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Oligonucleotide synthesis

 
Wikipedia: Oligonucleotide synthesis
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Oligonucleotide synthesis is the chemical synthesis of relatively short fragments of nucleic acids with defined chemical structure (sequence). The technique is extremely useful in current laboratory practice because it provides a rapid and inexpensive access to custom-made oligonucleotides of the desired sequence. Whereas enzymes synthesize DNA and RNA in a 5' to 3' direction, chemical oligonucleotide synthesis is carried out in the opposite, 3' to 5' direction. Currently, the process is implemented as solid-phase synthesis using phosphoramidite method and A, C, G, T (2'-deoxy only), and U (ribo only) nucleoside phosphoramidites or 2'-deoxynucleoside phosphoramidites as building blocks. To obtain the desired oligonucleotide, the building blocks are sequentially coupled to the growing oligonucleotide chain in the order required by the sequence of the product (see Synthetic Cycle below). The process has been fully automated in the late 1970s. Upon the completion of the chain assembly, the product is released from the solid phase to solution, deprotected, and collected. The occurrence of side reactions sets practical limits for the length of synthetic oligonucleotides (up to about 200 nucleotide residues) because the number of errors accumulates with the length of the oligonucleotide being synthesized.[1] Products are often isolated by HPLC to obtain the desired oligonucleotides in high purity. Typically, synthetic oligonucleotides are single-stranded DNA or RNA molecules around 15–25 bases in length. They are most commonly used as antisense oligonucleotides, small interfering RNA, primers for DNA sequencing and amplification, probes for detecting complementary DNA or RNA via molecular hybridization, tools for the targeted introduction of mutations and restriction sites, and for the synthesis of artificial genes.

Contents

History

The evolution of oligonucleotide synthesis saw four major methods of the formation of internucleosidic linkages and has been reviewed in the literature in great detail.[2][3][4]

Early work and contemporary H-phosphonate synthesis

Scheme. 1. i: N-Clorosuccinimide; Bn = -CH2Ph

In the early 1950’s, Alexander Todd’s group pioneered H-phosphonate and phosphate triester methods of oligonucleotide synthesis.[5][6] The reaction of compounds 1 and 2 to form H-phosphonate diester 3 is an H-phosphonate coupling in solution while that of compounds 4 and 5 to give 6 is a phosphotriester coupling (see Phosphotriester synthesis below).

Scheme. 2. Synthetic cycle in H-phosphonate method of oligonucleotide synthesis. X = O, S

Thrity years later, this work inspired, independently, two research groups to adopt the H-phosphonate chemistry to the solid-phase synthesis using nucleoside H-phosphonate monoesters 7 as building blocks and pivaloyl chloride, 2,4,6-triisopropylsulfonyl chloride (TPS-Cl), and other compounds as activators.[7][8] The practical implementation of H-phosphonate method resulted in a very short and simple synthetic cycle consisting of only two steps, detritylation and coupling (Scheme 2). Oxidation of internucleosidic H-phosphonate diester 8 to phosphodiester 9 with a solution of iodine in aqueous pyridine is carried out at the end of the chain assembly rather than as a step in the synthetic cycle. Alternatively, 8 can be converted to phosphorothioate 9 (X =S).

Phosphodiester synthesis

Scheme. 3 Oligonucleotide coupling by phosphodiester method; Tr = -CPh3

In the 1950s, Khorana and co-workers developed a phosphodiester method where 3’-O-acetylnucleoside-5’-O-phosphate 2 was activated with N,N’-dicyclohehylcarbodiimide (DCC) or 4-toluenesulfonylchloride (Ts-Cl) and a 5’-O-protected nucleoside 1 was reacted with the activated species to give a protected dinucleoside monophosphate 3.[9] 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 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 method seems to be a step back from the more selective chemistry described earlier; however, at that 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 to achieve the ultimate goal of the study.[2]

Phosphotriester synthesis

Scheme 4. Oligonucleotide coupling by phosphotriester method; MMT = -CPh2(4-MeOC6H4).

