Oligonucleotide synthesis chemistry

Background

An oligonucleotide is a linear polymer consisting of nucleoside units connected via phosphodiester linkages, where each internucleotide bridge is a phosphodiester monoanion formed between the 3'-hydroxyl of one sugar moiety and the 5'-hydroxyl of the adjacent nucleoside through a central phosphorus(V) atom bearing a non-bridging oxygen with a formal negative charge at physiological pH. The backbone comprises alternating β-D-2'-deoxyribofuranose (or ribofuranose for RNA) residues and phosphate groups in the canonical 3'→5' regioisomeric orientation, with each pentose C1' position bearing a nucleobase (adenine, guanine, cytosine, thymine, or uracil) via an N-glycosidic bond that adopts an anti-conformation in B-form duplexes.

During solid-phase synthesis, we construct the oligonucleotide architecture iteratively by detritylating the 5'-O-DMT protecting group, coupling a nucleoside-3'-O-(2-cyanoethyl-N,N-diisopropyl)phosphoramidite using tetrazole or similar activators to form a phosphite triester, oxidizing the P(III) intermediate to the P(V) phosphotriester with iodine/water/pyridine, and capping any unreacted 5'-hydroxyls with acetic anhydride.

Synthetic oligonucleotides produced by solid-phase synthesis are chemically identical to natural DNA segments of equivalent sequence once all protecting groups are removed, though synthetic oligos may occasionally retain trace protecting groups or linkage isomers (such as residual cyanoethyl groups on the phosphate or rare 2'→5' linkages from synthesis errors), and they lack the higher-order chromatin organization, epigenetic modifications (like 5-methylcytosine), and protein associations found in genomic DNA within cells. Thus, a purified synthetic oligonucleotide and a natural DNA fragment of the same sequence are (usually) structurally indistinguishable at the molecular level, differing only in context, length conventions, and potential minor impurities from synthesis.

The evolution of oligonucleotide synthesis methodologies progressed through multiple phosphorus-based chemistries that revolutionized nucleic acid research. The phosphodiester approach, pioneered by Khorana and colleagues in the 1950s-1960s, employed solution-phase P(V) chemistry using dicyclohexylcarbodiimide (DCC) activation to condense fully protected nucleotide monomers, enabling the first chemical gene synthesis despite laborious purification and modest yields. The phosphotriester method, developed by Letsinger in the 1960s and refined by Reese in the 1970s, utilized fully protected phosphate intermediates as neutral triesters (with aryl groups like 2-chlorophenyl or simple alkyl groups such as methyl protecting the non-bridging oxygen), permitting organic solvent-based synthesis with improved coupling rates compared to phosphodiester chemistry but requiring post-synthetic removal of phosphate protecting groups.

Historical Overview of Oligonucleotide Synthesis

The chemical evolution of oligonucleotide synthesis represents a trajectory from labor-intensive solution-phase couplings of pentavalent phosphorus—plagued by ionic shielding and sluggish substitution kinetics—to high-efficiency solid-phase methodologies exploiting the superior electrophilicity of trivalent phosphorus. The early era was defined by the Phosphodiester Approach (Khorana), which utilized unprotected phosphate monoesters and carbodiimide-mediated activation, enabling the first gene synthesis but suffering from pyrophosphate formation and extensive ion-exchange purification requirements. To mitigate the solubility and side-reaction limitations imposed by the anionic backbone, the Phosphotriester Approach (Letsinger) introduced fully protected, neutral phosphate triesters, facilitating the use of organic solvents and powerful sulfonyl condensing agents. A paradigm shift occurred with the introduction of P(III) chemistry; the Phosphite Triester Approach (Letsinger) demonstrated that trivalent phosphorus species, specifically phosphorochloridites, coupled orders of magnitude faster than their P(V) counterparts due to the lower activation energy required to access the trigonal bipyramidal transition state. This laid the groundwork for the Phosphoramidite Approach (Beaucage & Caruthers), which stabilized the P(III) center via a P–N bond, yielding the stable monomers that drive modern automated synthesis. Concurrently, the H-Phosphonate Approach emerged as a P(III) alternative exploiting the tautomeric equilibrium of phosphonate monoesters to allow global oxidation, while intrinsic backbone modifications were realized via the Methylphosphonate Approach. By transitioning from P(V) condensation to P(III) coupling followed by in situ oxidation, the field achieved the stepwise yields (>99%) required to assemble long genomic sequences with high fidelity.

Preparation of a dithymidinyl nucleotide (Michelson and Todd, 1955)

The first published account of the directed chemical synthesis of an oligonucleotide occurred in 1955 when Michelson and Todd reported the preparation of a dithymidinyl nucleotide (Michelson and Todd, 1955).

In their report, the phosphate link between two thymidine nucleosides was made by first preparing the 3' phosphoryl chloride of a 5' benzoyl protected thymidine, using phenylphosphoryl dichloride. This was then reacted with the 5' hydroxyl of a 3' protected thymidine. The chemistry worked reasonably well, albeit slowly. Additionally, the phosphoryl chloride intermediate was not stable, being susceptible to hydrolysis.

Phosphate Triester chemistry

In the early 1950s, Alexander Todd's group pioneered H-phosphonate and phosphate triester methods of oligonucleotide synthesis. Thirty years later, the work on H-phosphonate from Todd's group inspired, independently, two research groups to adopt the H-phosphonate chemistry to the solid-phase synthesis using nucleoside H-phosphonate monoesters as building blocks and pivaloyl chloride, 2,4,6-triisopropylbenzenesulfonyl chloride (TPS-Cl), and other compounds as activators.

On/off protection scheme (Khorana)

Khorana introduced two concepts to the field that made possible the convenient synthesis of oligonucleotides more than just a few bases long. One concept, the on-off protection scheme necessary for sequential oligonucleotide synthesis, is still widely used today by oligonucleotide chemists, virtually unmodified from Khorana's initial publications (Schaller, et. al., 1963; Smith, et. al., 1961). The second major concept was the phosphodiester method.

Exocyclic amine protecting groups (Khorana)

Khorana also introduced the protecting groups for the nucleosidyl exocyclic amines that are today known as the standard protecting groups; isobutyryl for guanosine and benzoyl for adenosine and cytidine (Schaller, et. al., 1963; Brown, et. al., 1979). Although others exist, these are the most commonly used groups today, with the possible exception of acetyl-protected cytidine, which is more readily removed compared to benzoyl.

Professor Khorana influenced many with his work, both through his publications and through his labs, where many of the great names in oligonucleotide chemistry passed as graduate students, post-docs, or visiting scholars. Those names include Marvin Caruthers, and Robert Letsinger, who worked nearby at Northwestern University and developed two important steps in the field: solid phase synthesis and phosphite-triester chemistry.

Solid-Phase Peptide Synthesis (Merrifield approach)

While not an oligonucleotide chemistry per se, the solid-phase peptide synthesis (SPPS) methodology introduced by R.B. Merrifield in 1963 provided the conceptual and mechanical framework upon which solid-phase oligonucleotide synthesis was subsequently built. Departing from solution-phase methods where every intermediate required isolation, Merrifield utilized an insoluble cross-linked polystyrene-divinylbenzene resin functionalized with chloromethyl groups (chloromethyl polystyrene). The carboxyl-terminal amino acid was anchored to this support via a benzyl ester linkage, allowing the peptide chain to grow in the C-to-N direction—opposite to the biological ribosomal translation. The alpha-amino group of the incoming amino acid required temporary protection, initially achieved using the acid-labile tert-butoxycarbonyl (Boc) group.

