mL, divide the slope by 1000 first so it is in AU per mg/mL.
  6. Enter the dilution factor you applied before reading — 10 for a 1:10 dilution. The output is scaled back to the original sample.
  7. Enter a molecular weight if you want the µM output. It is not needed for mg/mL in the E1% or standard-curve modes, but the calculator uses it for the molar figure in all five.
  8. Read the interpretation line. It names the method and the source of its constants, and flags a reading that sits outside the roughly 0.1–1.5 AU range over which bench spectrophotometers are linear.
  9. Before you trust an A280 number, check the A260/A280 ratio for nucleic-acid carryover and the A320 baseline for light scattering. This page does not correct for either.

The formula.

c = A ⁄ (ε × l)
A · absorbance at 280 nm (dimensionless)ε · molar absorption coefficient (M⁻¹cm⁻¹)l · path length (cm)c · molar concentration (M); mg/mL = c × MWPace 1995 · ε = 5500·nTrp + 1490·nTyr + 125·nCystine (native, water)Gill & von Hippel 1989 · ε = 5690·nTrp + 1280·nTyr + 120·nCystine (6 M GuHCl)E1% route · c(mg/mL) = 10A ⁄ (E1% × l)standard curve · c(mg/mL) = (A − intercept) ⁄ slope

The Beer–Lambert law states that absorbance is the product of the molar absorption coefficient, the concentration and the path length: A = ε·c·l. Rearranged, c = A ÷ (ε·l), which gives a molar concentration when ε is in M⁻¹cm⁻¹ and l is in centimetres. Multiplying that molarity by the molecular weight in g/mol gives grams per litre, which is numerically the same as milligrams per millilitre.

Two directional consequences are worth stating explicitly because one of them is counter-intuitive. Doubling the path length halves the concentration you infer from the same reading, which is why a 1 mm cell and a 1 cm cuvette are not interchangeable. And a LARGER ε means the same absorbance corresponds to LESS protein, because a strongly absorbing protein produces more signal per molecule. That is why swapping Pace's coefficients for Gill and von Hippel's — which give the smaller ε of 39,020 rather than 42,925 for the default composition — raises the reported concentration from 1.3155 to 1.4472 mg/mL rather than lowering it.

The composition modes are simple additive sums. Pace and colleagues give ε(280) = (#Trp)(5,500) + (#Tyr)(1,490) + (#cystine)(125). For a protein with 2 tryptophans, 20 tyrosines and 17 disulfide bridges that is 11,000 + 29,800 + 2,125 = 42,925 M⁻¹cm⁻¹, which is the default this page uses in the supply-your-own-ε mode so you can check the arithmetic yourself. Gill and von Hippel's set gives 2×5,690 + 20×1,280 + 17×120 = 11,380 + 25,600 + 2,040 = 39,020 M⁻¹cm⁻¹ for the same protein.

The E1% route avoids ε entirely. E1% is the absorbance of a 1% weight-per-volume solution, which is 10 mg/mL, over a 1 cm path. So c in mg/mL is 10A ÷ (E1% × l). With A = 0.85 and E1% = 6.6, that is 8.5 ÷ 6.6 = 1.2879 mg/mL. The familiar rule of thumb that 1 absorbance unit equals 1 mg/mL is the special case E1% = 10, and it is only a rough average across proteins.

The standard-curve route inverts a linear calibration: c = (A − intercept) ÷ slope. With A = 0.85 on a curve of slope 0.75 AU per mg/mL and intercept 0.08 AU, that is 0.77 ÷ 0.75 = 1.0267 mg/mL. If your reading falls below the intercept, the algebra returns a negative concentration; the calculator refuses that rather than printing it, because a negative protein concentration is not a measurement, it is a sign that the blank or the detection limit needs attention.

Rounding: every step runs in arbitrary-precision decimal arithmetic, and rounding happens exactly once, at the final result, to ten decimal places. The predicted ε is not rounded before it enters the division. Report your own results to two or three significant figures — a spectrophotometer's absorbance is good to about ±0.005 AU, and ε itself carries several percent of uncertainty even before you choose between the two coefficient sets.

A worked example.