In the 1960s, groups led by R. Letsinger[10] and C. Reese[11] developed a phosphotriester approach. The defining difference from the phosphodiester approach was the protection of the phosphate moiety in the building block 1 and in the product 2 with 2-cyanoethyl group. This precluded the formation of oligonucleotides branched at the internucleosidic phosphate. The higher selectivity of the method allowed the use of more efficient coupling agents and catalysts,[12][13] which dramatically reduced the length of the synthesis. The method, initially developed for the solution-phase synthesis, was also implemented on low-cross-linked “popcorn” polystyrene,[14] which initiated a massive research effort in solid-phase synthesis of oligonucleotides and eventually led to the automation of the oligonucleotide chain assembly.

Phosphite triester synthesis

In the 1970s, substantially more reactive P(III) derivatives of nucleosides, 3'-O-chlorophosphites, were successfully used for the formation of internucleosidic linkages.[15] This led to the discovery of the phosphite triester methodology. The group led by M. Caruthers took the advantage of less aggressive and more selective 1H-terazolidophosphites and implemented the method on solid phase.[16] Very shortly after, the workers from the same group further improved the method by using more stable nucleoside phosphoramidites as building blocks.[17] The use of 2-cyanoethyl phosphite-protecting group in place of a less user-friendly methyl group led to the nucleoside phosphoramidites currently used in oligonucleotide synthesis (see Phosphoramidite building blocks below).[18] Many later improvements to the manufacturing of building blocks, instrumentation, and synthetic protocols made the phosphoramidite chemistry a very reliable and expedite method of choice for the preparation of synthetic oligonucleotides.[1]

Phosphoramidite building blocks

Protected 2'-deoxynucleoside phosphoramidites.

Oligonucleotides are chemically synthesized using nucleoside phosphoramidites. A nucleoside phosphoramidite is a derivative of natural or synthetic nucleosides with protection groups added to its reactive exocyclic amine and hydroxy groups. As mentioned earlier (see Phosphodiester synthesis above), the naturally occurring nucleotides (nucleoside-3'- or 5'-phosphates) are insufficiently reactive to afford the synthetic preparation of oligonucleotides. A dramatically more reactive N,N-diisopropyl phosphoramidite group is therefore attached to the 3'-hydroxy group of a nucleoside to form nucleoside phosphoramidite. To prevent undesired side reactions, all other functional groups of 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. Below, the protecting groups currently used in commercially available and most common nucleoside phosphoramidite building blocks[19][20][21][22][23] are briefly reviewed:

  • The 5'-hydroxyl group is protected by an acid-labile DMT (dimethoxytrityl) group.
  • The exocyclic amino group of nucleic bases adenine, cytidine, and guanine are protected with base-labile groups. Nucleic bases of thymidine and uridine do not have exocyclic amino groups and hence do not require any protection. Most often, two protection schemes are used.
  • In the first, the standard and more robust scheme (Figure), Bz (benzoyl) protection is used for A, dA, C, and dC, and G and dG are protected with isobutyryl group. More recently, Ac (acetyl) group is often used to protect C and dC as shown in Figure.[24]
  • In the second, mild protection scheme, A and dA are protected with isobutyryl[25] or phenoxyacetyl groups (PAC).[26] C and dC bear acetyl protection,[24] and G and dG are protected with 4-isopropylphenoxyacetyl (i-Pr-PAC)[27] or dimethylformamidino (dmf)[28] 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.
2'-O-Protected ribonucleoside phosphoramidites.
  • In RNA synthesis, the 2'-hydroxy group is protected with TBDMS (t-butyldimethylsilyl) group or with TOM (t-butyldimethylsilyloxymethyl) 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 (see "Step 2: Coupling" below).