The synthetic cycle consists of repetitive deprotection and coupling steps: the Boc group is removed with trifluoroacetic acid to expose the alpha-amine, neutralized with triethylamine, and coupled to the next C-activated Boc-amino acid using dicyclohexylcarbodiimide (DCC) to form the amide bond. A critical evolution in this field was the introduction of the Fmoc/tBu strategy by Carpino, which employs the base-labile 9-fluorenylmethoxycarbonyl (Fmoc) group for alpha-amine protection, removal via piperidine, and acid-labile tert-butyl ethers/esters for side-chain protection. This orthogonal protection scheme avoids the repetitive acidolysis of the Boc method and allows final cleavage from the resin using trifluoroacetic acid rather than the hazardous liquid hydrogen fluoride required for Boc chemistry. It was the successful automation of these heterogeneous phase reactions—driving equilibria with excess reagents and removing byproducts via simple filtration—that inspired Itakura and Riggs to adapt the concept for nucleic acids, fundamentally shifting oligonucleotide synthesis from solution to the solid phase.

video: A lecture on solid phase peptide synthesis

Solid phase peptide synthesis (Letsinger)

Letsinger's first support for peptide synthesis was described in papers published in 1963 and 1964 (Letsinger and Kornet, 1963; Letsinger, et. al., 1964). The support consisted of what was called a "popcorn" polymer, a styrene-divinylbenzene polymer that had the unfortunate property of swelling in some solvents. In 1965 he published the first paper describing the solid-phase synthesis of dimer and trimer oligonucleotides using the same support (Letsinger and Mahadevan, 1965). In the initial report, 2' deoxycytidine (dC) was attached through the amine at the 4 position of the base itself to acid chloride modified support and forming an amide bond that was cleaved with ammonium hydroxide. The 3' hydroxyl of the dC was protected with a benzoyl group and the 5' position with a dimethoxytrityl (DMT) group. The DMT group was then removed with mild acid to prepare the support bound nucleoside for oligonucleotide synthesis. The attachment was made to the support, which was activated to an acid chloride, thus forming an amide bond that was cleavable with base (see #coupling-of-dC-to-polymer-support).

Letsinger began his career at Northwestern University in the late 1940s as a boron chemist. He was a significant player in that field, but in the early 1960s he turned his sights onto biomacromolecule synthesis. At that time, the target was peptide synthesis. However, a twist of fate moved Letsinger from peptide to oligonucleotide chemistry in the mid-1960s.

Letsinger was developing a peptide synthesis scheme using solid phase chemistry that had originated mainly for the support of catalysis. Letsinger utilized flow-through technology with a cyclic chemistry scheme of adding units sequentially. When applied to peptide synthesis, it added an internal filtering system that proved to be an incredibly important step forward. However, he wasn't the only researcher following this lead. Another scientist, Bob Merrifield, was also investigating the synthesis of peptides using solid phase technology. At the time they were neck and neck in the process of discovery and struggling to publish their findings as soon as possible. Bob Merrifield submitted his seminal paper describing the solid phase synthesis of peptides first and eventually won the Nobel Prize for his work. This unexpected scoop prompted Letsinger to regroup and focus his attention on another nascent chemistry: oligonucleotide synthesis. He rapidly converted his method using solid phase synthesis for peptides to the improvement of the oligonucleotide synthesis procedures taught by Khorana, thus converting a stroke of hard luck into scientific advancements that benefited an entire industry.

Letsinger made three major contributions to the field. First, he introduced solid phase chemistry as stated above. Secondly, he introduced the phosphotriester method of synthesis, an important improvement on Khorana's phosphodiester method. Finally, he introduced a radical departure, the phosphorus(III) based phosphite-triester method, which is the root of Marvin Caruthers' phosphoramidite method.

Through the 1960s he continued to explore the solid phase synthesis technique. He quickly determined that the best approach to solid phase synthesis was to attach the 3' hydroxyl to the support, as is done today. In fact, the graduate student instrumental in that work was Marvin Caruthers. Letsinger explored a number of polymer formulations, but never found a solution to the problem of swelling that was so detrimental to the chemistry. That role fell to his former student, Caruthers.

The Phosphodiester Approach

The phosphodiester approach, developed by Khorana and colleagues during the 1950s and 1960s, represents the foundational methodology for chemical oligonucleotide synthesis. This method relies on phosphorus(V) chemistry in solution, wherein fully protected nucleoside-3'-phosphates are condensed with 5'-hydroxyl-bearing nucleosides.

The activation strategy is mediated specifically by carbodiimide coupling reagents, most commonly dicyclohexylcarbodiimide (DCC) or later variants like 1-ethyl-3-(3-dimethylaminopropyl)carbodiimide (EDC). The mechanism proceeds through the activation of the phosphate monoester to form a highly reactive phosphoric anhydride-type intermediate (or acyl phosphate). The 5'-hydroxyl group of the acceptor nucleoside then attacks this electrophilic phosphorus center to form the internucleotide phosphodiester linkage.

A critical limitation of this chemistry is that the resulting internucleotide phosphate oxygen remains unprotected and anionic during synthesis (with a pKa of approximately 1). This anionic charge creates a "shielding" effect that repels incoming nucleophiles, drastically reducing reaction rates, and mandates the use of polar solvents like pyridine to maintain solubility. Furthermore, the unprotected phosphate oxygen remains nucleophilic, competing with the 5'-hydroxyl for the activated monomer. This leads to complicated side reactions, including the formation of 3'-to-3' pyrophosphate branches or cyclic triphosphates.

To ensure successful synthesis, robust protection strategies were required, including N-acyl groups on exocyclic amines of nucleobases (benzoyl for adenine and cytosine, isobutyryl for guanine) and acid-labile groups like monomethoxytrityl (MMT) for primary hydroxyls. Ultimately, the presence of branched impurities necessitated laborious isolation procedures using anion-exchange chromatography to resolve the desired linear product.

In the 1950s, Har Gobind Khorana and co-workers developed a phosphodiester method where 3'-O-acetylnucleoside-5'-O-phosphate was activated with N,N'-dicyclohexylcarbodiimide (DCC) or 4-toluenesulfonyl chloride (Ts-Cl). The activated species were reacted with a 5'-O-protected nucleoside to give a protected dinucleoside monophosphate. Upon the removal of 3'-O-acetyl group using base-catalyzed hydrolysis, further chain elongation was carried out. Following this methodology, sets of tri- and tetradeoxyribonucleotides were synthesized and were enzymatically converted to longer oligonucleotides, which allowed elucidation of the genetic code.

The major limitation of the phosphodiester method consisted in the formation of pyrophosphate oligomers and oligonucleotides branched at the internucleosidic phosphate. The phosphodiester method seems to be a step back from the more selective H-phosphonate chemistry described earlier, but at the time most phosphate-protecting groups available now had not yet been introduced. The lack of the convenient protection strategy necessitated taking a retreat to a slower and less selective chemistry. Without the phosphate-protecting group developments, there was an absence of phosphate-masking moieties that facilitate the temporary conversion of charged diesters into neutral phosphotriesters, most notably the 2-cyanoethyl group and various substituted aryl protecting groups like 2-chlorophenyl or 4-chlorophenyl. The 2-cyanoethyl group is particularly valuable because its electron-withdrawing character renders the beta-protons sufficiently acidic to permit facile deprotection via a base-catalyzed beta-elimination mechanism, releasing acrylonitrile and leaving the phosphodiester intact without attacking the internucleotide linkage. Similarly, the unavailability of aryl adjuncts such as the 2-chlorophenyl group precluded the development of the phosphotriester method at that time; these groups are chemically significant because they effectively cap the nucleophilic phosphate oxygen, thereby preventing the formation of pyrophosphate branches and cyclic triphosphates, while simultaneously masking the anionic charge to enhance solubility in organic solvents and enable purification via standard normal-phase silica chromatography rather than laborious anion-exchange protocols.

