A Continuous Strand of Nucliec Acids

Nucleic Acid

Geochemistry, Organic

Jürgen Rullkötter , in Encyclopedia of Physical Science and Technology (Third Edition), 2003

I.B.1 Nucleic Acids and Proteins

Nucleic acids, such as ribonucleic acids (RNA) or desoxyribonucleic acids (DNA), are biological macromolecules that carry genetic information. They consist of a regular sequence of phosphate, sugar (pentose), and a small variety of base units, i.e., nitrogen-bearing heterocyclic compounds of the purine or pyrimidine type. During biosynthesis, the genetic information is transcribed into sequences of amino acids which occur as peptides, proteins, or enzymes in the living cell. These macromolecules vary widely in the number of amino acids and thus in molecular weight. They account for most of the nitrogen compounds in the cell and serve in such different functions as the catalysis of biochemical reactions and the formation of skeletal structures (e.g., shells, fibers, muscles).

During sedimentation of decayed organisms, nucleic acids and proteins are readily hydrolysed chemically or enzymatically into smaller, water-soluble units. Amino acids occur in rapidly decreasing concentrations in Recent and subrecent sediments, but may also survive in small concentrations in older sediments. A certain proportion of the nucleic acids and proteins reaching the sediment surface may be bound into the macromolecular kerogen network of the sediments and there become protected against further rapid hydrolysis.

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Organized Monolayers and Assemblies: Structure, Processes and Function

Dev Kambhampati , Wolfgang Knoll , in Studies in Interface Science, 2002

2.1 Materials

Biotinylated DNA probes and MR-121 labeled fluorescent DNA targets were procured from Boehringer Mannheim. Cy5 labeled DNA targets (T2, T3, T4) were obtained from MWG Biotech. The sensor surface regeneration solutions, viz. sodium hydroxide (50   mM, Biacore AB) and HEPES buffer (pH   4.5, Aldrich) were used directly as received. PBS Buffer (10   mM phosphate, 2.7   mM KCl, 150   mM NaCl, pH 7.4, Sigma) was used to dissolve the nucleic acid solutions and was also used as the hybridization buffer.

Nucleic Acid Probe Sequences:

DNA Probe (15   T spacer):	5'-T15 – TGTACATCA CAA CTA – 3'	(P1)
DNA Probe (30   T spacer):	5'-T30 – TGT ACA TCA CAA CTA – 3'	(P2)

Nucleic Acid Target Sequences:

15mer Mismatch 1- 3'– ACA TGC AGT GTT GAT – MR 121 –5'	(T1)
25mer Mismatch 1–3' – T5 ACA TGC AGT GTT GAT T5 – Cy 5–5'	(T2)
45mer Mismatch 1- 3' – T15 ACA TGC AGT GTT GAT T15 – Cy5 – 5'	(T3)
75mer Mismatch 1- 3' – T30 ACA TGC AGT GTT GAT T30 – Cy5 – 5'	(T4)
45mer Mismatch 0- 3' – T15 ACA TGT AGT GTT GAT T15 – Cy5 – 5'	(T5)
75mer Mismatch 0- 3'–T30 ACA TGT AGT GTT GAT T30 – Cy5 – 5'	(T6)
15mer Mismatch 0-3' – ACA TGT AGT GTT GAT – MR 121–5'	(T7)

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Nucleic Acids and Nucleotides Studied Using Mass Spectrometry

Tracey A. Simmons , ... Patrick A. Limbach , in Encyclopedia of Spectroscopy and Spectrometry, 1999

Nucleic acids and nucleotides

Nucleic acids are high-molecular-mass biopolymers composed of repeating units of nucleotide (nt) residues ( Figure 1). The three major substituents of a nucleotide residue are the heterocyclic base, a sugar and a phosphate group. The five most common heterocyclic bases are adenine (Ade), cytosine (Cyt), guanine (Gua), thymine (Thy) and uracil (Ura). Cytosine, thymine and uracil are classified as pyrimidine bases, and adenine and guanine are classified as purine bases. Adenine, cytosine, guanine and thymine are the major bases found in deoxyribonucleic acids (DNA), and adenine, cytosine, guanine and uracil are the major bases found in ribonucleic acids (RNA). Nucleic acids are identified as either RNA or DNA, depending on the identity of the sugar. The sugar is a 2′-deoxy-d-ribose in DNA and a d-ribose in RNA. The phosphate group is usually attached through the 5′ or 3′ hydroxyl groups of the sugar.

Figure 1. Subunit structures of the major nucleotides from deoxyribonucleic acid (DNA) and ribonucleic acid (RNA). T, C and U nucleobases are pyrimidine derivatives, and A and G nucleobases are purine derivatives.

Mononucleotides are typically represented by a shorthand notation that identifies the sugar, the base and the number of phosphate groups. For example, dNMP represents any 2′-deoxynucleoside monophosphate, dCDP represents 2′-deoxycytidine diphosphate and GTP represents guanosine triphosphate. Unfortunately, this convention is rarely followed in the mass spectrometry literature, where nucleosides and mononucleotides are denoted by their single letter base abbreviation and a preceding or following 'p' to represent the location of the phosphate group on the mononucleotides (for example, dC, dCp, pT and T for 2′-deoxycytidine, 2′-deoxycytidine 3′-monophosphate, thymidine 5′-monophosphate and thymidine respectively). This designation may be a source of confusion for those not familiar with this nomenclature and care should be used whenever such designations are used, especially when one wishes to distinguish nucleobases, nucleosides and nucleotides.

