The protein concentration calculator above works two ways. In A280 mode it applies the Beer-Lambert relationship to convert an ultraviolet absorbance into a concentration using your protein's molar extinction coefficient, molecular weight and the cuvette path length. In colorimetric mode it reads a Bradford or BCA absorbance off the slope and intercept of a standard curve you have already fitted. Both modes apply a dilution factor and report the total protein in the tube.
Arb Digital publishes this as a free bench tool that runs entirely in your browser. The two methods are on one page deliberately, because the choice between them is the real decision: A280 is fast, non-destructive and needs no reagents but depends on knowing a coefficient, while a dye-binding assay needs standards and consumes sample but works on proteins whose sequence you do not know.
What This Protein Concentration Calculator Does
The A280 method rests on the fact that tryptophan, tyrosine and disulphide-bonded cystine absorb ultraviolet light at 280 nm while the rest of a protein does not. The ExPASy ProtParam documentation sets out the standard calculation: the molar extinction coefficient of a protein is the number of tyrosines multiplied by 1,490, plus the number of tryptophans multiplied by 5,500, plus the number of cystines multiplied by 125.
Those per-residue values come from the work described in How to measure and predict the molar absorption coefficient of a protein by Pace and colleagues, published in Protein Science in 1995, which calibrated the prediction against measured coefficients for a large set of proteins. ProtParam notes that it uses the Edelhoch method with these Pace coefficients, which is why extinction coefficients from different tools generally agree.
Colorimetric mode does something completely different. It has no model of the protein at all; it simply inverts a straight line fitted through standards of known concentration. That makes it agnostic to sequence and correspondingly dependent on the quality of your curve.
How to Use It
- Choose the method that matches your measurement. A 280 nm reading on a spectrophotometer or a microvolume instrument goes in A280 mode; a 595 nm or 562 nm assay reading goes in curve mode.
- Enter the blank-corrected absorbance. Subtracting the buffer blank is not optional, particularly for BCA where the reagent itself develops colour.
- Supply the extinction coefficient and molecular weight for your specific protein, computed from its sequence.
- In curve mode, enter the fitted slope and intercept from your own standards, run on the same plate on the same day.
- Set the dilution factor last, and check the pre-dilution figure in the grid so you can see what the instrument actually measured.
The Formula and How It's Calculated
Beer-Lambert states that absorbance equals the molar extinction coefficient multiplied by the molar concentration multiplied by the path length. Rearranged for concentration in mass terms, concentration in mg/mL equals absorbance multiplied by molecular weight, divided by the product of extinction coefficient and path length. With an absorbance of 0.500, a molecular weight of 66,463 daltons, a coefficient of 43,824 and a 1 cm path, that is 0.500 × 66,463 ÷ 43,824 = 0.758 mg/mL. The molar concentration is 0.500 ÷ 43,824 = 1.14 × 10⁻⁵ M, or 11.41 micromolar, and a 1 mL sample therefore contains 0.758 mg of protein.
The absorbance of a 1 mg/mL solution, often written A at 0.1 per cent and reported for a 1 cm path, is simply the extinction coefficient divided by the molecular weight: 43,824 ÷ 66,463 = 0.659. That figure is handy because it lets you read concentration straight off an absorbance without touching the molar arithmetic.
Colorimetric mode inverts the fitted line: concentration equals absorbance minus intercept, divided by slope. A reading of 0.500 against a curve with a slope of 0.65 and an intercept of 0.02 gives (0.500 − 0.020) ÷ 0.65 = 0.738 mg/mL. Either result is then multiplied by the dilution factor.
The Assumptions A280 Quietly Makes
Three of them matter. First, the extinction coefficient must be right for your protein in its current state — the value predicted from sequence applies to a folded protein in water, and denaturants shift it. Second, nothing else in the sample absorbs at 280 nm; nucleic acid contamination is the classic offender, since DNA and RNA absorb strongly there and inflate the reading. Third, the sample must be clear, because scattering from aggregates or particulates adds apparent absorbance that has nothing to do with concentration.
The 260 over 280 ratio is the standard check on the second point. A pure protein sits near 0.6, while a value climbing toward 1.0 or above indicates significant nucleic acid. Scattering is diagnosed by reading at 320 nm, where protein should not absorb at all; anything meaningful there should be subtracted or the sample cleared before measurement. Neither correction is applied automatically here, because both should be decisions you make knowingly.
When a Standard Curve Is the Better Answer
A colorimetric assay needs no sequence information, which makes it the only option for a crude lysate, a mixed sample, or a protein you have not identified. It is also far more sensitive than A280 at low concentrations. The cost is that the answer is only ever relative to the standard you used, and the standards are chosen for convenience rather than resemblance to your protein.
