Protein Mass & Peptide Molecular Weight Calculator

A Free Protein Mass & Peptide Molecular Weight Calculator

This calculator turns an amino acid sequence into the numbers you actually need for lab work: the molecular weight of the peptide or protein, its UV extinction coefficient, and how much of it to weigh out for a target concentration. Whether you're checking a peptide synthesis order, reading an SDS-PAGE gel, planning a dilution series, or reconstituting a lyophilized protein stock, this tool covers the calculation in one place instead of spreading it across three different spreadsheets.

Paste in any sequence using standard 1-letter amino acid codes and the calculator instantly works out the average molecular weight, the monoisotopic mass used in mass spectrometry, and a full breakdown of the amino acid composition. From there, it can estimate a UV extinction coefficient and back out a protein concentration from a simple A280 absorbance reading, or convert freely between mass, volume, and molar concentration for dosing, dilution, and reconstitution.

What Is Molecular Weight for a Peptide or Protein?

A protein or peptide is a chain of amino acids linked together by peptide bonds. Every time two amino acids join, the reaction releases one small water molecule — so a chain of amino acids weighs slightly less than the simple sum of its individual, free amino acids. The molecular weight of the finished chain is what's left after accounting for all of that lost water.

In practice, biochemists use a standard shortcut: each amino acid has a known 'residue mass' — its own weight already minus one water. Add up the residue masses for every amino acid in the chain, then add back exactly one water molecule for the free ends of the chain (the N-terminal amine and the C-terminal carboxyl group), and you get the correct total molecular weight. That's exactly the method this calculator uses.

How to Use the Sequence → MW Calculator

Paste your amino acid sequence into the text box using the 20 standard 1-letter codes (for example, MKTAYIAK...). FASTA-format headers — lines starting with a > symbol — are stripped out automatically, along with any spaces, line numbers, or stray formatting, so you can paste directly from most sequence databases or synthesis reports without cleaning it up first.

The calculator reports two numbers side by side: the average molecular weight and the monoisotopic mass. It also breaks down exactly which amino acids appear and how often, in a simple composition chart, so you can spot-check the sequence at a glance.

Average Mass vs Monoisotopic Mass — What's the Difference?

The average molecular weight uses each element's naturally occurring average atomic weight, blending together all of its stable isotopes in the ratio found in nature. This is the number you use for everyday lab work — weighing out a bulk sample on a balance, sizing a band on an SDS-PAGE gel, or reporting a protein's size in a paper or datasheet.

The monoisotopic mass, on the other hand, uses only the mass of the single most common isotope of each element (carbon-12, nitrogen-14, and so on). This is the number mass spectrometry instruments actually measure, since a mass spec resolves individual isotope peaks rather than an averaged blend. For small peptides especially, always match monoisotopic mass against monoisotopic mass when checking a mass spec result — mixing the two up is one of the most common sources of confusion in peptide analysis.

The UV Extinction Coefficient Method (Advanced)

One of the fastest, cheapest ways to estimate protein concentration is to shine 280 nm ultraviolet light through a sample and measure how much of it gets absorbed. Only three amino acid features absorb meaningfully at that wavelength: tryptophan (Trp/W), tyrosine (Tyr/Y), and cystine — the disulfide bond formed when two cysteine (Cys/C) residues pair up.

This calculator uses the widely cited method from Pace, Vajdos, Fee, Grimsley, and Gray (1995, Protein Science) — the same approach behind tools like ExPASy's ProtParam — to estimate the molar extinction coefficient, ε280, directly from those residue counts:

ε280 (M⁻¹cm⁻¹) = (number of Trp x 5500) + (number of Tyr x 1490) + (number of cystine pairs x 125)

Once ε280 is known, the Beer-Lambert law lets you go straight from a measured absorbance reading to a protein concentration, without needing a dye-based assay, a standard curve, or a reference protein.

From A280 to Concentration

The Beer-Lambert law states that absorbance equals the extinction coefficient times concentration times path length: A = ε x c x l. Rearranged for a protein sample measured in a standard 1 cm cuvette, concentration in mg/mL works out to:

Concentration (mg/mL) = (A280 x Molecular weight) / (ε280 x path length in cm)

This calculator's Extinction & A280 tab pulls the Trp, Tyr, and Cys counts straight from whatever sequence you entered on the Sequence → MW tab (or you can enter counts manually), applies the Pace formula, then converts your measured absorbance into both a mass concentration (mg/mL) and a molar concentration (µM) automatically.

