ATDBio - Solid Phase Synthesis of Oligonucleotides (2023)

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Millions of oligonucleotides are synthesized each year for use in laboratories around the world. Very small amounts of DNA are required for most applications, and oligonucleotide synthesis is mainly performed on a scale of 40 nmol or less. This provides enough for most biochemical and biological experiments. Larger amounts of DNA (10 μmol or more) can be prepared for biophysical studies (NMR and X-ray crystallography), and in extreme cases, solid-phase methods have been developed to synthesize kilo-oligonucleotides fordrug molecule(eg antisense oligonucleotides). For all these purposes, oligonucleotides are produced almost exclusively using automated solid-phase methods.

Advantages of solid phase synthesis

Solid-phase synthesis is widely used in peptide synthesis, oligonucleotide synthesis, oligosaccharide synthesis, and combinatorial chemistry. Solid-phase chemical synthesis was invented by Bruce Merrifield in the 1960s and was so important that he received an awardNobel Prize in Chemistry in 1984.

Solid-phase synthesis is performed on a solid substrate between filters, in a column that allows free flow of all reagents and solvents. Solid-phase synthesis has several advantages over solution synthesis:

• A large excess of reagents can be used in the solution phase to rapidly complete the reaction
• Contaminants and excess reagents are washed away, eliminating the need for cleaning after each step
• The procedure is suitable for automation on a computer-controlled solid-phase synthesizer.

solid support

Solid supports (also called resins) are insoluble particles, typically 50-200 µm in diameter, to which oligonucleotides are attached during synthesis. Various types of solid supports have been used, but controlled pore glass (CPG) and polystyrene have proven to be the most useful.

Controlled Porous Glass (CPG)

Controlled pore glass is rigid and non-expandable, with deep pores where oligonucleotides are synthesized. Glass substrates with 500 Å (50 nm) pores are mechanically strong and are commonly used for the synthesis of short oligonucleotides. However, when oligonucleotides larger than 40 bases were prepared on a 500 Å pore size resin, the synthesis yield was drastically reduced. This is because the growing oligonucleotides block the pores and limit the diffusion of the reagent through the matrix. Although macroporous resins are fragile, 1000 Å thick CPG resins have been shown to be suitable for the synthesis of oligonucleotides up to 100 bases long, and 2000 Å supports can be used for longer oligonucleotides.

Polystyrene (PS)

Highly cross-linked polystyrene beads are characterized by good moisture resistance and allow very efficient synthesis of oligonucleotides, especially at small scales (e.g. 40 nmol).

Solid supports for traditional oligonucleotide synthesis are typically loaded with 20-30 μmoles of nucleosides per gram of resin. Synthesis of oligonucleotides is less efficient at high loadings due to steric hindrance between adjacent resin-bound DNA strands, however, some applications, especially short oligonucleotides, take advantage of the ability to load up to 350 μmol/g polystyrene and the ability to synthesize large amts. of oligonucleotides.

Phosphoramidite method

Various methods of solution-phase oligonucleotide synthesis have been developed over the years, from the early experiments with the H-phosphonate and phosphotriester methods of Michelson and Todd and the Khorana phosphodiester method in the 1950s, to the reexamination of the phosphotriester method and the development of the Trieste phosphite process in the 1960s and 1970s. All these methods have their problems. Today the method of choice is the phosphoramidite method, pioneered by Marvin Caruthers in the early 1980s and perfected using solid phase technology and automation.

Synthesis of phosphoramidine oligonucleotides proceeds in the 3' to 5' direction (as opposed to the 5' to 3' direction of DNA biosynthesis)DNA replication). One nucleotide is added per round of synthesis. The DNA synthesis cycle of phosphoramidites consists of a series of steps described below:Figure 1.

Phosphoramidite Oligonucleotide Synthesis Cycle: Typical Time for Synthesis of DNA Oligonucleotides in the Phosphoramidite Cycle |

Detritylation of carrier-bound 3'-nucleosides

When starting oligonucleotide synthesis, the first protected nucleoside is pre-attached to the resin and the operator selects the synthesis column A, G, C or T based on the desired nucleoside at the 3' end of the oligonucleotide. The nucleoside bound to the substrate has a 5'-DMT (DMT = 4,4'-dimethoxytrityl) protecting group that prevents polymerization during resin functionalization and must be removed from the substrate Protecting group (detritylation) - binds to nucleosides before oligonucleotide synthesis can proceed. The detritylation mechanism is shown in the figureFigure 2.

