How Research Peptides Are Made: Solid-Phase Peptide Synthesis Explained

A vial of lyophilized research peptide gives no hint of how it was made, but the process behind it is one of the more elegant inventions in twentieth-century chemistry. Before 1963, assembling even a short peptide in solution required months of work, with each intermediate isolated and purified by hand. Bruce Merrifield’s idea was to attach the first amino acid to a plastic bead, build the chain outward while it stayed tethered, and simply wash away everything that was not attached. The method, solid-phase peptide synthesis, made it possible to produce any sequence of modest length on demand, and it remains the way almost every peptide in the research supply chain is manufactured.^1,2 Understanding it explains where impurities come from, why some sequences are harder to make than others, and what a purity figure on a certificate of analysis is actually measuring.

The problem Merrifield solved

Joining two amino acids is chemically simple: activate the carboxyl group of one and let the amino group of the other attack it. The difficulty is control. Each amino acid has both a carboxyl and an amino group, so without protection they will react with themselves in every possible combination. Solution-phase chemists handled this by masking the groups they did not want to react, coupling, purifying the product, unmasking the next reactive site, purifying again, and repeating. Every purification cost material and time, and yields fell geometrically with chain length.

Merrifield’s 1963 paper described a tetrapeptide built on a chloromethylated polystyrene resin.¹ The C-terminal amino acid was attached covalently to the bead. Excess reagents could then be used to drive each step to completion, and the excess was removed by filtration and washing rather than by chromatography. The peptide was never isolated until the very end, when it was cleaved from the resin. The 1984 Nobel Prize in Chemistry recognized that this simplification changed what was practical: the citation noted that the method had made peptide synthesis fast enough to be automated.²

Protecting groups: the grammar of the method

SPPS depends on two tiers of protecting groups that can be removed under different conditions. The temporary group sits on the alpha-amino nitrogen of the incoming amino acid and is removed at the start of each cycle to expose the site for the next coupling. The permanent groups sit on reactive side chains, such as the amine of lysine, the carboxylate of aspartate or the thiol of cysteine, and stay in place until the final cleavage.³

Merrifield’s original chemistry used the Boc group (tert-butyloxycarbonyl) as the temporary protection, removed with trifluoroacetic acid, with side-chain groups and the resin linkage requiring anhydrous hydrogen fluoride for final cleavage. In 1972 Carpino and Han introduced the Fmoc group (9-fluorenylmethoxycarbonyl), which is removed by a mild base, typically piperidine, rather than acid.⁴ This allowed an orthogonal scheme: Fmoc comes off with base, side-chain groups and the resin linker come off with acid, and neither condition disturbs the other. Fmoc/tBu chemistry is now the dominant approach in both research and manufacturing because it avoids hydrogen fluoride and is friendlier to automation.³

One cycle, step by step

With the first amino acid anchored to the resin through a linker, the chain grows from C-terminus to N-terminus, the reverse of the direction in which a ribosome builds a protein. Each cycle follows the same pattern.^3,5

1. Deprotection. A solution of piperidine in DMF removes the Fmoc group from the N-terminus of the resin-bound chain, releasing dibenzofulvene and exposing a free amine. The dibenzofulvene adduct absorbs UV light, and automated synthesizers monitor its release to confirm the step went to completion.
2. Washing. Repeated solvent washes remove piperidine and by-products. Residual base at the coupling stage would cause side reactions.
3. Activation and coupling. The next Fmoc-protected amino acid is activated at its carboxyl group with a coupling reagent, most commonly a uronium or phosphonium salt such as HBTU, HATU or PyBOP together with a tertiary base, or a carbodiimide with an additive such as OxymaPure. The activated ester reacts with the free amine on the resin to form the new peptide bond. The amino acid is used in several-fold excess to push the reaction toward completion.⁶
4. Washing and monitoring. Excess reagent is washed out. A qualitative test for unreacted amines, such as the ninhydrin-based Kaiser test, confirms whether coupling was complete; if free amines remain, the coupling is repeated or the remaining sites are capped with acetic anhydride so they cannot grow further.⁷

The cycle then repeats for the next residue. A 15-residue peptide such as BPC-157 requires 15 cycles, a 29-residue peptide such as sermorelin requires 29, and each cycle takes roughly one to two hours on a conventional instrument or minutes on a microwave-assisted synthesizer.

Solid-phase synthesis does not make peptides perfectly; it makes them predictably, which is what allows the imperfections to be found and removed.

