A bench-ready protocol for manual Fmoc solid-phase peptide synthesis, from swelling dry resin in DMF through the piperidine deprotection and HBTU coupling cycles, Kaiser test monitoring, TFA test and full cleavage, ether precipitation, and preparative C18 HPLC purification, with scale, reagent…
Fmoc solid-phase peptide synthesis SPPS can be run by hand with a fritted cartridge, a vacuum line, a shaker, and a standard set of reagents. The procedure below takes a peptide from dry resin to lyophilized crude product, and then to purified material: swell the resin, deprotect, couple, repeat, cleave, precipitate, and purify by HPLC. It is written for a researcher who has trained in organic chemistry laboratory safety and wants a complete, bench-ready sequence with expected yields at each stage.
SPPS builds a peptide from its C-terminus to its N-terminus, the opposite direction from ribosomal protein synthesis. The C-terminus is anchored to insoluble resin beads through a linker, and the chain grows one amino acid at a time. Because the peptide stays on the solid support, every excess reagent is removed by draining and washing rather than by chromatography. The resin used throughout this protocol is Rink amide resin , whose linker produces a C-terminal primary amide when cleaved with acid.
The Fmoc group 9-fluorenylmethoxycarbonyl is the most commonly used protecting group for the alpha-amine in SPPS. Fmoc chemistry is base-labile for deprotection, which leaves acid-labile side-chain protecting groups and the resin linker intact until the final cleavage step. Boc chemistry, which uses acid for every deprotection, is an alternative strategy but is not part of this protocol.
Deprotection removes the Fmoc group. Piperidine in DMF acts as a secondary amine base that abstracts the acidic proton on the fluorene ring, triggering beta-elimination. The products are dibenzofulvene, carbon dioxide, and a free primary amine at the peptide N-terminus. Piperidine also scavenges dibenzofulvene to form a stable adduct, preventing the reactive alkene from alkylating the growing peptide.
Coupling activates the carboxyl group of the incoming Fmoc-amino acid. HBTU , a benzotriazole-based coupling reagent, together with the tertiary amine base DIEA N,N-diisopropylethylamine , converts the carboxylic acid into an activated benzotriazol-1-yl OBt ester in situ. The resin-bound N-terminal amine attacks this ester to form the peptide bond. Excess reagents are used because no coupling is perfectly quantitative, and the excess is washed away after each step.
DMF is the standard solvent for this chemistry because it swells polystyrene resin well and dissolves Fmoc-amino acids, HBTU, and DIEA at the concentrations used. The 20% piperidine concentration in DMF is a workable compromise: high enough to deprotect within minutes and to scavenge the dibenzofulvene released by the reaction.
The final step is acidolytic cleavage . Trifluoroacetic acid severs the Rink amide linker and removes side-chain protecting groups such as tert-butyl ethers and esters. Those groups release carbocations, which can alkylate the peptide; scavengers such as water and triisopropylsilane TIPS trap the cations instead.
The simplest reaction vessel is a disposable solid-phase extraction cartridge fitted with a polyethylene frit. The frit pore size is 20 µm. A 3 mL cartridge suits roughly 100 mg of resin with 1 mL of solvent. A 6 mL cartridge is the better choice when solvent volumes exceed 1 mL or when the solvent is volatile. The cartridge fits on a vacuum manifold or a side-arm flask with a waste trap; a water aspirator draws solvent through the frit. Both ends are capped during agitation.
| Vessel | Resin scale | Solvent volume | Notes |
|---|---|---|---|
| 3 mL fritted cartridge | about 100 mg | 1 mL | standard for this protocol |
| 6 mL fritted cartridge | about 100 mg | above 1 mL | preferred for volatile solvents |
| 10 mL fritted syringe | up to 200 µmol | 2 mL minimum | suited to parallel synthesis |
A 10 mL fritted syringe is a practical alternative for syntheses up to about 200 µmol, and a nutating mixer handles as little as 2 mL of solution in a 10 mL vessel. Syringe setups adapt naturally to parallel synthesis of many peptides on a manifold. One constraint applies to typical vacuum manifolds: the supplied stopcocks are not stable to organic solvents, and the manifold reservoir has not been tested with trifluoroacetic acid. Replace the stopcocks with solvent-stable valves before the first run, and keep TFA out of the manifold body.
Agitation is a frequent source of ruined resin. Magnetic stirring crushes the beads and must not be used. Orbital shaking, a vortex mixer with a microplate tray, or a nutating mixer are all acceptable.
Scale sets expectations for yield. Resin at 0.6 to 0.8 mmol/g loading, at 100 mg, carries about 0.06 to 0.08 mmol of synthesis sites. For a pentapeptide near 800 g/mol, complete conversion gives roughly 50 mg of crude peptide after cleavage. Purification of a peptide made from standard Fmoc amino acids typically recovers 30 to 40% of the crude mass, which is 15 to 25 mg of pure peptide from this example. The loss is mostly deletion peptides and truncated byproducts from couplings that did not go to completion.
