Protein Concentration Calculator

Q: What A280 reading equals 1 mg/mL of protein?

It depends on the protein. The 1 mg/mL value is called the 0.1% extinction coefficient (E1%) or A280 of a 1 mg/mL solution. BSA: about 0.66 IgG: about 1.4 Lysozyme: about 2.7 To get it yourself, divide ε (M⁻¹·cm⁻¹) by the molecular weight (g/mol). For BSA: 43,824 ÷ 66,430 = 0.66.

Q: What is the linear range for A280 protein readings?

Most spectrophotometers are reliable between about 0.1 and 1.5 AU , and some stretch to 2.0 AU. Above 2.0 AU, too little light reaches the detector and the reading flattens out, so you get a number that is too low. Below 0.05 AU, noise from the buffer and the machine is bigger than the signal, so the error is large. Dilute a high sample, or concentrate a low one, so the reading lands near the middle.

Q: Which protein assay should I use: Bradford, BCA, or Lowry?

Pick based on what is in your buffer: Bradford : fastest (about 5 minutes) and works with reducing agents like DTT, but detergents such as SDS and Triton ruin it. Response varies a lot between proteins. BCA : tolerates detergents well and gives more even results across proteins, but needs 30 minutes at 37°C and is spoiled by reducing agents. Lowry : very sensitive but slow, with multiple steps and many interfering chemicals. BCA is the usual choice for lysates with detergent. Bradford is the choice when you need a fast answer.

Protein Concentration Calculator

Introduction

This Protein Concentration Calculator tells you how much protein is in your sample. You can work two ways. Use the A280 method, which reads light at 280 nm and uses the Beer–Lambert law. Or use a standard curve from a Bradford, BCA, or Lowry assay.

For the A280 mode, type in your absorbance reading, the extinction coefficient (ε), and the path length of your cuvette. Pick a common protein like BSA, IgG, or lysozyme and the ε and molecular weight fill in for you. You can also paste an amino acid sequence or a UniProt ID, and the tool will work out ε and MW for you.

For the standard curve mode, enter the slope and intercept of your fit, or just two standard points. Add your sample signal and the calculator reads the concentration off the line.

Results come back in mg/mL, µg/µL, g/L, and µM. You can add a blank reading and a dilution factor so the number matches your real stock. The tool also checks your A280/A260 ratio to flag DNA or RNA in your sample, and warns you if a reading is too high or too low to trust. Step-by-step math and a chart are shown, so you can check every number and use it in your lab notebook.

How to use our Protein Concentration Calculator

Enter your absorbance reading or assay signal, plus a few sample details, and the calculator returns your protein concentration in mg/mL, µg/µL, g/L, and µM, with a gauge, warnings, and full step-by-step math.

Quick Pick Presets: Click a preset button to load a ready-made example, like an IgG A₂₈₀ reading or a Bradford standard curve.

Calculation Mode: Pick "A₂₈₀ — Beer–Lambert Law" if you read your sample on a spectrophotometer. Pick "Standard Curve" if you ran a Bradford, BCA, or Lowry assay.

Protein: Choose your protein from the list. This fills in the extinction coefficient (ε) and molecular weight for you. Pick "Custom Protein" to enter your own.

Amino Acid Sequence: For a custom protein, paste the single-letter sequence and click the button. The tool works out ε and MW from the Trp, Tyr, and Cys content.

UniProt Accession: Type an ID like P02769 and click fetch. The sequence is pulled from UniProt, then ε and MW are set for you.

Absorbance (A₂₈₀): Type the 280 nm reading from your spectrophotometer.

Blank / Reference Absorbance: Type your buffer blank reading. It is taken off the sample reading. Leave it at 0 if you have none.

Absorbance at 260 nm: Optional. Add it to get the A₂₈₀/A₂₆₀ purity ratio. It does not change the concentration.

Molar Extinction Coefficient (ε): Enter the value in M⁻¹·cm⁻¹ from your protein datasheet, or let the list or sequence tool fill it in.

Pathlength: Enter the light path of your cuvette and pick cm or mm. Most cuvettes are 1 cm; microvolume readers are often 0.1 mm.

Assay Type: Choose Bradford, BCA, Lowry, or Other. This is just a label for your results.

Sample Signal: Enter the reading for your sample: absorbance, OD, or RFU.

Blank / Background Signal: Enter your blank well reading. It is subtracted before the concentration is read off the curve.

Curve Concentration Unit: Pick the unit your standards were made in: mg/mL, µg/mL, g/L, or µM. If you pick µM, you must also enter a molecular weight.

Standard Curve Input Method: Choose "Slope & intercept" if you already have a linear fit, or "Two-point entry" to build the line from two standards.

Slope (m): Enter the signal per unit of concentration from your fit line.