Activation and pairing (step 1)

After detritylation, the carrier-bound nucleoside is ready to react with the next base, which is added as a nucleoside phosphoramidite monomer. Mix a large excess of the appropriate phosphoramidite nucleoside with the activator (tetrazole or derivative) and dissolve both in acetonitrile (a good solvent for nucleophilic substitution reactions). The diisopropylamino group of the phosphoramidite nucleoside is protonated by the activator, thus turning it into a good leaving group. It is rapidly removed by attacking the 5'-hydroxyl of the substrate-bound nucleoside at the adjacent phosphorus atom and forming a new phosphorus-oxygen bond, resulting in the support-bound phosphite triester (picture 3)

Nucleoside phosphoramidites are quite stable in an inert atmosphere and can be prepared in large quantities, shipped worldwide, and stored as a dry solid for several months before use. Nucleoside phosphoramidites become reactive only after protonation.

Coverage (step 2)

It is not an exaggeration to expect a 99.5% yield for each coupling step, but even with the most efficient chemistry and the purest reagents, it is impossible to achieve 100% reaction of the carrier-bound nucleosides with the incoming phosphoramidites. This means that there will be some unreacted 5'-hydroxyl groups on the resin-bound nucleotides. If left unchecked, these 5'-hydroxyl groups will be available in the next coupling step to react with incoming phosphoramidites. The resulting oligonucleotide will be nonsense and will correspond to the deletion mutation of the desired oligonucleotide (Figure 4). If these deletion mutations are not controlled, they will accumulate in each successive cycle and the end product will be a complex mixture of oligonucleotides, most of which will contain incorrect genetic information and will be difficult to purify. This would destroy any subsequent biochemical experiments.

Deletion mutations can be avoided by introducing a "capping" step after the coupling reaction to block the unreacted 5'-hydroxyl. Two capping solutions are used in the synthesizer: acetic anhydride andnitrogen- Methylimidazole (NMI). These two reagents (dissolved in tetrahydrofuran with the addition of a small amount of pyridine) are mixed in the DNA synthesizer and delivered to the synthesis column. The electrophilic mixture rapidly acetylates the alcohol and the pyridine maintains an alkaline pH to prevent detritylation of the nucleoside phosphoramidites by the acetic acid formed by the reaction of acetic anhydride with NMI (Figure 5). Acetylation of the 5'-hydroxyl renders it inactive in subsequent reactions.

Oxidation (step 3)

The phosphorous triester (P(III)) formed in the coupling step is unstable in an acidic environment and must be converted to stable (P(III)w)) before the next TCA detritylation step. This is achieved by oxidizing iodine in the presence of water and pyridine (Figure 6). The resulting phosphotriesters are actually DNA backbones protected with 2-cyanoethyl. The cyanoethyl group prevents unwanted reactions with phosphorus in subsequent synthetic cycles.

In some DNA synthesizers, oxidation of iodine is followed by a second capping step. This is to dry the resin as the remaining water in the oxidizing mixture will remain and inhibit the subsequent coupling reaction. Excess water reacts with the acylating agent to form acetic acid, which is eluted with a THF/pyridine solvent mixture.

Detritylation (step 4)

After coupling, capping, and oxidation of the phosphoramidite, the DMT protecting group at the 5' end of the resin-bound DNA strand must be removed for the primary hydroxyl group to react with the next phosphoramidite nucleotide. Deprotection with trichloroacetic acid in dichloromethane is rapid and quantitative. The orange color is due to fragmentation of the DMT carbocation, which absorbs in the visible spectrum at 495 nm. The intensity of this absorption is used to determine the coupling efficiency. Most commercially available DNA synthesizers are equipped with equipment to measure and record trityl production in each cycle so that you can monitor synthesis efficiency in real time.

This cycle is repeated once for each base to produce the desired oligonucleotide.

Effect of coupling efficiency on performance

The importance of high average step yield (coupling efficiency) and its effect on the overall yield of oligonucleotide synthesis is shown below.

The cumulative effect of a series of bad couplings is twofold and leads to:

• Poor overall yield of required oligonucleotides
• The product is extremely difficult to clean.

It is clear from the table that a very high step yield is required for the synthesis of oligonucleotides longer than 100 bases. In practice, 98.5% is easy to achieve, and average stage yields of 99.5% or better can be achieved as long as all reagents are pure. Well-dried anhydrous solvents should be used as the phosphoramidite coupling reaction is very sensitive to moisture.

detached from solid support

A linker is a chemical entity that connects the 3' end of an oligonucleotide to a solid substrate. It must be stable against all reagents used to assemble oligonucleotides on the solid phase, but must be able to be degraded under certain conditions at the end of the synthesis. The most commonly used ligand in oligonucleotide synthesis is the electrical ligand. It can be readily decomposed by treatment with concentrated ammonium hydroxide for one hour at room temperature (Figure 7)

In some synthesizers, the cleavage reaction is performed automatically and the ammonia solution containing the oligonucleotide is delivered to a glass vial. Alternatively, the solution can be performed manually by removing the column from the synthesizer and washing it with a syringe containing ammonium hydroxide.

