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Peptide Synthesis 3.Strategies for Peptide Synthesis

Author:N/A    | Post time:2012-05-26

3. Strategies for Peptide Synthesis
Top of page
1. Introduction
2. Fundamentals of Peptide Synthesis
3. Strategies for Peptide Synthesis
4. Chain-Growing and Side-Chain Protective Groups
5. Coupling Methods
6. Side Reactions
7. Summary and Outlook
Nowadays, all most commonly applied methods for peptide synthesis are based on the Nobel prize (1984) awarded solid-phase peptide synthesis (SPPS), which Merrifield invented in the beginning of the 1960s [2]. The previously used liquid-phase synthesis methods were associated with time-consuming purification steps, comparably low coupling yields, and difficult automation, and, therefore, are rarely used today.
 
3.1. Solid-Phase Peptide Synthesis
The SPPS is the today\'s method of choice to synthesize peptides and small proteins in bench and industrial scale. In this method, amino acid monomers are consecutively coupled to a growing peptide chain, which is immobilized on a solid support (Fig. 3). In every cycle, the free N-terminal amino group of the peptide attached to the solid-phase peptide reacts with the activated C-terminal group from a freely diffusing amino acid unit. These are added in 5–6-fold molar excess, which gives typical yields of 95–99%.
 
Figure 3. Principle of SPPS

 , à = Amino acid monomer;   = N-terminal transient protection

However, when compared to the oligonucleotide synthesis (  Nucleic Acids – Chemical Synthesis), the coupling yield is much more sequence-dependent. A stretch of two adjacent prolines is only one especially prominent example of “difficult” peptide sequences that result in low coupling yields, probably because of sterical hindrance or low solubility of the growing peptide chain that is attached to the solid phase.
Coupling yields below 100% always result in peptide fragments which do not react with the monomer and therefore are routinely “capped” by a very large molar excess of acetic acid anhydride, which permanently blocks the N-terminal end of these fragments and efficiently prohibits any chain elongation in further coupling steps (not shown in Fig. 3). Otherwise, a whole panel of closely related peptides would be generated. These would either cause problems in the interpretation of experimental results, or in the purification of the desired product.
As already discussed above, it is mandatory to protect all the reactive side chains (not shown in Fig. 3) and the N-terminal end during SPPS to avoid unwanted side reactions. After each coupling and capping step, the N-terminal transient protective group must be efficiently removed under conditions that leave the side-chain protection unaffected, and, thus, generate a new N-terminal amino group that binds to other amino acids in subsequent cycles. This principle of selective deprotection is called the concept of “orthogonality” (Chap. Chain-Growing and Side-Chain Protective Groups)
The most commonly used polymeric supports used during solid phase synthesis are
polystyrene-based resins (originally introduced by Merrifield),
polyamide supports,
poly(ethylene glycol) (PEG)-based resins (developed in the 1980s and commercial available as Tentagel™ or Argogel™ beads [3]).
All of these solid supports have one characteristic in common: they extensively swell when incubated in the solvents typically used for peptide synthesis. The most commonly used solvents for this purpose are dimethylformamide (DMF) or 1-methyl-2-pyrrolidone (NMP). Extensive swelling of a two-dimensional surface leads to strong shear forces. This is the reason why most solid supports for peptide synthesis are in the form of beads that can swell in all three dimensions of space, typically with a bead diameter of 100–200 µm.
In order to finally cleave these peptides from the synthesis beads, the bond between the solid support and the peptide must be prone to hydrolysis. This can be achieved by specialized linker molecules that attach the C-terminus of the peptide to the polymeric support via a labile benzyl ester or an amide linkage. The most commonly used linker molecules are the Rink amide linker and the Wang linker (Fig. 4). When incubated in concentrated trifluoroacetic acid, the synthesized peptide is cleaved from the support. Usually, this cleaving step also removes the acid-sensitive permanent side-chain protection (see Section Side-Chain Protective Groups (“Permanent Protection”)).
 
Figure 4. Linker molecules in SPPS synthesis
Synthesizing peptides on a solid support can also be important in some analytical settings (see Section Peptide Arrays). Here, the only requirement on the solid support is an amino or hydroxyl group where the amino acid building blocks attach to through their activated C-terminus. The chemical moieties most commonly used for this purpose are PEG [4] and cellulose [5].
The advantages of cellulose over PEG are 1. an enormous density of reactive hydroxyl groups, which are commonly converted to amine functionalities with  -alanine via esterification; and 2. that the peptide is easily cleaved directly from the solid support without a specialized and expensive linker molecule. The cleavage is done by the addition of a strong base that hydrolyzes the bond between the hydroxyl group of the glucose and the C-terminus of the peptide. Many different peptides can be synthesized in parallel on a sheet of cellulose. The peptides are deprotected under acidic conditions and finally cleaved from the cellulose in ammonia vapor. This procedure precipitates the free peptides on the cellulose fibers, resulting into hundreds or even thousands of different peptides that can easily be separated on the paper sheet with a pair of scissors [6]. It is noteworthy that this procedure results into a C-terminally amidated sequence.
The two transient protective groups mainly used today (2009) are 9-fluorenylmethoxycarbonyl (Fmoc) and tert-butoxycarbonyl (tBoc). Both groups are used together with their corresponding side-chain protection agents (Chap. Chain-Growing and Side-Chain Protective Groups).
 