Cleavage and deprotection

When the last residue has been added and its Fmoc group removed, the resin is treated with a cleavage cocktail. For Fmoc/tBu chemistry this is concentrated trifluoroacetic acid (TFA), usually 90–95 percent, with scavengers such as water, triisopropylsilane and dithiothreitol or thioanisole. The acid simultaneously breaks the linker between peptide and resin and strips the acid-labile side-chain protecting groups. The scavengers intercept the reactive carbocations released from those groups, which would otherwise alkylate sensitive residues such as tryptophan, methionine and cysteine.^3,5

The peptide is precipitated from the TFA solution with cold ether, collected, dissolved in aqueous buffer and, in most cases, lyophilized to give a crude product. At this stage the material is typically between 50 and 85 percent the desired sequence by HPLC, depending on length and difficulty. The remainder is the reason purification exists.

Where impurities come from

Every step of SPPS is efficient but not perfect, and the imperfections accumulate. If each coupling proceeds at 99 percent, a 20-residue peptide will emerge with only about 82 percent of chains having every residue in place (0.99 raised to the twentieth power). At 98 percent per step the figure drops to 67 percent. The characteristic impurities are well catalogued in the analytical literature.⁸

Impurity class	Origin	Mass difference from target
Deletion sequence	Incomplete coupling at one position, chain continues without that residue	Minus one residue
Truncated sequence	Chain capped or permanently blocked mid-synthesis	Missing N-terminal segment
Insertion sequence	Incomplete Fmoc removal followed by double coupling in the next cycle	Plus one residue
Incompletely deprotected peptide	Side-chain protecting group survives cleavage	Plus the group (e.g. +56 for tBu, +100 for Boc)
Oxidized peptide	Methionine or tryptophan oxidation during cleavage or storage	Plus 16
Racemized peptide	Epimerization at the alpha carbon during activation, especially histidine and cysteine	None (same mass, different retention time)
Aspartimide-derived products	Base-catalyzed cyclization at Asp-Gly or Asp-Ser during Fmoc removal	Minus 18 or isomeric

The racemization row deserves emphasis. A peptide containing one D-amino acid in place of the L-form has exactly the same mass as the target, so mass spectrometry alone cannot detect it. Only a chromatographic method that separates the two diastereomers will show it, which is one reason HPLC and mass spectrometry are used together on a certificate of analysis rather than either alone.⁸

Difficult sequences

Not all peptides are equally easy to make. Long hydrophobic stretches can cause the growing chains to aggregate on the resin, folding into beta-sheet structures that block reagent access and cause coupling efficiency to collapse.^5,9 Sequences rich in beta-branched residues (valine, isoleucine, threonine) are prone to this, and so are peptides with several consecutive alanines or glycines. Chemists address these problems with pseudoproline dipeptides that temporarily kink the backbone, with backbone-protecting groups such as Dmb or Hmb, with elevated temperature or microwave heating, and with alternative solvents.^5,9 The practical consequence for a researcher is that crude purity varies widely by sequence, and two suppliers can legitimately report different pre-purification yields for the same peptide.

Purification and final form

The crude peptide is purified almost universally by preparative reversed-phase HPLC, using a water–acetonitrile gradient with 0.1 percent TFA as the ion-pairing agent. Fractions are collected across the main peak, analyzed, and pooled according to the purity target. The pooled fractions are then lyophilized. Because the mobile phase contained TFA, the peptide comes out of this step as a trifluoroacetate salt, with TFA counter-ions balancing the positive charges on lysine, arginine and the N-terminus. Some applications require exchanging TFA for acetate or chloride, a separate step that many research-grade products do not undergo.¹⁰ This is why the mass of powder in a vial is not the same as the mass of peptide, a distinction our note on purity and the planned note on net peptide content explore.

The lyophilized solid is then weighed into vials, capped and sealed. Our lyophilization note explains why the dry form is preferred for shipping and storage.

Scale, cost and sustainability

SPPS is reagent-hungry. The large excesses of amino acid and coupling reagent that make each step reliable, together with the extensive washing, mean that producing one kilogram of peptide can consume several thousand kilograms of solvent, mostly DMF and acetonitrile.¹¹ Manufacturing scale-up therefore involves trade-offs between speed, purity and waste, and pharmaceutical process chemists have published extensively on greener solvents, reduced excesses and hybrid solid-phase/solution-phase strategies for long peptides.¹¹ None of this changes the chemistry a researcher receives, but it does explain why longer peptides cost disproportionately more per milligram than shorter ones.

