Overview of Peptide Synthesis
Proteins are present in every living cell and possess a variety of biochemical activities. They appear as enzymes, hormones, antibiotics, and receptors. They compose a major portion of muscle, hair, and skin. Consequently, scientists have been very interested in synthesizing them in the laboratory. This interest has developed into a major synthetic field known as Peptide Synthesis. The major objectives in this field are four-fold:
The fundamental premise of this technique involves the incorporation of N-a-amino acids into a peptide of any desired sequence with one end of the sequence remaining attached to a solid support matrix. While the peptide is being synthesized usually by stepwise methods, all soluble reagents can be removed from the peptide-solid support matrix by filtration and washed away at the end of each coupling step. After the desired sequence of amino acids has been obtained, the peptide can be removed from the polymeric support.
The general scheme for solid phase peptide synthesis is outlined in Figure 1. The solid support is a synthetic polymer that bears reactive groups such as -OH. These groups are made so that they can react easily with the carboxyl group of an N-a-protected amino acid, thereby covalently binding it to the polymer. The amino protecting group (X) can then be removed and a second N-a-protected amino acid can be coupled to the attached amino acid. These steps are repeated until the desired sequence is obtained. At the end of the synthesis, a different reagent is applied to cleave the bond between the C-terminal amino acid and the polymer support; the peptide then goes into solution and can be obtained from the solution.
The crucial link in any polypeptide chain is the amide bond, which is formed by the condensation of an amine group of one amino acid and a carboxyl group of another. Generally, an amino acid consists of a central carbon atom (called the a-carbon) that is attached to four other groups: a hydrogen, an amino group, a carboxyl group, and a side chain group. The side chain group, designated R, defines the different structures of amino acids. Certain side chains contain functional groups that can interfere with the formation of the amide bond. Therefore, it is important to mask the functional groups of the amino acid side chain.
The general scheme which outlines the strategy of Fmoc synthesis is shown in Figure 2. Initially, the first Fmoc amino acid is attached to an insoluble support resin via an acid labile linker. Deprotection of Fmoc, is accomplished by treatment of the amino acid with a base, usually piperidine. The second Fmoc amino acid is coupled utilizing a pre-activated species or in situ activation. After the desired peptide is synthesized, the resin bound peptide is deprotected and detached from the solid support via TFA cleavage.
The removal of peptides in solid phase peptide synthesis is primarily done by acidolysis. The Fmoc chemistry employs the use of weak acids such as TFA or TMSBr. Various scavengers are included to protect the peptide from carbocations generated during cleavage which can lead to side reactions. These additives usually include thiol compounds, phenol, and water.
The following protecting groups are compatible with TFA and TMSBr cleavage:
Depending on the type of protecting groups present, certain combinations of scavengers must be used. For instance, when either Boc and t-Butyl groups are present, their carbocation counterparts (t-butyl cations and t-butyltrifluoroacetate) can react with Trp, Tyr, and Met to form their t-butyl derivatives. While EDT is a very efficient scavenger for t-butyl trifluoroacetate, it does not protect Trp from t-butylation. Therefore, water must be added in order to suppress alkylation. The indole ring of Trp and the hydroxyl group of Tyr are especially susceptible to the reactivity of the cleaved Pmc group. Again, water has been shown to be effective in suppressing this reaction. Similar occurrences can happen with the Trt and Mtr groups. Therefore, scavengers in the appropriate combination will greatly reduce the amount of side reactions.
The general scheme which outlines the strategy of Boc synthesis is shown in Figure 3. Initially, the first Boc amino acid is attached to an insoluble support resin via a HF cleavable linker. Deprotection of Boc, is accomplished by treatment of the amino acid with TFA. The second Boc amino acid is coupled utilizing a pre-activated species or in situ activation. After the desired peptide is synthesized, the resin bound peptide is deprotected and detached from the solid support via HF cleavage.
The Boc chemistry employs the use of strong acids such as HF, TFMSOTf, or TMSOTf. Various additives, usually thiol compounds are added to protect the peptide from the carbocations generated during cleavage.
The following protecting groups are compatible with HF cleavage:
The following protecting groups are compatible with TFMSOTf cleavage:
The following protecting groups are compatible with TMSOTf cleavage:
Coupling reactions in SPPS require the acylation reactions to be highly efficient to yield high-purity peptides.