A Note on Cystine and Disulfide Bonds

Free cysteine residues (with an unpaired thiol group) don't meaningfully absorb at 280 nm — only cystine, the disulfide-bonded pair of two cysteines, does. Because it's often hard to know in advance exactly how many disulfides have actually formed in a given sample, this calculator offers a toggle: assume all cysteines are paired (a common, simple estimate that divides the Cys count by two), or turn it off to exclude cystine entirely from the calculation if you know the protein is fully reduced or has no disulfide bonds at all.

For a protein whose disulfide-bond count is known precisely — from a crystal structure or biochemical characterization — the most accurate approach is to treat that known number of cystine pairs as the input, which you can do here using the manual entry option.

Mass ⇄ Molar Concentration Conversion

Many downstream calculations — enzyme kinetics, receptor binding assays, dosing regimens — need concentration in molar units (µM, nM) rather than mass units (mg/mL, µg/mL), because it's the number of molecules present, not their total weight, that determines how a biological reaction behaves. This calculator's Mass ⇄ Molarity tab handles that conversion directly: enter a mass and a volume, along with the molecular weight (pulled automatically from the sequence tab, or entered manually), and it returns the mass concentration, the molar concentration, and the total molar amount present.

Concentration (mg/mL) = mass (mg) / volume (mL); Molar concentration (µM) = [mass concentration (mg/mL) / molecular weight (g/mol)] x 10⁶. Because mg/mL is numerically identical to g/L, dividing by the molar mass in g/mol converts directly to mol/L, and multiplying by a million converts to micromolar.

Reconstituting a Lyophilized Peptide or Protein

A common real-world question is the reverse problem: given a freeze-dried (lyophilized) vial of peptide or protein, exactly how much buffer or water should you add to reach a specific working concentration? Switch the Mass ⇄ Molarity tab to 'Target Molarity → Mass' mode, enter the concentration you want and the final volume you're dissolving into, and the calculator works out exactly how much mass — in both milligrams and micrograms — that target represents.

This is especially useful when a vendor's certificate of analysis reports only the total mass or the molecular weight of a custom peptide, and you need to know precisely how much diluent to add to hit a round-number stock concentration like 1 mM or 100 µM.

Where These Numbers Come From

Residue masses used here follow the standard average and monoisotopic values used across bioinformatics tools such as ExPASy's Compute pI/Mw and ProtParam. These are the same reference values taught in biochemistry courses and used in peptide synthesis quality control, so results here should match what you'd get from those tools for the same sequence.

Common Uses for These Calculations

These three calculations show up constantly across molecular biology, biochemistry, and pharmaceutical work:

• Checking a custom peptide synthesis or recombinant protein expression result against its expected molecular weight
• Reading and interpreting SDS-PAGE gel band sizes against a known protein's calculated MW
• Matching an intact mass spectrometry peak against the correct monoisotopic mass
• Estimating protein concentration from a quick NanoDrop or spectrophotometer A280 reading, without running a full BCA or Bradford assay
• Converting a stock's mg/mL concentration (often what's printed on a vendor's label) into the molarity needed for an enzyme assay, binding study, or cell treatment protocol
• Working out exactly how much lyophilized material to weigh out, or how much buffer to add, to hit a specific target concentration

Common Mistakes to Avoid

The most common mistake is forgetting to add back the one extra water molecule for the free N- and C-termini — using just the raw sum of residue masses understates the true molecular weight by about 18 Da, which barely matters for a large protein but can meaningfully shift the calculated mass of a short peptide.

A second common mistake is comparing an average mass directly against a mass spec result, or a monoisotopic mass against a gel migration estimate — always match average-to-average and monoisotopic-to-monoisotopic. A third is assuming every cysteine forms a disulfide bond by default; for proteins expressed and purified under reducing conditions, that assumption can noticeably overestimate the extinction coefficient and therefore underestimate the true concentration.

Protein Mass & Peptide Molecular Weight: Quick Reference Summary

Average/monoisotopic MW = sum of residue masses + one water (18.02 Da average, 18.011 Da monoisotopic). Extinction coefficient: ε280 = (Trp x 5500) + (Tyr x 1490) + (cystine pairs x 125). Concentration from A280: mg/mL = (A280 x MW) / (ε280 x path length). Molar conversion: µM = (mg/mL / MW) x 10⁶.

This free calculator is built to support coursework, lab planning, and everyday biochemistry calculations. For results tied to a regulated process, a certified assay, or a publication, always confirm against your lab's validated method and reference standards.