What Is Native Chemical Ligation? Joining Peptide Fragments Into Proteins

Published October 2026 · Patriot Labs Research Team · ~18 min read

The short answer

Native chemical ligation is a reaction that joins two unprotected peptides into one. One piece ends in a thioester; the other begins with a cysteine. When they meet in neutral water, the cysteine’s sulfur captures the thioester and then rearranges, leaving an ordinary peptide bond at the junction. The result is a single, longer chain with a normal amide bond where the two pieces met — indistinguishable from a chain that had been made in one go.

What makes the reaction so useful is that neither peptide has to be protected. The two pieces can carry every one of their normal side chains — acids, bases, alcohols, even other cysteines — and still only react at the one intended spot. That selectivity, in plain water near neutral pH, is what let chemists stop building proteins atom by atom and start building them in large blocks instead.

Native chemical ligation was introduced in 1994 by Philip Dawson, Tom Muir, Ian Clark-Lewis and Stephen Kent.^[2] In the three decades since, it has become the central reaction of chemical protein synthesis: the standard way to assemble a protein, or a large modified peptide, from pieces that a synthesizer can make. The rest of this guide takes the reaction apart — why it was needed, how it works, and how it has been pushed well beyond its original limits.

Why joining peptides is the hard part

Almost every synthetic peptide is made by solid-phase peptide synthesis (SPPS), which adds one amino acid at a time to a chain anchored on a resin bead. Each addition is efficient but not perfect, and the tiny fraction that fails at each step compounds. By the time a chain reaches fifty residues or so, the accumulated deletions and side products make the target peptide a smaller and smaller share of the crude mixture, and purifying it away from close-in-size impurities becomes the limiting problem. In practice, that puts a soft ceiling of roughly fifty amino acids on what a single SPPS run can deliver in good purity.^[13]

Proteins are much longer than that — a small enzyme runs to a hundred residues or more. The obvious idea is to make the protein in pieces and join the pieces, a strategy called segment or fragment condensation. The trouble is the join itself. A peptide is a string of amino acids, each carrying reactive side chains, and simply activating one piece’s C-terminus to couple it to another piece’s N-terminus invites reactions everywhere at once: side-chain acids and amines compete, and the activated end is prone to racemization, scrambling the stereochemistry at the junction. The classical answer was to keep every side chain protected during the coupling, but fully protected peptide segments are notoriously insoluble and hard to handle, and the approach never became general.^[13]

What was needed was a way to join two unprotected peptides — side chains and all — so that the reaction happened at the junction and nowhere else, cleanly, in water. That is exactly the problem native chemical ligation solved.

From a 1953 clue to a 1994 reaction

The chemistry at the heart of native chemical ligation was glimpsed four decades before it was turned into a tool. In 1953, Theodor Wieland and coworkers reported that a valine thioester, mixed with cysteine in water, gave the dipeptide valyl-cysteine — and that the reaction ran through a transient bond between the thioester and the cysteine’s sulfur before settling into a peptide bond.^[3] That sulfur-to-nitrogen hand-off is the same step NCL relies on, but at the time it was a curiosity about a single dipeptide, not a method for proteins.

The idea of joining whole unprotected peptides arrived with Kent’s group in 1992. In building a version of HIV-1 protease, Martina Schnölzer and Stephen Kent coupled two unprotected synthetic peptides directly in solution, forming the protein by what they called “chemical ligation.”^[1] The join worked and the protein folded, but the bond at the junction was not a natural peptide bond — it was a non-native linkage that left a chemical scar in the backbone. The method proved that unprotected segments could be joined chemoselectively; it just did not yet give a native product.

The last step was to make the junction a real peptide bond. In 1994, Dawson, Muir, Clark-Lewis and Kent combined the chemoselective thioester capture with Wieland’s acyl shift, so that the intermediate rearranged into a native amide bond at the join. They named the reaction native chemical ligation and used it to assemble a functional protein from two synthetic pieces.^[2] The combination — unprotected peptides, water, neutral pH, and a scar-free bond — is what made it general.

How the reaction works, step by step

Native chemical ligation needs two chemical handles, one on each peptide: a thioester at the C-terminus of the first piece, and an N-terminal cysteine at the start of the second. Everything else about the two peptides can be ordinary and unprotected. The reaction then proceeds in two distinct steps.^[2][15]

Step one: transthioesterification. The thiol on the N-terminal cysteine’s side chain, present as its reactive ionized form in neutral buffer, attacks the thioester carbonyl of the first peptide. This swaps one thioester for another: the leaving thiol departs, and the two peptides are now tethered through a new thioester that links the first peptide’s C-terminus to the cysteine’s sulfur. This step is reversible — it can run backward just as easily — which turns out to be essential, because it lets the reaction correct any unproductive captures rather than getting stuck.

