The structure of chemical structure of DNA and RNA; Nucleosides and nucleotides; A-DNA, B-DNA and Z-DNA.

Nucleic acid structure


# 2'-Deoxyribonucleic acid (DNA)

DNA (2'-deoxyribonucleic acid) is the molecular store of genetic information in nearly all living systems. It is a large polymeric molecule composed of monomers known as _nucleotides_. Each nucleotide consists of a heterocyclic base, a pentose sugar (2'-deoxy-§d§-ribofuranose), and a phosphate group. There are four heterocyclic bases in DNA: adenine (A), guanine (G), cytosine (C) and thymine (T). Their structures and numbering system are shown in [figure: DNA-heterocyclic-bases].

{{> figure
    filename    DNA-heterocyclic-bases.svg
    ::title     Chemical structures of the heterocyclic bases of DNA
    ::subtitle  The natural bases of DNA are A, C, G and T
    width       388
    height      99
<}}

Adenine and guanine are purines and cytosine and thymine are pyrimidines ([figure: purine-pyrimidine]).

{{> figure
    filename    purine-pyrimidine.svg
    width       168
    height      74
    ::title     Structures of the purine and pyrimidine heterocyclic ring systems
<}}

The deoxyribose sugar is shown in [figure: deoxyribose-mutarotation]. As a free sugar it can _mutarotate_ under certain conditions, adopting furanose, acyclic and pyranose forms; but in DNA it is fixed as a furanoside.

{{> figure
    filename    deoxyribose-mutarotation.svg
    width       410
    height      82
    ::title     Mutarotation of 2'-deoxy-D-ribose allows interconversion between furanose, acyclic and pyranose forms.
<}}

The phosphate group can be found at the 5'- or 3'-position of the sugar depending on the method used to break down DNA to produce the nucleotides. Removal of the phosphate gives rise to a nucleoside. The heterocyclic base is linked to the 1'-position of the sugar. The chemical structures of a deoxynucleoside and a deoxynucleotide are shown in [figure: deoxynucleoside-deoxynucleotide].

{{> figure
    filename    deoxynucleoside-deoxynucleotide.svg
    width       305
    height      170
    ::title     Structures of a deoxynucleoside and a deoxynucleotide
<}}

The bond joining the 1'-carbon of the deoxyribose sugar to the heterocyclic base is the _N_-glycosidic bond. Rotation about this bond gives rise to _syn_ and _anti_ conformations ([figure: syn-anti-nucleoside-conformations]). Rotation about this bond is restricted and the _anti_ conformation is generally favoured, partly on steric grounds.

{{> figure
    filename    syn-anti-nucleoside-conformations.svg
    width       540
    height      180
    ::title     Structures of _syn_ and _anti_ nucleoside conformations
<}}

The nucleosides in the DNA duplex adopt the _anti_ conformation (there are very few exceptions to this rule, one of which is guanosine monophosphate, in which the guanine base adopts the _syn_ conformation about the glycosidic bond).

The 1'-position of the deoxyribose sugar is the _anomeric centre_. If a substituent attached to the 1'-carbon lies on the same face of the sugar ring as the 5'-hydroxyl group, it is known as the β-anomer; if the substituent is on the opposite side of the sugar ring it is the α-anomer ([figure: nucleoside-anomers]). All of the nucleosides in DNA are in the β-configuration.

{{> figure
    filename    nucleoside-anomers.svg
    width       368
    height      144
    ::title     Structures of nucleoside α- and β-anomers
<}}

# Ribonucleic acid (RNA)

Some organisms, for example retroviruses, use ribonucleic acid (RNA) instead of DNA as their store of genetic information. RNA is chemically very similar to DNA but there are two important differences:

- RNA has a hydroxyl group attached to the 2'-position of the sugar, and
- the pyrimidine uracil (U; [figure: uridine-thymidine]) replaces thymine in RNA.

The key biological role of RNA is as a messenger: it reads the genetic code in DNA ([transcription](/nucleic-acids-book/Transcription-Translation-and-Replication#Transcription)) and transports it to the ribosome, where it is decoded into the sequence of a protein ([translation](/nucleic-acids-book/Transcription-Translation-and-Replication#Translation)).

{{> figure
    filename    uridine-thymidine.svg
    width       291
    height      148
    ::title     Structures of uridine and thymidine nucleosides
    ::subtitle  Uridine occurs in RNA; thymidine in DNA
<}}

# Oligonucleotides

A dinucleotide (dimer) of DNA or RNA is formed by covalently linking the 5'-phosphate group of one nucleotide to the 3'-hydroxyl group of another to form a _phosphodiester bond_. An oligonucleotide (oligomer) is formed when several such bonds are made, and naturally-occurring nucleic acids are linear, high molecular weight molecules of this kind. At physiological pH (7.4) each phosphodiester group exists as an anion (hence the term _nucleic acid_), and nucleic acids are therefore highly charged polyanionic molecules ([figure: GCAT-TACG-oligos]).

{{> figure
    filename    GCAT-TACG-oligos.svg
    width       616
    height      313
    ::title     Chemical structures of the oligonucleotides dGCAT (left) and dTACG (right)
<}}

One end of a nucleic acid strand has a 5'-hydroxyl group (primary hydroxyl) and the other end has a 3'-hydroxyl group (secondary hydroxyl). The nucleic acid chain therefore has directionality. The tetranucleotide in [figure: GCAT-TACG-oligos]a has the sequence 5'-GCAT-3' and the tetranucleotide in [figure: GCAT-TACG-oligos]b has the sequence 5'-TACG-3'. By convention, the prefixes 5'- and 3'- are not written, and nucleic acid sequences are written in the 5' to 3' direction. The oligonucleotides GCAT and TACG are distinct molecules with different chemical and biophysical properties.

# Nucleic acid duplexes

The chemical structure of a single strand of DNA gives little insight into its biological function as a carrier of genetic information. However, when James Watson and Francis Crick showed in 1953 that DNA adopts a double-stranded structure (duplex), the mechanism of [DNA replication](/nucleic-acids-book/Transcription-Translation-and-Replication#DNA-replication) (copying) became apparent. The double-helical structure was principally elucidated from X-ray fibre diffraction data (acquired by Rosalind Franklin and Maurice Wilkins) and Chargaff's rules. Erwin Chargaff discovered that the molar amount of adenine in DNA was always equal to that of thymine and the same was true for guanine and cytosine (i.e. number of moles of G = number of moles of C). Watson and Crick were able to explain this by building models to show that the two strands of DNA are held together by hydrogen bonds between individual bases on opposite strands. The purine base A always pairs with the pyrimidine T and the purine G always pairs with the pyrimidine C ([figure: AT-CG-base-pairs]).

{{> figure
    filename    AT-CG-base-pairs.svg
    width       319
    height      110
    ::title     Hydrogen bonding in the A·T and C·G Watson-Crick DNA base pairs
<}}

Not surprisingly A·T and G·C are known as Watson-Crick base pairs. They are pseudosymmetric and if an A·T base pair is laid over any other base pair (T·A, G·C or C·G) the phosphodiester backbones fall on top of each other ([figure: GConAT-animated] and [figure: TA-AT-and-GC-AT-overlays]). Thus all four base pairs fit neatly into the double helix.

{{> figure
    filename    GConAT-animated.gif
    width       588
    height      425
    ::title     Animated overlay of G·C and A·T base pairs
<}}

{{> figure
    filename    TA-AT-and-GC-AT-overlays.svg
    width       373
    height      130
    ::title     Base pair overlays
    ::subtitle  Structures of the T·A base pair overlaid on the A·T base pair, and the G·C base pair overlaid on the A·T base pair
<}}

The sequence of one strand of DNA precisely defines the sequence of the other; the two strands are said to be _complementary_, and are sometimes called _reverse complements_ of each other. The two strands are antiparallel, with the 5'-end of one strand adjacent to the 3'-end of the other. The two strands coil around each other to form a right-handed double helix, with the hydrophobic base pairs in the centre and the sugars and negatively charged phosphates forming the external hydrophilic backbone. The term "right-handed" indicates that the backbone at the front of the molecule facing the observer slopes down from top right to bottom left. The planar heterocyclic bases stack one on another and the separation between successive base pairs along the helix axis is around 0.34 nm. One helical turn (a full 360° turn of the double helix) is repeated every 10 to 11 base pairs. The stability of the duplex is derived from both [base stacking](DNA-duplex-stability#Base-stacking) and [hydrogen bonding](DNA-duplex-stability#Hydrogen-bonding). The principal form of double helical DNA, B-DNA ([figure: A-DNA-B-DNA-Z-DNA], middle), has a wide major groove and a narrow minor groove running around the helix along the entire length of the molecule. Proteins interact with the DNA in these grooves (principally in the major groove) and some small drug molecules (e.g. netropsin, distamycin) bind in the minor groove (see [Nucleic acid-drug interactions](/nucleic-acids-book/Nucleic-acid-drug-interactions)).

{{> figure
    filename    A-DNA-B-DNA-Z-DNA.png
    width       1123.5
    height      532
    ::title     Three-dimensional structures of A-DNA, B-DNA and Z-DNA
<}}

RNA can also form a right-handed duplex using the same base pairing rules (A·U and G·C), but the RNA duplex has a distinctive shape (the A-form) in which the major groove is deep and the minor groove very shallow. Under conditions of low humidity DNA can also adopt the A-form ([figure: A-DNA-B-DNA-Z-DNA], left), and there are a number of other conformations of DNA and RNA, most of which are subtle variations on the A- and B-forms.

One drastically different DNA conformation, Z-DNA, ([figure: A-DNA-B-DNA-Z-DNA], right) was determined from an X-ray crystal structure of a chemically synthesized DNA strand of the alternating sequence dCGCGCG, which spontaneously forms a duplex in aqueous buffer. Z-DNA is left-handed and has a dinucleotide repeat unit, so the backbone is not smooth but appears to “zig-zag”. It is not clear whether Z-DNA has biological relevance.

# Techniques for determining nucleic acid structure

## Nuclear magnetic resonance (NMR)

NMR is a highly developed and powerful spectroscopic technique that is valuable in the investigation of the structural, thermodynamic and kinetic properties of nucleic acids. The technique can be used to study DNA duplexes, triplexes, quadruplexes, hairpin loops, RNA duplexes and other secondary and tertiary RNA structures. Primarily, ^1^H NMR is used in aqueous buffers with water suppression. ^31^P NMR is also useful to study the environment of the nucleic acid phosphodiester backbone. NMR is most useful for studying nucleic acid structure at the local level, i.e. the nature of base pairs and triplets, nucleic acid-drug interactions and nucleic acid-protein interactions. It is less powerful at giving information on global properties (e.g. A versus B helix, DNA duplex bending) because local interactions have to be used as an indication of global properties. NMR should always be used in conjunction with other physical techniques such as circular dichroism (indicative of conformation, A, B, Z), Ultraviolet melting of duplexes, triplexes and quadruplexes (thermodynamic information), X-ray diffraction (high resolution structure analysis), and fluorescence resonance energy transfer (distance measurements). Typical NMR chemical shifts of base and sugar protons are shown in [figure: DNA-base-proton-NMR-shifts] and [figure: DNA-sugar-proton-NMR-shifts].