Besides the Khorana contribution of the on-off protection scheme, the other major Khorana contribution was the first use of a stable phosphorylated nucleoside that coupled to the desired nucleoside when activated (Khorana, et. al., 1956). This protocol is the same cyclic scheme used today with the exception of the addition of one step, oxidation. In place of the hydrolysable phosphoryl chloride, he prepared 3' phosphates of the 5' protected nucleoside using phosphorochloridates that then hydrolyzed to the phosphomonoester. These 5' protected nucleoside 3' phosphates were subsequently activated using a condensation reagent, such as dicyclohexyl carbodiimide (DCC), to couple to the 5' hydroxyl of another 3'-protected nucleoside. This method was revolutionary at the time and produced a truly remarkable feat: the synthesis of an active 72-mer tRNA molecule, which was published in Nature (Khorana, 1970).

Like most archetypes, the method did have shortcomings. Because the phosphate itself was not protected, branching at the internucleotide phosphate linkages of the previous couplings was a major problem. As a result, it was necessary to follow a very arduous multi-step purification process in which the branched contaminants were removed. However, as the oligonucleotide length increased, the percentage of branching also increased, making purification even more challenging. The solution phase chemistry made the process very slow, because the oligonucleotide had to be purified or precipitated between steps to remove excess reagents. When one considers the magnitude of the task, the accomplishment of preparing an active tRNA molecule becomes even more remarkable.

Khorana's most lasting contribution, however, was in the area of nucleoside protecting groups. The key to developing an efficient, cyclic, step-wise synthesis is a good protecting group scheme that allows the selective removal of a specific protecting group at the desired time. To make matters more challenging, the protecting groups must be removable almost quantitatively. Otherwise, the yield of desired product will be low and the product itself may be irresolvable from contaminants. To raise the bar even further, purines are susceptible to depurination under mildly to moderately acidic conditions (pH 4-5 for extended periods, pH 1-3 for fairly short periods), so strong acids should be avoided.

The solution Khorana offered for 5' hydroxyl protection, the dimethoxytrityl (DMT) protecting group (Smith, et. al., 1961), is ubiquitous in oligonucleotide chemistry today. The combination of good general stability and easy removal with mild acid has been unbeatable. Several options are available, such as leuvenyl and fluorenylmethyloxycarbonyl (FMOC), but none are as popular as the unique trityl family.

The reason this triphenyl methyl ether cleaves so readily under acidic conditions lies in the fact it is one of the few molecules that actually likes to form a carbocation. The back bonding of the pi electron cloud system formed by the three phenyl groups is sufficient to allow the methyl carbon to remain stable as a positively charged species under very mildly acidic conditions. Like many carbocations, the trityls have a distinctive color when ionized, which has turned into an extremely useful diagnostic tool. The dimethoxytrityl (DMT) carbocation has a very strong orange color in mild acid that has a high extinction coefficient, which means that even at very low concentrations it can still be accurately measured optically. The monomethoxytrityl (MMT) carbocation has a yellow color, while the parent trityl (Tr) itself is deep red (see #tritylabsorbance). The efficiency of each cycle of nucleoside addition can be allowed by measuring the absorbance of the released DMT and comparing it to the previous step.

The Phosphotriester Approach (Letsinger)

The phosphotriester approach (Letsinger) uses protected phosphate triesters with aryl or alkyl groups that are removed after chain assembly, developed by Robert Letsinger at Northwestern University as an improvement over phosphodiester chemistry.

The phosphotriester approach, pioneered by Robert Letsinger in the 1960s and advanced by Colin Reese in the 1970s, represents a pivotal evolution in oligonucleotide synthesis by employing pentavalent phosphorus(V) chemistry from the outset, utilizing neutral, fully protected phosphate triester monomers to facilitate condensation in aprotic organic solvents, thereby circumventing the aqueous incompatibilities and side reactions plaguing the earlier phosphodiester method of Khorana, which relied on direct P(V) activation of phosphomonoesters prone to hydrolysis, pyrophosphate formation, and N-acyl migration. In this methodology, 5'-hydroxyl-terminated oligonucleotides (with nucleobases bearing standard acyl protections such as benzoyl for adenine/cytosine and isobutyryl for guanine, and 5'-O-monomethoxytrityl or similar acid-labile groups) are condensed with incoming 3'-O-protected nucleoside phosphorotriesters—wherein the phosphate bears two non-bridging protecting groups, typically an aryl moiety like 2-chlorophenyl (removable via base-catalyzed hydrolysis or sulfide reduction) and a smaller alkyl such as methyl or (later in the phosphoramidite era) 2-cyanoethyl for orthogonal deprotection—via activation of the triester phosphorus to an electrophilic species using condensing agents like mesitylenesulfonyl tetrazole, 2,4,6-triisopropylbenzenesulfonyl chloride, or arylsulfonyl azoles (or possibly other related sulfonyl azoles like MSNT?), enabling nucleophilic displacement at the 3'-position to form a new internucleotide phosphate triester linkage with coupling yields of 90-95% per step, though limited by steric bulk and diastereomeric mixtures at phosphorus. Unlike the phosphite triester (phosphoramidite) method, which constructs the backbone through transient trivalent P(III) phosphite triesters subsequently oxidized to P(V) phosphates for enhanced reactivity and scalability, the phosphotriester strategy maintains P(V) throughout chain assembly, offering inherent stability against autoxidation but necessitating rigorous post-synthetic deprotection protocols (e.g., removal of aryl groups using oximolysis (like with aldoximes) or thiophenolate-based methods, or β-elimination for cyanoethyl) to unmask the anionic phosphodiester backbone, alongside global removal of exocyclic and 2'-O protections via ammonolysis; this approach, while enabling solid-phase adaptations in the 1970s, yielded to phosphoramidite dominance due to slower kinetics and greater susceptibility to β-elimination side products, yet it laid foundational principles for modified backbone synthesis, including phosphorothioates via thioester intermediates, and remains relevant for specialized applications requiring pre-oxidized phosphate scaffolds.

((Note: The hallmark of the phosphotriester era was predominantly aryl (and related) phosphate protections. 2‑cyanoethyl became the dominant non‑bridging phosphate protecting group with the rise of phosphoramidite chemistry; while CE can be used in triester contexts, it was not “typical” of classic triester synthesis.))

Developed by Robert Letsinger and subsequently refined by Colin Reese to resolve the solubility complications caused by charged ions in the earlier phosphodiester method, this chemical approach preserves the phosphorus atom in its pentavalent oxidation state. However, it disguises the negatively charged oxygen atom by attaching a temporary protecting group. This group is usually an aryl moiety—such as a 2-chlorophenyl or 2,5-dichlorophenyl group—or an alkyl group like trichloroethyl.

The fundamental building block used in this process is a nucleoside three-prime phosphate diester. This molecule is chemically activated by "super-condensing agents," specifically compounds like 1-(mesitylene-2-sulfonyl)-3-nitro-1,2,4-triazole or 2,4,6-triisopropylbenzenesulfonyl chloride, in the presence of nucleophilic catalysts such as N-methylimidazole. This reaction generates a highly energetic sulfonyl-phosphate triester intermediate that is primed to accept a chemical bond from the hydroxyl group located at the five-prime position of the target molecule.

The resulting product is a fully protected phosphate triester that carries no net electrical charge and dissolves easily in lipids. This lipophilic nature allows the synthesis to proceed using standard organic solvents, such as dichloromethane and chloroform, while effectively preventing the formation of unwanted pyrophosphate branches. However, the coupling proceeds via an associative bimolecular nucleophilic substitution mechanism at the phosphorus center rather than a direct displacement. This process requires the nucleophile to attack the tetrahedral phosphorus to form a crowded pentacoordinate (trigonal bipyramidal) transition state, a geometric constraint that contributes to reaction kinetics significantly slower than those observed in trivalent phosphorus chemistry. Furthermore, removing the protective groups after synthesis presents a significant challenge; the aryl protecting groups typically generally require oximolysis to be removed. This process employs reagents such as the syn-isomer of 2-nitrobenzaldoxime or pyridine-2-aldoxime ions to trigger a nucleophilic attack on the phosphorus atom, thereby causing the structure to fragment into the desired phosphodiester.