Step two: the S-to-N acyl shift. Once the two peptides are tethered, the free α-amino group of that same N-terminal cysteine sits just five atoms away from the new thioester carbonyl. It swings in through a five-membered ring transition state and takes over the acyl group, converting the thioester into an amide. The cysteine’s side-chain thiol is handed back, free and unchanged, and what remains is a native peptide bond joining the two pieces — with a cysteine as the first residue of the second segment. This rearrangement is fast and, unlike step one, essentially irreversible, so it continually drains product out of the reversible first step and drives the whole process to completion.^[4]

Two features of this mechanism explain why the reaction is so clean. First, it is chemoselective: only a cysteine at the very start of a peptide has both a thiol and an adjacent α-amine arranged to close the ring and make the amide. A cysteine buried anywhere else in either chain can still capture the thioester in step one, but it has no neighbouring α-amine to complete the shift, so that capture simply reverses and the internal cysteine is left untouched.^[2] Second, it runs under mild, aqueous conditions — typically a neutral, pH 7 buffer at room temperature, with a denaturant such as guanidine hydrochloride added to keep the unfolded fragments soluble.^[15] No side chains need protecting, and acids, bases, alcohols and amides elsewhere on the peptides are all spectators.

The thiol catalyst that makes it practical

A ligation runs faster and cleaner with a small thiol additive in the buffer, and in practice almost every reaction includes one. The additive acts as a catalyst: it exchanges with the starting thioester to generate a more reactive aryl thioester in situ, which the cysteine then captures more readily, and because the whole first step is reversible, the catalyst keeps the system turning over toward product rather than toward dead ends.^[4]

The early additives were small aromatic thiols such as thiophenol and benzyl mercaptan. They worked, but they are volatile and foul-smelling, which made the reaction unpleasant at scale. The modern standard is 4-mercaptophenylacetic acid (MPAA), introduced through mechanistic work by Johnson and Kent in 2006: it is water-soluble, far less odorous, and an effective catalyst, and it has become the default thiol for routine ligations.^[4] The choice of catalyst, its concentration and the buffer pH are the main levers a chemist turns to get a sluggish junction to react in reasonable time.

Making the thioester half

The N-terminal cysteine half of a ligation is easy: cysteine is a standard amino acid, so a synthesizer simply puts it at the start of the second peptide. The thioester half is the harder one to prepare, and much of the method development in native chemical ligation has gone into making thioesters conveniently.

The original route used Boc-chemistry SPPS, which can deliver a C-terminal thioester directly because its strongly acidic deprotection and cleavage steps do not harm the thioester.^[13] The difficulty is that the field as a whole moved to the milder Fmoc chemistry, which removes each protecting group with a base — and thioesters do not survive repeated exposure to base. So a plain Fmoc synthesis cannot carry a thioester through to the end.^[5]

The workaround is to build a stable precursor during the Fmoc synthesis and convert it to the thioester only afterward. Several such surrogates are now routine:

• N-acyl-urea (Nbz) linkers. Blanco-Canosa and Dawson introduced an Fmoc-compatible 3,4-diaminobenzoyl handle that survives the synthesis and is activated to an acyl donor that reacts like a thioester under ligation conditions.^[5]
• Peptide hydrazides. A C-terminal hydrazide, made easily by Fmoc SPPS, is oxidized just before ligation to a reactive acyl species that is trapped by the thiol catalyst to form the thioester in situ.^[15]
• N,S-acyl shift devices. Groups such as bis(2-sulfanylethyl)amido (SEA) are stable during synthesis but rearrange on demand to expose a C-terminal thioester, so a SEA peptide can be ligated to a cysteine peptide directly.^[6]

All of these end at the same place — a reactive thioester or its equivalent at the moment of ligation — and the choice among them is a matter of which is easiest to make for a given sequence.

Ligating where there is no cysteine

The one real constraint on the original reaction is that it leaves a cysteine at the junction, and cysteine is one of the rarer amino acids — many target proteins simply do not have one where a convenient break in the chain falls. Two ideas removed that constraint and are now central to how the method is used.