{{> figure
    filename    DNA-base-proton-NMR-shifts.svg
    width       412
    height      284
    ::title     NMR chemical shifts of the key residues observed in 1H NMR spectra of nucleic acid bases
    ::subtitle  Data from Wijmenga and van Buuren, _Progress in Nuclear Magnetic Resonance Spectroscopy_ 32, 287–387 (1998).
<}}

{{> figure
    filename    DNA-sugar-proton-NMR-shifts.svg
    width       264
    height      179
    ::title     NMR Chemical shifts of the key residues observed in 1H NMR spectra of the deoxyribose sugar in nucleic acids
    ::subtitle  Data from Wijmenga and van Buuren, _Progress in Nuclear Magnetic Resonance Spectroscopy_ 32, 287–387 (1998).
<}}

## Crystallization and X-ray diffraction

Advances in [solid-phase oligonucleotide synthesis](/nucleic-acids-book/Solid-phase-oligonucleotide-synthesis) and [purification](Purification-of-oligonucleotides) have allowed a large number of oligonucleotides to be crystallized and their molecular structures solved. This has provided important information on both the B and A forms of DNA, confirming predictions made from earlier fibre-diffraction studies, and providing high-resolution information on the sequence dependence of nucleic acid structure. X-ray diffraction of single crystals such as those shown in [figure: DNA-crystals] provides an accurate picture of macromolecular structure with a resolution that exceeds that of other techniques.

When a crystal is placed in the path of an X-ray source it will generate a diffraction pattern, from which, with the help of some complicated mathematics, a three-dimensional picture of the unit cell and the molecules present can be generated. X-ray crystallography requires highly ordered crystals, and obtaining suitable crystals of nucleic acids is normally the slowest part of the process. A highly concentrated solution of an oligonucleotide is required to provide a thermodynamic driving force for crystal formation. The solubility limit is exceeded by gradually removing water from the oligonucleotide solution using an external dehydrating agent, or diffusing a precipitant such as isopropanol into the solution. High-resolution structures of A, B and Z-DNA have been obtained by X-ray crystallography and the technique has been used to study DNA-drug complexes in atomic detail. In most cases the resolution of the structure does not extend beyond 2 Å (0.2 nm), but this is sufficient to provide a reasonably clear picture of the heterocyclic bases and sugar-phosphate backbone.

{{> figure
    filename    DNA-crystals.jpg
    width       302
    height      241
    ::title     DNA crystals
    ::subtitle  Photograph of crystals of a DNA oligonucleotide
<}}


Transcription, Translation and Replication from the perspective of DNA and RNA; The Genetic Code; Evolution (DNA replication is not perfect).

Transcription, Translation and Replication


# DNA, RNA and protein synthesis

The genetic material is stored in the form of DNA in most organisms. In humans, the nucleus of each cell contains 3 × 10^9^ base pairs of DNA distributed over 23 pairs of chromosomes, and each cell has two copies of the genetic material. This is known collectively as the human genome. The human genome contains around 30 000 genes, each of which codes for one protein.

Large stretches of DNA in the human genome are transcribed but do not code for proteins. These regions are called _introns_ and make up around 95% of the genome. The nucleotide sequence of the human genome is now known to a reasonable degree of accuracy but we do not yet understand why so much of it is non-coding. Some of this non-coding DNA controls gene expression but the purpose of much of it is not yet understood. This is a fascinating subject that is certain to advance rapidly over the next few years.

The _Central Dogma of Molecular Biology_ states that _DNA makes RNA makes proteins_ ([figure: central-dogma]).

{{> figure
    filename    central-dogma.svg
    width       577.22
    height      103.49
    ::title     The Central Dogma of Molecular Biology
    ::subtitle  DNA makes RNA makes proteins
<}}

The process by which DNA is copied to RNA is called [transcription](#Transcription), and that by which RNA is used to produce proteins is called [translation](#Translation).

# DNA replication

Each time a cell divides, each of its double strands of DNA splits into two single strands. Each of these single strands acts as a template for a new strand of complementary DNA. As a result, each new cell has its own complete genome. This process is known as _DNA replication_. Replication is controlled by the Watson-Crick pairing of the bases in the template strand with incoming deoxynucleoside triphosphates, and is directed by DNA polymerase enzymes. It is a complex process, particularly in eukaryotes, involving an array of enzymes. A simplified version of bacterial DNA replication is described in [figure: DNA-replication].

{{> figure
    filename    DNA-replication.svg
    width       726
    height      253
    owidth      581
    oheight     202
    ::title     DNA replication in bacteria
    ::subtitle  Simplified representation of DNA replication in bacteria
<}}

DNA biosynthesis proceeds in the 5'- to 3'-direction. This makes it impossible for DNA polymerases to synthesize both strands simultaneously. A portion of the double helix must first unwind, and this is mediated by _helicase_ enzymes.

The leading strand is synthesized continuously but the opposite strand is copied in short bursts of about 1000 bases, as the lagging strand template becomes available. The resulting short strands are called _Okazaki fragments_ (after their discoverers, Reiji and Tsuneko Okazaki). Bacteria have at least three distinct DNA polymerases: Pol I, Pol II and Pol III; it is Pol III that is largely involved in chain elongation. Strangely, DNA polymerases cannot initiate DNA synthesis _de novo_, but require a short primer with a free 3'-hydroxyl group. This is produced in the lagging strand by an RNA polymerase (called DNA primase) that is able to use the DNA template and synthesize a short piece of RNA around 20 bases in length. Pol III can then take over, but it eventually encounters one of the previously synthesized short RNA fragments in its path. At this point Pol I takes over, using its 5'- to 3'-exonuclease activity to digest the RNA and fill the gap with DNA until it reaches a continuous stretch of DNA. This leaves a gap between the 3'-end of the newly synthesized DNA and the 5'-end of the DNA previously synthesized by Pol III. The gap is filled by DNA ligase, an enzyme that makes a covalent bond between a 5'-phosphate and a 3'-hydroxyl group ([figure: DNA-replication-polymerases]). The initiation of DNA replication at the leading strand is more complex and is discussed in detail in more specialized texts.

{{> figure
    filename    DNA-replication-polymerases.svg
    width       723
    height      196
    owidth      578
    oheight     157
    ::title     DNA polymerases in DNA replication
    ::subtitle  Simplified representation of the action of DNA polymerases in DNA replication in bacteria
<}}

## Mistakes in DNA replication

DNA replication is not perfect. Errors occur in DNA replication, when the incorrect base is incorporated into the growing DNA strand. This leads to _mismatched_ base pairs, or _mispairs_. DNA polymerases have proofreading activity, and a [DNA repair](/nucleic-acids-book/Mutagenesis-and-DNA-repair#DNA-repair-mechanisms) enzymes have evolved to correct these mistakes. Occasionally, mispairs survive and are incorporated into the genome in the next round of replication. These mutations may have no consequence, they may result in the death of the organism, they may result in a genetic disease or cancer; or they may give the organism a competitive advantage over its neighbours, which leads to evolution by natural selection.

# Transcription

Transcription is the process by which DNA is copied (_transcribed_) to mRNA, which carries the information needed for protein synthesis. Transcription takes place in two broad steps. First, pre-messenger RNA is formed, with the involvement of RNA polymerase enzymes. The process relies on Watson-Crick base pairing, and the resultant single strand of RNA is the reverse-complement of the original DNA sequence. The pre-messenger RNA is then "edited" to produce the desired mRNA molecule in a process called _RNA splicing_.

## Formation of pre-messenger RNA

The mechanism of transcription has parallels in that of [DNA replication](#DNA-replication). As with DNA replication, partial unwinding of the double helix must occur before transcription can take place, and it is the RNA polymerase enzymes that catalyze this process.

Unlike DNA replication, in which both strands are copied, only one strand is transcribed. The strand that contains the gene is called the _sense_ strand, while the complementary strand is the _antisense_ strand. The mRNA produced in transcription is a copy of the sense strand, but it is the antisense strand that is transcribed.

Ribonucleoside triphosphates (NTPs) align along the antisense DNA strand, with Watson-Crick base pairing (A pairs with U). RNA polymerase joins the ribonucleotides together to form a pre-messenger RNA molecule that is complementary to a region of the antisense DNA strand.wxh Transcription ends when the RNA polymerase enzyme reaches a triplet of bases that is read as a "stop" signal. The DNA molecule re-winds to re-form the double helix.

{{> figure
    filename    transcription.svg
    width       491
    height      405
    owidth      393
    oheight     324
    ::title     Transcription
    ::subtitle  Simplified representation of the formation of pre-messenger RNA (orange) from double-stranded DNA (blue) in transcription
<}}

## RNA splicing

The pre-messenger RNA thus formed contains introns which are not required for protein synthesis. The pre-messenger RNA is chopped up to remove the introns and create messenger RNA (mRNA) in a process called RNA splicing ([figure: RNA-splicing]).

{{> figure
    filename    RNA-splicing.svg
    width		649
    height		156
    owidth      519
    oheight     125
    ::title     RNA splicing
    ::subtitle  Introns are spliced from the pre-messenger RNA to give messenger RNA (mRNA)
<}}

## Alternative splicing

In alternative splicing, individual exons are either spliced or included, giving rise to several different possible mRNA products. Each mRNA product codes for a different protein isoform; these protein isoforms differ in their peptide sequence and therefore their biological activity. It is estimated that up to 60% of human gene products undergo alternative splicing. Several different mechanisms of alternative splicing are known, two of which are illustrated in [figure: alternative-splicing].

{{> figure
    filename    alternative-splicing.svg
    width       420
    height      315
    owidth      336
    oheight     252
    ::title     Alternative splicing
    ::subtitle  Several different mechanisms of alternative splicing exist - a cassette exon can be either included in or excluded from the final RNA (top), or two cassette exons may be mutually exclusive (bottom).
<}}

Alternative splicing contributes to protein diversity - a single gene transcript (RNA) can have thousands of different splicing patterns, and will therefore code for thousands of different proteins: a diverse proteome is generated from a relatively limited genome. Splicing is important in genetic regulation (alteration of the splicing pattern in response to cellular conditions changes protein expression). Perhaps not surprisingly, abnormal splicing patterns can lead to disease states including cancer.

