Peptide Bond: Formation, Structure and Hydrolysis

A peptide bond, also called a peptide linkage, is the amide bond (-CO-NH-) that joins the carboxyl carbon of one amino acid to the amino nitrogen of the next. It forms by a condensation reaction, often called dehydration synthesis, that releases one molecule of water (18.01 Da) per bond. Resonance gives the C-N bond partial double-bond character, so it is short (about 1.33 Å), flat and slow to rotate. It is almost always trans, except in front of proline, and it is slow to break in neutral water without a catalyst.

This article goes deeper on the bond itself: how it forms, why it is planar, how it breaks, how chemists make it in the laboratory and how it shows up in an analysis. For how bonds add up into a chain and a fold, see our article on peptide structure.

Peptide bond at a glance

Source: standard bond geometry for amides; IR band ranges per references [12, 13]; fragment nomenclature per references [14, 15].

What is a peptide bond?

Every amino acid carries an amino group and a carboxyl group on the same alpha carbon (see what an amino acid is). When the carboxyl group of one amino acid reacts with the amino group of another, the product is an amide: the carbonyl carbon is now bonded directly to nitrogen. Chemists call this specific amide a peptide bond because it is the link that repeats along every peptide and protein backbone.

It is a covalent bond, and in nearly every residue a secondary amide: the nitrogen carries one hydrogen and two carbon groups. Proline is the exception. Its side chain is bonded to its own nitrogen, so the bond in front of proline is a tertiary amide with no N-H, which changes how it behaves.

Peptide bond formation: dehydration synthesis

Peptide bond formation is a condensation reaction. The carboxyl group loses -OH, the amino group loses -H, and together they leave as water. Biology textbooks call the same reaction dehydration synthesis, the general name for building a larger molecule by removing water.

The water leaves a mark on the mass. Two glycine molecules weigh 75.0320 Da each (monoisotopic). Glycylglycine, the dipeptide, is 132.0535 Da: the sum of the two (150.0640 Da) minus one water (18.0106 Da). Every additional bond removes another 18.0106 Da, which is why a peptide’s mass is calculated from residue masses plus one water for the two free ends.

Why the reaction does not run on its own

Mixing two amino acids in water does not give a dipeptide in useful amounts. In neutral water the amino group is protonated (-NH3+) and the carboxyl group ionised (-COO-), so the nitrogen has no free lone pair to attack with. The equilibrium also lies on the side of the free amino acids, because water is present in vast excess.

Every practical route therefore activates the carboxyl group first. Cells use aminoacyl-tRNA esters. Nissen and colleagues solved structures of the large ribosomal subunit bound to substrate analogs and reported that only conserved ribosomal RNA, not protein, contacts the substrates at the site where the bond is made. No protein side-chain atom came closer than about 18 Å, which established the ribosome as a ribozyme [1]. Chemists do the same job with coupling reagents, covered below.

Resonance and planarity of the peptide bond

The nitrogen’s lone pair is shared with the neighbouring carbonyl group. The bond can be drawn as two resonance forms: a C-N single bond with a C=O double bond, and a C=N double bond with a negative charge on oxygen and a positive charge on nitrogen. The real bond sits between the two. Four practical consequences follow:

• It is short. At about 1.33 Å the C-N bond is noticeably shorter than the N-Cα single bond next to it (about 1.46 Å).
• It is flat. The carbonyl carbon, oxygen, nitrogen, amide hydrogen and the two flanking alpha carbons lie close to one plane. Pauling, Corey and Branson built this planar amide into their 1951 models of hydrogen-bonded helical configurations of the polypeptide chain [2].
• It resists rotation. Turning the C-N bond breaks the overlap that gives it double-bond character. For simple amides the rotation barrier is typically quoted at roughly 75 to 90 kJ/mol, high enough that rotation is slow at room temperature.
• It is polar. The oxygen carries partial negative charge and the N-H partial positive charge, so the group is both a hydrogen-bond acceptor (C=O) and donor (N-H). Backbone hydrogen bonds between these groups are what hold helices and sheets together.

Resonance is also why a backbone amide nitrogen is not basic: its lone pair is tied up, so it does not pick up a proton in water the way a free amine does.

Trans and cis peptide bonds, and why proline is different

A planar bond can still be arranged two ways. In the trans form the two alpha carbons sit on opposite sides of the C-N bond. In the cis form they sit on the same side and crowd each other. For an ordinary secondary amide, trans wins by a wide margin. Zhang and Germann measured cis bonds directly in short glycine- and isoleucine-containing oligopeptides by two-dimensional NMR exchange spectroscopy in DMSO and found 0.13 to 0.23% cis at 296 K, with trans-to-cis rate constants of 4 to 9 × 10-3 s-1 [3].

Proline changes the balance. Because its nitrogen carries a ring carbon instead of a hydrogen, the cis and trans forms are much closer in energy, and a few percent to a few tens of percent cis is common in short proline peptides in water. The switch is slow. Reimer and colleagues studied the pentapeptide series Ac-Ala-Xaa-Pro-Ala-Lys-NH2, with every standard amino acid in the Xaa position, by NMR and CD [4]. Both isomers were unstructured in water, so the measurements reflected the residue in front of proline alone. Cis-to-trans rate constants at 4 °C ranged from 3.2 × 10-3 s-1 (Xaa = Ala) to 0.5 × 10-3 s-1 (Xaa = Trp). Tyrosine and histidine slowed the switch severalfold when their side chains were deprotonated [4].