The first is desulfurization. In 2001 Yan and Dawson showed that after a normal ligation at cysteine, the extra sulfur can be chemically stripped off, converting the cysteine into an alanine.^[7] Alanine is common, so a chemist can plan a disconnection at an alanine, install a temporary cysteine there to run the ligation, and then desulfurize to leave the intended alanine with no trace of the join. The step became far more practical in 2007, when Wan and Danishefsky introduced a metal-free, radical-based desulfurization that is gentle and selective; any cysteines meant to survive in the final protein are simply protected so they are not touched.^[8]

The second idea generalizes the first. Chemists have made “thiol-derivatized” versions of many ordinary amino acids — valine, threonine, leucine, phenylalanine, lysine and others — each carrying a temporary thiol on its side chain in just the right place to drive the acyl shift. A ligation at one of these behaves like a cysteine ligation, and a final desulfurization returns the normal residue.^[15] Taken together, these tricks mean a junction can be placed at a wide range of positions, not only at the handful of cysteines a sequence happens to contain. A related variant uses selenocysteine in place of cysteine, which ligates and is then “deselenized” by analogous chemistry.^[9]

Expressed protein ligation: meeting biology halfway

Native chemical ligation does not care whether a peptide was made by a chemist or by a cell — it only needs the two handles. That observation led, in 1998, to expressed protein ligation (EPL), reported by Muir, Sondhi and Cole.^[10] EPL supplies the thioester half from a living organism rather than from a synthesizer.

The trick is a natural protein element called an intein, which normally splices itself out of a host protein. Engineered versions can be arrested midway through their chemistry so that, instead of splicing, they release the protein attached to them as a C-terminal thioester. That recombinant thioester — a full protein domain grown in bacteria — is then ligated to a synthetic peptide bearing an N-terminal cysteine, in exactly the reaction described above.^[10] The reverse pairing works too: a recombinant protein can be made with an N-terminal cysteine and joined to a synthetic thioester.

The point of EPL is reach. A cell can make a large protein easily but cannot install a precise chemical modification at a chosen spot; a synthesizer can make a small, exactly modified segment but not a whole large protein. EPL joins the two, letting a short synthetic piece — carrying a label, a specific modification, or a non-natural building block — be welded onto a large, cheaply expressed protein. It sits on the boundary between the two production routes described in recombinant vs synthetic peptides.

What native chemical ligation has built

Native chemical ligation is the reason total chemical synthesis of proteins is a routine research activity rather than a heroic exception. By joining several synthetic segments in sequence, it gives access to proteins and large modified peptides that are out of reach for a single synthesis run — and because the pieces are synthetic, the chemist controls every atom.^[12] A few families of use stand out.

Whole proteins and enzymes. The method is generally described as the most effective way to make native or modified proteins up to roughly 300 amino acids, usually by stringing together two or more fragments.^[12] One-pot sequential and kinetically controlled ligations let several segments be joined in a defined order in a single vessel, which is how longer targets are assembled.

Mirror-image proteins. Because the fragments are built from scratch, they can be made entirely from D-amino acids, the mirror images of the natural L forms, and then ligated into a mirror-image protein. Kent’s group demonstrated the principle by chemically synthesising both the natural L and the mirror-image D forms of HIV-1 protease and showing that each enzyme acted only on the matching mirror-image substrate.^[11] Mirror-image proteins are now used as tools in drug discovery.

Precisely modified proteins. A synthetic segment can carry a modification installed exactly where the chemist wants it — a phosphate (see phosphorylation), a sugar (see glycosylation), an isotope label or a non-coded residue. Ligating that segment into the rest of the protein yields a single, homogeneous modified species, which is far harder to obtain from biological expression alone.^[14] The same ability to form a bond between a peptide’s two ends also makes NCL a route to backbone-cyclized peptides (see cyclic peptides).

These are research applications — building and studying proteins. Most catalogue research peptides are short enough to come straight off a synthesizer and never need ligation; native chemical ligation belongs to the harder problem of assembling whole proteins. Whatever route makes a peptide, its identity and purity still have to be shown once it is finished, by the HPLC and mass-spectrometry testing described in understanding peptide COA testing.

What this guide does not establish

This guide explains a laboratory reaction used to build proteins. It is chemistry, and several limits are worth stating plainly.

