HPLC, mass spec and the other tests behind a peptide's purity

Built one link at a time

Imagine typing a 30-character password where every keystroke is 99% reliable. The chance of getting the whole thing right is only about 74%. Peptide chemists face the same arithmetic every time they build a chain.

Before the 1960s, peptides were made in solution, and the product had to be separated from by-products after every single step. The Nobel committee later spelled out why that was hopeless for long chains: at a very good 90% yield per step, a 100-step synthesis ends with 0.003% of the product ^[2].

Bruce Merrifield, at Rockefeller University, changed the game by anchoring the first amino acid to a solid polymer bead ^[1]. Each new amino acid is added in a cycle: remove the protecting group from the end of the chain, couple the next residue, then simply wash away everything that did not react. Only at the very end is the finished peptide cut off the bead. With step yields of 99.5% or better, the same 100-step synthesis could reach about 61%, and the whole process could be automated ^[2]. Merrifield received the 1984 Nobel Prize in Chemistry ^[2], and most peptides today are still made this way ^[3].

What goes wrong

Even at 99.5% per step, a crude peptide is a crowd of close relatives. The main families are ^[3,4]:

• Deletion sequences. One residue is missing because a deprotection or coupling step was incomplete.
• Insertions. One residue too many, from excess reagent.
• Truncations. Chains that stopped growing early.
• Racemisation. An amino acid flips into its mirror-image (D-) form, producing a diastereomer with exactly the same mass.
• Protecting-group adducts. Side-chain protecting groups that were not fully removed.
• Oxidation. Sensitive side chains pick up oxygen.
• Residual trifluoroacetate. TFA from synthesis and purification stays behind as a counter-ion.

Peptides can also degrade later, through reactions such as pyroglutamate or succinimide formation ^[3]. Our guide on storing research peptides covers that side of the story.

Reverse-phase HPLC: a race through a sticky tunnel

High-performance liquid chromatography is the workhorse purity test. Picture runners racing through a tunnel lined with velcro. The more "greasy" (hydrophobic) a runner is, the more it sticks to the walls and the later it reaches the finish.

In reverse-phase HPLC, the tunnel is a column packed with water-repelling material, and the runners are carried by a gradually rising percentage of acetonitrile in water, usually with TFA added ^[5]. Peptides come off the column in order of increasing hydrophobicity ^[5]. A UV detector set at 210–220 nm, where the peptide bond absorbs, records each one as a peak ^[5].

The output, a chromatogram, shows the target peptide as the dominant peak and its impurities as smaller peaks on either side. Purity is reported as the main peak's share of the total peak area.

HPLC has limits. Two molecules that stick to the column equally come out together and look like one peak, which is why a consortium of proteomics labs convened by the US National Cancer Institute recommended shallow gradients of no more than 2% organic solvent per minute ^[4]. Some impurities, such as certain diastereomers, are hard to separate at all.

Mass spectrometry: weighing molecules

A mass spectrometer answers a different question: does this molecule weigh what the sequence says it should? It works on ions, so the first challenge is getting large, fragile peptides into the gas phase with a charge and without breaking them. Two approaches solved that, and the work was recognised with half of the 2002 Nobel Prize in Chemistry, shared by John Fenn and Koichi Tanaka ^[6].

• Electrospray ionisation (ESI). The sample is sprayed through a strong electric field into charged droplets. As the solvent evaporates, bare, charged molecules are left floating free ^[6]. Each molecule can carry a different number of charges, so one peptide produces a characteristic series of peaks ^[6]. ESI couples naturally to HPLC, which is why "LC-MS" is so common, and it is the method USP scientists used to confirm the mass and sequence of reference peptides ^[7].
• Laser desorption and MALDI. Tanaka showed that an intense laser pulse could lift large molecules such as proteins off a surface as intact ions ^[6]. In today's standard version, MALDI (matrix-assisted laser desorption/ionisation), the sample is first mixed into a light-absorbing matrix. It is fast and usually gives simpler, singly charged spectra.

Breaking the ion into fragments inside the instrument (tandem MS, or MS/MS) reads out the sequence and helps identify what each impurity is ^[4]. But mass spectrometry cannot tell leucine from isoleucine, which weigh exactly the same ^[7], and it does not see mirror-image residues.

A correct mass and a 99% HPLC peak can both describe a vial in which a large share of the powder is salt and water. Only a content measurement catches that.

Amino-acid analysis: taking the peptide apart

Amino-acid analysis (AAA) does the opposite of synthesis. The peptide is heated in strong acid, for example 6 M hydrochloric acid at 110 °C for about 18 hours, until it falls apart into individual amino acids, which are then separated and counted ^[7]. The ratios confirm the composition, and AAA is one of the few routine ways to tell leucine from isoleucine ^[7].

Because AAA measures the absolute amount of each amino acid, it is also the standard way to determine net peptide content: the share of the weighed powder that is actually peptide rather than water and counter-ions ^[4]. Some labs instead measure total nitrogen by elemental analysis. Trifluoroacetate, acetate and water contain no nitrogen, so the nitrogen found can be attributed to the peptide and converted into a content figure from its formula.

Pharmacopoeias combine these results. A USP reference standard, for example, gets its value from a mass balance: HPLC purity, counter-ion content, residual solvents, ash and water, each measured separately ^[7].

Endotoxin: the horseshoe crab test

Endotoxins are fragments of the outer membrane of Gram-negative bacteria. They can cause fever and inflammation even when no live bacteria remain, which is why injectable products, and many cell experiments, are tested for them ^[10].

In 1964, Jack Levin and Frederik Bang reported that the blood of the Atlantic horseshoe crab clots in the presence of bacterial endotoxin ^[8]. For decades, drug testing has relied on a reagent made from those blood cells, Limulus amebocyte lysate (LAL). In 2023, Science reported on a proposal that would let synthetic, recombinant proteins replace the harvested reagent ^[9]. FDA guidance, revised in 2026, describes the gel-clot, photometric and kinetic test methods used under the US Pharmacopeia's chapter on bacterial endotoxins ^[10].

Why independent testing matters

A manufacturer testing its own product has an obvious conflict of interest. The NCI consortium also warned that many companies selling custom peptides do not make them, but resell material made elsewhere, so batch-to-batch consistency is hard to trace ^[4]. Its advice was to insist on MS and HPLC data for every lot, with the testing procedures available alongside ^[4].

An independent laboratory, ideally accredited for the methods it runs, has nothing to gain from a pass. Our guide on how to read a certificate of analysis shows what to look for on the final report, and our primer on what a peptide is covers the basics. At Synthos, every lot is tested by an independent laboratory and ships with its certificate of analysis.

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