5.10: Solid Phase Synthesis

1. Loading the Resin

1. To a 100mL peptide synthesis vessel, add 2-chlorotrityl chloride (CTC) resin (1.1 mmol/g, 0.360 g, 0.400 mmol). Add 20 mL DMF and allow them to swell for 30 min under N_2.
2. Drain the beads under vacuum and add 10 mL DMF.
3. Add 500 mg Fmoc-Ala-OH (1.60 mmol) and 2.5 mL i-Pr₂EtN, and mix under N₂ for 15 min.
4. Drain the solvent under vacuum and repeat the loading with Fmoc-Ala-OH for 15 min.
5. Drain the solvent under vacuum and wash the beads with 10 mL DMFunder N₂, and drain under vacuum 3x.

2. Deprotection of the Fmoc Group

1. Add 10 mL 20% 4-methylpiperidine in DMF and stir the beads under N₂ for 15 min.
2. Drain the solvent under vacuum and repeat the deprotection.
3. Wash the beads with 10 mL DMF under N₂ and drain under vacuum 3x.

3. Performing the Kaiser Test

1. Perform the Kaiser test by adding 1-2 drops of solution A (0.5 mL 0.01 M KCN, 24.5 mL pyridine), solution B (1 g ninhydrin, 20 mL n-butanol), and solution C (20 g phenol, 10 mL n-butanol) each into two test tubes. One test tube will be the control while the other will monitor the reaction.
2. Add a few beads of the resin from the reaction vessel to reaction test tube and heat up the two test tubes to 110 °C.
3. If the deprotection is complete, the contents of the test tube will turn a dark blue/purple color. If the deprotection is incomplete or failed, the solution will remain yellow. Compare the reaction test tube with the control test tube.

4. Coupling the Next Building Blocks

1. Drain the solvent under vacuum.
2. Wash the beads with 10 mL N-methyl-2-pyrrolidone under N₂ and drain the solvent under vacuum.
3. To begin the next coupling, add 10 mL NMP, 620 mgFmoc-Phe-OH (1.6 mmol), 610 mg HBTU (1.6 mmol), and 2.5 mLi-Pr₂EtN, and allow the resin to bubble under N₂ for 30 min.
4. Drain the solvent under vacuum.
5. Wash the beads with 10 mL DMF under N₂ and drain under vacuum 3x.
6. Perform the Kaiser test (see steps 3.1-3.3) to look for the completion of the coupling. The beads and solution in the test tube should be yellow.

5. Cleaving the Peptide Off the Resin

1. Cleave the remaining Fmoc group using steps 2.1-2.3.
2. After the solvent is drained under vacuum, add 40 mL cleavage solution (95% TFA, 2.5% H₂O, 2.5% TIPS) to the resin and bubble under N₂ for 3 h.
3. Place a new receiving flask on the peptide synthesizer and drain theTFA solution containing the desired peptide under vacuum into the new flask.

6. Precipitation and I solation of the Peptide

1. Separate the TFA solution into 4 conical vials and add 25 mL cold ether (−20 °C) to each vial to precipitate the peptide.
2. Centrifuge the vials (3,000 rpm, 0-4 °C) for 20 min. Decant the remaining TFA and ether solution from the conical vials and concentrate the peptide precipitate to afford the desired dipeptide as a white solid.

Solid phase synthesis is a method in which the product is synthesized while bound to an insoluble material.

Solid phase synthesis is often used to produce biological oligomers and polymers such as peptides, nucleic acids, and oligosaccharides. These molecules are composed of chains of smaller molecular subunits, called monomers. Synthesizing an oligomer or polymer takes many steps, as the monomers must be added in the correct order.

An issue with multi-step syntheses is that purification and isolation of the stable products of each step, called intermediate products, decreases the overall yield. In solid phase synthesis, the intermediate product remains bound to the solid support throughout synthesis. This allows solution-phase reagents, solvents, and byproducts to be washed away, eliminating the need to purify and isolate each intermediate product between steps.

This video will illustrate the procedure for solid phase peptide synthesis and introduce a few applications of solid phase synthesis in chemistry.

In solid-phase synthesis, a molecule is synthesized on a solid support in a sequence of reactions. For instance, an oligomer or polymer will be synthesized one monomer at a time to form the final product. The growing oligomer or polymer remains strongly bound to the solid support until it is separated, or cleaved, from the support with reagents.

Each monomer must have at least two binding sites to be part of the polymer chain, but only one binding site can be available at a time to ensure that the monomer binds to the correct atom. This is achieved with protecting groups, which are functional groups that are not reactive during one or more steps of the synthesis. The binding site is restored, or deprotected, by treating the molecule with specific reagents to convert the protecting group to a reactive functional group.

