What the sequence tells you about how a peptide will fail

What makes the chemistry worth a reader’s attention is that it is predictable from the sequence. Given a primary structure you can say, without any experiment, which pathways are available and roughly in what order to expect them. An asparagine followed by a glycine is the fastest deamidation motif known. A surface methionine is an oxidation target. A free cysteine is a disulphide problem waiting for oxygen. This is why the same storage conditions produce quite different outcomes for two peptides of similar size, and why generic storage advice is worth so little.

Deamidation, and the isomer with the same mass

Deamidation of asparagine proceeds through nucleophilic attack by the backbone nitrogen of the following residue on the asparagine side-chain carbonyl, forming a five-membered succinimide intermediate which then hydrolyses to a mixture of aspartate and isoaspartate, conventionally in a ratio favouring the isomer roughly three to one. Glutamine deamidates by an analogous route, far more slowly, through a six-membered intermediate.

Three factors govern the rate. Sequence is dominant: the residue immediately following the asparagine determines how readily the intermediate forms, and asparagine-glycine is the fastest motif known, with serine, histidine and alanine following. Solution pH matters, with the rate minimal in the mildly acidic region and rising steeply above neutrality as the backbone nitrogen becomes more nucleophilic. Temperature and water activity set the overall pace, which is why the solid state helps so much.

The analytical problem is that isoaspartate has the same elemental composition and therefore the same molecular mass as the parent. Identity confirmation by molecular ion alone cannot distinguish them, and a preparation that is substantially deamidated will present as the intended compound. The isomers usually separate on a sufficiently shallow reversed-phase gradient, and specific methods exist, but only a method designed for the question will find the answer.¹

Oxidation, and where the oxidant came from

Methionine oxidises to the sulfoxide and, under harsher conditions, the sulfone. Tryptophan oxidises through a series of products including kynurenine derivatives. Histidine and tyrosine are susceptible under metal-catalysed conditions, and free cysteine oxidises readily to disulphide. Each of these products differs from the parent by a defined mass increment, which makes oxidation the pathway most reliably detected by mass spectrometry: the sulfoxide is sixteen mass units heavier and unmistakable.

The interesting question is usually where the oxidant came from, and the answers are mundane. Trace transition metals leached from glass, stainless steel or a stopper catalyse oxidation of several residues. Peroxides accumulate in polysorbate surfactants during storage and are a well-documented source of methionine oxidation in formulated products. Dissolved oxygen in the diluent contributes. Light drives it, particularly for tryptophan, and light exposure during handling is entirely undocumented in this trade.

Practical consequences follow that are not obvious. A formulation containing a surfactant that has itself been stored warm for a year may oxidise a peptide that would have been perfectly stable in a plain aqueous vehicle. Headspace composition matters: vials backfilled with nitrogen behave differently from vials sealed under air, and the difference is a manufacturing choice recorded nowhere on the label.²

Hydrolysis, isomerisation and the slower routes

Backbone hydrolysis cleaves an amide bond outright and produces two fragments, each of which is a distinct chromatographic species and each of which is detectable by mass. It is generally slower than deamidation at ordinary storage conditions but becomes dominant at low pH and elevated temperature, which is one reason accelerated stability data for peptides extrapolates so poorly: the pathway that dominates at forty degrees may be irrelevant at five.

Certain positions are much more labile than others. Aspartate-proline and aspartate-glycine bonds hydrolyse relatively readily under acidic conditions. N-terminal glutamine can cyclise to pyroglutamate, losing ammonia. Peptides with an N-terminal sequence of the right geometry can form a diketopiperazine and shed the first two residues as a cyclic dipeptide, a route that is fast enough at neutral pH to matter for some sequences.

Racemisation at susceptible residues produces epimers that are chemically identical in composition and differ only in stereochemistry. They are among the hardest impurities to detect, requiring either a chiral method or a sufficiently discriminating reversed-phase separation, and they are essentially never reported. A vial can be nominally pure by every measurement on its certificate and contain a percentage of a diastereomer with unknown biological behaviour.

What makes a method stability-indicating

A stability study is only as good as the analytical method behind it, and the requirement has a name: the method must be stability-indicating, meaning it must resolve the parent compound from its degradation products and quantify the change. Establishing that is done by forced degradation — deliberately stressing the material with acid, base, oxidant, heat and light — and demonstrating that the resulting products are separated from the parent and from each other with adequate peak purity.

Almost nothing sold as a purity determination in this market has been validated that way. A generic peptide gradient run for twelve minutes may perfectly well resolve the parent from its two largest process impurities and entirely fail to resolve it from its isoaspartate isomer or a closely related oxidation product. The number it returns is a purity figure, not a stability measurement, and using a series of such figures to argue that a product has not degraded is a category error.

The compendial guidance on analytical validation is explicit about specificity, and about demonstrating it against the degradation products the molecule can actually form. The gap between that expectation and practice in this trade is not a matter of dishonesty. It is that the method being sold was designed for a different purpose and is being asked a question it was not built to answer.³

Light, and the exposure nobody records

Photostability has its own guideline, its own defined light source options and its own exposure requirement expressed in lux hours of visible light and watt hours per square metre of near ultraviolet. Products are tested in the immediate container, and where they fail, in the marketing pack, and where they fail again the label carries a protection instruction. The chemistry is real: tryptophan and tyrosine absorb in the near ultraviolet and photo-oxidise, and photolytic disulphide cleavage is well documented.

Nothing about light exposure is recorded anywhere in the research-peptide supply chain. Vials are frequently supplied in clear glass. Photographs for listings are taken under studio lighting. Parcels are opened on kitchen counters. A reconstituted vial may sit on a shelf under a window for weeks. The cumulative exposure is unknown and unknowable, and it is plausibly a larger contributor to degradation than the transit excursions that attract all the attention.

The Journal makes one narrow observation rather than a recommendation, because recommendations are not this publication’s business. Amber glass, or a secondary carton, costs a fraction of a cent per unit and removes an uncontrolled variable entirely. Several of the twenty companies we track already ship in amber vials; most do not, and the ones that do not have not, as far as we can establish, generated any data suggesting it does not matter.⁴

The customs leg remains the part of this story we cannot report properly, and it deserves saying every time the subject comes up: there is a segment of every cross-border journey during which nobody measures and nobody has authority to intervene. Any claim of end-to-end control across that segment is a claim about something unobserved.