3.2. Microwave-Assisted Peptide Synthesis
Peptide synthesis is a multimillion dollar business with an ever-increasing pressure on speed, quality, and pricing. Shorter coupling times can obviously be achieved with higher temperature, with microwave-derived energy being a source of energy that evenly distributes the energy throughout the whole vessel, and, thus, minimizes unwanted side reactions due to unevenly distributed heat. Introduced at the end of the 1990s, microwave synthesizers were available for peptide synthesis that considerably speeded up the synthesis reactions. Different to conventional microwaves, the peptide synthesis microwaves use one single frequency in order to provide a maximum penetration depth of the sample and uniform heating. Initially, this technique was regarded with suspicion, especially concerning the degree of racemization. Since the 1990s most commercially available peptides are synthesized by microwave peptide synthesizers. Microwave irradiation has been used to complete long peptide sequences with high yield and low degrees of racemization. However, when activated in situ at their C-terminus, cysteine and especially histidine are susceptible to racemization during the microwave-assisted synthesis at 80 °C. Aspartimide formation and subsequent racemization of aspartic acid can be reduced by the addition of hydroxybenzotriazole (HOBt, Section Diisopropylcarbodiimide (DIC)/N-Hydroxybenzotriazole (HOBt) Activation) to the deprotection solution and/or use of piperazine in place of piperidine [7]. Remaining susceptible amino acids are either coupled at 50°C, or used in an activated form that is not prone to racemization, namely the (expensive) ortho-pentafluoropheny-l (OPfp)-esters or the corresponding chlorides see Chap. Side Reactions) [8, 9]. Microwave irradiation during the coupling might have yet another feature that improves peptide synthesis: The alternating electromagnetic radiation is continuously absorbed by the growing peptide, which might prevent aggregation and thus increase the yield of the final product. Of course, also microwave-assisted peptide synthesis is based on the concept of solid-phase peptide synthesis.
 
3.3. Combinatorial Synthesis of Peptides
Decades after the first report, Merrifield\'s SPPS inspired yet another field of chemistry. The “one-bead–one-compound” method (  Combinatorial Chemistry – Split – Pool Synthesis Towards Combinatorial Libraries) adds one elegant trick to Merrifield\'s solid-phase synthesis. Specifically, huge peptide libraries are built by consecutive cycles of splitting beads for solid-phase synthesis among 20 different reaction vessels, with each reaction adding one of the 20 different amino acids to the growing, bead-bound peptide chain. Afterwards, beads are pooled before the next splitting cycle begins. This procedure imparts an individual history of sequential stopovers in the 20 vessels to each bead, and this history is reflected by the peptide sequence contained on each bead. Upon completion of the synthesis, nearly every bead displays a unique peptide, and each bead always contains only one kind of peptide (Fig. 5) [10]. This and similar approaches expanded the Merrifield synthesis to establish the field of combinatorial chemistry, which is characterized by a parallel processing of multiple reaction spheres to synthesize many different peptides or peptoids [11].
 
Figure 5. One-bead–one-compound method (beads are depicted as a flat surface)

 , à = Amino acid monomer;   = N-terminal transient protection

The extremely large number of different compounds generated by the one-bead–one-compound method has been successfully used to screen for peptide binders [12]. This chemical method is not restricted to the 20 natural l-amino acids that all biological systems rely on. The experimenter can also use hundreds of different commercially available building blocks. This feature is especially advantageous when searching for binding molecules for therapeutic intervention. Natural peptides are prone to degradation inside the human body, which is rich in proteases. On the other hand, at least partially non-natural molecules typically display superior pharmacokinetics, as their degradation is slower and they patrol the body for a longer time.
However, due to the random distribution of beads among the 20 different reaction vessels, it is nearly impossible to avoid the synthesis of “problematic” peptides during library preparation. These peptides bind to any protein, and, thus, elicit a strong background of false-positive binders. For example, hydrophobic patches are generated by in situ precipitated, insoluble peptides that usually stick to everything. Another drawback of the one-bead–one-compound method is the labor-intensive decoding that is necessary to obtain the sequences of library peptides binding to the target protein. Related to this drawback, only a restricted amount of information can be gained: The sequences of only a few binders can be characterized, while the majority of binders will be neglected. Information about which peptides do not bind is also missing. In some cases, these missing pieces of information are critical. For instance, before screening for different DNA-binding peptide modules that, when assembled, selectively target HIV-specific sequences, a full characterization of all potential binding partners should be performed to avoid the possibility that endogenous human DNA is targeted.
 