## Reverse transcription

In reverse transcription, RNA is "reverse transcribed" into DNA. This process, catalyzed by reverse transcriptase enzymes, allows retroviruses, including the human immunodeficiency virus (HIV), to use RNA as their genetic material. Reverse transcriptase enzymes have also found applications in biotechnology, allowing scientists to convert RNA to DNA for techniques such as [PCR](/nucleic-acids-book/Sequencing-forensic-analysis-and-genetic-analysis#The-Polymerase-Chain-Reaction-PCR).

# Translation

The mRNA formed in transcription is transported out of the nucleus, into the cytoplasm, to the ribosome (the cell's protein synthesis factory). Here, it directs protein synthesis. Messenger RNA is not directly involved in protein synthesis - transfer RNA (tRNA) is required for this. The process by which mRNA directs protein synthesis with the assistance of tRNA is called _translation_.

The ribosome is a very large complex of RNA and protein molecules. Each three-base stretch of mRNA (triplet) is known as a _codon_, and one codon contains the information for a specific amino acid. As the mRNA passes through the ribosome, each codon interacts with the _anticodon_ of a specific transfer RNA (tRNA) molecule by Watson-Crick base pairing. This tRNA molecule carries an amino acid at its 3'-terminus, which is incorporated into the growing protein chain. The tRNA is then expelled from the ribosome. [figure: translation] shows the steps involved in protein synthesis.

{{> figure
    filename    translation.svg
    width		700
    height		839
    owidth      560
    oheight     671
    ::title     Translation
    ::subtitle  (a) and (b) tRNA molecules bind to the two binding sites of the ribosome, and by hydrogen bonding to the mRNA; (c) a peptide bond forms between the two amino acids to make a dipeptide, while the tRNA molecule is left uncharged; (d) the uncharged tRNA molecule leaves the ribosome, while the ribosome moves one codon to the right (the dipeptide is translocated from one binding site to the other); (e) another tRNA molecule binds; (f) a peptide bond forms between the two amino acids to make a tripeptide; (g) the uncharged tRNA molecule leaves the ribosome
<}}

# Transfer RNA

Transfer RNA adopts a well defined tertiary structure which is normally represented in two dimensions as a cloverleaf shape, as in [figure: translation]. The structure of tRNA is shown in more detail in [figure: tRNA].

{{> figure
    filename    tRNA.svg
    width		414
    height		564
    owidth      331
    oheight     451
    ::title     Two-dimensional structures of tRNA (transfer RNA)
    ::subtitle  In some tRNAs the DHU arm has only three base pairs.
<}}

Each amino acid has its own special tRNA (or set of tRNAs). For example, the tRNA for phenylalanine (tRNAPhe) is different from that for histidine (tRNAHis). Each amino acid is attached to its tRNA through the 3'-OH group to form an ester which reacts with the α-amino group of the terminal amino-acid of the growing protein chain to form a new amide bond (peptide bond) during protein synthesis ([figure: protein-synthesis-tRNA]). The reaction of esters with amines is generally favourable but the rate of reaction is increased greatly in the ribosome.

{{> figure
    filename    protein-synthesis-tRNA.svg
    width       516
    height      418
    ::title     Protein synthesis
    ::subtitle  Reaction of the growing polypeptide chain with the 3'-end of the charged tRNA. The amino acid is transferred from the tRNA molecule to the protein
<}}

Each transfer RNA molecule has a well defined tertiary structure that is recognized by the enzyme aminoacyl tRNA synthetase, which adds the correct amino acid to the 3'-end of the uncharged tRNA. The presence of modified nucleosides is important in stabilizing the tRNA structure. Some of these modifications are shown in [figure: tRNA-modified-bases].

{{> figure
    filename    tRNA-modified-bases.svg
    width       464
    height      355
    ::title     Modified bases in tRNA
    ::subtitle  Structures of some of the modified bases found in tRNA
<}}

# The Genetic code

The genetic code is almost universal. It is the basis of the transmission of hereditary information by nucleic acids in all organisms. There are four bases in RNA (A,G,C and U), so there are 64 possible triplet codes (4^3^ = 64). In theory only 22 codes are required: one for each of the 20 naturally occurring amino acids, with the addition of a start codon and a stop codon (to indicate the beginning and end of a protein sequence). Many amino acids have several codes (_degeneracy_), so that all 64 possible triplet codes are used. For example Arg and Ser each have 6 codons whereas Trp and Met have only one. No two amino acids have the same code but amino acids whose side-chains have similar physical or chemical properties tend to have similar codon sequences, e.g. the side-chains of Phe, Leu, Ile, Val are all hydrophobic, and Asp and Glu are both carboxylic acids (see [genetic code](#genetic-code)). This means that if the incorrect tRNA is selected during translation (owing to mispairing of a single base at the codon-anticodon interface) the misincorporated amino acid will probably have similar properties to the intended tRNA molecule. Although the resultant protein will have one incorrect amino acid it stands a high probability of being functional. Organisms show "codon bias" and use certain codons for a particular amino acid more than others. For example, the codon usage in humans is different from that in bacteria; it can sometimes be difficult to express a human protein in bacteria because the relevant tRNA might be present at too low a concentration.

<table id="genetic-code">
    <thead>
    <tr>
        <th>First base (5'-end)</th>
        <th>Middle base</th>
        <th colSpan="4">Third Base ('3-end)</th>
    </tr>
    <tr>
        <th></th>
        <th></th>
        <th>U</th>
        <th>C</th>
        <th>A</th>
        <th>G</th>
    </tr>
    </thead>
    <tbody>
        <tr>
            <td>U</td>
            <td>U</td>
            <td>Phe</td>
            <td>Phe</td>
            <td>Leu</td>
            <td>Leu</td>
        </tr>
        <tr>
            <td></td>
            <td>C</td>
            <td>Ser</td>
            <td>Ser</td>
            <td>Ser</td>
            <td>Ser</td>
        </tr>  
        <tr>
            <td></td>
            <td>A</td>
            <td>Tyr</td>
            <td>Tyr</td>
            <td><b>Stop</b></td>
            <td><b>Stop</b></td>
        </tr>                
        <tr>
            <td></td>
            <td>G</td>
            <td>Cys</td>
            <td>Cys</td>
            <td><b>Stop</b></td>
            <td>Trp</td>
        </tr>   
        <tr>
            <td>C</td>
            <td>U</td>
            <td>Leu</td>
            <td>Leu</td>
            <td>Leu</td>
            <td>Leu</td>
        </tr>
        <tr>
            <td></td>
            <td>C</td>
            <td>Pro</td>
            <td>Pro</td>
            <td>Pro</td>
            <td>Pro</td>
        </tr>  
        <tr>
            <td></td>
            <td>A</td>
            <td>His</td>
            <td>His</td>
            <td>Gln</td>
            <td>Gln</td>
        </tr>                
        <tr>
            <td></td>
            <td>G</td>
            <td>Arg</td>
            <td>Arg</td>
            <td>Arg</td>
            <td>Arg</td>
        </tr>                  
        <tr>
            <td>A</td>
            <td>U</td>
            <td>lle</td>
            <td>lle</td>
            <td>lle</td>
            <td><i><b>Met</b></i></td>
        </tr>
        <tr>
            <td></td>
            <td>C</td>
            <td>Thr</td>
            <td>Thr</td>
            <td>Thr</td>
            <td>Thr</td>
        </tr>  
        <tr>
            <td></td>
            <td>A</td>
            <td>Asn</td>
            <td>Asn</td>
            <td>Lys</td>
            <td>Lys</td>
        </tr>                
        <tr>
            <td></td>
            <td>G</td>
            <td>Ser</td>
            <td>Ser</td>
            <td>Arg</td>
            <td>Arg</td>
        </tr>  
        <tr>
            <td>G</td>
            <td>U</td>
            <td>Val</td>
            <td>Val</td>
            <td>Val</td>
            <td>Val</td>
        </tr>
        <tr>
            <td></td>
            <td>C</td>
            <td>Ala</td>
            <td>Ala</td>
            <td>Ala</td>
            <td>Ala</td>
        </tr>  
        <tr>
            <td></td>
            <td>A</td>
            <td>Asp</td>
            <td>Asp</td>
            <td>Glu</td>
            <td>Glu</td>
        </tr>                
        <tr>
            <td></td>
            <td>G</td>
            <td>Gly</td>
            <td>Gly</td>
            <td>Gly</td>
            <td>Gly</td>
        </tr>                          
    </tbody>
</table>

## An exercise in the use of the genetic code

One strand of genomic DNA (strand A, coding strand) contains the following sequence reading from 5' to 3':

TCGTCGACGATGATCATCGGCTACTCGA

This strand will form the duplex

5'-TCGTCGACGATGATCATCGGCTACTCGA-3'
3'-AGCAGCTGCTACTAGTAGCCGATGAGCT-5'

The sequence of bases in the other strand of DNA (strand B) written 5' to 3' is therefore

TCGAGTAGCCGATGATCATCGTCGACGA

In the mRNA transcribed from strand A of DNA, the sequence of bases written 5' to 3' is

UCGAGUAGCCGAUGAUCAUCGUCGACGA

resulting in an amino acid sequence

Ser-Ser-Ser-Arg-STOP

However, if DNA strand B is the coding strand the mRNA sequence will be

UCGUCGACGAUGAUCAUCGGCUACUCGA

and the amino-acid sequence will be

Ser-Ser-Thr-Met-Ile-Ile-Gly-Tyr-Ser-

# The Wobble hypothesis

Close inspection of all of the available codons for a particular amino acid reveals that the variation is greatest in the third position (for example, the codons for alanine are GCU, GCC, GCA and GCG). Crick and Brenner proposed that a single tRNA molecule can recognize codons with different bases at the 3'-end owing to non-Watson-Crick base pair formation with the third base in the codon-anticodon interaction. These non-standard base pairs are different in shape from A·U and G·C and the term _wobble hypothesis_ indicates that a certain degree of flexibility or "wobbling" is allowed at this position in the ribosome. Not all combinations are possible; examples of "allowed" pairings are shown in [figure: RNA-wobble-base-pairs].

{{> figure
    filename    RNA-wobble-base-pairs.svg
    width       396
    height      269
    ::title     Structures of wobble base pairs found in RNA
<}}

The ability of DNA bases to form wobble base pairs as well as Watson-Crick base pairs can result in [mismatches](/nucleic-acids-book/Mutagenesis-and-DNA-repair#Mismatches-in-DNA-bases) occurring during DNA replication. If not repaired by DNA repair enzymes, these mismatches can lead to genetic diseases and cancer.


How genetic mistakes that occur during replication are kept in check; DNA duplex stability; DNA damage and its consequences.

Mutagenesis and DNA repair

Groove binders sit in the minor groove; intercalators sandwich between base pairs; alkylators chemically react with DNA; DNA cleavage agents break DNA chains.

