Peptide Sequencing Explained: How the Order of Amino Acids Is Determined

Published October 2026 · Patriot Labs Research Team · ~16 min read

The short answer

A peptide’s sequence — the order of its amino acids — is determined by two main methods. Edman degradation removes and identifies one residue at a time from the N-terminus; tandem mass spectrometry breaks the peptide into fragments and reads the sequence from their masses. Both reveal the order of the residues, not just which ones are present.^[1][7][11]

The two methods come at the same question from opposite directions. Edman chemistry is a stepwise chemical reaction run on an intact chain: it labels the end amino acid, snips it off, identifies it, and repeats, so it reads the sequence directly and in order from one terminus.^[1][3] Mass spectrometry is a measurement, not a reaction on the whole chain: it weighs fragments of the peptide and infers the order from the pattern of masses.^[7][11] Edman degradation ruled sequencing for four decades; tandem mass spectrometry is the dominant method today because it is faster, more sensitive, works on mixtures, and does not need a free chain end.^[11][12]

This guide takes the subject in order: what sequencing actually measures and why it is distinct from purity or mass, the history that produced these tools, the chemistry of each method, how a mass spectrum is turned into letters, the residues that remain hard to tell apart, and how sequence fits into the wider picture of research-peptide identity.

What “sequencing” actually measures

Sequencing answers one specific question: in what order are the amino acids joined along the chain? That is a different question from the ones the other analytical methods answer, and the distinction matters.

A peptide is a chain of amino acids linked by peptide bonds. Four separate properties can be measured about such a chain, and each has its own method. Composition — how much of each kind of amino acid is present — is measured by amino acid analysis, which hydrolyzes the chain into free amino acids and counts them, but destroys the order in the process. Mass — the molecular weight of the intact molecule — is measured by mass spectrometry and confirms identity against an intended formula. Purity — what fraction of the sample is the target rather than related substances — is measured by HPLC as a percentage of peak area. Sequence is the fourth property: the actual order of residues.

Composition and mass can both be correct while the sequence is wrong. Two peptides that are anagrams of each other — the same residues in a different order — have identical amino acid composition and identical total mass, yet are different molecules with different behavior. Only sequencing distinguishes them. This is why sequencing sits at the top of the characterization hierarchy: it is the measurement that pins down the primary structure itself, rather than a summary statistic about it.

From Sanger’s insulin to the sequenator

The first protein ever sequenced was insulin, worked out by the English biochemist Frederick Sanger in the years around 1951–1955. That achievement proved that a protein has a single, defined amino acid sequence — not a vague or variable mixture — and it earned Sanger the 1958 Nobel Prize in Chemistry.^[6]

Sanger had no stepwise machine. His approach combined an end-group label with brute-force reassembly. “Sanger’s reagent,” fluorodinitrobenzene (FDNB), reacts with the free N-terminal amino group to form a yellow dinitrophenyl derivative, marking which residue sat at the chain’s end. He then broke insulin into short peptides with acid or with enzymes such as trypsin, identified the pieces, and fit the overlapping fragments back together like a jigsaw to deduce the whole.^[6] He dated the sequences of the two insulin chains to 1951 and 1952 and the assignment of the disulfide bridges between them to 1955.^[6] The work took the better part of a decade for a single small protein.

The method that made sequencing routine came from the Swedish biochemist Pehr Victor Edman (1916–1977). In 1950, working at the University of Lund, he published a reaction that removes just the N-terminal residue of a peptide and leaves the rest of the chain intact, so the process can be repeated cycle after cycle.^[1][5] Seventeen years later, in 1967, Edman and his assistant Geoffrey Begg described an automated instrument — the “sequenator” — that carried out the chemistry mechanically, residue after residue, without an operator running each step by hand.^[2][5] Automated Edman sequencing spread quickly; within a few years scores of the machines were in use worldwide.^[3]

The next shift was instrumental rather than chemical. In 1986, Donald Hunt and John Yates and their colleagues demonstrated that peptides could be sequenced by tandem mass spectrometry — fragmenting ionized peptides inside the instrument and reading the sequence from the fragment masses.^[7] As soft ionization methods and faster, more accurate analyzers matured, mass spectrometry displaced Edman chemistry as the default, especially for the large-scale, mixture-heavy work of modern proteomics.^[11][12]

Edman degradation: reading from the N-terminus

Edman degradation sequences a peptide chemically, one residue at a time, starting from the N-terminal end. Its defining feature is selectivity: in each cycle, only the single residue at the free amino end is labeled and removed, and every other peptide bond in the chain is left untouched.^[3][4] That is what lets the cycle repeat on the shortened peptide and walk down the sequence.

