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Coupling of two amino acids in solution. The unprotected amine of one reacts with the unprotected carboxylic acid group of the other to form a peptide bond. The second reactive group (amine/acid) in each of the starting materials bears a protecting group.

In organic chemistry, peptide synthesis is the production of peptides, compounds where multiple amino acids are linked via amide bonds, also known as peptide bonds. Peptides are chemically synthesized by the condensation reaction of the carboxyl group of one amino acid to the amino group of another. Protecting group strategies are usually necessary to prevent undesirable side reactions with the various amino acid side chains.[1] Chemical peptide synthesis most commonly starts at the carboxyl end of the peptide (C-terminus), and proceeds toward the amino-terminus (N-terminus).[2] Protein biosynthesis (long peptides) in living organisms occurs in the opposite direction.

The chemical synthesis of peptides can be carried out using classical solution-phase techniques, although these have been replaced in most research and development settings by solid-phase methods (see below).[citation needed] Solution-phase synthesis retains its usefulness in large-scale production of peptides for industrial purposes however.

Chemical synthesis facilitates the production of peptides which are difficult to express in bacteria, the incorporation of unnatural amino acids, peptide/protein backbone modification, and the synthesis of D-proteins, which consist of D-amino acids.

Solid Peptide Synthesis Reactor


Solid-phase synthesisEdit

Solid-phase synthesis of a dipeptide using an (amine-functionalized) amide resin. The N-terminal protecting group (PG) can be Fmoc or Boc, depending on the protecting group scheme used (see below). The amino acid side chains (R1, R2 etc.) are orthogonally protected (not shown).

The established method for the production of synthetic peptides in the lab is known as solid-phase peptide synthesis (SPPS).[2] Pioneered by Robert Bruce Merrifield,[3][4] SPPS caused a paradigm shift within the peptide synthesis community, and allows the rapid assembly of a peptide chain through successive reactions of amino acids on an insoluble porous support.

The solid support consists of small, polymeric resin beads functionalized with reactive linker groups (such as amine or hydroxyl groups) on which peptide chains can be built.[2] The peptide remains covalently attached to the support throughout the synthesis. This allows for removal of excess reagents and side products to be removed by washing and filtration. This approach circumvents the comparatively time-consuming isolation of the product peptide from solution after each reaction step, which would be required when using conventional solution-phase synthesis.

Each amino acid to be coupled to the peptide chain N-terminus must be protected on its N-terminus and side chain using appropriate protecting groups such as Boc (acid-labile) or Fmoc (base-labile), depending on the side chain and the protection strategy used (see below).[1] This avoids unwanted reaction at those groups during the peptide coupling step, which occurs between the peptide N-terminal amine and the C-terminal carboxylate of the amino acid to be coupled.

The general SPPS procedure is one of repeated cycles of alternate N-terminal deprotection and coupling reactions, with resin washes between each step.[2] In the first step, the first amino acid is coupled to the resin. Subsequently, the amine is deprotected, and then coupled with the free acid of the second amino acid. This cycle repeats until the desired sequence has been synthesized. SPPS cycles may also include capping steps which block the ends of unreacted amino acids from reacting. At the end of the synthesis, the crude peptide is cleaved from the solid support while simultaneously removing all protecting groups using a reagent strong acids like trifluoroacetic acid or a nucleophile.[2] The crude peptide can be precipitated from a non-polar solvent like diethyl ether in order to remove organic soluble by products. The crude peptide can be purified using reversed-phase HPLC.[5] The purification process, especially of longer peptides can be challenging, because small amounts of several byproducts, which are very similar to the product, have to be removed. For this reason so-called continuous chromatography processes such as MCSGP are increasingly being used in commercial settings to maximize the yield without sacrificing on purity levels.[6]

SPPS is limited by reaction yields, and typically peptides and proteins in the range of 70 amino acids are pushing the limits of synthetic accessibility.[2] Synthetic difficulty also is sequence dependent; typically aggregation-prone sequences such as amyloids[7] are difficult to make. Longer lengths can be accessed by using ligation approaches such as native chemical ligation, where two shorter fully deprotected synthetic peptides can be joined together in solution.