Nucleic acid-drug interactions

All oligonucleotides are made by phosphoramidite solid-phase synthesis. The steps in solid phase synthesis, and the preparation of reagents.

Solid-phase oligonucleotide synthesis

RNA has an extra OH group, making it more reactive than DNA, and its synthesis more involved. Protecting groups, and avoiding 2'- to 3'-phosphate migration.

RNA oligonucleotide synthesis


# Introduction

The primary structures of RNA and DNA are similar, with two important differences: RNA has a ribose sugar rather than 2'-deoxyribose; and uracil replaces thymine as one of the heterocyclic bases. Not surprisingly, solid phase synthesis of RNA is based on [solid-phase DNA synthesis](Solid-phase-oligonucleotide-synthesis), though considerable attention has to be given to the choice of protecting group for the 2'-hydroxyl function of ribose.

# Solid-phase synthesis of RNA

The 2'-hydroxyl protecting group must be retained throughout the solid-phase synthesis cycle and then be removed cleanly at the end of the synthesis. It is imperative that there is no migration of the protecting group from 2'-OH to 3'-OH during synthesis of the phosphoramidite monomer, and no 3' to 2' phosphate migration during oligonucleotide synthesis or deprotection (see [figure: RNA-chain-cleavage-phosphate-migration]).

A further important consideration in choosing appropriate protection for the 2'-OH is the influence that this group exerts on the phosphoramidite group in the 3'-position. Bulky protecting groups give reduced coupling efficiency, owing to steric hindrance. In practice, the choice of RNA synthesis methodology is dictated by the availability of RNA phosphoramidite monomers of high quality and reasonable cost. The most common method of 2'-OH protection for RNA synthesis is the _tert_-butyldimethylsilyl (TBS) method. During synthesis of the phosphoramidite monomers, the TBS group is added to the 2'-OH of the 5'-DMT-protected nucleoside, to afford the desired 2'-TBS protected nucleoside ([figure: TBS-RNA-monomer-synthesis]). However, this reaction is not regiospecific and a mixture of 2' and 3'-TBS-protected nucleosides are obtained that require careful separation by silica gel column chromatography. The desired 2'-TBS isomer is converted to the phosphoramidite monomer by reaction with 2-cyanoethyl diisopropylaminophosphorochloridite in the presence of the non-nucleophilic base _N_,_N_-diisopropylethylamine (DIPEA).

{{> figure
    filename    TBS-RNA-monomer-synthesis.svg
    width       472
    height      299
    ::title     TBS-protected RNA monomer synthesis
    ::subtitle  Synthesis of 2'-_tert_-butyldimethylsilyl (TBS)-protected phosphoramidite monomers for use in RNA synthesis
<}}

After oligonucleotide synthesis and deprotection with ammonia, the 2'-TBS groups are removed from the oligoribonucleotide using a source of fluoride ion (e.g. tetrabutylammonium fluoride or Et~3~N · 3 HF). Removal of TBS protecting groups proceeds smoothly without cleavage or isomerization of the phosphodiester linkage.

In RNA synthesis, different heterocyclic base protecting groups are used from those employed in DNA synthesis. The _tert_-butylphenoxyacetyl group is used to protect the exocyclic amino groups of A, G and C. This group is useful for two reasons: (i) it is quickly removed in ammonia under conditions that do not cleave the 2'-TBS group (the carbonyl group of the amide is not sterically hindered), and (ii) the _tert_-butyl moiety increases the solubility of the monomer in organic solvents such as acetonitrile.

{{> figure
    filename    TBS-RNA-monomers.svg
    width       434
    height      412
    ::title     TBS-protected RNA monomers
    ::subtitle  Structures of 2'-_tert_-butyldimethylsilyl (TBS)-protected phosphoramidite monomers used in solid-phase RNA synthesis
<}}

The synthesis of oligoribonucleotides using 2'-TBS phosphoramidite monomers follows a similar synthesis cycle to that of standard [solid-phase DNA synthesis](Solid-phase-oligonucleotide-synthesis). The differences are:

- phosphoramidite coupling times are longer in RNA synthesis (up to 6 min) to allow for the steric bulk of the TBS group;
- the capping reagent in RNA synthesis is _tert_-butylphenoxyacetic anhydride, rather than acetic anhydride, to negate the effect of acyl exchange at the protected amino groups of the bases.

After synthesis, the protected oligoribonucleotide is released from the solid support using concentrated aqueous ammonia/ethanol (3:1), and the protecting groups are then removed from the bases by heating at 55 °C in the same mixture for one hour. Finally, the 2'-TBS groups are removed using tetrabutylammonium fluoride in tetrahydrofuran.

{{> figure
    filename    TBS-fluoride-deprotection.svg
    width       295
    height      114
    ::title     Deprotection of the TBS group
    ::subtitle  Mechanism of deprotection of the 2'-_tert_-butyldimethylsilyl (TBS) group with fluoride
<}}

The preparation of synthetic RNA has a number of limitations. The TBS group is not 100% stable to the basic conditions used to removed the _tert_-butylphenoxyacetyl protecting groups, and loss of the TBS group at this stage can lead to phosphodiester chain cleavage and (more seriously) 3'- to 2'-phosphate migration ([figure: RNA-chain-cleavage-phosphate-migration]). This is a particularly severe problem for long oligoribonucleotides as the probability of any given RNA chain containing at least one 5'-2' linkage increases with oligonucleotide length.

It is imperative that RNA is handled with care to avoid contamination with RNAse enzymes that might otherwise result in degradation. RNAses are ubiquitous and sterile conditions are necessary when handing RNA. Prification of RNA is carried out by anion-exchange HPLC and/or reversed-phase HPLC and RNA molecules can be analysed by capillary or gel electrophoresis.

{{> figure
    filename    RNA-chain-cleavage-phosphate-migration.svg
    width       509
    height      328
    ::title     RNA chain cleavage and phosphate migration
    ::subtitle  Mechanisms of RNA chain cleavage (top) and 3'- to 2'-phosphate migration (bottom)
<}}

# Alternative RNA protecting group strategies

Improvements in the synthesis, deprotection and purification of RNA have produced more robust methods that enhance the yield and purity of the final product.

## 2-O-Triisopropylsilyloxymethyl (TOM) protecting groups

The 2-_O_-triisopropylsilyloxymethyl protecting group (TOM, [figure: tom-rna-monomers]) is a simple modification of the widely used 2'-_O_-_tert_-butyldimethylsilyl (TBDMS/TBS) group and therefore is compatible with the currently established RNA synthesis chemistries.

{{> figure
    filename    tom-rna-monomers.svg
    width       483
    height      464
    ::title     TOM-protected RNA monomers
    ::subtitle  Structures of 2-_O_-triisopropylsilyloxymethyl (TOM)-protected phosphoramidite monomers, used in solid-phase RNA synthesis
<}}

The spacer between the 2'-position of the nucleoside and the silyl group overcomes the problem of steric hindrance encountered during the phosphoramidite coupling step of TBS-based RNA synthesis. The TOM protecting group is stable to the conditions required to remove the protecting groups from the heterocyclic bases and phosphodiesters (10 M methylamine in ethanol/water (1:1)). The TOM group is cleaved using 1 M TBAF in THF, without any degradation of the RNA oligonucleotide, and is stable to both basic and weakly acidic conditions. Importantly, the acetal group prevents 2'- to 3'-silyl migration in the nucleoside, which can occur with TBS protection, leading (after phosphitylation) to isomeric RNA monomers ([figure: TBS-RNA-monomer-synthesis]). Using the TOM protecting strategy, high coupling yields are achieved, in shorter reaction times, facilitating the synthesis of long RNA molecules.

## 2'-bis(2-Acetoxyethoxy)methyl (ACE) protecting groups

By changing the 5'-_O_-DMT protecting group to a silyl ether, the 2'-bis(2-acetoxyethoxy)methyl (ACE) orthoester group has been used to protect the 2'-OH of RNA monomers ([figure: ACE-RNA-monomers]). The 5'-_O_-silyl ether can be removed with fluoride under neutral conditions that are compatible with the acid-labile 2'-_O_-ACE group. The ACE esters are stable to the conditions of oligonucleotide synthesis but are deacylated during the base deprotection step, resulting in a 2'-bis(2-hydroxyethoxy)methyl orthoester, which can be removed using weakly acidic conditions (pH 3, 10 min, 55 °C). This method also produces very high coupling yields with short reaction times. A major advantage is that the RNA can be purified with the 2'-protecting group attached. The 2-ACE-protected RNA is resistant to RNase degradation, and so can be stored for extended periods and deprotected prior to use.

{{> figure
    filename    ACE-RNA-monomers.svg
    width       432
    height      508
    ::title     ACE-protected RNA monomers
    ::subtitle  Structures of 2'-Bis(2-acetoxyethoxy)methyl (ACE)-protected phosphoramidite monomers, used in solid-phase RNA synthesis
<}}

## 2'-Thiomorpholine-4-carbothioate (TC) protecting groups

The use of 2'-thiomorpholine-4-carbothioate (TC) protecting groups ([figure: TC-RNA-monomers]), which can be removed in the same conditions used in the deprotection of the nucleobase protecting groups, allows the one-step deprotection of all protecting groups. This is in contrast to the other methods described here, all of which require separate deprotection steps for the 2'-OH protecting groups and nucleobase protecting groups.

{{> figure
    filename    TC-RNA-monomers.svg
    width       486
    height      460
    ::title     TC-protected RNA monomers
    ::subtitle  Structures of 2'-thiomorpholine-4-carbothioate (TC)-protected phosphoramidite monomers, used in solid-phase RNA synthesis
<}}

A solution of anhydrous ethylenediamine in toluene is used for this global deprotection of TC and base protecting groups ([figure: TC-carbothioate-deprotection]).

{{> figure
    filename    TC-carbothioate-deprotection.svg
    width       775
    height      559
    ::title     Deprotection of the TC group
    ::subtitle  Mechanism of deprotection of the 2'-thiomorpholine-4-carbothioate (TC) protecting group (1) with ethylenediamine, via one of two mechanisms ((i) or (ii)), to form the deprotected product (4).
<}}


Description of methods for oligo purification: gel filtration, HPLC (ion exchange and reversed phase), polyacrylamide gel electrophoresis.

Purification and characterisation of oligonucleotides


# Introduction

Oligonucleotide synthesis involves a large number of individual reactions, which leads inevitably to the accumulation of impurities such as truncated oligonucleotides and sequences containing chemically-modified bases. These impurities must be removed after synthesis.

Chromatography is a technique for the separation of the components in a mixture by differential adsorption onto an adsorbent surface (stationary phase). The components are then eluted by a solvent or mixture of solvents (mobile phase).

In addition, it can be desirable to characterise the final product before use. A powerful method for this is mass spectrometry, often combined with chromatography.

