Protein Sequencing

5 min read Updated Apr 18, 2026

Once you have a pure protein, you can ask what amino acids are in it and in what order. Two main techniques, with mass spectrometry now dominating in the real world and Edman degradation classic enough to remain on the MCAT.

Edman Degradation - One Residue at a Time

Pehr Edman developed this method in 1950. A chemical reagent (phenyl isothiocyanate, PITC) reacts specifically with the N-terminal amino acid. A second step cleaves that amino acid off as a phenylthiohydantoin (PTH) derivative, which is then identified by HPLC (by comparing its retention time to known PTH standards). The remaining protein has a fresh N-terminus one residue shorter.

Repeat. Each cycle releases the next amino acid in the sequence. Cycle 1 gives residue 1, cycle 2 gives residue 2, and so on.

Limitations

Only works on the N-terminus. If the protein has a blocked N-terminus (common in eukaryotic proteins with N-terminal acetylation), Edman fails.
Becomes noisy beyond ~30-50 residues per run. To sequence a larger protein, you typically cleave it into fragments (with trypsin, CNBr, etc.), sequence each fragment, and then piece together the overlaps.
Each cycle takes ~30-60 minutes, so it is slow.

Edman degradation has largely been replaced by mass spectrometry in modern labs. It still appears on the MCAT because the principle - label-cleave-identify at the N-terminus - is an instructive chemistry example.

Mass Spectrometry

Mass spec measures mass-to-charge ratio (m/z). For proteins, the workflow is usually:

Digest the protein into peptides with a sequence-specific protease (often trypsin, which cuts after Arg or Lys).
Ionize the peptides and measure their m/z values.
Select one peptide at a time and fragment it inside the mass spec (tandem MS). The fragment pattern reveals the amino acid sequence.
A computer matches the observed masses to a database of all known protein sequences and identifies the protein.

MALDI-TOF and Electrospray

Two common ionization methods you should recognize:

MALDI (matrix-assisted laser desorption/ionization): peptides are embedded in a matrix that absorbs laser energy, releasing ions.
ESI (electrospray ionization): peptides are sprayed from a fine capillary under high voltage, generating droplets that evaporate to leave ions.

Both are then pushed through a mass analyzer. TOF (time-of-flight) analyzers separate ions by how long they take to cover a fixed distance - lighter ions move faster.

Why Mass Spec Wins

Mass spec can identify a protein from a tiny sample (femtomoles), can identify all proteins in a complex mixture (proteomics), and can detect post-translational modifications by the mass shift they add (phosphorylation = +80 Da, methylation = +14 Da, ubiquitination = +8564 Da). None of this is possible with Edman.

Total Amino Acid Composition

Sometimes you just want to know how many of each amino acid are in a protein, not the order. Acid hydrolysis (6 M HCl at 110°C for 24 hours) breaks every peptide bond. The resulting mixture of free amino acids can be quantified to give percent composition. Tryptophan is destroyed by acid hydrolysis and must be measured separately. This method gives composition but not sequence.

Quantifying Total Protein: The Bradford Assay

Before any sequencing or characterization experiment, you need to know how much protein you actually have. The Bradford assay is the MCAT-canonical method.

🔍 How Bradford Works

Reagent: Coomassie Brilliant Blue G-250 in acidic solution. In the free form, the dye is brown with absorbance maximum at 465 nm.

On binding to protein (especially to arginine and other basic side chains), the dye shifts to a stable blue form with absorbance maximum at 595 nm. The shift is caused by stabilization of the anionic form of the dye by the protein's positively charged residues.

Measurement: Read A₅₉₅ on a spectrophotometer. Absorbance is proportional to protein concentration (Beer-Lambert: A = εcℓ). Compare against a standard curve (usually built with bovine serum albumin, BSA) to get µg/mL.

Strengths: fast (~5 min), cheap, sensitive (1-20 µg range), compatible with most buffers.
Limitations: detergents (SDS, Triton) interfere; proteins with few basic residues (e.g., some IgGs) underestimate; readings are not strictly linear across a wide range.

Alternative methods the MCAT occasionally mentions: BCA assay (cheaper in bulk, uses Cu⁺ reduction), Lowry assay (older, similar principle to BCA), and UV absorbance at 280 nm (which reads tryptophan and tyrosine absorbance, no dye needed).

Which end of a peptide does Edman degradation sequence, and what reagent is used?

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The N-terminus. Phenyl isothiocyanate (PITC, Edman's reagent) reacts with the N-terminal amino acid's free amino group. A subsequent cleavage releases that residue as a phenylthiohydantoin (PTH) derivative, which is identified by HPLC. The process exposes a new N-terminus for the next cycle.

Why is trypsin commonly used to digest proteins before mass spectrometry?

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Trypsin is a serine protease that cleaves peptide bonds C-terminal to lysine or arginine (unless the next residue is proline). This produces peptides of a convenient size (~8-20 residues) with predictable C-terminal residues, which simplifies mass spec analysis and improves identification from sequence databases.

How can mass spectrometry detect post-translational modifications like phosphorylation?

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Each modification adds a specific mass. Phosphorylation adds 80 Da (the mass of HPO3). If the observed peptide mass is 80 Da larger than the predicted mass for the unmodified sequence, the peptide is likely phosphorylated. Tandem MS can further localize which residue is phosphorylated by examining the fragment ions.