Example

You have purified a 66,433 Da protein with 2 tryptophans, 20 tyrosines and 17 disulfide bridges. The neat stock is too concentrated to read directly, so you dilute it 1:10 into buffer and get an A280 of 0.430 in a 1 cm cuvette. Choose the Pace 1995 composition mode, enter the three residue counts, the absorbance, a dilution factor of 10 and a sample volume of 1000 µL. The calculator first predicts ε = 2×5500 + 20×1490 + 17×125 = 42,925 M⁻¹cm⁻¹, then applies Beer–Lambert: 0.430 ÷ 42,925 = 1.00175 × 10⁻⁵ M in the cuvette, which is 0.6655 mg/mL there. Multiply by the ten-fold dilution and the original stock is 6.6549 mg/mL, or 100.17 µM, and the 1000 µL you have left contains 6.6549 mg of protein. The reading of 0.430 AU sits comfortably inside the roughly 0.1–1.5 AU range over which bench spectrophotometers are linear, so the dilution was a sensible one. Now switch the mode to Gill & von Hippel 1989 without changing anything else: ε drops to 39,020 M⁻¹cm⁻¹ and the answer rises to 7.32 mg/mL — about ten percent higher from the identical cuvette. Neither number is wrong; they answer the question under different published conventions, and whichever you report you should name the paper beside it.

absorbance0.43
cystine Count17
sample Volume Ul1,000
dilution Factor10
molecular Weight66,433
tryptophan Count2
tyrosine Count20
path Length Cm1
solve Fora280CompositionPace

Frequently asked questions.

Which extinction coefficient set should I use — Pace or Gill & von Hippel?
Match the coefficient set to the state your protein is in when you read it. Gill and von Hippel measured their coefficients in 6 M guanidine hydrochloride, on denatured protein, and claim about five percent accuracy; their values are 5690 per tryptophan, 1280 per tyrosine and 120 per cystine. Pace and colleagues re-derived the coefficients for native protein in water from 116 measured values across 80 proteins, giving 5500, 1490 and 125. If you are reading a folded protein in an aqueous buffer, which is the normal case, use Pace. If you are reading in 6 M GuHCl — which is exactly what the Edelhoch method does — Gill and von Hippel is the matched set. The gap is not trivial: for a protein with 2 Trp, 20 Tyr and 17 cystines the two give 42,925 and 39,020 M⁻¹cm⁻¹, and that nine percent propagates straight into your concentration. Whichever you use, name it in your methods section.
Why does a smaller extinction coefficient give a higher concentration?
Because ε is in the denominator: c = A ÷ (ε·l). A large ε means each molecule absorbs strongly, so a given absorbance is produced by relatively few molecules; a small ε means each molecule absorbs weakly, so the same absorbance requires more of them. It catches people out because 'bigger coefficient' feels like it should mean 'more protein'. Concretely, with A280 = 0.85 in a 1 cm cuvette, Pace's ε of 42,925 gives 1.3155 mg/mL while Gill and von Hippel's smaller ε of 39,020 gives 1.4472 mg/mL. The same logic applies to path length: a longer path collects more absorbance from the same solution, so a reading taken in a 1 cm cuvette implies ten times less protein than the identical reading taken in a 1 mm cell.
Why do Bradford and BCA give different answers for the same sample?
Because they are chemically different measurements, not two attempts at the same one. The Bradford assay works by Coomassie Brilliant Blue binding to protein, and dye binding depends strongly on basic and aromatic residues — arginine especially. The BCA assay works by protein reducing Cu²⁺ to Cu⁺, which then chelates bicinchoninic acid, and that reduction depends on a different residue set including cysteine, tryptophan and tyrosine as well as the peptide backbone. A protein rich in arginine reads high on Bradford relative to BCA; a protein rich in cysteine does the opposite. On top of the chemistry, the answer depends on the standard: a curve calibrated against bovine serum albumin and one calibrated against immunoglobulin G give different numbers for the same unknown, because neither standard has your protein's composition. This is why a protein concentration should always be reported with the assay and the standard named, and why comparing a Bradford value from one paper against a BCA value from another is not a like-for-like comparison.
Is A280 = 1.0 really 1 mg/mL of protein?
Only as a rough average, and it can be wrong by a factor of two or more for an individual protein. The rule of thumb corresponds to an E1% of 10, meaning a 1% weight-per-volume solution would read 10 absorbance units over 1 cm. Real proteins vary enormously because ε depends on how many tryptophans and tyrosines they carry: a tryptophan-rich protein absorbs strongly and its E1% may be 20 or more, while a protein with no tryptophan at all may sit near 3 or lower. The default worked example on this page has an E1% of about 6.6, so 1 absorbance unit is about 1.5 mg/mL of it, not 1. Use the rule of thumb only for a crude 'is there protein here at all' check on an unknown mixture, and use a sequence-derived or measured ε whenever the number will end up in a methods section.
What if my protein has no tryptophan?
Then A280 is a weak method for it and the prediction is unreliable — Pace and colleagues say so directly, describing the coefficients as 'quite reliable for proteins containing Trp residues, and less reliable for proteins that do not'. Tryptophan contributes about 5500 M⁻¹cm⁻¹ each against tyrosine's 1490, so a protein with no tryptophan has a small ε and a correspondingly small, noisy absorbance at any workable concentration. If your protein also has no tyrosine and no disulfides, ε works out to zero and the division is undefined; this calculator raises a field error there rather than returning infinity. The practical answers are to use a colourimetric assay instead, to read at 205 or 214 nm where the peptide bond itself absorbs, or — following Pace's own recommendation — to measure ε by the Edelhoch method rather than predicting it.
Does this calculator correct for nucleic acid contamination?
No, deliberately. Nucleic acid absorbs strongly at 260 nm and still substantially at 280 nm, so DNA or RNA carried through a purification inflates A280 and makes your protein look more concentrated than it is. Some tools apply a Warburg–Christian style correction that subtracts an estimated nucleic-acid contribution using the A260 and A280 readings together; that correction rests on assumptions about the contaminant that are usually not verifiable for your particular sample, and applying it silently would hide the problem rather than solve it. The honest workflow is to check the A260/A280 ratio first — a clean protein preparation sits near 0.57, and a value approaching 1.0 or above means substantial nucleic acid — and to remove the contaminant or switch to a colourimetric assay, which is far less sensitive to it. Also check the A320 baseline: a reading well above zero means light scattering from aggregates or particulates, which inflates every wavelength including 280 nm.
What happens if my absorbance is very low or very high?
The calculator still returns a number, but the interpretation line flags it. Below about 0.1 absorbance units, instrument noise becomes a large fraction of the signal, so the answer carries far more uncertainty than its digits suggest; concentrate the sample, use a longer path length, or switch to a more sensitive colourimetric assay. Above about 1.5 absorbance units, Beer–Lambert linearity fails on most bench spectrophotometers because too little light reaches the detector, and the reading systematically underestimates the true concentration; dilute and re-read rather than trusting it. That 0.1 to 1.5 range is instrument convention rather than a published standard, and the verdict text says so — check your own instrument's specified photometric range, which for a good double-beam unit may extend higher. An absorbance of exactly zero is accepted and returns exactly zero protein; a negative absorbance is refused, because it means the reference was brighter than the sample and the instrument needs re-blanking.