The three chemical steps of one cycle

Each Edman cycle is three reactions run in sequence: coupling, cleavage and conversion.^[3][4]

Coupling. Phenyl isothiocyanate (PITC, the Edman reagent) reacts with the uncharged N-terminal amino group under mildly alkaline conditions, forming a phenylthiocarbamoyl (PTC) derivative on the end residue.^[3][4] Only a free, neutral α-amino group reacts, which is why the reaction targets the chain terminus and not the side chains along the backbone.

Cleavage. Treatment with anhydrous acid — trifluoroacetic acid is the usual choice — cleaves the derivatized residue from the rest of the chain. It comes off as a cyclic anilinothiazolinone (ATZ) derivative, and crucially the acid conditions cut only the bond to the labeled end residue while sparing the remaining peptide bonds, so a chain-shortened peptide is left behind intact.^[3][4]

Conversion. The ATZ derivative is unstable, so a further acid-catalyzed rearrangement in aqueous acid converts it into the stable phenylthiohydantoin (PTH) form of that amino acid.^[3][4] The PTH–amino acid is the thing that is actually identified. The shortened peptide, meanwhile, now presents a new N-terminal residue and is ready for the next cycle.

The read-out, the cycle limit and where it fails

Each cycle yields one PTH–amino acid, and it is identified by comparing how it moves in chromatography against reference PTH derivatives of the twenty common amino acids.^[3] Repeating coupling–cleavage–conversion over and over produces the sequence one letter at a time, from the N-terminus inward. Modern instruments achieve very high efficiency per cycle — well over 99% — and need only picomole quantities of peptide.^[3][4]

The method has a practical ceiling. Because the derivatization never goes perfectly to completion, a little of the previous residue is carried over each cycle, and these errors accumulate. In practice a clean, unambiguous read runs to roughly 30 residues, with an upper bound in the 50–60 range on favorable samples before by-product build-up obscures the signal.^[3][4] A long protein is sequenced by cutting it into shorter peptides first and sequencing those.

Edman degradation also has a hard requirement that defines when it cannot be used at all: the peptide must have a free N-terminus. If the N-terminal amino group has been chemically blocked — by an N-terminal acetyl cap, by a pyroglutamate residue, or by another modification — PITC has nothing to couple to and the first cycle fails.^[3] The reaction also stalls if it reaches a residue it cannot process, such as a non-α-linked amino acid.^[3] These blind spots are a large part of why a mass-based method was eventually needed.

Tandem mass spectrometry: sequencing by mass

Tandem mass spectrometry sequences a peptide by weighing its pieces. Rather than react on an intact chain, it ionizes many copies of the peptide, isolates one species by mass, breaks the backbone at various points across that population, and measures the masses of the fragments that result.^[7][11] The sequence is read out of the pattern of those masses.

The workflow has three stages inside the instrument. First, the peptide is ionized — soft ionization methods such as electrospray ionization (ESI) and matrix-assisted laser desorption/ionization (MALDI) put intact peptide molecules into the gas phase as charged ions.^[11] Second, a single precursor ion of one mass-to-charge value is selected in the first stage of analysis (MS1). Third, that precursor is fragmented and the fragment masses are recorded in the second stage (MS2) — the “tandem” in tandem MS, two stages of mass analysis with a fragmentation step between them.^[11] A fragment is only detected if it carries at least one charge, so a single molecule contributes one fragment ion, but a population of identical peptide molecules breaking at different backbone positions produces a whole family of fragments that together cover the chain.^[10][11]