Intercept (b): Enter the signal at zero concentration. Leave it blank to use 0.

Standard 1 and Standard 2: For two-point entry, type the known concentration and the measured signal for each standard. The two concentrations must be different.

Molecular Weight: Enter your protein's MW in g/mol or kDa. This unlocks the mass units in A₂₈₀ mode and the µM result in curve mode.

Dilution Factor: If you diluted your sample 10× before reading, enter 10. Use 1 for an undiluted sample.

Round results and Decimal places: Turn rounding on or off, and pick 2, 3, 4, or 6 decimal places for the output.

Show step-by-step calculation: Tick this box to see every formula and number used to get your answer.

Calculate and Reset: Click Calculate to see your results, or Reset to clear everything back to the default values.

What Is Protein Concentration?

Protein concentration tells you how much protein is in a set amount of liquid. Lab workers write it as mg/mL, µg/µL, g/L, or µM (micromolar). The first three are mass units, so they count the weight of protein. µM is a molar unit, so it counts the number of protein molecules. You need to know this number before you run a gel, set up an enzyme test, load a Western blot, or store a purified sample.

Two Common Ways to Measure Protein

1. A280 and the Beer–Lambert Law

Protein soaks up UV light at 280 nm. That happens because of three amino acids: tryptophan (W), tyrosine (Y), and cystine (paired cysteines). A spectrophotometer shines light through your sample and reports the absorbance, called A₂₈₀.

The Beer–Lambert law links that reading to concentration:

c = A ÷ (ε × ℓ)

A = absorbance at 280 nm, after you subtract the blank
ε = molar extinction coefficient (M⁻¹·cm⁻¹), which is how strongly that protein grabs light
ℓ = pathlength, how far the light travels through the sample. Normal cuvettes are 1 cm. Microvolume readers like a NanoDrop use about 0.1 mm.

A280 is fast and does not destroy your sample, but it only works if you know ε for your exact protein. You can look it up, or work it out from the amino acid sequence with the Pace method: ε = (5500 × Trp) + (1490 × Tyr) + (125 × disulfide bonds).

2. Standard Curve Assays (Bradford, BCA, Lowry)

These are color assays. You mix a dye or reagent with your protein and the tube changes color. More protein means more color. You also test a set of known standards, usually BSA, and plot signal against concentration. That straight line is your standard curve:

signal = m × C + b, so C = (signal − b) ÷ m

Bradford: Coomassie dye, read near 595 nm. Quick, but reacts unevenly with different proteins and hates detergents.
BCA: copper based, read near 562 nm. Handles detergents well, but needs warm incubation.
Lowry: the older copper and Folin method. Sensitive, but slower and fussier.

Color assays work even when you do not know ε, but your answer is only as good as your standards. Always read your sample inside the range your standards cover.

Things That Change Your Answer

Blank subtraction. Buffer and reagents absorb light too. Always subtract a blank.
Dilution factor. If you diluted 10× before reading, multiply the result by 10 to get the real stock concentration.
Molecular weight (MW). MW links mass units to molar units. Without MW you cannot switch between mg/mL and µM.
Linear range. Readings above about 2.0 AU are often too high to trust. Readings below about 0.05 AU are mostly noise. Dilute or concentrate your sample to land in the middle.
A280/A260 ratio. DNA and RNA absorb strongly at 260 nm. A pure protein sample usually gives a ratio near or above 1.7. A lower ratio hints at nucleic acid contamination, and your A280 number will read too high.

Quick Unit Facts

1 mg/mL = 1 g/L = 1 µg/µL. These three are the same number, just written differently. To get molarity: µM = (mg/mL ÷ MW in g/mol) × 10⁶. A big protein like IgG (about 150,000 g/mol) gives far fewer molecules per mg than a small one like insulin (about 5,808 g/mol).

Which Method Should You Pick?

Use A280 when your protein is pure, you know its sequence or ε, and you want the sample back. Use a standard curve assay when your protein is in a crude lysate, when you do not know ε, or when the amount is too small for a clean UV reading.

Formulas used

Blank-corrected absorbance

A_{\mathrm{corr}} = A_{280} - A_{\mathrm{blank}}

Molar concentration from Beer-Lambert law (with dilution factor)

c = \frac{A_{\mathrm{corr}}}{\varepsilon \, \ell} \times DF

Mass concentration from molar concentration

c_{\mathrm{mass}}\ (\mathrm{g/L}) = c\ (\mathrm{mol/L}) \times MW\ (\mathrm{g/mol})

Protein purity ratio

R = \frac{A_{280}}{A_{260}}

Concentration interpolated from a linear standard curve

C = \frac{(y - y_{\mathrm{blank}}) - b}{m} \times DF

Slope and intercept from two standard points

m = \frac{y_2 - y_1}{C_2 - C_1}, \qquad b = y_1 - m\,C_1

Molar extinction coefficient from sequence (Pace method)

\varepsilon = 5500\,n_{\mathrm{Trp}} + 1490\,n_{\mathrm{Tyr}} + 125 \left\lfloor \frac{n_{\mathrm{Cys}}}{2} \right\rfloor

Molecular weight from sequence residue masses

MW = \sum_{i} m_{\mathrm{residue},i} + 18.01528

Frequently asked questions

How do you convert mg/mL to µM for a protein?