Coupling Methods in Fmoc SPPS
The most widely used coupling method in Fmoc SPPS is the activated ester method either pre-formed (pre-activated species) or in situ (without pre-activation). Initially, the p-nitrophenyl and N-hydroxysuccinimide (ONSu) activated esters were the predominantly used forms (1-2). However, even in the presence of HOBt, the coupling reactions tended to be slow. In addition, ONSu esters of Fmoc amino acids were prone to the formation of the side product succinimido-carbonyl-b-alanine-N-hydroxysuccinimide ester (3-4). The most commonly used activated esters presently are the pentafluorophenyl (OPfp) ester and the 3-hydroxy-2,3-dihydro-4-oxo-benzo-triazone (ODhbt) ester (5-7). In the presence of HOBt, the rate of reaction is very rapid and the reaction is efficient with minimal side product formation. On the other hand, many coupling reactions can be done in situ using activating reagents such as DCC, HBTU, TBTU, BOP, or BOP-Cl. The direct addition of carbodiimide is considered to be the best choice (8-13). HBTU and TBTU would rank second, followed by BOP and finally BOP-Cl. With regards to ester coupling, the following order was found: BOP/HOBt > carbodiimide/HOBt ~ carbodiimide/ODhbT > DCC/OPfp (14-15).
More recently, 1-hydroxy-7-azabenzotriazole (HOAt) and its corresponding uronium salt analog O-(7-azabenzotrizol-1-yl)-1,1,3,3, tetra-methyluronium hexafluorophosphate (HATU) have been developed and found to have a greater catalytic activity than their HOBt and HBTU counterparts. The use of HOAt and HATU enhances coupling yields, shortens coupling times, and reduces racemization. Consequently, these reagents are suitable for the coupling of sterically hindered amino acids, thereby ensuring greater success in the synthesis of difficult peptides (16-17).
Coupling Methods in Boc SPPS
The carbodiimides, primarily DCC, were the coupling reagents of choice for many years (18). The major drawbacks encountered were the precipitation of dicyclohexylurea during the activation and acylation processes and the numerous side reactions associated with its usage. Several carbodiimides which produced soluble ureas were developed, such as diisopropylcarbodiimide (DIC), t-butyl methyl- and t-butylethyl-carbodiimides (19-22), but these did not resolve the problem of side reactions. Consequently, new types of activating agents were developed. The first of these was BOP (23), PyBroP (24-25) PyBOP (26), HBTU (27), TBTU (28), and HATU (29). All of these reagents require bases for activation.
All of the DCC and DCC-related derivatives discussed previously work by the formation of the symmetrical anhydride. The symmetrical anhydrides are usually very reactive and have been used extensively in SPPS, especially in Boc synthesis (30-33). Attempts at incorporating symmetrical anhydrides to Fmoc amino acids were met with some difficulties (34-36). For instance, symmetrical anhydrides prepared from N-(3-dimethylamino propyl)-N'-ethyl-carbodiimide•HCl, upon formation of the 2-alkoxy-5(4H)-oxazolone intermediate, rearranged in the presence of carbodiimides and tertiary amines (37). Also, not all of the Fmoc symmetrical anhydrides are soluble in DCM or remain insoluble regardless of the solvent used (38).
An alternative to the symmetrical anhydride is the mixed anhydride which is a carboxylic-carbonate or carboxylic-phosphinic mixed anhydride. Typically, these anhydrides are prepared by reacting either isobutyl- or isopropyl-chloroformate and substituted phosphinic chlorides with the N-a-protected amino acid (39-42). The reaction is typically rapid with little or no side reactions (43-46).
A type of mixed anhydride, N-carboxyanhydrides (NCA's), also known as Leuchs' anhydride have been widely used for the preparation of polyamino acids (47). This class of compounds combines N-a-protection with carboxyl group activation. Once reacted with another amino acid or peptide residue, the NCA releases carbon dioxide as its only by-product. NCA derivatives are easily prepared by treating a-amino acids with phosgene (48-51). The resulting NCA derivatives usually crystallize out and are ready for use under strictly defined conditions. These conditions require the pH to be carefully controlled during synthesis. At pH < 10, the peptide-carbamate (produced by the reaction between the NCA and the peptide or amino acid residue) tends to lose carbon dioxide with the generation of a free a-amino end group with resulting polymerization. At pH 10.5, hydrolytic decomposition of the NCA occurs. Therefore, the reaction is performed at pH 10.2. Another required condition is that the reaction proceeds for 2 minutes at 0°C with vigorous stirring. The resulting product is free of racemization and bears a free a-amino group that can be extended by addition of another anhydride.