Colin Reese had a central role in developing aryl-protected triester chemistry which was important for phosphotriester method development. While Letsinger introduced the phosphotriester concept, the robust aryl phosphate protecting groups (2-chlorophenyl, etc.) and the sulfonyl azole coupling agents that made the method practical were worked out by Colin Reese’s group in the 1970s.

In the late 1960s, Letsinger published the first paper on the phosphotriester method of oligonucleotide synthesis (Letsinger, et. al., 1969). The key advance of this method was the protection of the phosphate group to prevent the branching that plagued the phosphodiester approach. The protecting group most commonly used was the beta-cyanoethyl group that is easily removed with ammonium hydroxide (Letsinger and Ogilvie, 1969). An o-chlorophenyl was also used but it required a more complicated deprotection mixture. It turned out, however, that the key to pushing the efficiency of the reaction, which reached levels in excess of 95% per step, was the selection of a proper activator. Mesityl sulfonyl chloride (MSCl) and mesityl sulfonyl nitrotriazole (MSNT) were by far the most popular (Devine and Reese, 1986; Letsinger and Ogilvie, 1969).

This was the first chemistry that was simple enough to reproduce successfully in many labs. The combination of a chemistry that worked relatively easily with solid phase methodology led to the creation of the first viable automated and semi-automated DNA synthesizers, exemplified by the early instruments developed by Vega Biotechnologies. Another early entrant was Ron Cook and his company Biosearch. He introduced the SAM I in the late 1970s, which was based on phosphotriester chemistry and was the most popular instrument of its era. These instruments allowed non-chemists to prepare simple oligonucleotides, and created the ability to probe genes with radio-labeled oligonucleotides prepared with the exact sequence desired. Thus equipped, the industry was primed for the emerging techniques of gene mapping, polymerase chain reaction (PCR), and target validation.

However, the phosphotriester chemistry still suffered from critical drawbacks. Among them was the fact that despite the years of work by a number of research groups, the average step-wise efficiency could never reproducibly be raised above 97%, and often failed to reach 95%. This limited the method to the routine synthesis of oligonucleotides less than 20 bases in length. Another problem was the extensive coupling time, which resulted in cycle times that commonly ran longer than an hour and a half.

The phosphotriester method, initially developed for the solution-phase synthesis, was also implemented on low-cross-linked "popcorn" polystyrene, and later on solid support material controlled pore glass (CPG), which then initiated a massive research effort in solid-phase synthesis of oligonucleotides and eventually led to the automation of the oligonucleotide chain assembly.

SES group

The 2-(trimethylsilyl)ethanesulfonyl (SES) group has been used as a phosphate-protecting group in oligonucleotide synthesis, offering orthogonality to other protecting groups through its fluoride lability. The SES group can be removed with fluoride sources like TBAF, providing an alternative to the standard β-cyanoethyl (CE) protecting group, though CE remains dominant due to its compatibility with ammonia deprotection and lower cost.

SES protecting group is quite niche compared to 2-cyanoethyl.

The Solid-Phase Triester Approach (Itakura-Riggs)

The operationalization of phosphotriester chemistry into a robust solid-phase platform was achieved in the mid-1970s through the seminal work of Keiichi Itakura and Arthur Riggs at the City of Hope National Medical Center, a methodology that enabled the first total chemical synthesis of a functional gene, specifically that encoding the hormone somatostatin. Merging the principles of Letsinger’s neutral backbone strategy with the solid-support concepts pioneered by Merrifield for peptides, this approach utilized a copolymer of styrene and divinylbenzene as the stationary phase, to which the 3'-terminal nucleoside was tethered via a succinate linker. The synthesis relied on the stepwise addition of nucleoside 3'-(p-chlorophenyl)phosphate diesters, which were activated in situ using powerful arylsulfonyl azoles, such as 2,4,6-triisopropylbenzenesulfonyl chloride (TPS-Cl) or related sulfonyl azoles such as 1-(mesitylene-2-sulfonyl)-3-nitro-1,2,4-triazole (MSNT). These condensing agents were critical for overcoming the kinetic sluggishness of the pentavalent phosphorus center within the heterogeneous matrix of the solid support, as they generated highly reactive sulfonyl triazole intermediates that were less prone to sulfonating the 5'-hydroxyl nucleophile than the sulfonyl chlorides used in solution phase. The method utilized dimethoxytrityl groups for 5'-protection, establishing the protic acid deprotection cycles that remain standard today, while the internucleotide p-chlorophenyl protecting groups were retained until the final cleavage step to suppress backbone hydrolysis. Although the coupling times remained significant—often requiring 30 to 60 minutes per nucleotide to drive the second-order nucleophilic substitution to completion—and necessitated extensive high-performance liquid chromatography purification to remove truncated failure sequences, this technology provided the first scalable route to synthetic DNA. This culminated in the successful assembly of the 14-amino acid somatostatin gene and subsequently the human insulin gene, fundamentally launching the era of recombinant DNA technology and modern biotechnology before the eventual dominance of phosphoramidite chemistry.

The H-Phosphonate Approach

The H-phosphonate approach to oligonucleotide synthesis, pioneered by Froehler and Matteucci, represents a distinct mechanistic paradigm compared to phosphoramidite chemistry, primarily characterized by the deferral of phosphorus oxidation until the completion of chain assembly. Central to this methodology is the use of nucleoside 3'-H-phosphonate monoesters. Contrary to erroneous reports suggesting oxidative synthesis via sulfuryl chloride, these monomers are correctly prepared through the phosphitylation of 5'-O-protected nucleosides using reagents derived from phosphorus trichloride and limited base (typically imidazole) or via transesterification with diphenyl phosphite, followed by mild aqueous hydrolysis. This sequence yields the stable H-phosphonate monoester anion.

H‑phosphonate monoesters are tetracoordinate P(V) species. While they are often discussed in the context of P(III) due to the presence of the P-H bond and their synthetic origin, structurally they exist as the phosphonate tautomer.

(The oxidation-state / tautomerism picture is controversial in the literature. Many authors treat H-phosphonates as P(III) derivatives with a P–H bond and describe a phosphite ↔ phosphonate tautomeric equilibrium; others lean toward a dominant P(V) depiction.)

The H-phosphonate structure is:

    O
    ‖
RO–P–H
    |
    O⁻ (or OH)

This is a tetra-coordinate phosphorus with: - One P-H bond - One P-O(R) bond - One P=O bond - One P-O⁻ (or P-OH) bond

The H-phosphonate monoester adopts a tetrahedral geometry. The phosphite-phosphonate tautomerism involves a rare phosphite form—P(OR)(OH)₂—which is truly tricoordinate with a lone pair. However, the equilibrium overwhelmingly favors the dominant H-phosphonate form (HO)(R-O)P(=O)H. Unlike the tricoordinate phosphite, the stable H-phosphonate is a tetracoordinate P(V) species where the "lone pair" has formed the phosphoryl (P=O) bond. Therefore, the phosphorus center is saturated with four substituents and does not possess a stereochemically active lone pair in its ground state.

The synthetic cycle relies on the chemoselective activation of the H-phosphonate monoester using sterically hindered acid chlorides, such as pivaloyl chloride or adamantoyl chloride, in solvents like pyridine or acetonitrile/pyridine mixtures. This reaction generates a highly reactive mixed carboxylic-phosphonic anhydride (or an acyl-phosphonate intermediate). This electrophile undergoes nucleophilic substitution by the 5'-hydroxyl group of the support-bound growing chain. The resulting internucleotide linkage is a dinucleoside H-phosphonate diester. Crucially, the phosphorus-hydrogen bond provides sufficient steric and electronic protection to the phosphorus center, rendering it resistant to the coupling conditions and eliminating the need for transient protection of the internucleotide phosphate oxygen.