Oligonucleotides are most commonly joined together through the 3′ and 5′ sites of each nucleoside by a phosphodiester linkage. The primary sequence of an oligonucleotide is by definition determined from the 5′ to the 3′ end. Usually the sequence is written in shorthand notation, using the single letter abbreviations for the nucleobases; e.g., 5′-d(pTCAG)-3′ as in Figure 2. In the case where the base composition is known, but the primary sequence is not, the unknown sequence regions are enclosed in parenthesis; e.g. 5′-dACT(GGCT)AAT-3′. Oligonucleotides of a specific length are typically referred to as 'n-mers', where n is the number of oligonucleotide residues.

Figure 2. Shorthand notations of oligonucleotides: (A) line notation of oligonucleotides, and (B) one-letter sequence notation.

Modified nucleosides are designated in the overall sequence by their chemical symbol (e.g., Am for 2′-O-methyladenosine). Unknown modified nucleosides often are designated by an 'X' in the overall sequence. Modified linkages are generally identified by describing the type of modified oligonucleotide followed by the usual sequence notation: a methylphosphonate 10-mer d(ACACGTTGAC) or a phosphorothioate 12-mer d(GCGCATATGCGC). Occasionally, phosphorothiates are identified by an 's' internucleotide linkage, such as d(AsCsTsAsG).

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BIOMOLECULES, BIOINTERFACES, AND APPLICATIONS

Shyamalava Mazumdar , in Handbook of Surfaces and Interfaces of Materials, 2001

9 BINDING OF SURFACTANTS TO NUCLEIC ACIDS

The nucleic acids, such as DNA, RNA, etc., consist of a ribose sugar phosphate backbone, which makes the surface of the molecule highly negatively charged ( Fig. 4). The intrastrand space in a DNA is occupied by base pairs hydrogen-bonded to each other and forms a hydrophobic domain between the base pairs in the intrastrand region. The highly charged outer surface of the nucleic acid generally consists of sodium or magnesium ions, the counter cations. ²³Na-NMR studies [209] on DNA in the presence of a series of cationic surfactants, with quaternary ammonium ions forming the head group, were used to probe the ionic composition in the immediate vicinity of the DNA molecule. Analysis of the line shapes of quadripole split ted ²³Na-NMR lines under slow modulation provided information on the relative fraction of sodium counterions neutralizing the phosphate sites on DNA. Displacement of the sodium ions caused by the surfactants was thus used to determine the relative affinities of various surfactant ions for DNA. The results showed that the binding affinity of the detergent molecules for DNA increases with increasing chain length of the surfactant, indicating that the hydrophobic interactions are important in the binding of surfactants to the nucleic acid. The stability of the DNA double helix is determined from melting of the nucleic acid, monitored by absorption spectroscopy. The role of hydrophobicity and surface charge in the interaction of surfactants with DNA was reported [210] in spectroscopic studies in which the nucleic acid was melted in the presence of 9-(anthrylmethyl) trimethylammonium chloride (ATAC). The calf-thymus DNA, as well as Escherichia coli genomic DNA, was shown to strongly bind ATAC with a binding constant of ∼5 × 10⁴ M⁻¹ (in base molarity). The results showed that addition of cationic surfactants caused structural changes in the ATAC-DNA leading to release of ATAC from the complex. In contrast the anionic surfactants were shown to have no such effect on the ATAC-DNA complex [210].

Agarose gel electrophoresis of the association complexes formed between the cationic surfactants and the plasmid DNA pTZ19R did [210] not show UV luminescence in ethidium bromide staining, presumably because of cationic surfactant-induced condensation of DNA. Inclusion of surfactants in the DNA was shown to generally enhance the DNA melting temperatures of the nucleic acid, and the DNA melting profiles were shown to become broadened in the presence of high concentrations of the surfactant.

Detailed studies on the thermodynamic properties of interaction between DTAB and calf-thymus DNA have been made [211] by various techniques. Potentiometric studies using a DTAB-selective plastic membrane electrode, isothermal titration microcalorimetry, and UV spectrophotometric methods were used to determine the binding isotherms. The results showed that the binding of the cationic surfactant to the nucleic acid takes place initially through 1:1 electrostatic complexation through the phosphate moiety of the DNA. The second DTAB molecule bound to DNA (base molarity) also binds through electrostatic interaction, which was shown to cause a significant conformational change in the DNA backbone. Subsequently, up to 20 DTAB molecules, below the CMC of DTAB, were shown to bind to the nucleic acid through hydrophobic interactions. Studies on the binding of DTAB, CTAB, and myristyltrimethyl ammonium bromide (MTAB) to calf-thymus DNA and its complex with bovine serum albumin (BSA) [212] with the use of equilibrium dialysis also suggested that the chain length of the surfactants plays a significant role in the extent of binding under identical solution conditions.