Divide the mass concentration by the protein's molecular weight, then multiply by one million.

µM = (mg/mL ÷ MW in g/mol) × 1,000,000

Example: 2 mg/mL of BSA (MW 66,430 g/mol) = (2 ÷ 66,430) × 1,000,000 = 30.1 µM.

The same mass of a small protein gives many more molecules, so a smaller MW always gives a bigger µM number.

What A280 reading equals 1 mg/mL of protein?

It depends on the protein. The 1 mg/mL value is called the 0.1% extinction coefficient (E1%) or A280 of a 1 mg/mL solution.

BSA: about 0.66
IgG: about 1.4
Lysozyme: about 2.7

To get it yourself, divide ε (M⁻¹·cm⁻¹) by the molecular weight (g/mol). For BSA: 43,824 ÷ 66,430 = 0.66.

How do you calculate the extinction coefficient from an amino acid sequence?

Count the tryptophan (W), tyrosine (Y) and cysteine (C) residues, then use the Pace formula:

ε = (5500 × Trp) + (1490 × Tyr) + (125 × disulfide bonds)

Each disulfide bond uses two cysteines, so divide your Cys count by 2 and round down. If your protein is fully reduced, drop the disulfide term.

A protein with no Trp and no Tyr has ε near 0, so A280 will not work for it.

Why is my A280/A260 ratio below 1.7?

DNA and RNA soak up light strongly at 260 nm. A low ratio means nucleic acid is in your sample.

Pure protein usually reads 1.7 or higher. Below that, your A280 number is too high because the DNA adds extra absorbance at 280 nm too.

Fix it by treating the sample with nuclease, running it over a column, or switching to a Bradford or BCA assay, which do not react with nucleic acids.

What is the linear range for A280 protein readings?

Most spectrophotometers are reliable between about 0.1 and 1.5 AU, and some stretch to 2.0 AU.

Above 2.0 AU, too little light reaches the detector and the reading flattens out, so you get a number that is too low.

Below 0.05 AU, noise from the buffer and the machine is bigger than the signal, so the error is large.

Dilute a high sample, or concentrate a low one, so the reading lands near the middle.

Why does a NanoDrop use a 0.1 mm pathlength?

Microvolume readers hold a tiny drop between two pedestals instead of filling a cuvette. That drop is only about 1 mm or 0.1 mm thick.

A shorter path means less light is absorbed, so you can read concentrated samples without diluting them. A 1 cm cuvette is 100 times longer than 0.1 mm, so the same sample reads 100 times higher in a cuvette.

Most instruments auto-correct back to a 1 cm equivalent, so check whether your reported number is already normalized before you do the math yourself.

Which protein assay should I use: Bradford, BCA, or Lowry?

Pick based on what is in your buffer:

Bradford: fastest (about 5 minutes) and works with reducing agents like DTT, but detergents such as SDS and Triton ruin it. Response varies a lot between proteins.
BCA: tolerates detergents well and gives more even results across proteins, but needs 30 minutes at 37°C and is spoiled by reducing agents.
Lowry: very sensitive but slow, with multiple steps and many interfering chemicals.

BCA is the usual choice for lysates with detergent. Bradford is the choice when you need a fast answer.

Why do Bradford and A280 give different results for the same sample?

They measure different things. A280 counts tryptophan and tyrosine. Bradford counts how much Coomassie dye binds, which depends mostly on arginine and basic residues.

Bradford curves are usually built with BSA standards. If your protein has a different residue mix than BSA, the dye binds differently and the answer shifts. Differences of 2-fold or more are common.

Use the same method every time so your numbers can be compared. Do not mix A280 and Bradford values in one experiment.

How do you apply a dilution factor to a protein concentration?

Multiply the measured concentration by the dilution factor to get the original stock.

If you added 10 µL of sample to 90 µL of buffer, that is a 10× dilution. A reading of 0.4 mg/mL means your stock is 0.4 × 10 = 4 mg/mL.

The dilution factor is total volume ÷ sample volume. So 10 µL into 90 µL buffer gives 100 ÷ 10 = 10.

Is mg/mL the same as µg/µL?