Stepwise condensation is based on the repetitive addition of single N-a-protected amino acids to a growing amino component, generally starting from the C-terminal amino acid of the chain to be synthesized. The process of coupling individual amino acids can be accomplished through employment of the carbodiimide (52-53), the mixed carbonic anhydride (54-55), or the N-carboxyanhydride methods (56-57). The carbodiimide method involves coupling N- and C- protected amino acids by using DCC as the coupling reagent. Essentially, this coupling reagent promotes dehydration between the free carboxyl group of an N-protected amino acid and the free amino group of the C-protected amino acid, resulting in the formation of an amide bond with precipitation of the by- product, N,N'-dicylcohexylurea. This method, however, is hampered by side reactions which can result in racemization (58-59) or in the presence of a strong base, the formation of 5(4H)-oxazolones (60) and N-acylureas (61). Fortunately, these side reactions can be minimized, if not altogether eliminated, by adding a coupling catalyst such as N-hydroxysuccinimide (HOSu) or 1-hydroxybenzotriazole (HOBt). In addition, this method can be employed to synthesize the active ester derivatives of N-protected amino acids (62). In turn, the resulting activated ester will react spontaneously with any other C-protected amino acid or peptide to form a new peptide.
In cases where separation of the activated ester from the by-product dicyclohexylurea proves to be difficult, the mixed carbonic anhydride method can be employed. This method consists of two stages: the first stage involves activating the carboxyl group of an N-a-protected amino acid with an appropriate alkyl chlorocarbonate, such as ethylchlorocarbonate (63), or preferably isobutylchlorocarbonate (64). Activation occurs in an organic solvent in the presence of a tertiary base. The second stage involves reacting the carbonic anhydride with a free amine component of an amino acid or a peptide unit. The carbonic anhydride is usually added at a 14-fold excess over the amine component.
The mixed carbonic anhydride method is noted for being highly effective at low temperatures, resulting in high yields and pure products. However, it does have its short-comings. For instance, there is a tendency for the anhydride derivative to undergo racemization as a result of the strong activation of the carbonyl group. This problem does not occur when N-a-urethane protecting groups, such as Cbz or t-Boc, are employed (65-66). Furthermore, as a result of their high reactivity, mixed carbonic anhydrides are prone to the formation of 5(4H)-oxazolones (67), urethanes (68-69), diacyimides (70-71), esters (72), and are subject to disproportion (73-74). Conditions which prompt such side reactions to occur are high temperatures, prolonged activation times (the time interval between the addition to the alkylchlorocarbonate and the amine component after the mixed anhydride is formed), steric bulk of the amine component, and incomplete formation of the mixed anhydride. Fortunately, most of these side reactions, except for oxazalone and urethane formation, can be substantially reduced by performing the reaction at low temperature (~ -15°C) and allowing for shorter activation times (~ 1-2 min). To minimize the formation of oxazolone and urethane derivatives, the following conditions must be implemented: 1) dried organic solvents such as ethyl acetate, tetrahydrofuran, t-butanol, or acetonitrile must be used (75); 2) the tertiary base, N-methylmorpholine, should be used (76); and 3) Cbz- or Boc-N-a-protected amino acids must be utilized (77).
Although isobutyl- and ethylchlorocarbonate are typically used to form carbonic anhydrides, other coupling reagents do exist. For example, N-ethyloxycarbonyl-2-ethyloxy-1,2-dihydroquinoline (EEDQ) (78) and N-isobutyloxy-carbonyl-2-isobutyloxy-1,2-dihydroquinoline (IIDQ) (79) were developed to react with the carboxyl component to form the ethyl- or isobutylcarbonate derivative. Unlike the classical anhydride procedure, EEDQ and IIDQ do not require base nor low reaction temperatures. Typically, the procedure involves reacting equimolar amounts of the carboxyl and amine components in an organic solvent (a wide variety of solvents can be used) (80) at 0.1 M to 0.4 M concentrations. Then EEDQ or IIDQ is added in 5-10% excess and the mixture is allowed to stir for 15-24 hours at room temperature. After removal of the solvent, in vacuo, the residue is dissolved in ethyl acetate and washed with 1N NaHCO3, 10% citric acid, and salt water, then dried with Na2SO4 (anhydrous), and evaporated. The product can then be recrystallized or purified by chromatography. While this method circumvents the use of base, it is still subject to racemization and urethane side product formation at levels comparable to those found in the classical anhydride approach. Consequently, its only advantage may be that it is easy and convenient to use. It should be noted that a detailed comparison of the two methods has not been carried out to this date.
Analytical HPLC utilizes columns and pumping systems that can withstand and deliver very high pressures enabling the use of very fine particles (3-10 microns) as packing material. Consequently, peptides can be resolved with a high degree of resolution in a short time interval (i.e., minutes).