The cooperative nature of the interaction of cationic surfactant with oligonucleotides was shown to stabilize the duplex DNA structure [213]. Interaction of CTAB with DNA induces condensation and subsequent precipitation of the condensate. Light scattering studies on oligonucleotides GGAAAAAACTTCGTGC and GCACGAAGTTTTTTCC (ODN-1 and ODN-2) in the presence of different mole ratios of CTAB showed [213] an enhanced scattering due to an increase in the size of the complex. At a ratio of CTAB to phosphate (of ODNs) of about 0.75–0.8, the solution was shown to turn cloudy with a decrease in scattering due to the onset of precipitation of the complex. Thermal melting studies of the ODN duplex and the ODN–surfactant complex monitored [213] by absorption at 260 nm showed that the T _m of the ODN duplex was ∼41°C in the absence of the surfactant, and the addition of surfactant initially caused a small increase (1°C) in the T _m of the duplex. At a CTAB concentration of >12 μM, a new melting phase corresponding to the ODN–surfactant complex was observed that showed broad melting curves with T _m > 54°C in the presence of 10 mM NaCl. The results showed that at submicellar concentrations the association of CTAB with the oligonucleotides was initiated by electrostatic interaction of the cationic head group of the surfactant with the DNA that aligns the surfactant molecules on the DNA. This is followed by binding of more surfactants to the initial complex, driven cooperatively by hydrophobic forces, leading to the formation of surfactant-bound DNA duplexes. Detailed thermodynamic aspects of the binding of cationic surfactants to DNA was investigated [214], and selective stabilization of triplex DNA by poly(ethylene glycols) was also reported [215]. These surfactant–DNA complexes showed higher melting temperatures, indicating the higher stability conferred on the duplex structure by the surfactants. These studies have important implications for the rational design of DNA-binding drugs and DNA delivery systems.

Electrokinetic studies using microelectrophoresis techniques reported [216] on the design of a model DNA delivery system. The self-assembling lipid–DNA complex of a cationic surfactant, N,N-diolcoyl-N,N-dimethylammonium chloride (DODAC), was shown to be incorporated into liposomes prepared with 1,2-dioleoyl-i-glycero-3-phosphoethanolamine (DOPE) or 1,2-dioleoyl-sn-glycero-3-phosphocholine (DOPC). The affinity of DNA molecules for biological membranes was shown to be enhanced upon complexation with hydrophobic polycations [217]. The interaction of negatively charged phosphatidylcholine–cardiolipin liposomes with water-soluble negatively charged DNA–cetylpyridinium bromide and DNA–poly(N-alkyl-4-vinylpyridinium bromide) complexes was investigated. The results showed [217] that the DNA–cetylpyridinium bromide complex, while interacting with the liposomes, is destroyed to incorporate the cationic surfactant into the liposomal membrane, leaving the free DNA in the solution. The DNA–poly-(N-ethyl-4-vinylpyridinium bromide) complex, on the other hand, did not interact at all with the liposomes. A complex of DNA with the poly(vinylpyridinium) cation carrying a small amount of N-cetyl groups was shown to be adsorbed on the membrane as a whole, indicating that complexation of DNA with hydrophobized polycations can be used the enhance DNA affinity for biological membranes [217].

Investigations of the factors affecting transport of plasmid DNA into cells by cationic synthetic amphiphiles have also been conducted [218] with the aim of obtaining insight into the mechanism of DNA translocation. The interaction of micelles composed of the cationic amphiphile dioleyloxypropy ltrimethylammonium chloride (DOTMA) with cultured cells was investigated. The results showed that optimal transfection depends on the concentration of the amphiphile, which determines the efficiency of the penetration of the target cell membrane, as well as the toxicity of the amphiphiles toward the cell. A low surfactant to DNA ratio was shown [218] to prevent the complex from interacting with the cell surface, whereas at a relatively high amphiphile concentration the complex can become toxic to the cell. A mechanism of DNA entry involving translocation of the nucleic acids through pores across the membranes rather than delivery via fusion or endocytosis was proposed from these studies. The results also suggested that the phospholipid, dioleoylphosphatidylethanolamine (DOPE) strongly facilitates this pore formation by DOTMA ('lipofectin'). Plasmid DNA has also been reported [219, 220] to be entrapped in 'stabilised plasmid-lipid particles' (SPLP) containing fusogenic lipid dioleoylphosphatidylethanolamine (DOPE), low levels (5–10 mol%) of cationic lipid, dioleoydimethylammonium chloride (DODAC) and stabilized by a polyethyleneglycol (PEG) coating. Translocation of the DNA was also found to be effectively prevented [219] when the cells were pretreated with Ca²⁺ or pronase, which suggested that Ca²⁺-sensitive cell surface proteins may play a role in the amphiphile-mediated DNA translocation process. These studies showed that genetic materials such as DNA or plasmids can be incorporated into living cells using a suitable cataionic surfactant in conjunction with a phospoholipid, by forming a membrane-penetrating complex which do not affect the membrane structure of the cell. These results are important for the design of delivery system for the genetic materials into living cells for genetic engineering as well as for therapeutic purposes.