Peptide coupling reagentsEdit

An important feature that has enabled the broad application of SPPS is the generation of extremely high yields in the coupling step.[2] To illustrate the impact of suboptimal coupling yields for a given synthesis, consider the case where each coupling step were to have at least 99% yield: this would result in a 77% overall crude yield for a 26-amino acid peptide (assuming 100% yield in each deprotection); if each coupling were 95% efficient, the overall yield would be 25%. Thus, highly optimized amide bond formation conditions are required, employing highly efficient coupling reagents[8][9] and adding an excess of each amino acid (between 2- and 10-fold). The minimization of amino acid racemization during coupling is also of vital importance to avoid epimerization in the final peptide product.

Despite being thermodynamically favorable, direct amide bond formation between an amine and carboxylic acid suffers from a high activation energy, and as such often requires high temperatures. In light of the requirement for highly efficient coupling reactions discussed above, coupling reagents or 'activators' are therefore used for amide bond formation during peptide synthesis. Activation of the carboxyl group of the amino acid to be coupled greatly increases reaction efficiency, often via the formation of a more reactive 'active ester' species in situ. Many peptide coupling reagents exist,[8][9] a selection of which are described below.


Amide bond formation using DIC/HOBt.[9]

Carbodiimides such as dicyclohexylcarbodiimide (DCC) and diisopropylcarbodiimide (DIC) are frequently used for amide bond formation.[9] The reaction proceeds via the formation of a highly reactive O-acylisourea formed by nucleophilic attack of the carboxylate oxygen on the carbodiimide carbon. Subsequent aminolysis of this reactive species by the peptide N-terminal amine forms a peptide bond. Formation of the O-acylisourea is relatively slow and proceeds fastest in non-polar solvents such as dichloromethane. It can be speed up by the use of increased temperature and the presence of excess carbodiimide. [10]

DIC is particularly useful for SPPS as it is easily handled as a liquid, and the urea byproduct formed is soluble in most organic solvents, allowing facile removal during resin washes. Conversely, the related carbodiimide 1-Ethyl-3-(3-dimethylaminopropyl)carbodiimide (EDC) is often used for solution-phase peptide couplings as the urea byproduct in this case is water-soluble, and can therefore be removed easily by washing during aqueous work-up.[9]

Neighbouring group effect of HOAt

A potential disadvantage of carbodiimide activation is the potential for racemization of the activated amino acid.[9] This can be circumvented through the addition of 'racemization suppressing' additives such as the triazoles 1-hydroxy-benzotriazole (HOBt) and 1-hydroxy-7-aza-benzotriazole (HOAt). These work by attacking the O-acylisourea intermediate to form an active ester in situ, which subsequently reacts with the peptide to form the desired amine bond. HOAt is particularly reactive due to a neighbouring group effect involving the pyridyl nitrogen atom.[11]

Ethyl cyanohydroxyiminoacetate (Oxyma) is a more recently developed additive for carbodiimide coupling, and acts as an alternative to the potentially explosive triazole reagents with comparable coupling efficiency to HOAt.[12]

Aminium/uronium and phosphonium saltsEdit

Uronium-based peptide coupling reagents

More recently developed and commonly-used coupling reagents omit the carbodiimide completely and incorporate the HOAt/HOBt moiety as an aminium/uronium or phosphonium salt of a non-nucleophilic anion (tetrafluoroborate or hexafluorophosphate).[8] Examples of aminium/uronium reagents include HATU (HOAt), HBTU/TBTU (HOBt) and HCTU (6-ClHOBt). HBTU and TBTU differ only in the choice of anion. Phosphonium reagents include PyBOP (HOBt) and PyAOP (HOAt).

It should be noted that these reagents form the same active ester species as the carbodiimide activation conditions described above, but differ in the rate of the initial activation step, which is determined by nature of the carbon skeleton of the coupling reagent.[13] Furthermore, aminium/uronium reagents are capable of reacting with the peptide N-terminus to form an inactive guanidino by-product, whereas as phosphonium reagents are not.

COMU is a novel uronium reagent based on Oxyma and incorporating a morpholino group.[14]

Solid supportsEdit

Polystyrene cross-linked with divinylbenzene. This is the most common solid support used in SPPS, and was the support pioneered by R. Bruce Merrifield.

