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A DNA ligase is an enzyme that joins appropriately juxtaposed DNA ends by forming phosphodiester bonds.

A nucleic acid is a polymer built from linked nucleotides. The two principal kinds are DNA and RNA. Each nucleotide contributes a sugar, a nucleobase, and phosphate to the chain; the nucleobase is only the letter-bearing component, not the entire nucleotide. DNA uses deoxyribose, whereas RNA uses ribose. A chain of six nucleotides is therefore not six free nucleobases placed beside one another. See NHGRI: nucleic acids.

The chemical scaffold and the variable lettering do different jobs. Covalent bonds maintain the sugar–phosphate backbone, while the order of attached nucleobases specifies the nucleic acid sequence. A phosphodiester bond links the $3^{\prime }$ position of one sugar to the $5^{\prime }$ position of the next through phosphate. These position numbers refer to carbon chemical atoms in the sugar; they define a strand's direction even when it bends or forms a circle. See Brown, Genomes, 2nd edition: the human genome.

This division is useful: the backbone supplies a connected physical record, and base pairing allows its letters to recognize partners. DNA replication copies that record, and transcription uses a DNA template to make RNA. RNA can also fold into functional shapes through base pairing within one chain; a nucleic acid sequence specifies chemical order, not a requirement for a straight shape. The beauty is that the same kind of polymer can carry a message and form a structure that acts on messages.

The central dogma of molecular biology concerns sequence information: DNA can be copied by DNA replication; transcription makes RNA from DNA; and translation makes a protein from messenger RNA. The familiar $\mathrm{DNA}\to \mathrm{RNA}\to \mathrm{protein}$ diagram is not an assertion that all information transfer is one-way: reverse transcription and RNA replication are compatible with the dogma. Its key exclusion is transfer of a protein's amino-acid sequence back into nucleic-acid sequence by a general templating mechanism.

DNA is a polymer of deoxyribonucleotides with a sugar–phosphate backbone joined by phosphodiester bonds. Its usual DNA double helix contains antiparallel strands with complementary bases, adenine paired with thymine and guanine with cytosine. The base sequence stores inherited information; complementary pairing supports accurate copying and repair. A strand has distinct $5^{\prime }$ and $3^{\prime }$ ends, and ordinary chain synthesis extends the $3^{\prime }$ end. Packaging into chromosomes compacts and regulates access to the genetic material.

RNA uses ribose, with a $2^{\prime }$ hydroxyl group, and uracil in place of thymine. It is usually single-stranded but forms extensive paired secondary and tertiary structures. Messenger RNA conveys coding information, transfer RNA couples codons to amino acids, and ribosomal RNA provides both structural organization and catalysis within the ribosome. Other RNA molecules regulate expression or catalyse reactions. Thus RNA is not merely a passive intermediate.

Proteins are polypeptides of amino acids linked by peptide bonds, read from an amino terminus to a carboxyl terminus. Their primary sequence guides protein folding; backbone hydrogen bonds organize helices and sheets, while hydrophobic, electrostatic and other interactions produce tertiary and sometimes quaternary protein structure. Their functions include catalysis, transport, signalling, mechanical support and molecular movement. Sequence alone does not guarantee function: folding, modifications, partners and cellular conditions matter. During translation, a ribosome reads successive triplet codons in messenger RNA, with charged transfer RNA molecules supplying the corresponding amino acids; initiation and termination define the reading frame.

DNA is a directional polymer with a sugar-phosphate backbone linked by phosphodiester bonds. Its deoxyribose sugars carry adenine, thymine, guanine or cytosine. In the usual cellular duplex, antiparallel strands form a double helix with complementary A–T and G–C pairing, stabilized by base stacking as well as hydrogen bonding. The sequence can vary without changing the basic backbone, allowing a large information capacity. Complementarity provides a template for copying and repair; the two strands are not merely two redundant lists stored without a copying mechanism. Major and minor grooves permit proteins to recognize sequence-dependent chemical patterns. Eukaryotic chromosomes package DNA around histones, while bacterial chromosomes are usually supercoiled and associated with other proteins; not all cellular DNA is nuclear or linear.