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Nucleic Acid Synthesis

Sankar Mitra , ... Tadahide Izumi , in Encyclopedia of Physical Science and Technology (Third Edition), 2003

II.A Similarity of DNA and RNA Synthesis

All nucleic acids are usually synthesized by DNA template-guided polymerization of nucleotides—ribonucleotides for RNA and deoxy(ribo)nucleotides for DNA. The reactant monomers are 5′ ribonucleoside (or deoxyribonucleoside) triphosphates. These can be described in the following chemical equations:

$\text{DNA}+\text{ndNTP}\rightleftarrows \text{DNA}+{\text{nPP}}_{\text{i}}$

and

$\left(\text{DNA}\right)+\text{nNTP}\rightleftarrows \left(\text{DNA}\right)+\text{RNA}+{\text{nPP}}_{\text{i}}.$

Enzymatic polymerization is carried out by DNA and RNA polymerases, both of which carry out pyrophosphorolysis, i.e., cleavage of a high energy pyrophosphate bond coupled to esterification of 5′ phosphate linked to the 3′-OH of the previous residue. The reaction is reversible, although it strongly favors synthesis. Degradation of nucleic acids is not due to reversal of the reaction, but rather a hydrolytic reaction catalyzed by nucleases, namely, RNases and DNases, which generate nucleotides or deoxynucleotides, respectively.

Three distinct stages are involved in the biosynthesis of both DNA and RNA: initiation, chain elongation, and termination. Initiation denotes de novo synthesis of a nucleic acid polymer which is generally well regulated by complex processes, as described later. The key difference in initiation of a DNA vs RNA chain is that RNA polymerases can start a new chain, while all DNA polymerases require a "primer," usually a short RNA or DNA sequence with a 3′-OH terminus, to which the first deoxynucleotide residue is added. Elongation denotes continuing polymerization of the monomeric nucleotides, and termination defines stoppage of nucleotide addition to the growing polymer chain.

During synthesis the enzymes catalyzing the polymerization reaction are guided by nucleic acid templates that provide the complementary sequence for the incorporated nucleotides (Fig. 4). The basic catalytic enzyme in such reactions is called DNA or RNA polymerase. In cells the template for both DNA and RNA is genomic DNA. There are some exceptions to these general rules. Some DNA polymerases can synthesize homo- or heteropolymers of deoxynucleotides in vitro in the absence of a template; the substrate is restricted to one or two dNTPs. While it is unlikely that these homo- or heteropolymers, e.g., (dA•dT)_n or poly(dA)_n•poly(dT)_n, are formed in vivo, the availability of these polymers significantly advanced our understanding of the properties of DNA, before the age of chemical or enzymatic oligonucleotide synthesis.

FIGURE 4. Replication of circular DNA of prokaryotes and viruses, plasmids, and mitochondria. The basic steps of replication are shown. (A) Rolling circle mode of replication for single-stranded circular DNA: single-stranded (ss) DNA is replicated to the replicative form (RF), which then acts as the template for progeny ssDNA synthesis via a rolling circle intermediate. (B) Circular duplex DNA can be replicated at the ori site by formation of a θ intermediate. Replication could be bidirectional (as shown here) or unidirectional. 5′   →   3′ chain growth dictates that DNA synthesis is continuous on one side of the ori and discontinuous on the other side for each strand; (+) and (−) strands are shown to distinguish the strand types. (C) Replication of a linear genome with multiple origins.

There are some exceptions to the norm of DNA-dependent DNA or RNA synthesis, mostly in lower eukaryotes or viruses (Fig. 5). One example is RNA-dependent RNA synthesis in plant, animal, or bacterial viruses. In these cases, a single-stranded RNA template rather than double-stranded DNA guides synthesis of the complementary RNA strand, based on conventional base pairing. The polarity of RNA adds a level of complexity during synthesis. Thus, the RNA genome of a virus that can be directly read and thus provides the mRNA function is called the positive strand, as in polio virus. In this case, the viral genome RNA functions as the mRNA and encodes the RNA polymerase, which is synthesized like other viral proteins in the infected cell. This RNA polymerase subsequently synthesizes the complementary negative strand, which then serves as the template for synthesis of the progeny positive strand RNA. The progeny RNA is then packaged into mature progeny virus.

FIGURE 5. Replication of mammalian viral RNA genome. The basic steps of replication are shown for (A) a (+) strand genome, which acts as an mRNA for encoding viral proteins; (B) a (−) viral genome cannot encode protein and first has to be replicated by the RNA replicase (•) which is present in the virus particle. Once the complementary (+) strand which serves as mRNA is synthesized, viral-specific proteins are synthesized, including RNA replicase. (C) Replication of (+) stranded retroviral genomes first involves synthesis of the reverse transcriptase which directs synthesis of duplex DNA in two stages from the RNA template. Circularization of the DNA followed by its genomic integration allows synthesis of progeny viral RNA by the host transcription machinery.

In contrast, the genomic RNA of negative strand viruses (e.g., vesicular stomatitis virus) cannot function directly as mRNA and thus cannot guide synthesis of proteins, including the RNA replicase, by itself after the infection of host cells. These viruses carry their own RNA replicase within the virion capsids, which carry out (+) mRNA strand synthesis after infection (Fig. 5).

