From Wikipedia, the free encyclopedia
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 only in a
5' to 3' direction,
chemical oligonucleotide synthesis does not have this limitation,
although it is, most often, carried out in the opposite, 3' to 5'
direction. Currently, the process is implemented as
solid-phase synthesis using
phosphoramidite method and phosphoramidite building blocks derived from protected
2'-deoxynucleosides (
dA,
dC,
dG, and
T),
ribonucleosides (
A,
C,
G, and
U), or chemically modified nucleosides, e.g.
LNA or
BNA.
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 since 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
high-performance liquid chromatography
(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.
Oligonucleotides find a variety of applications in molecular biology and medicine. 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.
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.
Early work and contemporary H-phosphonate synthesis
Scheme. 1. i: N-Chlorosuccinimide; Bn = -CH2Ph
In the early 1950s,
Alexander Todd’s group pioneered H-phosphonate and
phosphate triester methods of oligonucleotide synthesis. 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. Oligonucleotide sythesis by the H-Phosphonate Method
Thirty 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-triisopropylbenzenesulfonyl chloride
(TPS-Cl), and other compounds as activators.
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 linkages in
8 to phosphodiester linkages in
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. If desired, the oxidation may be carried out
under anhydrous conditions. Alternatively,
8 can be converted to phosphorothioate
10 or phosphoroselenoate
11 (X = Se), or oxidized by
CCl4 in the presence of primary or secondary amines to phosphoramidate analogs
12.
The method is very convenient in that various types of phosphate
modifications (phosphate/phosphorothioate/phosphoramidate) may be
introduced to the same oligonucleotide for modulation of its properties.
Most often, H-phosphonate building blocks are protected at the
5'-hydroxy group and at the amino group of nucleic bases A, C, and G in
the same manner as phosphoramidite building blocks (see below). However,
protection at the amino group is not mandatory.
Phosphodiester synthesis
Scheme. 3 Oligonucleotide coupling by phosphodiester method; Tr = -CPh3
In the 1950s,
Har Gobind Khorana and co-workers developed a
phosphodiester method where 3’-
O-acetylnucleoside-5’-
O-phosphate
2 (Scheme 3) was activated with
N,N'-dicyclohexylcarbodiimide (DCC) or
4-toluenesulfonyl chloride (Ts-Cl). The activated species were reacted with a 5’-
O-protected nucleoside
1 to give a protected dinucleoside monophosphate
3. 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 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.
Phosphotriester synthesis
Scheme 4. Oligonucleotide coupling by phosphotriester method; MMT = -CPh2(4-MeOC6H4).
In the 1960s, groups led by R. Letsinger and C. Reese
developed a phosphotriester approach. The defining difference from the
phosphodiester approach was the protection of the phosphate moiety in
the building block
1 (Scheme 4) and in the product
3 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,
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,
and later on controlled pore glass (CPG, see "Solid support material"
below), 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. 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 1
H-tetrazolidophosphites and implemented the method on solid phase.
Very shortly after, the workers from the same group further improved
the method by using more stable nucleoside phosphoramidites as building
blocks. 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).
Many later improvements to the manufacturing of building blocks,
oligonucleotide synthesizers, and synthetic protocols made the
phosphoramidite chemistry a very reliable and expedient method of choice
for the preparation of synthetic oligonucleotides.
Synthesis by the phosphoramidite method
Building blocks
Nucleoside phosphoramidites
Protected 2'-deoxynucleoside phosphoramidites.
As mentioned above, the 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.
Below, the protecting groups currently used in commercially available and most common nucleoside phosphoramidite building blocks are briefly reviewed:
- 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.
In 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, these
approaches currently remain in the research stage. 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.
- In the first, the standard and more robust scheme (Figure), 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 as shown in Figure.
- In the second, 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 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.
2'-O-protected ribonucleoside phosphoramidites.
- 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 (see "Step 2: Coupling" below).
Non-nucleoside phosphoramidites
Non-nucleoside
phosphoramidites for 5'-modification of synthetic oligonucleotides. MMT
= mono-methoxytrityl,
(4-methoxyphenyl)diphenylmethyl.
Non-nucleoside
phosphoramidites are the phosphoramidite reagents designed to introduce
various functionalities at the termini of synthetic oligonucleotides or
between nucleotide residues in the middle of the sequence. In order to
be introduced inside the sequence, a non-nucleosidic modifier has to
possess at least two hydroxy groups, one of which is often protected
with the DMT group while the other bears the reactive phosphoramidite
moiety.
