Disulfide Bonds and Free Thiols in Peptides: A Laboratory Guide

Cysteine, thiols and the disulfide bond

The side chain of cysteine ends in a sulfhydryl group, –SH, also called a thiol. Two thiols can be oxidized to a single sulfur-sulfur bond, the disulfide bond, with each cysteine losing one hydrogen atom. A disulfide can join two cysteines in the same chain (intramolecular) or in two separate chains (intermolecular). Reviews of thiol chemistry describe disulfides as structural cross-links in peptides and proteins and as the main product of thiol oxidation.^[1]

Several familiar research compounds show the range. Oxytocin is a nine-residue amidated peptide whose two cysteines, at positions 1 and 6, are closed into a ring by one disulfide. Glutathione, the tripeptide γ-glutamyl-cysteinyl-glycine, is the reverse case: it is normally described in its reduced form with a free thiol, and two molecules oxidize to the disulfide-linked dimer GSSG.^[2] IGF-1 LR3, a longer analogue built on the IGF-1 sequence, carries three disulfides. Knowing which form a sequence is supposed to have is the first step in checking it; the sequence notation guide shows how a disulfide is written.

Why pH controls thiol chemistry

A thiol is a weak acid. The neutral –SH form reacts slowly, while the deprotonated thiolate, –S⁻, is a strong nucleophile and is the species that takes part in most oxidation and thiol-disulfide exchange reactions. For free cysteine the side-chain pKa is a little above 8, and in a peptide it shifts with the neighbouring residues. Raising the pH toward and past that value increases the share of thiolate and speeds up both oxidation and exchange; the kinetic review cited above discusses pH, pKa and the other parameters that set these rates in detail.^[1]

Thiol-disulfide exchange matters as much as oxidation. A free thiolate can attack an existing disulfide, swap partners and leave a new free thiol behind. In a peptide with several cysteines this can move disulfides to non-native positions, often called scrambling. Mass spectrometry work on disulfide mapping relies on the opposite effect: digestion at acidic pH is used because disulfide rearrangement is precluded under those conditions.^[3] The practical rule follows: acidic conditions tend to preserve whatever disulfide pattern is present, and neutral to alkaline conditions with a free thiol present allow it to change.

Counting free thiols with Ellman's assay

The standard colorimetric method for free thiols was described by Ellman in 1959.^[4] The reagent, 5,5′-dithiobis(2-nitrobenzoic acid) or DTNB, is itself a disulfide. A free thiol cleaves it and releases one equivalent of 2-nitro-5-thiobenzoate (TNB), an anion with a strong yellow colour that is read at 412 nm. A reexamination of the reagent by Riddles and colleagues refined the extinction coefficient and the conditions for the assay,^[5] and the value of 14,150 M⁻¹ cm⁻¹ at 412 nm for the TNB anion is the one most often used today.^[6]

The arithmetic is the Beer-Lambert law. With made-up numbers: a blank-corrected absorbance of 0.283 in a 1 cm cuvette gives 0.283 ÷ 14,150 = 2.0 × 10⁻⁵ M, or 20 µM thiol in the cuvette. If the sample was diluted tenfold into the assay, the original solution held 200 µM thiol. For a peptide with one cysteine that is 200 µM peptide in the reduced form; for glutathione it can be converted to a mass concentration with the molarity calculator using 307.32 g/mol. A result well below the expected value suggests that some of the thiol has already been oxidized.

• Blank. Run reagent in buffer without sample, and sample without reagent if it is coloured, and subtract both.
• pH. The TNB colour depends on its anionic form, so the assay is run in a buffered, slightly alkaline solution and the buffer is recorded.
• Reducing agents. Any reducing agent left in the sample reacts with DTNB and reads as thiol; TCEP reduces it rapidly and in a fixed ratio, which is why DTNB is used to measure TCEP itself.^[6]

Reducing agents: DTT and TCEP

When a disulfide has to be opened, for example to confirm it by mass or to keep a thiol peptide reduced, a reducing agent is added. Dithiothreitol (DTT) was introduced by Cleland in 1964 as a protective reagent for SH groups.^[7] It works because, after reducing a disulfide, it closes into a stable six-membered ring, which drives the reaction to completion. Like any thiol it works through the thiolate, so it is most active above about pH 7 and is also oxidized by air more readily as the pH rises.^[6]

Tris(2-carboxyethyl)phosphine (TCEP) is a phosphine, not a thiol. It is odourless and remains stable in both acidic and basic solutions. In one comparison it reduced the disulfide DTNB faster than DTT below pH 8, and it still reduced another disulfide at pH 1.5, where DTT was inactive.^[6] A direct comparison found that the two reagents preserved the activity of a thiol-sensitive enzyme equally well, that DTT had to be removed before maleimide labelling while TCEP could stay, that nickel ions carried over from affinity columns oxidized DTT rapidly without affecting TCEP, and that TCEP was the more stable of the two in storage unless metal chelators were present, in which case DTT was more stable. The authors concluded that the choice is application specific.^[8]

Confirming a disulfide by mass spectrometry

Each disulfide removes two hydrogen atoms, so it lowers the monoisotopic mass by 2.016 Da compared with the fully reduced molecule. For a small peptide the simplest check is to measure the intact mass, reduce an aliquot, and measure again: one intramolecular disulfide gives a shift of about +2.016 Da, two give about +4.03 Da. A covalent dimer, such as GSSG or two copies of a single-cysteine peptide joined by an intermolecular disulfide, appears near twice the monomer mass minus 2.016 Da and falls apart into monomers on reduction.

For molecules with several disulfides, the question is which cysteines are paired. A review of disulfide determination by mass spectrometry describes the usual approach: digest the molecule with its disulfides intact, compare the masses of non-reduced and reduced aliquots of the digest, and pair the cysteine-containing fragments, using tandem MS where a fragment carries more than one disulfide.^[3] Small mass differences of this kind need adequate resolution, which the mass spectrometry guide covers, and the expected mass of any sequence can be worked out with the peptide mass calculator.

Handling thiol-containing peptides

Most of the practical steps follow from the chemistry above: keep oxygen, metal ions and alkaline pH away from free thiols, and avoid free thiols when an existing disulfide pattern needs to be kept.

1. Record whether each cysteine in the sequence is expected to be a free thiol or part of a disulfide, and which cysteines are paired.
2. Use degassed buffers and keep headspace small for solutions of free-thiol peptides; air oxidation is the usual route to unintended disulfides and dimers.
3. A metal chelator such as EDTA in the buffer slows metal-catalysed oxidation; note it, because it also changes how DTT and TCEP behave in storage.^[8]
4. Slightly acidic solutions slow both oxidation and thiol-disulfide exchange, so they help keep whichever form is present.
5. Prepare single-use aliquots so a stock is not repeatedly exposed to air; the aliquoting guide covers the layout.
6. Remove or account for any reducing agent before a thiol assay or a labelling step that it would interfere with.

Oxidation of cysteine is one of several degradation routes; methionine and tryptophan oxidation, deamidation and hydrolysis are covered in the storage and degradation guide. This guide describes laboratory chemistry only and is not a statement about any particular material. Terms used here are defined in the glossary, and questions can be sent to support.