Retroviruses comprise diverse groups of viruses, including human immunodeficiency virus (HIV), which share a common mechanism of genome replication. The RNA genomes of these viruses encode an RNA-dependent DNA polymerase (reverse transcriptase or RT) which first generates the complementary (c) DNA of the viral genome. RT has also RNaseH (specific nuclease for degrading RNA from RNA–DNA hybrids) and DNA-dependent DNA polymerase activities. After copying the RNA template, the enzyme degrades the RNA and is able to convert the resulting single-stranded cDNA to duplex DNA. This is then integrated into the host cell genome as proviral DNA, from which the progeny viral RNA is eventually transcribed. Thus, the reverse transcriptase is an unusual polymerase because it can utilize both RNA and DNA templates (Fig. 5). There is strong evidence that such reverse transcription was involved in synthesis of "retrotransposons," a special class of mobile genetic elements, during the evolution of mammalian genomes. These mobile genetic elements, also known as transposons, when identified in bacteria and lower eukaryotes, consist of specific DNA sequences which can be relocated randomly in the genome. The transposition is mediated by enzymes called transposase, usually synthesized by a gene in the transposon. During transposition of retransposons, certain mRNAs are reverse transcribed and then integrated into the genome like the proviral sequence. The presence of specific flanking sequences allows these elements to relocate to other sites in the genome.

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Polymer Chain Mechanics

A. Ikai , in The World of Nano-Biomechanics (Second Edition), 2017

5.2 Polymer Chains

Proteins, nucleic acids, and polysaccharides are all polymeric substances and share fundamental properties with synthetic polymers such as polyethylene. In this chapter, some basic ideas that are necessary for understanding nanomechanical research are introduced.

I. Contour length: It is the physical distance along the main chain of a polymer molecule from the i-th to the j-th segment (L _ij ). The total contour length is the length from the first to the last segment (L ₀  = bN) where b is the length of monomeric unit and N is the total number of such unit in a chain.

II. End-to-end distance ( R ): It is the straight line distance between the first and the last segment of a polymer chain. A statistical average of root-mean-squared end-to-end distance is h  =   < R ²>^1/2.

III. Randomly coiled chain: Neighboring segments are connected by universal joints having no preferred restrictions for its rotation around the solid angle of 4π. Depending on the definition of segment, freely jointed chain (FJC) and wormlike chain (WLC) may be distinguished).

IV. Persistence length: It is the contour length from the i-th to the k-th segment where the directional correlation between two segmental vectors is lost.

V. Radius of gyration: It is the root-mean-squared distance weighted by the segmental mass of all the segments from the center of mass of a polymer chain.

VI. Entropic elasticity: It is the springlike behavior of a flexible polymer chain due to its entropic stability at equilibrium expansion. It is manifested when its conformation is disturbed either by forced extension or compression.

A polymer chain is basically a linear collection of n monomers of equal length l. Each monomer has two neighbors, except for those at the ends. A simple mathematical model of such a polymer chain assumes that neighboring monomers are connected with a universal joint, with no restraints on the relative rotational freedom as illustrated in Fig. 5.1.

Figure 5.1. A polymer chain model as a freely jointed chain (FJC). The rigid segment of length b is linearly connected at their ends by a joint (circled) having no preference for its rotation over the entire solid angle of 4π. The dotted line represents the end-to-end distance in this particular case.

This type of molecule has an interesting characteristic called rubberlike or entropic elasticity. Since at each joint the two independently variable angles between the neighboring segments, θ and φ (Fig. 5.2), can assume any value between 0 and π for the former and between 0 and 2π for the latter, the polymer has a large number of possible conformations, each of which is defined by different values of θ and φ. Although there are no preferred values for the two angles, the degeneracy for different values of θ is different and proportional to 2π·sinθdθ, which is largest for θ  = π/2. Thus < θ > =π/2. For larger or smaller values of θ, the degeneracy is less, which means that the conformational entropy of the chain is at its maximum for states with < θ > =π/2. The definition of the conformational entropy is S  = k _B lnZ, where Z is the number of different states with the same value of < θ >. Consequently, conformational states with either larger or smaller < θ > have smaller Z and S; thus these are less stable in terms of Gibbs energy (G  = H  − TS, where H and T are, respectively, enthalpy and temperature in K).

Figure 5.2. Chemical bonds in a polyethylene chain have a fixed bond length b and a bond angle θ. A single bond has the freedom of rotation around its axis, which is characterized by the angle φ.

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Biochemical Applications of Raman Spectroscopy

Peter Hildebrandt , Sophie Lecomte , in Encyclopedia of Spectroscopy and Spectrometry, 1999

Nucleic acids

Raman bands of nucleic acids originate from in-plane vibrations of the nucleic acid bases (adenine, guanine, cytosine, thymine and uracil) and from the furanose-phosphate backbone. In general, Raman spectra of DNA or RNA reveal structural information about base stacking and interbase hydrogen bonding interactions. Whereas base stacking is reflected solely by intensity variations of purine and pyrimidine bands, changes of interbase hydrogen interactions, for instance, due to helix formation or conformational transitions of the helix, are indicated by both intensity changes and frequency shifts. Raman spectroscopy offers the possibility to determine the conformation of the nucleic acid inasmuch as there are marker bands indicative for the A-, B- and Z-forms of DNA. The most characteristic marker bands originate from the backbone since the various helical structures differ with respect to the sugar–phosphate conformation. Based on the comparison of various DNAs of known crystal structure, it was found that the phosphodiester symmetric stretching vibration is at 811±3 cm ⁻¹ in A-DNA whereas it is upshifted to 835± 5 cm⁻¹ in the B-form. Furthermore, some ring vibrations of nucleic acid bases, particularly of guanine and cytosine, also serve to distinguish between the various DNA structures (Table 1). For quantitative analysis of the DNA structure, the relative intensities of these specific marker bands can be used to assess the percentage of A-, B- and Z-conformation using the conformation-insensitive band at 1100 cm⁻¹ (symmetric stretching of the PO₂ group) as a reference.