Non-nucleosidic phosphoramidites are used to introduce desired
groups that are not available in natural nucleosides or that can be
introduced more readily using simpler chemical designs. A very short
selection of commercial phosphoramidite reagents is shown in Scheme for
the demonstration of the available structural and functional diversity.
These reagents serve for the attachment of 5'-terminal phosphate (
1), NH
2 (
2), SH (
3), aldehydo (
4), and carboxylic groups (
5), CC triple bonds (
6), non-radioactive labels and
quenchers (exemplified by
6-FAM amidite 7 for the attachment of
fluorescein and dabcyl amidite
8, respectively), hydrophilic and hydrophobic modifiers (exemplified by
hexaethyleneglycol amidite
9 and
cholesterol amidite
10, respectively), and
biotin amidite
11.
Synthetic cycle
Scheme 5. 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 5) and consists of four chemical reactions:
Step 1: De-blocking (detritylation)
The DMT group is removed with a solution of an acid, such as 2%
trichloroacetic acid (TCA) or 3%
dichloroacetic acid (DCA), in an inert solvent (
dichloromethane or
toluene).
The orange-colored DMT cation formed is washed out; the step results in
the solid support-bound oligonucleotide precursor bearing a free
5'-terminal hydroxyl group.
It is worth remembering that conducting detritylation for an extended
time or with stronger than recommended solutions of acids leads to
depurination of solid support-bound oligonucleotide and thus reduces the yield of the desired full-length product.
Step 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,
1H-tetrazole, 5-ethylthio-1H-tetrazole, 2-benzylthiotetrazole, 4,5-dicyano
imidazole,
or a number of similar compounds. A more extensive information on the
use of various coupling agents in oligonucleotide synthesis can be found
in a recent review.
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 activated
phosphoramidite in 1.5 – 20-fold excess over the support-bound material
is then brought in contact with the starting solid support (first
coupling) or a support-bound 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. The coupling of 2'-deoxynucleoside
phosphoramidites is very rapid and requires, on small scale, about 20 s
for its completion. In contrast, sterically hindered 2'-
O-protected ribonucleoside phosphoramidites require 5-15 min to be coupled in high yields.
The reaction is also highly sensitive to the presence of water,
particularly when dilute solutions of phosphoramidites are used, and is
commonly carried out in anhydrous acetonitrile. Generally, the larger
the scale of the synthesis, the lower the excess and the higher the
concentration of the phosphoramidites is used. In contrast, 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.
Step 3: Capping
The capping step is performed by treating the solid support-bound material with a mixture of acetic anhydride and
1-methylimidazole or, less often,
DMAP as catalysts and, in the phosphoramidite method, serves two purposes.
- After the completion of the coupling reaction, a small
percentage of the solid support-bound 5'-OH groups (0.1 to 1%) remains
unreacted and needs 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. The unreacted
5'-hydroxy groups are, to a large extent, acetylated by the capping
mixture.
- It has also been reported that phosphoramidites activated with 1H-tetrazole react, to a small extent, with the O6 position of guanosine. Upon oxidation with I2 /water, this side product, possibly via O6-N7 migration, undergoes depurination. The apurinic sites
thus formed are readily cleaved in the course of the final deprotection
of the oligonucleotide under the basic conditions (see below) to give
two shorter oligonucleotides thus reducing the yield of the full-length
product. The O6 modifications are rapidly removed by treatment with the capping reagent as long as the capping step is performed prior to oxidation with I2/water.
- The synthesis of oligonucleotide phosphorothioates (OPS, see below) does not involve the oxidation with I2/water,
and, respectively, does not suffer from the side reaction described
above. On the other hand, if the capping step is performed prior to
sulfurization, the solid support may contain the residual acetic
anhydride and N-methylimidazole left after the capping step. The capping
mixture interferes with the sulfur transfer reaction, which results in
the extensive formation of the phosphate triester internucleosidic
linkages in place of the desired PS triesters. Therefore, for the
synthesis of OPS, it is advisable to conduct the sulfurization step prior to the capping step.
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. Oxidation may be carried out under anhydrous
conditions using
tert-Butyl hydroperoxide or, more efficiently, (1S)-(+)-(10-camphorsulfonyl)-oxaziridine (CSO). The step of oxidation may be substituted with a sulfurization step to obtain oligonucleotide phosphorothioates (see
Oligonucleotide phosphorothioates and their synthesis below). In the latter case, the sulfurization step is best carried out prior to capping.
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 liters. The overwhelming majority of oligonucleotides are synthesized on small scale ranging from 10 n
mol
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 per plate) became a method of choice for
parallel synthesis of oligonucleotides on small scale.