Those rates mean half-lives of minutes at low temperature. A proline-containing peptide can therefore show two sets of NMR signals, one per isomer, and occasionally a broadened or split chromatography peak when the two forms separate faster than they interconvert.

Hydrolysis: breaking a peptide bond

Hydrolysis is the reverse of condensation: water adds across the C-N bond and splits it back into a carboxyl group and an amine. Thermodynamically, hydrolysis is favoured in water. Kinetically, it is very slow at neutral pH and room temperature, which is why peptide chains are stable enough to purify, freeze-dry and analyse.

Measuring the uncatalysed rate is itself a research topic. Wolfenden’s review of benchmark reaction rates covers the spontaneous cleavage of C-N and other bonds in water and the uncatalysed reaction that corresponds to peptidyl transfer in the ribosome. Across these reactions, rates span about 19 orders of magnitude, with some half-lives above a million years, and the slowest reactions are the most sensitive to heat, with rates rising as much as 10⁷-fold between 25 and 100 °C [5]. Wolfenden and Snider used the same comparison to measure how proficient enzymes are as catalysts [6].

Laboratories use that heat sensitivity on purpose. Three routes are common:

• Acid hydrolysis. Classical amino acid analysis heats a sample in strong hydrochloric acid, typically 6 M at about 110 °C for around 24 hours, then separates and counts the free amino acids. Fountoulakis and Lahm describe this two-step method and faster microwave-assisted hydrolysis [7].
• Selective chemical cleavage. Bonds after aspartic acid, especially Asp-Pro, are relatively acid-labile.
• Enzymatic cleavage. Proteases cut at defined sites; trypsin cuts after lysine and arginine. Analysing the resulting peptides by LC-MS is the basis of peptide mapping.

How chemists form peptide bonds in the laboratory

Laboratory synthesis must activate the carboxyl group, block the wrong groups from reacting, and keep each alpha carbon’s stereochemistry.

Coupling reagents

Coupling reagents convert the carboxylic acid into a reactive intermediate that an amine attacks to form the amide. The main families are carbodiimides (such as DCC and DIC), usually used with an additive such as HOBt or Oxyma; aminium/uronium salts (such as HBTU and HATU); and phosphonium salts (such as PyBOP). Valeur and Bradley reviewed the choice among these “acronym” reagents for amide bond formation in general [8]. El-Faham and Albericio’s review covers the reagents used specifically for peptides [9].

Protecting groups and solid-phase synthesis

An amino acid has two reactive ends, and many have reactive side chains, so all but one reacting group must be blocked. In the common Fmoc/tBu approach, the alpha-amine carries a base-labile Fmoc group and reactive side chains carry acid-labile groups. Merrifield’s solid-phase method anchors the growing chain to a resin bead, so each coupling is followed by washing rather than purification [10]. The chain grows from the C-terminus toward the N-terminus, one residue at a time, by repeated rounds of deprotection and coupling.

Racemisation and step efficiency

Activation can let the neighbouring alpha carbon lose its proton, often through an oxazolone intermediate, converting some L residue to D. The product has the same mass as the target, so HPLC is read alongside mass data.

Step efficiency compounds quickly. If each coupling reaches 99%, about 83% of 20-residue chains and about 61% of 50-residue chains are full length before purification. At 99.9% per step the 50-residue figure rises to about 95%. So very long chains are often made as segments and joined. In native chemical ligation, two unprotected peptide segments react in water through a thioester-linked intermediate that rearranges to give a native peptide bond at the join [11]. Our article on how peptides are made covers cleavage, purification and freeze-drying.

How the peptide bond shows up in analysis

Ultraviolet absorbance

The amide group absorbs strongly below about 230 nm, so peptides are detected at 214 to 220 nm in reversed-phase HPLC, where every peptide bond contributes to the signal. Our explainer on HPLC peptide purity covers how those peaks are read.

Infrared amide bands

The peptide group gives characteristic infrared bands. The amide I band (about 1600-1700 cm-1) comes mainly from C=O stretching, and the amide II band (about 1510-1580 cm-1) from N-H bending coupled with C-N stretching. Because hydrogen bonding shifts the amide I position, the band shape carries information about secondary structure. Barth’s review of protein infrared spectroscopy covers the mid-infrared region [12], and Kong and Yu describe how amide band positions are correlated with helix, sheet and other structural components [13].

Mass spectrometry fragments

In tandem mass spectrometry, an energised peptide ion breaks mostly at its peptide bonds. Fragments that keep the N-terminus are called b ions and those that keep the C-terminus are called y ions, following the nomenclature proposed by Roepstorff and Fohlman [14]. Reading the mass gaps along each ladder recovers the residue order, as Steen and Mann describe [15].