Solid supports for peptide synthesis must be physically stable and permit the rapid filtration of liquids, such as excess reagents, must be inert to all reagents and solvents used during SPPS, must swell extensively in the solvents used to allow for penetration of the reagents, and must allow for the attachment of the first amino acid.[15]

There are three primary types of solid supports: gel-type supports, surface-type supports, and composites.[15] Improvements to solid supports used for peptide synthesis enhance their ability to withstand the repeated use of TFA during the deprotection step of SPPS.[16] Two primary resins are used, based on whether a C-terminal carboxylic acid or amide is desired. The Wang resin was, as of 1996, the most commonly used resin for peptides with C-terminal carboxylic acids.[17]

Protecting groups schemesEdit

As described above, the use of N-terminal and side chain protecting groups is essential during peptide synthesis to avoid undesirable side reactions, such as self-coupling of the activated amino acid leading to (polymerization).[1] This would compete with the intended peptide coupling reaction, resulting in low yield or even complete failure to synthesize the desired peptide.

Two principle orthogonal protecting group schemes exist for use in solid-phase peptide synthesis: so-called Boc/Bzl and Fmoc/tBu approaches.[2] The Boc/Bzl strategy utilizes TFA-labile N-terminal Boc protection alongside side chain protection that is removed using anhydrous hydrogen fluoride during the final cleavage step (with simultaneous cleavage of the peptide from the solid support). Fmoc/tBu SPPS uses base-labile Fmoc N-terminal protection, with side chain protection and a resin linkage that are acid-labile (final acidic cleavage is carried out via TFA treatment).

Both approaches, including the advantages and disadvantages of each, are outlined in more detail below.

Boc/Bzl SPPSEdit

Cleavage of the Boc group

The original method for peptide synthesis relied on tert-butyloxycarbonyl (or more simply 'Boc') as a temporary N-terminal α-amino protecting group. The Boc group is removed with acid, such as trifluoroacetic acid (TFA). This forms a positively charged amino group in the presence of excess TFA (note that the amino group is not protonated in the image on the right), which is neutralized and coupled to the incoming activated amino acid.[18] Neutralization can either occur prior to coupling or in situ during the basic coupling reaction.

The Boc/Bzl approach retains its usefulness in reducing peptide aggregation during synthesis..[19] In addition, Boc/Bzl SPPS may be preferred over the Fmoc/tBu approach when synthesizing peptides containing base-sensitive moieties (such as depsipeptides), as treatment with base is required during the Fmoc deprotection step (see below).

Permanent side-chain protecting groups used during Boc/Bzl SPPS are typically benzyl or benzyl-based groups.[1] Final removal of the peptide from the solid support occurs simultaneously with side chain deprotection using anhydrous hydrogen fluoride via hydrolytic cleavage. The final product is a fluoride salt which is relatively easy to solubilize. Scavengers such as cresol must be added to the HF in order to prevent reactive t-butyl cations from generating undesired products. A disadvantage of this approach is the potential for degradation of the peptide by hydrogen fluoride.

Fmoc/tBu SPPSEdit

Cleavage of the Fmoc group. Treatment of the Fmoc-protected amine with piperidine results in proton abstraction from the methine group of the fluorenyl ring system. This leads to release of a carbamate, which decomposes into carbon dioxide (CO2) and the free amine. Dibenzofulvene is also generated. This reaction is able to occur due to the acidity of the fluorenyl proton, resulting from stabilization of the aromatic anion formed. The dibenzofulvene by-product can react with nucleophiles such as the piperidine (which is in large excess), or potentially the released amine.[20]

The use of N-terminal Fmoc protection allows for a milder deprotection scheme than used for Boc/Bzl SPPS, and this protection scheme is truly orthogonal under SPPS conditions. Fmoc deprotection utilizes a base, typically 20–50% piperidine in DMF.[15] The exposed amine is therefore neutral, and consequently no neutralization of the peptide-resin is required, as in the case of the Boc/Bzl approach. The lack of electrostatic repulsion between the peptide chains can lead to increased risk of aggregation with Fmoc/tBu SPPS however. Because the liberated fluorenyl group is a chromophore, Fmoc deprotection can be monitored by UV absorbance of the reaction mixture, a strategy which is employed in automated peptide synthesizers.