Table 1. Conformation-sensitive nonresonant Raman bands of nucleic acid

Frequency (cm −1 )	Description
805–815	phosphodiester symmetric stretching: C-3′-endo conformation (A-DNA)
835 (weak)	phosphodiester symmetric stretching: C-2′-endo conformation (B-DNA)
870–880	phosphodiester symmetric stretching: C-DNA
682	guanine ring breathing: C-2′-endo-anti (B-DNA)
665	guanine ring breathing: C-3′ endo-anti (A-DNA)
625	guanine ring breathing: C-3′ endo-syn (Z-DNA)
1260 (weak)	cytosine band: B-DNA
1265 (strong)	cytosine band: Z-DNA
1318 (moderate)	guanine band: B-DNA
1318 (very strong)	cytosine band: Z-DNA
1334 (moderate)	guanine band: B-DNA
1355 (moderate)	guanine band: Z-DNA
1362 (moderate)	guanine band: B-DNA
1418 (weak)	guanine band: Z-DNA
1420 (moderate)	guanine band:B-DNA
1426 (weak)	guanine band: Z-DNA

In general, structural changes of DNA in biological processes such as transcription, replication or DNA packaging, are not global but are restricted only to a few nucleotides along the chain. Such changes can induce local melting of the secondary structure, reorientations and/or disruptions of the base stacking. Melting of RNA and DNA double helices always leads to the disappearance of the 814 cm⁻¹ and 835 cm⁻¹ bands, indicating a decrease in furanose conformational order and an increase of the backbone chain flexibility. Precise thermal melting profiles of both RNA and DNA double helical complexes have been determined based on the intensity variations of these bands. A careful analysis of the Raman spectra of nucleic acids allows the detection of subtle alterations of the helical organization. This has been demonstrated in studies of DNA packaging, which involves formation of DNA–protein complexes with a series of histone and nonhistone proteins. A backbone conformation of the B-type family was consistently observed for the nucleosome DNA complexes irrespective of the histone and nonhistone content. Furthermore, complex formation is reflected by the intensity decrease of the adenine and guanine ring modes at 1490 and 1580 cm⁻¹, respectively. Whereas the intensity attenuation of the 1580 cm⁻¹ band was attributed to the binding of histone proteins in the small grooves of double helical B-form DNA, the spectral changes of the 1490 cm⁻¹ band is related to the binding of nonhistone proteins in the large grooves.

Raman spectroscopy has also been applied to the analysis of the B–Z transformation in synthetic polymers induced by drug binding. For example, it was shown that binding of trans-dichlorodiamine platinium to poly(dG-dC)·poly(dG-dC) appeared to inhibit the right-to-left handed transition whereas the antitumour drug cis-dichlorodiamine platinium was found to reduce the salt requirement for adopting a left-handed modified Z-like conformation and rendered the transition essentially noncooperative.

A Raman spectroscopic method to obtain information about dynamic aspects of nucleic acid secondary structure monitors the deuterium exchange of the C-8 purine hydrogen. The exchange rates can be determined from the time-dependent intensity increase of the bands characteristic for the deuterated bases. As the exchange process depends on the chemical environment and the solvent accessibility for the individual bases, these data allow differentiation of helical structures.

Pyrimidine and purine bases of nucleic acids have electronic absorptions in the range between 200 and 280 nm. Using excitation lines in this spectral region, RR spectra of nucleic acids exclusively display bands of in-plane ring modes of the bases without any interference of bands from the backbone. However, a selective enhancement of modes originating from purine or pyrimidine bases is not possible so that the practical use of UV RR spectroscopy for structural investigations of nucleic acids is limited. On the other hand, RR spectroscopy can be employed to probe drug–DNA interactions if the electronic transition of the drug is shifted towards the near-UV and visible region. In this way, intercalation of adriamycin by DNA was studied using different excitation lines to probe either the DNA Raman spectrum (364 nm) or the RR spectrum of the bound drug (457 nm).

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Biomacromolecular Applications of Circular Dichroism and ORD

Norma J. Greenfield , in Encyclopedia of Spectroscopy and Spectrometry, 1999

Heteropolynucleotides

Circular dichroism spectra of nucleic acids are complex, compared to those of homopolynucleotides because four different bases contribute to the CD spectra, and the spectra are sequence dependent. In addition, nucleic acids display great conformational diversity and may form single-, double- or triple-stranded helices. For example, the spectra of poly(rA), poly(rU), the dimer poly(rA)·poly(rU) and the trimer poly(rU)·poly(rA)·poly(rU) are illustrated in Figure 9. Complex formation results in shifts in wavelength and intensity of the bands, compared to the addition of the spectra of the unmixed components.

Figure 9. The CD spectra of (——) Poly(rA); (·····) Poly(rU); (----) poly(rU)·poly(rA); (–·–·) poly (rU)·poly (rA)·poly (rU) at pH 7 in sodium phosphate buffer. Redrawn with permission from data in Steely T, Gray DM and Ratiff RL (1986) Nucleic Acid Research 14: 10 071–10 090. Copyright 1986 Oxford University Press.