# Gel filtration

Gel filtration is the separation of the components of a mixture on the basis of molecular size, and is the simplest form of chromatography for oligonucleotide purification. Cleaved protecting groups and short truncated sequences are retained in the gel matrix while larger oligonucleotide molecules elute quickly through the gel filtration column ([figure: gel-filtration]). Gel filtration also removes salts from oligonucleotides that have been purified by [HPLC](#High-performance-liquid-chromatography-HPLC).

{{> figure
    filename    gel-filtration.svg
    width		469
    height		333
    owidth      375
    oheight	    265
    ::title     Gel filtration
    ::subtitle  Schematic representation of gel filtration for the purification of oligonucleotides. Large molecules elute quickly through the column, while smaller molecules are retained in the gel matrix and elute more slowly.
<}}

While gel filtration is a very simple and useful technique, it does not remove impurities of a similar size to the desired product. It is recommended as the sole purification method only for very small (< 20 bases), unmodified oligos.

# High-performance liquid chromatography (HPLC)

High-performance liquid chromatography (HPLC) is an efficient method for the analysis and purification of synthetic oligonucleotides. It can be used for the separation of oligomers of similar size and is amenable to automation. HPLC systems typically consist of the following components: an injector, pumps for delivery of solvents through the column, an interchangeable HPLC column, a column oven, a solvent mixing system, and a detector (normally UV/visible), all controlled by a PC.

## Reversed-phase HPLC

Reversed-phase HPLC separates oligonucleotides and contaminants on the basis of differences in hydrophobicity. Crude oligonucleotides contain the desired product, failure sequences and chemically modified by-products, all of which will vary in hydrophobicity. For example, any oligonucleotide strands that have not been completely deprotected will be more hydrophobic than the fully deprotected product. Oligonucleotides with modifications such as fluorescent dyes are particularly suited to reversed-phase HPLC purification and reversed-phase HPLC is particularly effective in purifying short oligonucleotides (under 40 bases in length).

[figure: hplc] demonstrates how reversed-phase HPLC works. The mixture of oligonucleotides is injected onto a column that is made up of hydrocarbon chains (normally C~8~ or C~18~) chemically bound to a silica support of defined pore size, typically about 300 Å. Elution with aqueous ammonium acetate forces the oligonucleotide to interact strongly with the reversed-phase column. Further elution with a gradient of acetonitrile in aqueous ammonium acetate causes the individual components in the mixture to enter the mobile phase, which becomes more hydrophobic as the percentage of acetonitrile increases. Molecules with a greater degree of hydrophobicity elute more slowly, and this leads to separation of the components in the mixture.

{{> figure
    filename    hplc.svg
    width       273
    height      569
    ::title     Reversed-phase HPLC
    ::subtitle  Schematic representation of reversed-phase HPLC as used in the purification of oligonucleotides. Hydrophilic impurities (orange) elute from the column first, followed by the product (blue), and finally the hydrophobic impurities (blue).
<}}

Reversed-phase HPLC has certain limitations: the longer the oligonucleotide, the more difficult it is to separate the failure sequences of a similar length, and stable [secondary structure](#Secondary-structure-in-oligonucleotides) (e.g. hairpin loops) can cause an oligonucleotide to elute as a broad peak or even a series of peaks. Placing the HPLC column in a column oven at 60 °C can overcome this problem by (temporarily) destroying secondary structure.

pH and salt concentration (ionic strength), as well as other additives and variables, can also have an effect on reversed-phase HPLC. Under reversed-phase conditions, retention is longest when the oligonucleotide is most hydrophobic. Therefore, the pH at which the analyte is least ionised will lead to the longest retention.

The pH is often adjusted using a salt. This can additionally affect retention time by forming ion pairs (longer retention) and altering the interaction with the stationary phase.

## Anion-exchange HPLC

Anion-exchange HPLC separates oligonucleotides on the basis of charge differences. Each oligonucleotide in the crude mixture is a polyanion and has a particular net negative charge based on the number of phosphodiester groups in the molecule. The stationary phase on the anion-exchange column is a tertiary amine. Separation of the crude mixture is accomplished by slowly increasing the ionic strength of the mobile phase to weaken the interactions between the oligonucleotide (polyanion) and the cationic stationary phase. The longer, more highly charged oligonucleotides elute later than the shorter ones. This method is effective in separating oligonucleotides in the region of 5 to 50 bases in length from failure sequences.

## Which type of HPLC should be used to purify oligos?

Reversed-phase HPLC is used routinely in the purification of oligos.

Oligos with a high degree of secondary structure (e.g. hairpin loops) give rise to multiple peaks. This secondary structure can be eliminated in the presence of a highly alkaline pH, which eliminates hydrogen-bonding interactions. Because anion-exchange HPLC can tolerate high pH, anion-exchange HPLC is a useful technique in the purification of oligos with secondary structure.

Certain oligonucleotide sequences can be predicted to give rise to secondary structure; in simple terms, oligos containing a high proportion of G and C bases (high GC content) are likely candidates for anion-exchange HPLC purification. Anion-exchange HPLC is also useful for purifying longer oligos (40–100 bases).

Hydrophobic oligonucleotides can be purified by ion-exchange HPLC by including an organic solvent in the mobile phase.

## Trityl-on (DMT-on) purification

Oligonucleotides can be synthesized with the terminal 5'-DMT group attached (DMT on; trityl-on). This increases the hydrophobicity of the full-length oligonucleotide relative to all failure sequences (which will have been capped with acetic anhydride during synthesis and will not have a 5'-DMT group). Reversed-phase HPLC purification is very effective in separating the DMT-protected product from impurities. Finally, after purification, the 5'-DMT group is removed. Trityl-on purification is effective for the purification of long oligonucleotides (40 to 150 nucleotides). However, the removal of the DMT group (which requires acidic conditions) can lead to some depurination of the oligonucleotide.

## Large-scale HPLC

To save materials and time, it is common to optimise HPLC gradients on an analytical (small) scale and convert them to the preparative scale using a [calculator](https://www.waters.com/webassets/other/lp/prep_calc/AnalyticaltoPrep/NewPrepcalculator.htm). The conversion is done by taking into account geometrical considerations such as column dimensions, the size of the particles within the column, the flow rate and the dwell volume (volume between the buffer intake point and the detector) of the HPLC system.

# Fluorous affinity chromatography

Fluorous affinity chromatography exploits the concept of _fluorophilicity_: the high affinity of highly fluorinated organic compounds for other fluorinated compounds. Fluorous affinity chromatography requires that oligonucleotides are labelled with a fluorous tag (using a phosphoramidite with a 5'-DMT group derivatized with a perfluorinated alkyl chain; [figure: fluorous-phosphoramidite-oligonucleotide]) during [solid-phase oligonucleotide synthesis](Solid-phase-oligonucleotide-synthesis). As the fluorous label is added at the end of oligonucleotide synthesis, failure sequences will not have a fluorous label. Fluorous affinity chromatography is related to [trityl-on purification](#Trityl-on-DMT-on-purification).

{{> figure
    filename    fluorous-phosphoramidite-oligonucleotide.svg
    width       735
    height      204
    ::title     Structures of a fluorous nucleoside phosphoramidite monomer (left) and an oligonucleotide with a 5'-fluorous tag (right)
    ::subtitle  The fluorous tail is coloured blue.
<}}

After solid-phase synthesis, the oligo is dissolved in buffer and passed through a fluorous column (containing a polymer resin to which fluorinated organic groups are bound). The labelled oligonucleotide will bind to the column strongly, while the unlabelled impurities will not interact with the column and will pass through. Subsequent washing will not release the fluorinated oligo from the column, such is the strength of the fluorophilic interaction. After all the impurities have been removed, the desired oligonucleotide is detritylated on the column (using trifluoroacetic acid), which breaks the strong interaction between the oligo and the column. The deprotected oligo can now be eluted from the column ([figure: fluorous-affinity-chromatography]).

{{> figure
    filename    fluorous-affinity-chromatography.svg
    width		470
    height	    398
    owidth	    376
    oheight	    318
    ::title     Fluorous affinity chromatography
    ::subtitle  Schematic representation of fluorous affinity chromatography as used in the purification of oligonucleotides. Non-labelled impurities (black) elute quickly, while the fluorous-labelled desired oligonucleotide (light blue) remains bound strongly to the column. On-column detritylation frees the desired oligonucleotide (dark blue) from the column, and allows the desired oligo to be eluted.
<}}

Fluorous affinity chromatography is suitable for long oligonucleotides, owing to the strength of the fluorophilic interaction. This makes it a useful method for the purification of oligos that are difficult to purify using other methods. However, the incorporation of the 5'-fluorous tag during solid-phase synthesis, which requires special 5'-fluorous phosphoramidite monomers ([figure: fluorous-phosphoramidite-oligonucleotide], left), adds complexity and prevents the use of other 5'-modifications.

# Polyacrylamide gel electrophoresis (PAGE)

Electrophoresis separates molecules based on their electrical charge and hydrodynamic properties, which are a function of chain length. PAGE purification can be used for oligos of any length, but it recommended particularly for long oligos, of around 100 bases and longer, for which other purification methods present difficulties. PAGE purification typically results in oligos of very high purity, but at the expense of yield; and can be time-consuming.

# Secondary structure in oligonucleotides

The purification of certain oligonucleotides is complicated by the presence of secondary structures such as duplexes, hairpin loops ([figure: DNA-hairpin-loop-formation], [figure: DNA-hairpin-loop-3d]) and guanine quartets (quadruplexes). Secondary structure in an oligonucleotide will influence the choice of chromatographic purification method.

{{> figure
    filename    DNA-hairpin-loop-formation.svg
    width       217
    height      110
    ::title     Formation of a hairpin loop
    ::subtitle  Schematic representation of the formation of a hairpin loop from a linear DNA oligonucleotide
<}}

{{> figure
    filename    DNA-hairpin-loop-3d.webp
    width       545
    height      481
    ::title     Hairpin loop
    ::subtitle  3D representation of a hairpin loop in an AT-rich DNA oligonucleotide
<}}

Polymer-based anion-exchange HPLC columns are useful for purifying oligonucleotides that adopt extensive secondary structure. This is because the stationary phase is stable at high pH, so HPLC purification can be carried out at pH 12. At this pH, secondary structures in the oligonucleotide are destroyed because N(3) of thymine and N(1) of guanine are deprotonated, and inter-base hydrogen bonds are destabilized. Purification at high pH is not feasible for RNA because there is a risk of 3'- to 2'-phosphate migration and cleavage of the phosphodiester backbone (see [solid-phase synthesis of RNA](RNA-oligonucleotide-synthesis#Solid-phase-synthesis-of-RNA)).