RNA uses ribose with a reactive 2-prime hydroxyl and usually uracil in place of thymine. It is often a single chain, but intramolecular complementary segments form stems, loops and elaborate three-dimensional folds. Messenger RNA conveys coding information to ribosomes; transfer RNA combines an anticodon with an amino-acid attachment site; ribosomal RNA has structural and catalytic roles in translation. Regulatory and catalytic RNAs show why it is inadequate to describe RNA solely as a transient DNA copy. RNA's local pairing and folding let sequence encode both information and molecular machinery. DNA's chemical stability and duplex repair make it especially suitable for long-term inheritance.

Replication is highly faithful but imperfect. Heritable sequence changes, recombination and gene duplication create variation, and selection, Genetic drift and population history determine its spread. Thus the same chemistry explains both continuity of lineages and their divergence; mutation is not directed toward what an organism needs. Conserved genes, homologous sequences and the widely shared genetic code support common ancestry. RNA catalysis makes an early RNA-based stage of evolution a plausible hypothesis, not a direct historical observation or a deduction that all modern RNA preceded DNA in a demonstrated sequence. Structure constrains function and evolutionary possibilities without uniquely determining a species' history.^[ref]

PCR denaturation is the high-temperature stage that separates a DNA duplex into single-stranded templates for the next copying step. It disrupts the pairing and stacking arrangement of nucleobases; it does not require cutting the phosphodiester bonds along either strand. The distinction matters: breaking the backbone would damage the ordered nucleotide sequence that is meant to serve as a template.

For an original eight-base pair duplex, complete PCR denaturation exposes two eight-nucleotide strands. It leaves sixteen nucleotides altogether: separation has made the templates accessible without yet making more DNA. PCR extension, rather than heating alone, adds the new material. Already copied duplexes also need to separate at the beginning of subsequent cycles.

Separation is a condition-dependent process. Template composition and the buffer solution affect the required temperature and duration; heating longer is not automatically better because prolonged heating can reduce DNA polymerase activity. Incomplete separation leaves fewer accessible templates even when the PCR primers themselves are well chosen.^[ref]

A PCR thermal cycle separates three jobs. During PCR denaturation, heating separates the paired DNA strands without intentionally cutting their phosphodiester bonds. During PCR annealing, cooling permits base pairing between PCR primers and templates. During PCR extension, the chosen interval supports DNA polymerase synthesis from the paired $3^{\prime }$ ends. Each new nucleotide is supplied as a deoxynucleoside triphosphate; its incorporation lengthens the strand. The DNA polymerase is a catalyst, while these nucleotides are consumed substrates.^[ref]

Taq polymerase withstands repeated high-temperature strand separation better than a heat-sensitive DNA polymerase. Its extension activity still depends on temperature, buffer solution and magnesium ions. Heat resistance does not mean constant activity at every stage: the separation temperature serves the DNA, whereas the extension temperature serves the copying enzyme. As one institutional example, Addgene's Basic PCR Program specifies $94^{\circ }\mathrm{C}$ separation, $55^{\circ }\mathrm{C}$ primer binding and $72^{\circ }\mathrm{C}$ extension. These are example settings, not universal properties of all PCR primers or DNA polymerases.^[ref]

For timing arithmetic alone, those stipulated holds give $20+25+45=90\mathrm{s}$ per PCR thermal cycle, and three cycles give $270\mathrm{s}$. Adding an assumed $90\mathrm{s}$ initial separation and $180\mathrm{s}$ final extension gives $540\mathrm{s}=9\min$ of holds. A real thermal cycler also takes time to move between temperatures. The schedule tells us when conditions favour PCR denaturation, PCR annealing or PCR extension; it does not measure the fraction of DNA copied.