Solid supports

In solid-phase synthesis, an oligonucleotide being assembled is covalently bound, via its 3'-terminal hydroxy group, to a solid support material and remains attached to it over the entire course of the chain assembly. The solid support is contained in columns whose dimensions depend on the scale of synthesis and may vary between 0.05 mL and several litres. The overwhelming majority oligonucleotides are synthesized on small scale ranging from 40 nmol to 1 μmol. More recently, high-throughput oligonucleotide synthesis where the solid support is contained in the wells of multi-well plates (most often, 96 or 384 wells) became a method of choice for parallel synthesis of oligonucleotides on small scale.[citation needed] At the end of the chain assembly, the oligonucleotide is released from the solid support and eluted from the column or the well.

Solid Support Material

In contrast to organic solid-phase synthesis and peptide synthesis, the synthesis of oligonucleotides proceeds best on non-swellable solid supports. The two most often used solid-phase materials are Controlled Pore Glass (CPG) and macroporous polystyrene (MPPS).[29]

  • CPG is commonly defined by its pore size. In oligonucleotide chemistry, pore sizes of 500, 1000, 1500, 2000, and 3000 Å are used to allow the preparation of about 50, 80, 100, 150, and 200-mer oligonucleotides, respectively. To make native CPG suitable for further processing, the surface of the material is treated with (3-aminopropyl)triethoxysilane to give Aminopropyl CPG. The aminopropyl arm may be further extended to result in Long Chain Aminoalkyl (LCAA) CPG. The amino group is then used as an anchoring point for linkers suitable for oligonucleotide synthesis (see below).
  • MPPS suitable for oligonucleotide synthesis is a low-swellable, highly cross-linked polystyrene obtained by polymerization of divinylbenzene (min 60%), styrene, and 4-chloromethylstyrene in the presence of a porogeneous agent. The macroporous chloromethyl MPPS obtained is converted to aminomethyl MPPS.

Linker chemistry

Commercial solid supports for oligonucleotide synthesis.
Scheme 5. Mechanism of 3'-dephosphorylation of oligonucleotides assembled on universal solid supports.

To make the solid support material suitable for oligonucleotide synthesis, non-nucleosidic linkers or nucleoside succinates are covalently attached to the reactive amino groups in Aminopropyl CPG, LCAA CPG, or Aminomethyl MPPS. The remaining unreacted amino groups are capped with acetic anhydride. Typically, three conceptually different groups of solid supports are used.

  • Universal supports. In a more recent and more convenient method, the synthesis starts with the universal support where a non-nucleosidic linker is attached to the solid support material (Compounds 1 and 2). A phosphoramidite respective to the 3'-terminal nucleoside residue is coupled to the universal solid support in the first synthetic cycle of oligonucleotide chain assembly using the standard protocols. The chain assembly is then continued until the completion, after which the solid support-bound oligonucleotide is deprotected. The characteristic feature of the universal solid supports is that the release of the oligonucleotides occurs by the hydrolytic cleavage of a P-O bond that attaches the 3’-O of the 3’-terminal nucleotide residue to the universal linker as shown in Scheme 5. The critical advantage of this approach is that the same solid support is used irrespectively of the sequence of the oligonucleotide to be synthesized. The solid support 1 and several similar solid supports[30] require aqueous ammonium hydroxide, aqueous methylamine,[31] or their mixture[32] and are commercially available (Ref.[33][34] ). The solid support 2[35] requires a solution of ammonia in anhydrous methanol and is also commercially available.[36][37]
  • Nucleosidic solid supports. In a historically first and still popular approach, the 3'-hydroxy group of the 3'-terminal nucleoside residue is attached to the solid support via, most often, 3’-O-succinyl arm as in compound 3. The oligonucleotide chain assembly starts with the coupling of a phosphoramidite building block respective to the nucleotide residue second from the 3’-terminus. The 3’-terminal hydroxy group in oligonucleotides synthesized on nucleosidic solid supports is deprotected under the conditions somewhat milder than those applicable for universal solid supports. However, the fact that a nucleosidic solid support has to be selected in a sequence-specific manner reduces the throughput of the entire synthetic process and increases the likelihood of human error.
  • Special solid supports are used for the attachment of desired functional or reporter groups at the 3’-terminus of synthetic oligonucleotides. For example, the commercial[38] solid support 4[39] allows the preparation of oligonucleotides bearing 3’-terminal 3-aminopropyl linker. Many other special solid support suited for various applications are commercially available.