This produces a systematic effect that surprises people the first time they meet it: dye-binding responses vary with amino acid composition, so two proteins at genuinely identical concentrations can give measurably different readings in the same assay. Bradford in particular responds strongly to basic and aromatic residues. The result is precise and reproducible, but it is a concentration expressed in units of the standard, and comparing across assays or across standards is where errors creep in. Our calibration curve calculator handles fitting the line itself.
Detergents, Reducing Agents and Buffer Interference
Every colorimetric assay has a list of things that break it. Bradford is intolerant of detergents, so a lysate containing SDS or Triton will give a wrong answer or none at all. BCA copes with detergents but is disrupted by reducing agents such as dithiothreitol and beta-mercaptoethanol, which reduce copper and generate colour independently of protein. Chelating agents interfere with BCA for the opposite reason.
The practical defence is to make the standards up in exactly the buffer the sample sits in, so any interference affects both sides of the comparison equally. Where the interfering component is too concentrated for that, dilution before the assay is usually more reliable than a compatibility reagent. This is also a genuine argument for A280 where it is available: it has no reagents to interfere with, though it has its own sensitivity to anything absorbing at the same wavelength.
Why the Absorbance Window Matters
Spectrophotometers are least accurate at both ends of their range. Below about 0.1 absorbance units the signal is close to instrument noise and small blanking errors dominate; above about 1.0 the detector receives so little light that the relationship between concentration and reading starts to bend away from linear. The bar under the result shows where your reading sits in that window.
The fix in both directions is dilution or concentration of the sample rather than trusting a number outside the range, and the dilution factor field is there to make the correction explicit. Reading a diluted sample at 0.4 and multiplying by ten is far more reliable than reading the neat sample at 2.5. If you need to work out the dilution itself, the solution dilution calculator handles the stock and final volumes.
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Browse All Free Tools Contact Arb DigitalCommon Mistakes to Avoid
- Using a generic extinction coefficient instead of one computed from your protein's own sequence.
- Ignoring nucleic acid contamination, which absorbs strongly at 280 nm and inflates every A280 reading.
- Reusing a standard curve from a previous plate or a previous day, when reagent age and incubation both shift the response.
- Running Bradford on a detergent-containing lysate, or BCA on a sample containing reducing agents.
- Trusting an absorbance above about 1.0 rather than diluting the sample and applying the dilution factor.
Related Free Tools From Arb Digital
To get the extinction coefficient and molecular weight this page needs, run your sequence through the protein molecular weight calculator. For fitting the standards, use the calibration curve calculator; for the nucleic acid equivalent, the DNA concentration calculator. Reaction setup is covered by the molarity calculator and solution dilution calculator, and composition by the mass percent calculator. Browse the full free online tools hub for more.
Frequently Asked Questions
Concentration in mg/mL equals absorbance multiplied by molecular weight, divided by the molar extinction coefficient multiplied by the path length in centimetres. An absorbance of 0.500 for a 66,463 dalton protein with a coefficient of 43,824 in a 1 cm cuvette gives 0.758 mg/mL.
Predict it from the sequence. The standard calculation is the number of tyrosines multiplied by 1,490, plus tryptophans multiplied by 5,500, plus cystines multiplied by 125, which is the method the ExPASy ProtParam documentation describes.
It is the absorbance a 1 mg/mL solution of the protein would give in a 1 cm cuvette, and it equals the molar extinction coefficient divided by the molecular weight. It lets you convert an absorbance to mg/mL in one step.
Nucleic acid contamination and light scattering are the two usual causes. Check the ratio of absorbance at 260 nm to 280 nm, which sits near 0.6 for pure protein, and read at 320 nm to detect scattering from aggregates or particulates.
A280 is fast, reagent-free and non-destructive but needs a known extinction coefficient. Bradford and BCA work without sequence information and are more sensitive, but they measure relative to a standard and each is broken by different buffer components.
Because dye binding depends on amino acid composition, and Bradford responds particularly strongly to basic and aromatic residues. The result is a concentration expressed in units of whatever standard you used, not an absolute measurement.
Roughly 0.1 to 1.0 on most instruments. Below that the signal approaches noise and blanking errors dominate; above it the response bends away from linear. Dilute the sample and apply a dilution factor rather than trusting a reading outside the window.
This calculator provides general educational estimates from values you supply and does not replace validated laboratory procedure. Assay performance depends on buffer composition, instrument calibration and standard quality, and results should be confirmed against your own controls.