The ability of the Fmoc group to be cleaved under relatively mild basic conditions while being stable to acid allows the use of side chain protecting groups such as Boc and tBu that can be removed in milder acidic final cleavage conditions (TFA) than those used for final cleavage in Boc/Bzl SPPS (HF). Scavengers such as water and triisopropylsilane (TIPS) are added during the final cleavage in order to prevent side reactions with reactive cationic species released as a result of side chain deprotection. The resulting crude peptide is obtained as a TFA salt, which is potentially more difficult to solubilize than the fluoride salts generated in Boc SPPS.

Fmoc/tBu SPPS is less atom-economical, as the fluorenyl group is much larger than the Boc group. Accordingly, prices for Fmoc amino acids were high until the large-scale piloting of one of the first synthesized peptide drugs, enfuvirtide, began in the 1990s, when market demand adjusted the relative prices of Fmoc- vs Boc- amino acids.

Other protecting groupsEdit


The (Z) group is another carbamate-type amine protecting group, first used by Max Bergmann in the synthesis of oligopeptides.[21] It is removed under harsh conditions using HBr in acetic acid, or milder conditions of catalytic hydrogenation. While it has been used periodically for α-amine protection in peptide synthesis, it is almost exclusively used for side chain protection.

Alloc and miscellaneous groupsEdit

The allyloxycarbonyl (alloc) protecting group is sometimes used to protect an amino group (or carboxylic acid or alcohol group) when an orthogonal deprotection scheme is required. It is also sometimes used when conducting on-resin cyclic peptide formation, where the peptide is linked to the resin by a side-chain functional group. The Alloc group can be removed using tetrakis(triphenylphosphine)palladium(0).[22]

For special applications like synthetic steps involving protein microarrays, protecting groups sometimes termed "lithographic" are used, which are amenable to photochemistry at a particular wavelength of light, and so which can be removed during lithographic types of operations.

Regioselective disulfide bond formationEdit

The formation of multiple native disulfides remains one of the primary challenges of native peptide synthesis by solid-phase methods. Random chain combination typically results in several products with nonnative disulfide bonds.[23] Stepwise formation of disulfide bonds is typically the preferred method, and performed with thiol protecting groups.[24] Different thiol protecting groups provide multiple dimensions of orthogonal protection. These orthogonally protected cysteines are incorporated during the solid-phase synthesis of the peptide. Successive removal of these groups, to allow for selective exposure of free thiol groups, leads to disulfide formation in a stepwise manner. The order of removal of the groups must be considered so that only one group is removed at a time.

Thiol protecting groups used in peptide synthesis requiring later regioselective disulfide bond formation must possess multiple characteristics.[citation needed][verification needed] First, they must be reversible with conditions that do not affect the unprotected side chains. Second, the protecting group must be able to withstand the conditions of solid-phase synthesis. Third, the removal of the thiol protecting group must be such that it leaves intact other thiol protecting groups, if orthogonal protection is desired. That is, the removal of PG A should not affect PG B. Some of the thiol protecting groups commonly used include the acetamidomethyl (Acm), tert-butyl (But), 3-nitro-2-pyridine sulfenyl (NPYS), 2-pyridine-sulfenyl (Pyr), and trityl (Trt) groups.[citation needed] Importantly, the NPYS group can replace the Acm PG to yield an activated thiol.[25]

Using this method, Kiso and coworkers reported the first total synthesis of insulin in 1993.[26] In this work, the A-chain of insulin was prepared with following protecting groups in place on its cysteines: CysA6(But), CysA7(Acm), and CysA11(But), leaving CysA20 unprotected.[26]

Synthesizing long peptidesEdit

Stepwise elongation, in which the amino acids are connected step-by-step in turn, is ideal for small peptides containing between 2 and 100 amino acid residues. Another method is fragment condensation, in which peptide fragments are coupled. Although the former can elongate the peptide chain without racemization, the yield drops if only it is used in the creation of long or highly polar peptides. Fragment condensation is better than stepwise elongation for synthesizing sophisticated long peptides, but its use must be restricted in order to protect against racemization. Fragment condensation is also undesirable since the coupled fragment must be in gross excess, which may be a limitation depending on the length of the fragment.[citation needed]

A new development for producing longer peptide chains is chemical ligation: unprotected peptide chains react chemoselectively in aqueous solution. A first kinetically controlled product rearranges to form the amide bond. The most common form of native chemical ligation uses a peptide thioester that reacts with a terminal cysteine residue.[citation needed]