An actual institutional protocol makes the distinction concrete. New England Biolabs specifies mixing templates and PCR primers with its OneTaq master mix in a thin-walled tube; a quick spin collects liquid at the bottom. Its $25\mu \mathrm{L}$ preparation uses $12.5\mu \mathrm{L}$ of the twice-concentrated mix and $0.5\mu \mathrm{L}$ of each $10\mu \mathrm{M}$ PCR primer stock, with water completing the volume. Tubes enter a thermal cycler preheated to $94^{\circ }\mathrm{C}$; mineral oil is specified if there is no heated lid. The program gives thirty cycles of $94^{\circ }\mathrm{C}$ separation, $45$–$68^{\circ }\mathrm{C}$ binding and $68^{\circ }\mathrm{C}$ extension, with extension time scaled to product length. The binding setting must be selected for the PCR primers; the range is not a continuously swept setting.^[ref]

The dilution can be checked independently: each PCR primer has final concentration $10\times 0.5/25=0.2\mu \mathrm{M}$, while the mix is diluted twofold. For an endpoint readout, Addgene specifies examining a product aliquot by gel electrophoresis.^[ref] Pair that size comparison with the existing PCR controls: a known template and a no-template tube test different explanations of success or failure. Those comparisons are an analytical addition, not controls documented in the OneTaq schedule. An expected-sized band supports production of a length-compatible amplicon; it does not establish its nucleotide sequence, ideal PCR efficiency or successful separation during every cycle. Neither protocol supplies a measured temperature or yield curve for the schematic above.

A phosphodiester bond connects two groups through a phosphate diester. In a nucleic acid backbone, the bridging phosphate is connected through oxygen to the $3^{\prime }$ carbon position of one sugar and the $5^{\prime }$ carbon position of another. These covalent bonds connect successive nucleotides along a strand; they are distinct from the hydrogen bonds between paired nucleobases.

A ribozyme is an RNA molecule whose folded structure provides catalysis of a chemical reaction. A protein enzyme builds its active site from an amino acid chain; a ribozyme builds a working active site from RNA, sometimes with metal ions and associated proteins. Base pairing can help recognize a substrate, but recognition alone does not establish catalysis: the RNA must promote chemical conversion. A passive messenger RNA template being read by a ribosome is not thereby a ribozyme. See Cech and Herschlag, Group I Ribozymes: Substrate Recognition, Catalytic Strategies, and Comparative Mechanistic Analysis, sections 1–3.

The surprise is that an information-bearing polymer can also perform chemistry. Examples include the self-splicing group I intron of Tetrahymena thermophila and the catalytic RNA component of bacterial RNase P, discussed below. The ribosome supplies an everyday example: its peptide bond-forming active site is made of ribosomal RNA, although the complete machine also contains proteins. Nissen and colleagues examined structures of a large ribosome subunit with substrate analogues and found RNA contacting those analogues at the catalytic centre. This supports an RNA active site, not a claim that isolated ribosomal RNA can carry out all of biological translation. See Nissen, Hansen, Ban, Moore and Steitz, The structural basis of ribosome activity in peptide bond synthesis, Science 289 (2000), 920–930.

For the Tetrahymena group I intron, RNA splicing removes the intron and joins its flanking exons. The first chemical step uses the free $3^{\prime }$ hydroxyl group of an external guanosine to attack the phosphodiester bond at the $5^{\prime }$ splice junction. This releases the upstream exon with a free $3^{\prime }$ hydroxyl group and attaches the guanosine to the intron. That newly freed hydroxyl group then attacks the phosphodiester bond at the other splice junction, joining the exons and releasing the linear intron. Each step exchanges one backbone connection for another; it is not simply separation of base pairs. See Cech, Zaug and Grabowski, 1981: the phosphoester-transfer mechanism.

The motivating experiment began with an unexpected control. As Cech recounts, Zaug purified a ribosomal RNA precursor from Tetrahymena using a detergent and phenol extraction, then compared test tubes with and without added cell nucleus extract. Incubation in salts and nucleotides released the same amount of small intron RNA in both cases, detected by gel electrophoresis. Repetition made a missing ingredient or accidental omission less plausible. Activity also survived treatments intended to destroy proteins. Nevertheless, absence of added extract did not by itself exclude a tightly attached, resistant protein, or prove that the small RNA had been newly excised rather than released from an already processed complex. See Thomas R. Cech, 1989 Nobel lecture, pp. 655–659: the unexpected control and alternative explanations.