Synthetic cycle

Scheme 6. Synthetic cycle for preparation of oligonucleotides by phosphoramidite method.

Oligonucleotide synthesis is carried out by a stepwise addition of nucleotide residues to the 5'-terminus of the growing chain until the desired sequence is assembled. Each addition is referred to as a synthetic cycle (Scheme 6) and consists of four chemical reactions:

  • Step 1 - De-blocking (detritylation): The DMT group is removed with a solution of an acid, such as TCA or Dichloroacetic acid (DCA), in an inert solvent (dichloromethane or toluene) and washed out, resulting in a free 5' hydroxyl group on the first base.
  • Step 2 - Coupling: A nucleoside phosphoramidite (or a mixture of several phosphoramidites) is activated by an acidic azole catalyst, tetrazole, 2-ethylthiotetrazole, 2-bezylthiotetrazole, 4,5-dicyanoimidazole, or a number of similar compounds. This mixture is brought in contact with the starting solid support (first coupling) or oligonucleotide precursor (following couplings) whose 5'-hydroxy group reacts with the activated phosphoramidite moiety of the incoming nucleoside phosphoramidite to form a phosphite triester linkage. This reaction is very rapid and requires, on small scale, about 20 s for its completion. The phosphoramidite coupling is also highly sensitive to the presence of water and is commonly carried out in anhydrous acetonitrile. Unbound reagents and by-products are removed by washing.
  • Step 3 - Capping: After the completion of the coupling reaction, a small percentage of the solid support-bound 5'-OH groups (0.1 to 1%) remain unreacted and need to be permanently blocked from further chain elongation to prevent the formation of oligonucleotides with an internal base deletion commonly referred to as (n-1) shortmers. This is done by acetylation of the unreacted 5'-hydroxy groups using a mixture of acetic anhydride and 1-methylimidazole as a catalyst. Excess reagents are removed by washing.
  • Step 4 - Oxidation: The newly formed tricoordinated phosphite triester linkage is not natural and is of limited stability under the conditions of oligonucleotide synthesis. The treatment of the support-bound material with iodine and water in the presence of a weak base (pyridine, lutidine, or collidine) oxidizes the phosphite triester into a tetracoordinated phosphate triester, a protected precursor of the naturally occurring phosphate diester internucleosidic linkage. This step can be substituted with a sulfurization step to obtain oligonucleotide phosphorothioates (see below). In the latter case, the sulfurization step is carried out prior to capping.

Oligonucleotide phosphorothioates

Sp and Rp-diastereomeric internucleosidic phosphorothioate linkages.

Oligonucleotide phosphorothioates (OPS) are modified oligonucleotides where one of the oxygen atoms in the phosphate moiety is replaced by sulfur. Only the phosphorothioates having sulfur at a non-bridging position as shown in Figure are widely used and are available commercially. The replacement of the non-bridging oxygen with sulfur creates a new center of chirality at phosphorus. In a simple case of a dinucleotide, this results in the formation of a diastereomeric pair of Sp- and Rp-dinucleoside monophosphorothioates whose structures are shown in Figure. In a n-mer oligonucleotide where all (n - 1) internucleosidic linkages are phosphorothioate linkages, the number of diastereomers m is calculated as m = 2(n - 1). Being non-natural analogs of nucleic acids, OPS are substantially more stable towards hydrolysis by nucleases, the class of enzymes that destoy nucleic acids by breaking the bridging P-O bond of the phosphodiester moiety. This property determines the use of OPS as antisense oligonucleotides in in vitro and in vivo applications where the extensive exposure to nucleases is inevitable. Similarly, to improve the stability of siRNA, at least one phosphorothioate linkage is often introduced at the 3'-terminus of both sense and antisense strands. In chirally pure OPS, all-Sp diastereomers are more stable to enzymatic degradation than their all-Rp analogs.[40] However, the preparation of chirally pure OPS remains a synthetic challenge. In laboratory practice, mixtures of diastereomers of OPS are commonly used. Synthesis of OPS is very similar to that of natural oligonucleotides. The difference is that the oxidation step is replaced by sulfur transfer (sulfurization) and that the capping step is performed after the sulfurization. Of many reported reagents capable of the efficient sulfur transfer, only three are commercially available:

Commercial sulfur transfer agents for oligonucleotide synthesis.
  • 3-((Dimethylamino-methylidene)amino)-3H-1,2,4-dithiazole-3-thione, DDTT (3, Fig. 4) provides rapid kinetics of sulfurization and high stability in soltion.[41][42] The reagent is available from several sources.[43][44]
  • 3H-1,2-benzodithiol-3-one 1,1-dioxide (4, Fig. 4)[45] also known as Beaucage Reagent displays a better solubility in acetonitrile and fast reaction times. However, the reagent is of limited stability in solution and is less efficient in sulfurizing RNA linkages.[41][42]
  • N,N,N'N'-Tetraethylthiuram disulfide (TETD) is soluble in acetonitrile and is commercially available.[46] However, the sulfurization reaction of an internucleosidic DNA linkage with TETD requires 15 min,[47] which is more than 5 times as slow as that with compounds 3 and 4.

Automation

In the past, oligonucleotide synthesis was carried out manually using, as containers for the solid phase, miniature glass columns similar in their shape to low-pressure chromatography columns or syringes equipped with porous filters.[48] Currently, solid-phase oligonucleotide synthesis is carried out automatically using computer-controlled instruments (oligonucleotide synthesizers) and is technically implemented in column, multi-well plate, and array formats. The column format is best suited for research and large scale applications where a high-throughput is not required. Multi-well plate format is designed specifically for high-throughput synthesis on small scale to satisfy the growing demand of industry and academia for synthetic oligonucleotides. A number of oligonucleotide synthesizers are available commercially.

Synthesis of oligonucleotide microarrays

Main article: DNA microarray#Fabrication

One may visualize an oligonucleotide microarray as a miniature multi-well plate where physical dividers between the wells (plastic walls) are intentionally removed. With respect to the chemistry, synthesis of oligonucleotide microarrays is different from the conventional oligonucleotide synthesis in two respects:

5'-O-NPPOC-protected nucleoside phosphoramidite.
  • Oligonucleotides remain permanently attached to the solid phase, which requires the use of linkers that are stable under the conditions of the final deprotection procedure.
  • The absence of physical dividers between the sites occupied by individual oligonucleotides, a very limited space on the surface of the microarray (one oligonucleotide sequence occupies a square 25x25 μm)[49] and the requirement of high fidelity of oligonucleotide synthesis dictate the use of site-selective 5'-deprotection techniques. In one approach, the removal of the 5'-O-DMT group is effected by electrochemical generation of the acid at the required site(s).[50] In another approach, 5'-O-(α-methyl-6-nitropiperonyloxycarbonyl) (NPPOC) protecting group removable by UV-irradiation at 365 nm is used.[49]

Post-synthetic processing

After synthesis is complete, the fully protected, solid support-bound oligonucleotides are subjected to deprotection using one of the two general approaches.