Other methods applicable for covalently linking polypeptides in aqueous solution include the use of split inteins,[27] spontaneous isopeptide bond formation[28] and sortase ligation.[29]

In order to optimize synthesis of long peptides, a method was developed in Medicon Valley for converting peptide sequences.[citation needed] The simple pre-sequence (e.g. Lysine (Lysn); Glutamic Acid (Glun); (LysGlu)n) that is incorporated at the C-terminus of the peptide to induce an alpha-helix-like structure. This can potentially increase biological half-life, improve peptide stability and inhibit enzymatic degradation without altering pharmacological activity or profile of action.[30][31]

Microwave-assisted peptide synthesisEdit

Although microwave irradiation has been around since the late 1940s, it was not until 1986 that microwave energy was used in organic chemistry. During the end of the 1980s and 1990s, microwave energy was an obvious source for completing chemical reactions in minutes that would otherwise take several hours to days. Through several technical improvements at the end of the 1990s and beginning of the 2000s, microwave synthesizers have been designed to provide both low and high energy pockets of microwave energy so that the temperature of the reaction mixture could be controlled. Microwave-assisted peptide synthesis uses a single frequency which provides maximum penetration depth of the sample, in contrast to conventional kitchen microwaves.[citation needed]

In peptide synthesis, microwave irradiation has been used to complete long peptide sequences with high degrees of yield and low degrees of racemization. Microwave irradiation during the coupling of amino acids to a growing polypeptide chain is catalyzed not only by the increase in temperature but also by the alternating electric field of the microwave.[32] This is because the polar N-terminal amine group and peptide backbone continuously try to align with the alternating electric field, thus helping prevent aggregation and increasing access to the solid phase reaction matrix. This increases yields of the final peptide products.[citation needed] There is however no clear evidence that microwave is better than simple heating and some peptide laboratories regard microwave just as a convenient method for rapid heating of the peptidyl resin. Heating to above 50–55 degrees Celsius also prevents aggregation and accelerates the coupling.[citation needed]

Despite the main advantages of microwave irradiation of peptide synthesis, a drawback has historically been increased epimerization of sensitive residues such as cysteine and histidine while activated esters. However, in the case of cysteine this can be avoided by using carbodiimide coupling processes even at temperatures up to 100C by avoiding the requirement of strong bases (ex. DIEA) [33]. An additional advantage of microwave processes is that they can be rapidly turned on and off allowing sensitive residues to be coupled at room temperature with microwave heating subsequently turned back on later.

Cyclic peptidesEdit

On resin cyclizationEdit

Peptides can be cyclized on a solid support. A variety of cylization reagents can be used such as HBTU/HOBt/DIEA, PyBop/DIEA, PyClock/DIEA.[citation needed] Head-to-tail peptides can be made on the solid support. The deprotection of the C-terminus at some suitable point allows on-resin cyclization by amide bond formation with the deprotected N-terminus. Once cyclization has taken place, the peptide is cleaved from resin by acidolysis and purified.

The strategy for the solid-phase synthesis of cyclic peptides in not limited to attachment through Asp, Glu or Lys side chains. Cysteine has a very reactive sulfhydryl group on its side chain. A disulfide bridge is created when a sulfur atom from one Cysteine forms a single covalent bond with another sulfur atom from a second cysteine in a different part of the protein. These bridges help to stabilize proteins, especially those secreted from cells. Some researchers use modified cysteines using S-acetomidomethyl (Acm) to block the formation of the disulfide bond but preserve the cysteine and the protein's original primary structure.[citation needed]

Off-resin cyclizationEdit

Off-resin cyclization is a solid-phase synthesis of key intermediates, followed by the key cyclization in solution phase, the final deprotection of any masked side chains is also carried out in solution phase. This has the disadvantages that the efficiencies of solid-phase synthesis are lost in the solution phase steps, that purification from by-products, reagents and unconverted material is required, and that undesired oligomers can be formed if macrocycle formation is involved.[34]

The use of pentafluorophenyl esters (FDPP,[35] PFPOH[36]) and BOP-Cl[37] are useful for cyclising peptides.

See alsoEdit


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Further readingEdit

External linksEdit