Oligoprotection

The oligonucleotide is now dissolved in concentrated ammonia and heated to remove the protecting groups from the oligonucleotide.heterocyclic baseIPhosphodiester backbone(Number 8). The aqueous solution is then removed by evaporation and the oligonucleotide is ready for purification.

Deprotection of heterocyclic bases

The exocyclic primary amine groups of the heterocyclic bases (A, C, and G) are nucleophilic and therefore must be protected during oligonucleotide synthesis. In the final deprotection step, the protecting groups were quantitatively removed by treatment with concentrated ammonium hydroxide at 55°C for 5 h. The most commonly used heterocyclic base protecting groups are the followingFigure 9.

The benzoyl groups in A and C are rapidly cleaved in ammonium hydroxide, but the isobutyryl protecting group on guanine is more resistant to hydrolysis, and the rate-determining step in oligonucleotide deprotection is the isobutyryl group by guanine base cleavage. For some chemically modified oligonucleotides, heating in ammonia can cause degradation, so a more labile guanine protecting group is required in such cases. The most commonly used protecting group for labile guanine is dimethylformamide (dmf dG), which allows deprotection of oligonucleotides under milder conditions (concentrated ammonium hydroxide, 55°C for 1 h). For the synthesis of modified oligonucleotides with chemical groups that are highly sensitive to ammonia, a different set of “super” protected monomers must be used. The most popular of these are shown atFigure 10.

Extremely mild protecting groups can be removed with potassium carbonate in methanol or a mixture of 33% ammonia and 40% methylamine in water at room temperature. The reason for using acetyl dC as a protecting group is to avoid the side transamidation reaction of benzoyl dC with methylamine (Figure 11). Acetyl dC does not undergo transamidation reactions due to the very rapid hydrolysis of the acetyl group.

Deprotection of the phosphodiester backbone

Phosphate groups are protected as 2-cyanoethylphosphotriesters during oligonucleotide synthesis and must be deprotected after synthesis is complete. Because of the strong acidity of the hydrogen on the carbon atom adjacent to the electron-withdrawing cyano group, the cyanoethyl group is quickly removed to concentrated ammonium hydroxide. Its mechanism is β-elimination (Figure 12)

At the beginning of the synthesis of phosphoramidine oligonucleotides, the phosphate group is protected as a methyl triester and must be deprotected with thiophenol (Figure 13). Thiophenol is a foul-smelling, toxic liquid. Koester's development of β-cyanoethyl phosphoramidites is a particularly welcome development.

Adduct formation

Acrylonitrile, a byproduct of phosphodiester deprotection (Figure 12) is a Michael receptor. Under strongly basic conditions used for oligonucleotide deprotection, 2-cyanoethyl adducts can be formed with heterocyclic bases, particularly thymine (Figure 14)

If these cyanoethyl adducts are a problem, the steps of resin cleavage and deprotection of the phosphoramidine backbone can be reversed. If the substrate-bound oligonucleotide is treated with a solution of a weak base in an organic solvent (such as 10% diethylamine in acetonitrile or 1:1 triethylamine/acetonitrile), the cyanoethyl protecting group is released from the phosphate backbone, but the oligo is still bound from support (Figure 15)

Synthesis of nucleoside-phosphoamidine monomers

A, G, C and T phosphoramidites for oligonucleotide synthesis are produced on a large scale from free nucleosides derived from natural sources. Using these precursors avoids creating the required chirality in the molecule—not a simple task, because deoxynucleosides contain three stereocenters.

nucleoside protection

The synthesis of the four monomers begins with protection of the amine (actually the amidine and guanidine) of the heterocyclic base, deoxyadenosine (Figure 16)

Thymine does not require protection. The protected cytosine and guanine bases are shown inFigure 17.

Protected dA, dC and dG can also be made by transient protection of the alcohol function followed by benzoylation of the amino group (Figure 18). The advantage of this method is that silylation and benzoylation can be performed in a single reaction vessel without the need to isolate or purify silylated nucleoside intermediates ("one-pot" synthesis).

The amino group of deoxycytidine is sufficiently reactive to function with a reactive benzoate that does not react with the hydroxyl group. This provides a one-stage synthesisnitrogen(4)-Benzoil dC (Figure 19)

Trillization

Nucleosides (protected dA, dG and dC and unprotected T) for tritylation (Figure 20)

Phosphorylation

After purification, in the non-nucleophilic base diisopropylethylamine (DIPEA; DIPEA;Figure 21), to obtain a phosphoramidite monomer. After silica gel column chromatography, hexane precipitation, and microfilter filtration, the phosphoramidine monomer is ready for oligonucleotide synthesis.