References& sources.

  1. [1]Pace, C. N., Vajdos, F., Fee, L., Grimsley, G. & Gray, T. (1995). 'How to measure and predict the molar absorption coefficient of a protein.' Protein Science 4(11): 2411–2423. Source of the native-state coefficients implemented here: epsilon(280) (M-1 cm-1) = (#Trp)(5,500) + (#Tyr)(1,490) + (#cystine)(125), derived from 116 epsilon values across 80 proteins. Also the source of the on-page caveat that the prediction is 'quite reliable for proteins containing Trp residues, and less reliable for proteins that do not', and of the recommendation to measure rather than predict epsilon using the Edelhoch method. Open access via PubMed Central. Retrieved and read 2026-07-29.
  2. [2]Gill, S. C. & von Hippel, P. H. (1989). 'Calculation of protein extinction coefficients from amino acid sequence data.' Analytical Biochemistry 182(2): 319–326. doi:10.1016/0003-2697(89)90602-7, PMID 2610349. Independent second authority consulted as a cross-check: confirms the additive Trp/Tyr/cystine method but uses different coefficients — 5690, 1280 and 120 — measured in 6 M guanidine hydrochloride, with a stated accuracy of about ±5%. The two coefficient sets differ by roughly 9% for a typical protein, which is why both ship as separate modes on this page. An erratum to the original paper exists. Paywalled at the publisher; abstract open via PubMed. Verified 2026-07-29.
  3. [3]Bradford, M. M. (1976). 'A rapid and sensitive method for the quantitation of microgram quantities of protein utilizing the principle of protein-dye binding.' Analytical Biochemistry 72(1–2): 248–254. doi:10.1016/0003-2697(76)90527-3. The original Coomassie dye-binding assay, cited for the standard-curve mode and for the on-page statement that the response is protein-dependent because dye binding tracks basic and aromatic residues. Paywalled; listed bibliographically.
  4. [4]Smith, P. K. et al. (1985). 'Measurement of protein using bicinchoninic acid.' Analytical Biochemistry 150(1): 76–85. doi:10.1016/0003-2697(85)90442-7. The original BCA assay. Cited for the same assay-dependence point in a chemistry — Cu²⁺ reduction followed by bicinchoninic acid chelation — that responds to a different residue set than Bradford's dye binding, which is why the two assays legitimately disagree on the same sample. Paywalled; listed bibliographically.
  5. [5]IUPAC Compendium of Chemical Terminology, 3rd ed. (the 'Gold Book'), entry 'Beer–Lambert law' (doi:10.1351/goldbook.B00626). The formal definition of A = ε c l, the units of the molar absorption coefficient (M⁻¹cm⁻¹; formally m² mol⁻¹ in SI), and the linearity assumptions the law carries. Listed WITHOUT a link deliberately: goldbook.iupac.org returned HTTP 403 to automated retrieval on 2026-07-29, so this reference was not machine-verified and is given by DOI for a reader to check directly rather than shipped as a link that may look dead.
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