# Recommended methods of purification for different applications

The purification method you choose will depend on the sequence and scale of the oligo, but also on its intended use.

| Application           | Length      | Recommended purification method(s) |
| --------------------- | ----------- | ---------------------------------- |
| PCR primer            | < 20 bases  | Gel filtration only, or HPLC       |
| PCR primer            | 20–40 bases | HPLC                               |
| PCR probe             | any         | HPLC                               |
| X-ray crystallography | any         | Double HPLC or PAGE                |


Oligo labelling: components of a label, methods of labelling and common labels including phosphate, biotin, aminohexyl (aminolink), thiol; indirect enzymic labelling.

Synthesis and applications of chemically modified oligonucleotides

Introduction to fluorescence, and the need for fluorescent oligos. Common fluorescent dyes used with oligos, and how they are added.

Synthesis and properties of fluorescent oligonucleotides


# Introduction

Fluorophores are very useful and sensitive labels for biomolecules, and fluorescent labels are by far the most common labels used for synthetic oligonucleotides. Many different fluorophores can be readily attached to oligonucleotides and they form the basis of the detection systems used in DNA sequencing, forensic science and genetic analysis. The principles of fluorescence and the synthesis of fluorescent oligonucleotides are described below.

The major _applications_ of fluorescent and fluorogenic oligonucleotides are discussed in the section on [sequencing, forensic analysis and genetic analysis](Sequencing-forensic-analysis-and-genetic-analysis).

# Principles of fluorescence

Prior to excitation, a fluorescent molecule is in the lowest vibrational level of the electronic ground state S~0~. Excitation by the absorption of light of an appropriate wavelength produces a change to an excited vibrational level of the first excited singlet state S~1~. After a very short time period (10^−12^ s) the lowest vibrational level of S~1~ is reached by a series of non-radiative energy transfer processes known as internal conversion, in which the molecule collides with surrounding solvent molecules and loses energy as heat. The system remains at this level for a significantly longer period (10^−9^ s) before a transition occurs to the S~0~ state with the emission of a photon. This process is known as fluorescence ([figure: Jablonski-diagram]).

{{> figure
    filename    Jablonski-diagram.svg
    width       287
    height      334
    ::title     Jablonski diagram, showing the energy levels of a fluorescent molecule in solution.
<}}

Eventually photobleaching occurs and the molecule becomes non-fluorescent. Fluorescence emission is of lower energy than absorption and is therefore observed at a higher wavelength. The difference in frequency between excitation and emission is called the Stokes shift. A large Stokes shift facilitates the measurement of fluorescence and leads to a lower background signal.

# Common fluorophores

Many different fluorescent dyes are available. All these molecules are organic dyes that emit in the visible region of the electromagnetic spectrum. They can be conveniently excited by laser light.

| Dye         | Absorption wavelength / nm | Emission wavelength / nm | Colour        |
| ----------- | -------------------------- | ------------------------ | ------------- |
| FAM         | 495                        | 520                      | Green/yellow  |
| TET         | 525                        | 550                      | Orange/yellow |
| HEX         | 535                        | 565                      | Pink          |
| JOE         | 520                        | 548                      | Pink/Green    |
| Cyanine-3   | 550                        | 570                      | Dark pink     |
| TAMRA       | 550                        | 575                      | Red           |
| ROX         | 580                        | 605                      | Purple        |
| Cyanine-3.5 | 580                        | 595                      | Purple/blue   |
| Cyanine-5   | 650                        | 670                      | Blue          |
| Cyanine-5.5 | 675                        | 695                      | Blue          |

Some of the most commonly employed fluorescent dyes are 6-carboxyfluorescein (FAM), its tetra- and hexachlorinated analogues (TET and HEX), and carboxy-X-rhodamine (ROX) ([figure: FAM-HEX-TET-ROX-phosphoramidites]). However, their properties are not ideal; they have broad emission spectra, are non-fluorescent at low pH, and are susceptible to photobleaching. These inadequacies led to the development of alternatives such as the cyanine Dyes (Cyanine Dyes) which have the general structure shown in [figure: Cyanine-dyes]. Their spectral properties can be tuned by the appropriate choice of heterocyclic nuclei (X and Y) and the length of the polymethine chain (_n_) to provide fluorophores that emit in the range 500 to 750 nm. The groups R~1~ - R~4~ can be varied to provide the desired aqueous solubility.

{{> figure
    filename    Cyanine-dyes.svg
    width       410
    height      99
    ::title     Cyanine dyes
    ::subtitle  General structure of the cyanine dyes. The spectral properties of the Cyanine dyes can be tuned by altering the heterocyclic nuclei (X and Y) and the length of the polymethine chain (_n_). The groups R~1~ - R~4~ can be varied to provide aqueous solubility.
<}}

For further information see FAM (fluorescein), HEX, JOE, ROX, TAMRA, TET, Texas Red® and others] and Cyanine dyes. Other important fluorescent dyes include BODIPY®, Alexa® dyes, ATTO dyes and Yakima Yellow.

# Fluorescent base analogues

Standard nucleobases (A, G, C, T and U) are not fluorescent under standard conditions. Fluorescent base analogues are structural analogues of the standard bases that are fluorescent, while still forming hydrogen bonds with standard bases.

Fluorescent labels are commonly attached to the double helix at the end of a linker, which places the fluorophore relatively far from the DNA bases. This is useful in many circumstances, but sometimes it is necessary to incorporate a fluorophore closer to the DNA or RNA double helix, without perturbing the helix. As fluorescent base analogues are located rigidly in the double helix, their movement is restricted. This restricted movement results in a more predictable orientation of the fluorophore, an advantage in many applications such as fluorescence resonance energy transfer (FRET) and fluorescence anisotropy.

## 2-Aminopurine

2-Aminopurine ([figure: 2-aminopurine]) is the original, and most frequently used, fluorescent base analogue. It is highly fluorescent, and readily available.

{{> figure
    filename    2-aminopurine.svg
    width       380
    height      140
    ::title     2-Aminopurine
    ::subtitle  Structure of 2-aminopurine (2AP), the first fluorescent base analogue, and structures of base pairs between 2-aminopurine and thymine (T) and cytosine (C).
<}}

Like almost all fluorescent base analogues, 2-aminiopurine has a quantum yield that is heavily dependent on its environment. The DNA duplex quenches the fluorescence of more fluorescent base analogues, limiting their usefulness. This effect can be pronounced: 2-aminopurine has a good quantum yield of 0.68 in water, but this decreases to less than 0.02 in single-stranded DNA and less than 0.01 in double-stranded DNA.

## Tricyclic cytosine analogues

The tricyclic cytosine analogues ([figure: tricyclic-cytosine-analogues]), intended for use in antisense therapy but recently discovered to be strongly fluorescent, are the only fluorescent base analogues to have quantum yields that are not affected appreciably by the environment. The quantum yield of the tricyclic cytosine analogue 1,3-diaza-2-oxophenothiazine (tC) does not vary much whether it is in monomeric form, in single- and double-stranded DNA; and whatever bases surround it.

{{> figure
    filename    tricyclic-cytosine-analogues.svg
    width       432
    height      149
    ::title     Tricyclic cytosine fluorescent base analogues
    ::subtitle  Structures of cytosine (C), and the tricyclic cytosine analogues 1,3-diaza-2-oxophenothiazine (tC), 1,3-diaza-2-oxophenoxazine (tC^O^) and 7-nitro-1,3-diaza-2-oxophenothiazine (tC~nitro~) - the only fluorescent base analogues with quantum yields not appreciably affected by environment.
<}}

Like cytosine, the tricyclic cytosine analogues form hydrogen-bonding interactions with guanine ([figure: tC-hydrogen-bonding]), but not with adenine. If cytosine is replaced with tC in a DNA duplex, the DNA adopts the normal B-form, and only small distortions in the helix are observed around the base analogue: tC is therefore an excellent fluorescent base analogue.

{{> figure
    filename    tC-hydrogen-bonding.svg
    width       167
    height      155
    ::title     Structure of the G·tC base pair
<}}

Relative to cytosine, tC has makes increased base stacking interactions in the DNA duplex, which means that duplexes containing tC are slightly more stable than analogous unmodified DNA.

The oxygen derivative of tC, tC^O^ ([figure: tricyclic-cytosine-analogues]), is also an excellent base analogue and, with an extinction coefficient (_E_) more than double that of tC, tC^O^ is the brightest fluorescent base analogue. The 7-nitro analogue of tC, tC~nitro~ ([figure: tricyclic-cytosine-analogues]), is not fluorescent under standard conditions, but is used as a quencher, in combination with tC or tC^O^ in a FRET pair.

| Name            | λ~max~ / nm (absorption) | λ~max~ / nm (emission) | _E_ at λ~max~ | Φ (solution) | Φ (duplex) | τ / ns (solution) | τ / ns (duplex) |
| --------------- | ------------------------ | ---------------------- | ------------- | ------------ | ---------- | ----------------- | --------------- |
| 2-aminopyridine | 303                      | 371                    | 3 600         | 0.68         | < 0.01     | -                 | -               |
| tC              | 395                      | 505                    | 4000          | 0.20         | 0.16–0.21  | 3.7               | 6.3             |
| tC^O^           | 360                      | 465                    | 9000          | 0.30         | 0.22       | 3.4               | 4.1             |
| tC~nitro~       | 440                      | -                      | 5400          | -            | -          | -                 | -               |

_E_ = extinction coefficient; Φ = quantum yield; τ = fluorescence lifetime. Data from Wilhelmsson, Q. Rev. of Biophys. 43, 2 (2010), 159–183 and other literature sources cited therein, and www.glenresearch.com.

The tricyclic cytosine analogues tC, tC^O^, and tC~nitro~ can be incorporated into oligonucleotides during solid-phase synthesis via their commercially-available phosphoramidites ([figure: tC-phosphoramidites]).

{{> figure
    filename    tC-phosphoramidites.svg
    width       602
    height      247
    ::title     Tricyclic cytosine analogue phosphoramidites
    ::subtitle  Structures of tricyclic cytosine analogue phosphoramidites, for incorporation of the tricyclic cytosine analogues tC, tC^O^ and tC~nitro~ into oligonucleotides during phosphoramidite oligo synthesis
<}}

# Synthesis of fluorescent oligonucleotides

## Synthesis of oligonucleotides with the 5'-fluorescent dyes

### FAM, HEX, TET and ROX

The synthesis of 5'-ROX and TAMRA oligonucleotides by post-synthetic labelling using 5'-amino-modified oligonucleotides and active esters of ROX and TAMRA is discussed in the article on [chemically modified oligonucleotides](Synthesis-and-applications-of-chemically-modified-oligonucleotides/#5-Aminohexyl-oligonucleotides-and-post-synthetic-labelling). This method is used when the dye is unstable to oligonucleotide synthesis/deprotection conditions.

