Solid-phase peptide synthesis (SPPS): complete guide to Merrifield, Fmoc, Boc, resins, coupling reagents and quality control
Solid-phase peptide synthesis (SPPS) is the standard industrial and laboratory method for manufacturing modern synthetic peptides, from BPC-157 to semaglutide through retatrutide and cagrilintide. Invented in 1963 by Robert Bruce Merrifield at Rockefeller University (Nobel Prize in Chemistry 1984), SPPS rests on a simple but revolutionary principle: anchor the first amino acid on an insoluble support (polymeric resin) and grow the peptide chain residue by residue, removing excess reagents after each cycle by simple filtration. This logic transformed a craft process spanning several months into an automatable routine of a few hours to a few days.
This guide consolidates the entire scientific foundation of SPPS: Merrifield principle, Boc and Fmoc strategies, resins, coupling reagents, complete synthesis cycle, cleavage, purification by reverse-phase high-performance liquid chromatography (RP-HPLC), quality control by coupled mass spectrometry (LC-MS) and colorimetric tests (Kaiser, TNBS, chloranil), difficult peptides, modern innovations in flow chemistry and industrial outlook. All references are intended for research use only (RUO) in accordance with applicable requirements in France and Europe.
Merrifield principle: the founding invention
Before Merrifield, peptide synthesis was carried out in solution (LPPS, Liquid-Phase Peptide Synthesis): each step involved chromatographic separations or crystallizations that turned synthesizing a 10-residue peptide into a multi-month project with a cumulative yield often below 10%. In 1963, Merrifield published in the Journal of the American Chemical Society (JACS 85:2149) the synthesis of the tetrapeptide Leu-Ala-Gly-Val, starting from an amino acid anchored on a chloromethylated styrenic resin and successively adding each next residue through deprotection-coupling-wash cycles. This innovation earned Merrifield the Nobel Prize in Chemistry in 1984.
The operating principle is as follows. The first amino acid (C-terminal of the target sequence) is attached through its carboxyl function to a resin via a linker. Its alpha amino group (N-alpha) bears a temporary protecting group. The synthesis cycle comprises four steps: N-alpha deprotection (removal of the temporary protecting group), wash (elimination of residual reagents), coupling (activation of the incoming amino acid's carboxyl and formation of the peptide bond with the chain's free amine), wash again. This cycle is repeated for each residue of the sequence, from the C-terminal end to the N-terminal end. After complete assembly, the peptide is cleaved from the resin and side-chain protecting groups are removed.
Boc and Fmoc strategies: the two schools
Two N-alpha temporary protection strategies have historically coexisted.
The Boc strategy (tert-butyloxycarbonyl), developed by Merrifield, uses the Boc group on the alpha amine, removed by trifluoroacetic acid (TFA) at each cycle. Side chains are protected by groups stable to TFA (benzyl, cyclohexyl), and the final peptide is cleaved from the resin and deprotected using anhydrous hydrogen fluoride (HF) or trifluoromethanesulfonic acid. This strategy produces excellent-quality peptides but requires HF, a highly toxic and corrosive reagent, and has progressively been replaced in most academic laboratories.
The Fmoc strategy (9-fluorenylmethyloxycarbonyl), introduced by Louis Carpino in 1970 and popularized by Robert Sheppard from 1978, uses the Fmoc group on the alpha amine, removed by 20% piperidine in DMF (dimethylformamide) via a beta-elimination reaction. Side chains are protected by groups stable in basic medium but sensitive to TFA (tert-butyl, trityl, 2,2,4,6,7-pentamethyldihydrobenzofuran-5-sulfonyl Pbf for arginine). Final cleavage and complete deprotection are done simultaneously with a typical TFA cocktail (95% TFA, 2.5% water, 2.5% triisopropylsilane TIS, Reagent K or Reagent R cocktail). The absence of HF and modularity have made Fmoc-SPPS the dominant strategy since the 1990s.
Resins: the choice of support
The choice of resin determines the final peptide format (free acid or C-terminal amide) and the cleavage chemistry.