The hydrolytic stability of the P-H bond permits the accumulation of H-phosphonate diester linkages throughout chain elongation, bypassing the need for iterative oxidation cycles. Upon completion of the sequence, a single conversion step determines the final backbone structure, offering a unique versatility unavailable in phosphoramidite chemistry. For the generation of native phosphodiesters, oxidation is typically achieved using iodine in aqueous pyridine, where water serves as the essential nucleophilic oxygen donor. Conversely, the backbone can be modified under strictly anhydrous conditions to create analogues: sulfurization with elemental sulfur yields phosphorothioates, while oxidative amidation—utilizing Atherton-Todd conditions with carbon tetrachloride and primary or secondary amines—produces phosphoramidates. This capacity for divergent functionalization from a common H-phosphonate precursor makes the method unexcelled for the synthesis of complex, phosphate-modified oligonucleotides.

While the coupling kinetics are generally slower (5–30 minutes) compared to phosphoramidite chemistry, and the reagents are susceptible to hydrolytic degradation if not strictly anhydrous, the H-phosphonate method remains unexcelled for the synthesis of phosphate-modified backbones and analogues where orthogonal protection strategies would otherwise be prohibitive.

The Phosphite Triester Approach (chloridite chemistry) (Letsinger)

Letsinger (1976). The first use of highly reactive P(III) chloridites, drastically reducing coupling time, though the reagents were unstable. In the 1970s, substantially more reactive P(III) derivatives of nucleosides, 3'-O-chlorophosphites, were successfully used for the formation of internucleosidic linkages, which then lead to the phosphite triester method.

The phosphite triester method, introduced by Letsinger, represents the first successful exploitation of the reaction kinetics of trivalent phosphorus. Reagents are synthesized by reacting a nucleoside protected at the 5-prime position with phosphorus trichloride or an installable alkyl dichlorophosphite at low temperature to generate a nucleoside phosphorochloridite. The chlorine atom renders the trivalent phosphorus center intensely electrophilic. Upon addition to a nucleoside with a free 5-prime hydroxyl group and a tertiary amine base, coupling occurs almost instantaneously via nucleophilic substitution, forming a dinucleoside phosphite triester. This intermediate is distinct from the phosphate found in the natural backbone in that the trivalent phosphorus atom renders the internucleotide linkage chemically unstable and highly susceptible to acid cleavage. Therefore, it necessitates immediate oxidation—typically using a mixture of elemental iodine, water, and a base—to the pentavalent phosphorus state to stabilize the backbone against hydrolysis during subsequent cycles. While this validated the speed of trivalent phosphorus chemistry, the phosphorochloridite monomers are extremely hygroscopic and susceptible to hydrolysis and disproportionation, rendering them unsuitable for storage or use in automated delivery systems.

(The phosphotriester approach, which utilizes pentavalent phosphorus with protecting groups on all three oxygens from the onset, represents Letsinger's earlier methodology and is fundamentally different from approaches utilizing trivalent phosphorus intermediates. While both the original phosphite triester method (using chlorophosphites) and the later phosphoramidite chemistry share the strategy of using trivalent phosphorus that is subsequently oxidized to pentavalent phosphate, they are distinct technologies. Both capitalize on the fact that trivalent phosphorus reacts orders of magnitude faster than pentavalent species; however, modern automated synthesis specifically employs phosphoramidites rather than the unstable chlorophosphites used in the initial phosphite triester approach.)

In the mid-1970s, Letsinger published the first papers describing the phosphite-triester method of oligonucleotide chemistry (Letsinger, et. al., 1975; Letsinger and Lunsford, 1976). This chemistry is based on the use of reactive phosphorus in the phosphorus(III) state, instead of the classic phosphorus(V) phosphoryl chemistry. The scheme required an additional step in the synthesis cycle, oxidation, in order to prepare the natural phosphorus(V) backbone. The major advantage of this chemistry was the significant reduction in time required for coupling due to the highly reactive nature of the nucleoside phosphomonochloridite intermediate.

The fact that the phosphorus(III) intermediate is more reactive than the phosphorus(V) species is not intuitive. One would suspect, in the absence of data, that because of the doubly bonded oxygen, the phosphorus(V) would be more reactive to attack by a nucleophile based on its enhanced electronegativity. However, the determining factor of the reaction rate turns out to be the difference in the energy of formations for the transitional intermediates of the phosphorus(III) species versus phosphorus(V). As shown in #trigonal-bipyrimidal-intermediate, a trigonal bipyrimidal intermediate is formed. The doubly bonded oxygen hinders the transition from the tetrahedron configuration into the planar much more than the lone pair of electrons.

Oxidation of the phosphite-triester intermediate into a phosphotriester was needed in order to stabilize the backbone. This oxidation was required at each step of the cycle because of the instability of the phosphite-triester intermediate to the acid required to remove the DMT group. Fortunately, a very simple mixture of iodine, water and some base very efficiently and quantitatively oxidizes phosphorus within seconds.

The research community was quick to accept this new chemistry as a significant step forward. Not only could standard DNA be prepared faster, but the door was opened for the investigation into a variety of backbone modified oligonucleotides. Biologics, a company partially comprised of former Letsinger students, marketed an automated synthesizer based on this chemistry and another was in development by Vega Biotechnologies.

The early form of the phosphite-triester chemistry did indeed have major drawbacks. The most significant problem was the highly reactive nature of the nucleoside phosphomonochloridite intermediate. It was very susceptible to hydrolysis. The intermediate was not easy to store and therefore was best made just prior to each coupling. Another issue was that the formation of active intermediate was very tricky. The phosphodichloridite activating reagent had to be added to the 5' protected nucleoside in such a manner as to maximize the formation of desired intermediate while reducing the formation of 3'-3' dimer (fig9). The formation of this side-product did double damage in that it reduced the amount of desired material and increased the amount of unused phosphodichloridite that remained in solution. This unused reagent would very efficiently cap off the growing chain before the desired intermediate had time to couple. That was the reason that an excess of the reagent could not be used to reduce formation of the 3'-3' adduct. Using too few equivalents of the phosphodichloridite had a like-wise harmful effect in that too much 3'-3' adduct would be formed, reducing the concentration of active nucleoside reagent below a critical threshold. Increasing the concentration of the reagents to combat that only led to the opposite effect and an even less controllable reaction.

The protocols designed to optimize this reaction called for the slow addition of a very slight excess of solubilized 5' protected nucleoside to a solution of RO-PCl2 at extremely cold temperatures (-78 degrees C). As it turned out, the combination of the requirement for preparing the active reagent just prior to each coupling, and the need for arduous conditions during this activation, removed nearly all of the advantages brought about by the faster coupling time.

This problem was not solved until the early 1980s. A serendipitous discovery was made by a graduate student that showed if there was a rapid introduction of the phosphodichloridite reagent to the nucleoside at room temperature, it formed a useful active reagent without making too much of the 3'-3' adduct or leaving too much phosphodichloridite in the mix (Hogrefe, 1987). This improved method was later coupled with a scavenger system involving trityl alcohol that selectively removed any excess RO-PCl2 from the reaction mixture. It was this new protocol, which finally allowed the development of a practical automated DNA synthesizer with coupling times of 15 minutes or less. This instrument was also developed by Vega Biotechnologies in collaboration with Letsinger. Although this particular instrument was a significant improvement over the phosphotriester instruments described earlier, it was never sold. The phosphodichloridite method was soon eclipsed by a new chemistry discovered by Marvin Caruthers, the phosphoramidite method. It solved many of the problems that clouded the entry of the phosphodichloridite method into the market.