The reagent of the above phosphorylation reaction is a reaction with phosphorus trichloride and 2-cyanoethanol followed bynitrogen,nitrogen- Diisopropylamine in the presence of DIPEA (nitrogen,nitrogen- Diisopropylethylamine, a non-nucleophilic base.Figure 22). Other secondary amines can attach to phosphorus, but dimethylamine and diethylamine nucleoside phosphoramidites are rather unstable, and larger amines react very slowly during the coupling step in oligonucleotide synthesis.

Since phosphorus is tetrahedral and the phosphorus atom is the chiral center, the phosphoramidine monomer formed in the phosphorylation reaction is a mixture of two diastereomers (Figure 23). These are different compounds that give two spots on TLC, two peaks on the HPLC spectrum (if the resolution is good enough) and two peaks on the chromatogram.³¹P NMR spectrum.

Resin functionalization

Controlled pore (aminoalkyl) glass and polystyrene resins are commercially available in amounts of 20-100 micromoles per gram. These resins are suitable for oligonucleotide synthesis provided they have pores of 500 Å or larger to allow diffusion of the necessary reagents and the individual particles are larger than 500 Å (50 nm) in diameter. Smaller particles prevent the rapid flow of solvents and reagents through the synthesis column and can clog filters. The chemistry of nucleoside attachment is shown inFigure 24. Typical oligonucleotide synthesis requires four separate resins—one functionalized resin with each base (dA, dG, dC, and dT).

To synthesize these functional resins, the nucleoside 5'-DMTFigure 20Treatment with succinic anhydride in the presence of pyridine at room temperature. Pyridine is a good solvent for the acylation reaction and also prevents detritylation of the DMT ether by the succinic nucleoside acid formed in the reaction. A large excess of each nucleoside electron is then added to the amino-functional resin batch along with an imide coupling agent and an acidic alcohol such as 4-nitrophenol to form an active ester where it acts as a good leaving group. This is followed by a capping step to exclude any unreacted amino groups that would otherwise cause oligonucleotide synthesis problems (Figure 24)

Calculation of resin consumption

The nucleoside loading of the resin can be determined by trityl analysis. Treat a small amount of resin (1 mg) with a strong acid (e.g., a 1:1 mixture of concentrated hydrochloric acid and ethanol) to cleave the DMT group. Take a sample of the resulting orange solution and measure its absorbance at 495 nm in a UV/Vis spectrophotometer. This allows the amount of DMT cation to be determined (using the extinction coefficient of the DMT cation at 495 nm,But_{Chapter 495}= 71 700 M⁻¹centimeter⁻¹). The resin content (per 1 mg of resin) is equal to the following amount:

@load =(But_{Chapter 495}/ONE_{Chapter 495}) ×V× (1/eat）@

WhereVis the volume of the cuvette,eatis the fraction of the solution used to measure absorbance.

Synthesis of oligonucleotides with universal support

The first base of a typical solid support is pre-immobilized onto the resin. Universal scaffolds do not have a base, instead the first base is added at the first connection stage (Figure 25)

The advantages of a universal substrate are obvious: for the synthesis of any oligonucleotide, one type of column can be used, regardless of whether its 3' end is A, C, G or T, or a modified (unnatural) base, e.g. . deoxyuridine, 5-methyldeoxycytosine or fluorescent dyes. For standard mounts, a separate composition column must be used for each standard mount, and columns for modified mounts may not be readily available.

However, the different chemical properties of the universal supports require different conditions to cleave the oligonucleotides from the substrate, which can be problematic.

Separation from common support

Cleavage of oligonucleotides from typical substrates involves ester hydrolysis which occurs rapidly in an aqueous base at room temperature (eg, 1 hour in ammonia at room temperature). In contrast, cleavage from the general support involves two steps: first hydrolysis of the esters and then dephosphorylation (cleavage of the PO bond)Figure 26). The first is general support for the need to treat with a relatively strong base such as methylamine to achieve dephosphorylation. However, incomplete cleavage is a common problem even under such harsh conditions. Universal agents such as UnyLinker/UnySupport have stretched structures where the presence of neighboring groups (auxiliary chimerism) accelerates the dephosphorylation reaction. Then, as with standard supports, ammonia can be used as a base to break down the resin and remove the protecting group. However, when handling UnyLinker/UnySupport, an extended solution time (eg 18 hours at 55°C) should be used.

The extended cleavage time required when using a universal vector increases the time required for oligonucleotide synthesis and precludes the use of reagents that are unstable under these conditions (such as "very mild" reagents) and cannot be used for RNA synthesis.