• The reaction needs a thiol junction. Classic NCL requires a cysteine at the join; reaching other positions depends on desulfurization or thiol-derivatized residues, which add steps and their own constraints.^[7][8][15]
• Thioesters are delicate. They can hydrolyze under the aqueous conditions, and activating a peptide’s C-terminus as a thioester carries a risk of racemizing the final residue, which is part of why junctions are often chosen at glycine or other positions that minimize that risk.^[15]
• There is still a size ceiling. “Up to roughly 300 amino acids” is a practical guide, not a law; larger proteins usually require expressed protein ligation or other hybrid strategies.^[12]
• This is a synthesis method, not a biological process. NCL describes how proteins are made on the bench; it is not how the body makes peptides, which is covered in how peptides are made.
• Nothing here describes any use in a person or animal. The chemistry is described to explain how peptides and proteins are assembled in the laboratory, not what any compound does, and this guide does not describe the synthesis or manufacture of any specific product.

Frequently asked questions

What is native chemical ligation?

Native chemical ligation (NCL) is a reaction that joins two unprotected peptides into a single chain. One peptide ends in a thioester and the other begins with a cysteine; when they are mixed in neutral, water-based buffer, the cysteine’s sulfur attacks the thioester and the intermediate rearranges to leave an ordinary peptide (amide) bond at the junction. It is the central reaction of chemical protein synthesis, introduced by Dawson, Muir, Clark-Lewis and Kent in 1994.

How does native chemical ligation work?

In two steps. First, the thiol of the N-terminal cysteine on one peptide attacks the C-terminal thioester of the other, swapping one thioester for another and tethering the two peptides together; this step is reversible. Second, the free α-amino group of that same cysteine is now held next to the new thioester and undergoes a spontaneous sulfur-to-nitrogen acyl shift, forming a stable amide bond and freeing the sulfur again. The amide step is effectively irreversible, which pulls the whole reaction forward.

Why does native chemical ligation need a cysteine?

Because the reaction is driven by a cysteine’s two neighbouring groups: its side-chain thiol captures the thioester, and its α-amino group then closes a five-membered ring to make the amide. Only a cysteine sitting at the very start of a peptide has both groups positioned to do this, which is also why internal cysteines elsewhere in the chain do not interfere — they can grab the thioester briefly but cannot complete the rearrangement, so they let go again.

What is the difference between native chemical ligation and expressed protein ligation?

Expressed protein ligation (EPL) is native chemical ligation applied to a piece made by living cells. Instead of synthesising the thioester chemically, EPL uses a self-splicing protein element called an intein to produce a recombinant protein that carries a C-terminal thioester. That recombinant piece is then ligated to a synthetic cysteine peptide exactly as in NCL. EPL lets a small, precisely modified synthetic segment be attached to a large protein grown in bacteria.

Can native chemical ligation work without a cysteine?

Yes, indirectly. Chemists run the ligation at a cysteine as usual and then remove the extra sulfur afterward in a step called desulfurization, which converts the cysteine into an alanine and leaves no trace of the join. Related chemistry uses specially made amino acids — valine, threonine, leucine, phenylalanine and others — that carry a temporary thiol handle for the ligation and are desulfurized back to the normal residue, so a join can be placed at many different positions.

What is the largest protein that can be made by native chemical ligation?

There is no single hard limit, but native chemical ligation is generally described as the most effective way to chemically make proteins up to roughly 300 amino acids, usually by joining several synthetic segments one after another. Larger targets are reached by combining the method with recombinant protein pieces through expressed protein ligation.

Who invented native chemical ligation?

It was reported in 1994 by Philip Dawson, Tom Muir, Ian Clark-Lewis and Stephen Kent at The Scripps Research Institute, building on an earlier non-native “chemical ligation” method from Kent’s group in 1992 and on a sulfur-to-nitrogen acyl shift that Theodor Wieland and coworkers had observed in 1953.

Is native chemical ligation used to make research peptides?

Most catalogue research peptides are short — a few dozen residues at most — and come directly off a solid-phase synthesizer without any ligation. Native chemical ligation belongs to the harder job of assembling whole proteins and large modified peptides from several fragments. It is a protein-synthesis research technique rather than a routine step in making the short peptides sold for laboratory research.

