Almost every research peptide is made by solid-phase peptide synthesis (SPPS). The first amino acid is anchored to an insoluble resin bead, and the chain is built one residue at a time by repeating two reactions: removing a temporary protecting group from the chain's free end, then coupling the next protected amino acid to it. When the sequence is complete, the peptide is cut from the resin with trifluoroacetic acid, purified by reversed-phase HPLC, checked by mass spectrometry and freeze-dried into a powder.
The rest of this article walks through each stage of peptide synthesis, the chemistry behind it, where impurities come from, and what that means when reading the analytical data that comes with a vial.
What peptide synthesis is
Peptide synthesis is the chemical assembly of a peptide chain from amino acid building blocks. Each step joins the carboxyl group of one amino acid to the free amino group of the next, and that peptide bond formation is repeated until the peptide sequence is complete. Stawikowski and Fields summarize the concept as elongation by a coupling reaction between amino acids, followed by removal of a reversible protecting group.
Protecting groups are what make the synthesis of peptide bonds controllable. Every amino acid has at least two reactive ends, and many have a reactive side chain, so without protection the reagents would join in every possible order. Blocking all but one position at a time is what lets a chemist synthesize one defined sequence instead of a mixture.
There are two ways to run the chemistry. In solution-phase methods every intermediate stays dissolved and has to be isolated; the same authors note that solution synthesis is still valuable for large-scale manufacturing and specialized laboratory applications. In the solid-phase method the chain grows on a solid resin support, which is how most research peptides and custom peptide synthesis orders are made. The first automated peptide synthesizer, based on Boc chemistry, was built by Merrifield, Stewart and Jernberg.
Why peptides are made by chemistry
Cells make peptides on the ribosome, often by cutting them out of larger precursor proteins, but that route is poorly suited to producing a short, pure research compound. Chemical synthesis has several advantages for chains up to a few dozen residues:
- It makes exactly the sequence requested, with no host-cell proteins to remove.
- It can include residues and modifications that cells cannot easily install: D amino acids, non-standard amino acids, a C-terminal amide, N-terminal acetylation, fatty-acid side chains or metal-binding motifs.
- The same chemistry works at small and large scale, and automated synthesizers run it unattended.
For longer chains the balance shifts toward expression in cells, a trade-off discussed in peptides vs proteins.
A short history of solid-phase synthesis
Before the 1960s, peptides were made in solution: each intermediate had to be isolated and purified before the next residue was added, which made even short sequences slow to produce. In 1963, R. Bruce Merrifield described a different approach. He attached the first amino acid to an insoluble polymer support and built a tetrapeptide on it, using the carbobenzoxy (Cbz) group to protect the alpha-amine, N,N'-dicyclohexylcarbodiimide (DCC) to form each bond, and saponification or HBr to release the product. Because the growing chain was fixed to a solid, excess reagents could be washed away by filtration instead of by purification at every step. Merrifield received the 1984 Nobel Prize in Chemistry "for his development of methodology for chemical synthesis on a solid matrix."
Two chemical strategies followed. The Boc/Bzl strategy uses a tert-butyloxycarbonyl (Boc) group on the alpha-amine and benzyl-based side-chain protection, and requires anhydrous hydrogen fluoride for the final cleavage. The Fmoc/tBu strategy uses the 9-fluorenylmethoxycarbonyl (Fmoc) group, introduced by Carpino and Han, which is removed by a mild base, with tert-butyl-based side-chain protection that comes off in trifluoroacetic acid (TFA). Avoiding hydrogen fluoride made Fmoc chemistry easier to run in ordinary laboratories and on automated synthesizers. By 1994, a survey found 98% of participating laboratories were using Fmoc chemistry, and a 2016 review describes Boc SPPS as now used mainly for specialist applications.
The materials
| Component | Role | Common examples |
|---|---|---|
| Resin with linker | Anchors the C-terminal residue; the linker decides the C-terminal group | Wang resin (C-terminal acid), Rink amide (C-terminal amide), 2-chlorotrityl (acid, mild release) |
| Temporary alpha-amine protection | Blocks the chain end between couplings | Fmoc, removed with piperidine in DMF (commonly 20%) |
| Permanent side-chain protection | Keeps reactive side chains inert until the end | Pbf (Arg); Trt (Asn, Gln, Cys, His); OtBu (Asp, Glu); tBu (Ser, Thr, Tyr) |
| Coupling reagents | Activate the incoming carboxyl group | Carbodiimides (DCC, DIC) with HOBt; HBTU, HATU, TBTU |
| Cleavage mixture | Releases the peptide and strips side-chain groups | TFA with scavengers such as water, phenol, thioanisole, ethanedithiol |
Fmoc-protected amino acids are now available at high purity; the 2016 review in the Journal of Peptide Science notes that most are sold at RP-HPLC purity above 99%, which matters because an impurity in a building block is copied into every chain it reaches.
Step by step: building the chain
1. Anchoring the first residue
The C-terminal amino acid, with its alpha-amine protected by Fmoc and its side chain protected as needed, is attached to the resin through the linker. Peptides are therefore built backwards compared with the ribosome, from the C-terminus toward the N-terminus. The choice of linker is made at this point: a Wang linker gives a free acid at the C-terminus after cleavage, and a Rink amide linker gives an amide.
2. Deprotection
The resin is treated with piperidine in DMF, which removes the Fmoc group and exposes a free amine at the end of the chain. The by-product, dibenzofulvene and its piperidine adduct, is soluble and is simply washed away, which is a large part of why Fmoc works so well on a solid support.
3. Washing
Solvent washes remove the deprotection reagent. On a solid support this takes minutes, not a separate purification.