Fragment-ion nomenclature: a, b, c and x, y, z

Peptide fragments are named by which end of the chain they contain and which backbone bond broke. The scheme was proposed by Roepstorff and Fohlman in 1984 and later refined, and it is the common language of every peptide mass spectrum.^[8][9][10] Fragments that retain the N-terminal part of the chain are called a, b or c ions; fragments that retain the C-terminal part are called x, y or z ions.^[10][11]

The letter within each set records which of the three backbone bonds was cut. Reading across the repeating backbone, the cuts are: the a/x pair at the Cα–carbonyl bond, the b/y pair at the amide (peptide) bond itself, and the c/z pair at the amide nitrogen–Cα bond.^[11] Because the b/y pair comes from breaking the peptide bond — the weakest and most readily cleaved backbone bond under ordinary conditions — b and y ions are the most abundant in most spectra.^[10][11] The table below lays out the six ion types and their cleavage sites.

Reading the ladder of masses

The sequence is read from the mass differences between consecutive fragment ions in one series. Within a single series — say the b ions — each successive ion is larger than the one before it by exactly the mass of the one additional residue it contains.^[11] List the b ions in order of mass, take the difference between each neighboring pair, and each difference matches the residue mass of a specific amino acid. Those differences, read end to end, spell the sequence.

Having two series that count from opposite ends is what makes the read reliable. A b-ion ladder reads the chain from the N-terminus; a y-ion ladder reads the same chain from the C-terminus. The two are complementary — for a given peptide the masses of a matching b and y ion sum to the mass of the whole peptide plus a fixed constant — so one series confirms the other and fills gaps where a fragment is missing.^[11] When a sequence is reconstructed this way straight from the spectrum, without comparison to any database, the process is called de novo sequencing; when the spectrum is instead matched to an already-known sequence in a database, it is database searching.^[11][13] De novo is the method that applies to peptides no database contains — novel, engineered or synthetic sequences.^[11]

CID, HCD and ETD

How the backbone is broken determines which ion types appear, so the fragmentation method is chosen to suit the peptide. The three in common use are collision-induced dissociation (CID), higher-energy collisional dissociation (HCD) and electron-transfer dissociation (ETD).

CID accelerates the precursor ion into neutral gas molecules; the collisions deposit energy that breaks the most vulnerable backbone bond, the amide bond, so CID yields predominantly b and y ions.^[10][11] HCD is a higher-energy collisional variant used on modern trapping instruments; it likewise produces mainly b and y ions but with cleaner low-mass detail.^[12] ETD works by a different mechanism: an electron is transferred to the multiply-charged peptide, cleaving the N–Cα bond and producing c and z-type ions instead.^[11] ETD is especially valuable for larger peptides and for ones carrying fragile modifications, because it can break the backbone while leaving those modifications in place. Instruments often run complementary methods on the same peptide to cover gaps one method leaves.^[11]

The hard cases: Leu/Ile and Gln/Lys

Because mass spectrometry reads sequence from mass, residues that have the same or nearly the same mass are the method’s blind spots. Two pairs cause most of the trouble.

Leucine and isoleucine are isobaric: they are structural isomers with the identical chemical formula and therefore the identical residue mass, about 113.084 daltons.^[14] No ordinary mass measurement can separate them, however high the resolution, because there is no mass difference to measure. Telling them apart requires specialized fragmentation that probes the side chain (such as the w and d ions produced under high-energy conditions), or an orthogonal method.^[10] Glutamine and lysine are not identical but are very close: their residue masses are about 128.059 Da and 128.095 Da, a difference of only around 0.036 Da, which blurs together on low-resolution instruments and needs a high-resolution analyzer to resolve.^[14] The table summarizes the two problem pairs.

Edman degradation does not share these ambiguities, because it identifies each residue by its chemistry — the elution behavior of its PTH derivative — rather than by mass, and can distinguish both pairs directly.^[3] This complementarity is one reason the two methods have long been used alongside each other.

Edman degradation vs tandem MS

The two methods are complementary rather than redundant: each is strong exactly where the other is weak. Edman reads cleanly and unambiguously from the N-terminus but needs a free chain end and a purified single peptide; mass spectrometry is fast, sensitive, handles mixtures and blocked termini, but reads from mass and so stumbles on isobaric residues.^[3][11][12] The table sets the main properties side by side.