Caruthers took the advantage of less aggressive and more selective 1H-tetrazolidophosphites and implemented the phosphite triester method in solid phase (1981) with Matteucci. Shortly after this work, Caruthers made further improvements by using a 2-cyanoethyl phosphite-protecting group in place of a less user-friendly methyl group to make nucleoside phosphoramidites.

The Phosphoramidite Approach (Caruthers)

The phosphoramidite approach to oligonucleotide synthesis, conceptualized by Serge Beaucage and Marvin Caruthers in the late 1970s and refined for automated solid-phase implementation by the early 1980s, stands as the cornerstone of modern nucleic acid assembly. This methodology revolutionized the field by exploiting the distinct chemo-physical properties of trivalent phosphorus, specifically shifting away from the hard-to-handle chlorophosphites of earlier eras to the stable yet reactive N,N-dialkylphosphoramidite monomers. The fundamental unit of this chemistry is the nucleoside 3'-O-(2-cyanoethyl-N,N-diisopropyl)phosphoramidite. The brilliance of this molecular design lies in the diisopropylamino moiety, which stabilizes the phosphorus atom toward air and moisture under neutral conditions through significant steric shielding and the electron donation inherent in the P-N bond (but previously it was believed that the diisopropylamino group stabilizes P(III) via p‑π to d‑π orbital donation which is now an outdated belief). This stability allows for the robust storage and handling of monomers while maintaining a high susceptibility to protonation that is leveraged for activation during coupling. All of these reagents are handled in rigorously anhydrous conditions, like the other methods.

The synthetic cycle proceeds on a solid support, such as controlled pore glass or polystyrene, beginning with the selective deprotection of the 5'-dimethoxytrityl (DMT) group using dilute acid, typically trichloroacetic acid in dichloromethane. This releases a chromophoric trityl cation—often monitored at 498 nm to calculate stepwise yields—and exposes a free 5'-hydroxyl nucleophile. Coupling is initiated by introducing the 5'-DMT-protected phosphoramidite monomer in the presence of a weak acid activator. While 1H-tetrazole (pKa approximately 4.9) was the historical standard, sterically hindered sequences or specific secondary structures often necessitate more potent activators like 5-ethylthio-1H-tetrazole or 4,5-dicyanoimidazole. The activator protonates the aminophosphine nitrogen, converting the stable diisopropylamine into a facile leaving group and generating a highly electrophilic tetrazolyl-phosphite intermediate. The 5'-hydroxyl of the support-bound oligonucleotide then attacks this intermediate via associative nucleophilic substitution at the chiral phosphorus center, yielding a phosphite triester linkage. To ensure sequence fidelity, any unreacted 5'-hydroxyl species are immediately "capped" via acetylation with acetic anhydride and N-methylimidazole, effectively terminating failure sequences and preventing "n-minus-1" deletion impurities.

Following coupling and capping, the internucleotide linkage exists as a labile P(III) phosphite triester, which is susceptible to acid-catalyzed scission. To secure backbone integrity, this intermediate is oxidized in situ to the stable pentavalent phosphate triester. The standard oxidation utilizes iodine dissolved in a mixture of tetrahydrofuran, pyridine, and water. Mechanistically, iodine attacks the phosphorus lone pair to form a P-I phosphonium adduct (this is a bit simplified and incorrect, but basically there is a standard iodine/water/base oxidation), which is subsequently displaced by water to yield the phosphate. In sophisticated variants, such as the stereocontrolled synthesis of therapeutic antisense oligonucleotides, this oxidation step is replaced by sulfurization using reagents like 3H-1,2-benzodithiol-3-one to generate phosphorothioate backbones. Crucially, the phosphate remains protected by the 2-cyanoethyl group throughout the cycle. This specific protecting group was a significant innovation over simpler alkyls, as it facilitates clean removal via a beta-elimination mechanism (retro-Michael reaction) during the final ammonolysis deprotection. The advantage of CE is that it is cleanly removable under basic conditions via β‑elimination without harsh reagent (and the problem with earlier protections was harsh deprotection conditions and side reactions).

((For the P-I phosphonium adduct, a more satisfying mechanistic explanation would clarify that the reaction is driven by the inherent nucleophilicity of the trivalent phosphite, which attacks the diatomic iodine to generate a discrete iodophosphonium salt intermediate rather than a distinct P-I adduct. In this highly electrophilic cationic species, the phosphorus atom is primed for reaction with nucleophiles; water, present in the solvent mixture, attacks the phosphorus center to displace the iodide ion. To drive this equilibrium forward, the pyridine acting as the solvent base plays a dual thermodynamic role: it may stabilize the initial active iodine species or intermediate, but more critically, it acts as a proton scavenger to neutralize the hydrogen iodide generated during the hydrolysis of the iodophosphonium salt. This deprotonation facilitates the final collapse of the intermediate, irreversibly establishing the thermodynamically stable phosphorus-oxygen double bond characteristic of the pentavalent phosphate triester.))

This highly optimized chemistry achieves stepwise yields routinely exceeding 99%, allowing for the synthesis of DNA and RNA oligonucleotides up to 200 mers with high fidelity. The method accommodates a vast array of orthogonal protection schemes, such as tert-butyldimethylsilyl for the 2'-hydroxyl in RNA or base-labile acetyl, benzoyl, and isobutyryl groups on the exocyclic amines of nucleobases. Furthermore, the approach supports the incorporation of sensitive modifications, including 2'-fluoro substituents, locked nucleic acids, and azobenzene linkers, typically validated by anion-exchange HPLC and mass spectrometry. By resolving the historical challenges of P(III) instability through the ingenious use of amine leaving groups and rapid oxidation cycles, the Beaucage-Caruthers methodology provides the scalability and functional group tolerance required for the industrial production of primers, therapeutic siRNAs, and aptamers.

video: The bumbling chemist on phosphoramidite chemistry

video: Caruthers lecture on phosphoramidite chemistry including some history (2018) and a transcript of Caruthers' talk.

Nucleoside phosphoramidites

Naturally occurring nucleotides (nucleoside-3'- or 5'-phosphates) and their phosphodiester analogs are insufficiently reactive to afford an expeditious synthetic preparation of oligonucleotides in high yields. The selectivity and the rate of the formation of internucleosidic linkages is dramatically improved by using 3'-O-(N,N-diisopropyl phosphoramidite) derivatives of nucleosides (nucleoside phosphoramidites) that serve as building blocks in phosphite triester methodology. To prevent undesired side reactions, all other functional groups present in nucleosides have to be rendered unreactive (protected) by attaching protecting groups. Upon the completion of the oligonucleotide chain assembly, all the protecting groups are removed to yield the desired oligonucleotides.

The 5'-hydroxyl group is protected by an acid-labile DMT (4,4'-dimethoxytrityl) group.

Thymine and uracil, nucleic bases of thymidine and uridine, respectively, do not have exocyclic amino groups and hence do not require any protection.

Although the nucleic base of guanosine and 2'-deoxyguanosine does have an exocyclic amino group, its basicity is low to an extent that it does not react with phosphoramidites under the conditions of the coupling reaction. However, a phosphoramidite derived from the N2-unprotected 5'-O-DMT-2'-deoxyguanosine is poorly soluble in acetonitrile, the solvent commonly used in oligonucleotide synthesis. contrast, the N2-protected versions of the same compound dissolve in acetonitrile well and hence are widely used. Nucleic bases adenine and cytosine bear the exocyclic amino groups reactive with the activated phosphoramidites under the conditions of the coupling reaction. By the use of additional steps in the synthetic cycle or alternative coupling agents and solvent systems, the oligonucleotide chain assembly may be carried out using dA and dC phosphoramidites with unprotected amino groups. However, in routine oligonucleotide synthesis, exocyclic amino groups in nucleosides are kept permanently protected over the entire length of the oligonucleotide chain assembly.

The protection of the exocyclic amino groups has to be orthogonal to that of the 5'-hydroxy group because the latter is removed at the end of each synthetic cycle. The simplest to implement, and hence the most widely used, strategy is to install a base-labile protection group on the exocyclic amino groups. Most often, two protection schemes are used.

