Peptide synthesis: how Fmoc SPPS actually works

Solid-phase peptide synthesis (SPPS) is a method for assembling peptides one amino acid at a time on an insoluble polymer support. It was invented by Bruce Merrifield at Rockefeller University in 1963 and earned him the 1984 Nobel Prize in Chemistry. The fundamental insight is simple: if you anchor the growing peptide chain to a bead, you can wash away every excess reagent and byproduct after each step instead of purifying intermediates. That single change made it practical to manufacture peptides at the scale modern medicine depends on.

Today, essentially all commercial peptide manufacturing — from research compounds to GLP-1 analogs to antimicrobial drugs — uses some variant of SPPS. The chemistry has evolved in two directions: Fmoc (9-fluorenylmethyloxycarbonyl) chemistry has become the modern default because it avoids highly toxic anhydrous HF; Boc (tert-butyloxycarbonyl) chemistry is still preferred for certain difficult sequences. The rest of this article focuses on Fmoc SPPS.

The synthesis begins with a functionalized polymer resin — typically polystyrene crosslinked with divinylbenzene, or a polyethylene glycol composite. Three resin choices are standard:

• Wang resin — gives a free C-terminal carboxylic acid after cleavage. Standard for peptides terminating in -COOH.
• Rink amide resin — gives a C-terminal amide after cleavage. Required when the natural peptide is C-terminally amidated (very common in neuropeptides and hormones).
• 2-chlorotrityl chloride resin — mildly acid-labile, lets the chemist cleave the protected peptide off the resin under mild conditions. Useful for fragment condensation and head-to-tail cyclic peptides.

Resin loading is measured in millimoles of attachment site per gram of dry resin (typical 0.3 to 1.0 mmol/g). Loading directly determines the maximum theoretical yield of the synthesis, and over-loaded resin is one of the most common causes of difficult-sequence aggregation.

Once the C-terminal residue is attached to the resin, every subsequent amino acid is added by repeating the same three-step cycle: deprotect, couple, wash. The chain grows from C-terminus to N-terminus — the opposite direction from how ribosomes synthesize proteins.

1. Fmoc deprotection. The temporary Fmoc group on the alpha-amine of the most recent residue is removed with 20% piperidine in DMF. This frees up the amine for the next coupling.
2. Coupling. The next Fmoc-protected, side-chain-protected amino acid is activated with a coupling reagent (HBTU, HATU, DIC + Oxyma, or PyBOP) and a tertiary amine base (DIPEA or NMM). The activated carboxyl reacts with the resin-bound amine to form a new amide bond.
3. Wash. Excess reagents and byproducts are washed away with DMF and DCM.

A modern automated synthesizer runs this cycle in roughly 30 to 90 minutes per residue. A 30-residue peptide therefore takes 1 to 2 days of continuous synthesis before it is even cleaved from the resin.

Each coupling cycle is highly efficient but never perfect. If the average per-cycle efficiency is 99.5%, the maximum theoretical purity of the crude peptide before any side reaction is:

• 10 residues: 0.995⁹ = ~95.6%
• 30 residues: 0.995²⁹ = ~86.5%
• 50 residues: 0.995⁴⁹ = ~78.2%
• 80 residues: 0.995⁷⁹ = ~67.2%

And 99.5% per cycle is generous. Difficult sequences — long beta-sheet stretches, hindered residues like Val-Val-Val or Ile-Ile-Ile, or strongly aggregating sequences — can drop coupling efficiency to 95% or worse. This is the structural reason why long peptides need extensive purification and why single-letter sequence changes can dramatically alter the impurity profile of a finished product.

Even when each coupling proceeds, several reactions can damage the chain:

• Deletion sequences. A coupling that fails outright produces a chain missing one residue. Subsequent cycles continue normally, so the impurity differs from the target by exactly one amino acid — often very hard to separate by HPLC.
• Truncations. A chain that fails to deprotect is permanently capped. The result is a shorter peptide that elutes earlier than the target.
• Racemization. Activated amino acids can lose stereochemistry at the alpha-carbon, especially Cys and His. The resulting D-isomer has identical mass but different biological activity.
• Aspartimide formation. Asp-X bonds (especially Asp-Gly) cyclize during repeated piperidine treatments, producing a 5-membered ring that opens to alpha- and beta-aspartyl byproducts.
• Side-chain damage during cleavage. The TFA cleavage cocktail generates electrophilic cations that can alkylate Trp, Met, Cys, and Tyr unless properly scavenged.

These are the impurities that show up as extra peaks on an HPLC chromatogram and as satellite masses in LC-MS — and they are why an honest analytical lab is the only way to know what is actually in a finished peptide.

After the final residue is coupled, the peptide is still attached to the resin and still wears every side-chain protecting group it was built with. A single cocktail step does both jobs at once.

A typical Fmoc cleavage cocktail (Reagent K) is 82.5% TFA, 5% phenol, 5% water, 5% thioanisole, 2.5% EDT. The peptide is shaken with this cocktail for 1.5 to 4 hours at room temperature. TFA cleaves the resin linkage and removes all acid-labile protecting groups (Boc, tBu, Trt, Pbf, OtBu). The water, phenol, EDT, and thioanisole serve as scavengers, soaking up the reactive cations released from the Trt and Pbf groups before they can damage Trp, Cys, Met, or Tyr side chains.

The crude peptide is then precipitated by pouring the cocktail into cold diethyl ether (typically −20°C), where the peptide drops out as a solid while small organic byproducts stay in the ether. After centrifugation, decanting, and lyophilization, what remains is the crude peptide — ready for analytical HPLC and preparative purification.

Crude SPPS material is rarely above 70 to 90% pure. It contains the target sequence plus deletions, truncations, oxidized analogs, and side-chain modified species. To produce a usable peptide, the crude is purified by preparative reverse-phase HPLC, often with a counter-ion exchange step from TFA salt to the desired counter-ion (acetate, hydrochloride, or free base). The purified peptide is then lyophilized and characterized.

The next two articles in this series cover those steps in detail: peptide purification by reverse-phase HPLC and characterizing a peptide by HPLC, LC-MS, AAA, and NMR.