References

1. Schnölzer, M., & Kent, S. B. H. (1992). Constructing proteins by dovetailing unprotected synthetic peptides: backbone-engineered HIV protease. Science, 256(5054), 221–225. doi:10.1126/science.1566069
2. Dawson, P. E., Muir, T. W., Clark-Lewis, I., & Kent, S. B. H. (1994). Synthesis of proteins by native chemical ligation. Science, 266(5186), 776–779. doi:10.1126/science.7973629; PMID 7973629
3. Wieland, T., Bokelmann, E., Bauer, L., Lang, H. U., & Lau, H. (1953). Über Peptidsynthesen. 8. Mitteilung: Bildung von S-haltigen Peptiden bei der intramolekularen Umlagerung von Aminoacyl-mercaptanen. Justus Liebigs Annalen der Chemie, 583, 129–149. doi:10.1002/jlac.19535830110
4. Johnson, E. C. B., & Kent, S. B. H. (2006). Insights into the mechanism and catalysis of the native chemical ligation reaction. Journal of the American Chemical Society, 128(20), 6640–6646. doi:10.1021/ja058344i
5. Blanco-Canosa, J. B., & Dawson, P. E. (2008). An efficient Fmoc-SPPS approach for the generation of thioester peptide precursors for use in native chemical ligation. Angewandte Chemie International Edition, 47(36), 6851–6855. doi:10.1002/anie.200705471
6. Ollivier, N., Dheur, J., Mhidia, R., Blanpain, A., & Melnyk, O. (2010). Bis(2-sulfanylethyl)amino native peptide ligation. Organic Letters, 12(22), 5238–5241. doi:10.1021/ol102273u; PMID 20964289
7. Yan, L. Z., & Dawson, P. E. (2001). Synthesis of peptides and proteins without cysteine residues by native chemical ligation combined with desulfurization. Journal of the American Chemical Society, 123(4), 526–533. doi:10.1021/ja003265m
8. Wan, Q., & Danishefsky, S. J. (2007). Free-radical-based, specific desulfurization of cysteine: a powerful advance in the synthesis of polypeptides and glycopolypeptides. Angewandte Chemie International Edition, 46(48), 9248–9252. doi:10.1002/anie.200704195
9. McGrath, N. A., & Raines, R. T. (2011). Chemoselectivity in chemical biology: acyl transfer reactions with sulfur and selenium. Accounts of Chemical Research, 44(9), 752–761. doi:10.1021/ar200081s; PMID 21639109
10. Muir, T. W., Sondhi, D., & Cole, P. A. (1998). Expressed protein ligation: a general method for protein engineering. Proceedings of the National Academy of Sciences USA, 95(12), 6705–6710. doi:10.1073/pnas.95.12.6705; PMID 9618476
11. Milton, R. C. deL., Milton, S. C. F., & Kent, S. B. H. (1992). Total chemical synthesis of a D-enzyme: the enantiomers of HIV-1 protease show reciprocal chiral substrate specificity. Science, 256(5062), 1445–1448. doi:10.1126/science.1604320
12. Kent, S. B. H. (2009). Total chemical synthesis of proteins. Chemical Society Reviews, 38(2), 338–351. doi:10.1039/b700141j; PMID 19169452
13. Nilsson, B. L., Soellner, M. B., & Raines, R. T. (2005). Chemical synthesis of proteins. Annual Review of Biophysics and Biomolecular Structure, 34, 91–118. doi:10.1146/annurev.biophys.34.040204.144700; PMID 15869385
14. Conibear, A. C., Watson, E. E., Payne, R. J., & Becker, C. F. W. (2018). Native chemical ligation in protein synthesis and semi-synthesis. Chemical Society Reviews, 47(24), 9046–9068. doi:10.1039/c8cs00573g; PMID 30418441
15. Agouridas, V., El Mahdi, O., Diemer, V., Cargoët, M., Monbaliu, J.-M., & Melnyk, O. (2019). Native chemical ligation and extended methods: mechanisms, catalysis, scope, and limitations. Chemical Reviews, 119(12), 7328–7443. doi:10.1021/acs.chemrev.8b00712; PMID 31050890

For more on the chemistry this guide touches, see What Is Solid-Phase Peptide Synthesis? for how the fragments are built, Recombinant vs Synthetic Peptides for the two production routes expressed protein ligation bridges, What Is a Peptide Bond? for the bond the reaction forms, What Is a Disulfide Bond? for the cysteine chemistry it depends on, and How Peptides Are Made for the wider manufacturing picture.

All Patriot Labs products are sold strictly for in-vitro research and laboratory use only. Not for human or veterinary consumption. This guide is educational and describes a laboratory reaction used to synthesize peptides and proteins; it is not medical advice, does not describe how to use any product, and makes no claim about the synthesis or manufacturing method of any specific product.