4. Activation and coupling
The next Fmoc-amino acid is activated with a coupling reagent and added in excess. Its activated carboxyl reacts with the free amine on the resin to form a new peptide bond. The excess drives the reaction toward completion, and whatever does not react is washed off. The chemistry of that bond is covered in the peptide bond.
5. Repeat
Deprotection, washing, coupling and washing are repeated for every residue in the sequence. Some syntheses add a capping step after coupling, acetylating any chain that failed to couple so that it stops growing and is easier to separate later.
6. Cleavage and global deprotection
When the last residue is on, the resin is treated with a TFA mixture. The acid cuts the peptide from the linker and removes the side-chain protecting groups at the same time. The protecting groups leave as reactive cations that can attach themselves to tryptophan, methionine, tyrosine or cysteine, so the mixture includes scavengers to trap them. A well-known example, Reagent K, is TFA, phenol, thioanisole, 1,2-ethanedithiol and water in the ratio 82.5:5:5:2.5:5. The crude peptide is then precipitated, usually in cold ether, and collected.
7. Purification
The crude product is a mixture: the target sequence plus shorter and modified chains. It is purified by reversed-phase HPLC, which separates molecules by hydrophobicity on a C18 or similar column using a water and acetonitrile gradient. Fractions containing the target at the required purity are pooled.
8. Analysis
Each lot is checked in two ways. Analytical HPLC gives purity, expressed as the area of the principal peak as a percentage of the total area; the article on peptide purity explains the calculation. Mass spectrometry confirms that the main component has the expected molecular weight, and tandem mass spectrometry or Edman sequencing can confirm the sequence itself.
9. Freeze-drying
The pooled fractions are frozen and lyophilized, removing water and volatile solvents under vacuum to leave a solid. A dry peptide is far more stable than the same peptide in solution, because the water that drives hydrolysis is gone. Peptides purified with TFA in the mobile phase are usually isolated as trifluoroacetate salts, and some are exchanged to acetate or another counter-ion afterwards, which is why a certificate may list the salt form alongside the sequence.
Where impurities come from
No coupling is perfectly efficient, and the losses compound. As an illustration, if each of 20 coupling steps went to 99% completion, only about 82% of chains would carry the full sequence (0.99 multiplied by itself 20 times); at 99.5% per step, about 90% would. Most impurities in a synthetic peptide trace back to a few causes.
| Impurity type | How it forms | How it shows up |
|---|---|---|
| Deletion sequences | A coupling fails and the next residue is added to an uncapped chain | Mass lower by one residue; often elutes close to the target |
| Truncated sequences | Chain growth stops (for example after capping) | Shorter masses; usually easy to separate |
| Aspartimide products | Aspartic acid side chain attacks the backbone during base treatment | Same or near-same mass; can rearrange to beta-aspartyl forms |
| Racemized residues | Activation or base converts an L residue partly to D | Same mass; separated only by a well-resolved HPLC method |
| Incomplete deprotection | A side-chain group survives cleavage | Mass higher by the protecting group |
| Oxidation | Methionine or cysteine oxidized during handling | Mass higher by 16 (methionine sulfoxide) |
Aspartimide formation
The 2016 review in the Journal of Peptide Science calls aspartimide formation the most serious side reaction in Fmoc chemistry. Repeated exposure to piperidine lets the side chain of aspartic acid attack the neighboring backbone nitrogen, forming a ring that can reopen to the wrong isomer. The sequence Asp-Gly is the worst case; other susceptible pairs include Asp-Asp, Asp-Asn, Asp-Arg, Asp-Thr and Asp-Cys. Epitalon, the tetrapeptide Ala-Glu-Asp-Gly listed in the Anhydrolabs catalog as Epithalon, ends in exactly that Asp-Gly motif, so its synthesis needs conditions chosen to suppress the side reaction.
Racemization
Cysteine and histidine are the residues most prone to losing their L configuration during coupling. The same review notes that cysteine racemization is particularly high with base-mediated activation, microwave heating and preactivation, and that histidine can racemize significantly even under standard conditions.
Difficult sequences
Some sequences fold or aggregate on the resin, burying the chain end so that it couples slowly or not at all. Hydrophobic stretches and runs that form beta-sheet structure are the usual culprits. Chemists break up these structures with pseudoproline dipeptides (oxazolidine derivatives of serine or threonine that put a kink in the chain) or backbone protecting groups such as Dmb and Hmb, which are removed during the final cleavage.
How long can a synthetic peptide be?
A figure of around 50 residues is often quoted as the routine limit for stepwise SPPS, but the 2016 review points out that the number means little on its own, because some much shorter sequences are very hard to make and some longer ones are routine. Sequence matters more than length.
For longer targets, chemists join segments. In native chemical ligation, described by Dawson, Muir, Clark-Lewis and Kent in 1994, a peptide carrying a C-terminal thioester reacts with a second peptide that starts with cysteine, and the intermediate rearranges into a native peptide bond at the join. Their demonstration made a cytokine containing multiple disulfide bonds in one ligation step. Beyond a few hundred residues, recombinant expression in cells is usually the practical route.
What this means when you receive a peptide
Knowing how a peptide is made explains what its paperwork shows. The HPLC purity figure tells you what fraction of the UV-absorbing material is the main component; the mass spectrum tells you whether that main component has the right molecular weight. A deletion sequence or a D-isomer can sit close to the main component on the trace, which is why both measurements are needed. The article on how to read a peptide certificate of analysis goes through each field.
Anhydrolabs supplies its peptides as lyophilized powder in vacuum-sealed vials, and as 10-vial kits, each lot with its own analytical data. All are for in-vitro laboratory research only.