The Wang resin (4-hydroxymethyl-phenoxymethyl-polystyrene), introduced by Su-Sun Wang in 1973, is the most widely used for acid peptides in Fmoc-SPPS. The first amino acid is attached by esterification to the resin's hydroxyl group, and the final peptide is released as a C-terminal acid by TFA.
The Rink amide resin, based on the 4-(2',4'-dimethoxyphenyl-Fmoc-aminomethyl)phenoxy linker, is the standard for C-terminal amide peptides in Fmoc-SPPS. After TFA cleavage, the peptide bears a C-terminal CO-NH2 function, a frequent format for bioactive peptides (BPC-157, MT-II, ipamorelin).
The 2-chlorotrityl chloride resin, introduced by Kostas Barlos in 1989, offers a very mild anchoring cleavable by 1% TFA or hexafluoroisopropanol (HFIP), preserving side-chain protecting groups. It is preferred for fragment synthesis for convergent couplings (segmented synthesis) and for cyclic peptides with final cyclization.
The Merrifield resin (original chloromethylated polystyrene) remains used in Boc strategy and for some specialized resins.
Coupling reagents: activating the carboxyl
Peptide bond formation requires activation of the incoming amino acid's carboxyl group to make it reactive toward the growing free amine.
The carbodiimides (DIC, diisopropylcarbodiimide; DCC, dicyclohexylcarbodiimide) are the historical activators. They form an unstable O-acylisourea intermediate that reacts with the amine to form the peptide bond. Adding HOBt (1-hydroxybenzotriazole) or HOAt (1-hydroxy-7-azabenzotriazole) suppresses racemization and accelerates coupling via formation of an activated ester intermediate. DIC/HOBt remains the reference cocktail for standard peptides.
The aminium/uronium salts (HBTU, O-(benzotriazol-1-yl)-N,N,N',N'-tetramethyluronium hexafluorophosphate; HATU, the azabenzotriazole equivalent) are high-performance coupling reagents. HATU, introduced by Carpino in 1993, is particularly effective for difficult peptides, sterically hindered amino acids and cyclizations. HCTU and TCTU are chlorinated variants offering a cost-performance compromise.
The phosphonium salts (PyBOP, BOP, PyAOP) constitute an alternative family particularly suited to N-methyl amino acid couplings and macrocyclizations.
All these activators are typically used at 1.5-3 equivalents per cycle with a tertiary base (DIEA, diisopropylethylamine; NMM, N-methylmorpholine) at 3-6 equivalents, in DMF or NMP (N-methylpyrrolidone) as solvent.
Complete synthesis cycle
A typical Fmoc-SPPS cycle proceeds as follows:
Step 1 - Fmoc deprotection: 20% piperidine in DMF, 2 x 5 min. Fmoc removal releases dibenzofulvene and the growing chain's alpha amine.
Step 2 - Washes: DMF x 5, removing piperidine and dibenzofulvene residues.
Step 3 - Coupling: Fmoc-AA-OH (3-5 eq), HBTU or HATU (3-5 eq), DIEA (6-10 eq) in DMF, 30-60 min at room temperature. Carboxyl activation generates the OBt activated ester that reacts with the free amine.
Step 4 - Capping (optional but recommended): acetic anhydride (0.5 M) and DIEA in DMF, 5 min. Acetylation of uncoupled chains prevents truncated sequence formation (deletion peptides) in subsequent cycles.
Step 5 - Washes: DMF x 5, DCM x 3, preparation for the next cycle.
Coupling monitoring uses the Kaiser test (ninhydrin) detecting free primary amines: a negative test (yellow) indicates complete coupling, a positive test (intense blue) signals incomplete coupling requiring additional coupling (double coupling). For secondary amines (proline, N-methyl), the chloranil test is used.
Cleavage and final deprotection
In standard Fmoc-SPPS, cleavage and complete deprotection occur simultaneously with a TFA cocktail. Classic cocktails are:
Cocktail K (Reagent K): 82.5% TFA, 5% phenol, 5% thioanisole, 5% water, 2.5% ethanedithiol EDT. Used for peptides containing methionine and cysteine, where phenol, thioanisole and EDT trap released tert-butyl cations.