The various oligomerization states of the polynucleotides, moreover, can exist in multiple conformations. For example the A, B and Z forms of double-stranded nucleic acids have distinct CD spectra. Figure 10A illustrates the CD spectra of poly (dAdC)·poly(dGdT) in the A, B and Z conformations and Figure 10B illustrates CD spectra of a double-helical polynucleotide, poly(dGdC)·poly(dGdC) in the B form, the Z form and in a form in which the bases assume the Hoogsteen base-pairing conformation.

Figure 10. (A) The CD spectra of poly (dAdC)·poly(dGdT) in the (·····) A, (—) B and (---) Z conformations. Redrawn from data in Riazance-Lawrence JH and Johnson WC Jr (1992) Biopolymers 32: 271–276 by permission of John Wiley and Sons, Inc. (B) CD spectra of poly(dGdC)·poly(dGdC) in the (—) B-form, (---) Z-form and (·····) in a form in which the bases assume the Hoogsteen base-pairing conformation. Abstracted with permission from data of Seger-Nolten GMJ, Sijtsema NM and Otto C (1997) Biochemistry 36: 13 241–13 247. Copyright 1997 American Chemical Society.

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Biopolymers

E.Ann MacGregor , in Encyclopedia of Physical Science and Technology (Third Edition), 2003

II.C.1 Structure

The monomers from which nucleic acids are made are more complex than those of either proteins or polysaccharides. These are nucleoside triphosphates and they can be considered to consist of three parts ( Fig. 17)—a triphosphate group, a five-carbon sugar in the furanose ring form, and a cyclic base—that is a ring system with basic properties (which can associate with H⁺ in water). Five different bases are commonly found in nucleic acids.

FIGURE 17. Common nucleoside triphosphates from which nucleic acids are made. The numbering system for the carbon atoms of the sugar rings and the initial letters commonly used as abbreviations for the base names are shown.

In DNA the sugar is 2-deoxy-D-ribose. In RNA the sugar is D-ribose. From this difference came the names of the two polymers. The common bases are adenine, guanine, cytosine, thymine, and uracil; adenine and guanine are purines, while the others are pyrimidines. In each type of nucleic acid, DNA or RNA, only four bases occur commonly—in DNA these are guanine, adenine, cytosine, and thymine and in RNA they are guanine, adenine, cytosine, and uracil. Each base becomes attached to a sugar through a β-glycosidic link. Such a combination of sugar plus base is a nucleoside. The triphosphate is linked to carbon 5′ of the sugar; nucleoside phosphates such as shown in Fig. 17 are also known as nucleotides.

During nucleic acid synthesis monomers are joined together with elimination of a pyrophosphate (diphosphate) grouping as shown in Fig. 18a. From two nucleotides, a dinucleotide is formed and the new link, a phosphodiester link, connects carbon 3′ of one nucleotide sugar to carbon 5′ of the next. A third nucleotide can become joined at carbon 3′ of the right-hand sugar ring, and the process can continue to give a chain of several linked nucleotides—an oligonucleotide—or a chain of hundreds or even thousands of covalently bonded nucleotides, a polynucleotide (Fig. 18b). Most DNA molecules consist of two polynucleotide chains, while RNA molecules usually contain only one polynucleotide chain.

FIGURE 18. (a) Formation of a dinucleotide. (b) The chain structure of a polynucleotide or nucleic acid. X   =   H in DNA and OH in RNA; Y is often a triphosphate group.

At one end of a chain the group attached to carbon 5′ of the sugar ring is not involved in a phosphodiester link. This is referred to as the 5′ end of the chain and by convention is written at the left end of the chain. At the other end, the grouping on carbon 3′ of the sugar is not involved in a phosphodiester linkage; this is the 3′ end of the chain and is written on the right (Fig. 18b).

Nucleic acids differ from one another in the lengths of the polynucleotide chains of their molecules and in the sequence of bases along these chains. For any one nucleic acid, however, all the molecules are identical in size and base sequence. Any sequence of bases is possible in a nucleic acid. Abbreviations are often used to represent nucleic acid structure, and the simplest of these involves giving the base sequence of a chain in terms of the initial letters of the base names, starting from the 5′ end of the chain; in this convention sugar and phosphate groups are not mentioned. Thus if, in Fig. 18b, Base 1   =   guanine, Base 2   =   thymine, Base 3   =   adenine, and Base n   =   cytosine, the structure would be represented as GTA … C.

The two polynucleotide chains in DNA molecules are believed to be wound around each other to give a regular secondary structure, a right-handed double helix (Fig. 19). The two chains run in opposite directions with the phosphate and sugar groups on the outside of the helix and the bases in the interior. The proposal for this structure was first made by Watson and Crick and led to their being awarded the Nobel Prize in 1962. Since the bases are flat, they can stack on top of one another in any sequence almost at right angles to the helix axis. The two chains of the helix are held together by hydrophobic associations between the stacked bases and also by specific hydrogen bonding between pairs of bases, where one base of a pair is contributed by each chain. In an unstrained structure there are 10 base pairs per complete turn of the helix. The two chains of the helix are always approximately the same distance apart (the helix has a diameter of 20 Å) and so the base pair must always consist of one double-ring base, a purine (A or G), hydrogen bonded to a single-ring base, a pyrimidine (T or C). This base-pairing is specific; adenine on one chain always bonds to thymine on another, while guanine on one chain bonds to cytosine on the other. The result is that the sequence of bases on one chain determines the sequence on the second chain, if base-pairing is to take place. For example, if the base sequence of a chain segment is … ACTAGTC … then in the second chain T must bond to A, G to C, etc., and the double helix would have the sequence

FIGURE 19. The double helix of DNA.

