Why do I subtract the blank absorbance?

The reagent and buffer absorb a little on their own, which is what the blank measures. Subtracting it gives the absorbance due to protein alone, so the concentration is not inflated by the assay chemistry.

What path length do I use for a microplate well?

The well height from the plate datasheet, typically 0.3 to 0.6 cm rather than the 1 cm of a cuvette. Because absorbance scales with path length, using 1 cm for a 0.3 cm well overstates the concentration by more than three times.

How do I choose an extinction coefficient?

Use the published value for your protein if you have one, calculated from the sequence with tools such as ProtParam, or the standard figure for BSA if your sample is an unknown mix. If the protein is impure or aggregated, use a standard curve instead: it needs no assumption at all.

Why is my Bradford result lower than my A280 result?

A Bradford or BCA assay reacts with particular amino acids, so it responds to composition rather than to absorbance, and can under-read a protein rich in the residues it does not detect. A280 reads any aromatic residue plus nucleic acid contamination. Neither is automatically right; they should agree within about 20% on a clean sample.

Why do tables give so many different extinction coefficients for BSA?

Because they use different units. The same protein is 43,824 M^-1 cm^-1, 43.8 mM^-1 cm^-1, 5.5 in the 10^4 M^-1 cm^-1 convention, 0.66 A280 per mg/mL, or an A280 of about 0.58 for a 1 mg/mL solution measured at 279 nm rather than 280 nm. Pick one convention and stay in it. The calculator uses molar mM^-1 cm^-1 and labels every preset with it, so a 1 cm cell reading of 0.70 A against BSA gives about 1.06 mg/mL.

When is a standard curve better than Beer-Lambert?

Whenever the extinction coefficient is an assumption rather than a measurement: impure preparations, protein mixtures, colourimetric assays such as Bradford or BCA, aggregated proteins, and anything outside the linear range of a cuvette reading.

Why does the calculator warn about extrapolation?

A standard curve is only trustworthy between the concentrations you calibrated. If your sample reads above the top standard or below the zero-protein blank, the number is an extrapolation of the line, not a measurement, and it should be re-read against standards that bracket it.

Can I use this to make a specific dilution?

Yes. Enter your target concentration and the final volume you want, and the calculator returns the fold dilution plus the volume of stock and diluent to mix. It refuses a target above the stock concentration, because dilution cannot raise concentration.

Protein Concentration Calculator — Free Online Calculator

What Is the Protein Concentration Calculator?

The Protein Concentration Calculator converts a spectrophotometer reading into the concentration of a protein stock, in both molar and mass units, and back-corrects any dilution made before the sample was read.

Two methods cover most laboratory practice. Beer-Lambert needs an extinction coefficient and a known molecular weight, which makes it the fastest route for a reasonably pure protein at 280 nm. A standard curve needs no assumptions about the protein at all, which makes it the safer route for anything impure, aggregating or assayed by colour reaction.

The calculator also reports the mass of protein in the volume you actually loaded, which is the number that matters when you are deciding whether to pool fractions or re-run a gel.

How Does the Protein Concentration Calculator Work?

The raw reading always has the blank subtracted first. A blank of 0.02 A against a sample of 0.72 A gives a usable absorbance of 0.70 A.

For Beer-Lambert the molar concentration is c = A / (epsilon x l). The extinction coefficient is quoted for a 1 cm cell, so the path length divides the reading: the same protein at 0.5 cm reads half as much absorbance for the same concentration.

Mass concentration follows from the molecular weight: c (mg/mL) = c (mM) x MW (Da) / 1000, because 1 mM of a 66,430 Da protein weighs 6.643 mg per litre of molarity.

For the standard-curve method, two known concentrations give a straight line. The slope is the rise in absorbance over the rise in concentration, and the intercept absorbs whatever the reagent itself reads at zero protein.

The sample is then interpolated: c = (A - intercept) / slope. If the answer falls outside the range the standards covered, the result is an extrapolation and the tool says so.

The dilution factor is applied last. The instrument measures the diluted sample, so multiplying by the fold dilution recovers the stock you actually hold.

Protein Concentration Calculator Formula & Variables

The core mathematical equation utilized by this calculator is expressed as:

\text{Beer-Lambert}\; c = \frac{A}{\varepsilon \, l} \qquad \text{Standard curve}\; m = \frac{A_{2} - A_{1}}{c_{2} - c_{1}},\; b = A_{1} - m\,c_{1} \\ c = \frac{A - b}{m} \qquad \text{Dilution}\; \mathrm{DF} = \frac{c_{\text{stock}}}{c_{\text{target}}},\; V_{\text{stock}} = \frac{V_{\text{final}}}{\mathrm{DF}}

Variable Definitions

Symbol	Variable Meaning & Units
A	Blank-corrected absorbance at the assay wavelength
epsilon	Extinction coefficient in mM^-1 cm^-1, quoted for a 1 cm cell
l	Cuvette path length in centimetres
m	Slope of the standard curve in absorbance per mg/mL
b	Intercept of the standard curve
DF	Dilution factor applied to the stock before it was read

Beer-Lambert says absorbance is proportional to concentration multiplied by the path length and the extinction coefficient, so dividing the blank-corrected absorbance by the product of path length and extinction coefficient returns the molar concentration. Because the extinction coefficient is quoted for a 1 cm cell, the path length must divide the result: a 0.5 cm well reads half the absorbance of a 1 cm cuvette for the same concentration. Converting molar to mass concentration needs the molecular weight, since 1 mM of a 66,430 Da protein weighs 6.643 mg/mL. Where the extinction coefficient is unknown, two known standards define a straight line and the sample is interpolated on it, which is exactly what a Bradford or BCA plate does. The dilution factor is applied last, because the instrument reads the diluted sample while the stock is what sits in the freezer.