An independent chemical readout addressed the latter problem. Cech mixed precursor RNA with phosphorus-32-labelled guanosine bearing three phosphate groups (GTP), and used gel electrophoresis followed by exposure to X-ray film to locate the label. Label appeared in the excised intron with magnesium present; it was absent when magnesium was omitted or labelled ATP replaced the labelled guanosine reagent. This experiment is described in Cech, 1989 Nobel lecture, pp. 657–658: labelled-guanosine controls. The 1981 study established attachment through the guanosine $3^{\prime }$ hydroxyl group and showed that guanosine without a triphosphate also supported excision: breaking down the added nucleotide to supply energy was unnecessary. See Cech, Zaug and Grabowski, In vitro splicing of the ribosomal RNA precursor of Tetrahymena, Cell 27 (1981), 487–496: primary abstract. Separately, sequencing and terminal analysis found an extra $5^{\prime }$ guanosine absent from the corresponding DNA sequence. See Zaug and Cech, The intervening sequence excised from the ribosomal RNA precursor of Tetrahymena contains a 5-terminal guanosine residue not encoded by the DNA, Nucleic Acids Research 10 (1982), 2823–2838. Together, incorporation and sequence evidence support a new covalent bond, beyond a change in how RNA associates.

Kruger, Grabowski, Zaug, Sands, Gottschling and Cech's 1982 test removed dependence on a precursor purified from Tetrahymena. They cloned the intron and adjacent exons into a DNA template, made precursor RNA with purified bacterial RNA polymerase under conditions suppressing RNA splicing, and removed proteins by detergent–phenol extraction. Denaturing gel electrophoresis separated precursor species. Purified precursor was then incubated with the guanosine reagent, magnesium and salt; matched samples without this incubation supplied a precursor baseline. Their standard assay used $10\mathrm{mM}$ magnesium chloride, $0.1\mathrm{mM}$ of that guanosine reagent, $100\mathrm{mM}$ ammonium sulfate and a buffer solution at pH $7.5$, for $30$ minutes at temperature $37{}^{\circ }\mathrm{C}$, with specified variations in individual experiments. Gel electrophoresis and radioactive-film detection showed excised intron RNA; sequencing verified its $5^{\prime }$ junction. See Kruger et al., Self-Splicing RNA, Cell 31 (1982), 147–157, figures 3–4 and Experimental Procedures.

Excision alone could leave disconnected exons. To test joining, the authors paired labelled RNA with DNA spanning the joined-exon sequence, removed exposed RNA with enzymes, and separated the protected pieces by denaturing gel electrophoresis. Spliced samples yielded protected RNA long enough to span the junction. See Kruger et al., 1982, figure 6: independent evidence for exon ligation. The experimental logic matters: an intron-sized band identifies a candidate product, while sequence and junction assays test its chemical identity.

These controls support sufficiency of the prepared RNA under the assay conditions. They do not show that every RNA self-splices, or that proteins are dispensable inside a biological cell. Many group I introns use proteins to help attain an active fold. See Cech and Herschlag, Group I Ribozymes, section 6: protein facilitation. Nor does a failed assay necessarily identify an inactive nucleotide sequence: unsuitable ions, pH, or temperature can prevent a working active site from forming.

The distinction between self-processing and repeated use became especially clear with bacterial RNase P. In 1983 Guerrier-Takada, Gardiner, Marsh, Pace and Altman separated the RNA and protein components of RNase P from two bacteria. Purified RNA cleaved precursor transfer RNA under conditions containing $60\mathrm{mM}$ magnesium ions, or $10\mathrm{mM}$ magnesium ions plus $1\mathrm{mM}$ spermidine. The protein components alone had no catalytic activity. At $5$–$10\mathrm{mM}$ magnesium without spermidine, both components were needed. Thus the comparison demonstrated an RNA catalyst while also explaining why protein assistance could matter under other conditions. RNase P produces the mature $5^{\prime }$ end of transfer RNA by removing its leader; it acts on a separate substrate and can be used again. See Guerrier-Takada et al., The RNA moiety of ribonuclease P is the catalytic subunit of the enzyme, Cell 35 (1983), 849–857: primary abstract.