At the end of the chain assembly, the oligonucleotide is released from
the solid support and is 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 or
low-swellable solid supports. The two most often used solid-phase
materials are controlled pore glass (CPG) and macroporous
polystyrene (MPPS).
- 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 6. 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, more convenient,
and more widely used 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 6. The
critical advantage of this approach is that the same solid support is
used irrespectively of the sequence of the oligonucleotide to be
synthesized. For the complete removal of the linker and the 3'-terminal
phosphate from the assembled oligonucleotide, the solid support 1 and several similar solid supports require gaseous ammonia, aqueous ammonium hydroxide, aqueous methylamine, or their mixture and are commercially available. The solid support 2 requires a solution of ammonia in anhydrous methanol and is also commercially available.
- 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 solid support 4
allows the preparation of oligonucleotides bearing 3’-terminal
3-aminopropyl linker. Similarly to non-nucleosidic phosphoramidites,
many other special solid supports designed for the attachment of
reactive functional groups, non-radioactive reporter groups, and
terminal modifiers (e.c. cholesterol
or other hydrophobic tethers) and suited for various applications are
commercially available. A more detailed information on various solid
supports for oligonucleotide synthesis can be found in a recent review.
Oligonucleotide phosphorothioates and their synthesis
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 S
p- and R
p-dinucleoside monophosphorothioates whose structures are shown in Figure. In an
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 destroy 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. 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 reaction (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-(Dimethylaminomethylidene)amino-3H-1,2,4-dithiazole-3-thione, DDTT (3) provides rapid kinetics of sulfurization and high stability in solution. The reagent is available from several sources.
- 3H-1,2-benzodithiol-3-one 1,1-dioxide (4)
also known as Beaucage reagent displays a better solubility in
acetonitrile and short reaction times. However, the reagent is of
limited stability in solution and is less efficient in sulfurizing RNA
linkages.
- N,N,N'N'-Tetraethylthiuram disulfide (TETD) is soluble in acetonitrile and is commercially available. However, the sulfurization reaction of an internucleosidic DNA linkage with TETD requires 15 min, which is more than 10 times as slow as that with compounds 3 and 4.
Automation
In
the past, oligonucleotide synthesis was carried out manually in
solution or on solid phase. The solid phase synthesis was implemented
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.
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 for small scale synthesis and medium to large scale synthesis are available commercially.
First commercially available oligonucleotide synthesizers
In
March 1982 a practical course was hosted by the Department of
Biochemistry, Technische Hochschule Darmstadt, Germany. M.H. Caruthers,
M.J. Gait, H.G. Gassen, H.Koster, K. Itakura, and C. Birr among others
attended. The program comprised practical work, lectures, and seminars
on solid-phase chemical synthesis of oligonucleotides. A select group of
15 students attended and had an unprecedented opportunity to be
instructed by the esteemed teaching staff.
Along
with manual exercises, several prominent automation companies attended
the course. Biosearch of Novato, CA, Genetic Design of Watertown, MA,
were two of several companies to demonstrate automated synthesizers at
the course. Biosearch presented their new SAM I synthesizer. The Genetic
Design had developed their synthesizer from the design of its sister
companies (Sequemat) solid phase peptide sequencer. The Genetic Design
arranged with Dr Christian Birr (Max-Planck-Institute for Medical
Research)
a week before the event to convert his solid phase sequencer into the
semi-automated synthesizer. The team led by Dr Alex Bonner and Rick
Neves converted the unit and transported it to Darmstadt for the event
and installed into the Biochemistry lab at the Technische Hochschule. As
the system was semi-automatic, the user injected the next base to be
added to the growing sequence during each cycle. The system worked well
and produced a series of test tubes filled with bright red trityl color
indicating complete coupling at each step. This system was later fully
automated by inclusion of an auto injector and was designated the Model
25A.
History of mid to large scale oligonucleotide synthesis
Large
scale oligonucleotide synthesizers were often developed by augmenting
the capabilities of a preexisting instrument platform. One of the first
mid scale synthesizers appeared in the late 1980s, manufactured by the
Biosearch company in Novato, CA (The 8800). This platform was originally
designed as a peptide synthesizer and made use of a fluidized bed
reactor essential for accommodating the swelling characteristics of
polystyrene supports used in the Merrifield methodology. Oligonucleotide
synthesis involved the use of CPG (controlled pore glass) which is a
rigid support and is more suited for column reactors as described above.