Cocktail R (Reagent R): 90% TFA, 5% thioanisole, 3% anisole, 2% EDT. Variant for peptides containing tryptophan.
Standard cocktail: 95% TFA, 2.5% water, 2.5% triisopropylsilane TIS. The most frequent for peptides without sensitive residues. Easier handling and less aggressive odor than EDT.
Cleavage lasts 2 to 4 hours at room temperature. The crude peptide is precipitated by adding cold diethyl ether (which solubilizes scavengers and protecting groups but precipitates the peptide), centrifuged, washed several times, redissolved in water / acetonitrile / acetic acid, and lyophilized to obtain the crude peptide before purification.
Preparative RP-HPLC purification
The crude peptide from cleavage contains the majority target peptide, deletion peptides (truncated sequences), oxidized peptides, incomplete deprotection by-products, and sometimes racemized forms. Purification is achieved by preparative reverse-phase high-performance liquid chromatography (preparative RP-HPLC).
Classical conditions use C18 columns (or C8 for more hydrophobic peptides) with large cross-section (50-100 mm internal diameter) and acetonitrile gradients in TFA 0.1% acidified water (ion pairing). Eluted fractions are analyzed by analytical LC-MS to identify the fraction containing the target peptide, then pooled and lyophilized. The final purity required for research is typically > 98% by analytical HPLC, confirmed by high-resolution mass spectrometry.
For very hydrophilic peptides (short polyribonucleotides, acidic fragments), reverse-phase HPLC purification may be replaced by ion-exchange chromatography or size-exclusion chromatography.
Quality control: LC-MS and characterization
Quality control of a research-grade peptide comprises several standard analyses.
Analytical LC-MS: confirms the exact molecular mass and chromatographic purity. Electrospray ionization (ESI-MS) generates multi-charged ions characteristic of peptides; deconvolution yields the monoisotopic molecular mass. High-resolution mass spectrometry (Orbitrap, Q-TOF) achieves mass accuracy < 5 ppm, sufficient to distinguish deamidation or oxidation isomers.
Amino acid analysis (AAA): complete acid hydrolysis (HCl 6 N, 110 °C, 24 h) followed by amino acid quantification by HPLC with post-derivatization (ninhydrin, OPA) or pre-derivatization (AQC, PITC). Validates relative content in each residue.
N-terminal sequencing (Edman degradation): confirms the first 10 to 20 residues of the sequence. Useful for doubled or problematic synthetic peptides.
NMR analysis (proton and carbon): ultimate reference for complex cyclic peptides and to verify functional group integrity.
The Certificate of Analysis (CoA) accompanying each synthesized batch reports at minimum: molecular mass confirmed by MS, HPLC purity, net content (after water and salt correction), residual TFA, and reference batch.
Difficult peptides: aggregation and problem sequences
Some sequences are intrinsically difficult to synthesize by SPPS. Difficult peptides are those showing a tendency to aggregate on resin (beta-sheets in the growing chain), slowing coupling and deprotection. Typical symptoms are incomplete coupling cycles despite standard conditions, deletion peptides dominant in the crude.
Workaround strategies include: alternative solvents (NMP, DMF+DMSO, GVL) that disrupt aggregates, more powerful activators (HATU, COMU), systematic double couplings, incorporation of pseudoproline dipeptides (Ser(psiMe,Mepro)) that break beta-sheets by imposing a turn, strategic N-methylations, and microwave synthesis that accelerates kinetics and diffusion.
Long peptides (> 50 residues) often benefit from a segmented approach: synthesis of short fragments separately, purification, then native chemical ligation (NCL) via cysteine residues. This approach enables synthetic proteins of 200 to 300 residues with superior quality compared with direct linear synthesis.