  • (1) Most often, 5'-DMT group is removed at the end of the automatic synthesis. The oligonucleotides are then released from the solid phase and deprotected (base and phosphate) by treatment with aqueous ammonium hydroxide, aqueous methylamine, their mixtures,[24] gaseous ammonia or methylamine[51] or, less commonly, solutions of other primary amines or alkalies at ambient or elevated temperature. This removes all remaining protection groups from 2'-deoxyoligonucleotides, resulting in a reaction mixture containing the desired product. If the oligonucleotide contains any 2'-O-protected ribonucleotide residues, the deprotection protocol includes the second step where the 2'-O-protecting silyl groups are removed by treatment with fluoride ion. The fully deprotected product is used as is, or the desired oligonucleotide can be purified by a number of methods. Most commonly, the crude product is desalted using ethanol precipitation or size exclusion chromatography. To eliminate unwanted truncation products, the oligonucleotides can be purified via polyacrylamide electrophoresis or anion-exchange HPLC followed by desalting.
  • (2) The second approach is only used when the intended method of purification is reverse-phase HPLC. In this case, the 5'-terminal DMT group that serves as a hydrophobic handle for purification is kept on at the end of the synthesis. The oligonucleotide is deprotected under basic conditions as described above and, upon evaporation, is purified by reverse-phase HPLC. The collected material is then detritylated under aqueous acidic conditions and, finally, desalted.
  • For some applications, additional reporter groups may be attached to an oligonucleotide using a variety of post-synthetic procedures.

Characterization

As with any other organic compound, it is prudent to characterize synthetic oligonucleotides upon their preparation. In more complex cases (research and large scale syntheses) oligonucleotides are characterized after their deprotection and after purification. The ultimate approach to the characterization is sequencing, a relatively laborious and time consuming procedure. In day-by-day practice, it is sufficient to obtain the molecular mass of an oligonucleotide by recording its mass spectrum. Two methods are currently widely used for characterization of oligonucleotides: electrospray mass spectrometry (ES MS) and matrix-assisted laser desorption/ionization time-of-flight mass spectrometry (MALDI-TOF). Prior to submitting a sample to the analysis by either of the methods, it is very important to exchange all metal ions that might be present in the sample for ammonium or trialkylammonium (e.c. triethylammonium) ions.

Deconvoluted ES MS of crude oligonucleotide 5'-DMT-T20 (Calcd. 6324.26 Da).
  • In ES MS spectrum, a given oligonucleotide generates a set of ions that corresponds to different ionization states of the compound. Thus, the oligonucleotide with molecular mass M generates ions with masses (M - nH)/n where M is the molecular mass of the oligonucleotide in the form of free acid (all charges of internucleosidic phosphodiesters are neutralized with H+), n is the ionization state, and H is the atomic mass of hydrogen (1 Da). Most useful for characterization are the ions with n ranging from 2 to 5. Software supplied with the recently purchased instruments is capable of performing a deconvolution procedure that is, it finds peaks of ions that belong to the same set and derives the molecular mass of the oligonucleotide.
  • To obtain more detailed information on the impurity profile of oligonucleotides, liquid chromatography-mass spectrometry (LC-MS or HPLC-MS)[52] or capillary electrophoresis mass spectrometry (CEMS)[53] are used.


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Further reading

  • Comprehensive Natural Products Chemistry, Volume 7: DNA and Aspects of Molecular Biology. Kool, Eric T.; Editor. Neth. (1999), 733 pp. Publisher: (Elsevier, Amsterdam, Neth.)
  • Beaucage, S. L.; Iyer, R. P. Advances in the synthesis of oligonucleotides by the phosphoramidite approach. Tetrahedron 1992, 48, 2223-2311.
  • Beaucage, S. L.; Iyer, R. P. The functionalization of oligonucleotides via phosphoramidite derivatives. Tetrahedron 1993, 49, 1925-1963.
  • Beaucage, S. L.; Iyer, R. P. The synthesis of modified oligonucleotides by the phosphoramidite approach and their applications. Tetrahedron 1993, 49, 6123-6194.
  • Beaucage, S L. Oligodeoxyribonucleotides synthesis. Phosphoramidite approach. Methods in Molecular Biology (Totowa, NJ, United States) (1993), 20 (Protocols for Oligonucleotides and Analogs), 33-61.
  • Reese, C. B. The chemical synthesis of oligo- and poly-nucleotides: a personal commentary. Tetrahedron 2002, 58, 8893-8920.

Commercialization

See also


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