It is more convenient and efficient to add the fluorescent dye during solid-phase synthesis and a number of dye phosphoramidites have been synthesized to facilitate this. Fluorescein and its tetra- and hexachlorinated derivatives TET and HEX are sufficiently stable to survive oligonucleotide deprotection so they are added to oligonucleotides as phosphoramidite monomers ([figure: FAM-HEX-TET-ROX-phosphoramidites]).

{{> figure
    filename    FAM-HEX-TET-ROX-phosphoramidites.svg
    width       620
    height      311
    ::title     Fluorescein phosphoramidite monomer, and chlorinated derivatives
    ::subtitle  Structures of the fluorescein (FAM) phosphoramidite monomer, and its chlorinated derivatives HEX, TET and ROX.
<}}

The phenolic hydroxyl groups are protected as pivaloyl esters to prevent side-reactions during oligonucleotide synthesis, and a six-carbon spacer is inserted between the dye and the phosphoramidite group to distance the dye from the DNA and minimize the possibility of fluorescence quenching induced by the proximity of the dye to DNA bases.

### Cyanine dyes

The Cyanine dyes are particularly useful when high quantum yields and resistance to photobleaching are important, and Cyanine dye analogues are available that cover a wide range of the visible absorption and emission spectrum. Some Cyanine dyes are available as phosphoramidites ([figure: Cyanine-dye-phosphoramidite-monomers]) for addition to the 5'-end of oligonucleotides during solid-phase synthesis. These monomers can be incorporated within the nucleotide sequence of oligonucleotides as they have a DMT-protected hydroxyl group and a phosphoramidite group. However, this would destabilize any duplex formed between the oligonucleotide and a complementary strand, as the monomers do not possess a heterocyclic base, and therefore have no means of base pairing. Cyanine dye phosphoramidites are therefore typically added to the 5'-terminus of oligonucleotides, although 3'-labelling is also carried out for special applications.

{{> figure
    filename    Cyanine-dye-phosphoramidite-monomers.svg
    width       368
    height      184
    ::title     Cyanine dye phosphoramidite monomers
    ::subtitle  Structures of cyanine dye phosphoramidite monomers
<}}

Cyanine dyes are not stable to the standard conditions of oligonucleotide deprotection (concentrated aqueous ammonia for 5 hours at 55 °C) but do withstand brief treatment (1 hour at 55 °C) with this reagent. Therefore it is necessary to use the more labile "ultramild" dmf-protected dG monomer rather than the more common isobutyryl dG phosphoramidite in the synthesis of Cyanine dye-labelled oligonucleotides (see [Deprotection of heterocyclic bases](Solid-phase-oligonucleotide-synthesis#Deprotection-of-heterocyclic-bases)).

## Synthesis of oligonucleotides with internal fluorescent dyes

Fluorescent dyes can be incorporated within oligonucleotide sequences by attaching them to the heterocyclic bases at positions that do not interfere with inter-base hydrogen bonding. The most convenient loci are the 5-position of thymine and cytosine and the 7-position of 7-deaza dG and dA. In practical terms the easiest base to modify is thymine ([figure: fluorescein-phosphoramidite-monomer]) and a synthetic scheme for the synthesis of labelled thymidine phosphoramidite monomers is shown in [figure: fluorescein-phosphoramidite-monomer].

{{> figure
    filename    fluorescein-phosphoramidite-monomer.svg
    width       386
    height      263
    ::title     Fluorescein phosphoramidite monomer
    ::subtitle  Structure of the monomer used to label oligonucleotides with fluorescein at thymidine sites
<}}

2'-Deoxyuridine is converted to its 5-iodo derivative by treatment with iodine monochloride, and the 5'-DMT group is then added. In a parallel reaction sequence an aminoalkyl side chain is built up by reacting propargylamine with TFA-protected 6-aminohexanoic acid. This is coupled to the 5-position of 5'-DMT-5-iodo-2'-deoxyuridine in a Sonogashira reaction using tetrakistriphenylphosphine palladium (§0§), copper iodide and triethylamine in DMF. The product of this reaction is a very useful intermediate with a protected amino group attached via a linker to the 5-position of thymidine. The trifluoroacetyl protecting group can be removed by treatment with ammonia and a range of labels can be added as active esters or as carboxylic acids using a diimide coupling reagent ([figure: phosphoramidite-monomer-label-attachment]). The monomer in [figure: fluorescein-phosphoramidite-monomer] can be made by this method. It is used to add internal labels to oligonucleotides at thymidine sites. The fluorescein and linker moieties do not affect the ability of the labelled thymine to form base pairs.

{{> figure
    filename    phosphoramidite-monomer-label-attachment.svg
    width       507
    height      562
    ::title     Fluorescently labelled phosphoramidite monomer synthesis
    ::subtitle  Scheme showing the attachment of a fluorescent label to a nucleoside phosphoramidite
<}}


PCR; DNA sequencing; STR analysis (fingerprinting); SNPs; FISH; Real-time PCR: molecular probes (including FRET, molecular beacons, Scorpions).

Sequencing, forensic analysis and genetic analysis

Nucleic acids with non-natural backbones: PNA (peptide nucleic acid), LNA (locked nucleic acid), and triazole DNA (synthesized using click chemistry)

Nucleic acid analogues


# Peptide nucleic acid (PNA)

Peptide nucleic acid (PNA) is a nucleic acid analogue in which the normal phosphate linkage found in DNA and RNA is replaced by an neutral peptide-like _N_-(2-aminoethyl)glycine backbone ([figure: PNA-oligo], [figure: PNA-duplex-3d]).

{{> figure
    filename    PNA-oligo.svg
    width       304
    height      148
    ::title     Chemical structure of a PNA oligonucleotide
<}}

{{> figure
    filename    PNA-duplex-3d.webp
    width       737
    height      727
    ::title     Three-dimensional structure of a PNA duplex
<}}

## Synthesis

PNA synthesis is usually performed by automated solid-phase methods, as with [DNA](Solid-phase-oligonucleotide-synthesis) and [RNA](RNA-oligonucleotide-synthesis) oligonucleotide synthesis. The mechanism of PNA synthesis has much in common with solid-phase peptide synthesis.

### Solid-phase peptide synthesis

The coupling step in peptide synthesis is an amide bond coupling reaction between two amino acids ([figure: solid-phase-peptide-synthesis]). The growing peptide chain is attached to the resin via an ester or amide linkage, and the amine of the new unit is protected with a 9-fluorenylmethyloxycarbonyl (Fmoc) protecting group.

{{> figure
    filename    solid-phase-peptide-synthesis.svg
    width       543
    height      156
    ::title     Solid-phase peptide synthesis
<}}

The carboxylic acid must be activated in order for the coupling reaction to proceed. The carboxylic acid is first converted to a _O_-acylisourea by reaction with a carbodiimide. Formation of diisipropylurea drives the reaction to completion. The _O_-acylisourea, which is susceptible to rearrangement to the unreactive _N_-acylurea, is spontaneously converted to a reactive 1-hydroxybenzotriazole ester, by reaction with 1-hydroxybenzotriazole.}

{{> figure
    filename    diimide-activation-rearrangement.svg
    width       442
    height      315
    ::title     Formation of the activated hydroxybenzotriazole ester ester of a carboxylic acid
    ::subtitle  The carboxylic acid is first converted to a _O_-acylisourea (by reaction with a carbodiimide), which is spontaneously converted to a 1-hydroxybenzotriazole ester (by reaction with 1-hydroxybenzotriazole). This avoids formation of the unreactive _N_-acylurea, by rearrangement of the _O_-acylisourea.
<}}

### Solid-phase PNA synthesis

As in solid-phase peptide synthesis, the coupling step in PNA oligonucleotide synthesis is an amide bond coupling reaction between two amino acids ([figure: PNA-coupling-schematic]).

{{> figure
    filename    PNA-coupling-schematic.svg
    width       495
    height      259
    ::title     PNA coupling
    ::subtitle  Schematic representation of the coupling step in PNA oligonucleotide synthesis
<}}

In PNA synthesis, the carboxylic acid is activated using a benzotriazole uronium salt, such as _O_-(1*H*-benzotriazol-1-yl)-_N_,_N_,_N_',_N_'-tetramethyluronium hexafluorophosphate (HBTU), which, like the combination of HOBt and a diimide, yields a hydroxybenzotriazole ester. The driving force for the reaction is the formation of the by-product tetramethylurea ([figure: PNA-coupling-mechanism]).

{{> figure
    filename    PNA-coupling-mechanism.svg
    width       682
    height      364
    ::title     PNA coupling mechanism
    ::subtitle  Mechanism of the coupling step in PNA oligonucleotide synthesis
<}}

The Fmoc protecting groups are removed by treatment of the PNA oligonucleotide with a solution of 20% piperidine in DMF ([figure: Fmoc-PNA-deprotection]) at the end of each PNA synthesis cycle.

{{> figure
    filename    Fmoc-PNA-deprotection.svg
    width       534
    height      277
    ::title     9-Fluorenylmethyloxycarbonyl (Fmoc) amine deprotection
    ::subtitle  Mechanism of Fmoc deprotection of PNA peptide units in PNA oligonucleotide synthesis
<}}

### Structures of PNA monomers

As the chemistry of PNA oligonucleotide synthesis is different from that of DNA and RNA oligo synthesis, different types of protecting groups are needed for the heterocyclic bases. The benzhydryloxycarbonyl (Bhoc) protecting group is used to protect the primary amines of the heterocyclic bases in PNA oligo synthesis. ([figure: PNA-monomers]).

{{> figure
    filename    PNA-monomers.svg
    width       542
    height      309
    ::title     PNA monomers
    ::subtitle  Structures of monomers used in PNA synthesis
<}}

### Synthesis of PNA monomers, and resin functionalization

The Bhoc-protected heterocyclic base (or unprotected thymine) is alkylated at the _N_(2) position with bromoacetic acid. The resulting nucleobase acetic acid is then coupled with an Fmoc-protected 2-((2-aminoethyl)amino) acetate using a coupling reagent (e.g. a carbodiimide), to yield the monomer ([figure: PNA-monomer-synthesis-resin-functionalization]a). Functionalization of the resin with the PNA monomer is a simple matter of coupling the monomer with the free resin ([figure: PNA-monomer-synthesis-resin-functionalization]b).

{{> figure
    filename    PNA-monomer-synthesis-resin-functionalization.svg
    width       477
    height      198
    ::title     PNA monomer synthesis and resin functionalization
    ::subtitle  Synthesis of monomers for use in PNA synthesis
<}}

### Deprotection of heterocyclic bases

The Bhoc protecting groups on the heterocyclic bases, and the linker between the PNA and the solid support, are acid-labile, and so can be removed in a single step by treatment with trifluoroacetic acid at the end of PNA assembly ([figure: Bhoc-deprotection]).