Sequencing and research-peptide identity

In practice, routine batch release of a research peptide does not re-sequence the whole chain every time; it confirms identity against a known target sequence. Full sequencing is a characterization method — used to establish or verify what a sequence is — while routine quality testing checks that a given batch matches that established identity.

The two everyday checks are mass and purity. Mass spectrometry confirms that a batch’s measured molecular mass matches the intended sequence’s calculated mass, which is a strong identity check because a wrong residue, a deletion or an unintended modification shifts the mass. HPLC measures purity as peak area. Together with net peptide content and counterion data, these are the fields that appear on a batch Certificate of Analysis. Patriot Labs tests every batch at an independent US laboratory by HPLC and mass spectrometry, and the COA is posted on each product page. Where a measured mass matches but a sequence question remains — for example, distinguishing a target from a closely related isomer or a deletion-sequence impurity — tandem MS/MS fragmentation is the tool that resolves it, because it reads the order and not just the total mass.

What this guide does not establish

This is an educational chemistry explainer about how peptide sequences are determined. It describes analytical methods in general terms and makes no claim about any biological activity, effect or use of any peptide.

Several boundaries are worth stating plainly. The descriptions of Edman chemistry and mass-spectrometric fragmentation are simplified summaries of well-established methods, not laboratory protocols, and specific reagents, conditions and instrument settings vary by application. The residue masses given are standard monoisotopic values for the common amino acids; real measurements carry instrument error and depend on calibration. “Sequence confirmation” on a routine Certificate of Analysis generally means an identity check by mass and purity rather than a full residue-by-residue sequencing run, and a matching mass does not by itself prove every residue is in the intended position. Finally, the fact that a sequence can be measured says nothing about what a molecule does; sequence is primary structure only.

Frequently asked questions

How is a peptide’s amino acid sequence determined?

A peptide’s sequence is determined by two main methods. Edman degradation chemically removes and identifies one residue at a time from the N-terminus, reading the chain in order. Tandem mass spectrometry breaks copies of the peptide into fragments and reconstructs the sequence from the mass differences between those fragments. Both reveal the order of the amino acids, not merely which ones are present.

What is Edman degradation?

Edman degradation is a chemical method, introduced by Pehr Edman in 1950, that sequences a peptide one residue at a time from the N-terminus. Phenyl isothiocyanate reacts with the free N-terminal amino group, acid then cleaves that single residue as a thiazolinone, which is converted to a stable phenylthiohydantoin (PTH) amino acid and identified by chromatography. The shortened peptide is carried into the next cycle.

How does tandem mass spectrometry sequence a peptide?

The peptide is ionized, a single precursor ion is selected by mass, and that ion is fragmented inside the instrument so that the backbone breaks at many points across a population of molecules. The masses of the resulting fragment ions are measured, and because consecutive fragments differ by the mass of exactly one amino acid residue, the ladder of masses spells out the sequence.

What is the difference between b ions and y ions?

Both arise from cleavage of the peptide (amide) bond in tandem mass spectrometry. A b ion keeps the charge on the N-terminal part of the peptide; a y ion keeps the charge on the C-terminal part. Because they count the chain from opposite ends, a b-ion series and a y-ion series provide two independent reads of the same sequence. Under collision-induced dissociation, b and y ions dominate the spectrum.

Can you sequence a peptide with a blocked N-terminus?

Not by classical Edman degradation, which needs a free, uncharged N-terminal amino group; an N-terminal acetyl cap or a pyroglutamate residue stops the reaction before it starts. Tandem mass spectrometry has no such requirement, because it reads the sequence from fragment masses rather than from the N-terminus, so it is the usual route for N-terminally modified peptides.

Why can’t sequencing always tell leucine from isoleucine?

Leucine and isoleucine are isobaric: they have the same chemical formula and therefore the same residue mass, about 113.084 daltons, so a method that reads sequence from mass cannot distinguish them without specialized fragmentation. Glutamine (about 128.059 Da) and lysine (about 128.095 Da) differ by only about 0.036 Da and need high-resolution instruments to separate. Edman degradation, which identifies residues chemically rather than by mass, can tell these pairs apart.