VII $\begin{array}{c}5\prime \text{end}----\text{ACTAGTC}----3\prime \text{end}\\ 3\prime \text{end}----\text{TGATCAG}----5\prime \text{end}\end{array}$

The sequences of the two chains are said to be complementary to each other. The helices are not completely regular along their length, but small variations in helix architecture occur with variations in base sequence. These relatively minor changes are important for DNA-protein recognition.

The DNA molecules may be several thousand to over a million nucleotides long, and some segments of the base sequence code for protein structure, while others form control elements. Yet other sequences code for the structures of the r-RNA and t-RNA essential for protein synthesis. The long DNA molecules behave as flexible rods that can coil up if long enough, and in some bacteria the two ends of a molecule can join together to give a closed loop. There is some evidence that, for GC sequences, a stretch of left-handed double helix can form. The importance of this secondary structure of DNA, called Z-DNA, in biological systems is not yet clear.

Except for a few viral ribonucleic acids, all RNA molecules are single-stranded. The polynucleotide chains can fold up on themselves, and if base sequences of two stretches of a chain are complementary, a stretch of right-handed double helix, similar to a DNA double helix, can form. In an RNA double helix, there are approximately eleven base pairs per turn of the helix, and uracil, instead of the thymine of DNA, base-pairs with adenine. The single polynucleotide chains are folded to give short stretches of double helix separated by single-stranded nonhelical segments.

The secondary structures (helices) and tertiary structures (overall chain folding) of RNA are not uniform and differ with the type of RNA. The chain folding is often complex, involving short helices, loops, and even 3- or 4-way junctions of single-stranded chain. Messenger RNA molecules carry in their base sequences the information specifying protein amino acid sequence. There is one messenger for each protein, and so m-RNA molecules vary greatly in length and sequence; hence they differ in secondary and tertiary structure. Ribosomal ribonucleic acids form part of the structures known as ribosomes, where protein synthesis takes place in a living cell. In simple bacteria, each ribosome contains three sizes of r-RNA, designated 5   s, 16   s, and 23   s RNA. (The numbers 5   s, 16   s, and 23   s relate to the speed of movement of the RNA molecules through a solution spinning in a high speed centrifuge, and depend on the size of the RNA molecules). These molecules are typically 120, 1500, and 2900 nucleotides long, respectively. The base sequences of many r-RNAs are known, and segments of complementary base sequence have been observed, so that double helix formation is believed to occur within the ribosome. Transfer RNAs, which bring the amino acids to the sites of protein synthesis, are the smallest ribonucleic acids, being on average about 80 nucleotides long. The base sequences of many t-RNAs have been studied and it has been found that they contain many "unusual" bases, (i.e., bases other than A, G, U, and C). For example, thymine, a "normal" constituent of DNA, is found in t-RNA. All the t-RNAs examined so far have segments of complementary base sequence which can make up four short segments of double helix. Each t-RNA is specific for one amino acid, and so many t-RNAs exist, differing in the number and sequence of nucleotides. Despite this, it is believed all t-RNA molecules have approximately the same overall chain folding (i.e., tertiary structure) shown inFig. 20. Unusual base pairs (e.g., G–U pairs) are found in this structure as well as base triplets (i.e., three bases held together by hydrogen bonding).

FIGURE 20. Tertiary structure of a t-RNA. The four short double helical segments are shown shaded. [Adapted with permission from Rich, A., and RajBhandary, U. L. (1976) Annu. Rev. Biochem. 45, 805–860. Copyright 1976 Annual Reviews Inc.]

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Self-Assembly and Nanochemistry

S.O. Kelley , in Comprehensive Nanoscience and Technology, 2011

5.09.4.2 Oligomeric Nucleotides as Semiconductor Nanocrystal Ligands: Roles of Length and Sequence

The investigation of oligomeric nucleic acids as ligands for semiconductor quantum dots has yielded a few surprises that would not have been predicted from the results described above for monomeric nucleotides. A recent study of homopolymeric 8 and 20 nucleotide sequences used as ligands for CdS did not produce the same trends as those conducted with monomers [31]. For example, materials produced with pyrimidine oligomers exhibited emission intensities comparable or higher than those for materials made with purine oligomers. This observation may reflect that oligomeric ligands can leverage additive affinities of weakly bound ligands to provide enough binding energy to facilitate passivation. Decreased stabilities were noted for some pyrimidine-ligand materials in solutions with higher ionic strengths, indicating that a portion of the nanocrystal–DNA interaction may be electrostatic in nature. Nonetheless, it is clear that the role of sequence in dictating the function of an oligonucleotide as a ligand will reflect both the type of functionalities present in an oligomer, and the number of overall units present.

Another very interesting length-related effect detected in studies of CdS nanocrystals liganded by DNA oligomers relates to size control effected by sequence length. In a study of guanine-based oligomers of varying sizes [32], it was observed that the size of the nanocrystal was inversely related to the length of the oligomer used as a ligand. The source of this effect remains unknown, but it is possible that the longer oligomers are better able to control nanocrystal size by providing a complete ligand set, providing a self-contained microenvironment for nanocrystal growth.

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