Modern innovations: flow chemistry and microwave
Classical batch SPPS has been revolutionized by several recent innovations. Flow synthesis (flow SPPS), developed notably by Bradley Pentelute's team at MIT, reduces each coupling-deprotection cycle from 30-60 minutes to 40-90 seconds thanks to a continuous mixer and high flow rates. A 30-residue peptide can be synthesized in less than an hour instead of 24 hours. AFPS (Automated Fast-Flow Peptide Synthesis) machines combine speed, quantity and quality.
Microwave synthesis, commercialized by CEM (Liberty Blue) and Biotage, accelerates each step by controlled heating at 75-90 °C, reducing cycles to 4-8 minutes and improving quality for difficult peptides. It is particularly suited to peptides containing histidine, arginine or tryptophan, residues whose coupling is improved by thermal agitation.
AutoMAP (automated peptide manufacturing) and high-throughput synthesis platforms enable production of thousands of sequences in parallel for ligand identification, epitope mapping and library screening applications.
Industrial SPPS and commercial production
Industrial SPPS, practiced by specialized manufacturers (Bachem, PolyPeptide Group, PeptaNova, CordenPharma), operates at scales ranging from gram to kilogram per batch. Modern commercial peptides (semaglutide, tirzepatide, liraglutide) require industrial yields and GMP pharmaceutical purity (Good Manufacturing Practice, ICH Q7). Reactors can reach 50 to 200 liters, with resins loaded at 0.5-1 mmol/g.
Hybrid SPPS-LPPS transition is sometimes used for very long peptides: SPPS to assemble 15-25 residue fragments, then fragment couplings in liquid phase. Semaglutide, 31 residues, is produced by direct solid-phase SPPS. Tirzepatide, 39 residues, and retatrutide, 39 residues, use optimized hybrid strategies.
Conclusion
SPPS is the central technology of modern peptide pharmacology. Invented by Merrifield more than sixty years ago, it transformed peptide synthesis from an artisanal chemistry project into an industrial production line capable of delivering pharmaceutical quantities of complex molecules such as semaglutide, tirzepatide and retatrutide. Innovations in flow chemistry, microwave and AutoMAP continue to improve speed, quality and accessibility, positioning SPPS as one of the pillars of the 21st-century biotech industry. For any rigorous research peptide, mastery of the synthesis-cleavage-purification-quality control chain is the guarantee of scientific reproducibility.
Frequently asked research questions on solid-phase peptide synthesis
Why are peptides synthesized C-terminus to N-terminus and not the other way?
C-terminus to N-terminus synthesis is imposed by the need to protect the carboxyl to avoid self-couplings and oligomerizations. Starting from the C-terminal residue anchored by its carboxyl to the resin, only the incoming amino acid's carboxyl is activated at each cycle, and only the growing chain's free alpha amine can react. This directionality eliminates parasitic reactions and enables reproducible chemistry.
Why did the Fmoc strategy supplant the Boc strategy?
The Boc strategy imposes a final anhydrous HF cleavage, an extremely toxic and corrosive reagent requiring specialized equipment (Teflon reactors, double gloves, dedicated fume hoods). The Fmoc strategy uses piperidine (mild base) for cyclic deprotection and TFA for final cleavage, both manageable in standard laboratories. Increased modularity, compatibility with modern side chains and better robotic integration completed the switch.
What is the difference between a C-terminal acid peptide and a C-terminal amide peptide?
The difference lies in the terminal function: -COOH for the acid, -CONH2 for the amide. This choice depends on the target biological sequence. Many natural bioactive peptides (BPC-157, ipamorelin, MT-II) are C-terminal amidated, with physiological amidation catalyzed by peptidylglycine alpha-amidating monooxygenase (PAM). In SPPS, the resin choice (Wang vs Rink amide) determines the final format.
How is it verified that a synthesized peptide matches the target sequence?
Sequence confirmation combines several analyses: high-resolution LC-MS to verify the exact mass; enzymatic digestion (trypsin, chymotrypsin, Asp-N) followed by MS/MS to map internal fragments; amino acid composition analysis to confirm proportions; and Edman sequencing on the first 10-20 residues. Together, these analyses converge on an unambiguous confirmation of primary structure.