3.4. Peptide Arrays
When a scientist “arrays” different molecules on a two-dimensional surface, he exactly knows the position and the sequence of all of these different molecules. If such an array is then incubated, e.g., with a labeled protein, the diffusing protein probes all the different molecules on the array and eventually sticks to those with a complementary surface, i.e., specifically binds them. Thereby, with one single experiment, the experimenter immediately knows the sequences of those molecules that the particular protein interacted with and in addition the sequence of those that did not interact. Moreover, the information gained from this type of experiment increases with the number of arrayed molecules. This is the underlying reason why ever since the invention of the array by Ekins [13] scientists strive to achieve higher peptide or oligonucleotide densities on arrays. Another advantage of the array concept over, e.g., the one-bead–one-compound method is that the oligomers that nonspecifically bind to any protein are easily identified and simply omitted in the next array generation. Soon after the introduction of the array concept, the power of this approach began to be widely recognized in the field of peptide sciences, sparking many different attempts to assemble as many peptides in the array format as possible. Simulating the approach pioneered by Ekins, many companies presynthesized different peptides and spotted them onto support materials. The disadvantage of these methods is the high costs, which are only justified when many replicas of one standardized array are manufactured.
Similar to the approach pioneered by Southern to array oligonucleotides [15], the peptide array method invented by Frank in the beginning of the 1990s [14] entails patterning the 20 different activated amino acid derivatives as small droplets onto a derivatized cellulose sheet. There, consecutive layers react with the free amino groups of the many different growing peptide chains that are linked to the cellulose support during solid-phase synthesis. Again, each droplet defines a small reaction sphere (Fig. 6) [16, 17]. This SPOT synthesis method, for the first time, made large numbers of small proteins available to the scientific community. Over the years, the method has proven to be reliable and widely applicable and, thus, still dominates the field.
 
Figure 6. SPOT synthesis

 , à = Amino acid monomer;   = N-terminal transient protection

Many studies have used the SPOT synthesis to determine the exact binding motifs of monoclonal or polyclonal antibodies [18]. Peptide arrays have also revealed the binding motifs of proteins (e.g., chaperones) [19, 171]. This has also been achieved for binders that require a free C-terminal end, although conventional solid-phase peptide synthesis tethers the growing peptide chain to the solid support by its C-terminus. Boisguerin et al. synthesized peptide ligands, chemically turned around these ligands in the array format, which resulted into an array of peptides with freely accessible C-terminal ends, stained the array with PDZ protein domains, and determined the exact amino acid positions that, when phosphorylated, prevented peptide binding [20]. Others have used peptide arrays to profile kinase substrates [21, 22] or protease substrates [23]. In another application, arrayed peptides were cleaved from the cellulose support in a vapor of ammonia to screen for peptides that kill bacteria, an approach that might yield novel antibiotics [24].
In the beginning of the new century a method for the combinatorial synthesis of peptide arrays was developed by Breitling et al. which is based on 20 different sorts of solid amino acid particles. These particles (triboelectrically charged by mild friction) are positioned on a two-dimensional support by using electrical fields generated by either a laser printer [25] or a computer chip, whereby resolutions of up to 40 000 peptides per cm2 are possible [26]. Once positioned, the whole layer of amino acid particles is simultaneously melted to initiate the coupling reaction (Fig. 7). Washing and deprotection steps complete the cycle, resulting in the combinatorial synthesis of a peptide array, if repeated. This method uses the conventional Fmoc chemistry [27] (Chap. Chain-Growing and Side-Chain Protective Groups) and differs from the SPOT synthesis only in the use of an (at room temperature) solid solvent, which enables intermittent immobilization of amino acids within the particles.
 
Figure 7. Particle-based synthesis of peptide arrays

 , à = Amino acid monomer;   = N-terminal transient protection

 
3.5. Other Synthetic Methods
A chimeric approach uses cell-free translation mixtures that are readily available to the synthesis of peptides. These consist mainly of the ribosomes, different amino acids loaded on the tRNAs, and a mRNA that directs the synthesis of a specific peptide. The robust polymerases and ribosomes derived from thermophilic Archaebacteria [28] might be especially suited for the in situ translation of single peptides or even of an array of genes into a corresponding array of peptides or proteins. Although this approach, which uses the digitally programmed nanomachines from nature, is currently limited in terms of density and throughput, it has one major advantage over the available chemical methods: nature\'s ribosomes build very long peptides or even proteins [29].
However, the chemical methods strive to synthesize longer peptides, too. It is difficult to obtain peptides that are more than 70 amino acids long by SPPS because the yield of the desired product drops exponentially in consecutive coupling steps. The obvious solution to this difficulty is the condensation of two peptide fragments to one larger peptide. However, the number of fragment condensations must be limited because the C-terminal activation of a peptide (and also of an amino acid dimer) sensitizes the resulting peptide bond for racemization, see Chap. Side Reactions).
Yet another procedure for the production of longer peptides is called chemical ligation. In its most common form, it uses a thioester to connect two fragments via their cysteine residues.

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