What is de novo peptide sequencing?

De novo peptide sequencing is reading a sequence directly from a tandem mass spectrum, using only the fragment masses, without matching against a database of known sequences. It is the method used for peptides that are not in any database, including synthetic research peptides and novel or engineered sequences. Database searching, by contrast, matches a spectrum to a sequence that is already known.

References

1. Edman, P., Högfeldt, E., Sillén, L. G., & Kinell, P. O. (1950). Method for determination of the amino acid sequence in peptides. Acta Chemica Scandinavica, 4, 283–293. doi:10.3891/acta.chem.scand.04-0283
2. Edman, P., & Begg, G. (1967). A protein sequenator. European Journal of Biochemistry, 1(1), 80–91. doi:10.1111/j.1432-1033.1967.tb00047.x
3. Edman degradation. Wikipedia. https://en.wikipedia.org/wiki/Edman_degradation
4. Peptide Sequencing: The Edman Degradation. Organic Chemistry (NC State University / Pressbooks). https://ncstate.pressbooks.pub/organicchem/chapter/peptide-sequencing-the-edman-degradation/
5. Pehr Victor Edman. Wikipedia. https://en.wikipedia.org/wiki/Pehr_Edman
6. Frederick Sanger Sequences the Amino Acids of Insulin, the First Protein Sequenced (historical record of the 1951–1955 work and the 1958 Nobel Prize in Chemistry). HistoryofInformation.com. https://historyofinformation.com/detail.php?id=729
7. Hunt, D. F., Yates, J. R., Shabanowitz, J., Winston, S., & Hauer, C. R. (1986). Protein sequencing by tandem mass spectrometry. Proceedings of the National Academy of Sciences, 83(17), 6233–6237. https://pmc.ncbi.nlm.nih.gov/articles/PMC386476
8. Roepstorff, P., & Fohlman, J. (1984). Proposal for a common nomenclature for sequence ions in mass spectra of peptides. Biomedical Mass Spectrometry, 11(11), 601. (As cited in the Mascot peptide-fragmentation reference, ref. 10.)
9. Johnson, R. S., Martin, S. A., Biemann, K., et al. (1987). Modification of the a/b/c and x/y/z fragment-ion nomenclature for collision-induced peptide decomposition. (As cited in the Mascot peptide-fragmentation reference, ref. 10.)
10. Peptide fragmentation — fragment ion types and nomenclature. Matrix Science, Mascot Help. https://www.matrixscience.com/help/fragmentation_help.html
11. De novo peptide sequencing. Wikipedia. https://en.wikipedia.org/wiki/De_novo_peptide_sequencing
12. Steen, H., & Mann, M. (2004). The ABC’s (and XYZ’s) of peptide sequencing. Nature Reviews Molecular Cell Biology, 5(9), 699–711. https://www.nature.com/articles/nrm1468
13. Eng, J. K., McCormack, A. L., & Yates, J. R. (1994). An approach to correlate tandem mass spectral data of peptides with amino acid sequences in a protein database (SEQUEST). Journal of the American Society for Mass Spectrometry, 5(11), 976–989.
14. Monoisotopic and average amino acid residue masses. UW Proteomics Resource. https://proteomicsresource.washington.edu/protocols06/masses.php
15. After 75 years, an alternative to Edman degradation: a mechanistic and efficiency study of a base-induced method for N-terminal peptide sequencing (2025). PMC. https://pmc.ncbi.nlm.nih.gov/articles/PMC12721331/

For the analytical methods this guide connects to, see What Is Mass Spectrometry?, What Is HPLC? and What Is Amino Acid Analysis?. For the structures being read, see What Is a Peptide Bond? and What Is an Amino Acid?.

All Patriot Labs products are sold strictly for in-vitro research and laboratory use only. Not for human or veterinary consumption. This guide is educational and describes the chemistry and analytical methods of peptide sequencing in general terms; it is not medical advice, does not describe how to use any product, and makes no claim about outcomes in humans.