Two common HPLC purification methods are, ion exchange and reverse phase. Ion exchange HPLC is based on direct charge interactions between the peptide and the stationary phase. The column support is derivatized with an ionic species that maintains a particular charge over a certain pH range, while the peptide or peptide mixture exhibits an opposite charge which is dependent on its amino acid composition. Separation is dependent on charge interactions. The peptide is eluted by changing the pH, the ionic strength, or both. Typically, a solution of low ionic strength is used; the ionic strength of the solution is then gradually or step-wise increased until the peptide is eluted from the column. One example of ion exchange separation incorporates the use of strong cation exchange columns such as sulfoethylaspartimide which separates on the basis of positive charge at an acidic pH.
Reverse phase HPLC conditions are essentially the reverse of normal phase chromotography. The peptide binds on the column through hydrophobic interactions and is eluted by decreasing the ionic strength (i.e., increasing the hydrophobicity of the eluent). Generally, the column supports are composed of hydrocarbon alkane chains which are covalently attached to silica. These chains range from C4 to C18 carbon atoms in length. Since elution from the column is a function of the hydrophobicity, the longer chain hydrocarbon columns are better for small, highly charged peptides. On the other hand, large hydrophobic peptides elute better using short chain hydrocarbon supports. However, in general practice, these two types of columns can be used interchangeably with little significant differences. Other types of supports consist of aromatic hydrocarbons such as phenyl groups.
A typical run usually consist of two buffers, 0.1% TFA-H2O and 80% acetonitrile/0.1% TFA-H2O, which are mixed using a linear gradient with a flow rate which will give a 0.5% to 1.0% change per minute. Typical columns for analytical and purification runs are 4.6 x 250 mm (3-10 microns) and 22 x 250 mm (10 microns), respectively. If radial packed columns are used, then column sizes are 8 x 100 mm (3-10 microns) and 25 x 100 mm (10 microns), respectively. A variety of other buffers can contain many different types of reagents such as 0.1% heptafluorobutyric acid, 0.1% phosphoric acid, dilute HCl, formic acid (5-60%, pH 2-4), 10-100 mM NH4HCO3, sodium/ammonium acetate, TFA/TEA, sodium or potassium phosphate, or triethylammonium phosphate (pH 4-8). In addition, water miscible eluents can also be added such as methanol, propanol, and isopropanol. Therefore, many combinations of solvents and additives for a buffer are possible. It should be noted that silica-based reverse phase column packing must not be exposed to high pH's or even slightly basic pH's for extended periods of time because the column can be destroyed at those pH levels.
The crude peptide obtained from SPPS will contain many by-products which are a result of deletion or truncated peptides as well as side products stemming from cleaved side chains or oxidation during the cleavage and deprotection process. Earlier purification methods included ion exchange, partition, and counter current chromatography. Recent purification methods include reverse phase HPLC which is generally successful with peptides containing 60 residues or less. In conjunction, ion exchange HPLC can be used in cases where reverse phase HPLC does not work.
Typically, analytical HPLC results are used to determine the purification conditions. For example, if a peptide elutes out at 30% (0.1% TFA) aqueous acetonitrile (determined by analytical HPLC analysis), a buffer containing a lower concentration of acetonitrile is chosen such that the peptide peak will come out 4-5 minutes after the solvent peak under isocratic conditions (e.g., 28% (0.1% TFA) aqueous acetonitrile). The purification conditions will entail using a linear gradient of 16-35% (0.1% TFA) aqueous acetonitrile over one or two hours depending on the type of column chosen. The collected fractions will then be checked by analytical HPLC, using the buffer chosen for isocractic conditions.
Peptides have widely varying solubility properties. The main problem associated with the dissolution of a peptide is secondary structure formation. This formation is likely to occur with all but the shortest of peptides and is even more pronounced in peptides containing multiple hydrophobic amino acid residues. Secondary structure formation can be promoted by salts. It is recommended first to dissolve the peptide in sterile distilled or deionized water. Sonication can be applied if necessary to increase the rate of dissolution. If the peptide is still insoluble, addition of a small amount of dilute (approximately 10%) acetic acid (for basic peptides) or aqueous ammonia (for acidic peptides) can facilitate dissolution of the peptide.
For long-term storage of peptides, lyophilization is highly recommended. Lyophilized peptides can be stored for years at temperatures of -20°C or lower with little or no degradation. Peptides in solution are much less stable. Peptides are susceptible to degradation by bacteria so they should be dissolved in sterile, purified water.
Peptides containing methionine, cysteine, or tryptophan residues can have limited storage time in solution due to oxidation. These peptides should be dissolved in oxygen-free solvents. To prevent the damage caused by repeated freezing and thawing of peptides, dissolving the amount needed for the immediate experiment and storing the remaining peptide in solid form is recommended.