Short answer: Edman degradation is a chemical method of peptide sequencing that removes one amino acid at a time from the N-terminal end of a chain, identifies it, and then repeats the cycle on the shortened peptide [1]. The peptide is treated with phenyl isothiocyanate (PITC) and then trifluoroacetic acid, and each released residue is converted to a phenylthiohydantoin (PTH) derivative that is identified by its chromatographic elution time [1]. Automated sequencers can run as many as 50 cycles and can work from as little as 1 to 5 picomoles of sample [1].
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What Edman degradation is
Sequencing a peptide means finding the order in which its amino acids are linked [1]. Much of that work is now done by mass spectrometry, but Edman degradation remains a chemical method in common use [1].
The idea is simple: cleave one amino acid at a time from one end of the chain, separate and identify it, and then repeat the cleavage on the chain-shortened peptide until the whole sequence is known [1]. It works from the N-terminus, where PITC attaches to the –NH2 group of the first amino acid [1]. (Our N-terminus vs C-terminus guide explains how the two ends are defined.)
The chemistry is named after Per Edman, who introduced it in 1949 [2].
The Edman degradation mechanism, step by step
Each cycle has three chemical stages [1][2]:
- Coupling: phenyl isothiocyanate (PITC), C6H5–N=C=S, attaches to the –NH2 group of the N-terminal amino acid [1]. This reaction runs under mildly alkaline conditions [2].
- Cleavage: treatment with trifluoroacetic acid splits the N-terminal residue from the chain. This gives an anilinothiazolinone (ATZ) derivative plus the chain-shortened peptide [1]. (See our trifluoroacetic acid guide for more on TFA in peptide work.)
- Conversion: further acid-catalyzed rearrangement in aqueous acid converts the ATZ derivative into a phenylthiohydantoin (PTH) [1].
How the PTH amino acid is identified
Early work identified the released amino acids by paper chromatography, which was laborious [2]. Later gains in sensitivity came mainly from improving the on-line analysis of PTH amino acids by reverse-phase HPLC [4]. With microbore columns of 0.8 to 1.0 mm, standards can be run at the 0.5 to 1.0 pmol level and samples sequenced in the 1 to 5 pmol range [4]. (For background on reverse-phase separations, see our what is HPLC peptide testing guide.)
Some residues need extra handling. One example is cysteine: alkylating cysteine with acrylamide gives a derivative, Cys-S-β-propionamide, that is stable during automated Edman degradation, and its PTH form separates easily from the other PTH amino acids by HPLC [6].
From manual chemistry to automated sequencers
- The spinning cup sequenator (1967): Edman and Begg built an instrument they called the protein sequenator, which automated many steps of the process [2]. A spinning cup spread the protein along its surface and delivered a thin film of each reagent, which helped prevent the protein from washing out [2].
- The gas-liquid solid phase sequencer (1981): Hewick and colleagues described a miniaturized sequenator that used gas-phase reagents at the coupling and cleavage steps [3]. The sample sat in a Polybrene matrix dried onto a glass fiber disc [3]. It could give useful sequence data from as little as 5 pmol of protein and run extended sequences of more than 30 residues on subnanomole amounts [3]. It also had a cycle time of only 50 minutes [3]. This design improved sensitivity by a factor of 100 over the most advanced spinning cup sequenator [2].
- Blotting onto membranes: later work allowed proteins to be electroblotted onto PVDF, a high-capacity polymer, for direct Edman sequencing on the membrane [2].
Limits of Edman degradation
The method has several known limits.
- Read length: complete sequencing of large proteins is impractical because unwanted by-products build up [1]. Automated instruments allow up to about 50 cycles before those by-products interfere [1].
- Blocked N-termini: the method needs a free N-terminal amino group. If a protein's N-terminus is blocked, it has to be digested and the resulting peptides purified and sequenced individually [2]. That is the case for about half of all mammalian proteins, according to one historical review [2].
- Pyroglutamate: an N-terminal pyroglutamate is one kind of block. The enzyme pyroglutamate aminopeptidase can remove it, though this usually needs chaotropic and reducing agents and often has limited success [7]. One study removed it from antibody heavy chains with a thermostable enzyme, heat and polysorbate 20 before Edman sequencing, with deblocking that varied by sample (typically more than 50%) [7].
- Proline: proline residues often start overlaps during sequencer runs. The cause is an abnormally slow cleavage reaction, and its rate depends on where proline sits in the sequence [5].
- Sensitivity ceiling: alternative Edman reagents, including fluorescent chemistries, promised low-femtomole sensitivity, but none progressed into routine use [4].
Sequencing longer chains: fragment and overlap
To get around the read-length limit, a long chain is first cut into smaller fragments by partial hydrolysis [1]. Each fragment is sequenced, and the fragments are fitted together by matching their overlapping ends [1]. Protein chains with more than 400 amino acids have been sequenced this way [1].
- Acid hydrolysis is unselective and gives a more or less random mix of small fragments [1].
- Trypsin cuts only on the carboxyl side of arginine and lysine [1].
- Chymotrypsin cuts only on the carboxyl side of phenylalanine, tyrosine and tryptophan [1].
Edman degradation vs mass spectrometry
Mass spectrometry entered protein sequencing in 1959, when Klaus Biemann used it to analyze di- and tripeptides [2]. By the late 1970s, MS methods still needed much more starting material than Edman sequencing [2]. As tandem mass spectrometry improved, Edman methods struggled to push sensitivity past the gains of the gas-liquid instrument [2]. Peptide sequencing is now the key technology inside mass-spectrometry-based proteomics [8].
In practice the two approaches answer slightly different questions:
- Edman degradation reads residues one by one from a free N-terminus and identifies each by its PTH derivative [1].
- Mass spectrometry with ESI or MALDI ionization and a time-of-flight analyzer now handles much peptide sequencing [1]. Mass-spectrometry-based proteomics can also be used to determine post-translational modifications [8].
FAQ
What reagent is used in Edman degradation?
Phenyl isothiocyanate (PITC), followed by trifluoroacetic acid for the cleavage step [1].
What is the product of each Edman cycle?
An anilinothiazolinone (ATZ) derivative that is converted to a phenylthiohydantoin (PTH) amino acid, plus the peptide shortened by one residue [1].
Which end of the peptide does Edman degradation sequence?
The N-terminus. PITC attaches to the –NH2 group of the N-terminal amino acid [1].
How many residues can Edman degradation sequence?
Automated sequencers allow as many as 50 cycles before by-products interfere [1].
Why does Edman degradation fail on some proteins?
It needs a free N-terminal amino group. If the N-terminus is blocked, for example by pyroglutamate, the block has to be removed or the protein digested into peptides that are sequenced individually [2][7].
Bottom line
Edman degradation sequences a peptide from its N-terminus one residue per cycle: PITC coupling, TFA cleavage, and conversion to a PTH amino acid identified by chromatography [1]. Automated sequencers run up to about 50 cycles on 1 to 5 picomoles [1], and the 1981 gas-phase design pushed sensitivity to about 5 pmol [3]. Blocked N-termini, by-product buildup and slow proline cleavage are its main limits [1][2][5], which is why long chains are sequenced as overlapping fragments [1] and much sequencing has moved to mass spectrometry [1][2].
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