What the method means for research material

Three practical points follow from the chemistry. First, purity is a property of a specific lot, because crude purity and the choices made during preparative HPLC vary from run to run; that is why Wednesday reports results lot by lot in the COA library. Second, mass spectrometry confirms identity and HPLC confirms purity, and a complete report needs both. Third, the counter-ion and residual water contribute to the weight of what is in the vial, so purity and content are different questions. The next notes in this series, on purity and on HPLC, take up each of these in turn.

Frequently asked questions

How are research peptides made?

Almost all are made by solid-phase peptide synthesis. The first amino acid is attached to a resin bead, and each subsequent amino acid is added in a repeating cycle of deprotection, washing and coupling. When the sequence is complete the peptide is cleaved from the resin with acid, purified by preparative HPLC and freeze-dried.

What is Fmoc solid-phase peptide synthesis?

Fmoc SPPS is the version of the method that uses the base-labile Fmoc group to protect the N-terminus of each incoming amino acid. Fmoc is removed with piperidine, while side-chain protecting groups are removed later with trifluoroacetic acid. This orthogonal scheme, introduced by Carpino and Han in 1972, is now standard in both laboratory and industrial synthesis.

Why is peptide purity never 100 percent?

Each coupling step in synthesis is slightly incomplete, so a fraction of chains miss a residue or stop growing. These deletion and truncation sequences, together with oxidation and protecting-group by-products, must be separated out by HPLC. Purification removes most but never all of them, so purity is reported as a percentage rather than assumed to be complete.

Are synthetic peptides the same as natural ones?

A correctly synthesized peptide has the same sequence and covalent structure as its natural counterpart. Differences arise in the surrounding material: a synthetic peptide carries a counter-ion such as trifluoroacetate from purification and may contain closely related impurities, whereas a natural peptide is embedded in a biological matrix. Mass spectrometry confirms that the synthetic sequence matches the intended one.

Why do longer peptides cost more to make?

Each additional residue adds a full synthesis cycle, and because coupling is never perfectly efficient, crude purity falls as length increases. Longer peptides therefore need more reagent, more purification effort and yield less final material per gram of starting resin.

References & further reading

1. Merrifield RB. Solid phase peptide synthesis. I. The synthesis of a tetrapeptide. J Am Chem Soc. 1963;85(14):2149-2154. doi:10.1021/ja00897a025
2. The Nobel Prize in Chemistry 1984. Press release. Nobel Foundation, 17 October 1984. nobelprize.org
3. Behrendt R, White P, Offer J. Advances in Fmoc solid-phase peptide synthesis. J Pept Sci. 2016;22(1):4-27. doi:10.1002/psc.2836 / PMID 26785684
4. Carpino LA, Han GY. The 9-fluorenylmethoxycarbonyl amino-protecting group. J Org Chem. 1972;37(22):3404-3409. doi:10.1021/jo00795a005
5. Coin I, Beyermann M, Bienert M. Solid-phase peptide synthesis: from standard procedures to the synthesis of difficult sequences. Nat Protoc. 2007;2(12):3247-3256. doi:10.1038/nprot.2007.454
6. Albericio F, El-Faham A. Choosing the right coupling reagent for peptides: a twenty-five-year journey. Org Process Res Dev. 2018;22(7):760-772. doi:10.1021/acs.oprd.8b00159
7. Kaiser E, Colescott RL, Bossinger CD, Cook PI. Color test for detection of free terminal amino groups in the solid-phase synthesis of peptides. Anal Biochem. 1970;34(2):595-598. doi:10.1016/0003-2697(70)90146-6 / PMID 5443684
8. D’Hondt M, Bracke N, Taevernier L, et al. Related impurities in peptide medicines. J Pharm Biomed Anal. 2014;101:2-30. doi:10.1016/j.jpba.2014.06.012 / PMID 25044089
9. Mueller LK, Baumruck AC, Zhdanova H, Tietze AA. Challenges and perspectives in chemical synthesis of highly hydrophobic peptides. Front Bioeng Biotechnol. 2020;8:162. doi:10.3389/fbioe.2020.00162 / PMID 32195241
10. Roux S, Zékri E, Rousseau B, Paternostre M, Cintrat JC, Fay N. Elimination and exchange of trifluoroacetate counter-ion from cationic peptides: a critical evaluation of different approaches. J Pept Sci. 2008;14(3):354-359. doi:10.1002/psc.951 / PMID 18035848
11. Isidro-Llobet A, Kenworthy MN, Mukherjee S, et al. Sustainability challenges in peptide synthesis and purification: from R&D to production. J Org Chem. 2019;84(8):4615-4628. doi:10.1021/acs.joc.8b03001