Self-splicing demonstrates RNA chemistry, but one precursor completing one rearrangement is not yet evidence for catalytic turnover: repeated conversion of separate substrates by a regenerated catalyst. The self-splicing precursor changes its own connectivity and incorporates external guanosine. In 1986 Zaug and Cech showed that a shortened Tetrahymena intron RNA could cleave and rejoin separate short RNA substrates in a sequence-dependent reaction. This demonstrated an RNA enzyme acting beyond its own self-processing. See Zaug and Cech, The intervening sequence RNA of Tetrahymena is an enzyme, Science 231 (1986), 470–475: primary abstract.

Read the second figure as a counting argument. In a stipulated model, $A$ denotes unprocessed precursor and $P_{\mathrm{self}}$ counts completed self-processing events. Set $A(0)=R_0$, $P_{\mathrm{self}}(0)=0$ and $dA/d\tau =-A$. The exponential function solution gives $P_{\mathrm{self}}/R_0=1-e^{-\tau }$, never exceeding one. Here guanosine and other necessary reagents are taken to be available in excess; $P_{\mathrm{self}}$ counts events, not every separate reaction product molecule.

For a second stipulated model, let an unchanged ribozyme $R$ perform $R+S⟶R+P$ on a separate substrate $S$. Assume the effective law of mass action $dS/dt=-kRS$, no reverse reaction, no ribozyme loss and no inhibited or long-lived bound states. Set $R=R_0$, $S(0)=5R_0$, $P(0)=0$ and $\tau =k{R}_{0}t$. Then $S/R_0=5e^{-\tau }$ and $P/R_0=5(1-e^{-\tau })$, with $S+P=5R_0$. At $\tau =1$, self-processing gives about $0.632$ completed events per initial RNA, whereas the regenerated ribozyme gives about $3.161$ reaction product molecules per initial ribozyme. More than one substrate conversion per initial ribozyme, with appropriate background and identity controls, is evidence for turnover. Product accumulation below that threshold alone cannot distinguish a reusable catalyst from one-shot chemistry. These models illustrate that distinction; actual enzyme kinetics can include folding delays, binding, saturation and product release.

Sanger sequencing determines a DNA nucleotide sequence by extending a common sequencing primer and occasionally incorporating a dideoxynucleoside triphosphate. A normal deoxynucleoside triphosphate already has hydrogen in place of a $2^{\prime }$ hydroxy group; it retains the $3^{\prime }$ hydroxy group. A dideoxynucleoside triphosphate lacks that $3^{\prime }$ hydroxy group too. It can join the growing strand because the preceding nucleotide supplies the reacting $3^{\prime }$ hydroxy group. Once incorporated, its own $3^{\prime }$ end cannot form the next phosphodiester bond. Thus the blocked step is the next addition, not attachment of the terminator itself.^[ref]

In the historical four-reaction format, each tube receives copies of the same template, the same sequencing primer, a DNA polymerase and all four ordinary deoxynucleoside triphosphates. The tubes differ in their terminating substrate: ddATP, ddCTP, ddGTP or ddTTP. In the A reaction, ordinary dATP can allow copying to continue past one A position, whereas incorporation of ddATP stops another copy there. Competition produces a family of endpoints, rather than every strand stopping at the first A. Adjusting substrate proportions makes later endpoints detectable. A lane identifies the terminating nucleobase in the new strand, not the complementary template letter.^[ref]

The readout requires gel electrophoresis capable of separating single-stranded DNA fragments differing by one nucleotide. Historical denaturing polyacrylamide gel electrophoresis suppressed strand folding and resolved these adjacent lengths; radioactive labels were detected by autoradiography. An ordinary agarose gel electrophoresis comparison with a 100-base pair DNA ladder does not supply that resolution. Four separate reaction lanes carry the identities in this format.^[ref]