26.5 Amino Acid Analysis of Peptides

Amino Acid Analysis of Peptides

Amino acid analysis tells you what amino acids are in a peptide and how many of each are present. This is one of the first steps in characterizing an unknown peptide, and it lays the groundwork for full sequence determination later on.

The overall workflow has three stages: prepare the peptide (break it into free amino acids), separate those amino acids by chromatography, and then quantify the results.

Peptide Preparation for Analysis

Before you can analyze individual amino acids, you need to completely disassemble the peptide. That means dealing with two types of bonds: disulfide bonds between cysteine residues and the peptide (amide) bonds linking each amino acid.

Disulfide bond reduction

Cysteine residues can form disulfide bonds ($\text{–S–S–}$) that stabilize the peptide's three-dimensional structure. If you don't break these first, the peptide won't fully unfold, and hydrolysis will be incomplete.

1. Treat the peptide with a reducing agent such as dithiothreitol (DTT) or 2-mercaptoethanol to convert disulfide bonds into free thiol groups ($\text{–SH}$).
2. Alkylate the free thiols with an agent like iodoacetamide to cap them and prevent the disulfide bonds from re-forming.

Amide bond hydrolysis

Once the peptide is fully reduced and alkylated, you break every peptide bond to release free amino acids.

1. Heat the peptide in 6 M HCl at 110–165 °C for 12–72 hours (exact time depends on the peptide).
2. Complete hydrolysis yields the individual amino acids ready for separation.

One important caveat: acid hydrolysis isn't perfectly clean for every residue. Tryptophan is largely destroyed under these conditions. Serine, threonine, and tyrosine undergo partial degradation, so their measured amounts tend to be lower than the true values. Asparagine and glutamine are converted to aspartate and glutamate, respectively, so you can't distinguish the amide forms from the acid forms using this method alone.

Chromatography in Amino Acid Separation

After hydrolysis, you have a mixture of free amino acids that needs to be separated. Two main chromatographic approaches are used.

Ion-exchange chromatography

This method separates amino acids based on their net charge at a given pH.

• The stationary phase is a cation-exchange resin (typically sulfonated polystyrene), which binds positively charged species.
• Amino acids are eluted by gradually increasing the ionic strength or pH of the buffer. As the pH rises, amino acids lose their positive charge and release from the resin, each at a characteristic point.

Reverse-phase HPLC (RP-HPLC)

This separates amino acids based on hydrophobicity rather than charge.

• Because most amino acids don't absorb UV light strongly on their own, they're first derivatized with a chromophore or fluorophore (such as OPA or FMOC) to make them detectable.
• Elution uses a gradient of increasing organic solvent (commonly acetonitrile) in the mobile phase. More hydrophobic amino acids elute later.

Post-column derivatization

In classical ion-exchange analyzers, amino acids are separated first and then reacted with a derivatizing agent like ninhydrin as they come off the column. Ninhydrin produces a purple color (or yellow for proline) that can be measured by absorbance, allowing quantification.

Interpretation of Analyzer Results

Reading the chromatogram

• Each peak corresponds to a single amino acid.
• You identify each peak by comparing its retention time to that of known amino acid standards run under the same conditions.
• The area under each peak (or peak height) is proportional to the amount of that amino acid in the sample.

Quantification methods

• External standard method: You build a calibration curve by running known concentrations of amino acid standards. Then you read your unknown peak areas off that curve.
• Internal standard method: You add a known amount of a non-naturally occurring amino acid (commonly norleucine) to the sample before hydrolysis. This internal standard corrects for any losses during sample preparation, since it goes through the same process as your analytes.

Determining molar ratios

This is how you figure out the amino acid composition of the peptide:

1. Quantify the absolute amount (in moles) of each amino acid from the chromatogram.
2. Divide each value by the amount of the least abundant amino acid.
3. Round each ratio to the nearest whole number.

For example, if hydrolysis of a peptide gives you Gly : Ala : Leu in a ratio of 2.05 : 1.00 : 2.98, you'd round to 2 Gly : 1 Ala : 3 Leu. This tells you the composition but not the sequence.

Advanced Peptide Analysis Techniques

Amino acid analysis reveals composition, but to determine the actual order of residues, you need additional methods.

• Mass spectrometry measures the mass-to-charge ratio ($m/z$) of peptide fragments. Tandem MS (MS/MS) can fragment a peptide in predictable ways, generating data that reveal the sequence.
• Edman degradation sequentially removes and identifies the N-terminal amino acid one residue at a time. The peptide is treated with phenyl isothiocyanate (PITC), which reacts with the free $\alpha$-amino group. The labeled N-terminal residue is then cleaved as a PTH-amino acid and identified, and the cycle repeats. This method works well for peptides up to about 50 residues.
• Proteolytic enzymes (such as trypsin, chymotrypsin, or pepsin) cleave peptides at specific sites. Trypsin, for instance, cuts on the C-terminal side of Lys and Arg. By digesting a large peptide into smaller, overlapping fragments, you can sequence each fragment individually and then piece together the full sequence.