Before starting, confirm access to LCMS, a preparative HPLC system, and a lyophilizer. Cover the bench with aluminum foil or keep cartridges in plastic bags, because spills are routine. Empty the waste trap before each run.
Dry resin must swell before chemistry begins. Add about 1 mL of DMF per 100 mg of resin and agitate for 30 minutes. Swelling opens the polymer network so reagents can reach the internal sites. Drain the DMF and start the first cycle.
Deprotection uses 20% piperidine in DMF by volume. Add about 1 mL, agitate for 5 minutes, and drain. Add fresh 20% piperidine solution and agitate for 15 minutes. No wash is needed between the two piperidine treatments; the first portion removes most of the Fmoc groups and the second clears the remainder, so the new N-terminus is fully exposed. After the second treatment, wash the resin with 3 × 1 mL of DMF.
For the coupling, dissolve 3 equivalents of the Fmoc-amino acid in a small volume of DMF. Add 3 equivalents of HBTU and 6 equivalents of DIEA, and let the mixture activate with occasional mixing for about 5 minutes. Add the activated solution to the drained resin and agitate for about 1 hour. Drain and wash the resin with 3 × 1 mL of DMF.
| Step | Reagent | Volume for 100 mg resin | Time |
|---|---|---|---|
| Swell | DMF | 1 mL | 30 min |
| Deprotect 1 | 20% piperidine in DMF | 1 mL | 5 min |
| Deprotect 2 | 20% piperidine in DMF, fresh | 1 mL | 15 min |
| Wash | DMF | 3 × 1 mL | brief |
| Couple | Fmoc-AA 3 equiv , HBTU 3 equiv , DIEA 6 equiv in DMF | 1 mL | 1 hour after 5 min activation |
| Wash | DMF | 3 × 1 mL | brief |
Equivalents are relative to resin loading. At 100 mg of 0.6 to 0.8 mmol/g resin, 3 equivalents of amino acid means roughly 0.2 mmol per coupling; multiply by the Fmoc-amino acid molecular weight to find the required mass. The same molar basis applies to HBTU at 3 equivalents and DIEA at 6 equivalents. The excess is intentional: coupling kinetics depend on reagent concentration, and unreacted material is washed away.
During coupling, check that the cartridge is not leaking and that the resin stays suspended in the solution. If the resin beds down and stays down, the agitation is too gentle or the vessel is the wrong size. It is better to couple longer than to end the step while free amine remains, because a failed coupling propagates through every later round.
Repeat the pair of operations until the sequence is assembled. At larger scales, cap the cartridge and shake it during washes. Some protocols alternate DMF, methanol, and dichloromethane for washes; methanol shrinks the resin beads and is optional. After the final coupling, wash the resin with dichloromethane and let the residual solvent evaporate so the resin is dry before cleavage.
The Kaiser test is a qualitative colorimetric test for free primary amines. Ninhydrin reacts with free amine to produce Ruhemann's purple, a deep blue-purple color. In SPPS it is used after a coupling: strong color means free amine remains, so the coupling is incomplete; a pale yellow result means no detectable free amine. Run the sample against a reference tube containing the reagents but no resin, which controls for background color from the reagents themselves, and apply gentle heating. The three solutions are simple to prepare.
| Solution | Composition |
|---|---|
| A | 66% w/v potassium cyanide in water, diluted 1/50 into pyridine 1 mL of the aqueous KCN stock in 49 mL of pyridine |
| B | 5% w/v ninhydrin in butanol |
| C | 200% w/v phenol in butanol |
Add 3 drops of each solution to the sample and the reference tube, heat gently, and compare. Potassium cyanide and phenol accelerate the color development. That acceleration was characterized in photometric ninhydrin studies from 1953 Troll and Cannan, J. Biol. Chem. 200, 803, 811 and 1954 Moore and Stein, J. Biol. Chem. 211, 907, 913 . The color test as used in peptide synthesis was published by Kaiser et al. in 1970 Anal. Biochem. 34, 595, 598 , and quantitative ninhydrin monitoring of solid-phase synthesis by Sarin et al. in 1981 Anal. Biochem. 117, 147, 157 .
Kaiser results are qualitative. A positive test means free amine is present; it cannot distinguish a coupling that is simply slow from one that has stalled, and it does not identify which sequence failed. Treat a positive result after coupling as a signal to repeat the coupling or extend the reaction time. The test detects primary amines; an N-terminal proline, which is a secondary amine, gives an anomalous result and cannot be monitored this way.
Before cleavage, wash the resin with dichloromethane and dry it thoroughly. Make the cleavage cocktail fresh. For Rink amide resin, and only for peptides that contain neither cysteine nor methionine, the standard cocktail is 95:2.5:2.5 by volume TFA:water:TIPS. Water and TIPS are scavengers for the carbocations released when side-chain protecting groups are removed. Cysteine and methionine require different scavenger systems, so this cocktail does not cover them.