A worked example with Gly-His-Lys (GHK), using monoisotopic residue masses and a single charge:

Source: calculated by Vortex from monoisotopic residue masses; 1.0073 Da is the mass of a proton. A b ion carries no added water; a y ion carries the C-terminal water.

The gap between y1 and y2 (137.0589 Da) is a histidine residue, read across the His-Lys peptide bond. Our article on what peptides are covers the wider set of identity methods.

Frequently asked questions

What is a peptide bond in simple terms?

A peptide bond is the amide link that joins one amino acid to the next in a peptide or protein. It forms between the carboxyl carbon of one residue and the amino nitrogen of the next, and one water molecule is released for each bond.

Is dehydration synthesis the same as a condensation reaction?

Yes, for peptide bonds the two terms describe the same reaction. Two molecules join and a water molecule is removed. Biology texts tend to say dehydration synthesis, and chemistry texts tend to say condensation.

What type of bond is a peptide bond?

It is a covalent amide bond, specifically a secondary amide in most residues and a tertiary amide in front of proline. Resonance gives the C-N bond partial double-bond character, so the group is planar and rotation around it is restricted.

Why is the peptide bond stable in water?

Hydrolysis is thermodynamically favoured in water, but without a catalyst it is extremely slow at neutral pH and room temperature. Laboratories break peptide bonds deliberately with strong acid and heat, with strong base, or with protease enzymes.

Research materials: Vortex supplies peptides as sealed lyophilised research reagents. Browse the catalogue at /shop/. Published lab reports are at /coa/.

References

1. Nissen P, Hansen J, Ban N, Moore PB, Steitz TA. The structural basis of ribosome activity in peptide bond synthesis. Science. 2000;289(5481):920-30. PMID 10937990. doi:10.1126/science.289.5481.920
2. Pauling L, Corey RB, Branson HR. The structure of proteins; two hydrogen-bonded helical configurations of the polypeptide chain. Proc Natl Acad Sci U S A. 1951;37(4):205-11. PMID 14816373. doi:10.1073/pnas.37.4.205
3. Zhang J, Germann MW. Characterization of secondary amide peptide bond isomerization: thermodynamics and kinetics from 2D NMR spectroscopy. Biopolymers. 2011;95(11):755-62. PMID 21538331. doi:10.1002/bip.21642
4. Reimer U, Scherer G, Drewello M, Kruber S, Schutkowski M, Fischer G. Side-chain effects on peptidyl-prolyl cis/trans isomerisation. J Mol Biol. 1998;279(2):449-60. PMID 9642049. doi:10.1006/jmbi.1998.1770
5. Wolfenden R. Benchmark reaction rates, the stability of biological molecules in water, and the evolution of catalytic power in enzymes. Annu Rev Biochem. 2011;80:645-67. PMID 21495848. doi:10.1146/annurev-biochem-060409-093051
6. Wolfenden R, Snider MJ. The depth of chemical time and the power of enzymes as catalysts. Acc Chem Res. 2001;34(12):938-45. PMID 11747411. doi:10.1021/ar000058i
7. Fountoulakis M, Lahm HW. Hydrolysis and amino acid composition of proteins. J Chromatogr A. 1998;826(2):109-34. PMID 9917165. doi:10.1016/s0021-9673(98)00721-3
8. Valeur E, Bradley M. Amide bond formation: beyond the myth of coupling reagents. Chem Soc Rev. 2009;38(2):606-31. PMID 19169468. doi:10.1039/b701677h
9. El-Faham A, Albericio F. Peptide coupling reagents, more than a letter soup. Chem Rev. 2011;111(11):6557-602. PMID 21866984. doi:10.1021/cr100048w
10. Merrifield B. Solid phase synthesis. Science. 1986;232(4748):341-7. PMID 3961484. doi:10.1126/science.3961484
11. Dawson PE, Muir TW, Clark-Lewis I, Kent SB. Synthesis of proteins by native chemical ligation. Science. 1994;266(5186):776-9. PMID 7973629. doi:10.1126/science.7973629
12. Barth A. Infrared spectroscopy of proteins. Biochim Biophys Acta. 2007;1767(9):1073-101. PMID 17692815. doi:10.1016/j.bbabio.2007.06.004
13. Kong J, Yu S. Fourier transform infrared spectroscopic analysis of protein secondary structures. Acta Biochim Biophys Sin (Shanghai). 2007;39(8):549-59. PMID 17687489. doi:10.1111/j.1745-7270.2007.00320.x
14. Roepstorff P, Fohlman J. Proposal for a common nomenclature for sequence ions in mass spectra of peptides. Biomed Mass Spectrom. 1984;11(11):601. PMID 6525415. doi:10.1002/bms.1200111109
15. Steen H, Mann M. The ABC’s (and XYZ’s) of peptide sequencing. Nat Rev Mol Cell Biol. 2004;5(9):699-711. PMID 15340378. doi:10.1038/nrm1468

Methods note: written by the Vortex Research Editorial Team. Literature was selected from PubMed for chemistry and analytical topics, every PMID was checked against its PubMed record for title, journal and year, and each abstract (where PubMed holds one) was read against the sentence that cites it. Masses and step-efficiency figures were calculated by Vortex.

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