Yes. 1 mg/mL = 1 µg/µL = 1 g/L. All three are the same amount, just written with different prefixes.

1 mg = 1000 µg and 1 mL = 1000 µL, so the 1000s cancel out. A 5 mg/mL stock is also 5 µg/µL and 5 g/L.

This is handy for loading gels: if you need 20 µg of protein from a 5 µg/µL stock, load 4 µL.

Why is my calculated protein concentration negative?

A negative number means your blank was bigger than your sample reading, or your curve intercept was above your corrected signal.

Common causes: a bubble or fingerprint on the blank cuvette, a blank made from the wrong buffer, or a sample with almost no protein in it.

Re-blank with the exact buffer your sample is in, wipe the cuvette, and read again. If the sample really is near zero, you need to concentrate it or use a more sensitive assay.

Do I need a molecular weight to calculate protein concentration?

Not for mass units. A280 with ε gives molarity directly, and a Bradford curve in mg/mL gives mass directly.

You need MW only to switch between the two. Without it you cannot turn mg/mL into µM, or µM into mg/mL.

For an unknown protein mix like a cell lysate, there is no single MW, so report mass units only.

Why should my sample fall inside the range of my standards?

A standard curve is only proven straight between your lowest and highest standard. Outside that range you are guessing.

Most color assays flatten out at high protein, so reading above your top standard gives a number that is too low. Reading below your bottom standard sits in the noise.

If your sample reads outside the range, dilute it and run it again rather than extrapolating the line.

How much protein do I need to load on an SDS-PAGE gel?

For Coomassie staining, load 10 to 30 µg of total protein per lane on a mini gel. For silver stain, 1 to 5 µg is plenty. For a Western blot of an abundant target, 10 to 20 µg works.

Keep the volume under your well size, usually 15 to 25 µL including loading buffer. That is why knowing mg/mL matters: from a 2 mg/mL stock, 20 µg needs 10 µL.

What makes A280 read too high?

Several things absorb at 280 nm besides your protein:

Nucleic acids: DNA and RNA spill over from 260 nm
Cloudiness: aggregates and dust scatter light, which the machine records as absorbance
Buffer parts: imidazole, DTT, and some detergents absorb in the UV
Phenol red from culture media

Check A320 or A340; a clear sample should read near zero there. If it does not, your sample is cloudy and you should spin or filter it.

Protein Concentration Results

Rate this calculator

Introduction

How to use our Protein Concentration Calculator

What Is Protein Concentration?

Two Common Ways to Measure Protein

1. A280 and the Beer–Lambert Law

2. Standard Curve Assays (Bradford, BCA, Lowry)

Things That Change Your Answer

Quick Unit Facts

Which Method Should You Pick?

Formulas used

Frequently asked questions

How do you convert mg/mL to µM for a protein?

What A280 reading equals 1 mg/mL of protein?

How do you calculate the extinction coefficient from an amino acid sequence?

Why is my A280/A260 ratio below 1.7?

What is the linear range for A280 protein readings?

Why does a NanoDrop use a 0.1 mm pathlength?

Which protein assay should I use: Bradford, BCA, or Lowry?

Why do Bradford and A280 give different results for the same sample?

How do you apply a dilution factor to a protein concentration?

Is mg/mL the same as µg/µL?

Why is my calculated protein concentration negative?

Do I need a molecular weight to calculate protein concentration?

Why should my sample fall inside the range of my standards?

How much protein do I need to load on an SDS-PAGE gel?

What makes A280 read too high?

Protein Concentration Calculator

Protein Concentration Results

Rate this calculator

Introduction

How to use our Protein Concentration Calculator

What Is Protein Concentration?

Two Common Ways to Measure Protein

1. A280 and the Beer–Lambert Law

2. Standard Curve Assays (Bradford, BCA, Lowry)

Things That Change Your Answer

Quick Unit Facts

Which Method Should You Pick?

Formulas used

Frequently asked questions

How do you convert mg/mL to µM for a protein?

What A280 reading equals 1 mg/mL of protein?

How do you calculate the extinction coefficient from an amino acid sequence?

Why is my A280/A260 ratio below 1.7?

What is the linear range for A280 protein readings?

Why does a NanoDrop use a 0.1 mm pathlength?

Which protein assay should I use: Bradford, BCA, or Lowry?

Why do Bradford and A280 give different results for the same sample?

How do you apply a dilution factor to a protein concentration?

Is mg/mL the same as µg/µL?

Why is my calculated protein concentration negative?

Do I need a molecular weight to calculate protein concentration?

Why should my sample fall inside the range of my standards?

How much protein do I need to load on an SDS-PAGE gel?

What makes A280 read too high?

Related Biology calculators

Ligation Calculator

Peptide Mass Calculator

Tm Calculator