Bz (benzoyl) protection is used for A, dA, C, and dC, while G and dG are protected with isobutyryl group. More recently, Ac (acetyl) group is used to protect C and dC.

In a different scheme, a mild protection scheme, A and dA are protected with isobutyryl or phenoxyacetyl groups (PAC). C and dC bear acetyl protection, and G and dG are protected with 4-isopropylphenoxyacetyl (iPr-PAC) or dimethylformamidino (dmf) groups. Mild protecting groups are removed more readily than the standard protecting groups. However, the phosphoramidites bearing these groups are less stable when stored in solution.

The phosphite group is protected by a base-labile 2-cyanoethyl protecting group. Once a phosphoramidite has been coupled to the solid support-bound oligonucleotide and the phosphite moieties have been converted to the P(V) species, the presence of the phosphate protection is not mandatory for the successful conducting of further coupling reactions.

In RNA synthesis, the 2'-hydroxy group is protected with TBDMS (t-butyldimethylsilyl) group or with TOM (tri-iso-propylsilyloxymethyl) group, both being removable by treatment with fluoride ion.

The phosphite moiety also bears a diisopropylamino (iPr2N) group reactive under acidic conditions. Upon activation, the diisopropylamino group leaves to be substituted by the 5'-hydroxy group of the support-bound oligonucleotide.

Phosphoramidite Chemistry with Alternative Activators

Subsequent optimization of the standard Caruthers cycle (e.g., moving from tetrazole to stronger activators like ETT).

While 1H-tetrazole served as the standard activator for phosphoramidite coupling throughout the 1980s, the demand for higher coupling efficiencies—particularly for sterically hindered modifications like RNA (2'-O-TBDMS), LNA, or long-chain synthesis—necessitated the development of more potent activators. The limitation of 1H-tetrazole lies in its modest acidity (pKa approximately 4.9) and its crystalline nature, which can lead to precipitation in automated fluidics lines.

To enhance the reaction kinetics, the field moved toward activators with lower pKa values (higher acidity) and improved nucleophilicity to accelerate the formation of the reactive protonated phosphoramidite-activator complex. Prominent among these is 5-ethylthio-1H-tetrazole (ETT) (pKa approximately 4.3) and 5-benzylthio-1H-tetrazole (BTT), which offer superior solubility in acetonitrile and faster activation rates for RNA synthesis. A structurally distinct class of activator is 4,5-dicyanoimidazole (DCI). Although less acidic (pKa approximately 5.2) than tetrazole, DCI acts as a highly effective nucleophilic catalyst, displacing the diisopropylamine to form a reactive imidazolium-phosphite intermediate without the risk of acidic depurination or explosive hazards associated with tetrazoles. Furthermore, pyridinium salts, such as pyridinium hydrochloride or imidazolium triflate, function by pure acid catalysis and have been utilized in specific contexts, particularly for the activation of cyclic phosphoramidites or in coupling reactions where non-nucleophilic conditions are preferred to avoid side reactions with electrophilic centers on modified nucleobases.

In modern solid-phase synthesis, the 5'-O-dimethoxytrityl (DMT) group is the standard temporary protecting group.

For accurate information on modern RNA synthesis, consult literature on 2'-O-TBDMS RNA phosphoramidites with standard 5'-DMT protection and β-cyanoethyl phosphate protection, or specialized methods like 2'-ACE or 2'-TOM chemistry.

The Methylphosphonate Approach

This methodology is distinct because it generates a non-ionic, isosteric DNA analogue characterized by a direct phosphorus-carbon bond. The synthesis cannot utilize standard phosphorylating agents; instead, it employs methyldichlorophosphine or methylphosphonamidites as the starting source of phosphorus. This phosphorus-carbon bond is robust and withstands all standard biological and chemical degradation pathways.

The coupling process involves the reaction of a nucleoside methylphosphonamidite protected at the five-prime position (activated by the clinical agent tetrazole) with the five-prime hydroxyl group of the oligonucleotide chain bound to the solid support. This reaction forms a methylphosphonite intermediate containing a trivalent phosphorus atom. This intermediate is subsequently oxidized—typically utilizing tert-butyl hydroperoxide or elemental iodine—to create the final methylphosphonate linkage containing a pentavalent phosphorus atom.

A critical chemical feature is that the resulting phosphorus atom constitutes a chiral center, meaning it can exist in either the R-phosphorus or S-phosphorus configuration. Because standard chemical synthesis techniques are not stereoselective, an oligomer of length "n" will exist as a complex mixture of two raised to the power of "n" diastereomers. The isomers possessing the R-phosphorus configuration generally base-pair with ribonucleic acid (RNA) with higher affinity than those with the S-phosphorus configuration. Furthermore, the electrically neutral nature of the backbone significantly alters solubility—necessitating the use of hydrophobic interaction chromatography for purification—and modifies its cellular uptake properties.

A methylphosphonate linkage (where the phosphorus is bonded to a methyl group, a doubly bonded oxygen, and two different nucleoside oxygens) is chiral. Because standard synthesis does not control this stereochemistry, the resulting oligonucleotide is indeed a racemate (a mixture of diastereomers). This implies that they actually have massive diastereomeric complications.

Thiophosphoramidite Chemistry (Eckstein and colleagues)

The thiophosphoramidite chemistry, often synonymous in practical application with the synthesis of phosphorothioate oligonucleotides via modified phosphoramidite protocols, was systematically advanced by Fritz Eckstein and colleagues in the 1980s to address the rapid enzymatic degradation of natural phosphodiester DNA in biological milieus. This methodology departs from the canonical synthesis during the oxidation step; rather than converting the transient internucleotide phosphite triester P(III) intermediate into a phosphate diester using aqueous iodine, the protocol employs a sulfurizing agent to install a sulfur atom at the non-bridging position, yielding a chemically robust P(V) phosphorothioate linkage.

The pioneering protocols realized that the trivalent phosphorus of the phosphite triester is a soft nucleophile capable of attacking elemental sulfur (S8), historically dissolved in a mixture of carbon disulfide and pyridine or carbon disulfide and lutidine to facilitate solubility and kinetics. However, due to the flammability and toxicity of carbon disulfide, the field evolved to utilize more efficient, soluble, and rapid sulfur-transfer reagents such as 3H-1,2-benzodithiol-3-one 1,1-dioxide (Beaucage reagent), tetraethylthiuram disulfide (TETD), or phenylacetyl disulfide (PADS). These reagents function by delivering an electrophilic sulfur species to the P(III) center, affecting oxidative sulfurization with high conversion rates consistent with automated synthesis cycles.

A defining chemical consequence of this modification is the creation of a chiral center at the phosphorus atom, as the non-bridging sulfur and oxygen atoms render the phosphate non-prochiral. Consequently, the synthesis of a phosphorothioate oligonucleotide of length n results in a racemate mixture of 2 to the power of n diastereomers, comprising both Rp and Sp absolute configurations at each linkage. Eckstein's work was instrumental in characterizing that these stereoisomers possess distinct physicochemical properties; generally, the Rp diastereomer tends to pair (in a sequence-dependent manner) more faithfully with complementary RNA and mediates RNase H recruitment more effectively, while the Sp isomer displays greater resistance to specific nucleases. Although standard solid-phase synthesis produces a random stereochemical mixture (stereorandom), the introduction of the phosphorothioate backbone was the critical breakthrough that enabled the first generation of antisense nucleotide therapeutics (such as Fomivirsen) by conferring sufficient metabolic stability to survive serum nucleases and enter cells, largely due to the increased lipophilicity. Phosphorothioate linkages increase nonspecific protein binding and this high protein binding is a defining pharmacological feature of phosphorothioate oligos and contributes to their in vivo distribution and half-life.