A test cleavage on a few beads is always advisable. Take a small sample of resin, add 200 to 500 µL of cleavage cocktail, and agitate gently for about 30 minutes. Precipitate the crude peptide in about 1 mL of cold ether, then dissolve the pellet in 200 µL of 1:1 acetonitrile:water for LCMS analysis. Small peptides can remain soluble in diethyl ether, so do not discard the ether layer until LCMS confirms where the peptide is.
Full cleavage uses about 4 mL of cocktail on the dry resin with gentle agitation for about 2 hours. Filter the solution into a 50 mL falcon tube and add about 30 mL of cold ether to precipitate the peptide. The ether should be kept at -20°C for more than 2 hours before use. Centrifuge for 5 minutes to pellet the peptide, decant the ether carefully, and redissolve the pellet in 1:1 acetonitrile:water. Lyophilize for a crude weight, or load the solution directly onto the HPLC.
| Step | Test cleavage | Full cleavage |
|---|---|---|
| Cocktail volume | 200 to 500 µL | 4 mL |
| Agitation time | 30 min | 2 hours |
| Precipitation | 1 mL cold ether | 30 mL cold ether in a 50 mL tube |
| Workup | dissolve pellet in 200 µL 1:1 MeCN:H2O for LCMS | centrifuge 5 min, redissolve in 1:1 MeCN:H2O |
Cleavage is the most hazardous step. Prepare the cocktail in a fume hood, add it to the resin with the vessel capped, and keep the vessel closed during agitation. After filtration, add the TFA filtrate to the cold ether for precipitation, cap the tube during centrifugation, and open it only inside the hood, because residual TFA and volatile scavengers remain in the ether.
Crude peptide from TFA cleavage and ether precipitation is a mixture: full-length product, deletion peptides, truncated byproducts from failed couplings, and residual scavengers. The standard purification is preparative reverse-phase HPLC on a C18 column , using a gradient of acetonitrile in water. The purification guidance available for this protocol specifies the C18 column and the acetonitrile/water gradient but does not give the exact mobile-phase modification or gradient slope, so those conditions should be optimized for each peptide and column. Fractions eluted during a preparative run are checked against the analytical LCMS of the crude material before pooling, and the analytical conditions are the usual starting point for the preparative gradient.
Two problems recur. The first is UV-active cleavage scavengers such as phenol or thioanisole, which are difficult to remove completely during ether precipitation and appear as extra peaks in HPLC chromatograms. The second is solubility: large, hydrophobic, or aggregation-prone peptides may not stay dissolved in water and acetonitrile long enough to be purified. If a peptide will not dissolve or crashes out during the run, the dissolution solvent and loading conditions must be changed before the preparative run is attempted, not during it.
Purified yield expectations follow from the same economics as the synthesis: from the 100 mg resin example, the 50 mg of crude peptide yields 15 to 25 mg of pure peptide, assuming the typical 30 to 40% recovery through purification. After lyophilization, weigh the crude peptide and store it dry until purification.
The reagents in this protocol are not benign. HBTU is a sensitizer and allergen, and repeated exposure has been associated with anaphylaxis. Trifluoroacetic acid is corrosive, toxic, and reactive. Thiol scavengers such as ethanedithiol, which appear in cleavage cocktails for cysteine-containing peptides, are intensely malodorous. Piperidine, pyridine, and diethyl ether carry their own hazards. Work in a fume hood, use appropriate gloves and eye protection, and treat all waste as hazardous. Dispose of piperidine and TFA waste separately from chlorinated and non-chlorinated solvent waste, following the rules of your institution.
This protocol is intended for researchers who have already trained in organic chemistry laboratory safety. It is not a substitute for institutional safety policies, and the American Peptide Society disclaims liability for actions taken by users of such information.
The limits of this protocol should be stated plainly. The cleavage cocktail applies only to Rink amide resin and only to peptides without cysteine or methionine; cleavage chemistry must be adapted to the resin, linker, and sequence in use. Magnetic stirring is never acceptable. The vacuum manifold caveats, the ether solubility trap for small peptides, and the UV-active scavenger problem are all real and all easy to rediscover the hard way.
Several questions remain open in the wider practice of Fmoc SPPS. The exact preparative HPLC conditions behind this protocol were not specified in the available guidance. Alternative cleavage cocktails for cysteine- or methionine-containing peptides are not covered here. Greener solvents to replace DMF and dichloromethane are an active area of work. Difficult couplings, where the sequence is sterically hindered or prone to aggregation, have specific remedies that fall outside a beginner protocol. And large, hydrophobic, aggregation-prone peptides need special handling during dissolution and purification. Each of these is a separate problem, and each is best approached after the basic cycle in this protocol is running reliably.
Vendors referenced: Pure Peptide.
Related reading: API Contract Manufacturing: Process, Benefits, and Partner Selection, Peptide CoA: What HPLC Purity and LC-MS Actually Prove, Synthesizing Peptides Over 100 Amino Acids: Methods and Examples, Tag-Assisted Peptide Synthesis: How TAPS Works and Its Benefits.