To begin solid-phase synthesis, the starting material is bound to a specially designed resin or insoluble polymer at its only available binding site. Then, the bound starting material is deprotected to allow binding of the second monomer in the chain. Next, a solution of the second monomer in the chain is added, along with a coupling agent to facilitate bonding between the monomers.

Once the second monomer binds to the starting material, the resulting dimeric intermediate product is deprotected. This process is repeated until the target oligomer or polymer has formed. The product is cleaved from the solid support into solution, from which it can be purified, isolated, and analyzed.

Solid phase synthesis is often used for the synthesis of peptides, which are chains of amino acids. Amino acids have an amine group, a carboxyl group, and a substituent, or 'side chain'. The amine is initially protected. Once deprotected, the amine forms a peptide bond with the carboxyl group of the next amino acid.

Now that you understand the principles of solid phase synthesis, let's go through a procedure for solid phase peptide synthesis, in which we will demonstrate the addition of the first two amino acids.

To begin the procedure, connect a receiving flask for waste to a 100-mL manual peptide synthesis vessel. Then place 0.360 g of 2-chlorotrityl chloride resin into the vessel. Connect a nitrogen gas line to the vessel sidearm and a vacuum line to the serrated hose adapter.

Add 20 mL of dimethylformamide to the resin and allow the resin beads to swell for 30 min under a flow of nitrogen gas. Then, apply vacuum to drain the solvent.

Add 10 mL of DMF, 1.6 mmol of an Fmoc-protected amino acid, and 2.5 mL of N,N-diisopropylethylamine to the vessel. Bubble under the nitrogen gas, which mixes the solution, for 15 min to load the protected amino acid onto the resin.

Remove the solvent under vacuum and perform a second loading. After removing the solvent, agitate the loaded resin beads three times in 10-mL portions of DMF, draining each wash into the receiving flask.

Next, add to the loaded beads 10 mL of a 20% solution of 4-methylpiperidine in DMF. Bubble the mixture for 15 min to remove the Fmoc group.

Drain the solvent and repeat the deprotection procedure. Wash and drain the loaded resin three times, as before. Store the beads under solvent until they are ready for the next step.

To verify that the loaded compound was completely deprotected, first place 1 to 2 drops of each Kaiser test solution in two test tubes.

Place a few loaded beads in a test tube and heat both tubes to 110 degrees in an oil bath. Deprotection is complete if the resin mixture turns dark blue to purple, indicating the presence of amine groups in the mixture.

To begin the coupling step, first wash the beads with 10 mL of NMP under a flow of N2 gas.

Then, add 10 mL of NMP, 1.6 mmol of the next Fmoc-protected amino acid, 1.6 mmol of the coupling agent HBTU, and 2.5 mL of DIPEA to the loaded resin.

Bubble N2 gas through the resin mixture for 30 minutes, and then drain the solvent. Wash and drain the beads with 10-mL portions of DMF three times, as before.

Repeat the Kaiser test. Coupling has occurred successfully if the beads and solution turn yellow, indicating that no amine groups are present.

Next, cleave the new Fmoc group with 20% 4-methylpiperidine in DMF and wash the beads with 10-mL portions of DMF. Repeat the coupling and deprotection for each remaining amino acid in the target peptide.

After the last amino acid has been deprotected and the resin beads have been washed, add 40 mL of peptide cleavage solution to separate the peptide product from the resin.

Bubble nitrogen gas through the resin mixture for 3 h, and then replace the receiving flask. Transfer the solution from the resin mixture to the new receiving flask under vacuum.

To generate the final product, remove the solvent with a rotary evaporator.

Solid phase synthesis is widely used in biology and chemistry. Let's look at a few examples.

Solid-phase synthesis opened many new synthetic pathways to oligosaccharides, which are short chains of simple sugar monomers with important biological roles, such as energy storage. Unlike peptide bonds, each bond between sugars contains a stereocenter. To synthesize an oligosaccharide, not only must the monomers be in the correct order, but the bonds must also have the correct stereochemistry. Solid-phase synthesis techniques were developed to couple each monomer by a highly stereoselective process, which today is sufficiently refined to be automated.

Solid-phase synthesis is a common approach to combinatorial chemistry, which is the practice of synthesizing many variants of a compound in a single synthetic process. The loaded resin can easily be split into portions to react with different monomers or molecules. After each reaction, the portions are washed and recombined. This is repeated until the desired number of products has been generated. This technique is particularly useful in pharmaceutical research, as it can be used to generate new compounds or to evaluate the reactivity of a compound with a wide array of molecules.

You've just watched JoVE's introduction to solid phase synthesis. You should now understand the underlying principles of solid phase synthesis, the procedure for solid phase peptide synthesis, and a few examples of how solid phase synthesis is used in organic chemistry. Thanks for watching!