Peptide Nucleic Acids (PNA)

Introduced by Peter Nielsen and colleagues in 1991, Peptide Nucleic Acids (PNA) constitute a DNA mimic in which the entire sugar-phosphate backbone is replaced by an achiral, uncharged pseudopeptide scaffold composed of repeating N-(2-aminoethyl)glycine units. The nucleobases are attached to the secondary amine of the backbone via a methylenecarbonyl (acetyl) linker. Despite this profound chemical alteration, PNA retains the ability to hybridize with complementary DNA and RNA with remarkable specificity and thermal stability, adhering to Watson-Crick base-pairing rules.

From a synthetic perspective, PNA oligomerization is indistinguishable from solid-phase peptide synthesis and utilizes Fmoc (or occasionally Boc) chemistry rather than phosphoramidite nucleotide chemistry. The monomers are N-protected amino acids containing the nucleobase side chain. Synthesis proceeds via the activation of the monomer's carboxylic acid using peptide coupling reagents such as HATU (1-[Bis(dimethylamino)methylene]-1H-1,2,3-triazolo[4,5-b]pyridinium 3-oxid hexafluorophosphate) or HBTU in the presence of a base like diisopropylethylamine, followed by nucleophilic attack by the terminal primary amine of the growing chain. Because the backbone is strictly non-ionic, PNA/DNA duplexes lack the inter-strand electrostatic repulsion found in DNA/DNA duplexes, leading to significantly high melting temperatures that are largely independent of salt concentration. However, this extreme affinity and poor water solubility often necessitate the conjugation of solubility-enhancing amino acids (like lysine) or the use of shorter oligomers.

Morpholino Oligonucleotides

(Replaces the sugar with a morpholine ring and phosphodiester with phosphorodiamidate.)

Developed by James Summerton and Dwight Weller in 1997 to address the limitations of charged phosphate backbones, Morpholino oligonucleotides (PMO) represent a radical structural redesign where the deoxyribose sugar is replaced by a six-membered morpholine ring, and the anionic phosphodiester linkage is substituted by a non-ionic phosphorodiamidate group. The synthesis of these oligomers avoids the standard phosphoramidite coupling cycle. Unlike earlier misconceptions suggesting a simple oxidative conversion of ribose, PMO monomers are actually produced via elaborate multistep synthetic routes designed to construct the six‑membered morpholine scaffold and subsequently install the phosphorodiamidate functionality.

Assembly typically proceeds on a solid support (such as aminomethyl polystyrene), where the operative coupling chemistry involves the reaction of a 5'-hydroxyl-bearing morpholino subunit with an incoming monomer featuring an activated chlorophosphorodiamidate or fluorophosphoramidate tail group. Because the backbone is uncharged at physiological pH, morpholinos eliminate the electrostatic repulsion between the synthetic oligomer and the target RNA, often resulting in high binding affinity despite the structural deviation from natural nucleic acids. Furthermore, the replacement of the ester linkage with a phosphorodiamidate bond and the sugar with a morpholine ring renders the backbone completely resistant to nucleases and generally incapable of recruiting RNase H, making them steric blocking agents rather than catalytic degraders of RNA.

Locked Nucleic Acids (LNA)

(A rigid bicyclic sugar modification.)

Locked Nucleic Acids (LNA), also known as Bridged Nucleic Acids (BNA), were pioneered by Jesper Wengel's group (and independently by Imanishi) in 1998. The defining chemical feature of the LNA monomer is a bicyclic furanose unit containing a 2'-O,4'-C-methylene bridge. This covalent tether structurally constrains (or "locks") the ribose sugar into the C3'-endo (North) conformation, which is the pucker characteristic of A-form RNA helices.

Unlike PNA or morpholinos, LNA synthesis is fully compatible with standard DNA/RNA automated synthesis. The monomers are phosphoramidites, specifically 5'-O-DMT-LNA-nucleoside-3'-O-(2-cyanoethyl-N,N-diisopropyl)phosphoramidites. Due to the pre-organized structure of the sugar, the entropic penalty of duplex formation is substantially reduced. Consequently, the incorporation of LNA monomers into an oligonucleotide increases the thermal melting temperature (Tm) of the duplex by anywhere from 2 to 8 degrees Celsius per modified nucleotide. This high affinity allows for the use of much shorter probes (typically 8-15 mers) for successful hybridization. The synthesis follows the standard tetrazole-mediated coupling and iodine oxidation cycles, although the bulky bicyclic system can occasionally require extended coupling times to overcome steric hindrance at the 3'-phosphorus center.

Other methods

other stuff to think about:

• phosphodiester approach (Khorana, 1950s-1960s)
• phosphite triester chemistry
• methylphosphonate approach
• phosphoramidite approach
• H-phosphonate approach
• 2'-O-TBDMS RNA phosphoramidites with standard 5'-DMT protection and β-cyanoethyl phosphate protection,
• 2'-ACE or 2'-TOM chemistry
• 2-(trimethylsilyl)ethanesulfonyl (SES) group has been used as a phosphate-protecting group in oligonucleotide synthesis

Phosphoramidite method for oligonucleotide synthesis

Here are some notes on oligonucleotide synthesis using phosphoramidite chemistry.

Acid/base activators in phosphoramidite oligonucleotide synthesis

The nucleotide to be coupled to the growing oligomer is often presented as a phosphoramidite containing a diisopropylamine leaving group and cyanoethyl and 4,4′-dimethoxytrityl (DMTr) as P–O and 5′ hydroxyl protecting groups.

The phosphoramidite coupling reaction is often incorrectly referred to as phosphorylation rather than phosphitylation. The phosphoramidite coupling reaction is the nucleophilic substitution of the amine moiety of the nucleosidic phosphoramidite by the 5′ hydroxy function of the solid support bound nucleoside. This reaction must be in the presence of a suitable acid/base activator because the reactants are otherwise inert.

Although it has been suggested that protonation of the phosphoramidite occurs on phosphorous, it seems likely that for a reaction to occur, nitrogen protonation is required. Evidence supporting this has been reported by Korkin and Tvetkov who showed, by molecular modelling of H2P–NH2, that protonation on nitrogen lengthens and weakens the phosphorous nitrogen bond, whereas phosphorous protonation shortens and strengthens the phosphorous nitrogen bond. Nurminen has claimed that phosphorous protonation could be achieved with strong acids, although subsequent nucleophilic substitution of the amine was much slower than with the corresponding nitrogen protonated species.

"Herein we report mechanistic studies on the phosphitylation of nucleosidic species in acetonitrile using saccharin and N-methylimidazole as an activator, which is currently used as part of the manufacture of oligonucleotides on an industrial scale." pdf

Synthesis cycle steps

The following steps are for the addition of each nucleotide to the growing oligos. It's not the complete set of steps. Each addition of a nucleotide requires the following steps.

1. Deblocking (detritylation)

Detritylation occurs using an acid such as 2% trichloroacetic acid (TCA) or 3% dichloroacetic acid (DCA), in an inert solvent (dichloromethane or toluene).

• produces an orange-colored DMT cation which needs to be washed out
• oligonucleotide precursor has a free 5'-terminal hydroxyl group
• using a stronger acid or conducting detritylation for an extended time, leads to depurination of the oligonucleotide

2. Coupling

A 0.02–0.2 M solution of nucleoside phosphoramidite (or a mixture of several phosphoramidites) in acetonitrile is activated by a 0.2–0.7 M solution of an acidic azole catalyst (below). The mixing is usually very brief and occurs in fluid lines of oligonucleotide synthesizers (see below) while the components are being delivered to the reactors containing solid support. The extension reaction forms a phosphite triester linkage (below).

The concentration of the activator is primarily determined by its solubility in acetonitrile and is irrespective of the scale of the synthesis.

Upon the completion of the coupling, any unbound reagents and by-products are removed by washing.