What I am working with: 96.4% · Medutest · 95.2% · Janoshik.
Both of these get recommended confidently by different people, which suggests neither is obviously right.
My constraints are cost, measurement resolution and how much handling I am prepared to do — in roughly that order.
Is there a defensible reason to prefer one, or is this a coin flip?
96.4 and 95.2 are 1.2 percentage points apart, which sounds small until you restate it as impurity. Medutest is leaving 3.6 per cent of the detected area unassigned and Janoshik 4.8 per cent — a factor of 1.33 between them. The impurity fraction is the quantity that moves when a method changes, and it is the one worth arguing about; the headline is just its complement. Comparable means same column chemistry, same gradient slope, same detection wavelength, same integration convention. Until you have those four from both laboratories, 96.4 and 95.2 are two measurements of slightly different quantities that happen to share a unit.
To be exact about it, the single most important distinction is between what purity measures — the fraction of detected material that is your target — and what you actually want to know — how much of the material in the vial is your target.
Column pore size affects mass transfer — a 100 Angstrom packing on a 5 kDa peptide restricts diffusion, broadening the peak and potentially hiding small impurities in the shoulders.
| Δ mass (Da) | Most likely cause | Distinguishing feature |
|---|---|---|
| +1 | Deamidation (Asn or Gln) | New peak, slightly earlier retention |
| −17 | Loss of ammonia | Often with deamidation |
| −18 | Dehydration / succinimide | pH-dependent, reversible |
| +16 | Oxidation (Met, Trp) | Earlier retention, light-related |
| −128 | Missing Gln or Lys | Deletion sequence from synthesis |
| 0 | Isomer: racemisation or scrambling | Same mass, shifted retention |
Concretely, gradient slope controls resolution, and gentler slopes resolve co-eluting impurities into separate peaks — so the better method reports the worse purity number.
Published side-by-side method comparisons show that a two-point difference in purity on the same vial is easily explained by method choice alone.
One qualification: achieving purity above roughly 98 per cent on a 30-residue peptide is fighting the chemistry of synthesis, not the quality of the purification.
If you are ranking vendors, specify a method and have all samples tested at the same place.
Analytical standards and reagents with traceable certificates. Every quantitative result you read inherits the accuracy of the standard behind it.
Shop standardsThe underlying point is that understanding purity requires separating the chemistry from the method from the reporting convention, and the three are not independent.
The fraction of your main peak that is actually your target versus isomers, fragments or related sequences is invisible without complementary identity testing.
Mobile phase additive choice affects ionisation and peak shape — TFA gives sharp peaks but suppresses mass spectrometry signal, formic acid gives worse peaks but preserves signal.
Proline conformer interconversion kinetics are well-characterised and the half-life is of the same order as the chromatographic peak width at room temperature.
The practical summary: ask for the chromatogram and the method, and ignore the headline number until you have both.
To be exact about it, the method matters more than the vial, which is why specifying a method buys you far more than changing suppliers does.
Integration of the shoulder — whether you use perpendicular drop or tangent skim — determines what area gets assigned to the main peak versus the impurity table.
It helps to be literal here: retention time is sequence-specific and method-specific, so comparing your result to a supplier value using a different method is meaningless without method documentation.
The Arrhenius relationship for peptide degradation is the basis of accelerated stability testing and also governs how quickly methods drift with temperature.
Compare purity within a single laboratory on the same method, never across laboratories.
edited 4 Jun 2026 by n_takahashi — added a caveat about sampling
Mechanically, identity and purity are orthogonal, and a high purity says almost nothing about whether the peak is actually what you think it is.
Temperature affects the dynamics of molecular conformation, and if a peptide has proline residues that interconvert on the chromatographic timescale, the peak will split or shoulder at low temperature and collapse at high temperature.
Inter-laboratory studies on identical peptide material routinely find half-a-per-cent to a full-per-cent spreads in reported purity on the same sample.
The caveat is that purity without identity is only half an answer — a high purity does not mean the peak is actually what you think it is.
If you only pay for one test, pay for quantified content. Purity is the number everyone quotes and content is the number that changes what you do.
The honest answer is that the achievable range of plausible purity figures for a given vial is wider than most people expect.
Tailing factor measures peak shape, and a badly tailing peak spreads into the region where small impurities live, forcing tangent-skim integration that assigns tail area to the main peak.
The ICH Q3A impurity thresholds and the relevant pharmacopoeial chapters all specify method validation requirements that almost no research-grade certificate claims to meet.
In practice: ask for the chromatogram, check the method section, check the lot number against the vial, and set your accept threshold before you see the result rather than after.
Ask PeptideStack is a static archive. Posting is closed, but the norms are worth stating: answer the question that was asked, show your working, cite the trial or the certificate, and say plainly where the evidence runs out.