{{> figure
    filename    Bhoc-deprotection.svg
    width       285
    height      140
    ::title     Benzhydryloxycarbonyl (Bhoc) base deprotection
    ::subtitle  Mechanism of deprotection of Bhoc-protected heterocyclic bases in PNA oligonucleotide synthesis
<}}

## Properties and uses

PNA oligonucleotides have unique properties that have been utilized in many molecular biology and biochemical applications.

PNA oligonucleotides hybridize with complementary DNA and RNA oligonucleotides, by Watson-Crick base pairing. Perhaps surprisingly, PNA oligonucleotides hybridize to complementary DNA/RNA with higher affinity and specificity than normal DNA/RNA oligonucleotides. PNA is not a nucleic _acid_, but a neutral molecule. The lack of a negatively charged backbone reduces electrostatic repulsion between oligonucleotides in PNA:DNA and PNA:RNA hybrids, contributing to increased stability. In contrast to DNA:DNA duplexes, the stability of PNA:DNA duplexes is practically independent of salt concentration.

### PNA probes

Like DNA and RNA, PNA oligonucleotides can be labelled with fluorescent dyes during or after solid-phase synthesis. These PNA probes are used in diagnostics and detection.

The increased specificity of PNA oligonucleotides in forming hybrid duplexes with DNA and RNA means that mismatched base pairs in hydrid PNA:DNA duplexes are particularly unstable. PNA probes are therefore useful in identifying point [mutations](Mutagenesis-and-DNA-repair) and [SNPs](Sequencing-forensic-analysis-and-genetic-analysis#Single-nucleotide-polymorphisms-SNPs). The destabilizing effect of the mismatch can be increased by the use of very short PNA probes. DNA probes often fail to distinguish between certain regions of highly structured RNA, but PNA probes can be used to detect these regions, if the hybridization is carried out at low salt concentration so that the RNA is unstructured.

PNA probes are also used in [fluorescent in situ hybridization](Sequencing-forensic-analysis-and-genetic-analysis#Fluorescence-in-situ-hybridization-FISH) (PNA-FISH). In fluorescent in situ hybridization (FISH), short probes can be difficult to detect (owing to insufficient hybridization and low levels of labelling), while longer probes can induce non-specific background fluorescence. In PNA-FISH, shorter PNA probes can be used (because of the higher specificity of PNA), resulting in less background noise.

Because of their insensitivity to salt concentration, PNA probes can often be used in a wider range of conditions than DNA and RNA probes, which can be useful in applications such as [microarrays](Sequencing-forensic-analysis-and-genetic-analysis#DNA-microarrays).

### Therapeutic applications

PNA is an unnatural molecule, and PNA oligonucletides are not substrates for enzymes that modify DNA, RNA and proteins. DNA and RNA oligonucleotides are effective in [antisense therapy](Oligonucleotides-as-drugs#Antisense-oligonucleotides) in vitro, but are degraded quickly by intracellular nucleases in vivo, and this degradation destroys their antisense activity. PNA oligomers are resistant to degradation by nucleases and are therefore much more effective than DNA and RNA in therapeutic antisense gene silencing.

### PNA in molecular biology

The lack of reactivity of PNA oligonucleotides to nucleic acid- and protein modifying enzymes means that PNA oligonucleotides are incompatible with useful molecular biology techniques such as [PCR](Sequencing-forensic-analysis-and-genetic-analysis#The-Polymerase-Chain-Reaction-PCR), restriction enzymes and proteolysis. However, this allows PNA to used along with DNA in the presence of enzymes: while the DNA is modified by the enzyme, the PNA remains intact. PNA oligonucleotides can be used in the presence of nucleases to form an "artificial restriction" system, enabling the cutting of DNA at sites not accessible using restriction enzymes. PNA oligonucleotides can also be used in PCR, enhancing the amplification of specific DNA products.

# Locked nucleic acid (LNA)

Locked nucleic acid (LNA) is an unnatural conformationally restricted oligonucleotide analogue bearing a close structural resemblance to DNA, but possessing monomer units with a 2'-_O_-4'-_C_-methylene bridge. The bicyclic structure locks the molecule in a C3'-_endo_ sugar (N-type) configuration ([figure: LNA-structure]), ensuring the oligonucleotide adopts the A-form helix (which is associated with high duplex stability).

{{> figure
    filename    LNA-structure.svg
    width       305
    height      229
    ::title     Chemical structure of an LNA oligonucleotide (left), and three-dimensional structure of C(3)-endo LNA (right)
<}}

{{> figure
    filename    LNA-bases-3d.webp
    width       586
    height      532
    ::title     Three-dimensional structure of an LNA oligonucleotide
    ::subtitle  Two successive bases are shown, each attached to a locked bicyclic C3'-_endo_ sugar, with a phosphate group connecting the sugars
<}}

The original LNA oligomers were "oxy-LNAs". Subsequently "thio-LNA" (2'-_S_-CH2-4') and "amino-LNA" (2'-NH-CH2-4') analogues were prepared ([figure: oxy-LNA-thio-LNA-amino-LNA]).

{{> figure
    filename    oxy-LNA-thio-LNA-amino-LNA.svg
    width       364
    height      152
    ::title     Structures of thio- and amino-analogues of LNA (oxy-LNA)
<}}

## Synthesis

LNA oligonucleotides (and mixed LNA/DNA and LNA/RNA oligomers) can be synthesized on standard automated DNA synthesizers using commerically available LNA phosphoramidite monomers ([figure: LNA-monomers]). The only changes to the standard DNA synthesis cycle are increased reaction times for the coupling and oxidation steps. LNA oligonucleotides can be labelled with fluorescent dyes and other chemical modifications in the same way as DNA and RNA oligonucleotides.

{{> figure
    filename    LNA-monomers.svg
    width       494
    height      444
    ::title     Structures of phosphoramidite monomers used in the synthesis of LNA oligonucleotides
<}}

## Properties and uses

LNA forms stable hybrid duplexes with DNA and RNA. Duplexes formed between two LNA oligonucleotides exhibit very high affinity and specificity. LNA units can be incorporated into normal DNA or RNA oligonucleotides to increase the stability of probe-target hybrids, manifested as an increased melting temperature (_T_~m~) of the hybrid duplex. LNA oligonucleotides can be used for mismatch discrimination, their high binding affinity allowing short probes to be used. LNA oligos have been used in triplex forming oligonucleotides, antisense oligonucleotides and microarrays.

The incorporation of LNA bases into antisense DNA oligonucleotides provides both strong affinity towards the RNA target and stability to nucleases. One of these _gapmers_, EZN-3042, has proved effective in inhibiting tumour growth and is currently in clinical trials as a leukaemia drug.

# Unlocked nucleic acid (UNA)

Unlocked nucleic acid (UNA) is an analogue of RNA in which the C2'-C3' bond has been cleaved. Whereas LNA is conformationally restricted (relative to DNA and RNA), UNA is very flexible ([figure: UNA-structure]).

{{> figure
    filename    UNA-structure.svg
    width       120
    height      200
    ::title     Chemical structure of a UNA oligonucleotide
<}}

## Synthesis

UNA can be synthesized using an automated solid-phase approach, using UNA phosphoramidite monomers ([figure: UNA-monomers]).

{{> figure
    filename    UNA-monomers.svg
    width       450
    height      460
    ::title     Structures of phosphoramidite monomers used in the synthesis of UNA oligonucleotides
<}}

## Properties and uses

While the incorporation of LNA units into DNA and RNA oligonucleotides increases the stability of duplexes, UNA has a destabilizing effect. Incorporation of UNA units into an oligonucleotide probe lowers the _T_~m~ of the probe-target duplex. With their complementary effect on probe-target stability, LNA and UNA units can therefore be used to fine-tune the thermodynamic properties of DNA and RNA probes.

UNA bases have been incorporated into siRNA oligonucleotides, used in [RNA interference (RNAi)](Oligonucleotides-as-drugs#RNA-Interference-RNAi). While remaining potent, these UNA-modiﬁed siRNA (UsiRNA) molecules were found to be resistant to degradation in serum, have reduced toxicity and reduced off-target effects.

# Triazole-linked DNA

Recently, triazole-linked DNA oligonucleotides, in which one or more of the natural phosphate backbone linkages are replaced with triazole linkages, have been synthesized using [click chemistry](Click-chemistry-and-nucleic-acids).

The presence of a triazole linker does not perturb the global structure of double-stranded DNA relative to that of natural DNA. Hydrogen bonding between bases either side of the triazole linkage is not disrupted, and the overall structure of the helix is preserved. Small local changes are observed at the site of the triazole, but one of the nitrogen atoms of the triazole appears to mimic the phosphate group, a hydrogen bond acceptor, diminishing the effect of these local structural perturbations ([figure: triazole-linked-DNA-3d]).

{{> figure
    filename    triazole-linked-DNA-3d.webp
    width       386
    height      694
    ::title     Three-dimensional structure of a triazole-linked DNA:DNA duplex
    ::subtitle  One of the two DNA strands contains a single triazole linkage (coloured orange).
<}}

## Synthesis by click chemistry

Triazole-linked oligonucleotides are assembled from smaller oligonucleotides with terminal alkyne and azide groups, in the copper-catalyzed azide-alkyne cycloaddition (CuAAC) reaction, the canonical [click chemistry](Click-chemistry-and-nucleic-acids) reaction.

{{> figure
    filename    triazole-DNA-synthesis.svg
    width       360
    height      188
    ::title     Synthesis of triazole-linked DNA
    ::subtitle  Triazole-linked DNA is synthesized by click chemistry: DNA oligonucleotides with terminal alkyne and azide groups are ligated to form a larger oligonucleotide containing a triazole linkage.
<}}

## Properties and uses

Triazole-linked oligonucleotides form duplexes with complementary strands of natural DNA. Triazole-linked DNA is read through by DNA and RNA polymerases, and is functional as a PCR template in _E. coli_. Triazole-linked RNA has been used in the synthesis of an ribozyme: despite the presence of a triazole linkage at the active site, the triazole-linked ribozyme was found to be catalytically active.

The assembly of large oligonucleotides by "click" ligation of smaller oligonucleotides has the potential to overcome the practical size limit of around 200 bases that applies to solid-phase phosphoramidite oligonucleotide synthesis. Long oligonucleotides can be made by enzymatic ligation of shorter oligos, but this has problems: it does not work well on a large scale and does not tolerate epigenetic modifications. The chemical ligation of shorter oligos using click chemistry overcomes these problems, and could be used in the total synthesis of very long strands of DNA including synthetic genes.


Antisense oligonucleotides (hybridize to specific mRNAs inhibiting expression of particular proteins); triplexes and quadruplexes; RNAi (RNA interference).

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