The scale of the 8800 was limited to the flow rate required to fluidize
the support. Some novel reactor designs as well as higher than normal
pressures enabled the 8800 to achieve scales that would prepare 1 mmole
of oligonucleotide. In the mid 1990s several companies developed
platforms that were based on semi-preparative and preparative liquid
chromatographs. These systems were well suited for a column reactor
approach. In most cases all that was required was to augment the number
of fluids that could be delivered to the column. Oligo synthesis
requires a minimum of 10 and liquid chromatographs usually accommodate
4. This was an easy design task and some semi-automatic strategies
worked without any modifications to the preexisting LC equipment.
PerSeptive Biosystems as well as Pharmacia (GE) were two of several
companies that developed synthesizers out of liquid chromatographs.
Genomic Technologies, Inc.
was one of the few companies to develop a large scale oligonucleotide
synthesizer that was, from the ground up, an oligonucleotide
synthesizer. The initial platform called the VLSS for very large scale
synthesizer utilized large Pharmacia liquid chromatograph columns as
reactors and could synthesize up to 75 millimoles of material. Many
oligonucleotide synthesis factories designed and manufactured their own
custom platforms and little is known due to the designs being
proprietary. The VLSS design continued to be refined and is continued in
the QMaster synthesizer which is a scaled down platform providing milligram to gram amounts of synthetic oligonucleotide.
The current practices of synthesis of chemically modified oligonucleotides on large scale have been recently reviewed.
Synthesis of oligonucleotide microarrays
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-MeNPOC-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 25×25 μm)
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). Another approach uses 5'-O-(α-methyl-6-nitropiperonyloxycarbonyl)
(MeNPOC) protecting group, which can be removed by irradiation with UV
light of 365 nm wavelength.
Post-synthetic processing
After the completion of the chain assembly, the solid support-bound oligonucleotide is fully protected:
- The 5'-terminal 5'-hydroxy group is protected with DMT group;
- The internucleosidic phosphate or phosphorothioate moieties are protected with 2-cyanoethyl groups;
- The exocyclic amino groups in all nucleic bases except for T and U are protected with acyl protecting groups.
To furnish a functional oligonucleotide, all the protecting groups
have to be removed. The N-acyl base protection and the 2-cyanoethyl
phosphate protection may be, and is often removed simultaneously by
treatment with inorganic bases or amines. However, the applicability of
this method is limited by the fact that the cleavage of 2-cyanoethyl
phosphate protection gives rise to
acrylonitrile
as a side product. Under the strong basic conditions required for the
removal of N-acyl protection, acrylonitrile is capable of alkylation of
nucleic bases, primarily, at the N3-position of thymine and uracil
residues to give the respective N3-(2-cyanoethyl) adducts via
Michael reaction. The formation of these side products may be avoided by treating the
solid support-bound oligonucleotides with solutions of bases in an
organic solvent, for instance, with 50%
triethylamine in
acetonitrile or 10%
diethylamine in acetonitrile.
This treatment is strongly recommended for medium- and large scale
preparations and is optional for syntheses on small scale where the
concentration of acrylonitrile generated in the deprotection mixture is
low.
Regardless of whether the phosphate protecting groups were
removed first, the solid support-bound oligonucleotides are deprotected
using one of the two general approaches.
- (1) Most often, 5'-DMT group is removed at the end of the
oligonucleotide chain assembly. The oligonucleotides are then released
from the solid phase and deprotected (base and phosphate) by treatment
with aqueous ammonium hydroxide, aqueous methylamine, their mixtures, gaseous ammonia or methylamine
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 by various methods.
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, size exclusion chromatography, or reverse-phase HPLC. To eliminate unwanted truncation products, the oligonucleotides can be purified via polyacrylamide gel 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. On small
scale (less than 0.01–0.02 mmol), the treatment with 80% aqueous acetic
acid for 15–30 min at room temperature is often used followed by
evaporation of the reaction mixture to dryness in vacuo. Finally, the product is desalted as described above.
- For some applications, additional reporter groups may be attached to
an oligonucleotide using a variety of post-synthetic procedures.
Characterization
Deconvoluted ES MS of crude oligonucleotide 5'-DMT-T20
(calculated mass 6324.26 Da).
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. Although the ultimate
approach to the characterization is
sequencing,
a relatively inexpensive and routine procedure, the considerations of
the cost reduction preclude its use in routine manufacturing of
oligonucleotides. 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).
To obtain informative spectra, it is very important to exchange all
metal ions that might be present in the sample for ammonium or
trialkylammonium [
e.c. triethylammonium, (C
2H
5)
3NH
+] ions prior to submitting a sample to the analysis by either of the methods.
- In ES MS spectrum, a given oligonucleotide generates a set of
ions that correspond 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 a
free acid (all negative charges of internucleosidic phosphodiester
groups 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 more recently manufactured 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) or capillary electrophoresis mass spectrometry (CEMS) are used.