How to Use the Protein Concentration Calculator

Choose the method. Use Beer-Lambert if you know the extinction coefficient of your protein; use a standard curve if you calibrated one for this assay.
Enter the sample absorbance exactly as the instrument shows it, and the blank absorbance measured with the same buffer and reagent.
For Beer-Lambert, pick the protein preset that matches your sample and check the extinction coefficient and molecular weight against the literature value for your construct.
Set the path length. A standard 1 cm cuvette is the usual case; for a microplate well, use the well height from the plate datasheet.
For a standard curve, enter the concentration and reading of two standards that bracket your sample. A zero-protein blank and a single known standard are enough.
Enter the fold dilution you made before reading, or 1 if you read the sample neat.
Optionally enter the volume you loaded to get the total protein mass, and a target concentration with a final volume to get a dilution recipe.
Read the stock concentration first: that is the number you act on. The cuvette concentration is shown next to it for comparison with the assay range.

Step-by-Step Example Calculation

A five-fold diluted BSA stock read at 280 nm

Input Values:

method:a280_extinction

preset:bsa

absorbance:0.72

blankAbsorbance:0.02

pathLength:1

dilutionFactor:5

sampleVolume:0.2

Worked Steps: Blank-corrected absorbance = 0.72 - 0.02 = 0.70 A. Beer-Lambert with the BSA coefficient of 43.8 mM^-1 cm^-1 (43,824 M^-1 cm^-1) in a 1 cm cell gives 0.70 / 43.8 = 0.016 mM, which is 0.016 x 66,430 / 1000 = 1.0617 mg/mL in the cuvette. The 0.2 mL cuvette therefore held 0.2123 mg of protein. Adding back the five-fold dilution gives a stock of 5.3083 mg/mL. The same dilution calculated from the mass extinction coefficient of 0.66 A280 per mg/mL gives 0.70 / 0.66 = 1.0606 mg/mL, which agrees to about 0.1 percent.

Understanding Your Result

Stock concentration: the diluted reading multiplied by the dilution factor. This is the concentration in your vial.

Diluted sample concentration: what the instrument actually saw. Compare it with the linear range of the assay before trusting it.

Molar concentration: useful for stoichiometry, binding calculations and molarity-based buffers.

Mass in the sample volume: the total protein that went into the cuvette or well.

Standard curve: the slope and intercept, so you can read off any concentration on the same plate.

Standard range and extrapolation warning: whether the sample sat inside the range your standards covered.

Dilution plan: the fold dilution required and the stock and diluent volumes that reach your target concentration.

Range assessment: a plain warning when the concentration is too high or too close to background for a reliable reading.

Factors That Affect the Result

Extinction coefficient: it scales with the number of aromatic residues per dalton, so the same absorbance means very different concentrations for insulin and for IgG.
Path length: a microplate well is a fraction of a cuvette deep, so an absorbance from a plate cannot be read with a 1 cm assumption.
Purity: nucleic acids, free nucleotides and aromatic small molecules all add absorbance at 280 nm and inflate the result.
Buffer and reagent blank: a coloured or absorbing buffer shifts the baseline, which is why the blank must be made from the same mix.
Aggregation and self-association: proteins that oligomerise expose fewer aromatic residues per unit mass and can break Beer-Lambert behaviour.
Temperature and pH: both shift aromatic side-chain exposure slightly, and high salt or detergent can change the apparent extinction coefficient.

When Should You Use This Calculator?

After a purification step: converting a fraction reading into a concentration before pooling or storing it.
Before loading a gel or a cuvette: checking that the sample is in the assay range and not saturating the instrument.
Standard preparation: diluting a stock to a known working concentration for a calibration curve or an enzyme assay.
Mass balance: turning an absorbance into micrograms so you can size a column run or a dialysis.
Dilution planning: working out how much stock and diluent make a target concentration at a chosen final volume.
QC of a protein prep: comparing measured concentration with expected yield per litre of culture.

Assumptions & Limitations

Assumes Beer-Lambert behaviour, which holds only while the protein is monomeric, in range, and free of absorbing contaminants.
Two points define a straight line exactly, so a standard-curve result from two standards carries no measure of curve fit. Use three or more standards and a regression when the assay is nonlinear.
Does not correct for temperature, pH or detergent effects on the extinction coefficient.
Does not account for light scattering from turbid or precipitated samples, which adds a wavelength-dependent baseline.
Assumes the blank reading matches the sample buffer and reagent exactly.
Reports concentrations but not purity: a 260/280 or 280/230 ratio has to be read separately.

Frequently Asked Questions

Calculation Accuracy & Reference Note

The arithmetic is exact for the values you enter. The published coefficients used by the presets are the standard 280 nm figures for BSA, IgG and small peptides, rounded as they are normally quoted in the literature.

Standard Reference: Beer-Lambert law as applied to protein absorbance at 280 nm; standard extinction coefficients for BSA, IgG and small peptides; Bradford protein assay standard curve practice using a BSA standard of known concentration.