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Agarose gel electrophoresis uses a porous agarose polymer network. For linear double-stranded DNA within a suitable separation range, shorter fragments generally move farther in a fixed run. Raising the agarose concentration changes the mesh and the range of sizes that can be resolved.^[ref] A compact circular DNA molecule need not migrate like linear DNA of the same length: geometry affects passage through the mesh.^[ref]

A concrete laboratory example is Addgene's documented DNA separation protocol. Its standard 1% gel contains 1 g of agarose in 100 mL of TAE buffer. Dissolved agarose sets in a casting tray around a comb; removing the comb leaves sample wells. The gel is covered with the same buffer solution in a box connected to a voltage source. Loading buffer contains glycerol to increase sample density and a tracking dye to show run progress. A DNA ladder occupies a separate lane beside the samples. With wells near the negative cathode, DNA migrates toward the positive anode. After separation, bound stain makes the DNA visible by fluorescence under a UV transilluminator. The DNA ladder supplies the size reference; the tracking dye monitors migration but does not identify sample DNA.^[ref]

In the manufacturer's controlled comparison, the same DNA ladder runs in 1%, 2%, and 3% agarose gels with the same run time and other stated conditions. The changed gel band spacing shows why a calibration belongs to its particular gel conditions, rather than establishing a universal distance law.^[ref]

In interpreting an unknown sample, a blank lane tests for detectable background or carryover in that blank, while the DNA ladder checks migration and detection of its known fragments. An empty sample lane cannot by itself distinguish absent DNA from unsuccessful loading or insufficient detection. These controls answer different questions; neither makes matching migration a proof of matching nucleotide sequence.

A DNA ladder contains DNA fragments of known lengths in base pairs. Its gel bands supply size references. Match the sample's strandedness and shape, and use the specified lengths for that DNA ladder. A size reference need not provide a quantitative mass standard.^[ref]

Separation of charged molecules through a gel in an electric field. Denaturing conditions suppress folded structures, helping compare RNA lengths; matching migration alone does not establish an identical nucleotide sequence or identical covalent bonds. In the ribozyme experiments, labelled RNA was detected after separation by exposing photographic film.

In DNA agarose gel electrophoresis, the backbone phosphate groups carry negative electric charge under the usual mildly alkaline buffer solution conditions. DNA therefore moves toward the positive anode, away from the negative cathode.^[ref]

The electric field points the other way: the electric force $\mathbf{F}=q\mathbf{E}$ reverses direction when $q<0$. The apparatus is driven by an external source of electric potential difference; the positive anode convention here belongs to that operating mode. Direction follows electric charge, so it is not a rule that all molecules move toward the same electrode.

The useful result is a spatial comparison: a mixture becomes separated gel bands, which can be compared with a DNA ladder or recovered for subsequent analysis. Agarose gel electrophoresis of linear double-stranded DNA, gel electrophoresis of denatured RNA, and SDS-PAGE of proteins use different conditions to make migration informative about chain size. An electric field supplies motion; the gel supplies a porous obstacle course.

A gel electrophoresis size calibration relates migration distance to known fragment lengths on the same gel. Here is an explicitly chosen model for linear double-stranded DNA, not measured data or a universal law. Let $L$ be the numerical length in base pairs and $d$ the numerical migration distance in millimetres from the wells:The logarithm has a dimensionless argument: $L/100$ is the length divided by 100 base pairs. The chosen DNA ladder fragments $L=100,250,500,1000$ give $d=55.00,50.22,46.61,43.00$, respectively. The gel bands therefore appear in the reverse order of length along the direction of migration.

To invert the calibration, rearrange the logarithm:Our chosen unknown $L=375$ gives $d=48.1116248\dots$, between the 250-base pair and 500-base pair references. Inserting that distance into the inverse recovers 375; retaining only $d=48.11$ gives about $375.12$ base pairs. The extra digits reflect rounding in this model, not the resolving power of a real gel.

For linear interpolation between those two reference gel bands, use the logarithm of length. Here the fraction of the distance from the 500-base pair reference toward the 250-base pair reference is $t={\log }_{10}(500/375)/{\log }_{10}2\approx 0.4150$, giving $L=500(250/500{)}^t=375$. Direct linear interpolation of lengths instead gives approximately $396.24$ base pairs. This difference explains why the choice of calibration coordinates matters. A real calibration must be checked over its usable range; folded RNA or circular DNA does not automatically share the linear double-stranded DNA calibration.

Plasmid restriction mapping compares fragment lengths after a restriction digest with the pattern predicted by specified cut positions. Use the original 6000-base pair plasmid feature map. Abstract restriction enzyme A cuts once, at 1000. Abstract restriction enzyme B cuts twice, at 2500 and 4500. Assume complete cutting of identical circular copies at exactly those sites, with no loss of DNA. These assumptions define a length-accounting exercise; the labels A and B do not stand for particular commercial enzymes.

For the combined restriction digest, successive clockwise distances areand the final fragment crosses the chosen zero:Their sum is $1500+2000+2500=6000$. The single A cut opens the circle into one full-length linear fragment. It does not produce pieces of lengths 1000 and 5000, because coordinate zero was never an end.

For contrast, stipulate a different starting substrate: a linear 6000-base pair DNA fragment with physical ends at 0 and 6000 and the same three internal cut coordinates. Its complete restriction digest givesbase pairs, summing again to 6000. There are four fragments, including two of equal length. Even an ideal length-resolving comparison would show only three distinct sizes if both 1500-base pair fragments occupy the same gel band. Fragment number and visible gel band number need not agree.

The general counting argument is short. For $k\ge 1$ distinct complete cuts of a circle, each fragment runs from one cut to the next, so there are $k$ fragments. For a linear substrate with $k$ distinct internal cuts, its two pre-existing ends give $k+1$ intervals. Summing successive distances counts every original base pair once in either case. The rule assumes internal double-strand cuts and does not treat an uncut circle as a zero-length product.

In practice, compare released linear fragments with an appropriate DNA ladder using agarose gel electrophoresis. The original uncut circular DNA need not migrate like a linear fragment of equal length; the existing agarose gel electrophoresis discussion explains that shape-dependent limitation. Partial cutting also changes the expected pattern. In this example, a circular copy cut at 1000 and 2500 but not 4500 yields lengths 1500 and 4500. A 4500-base pair product would therefore be compatible with that missed cut rather than uniquely identifying a different construction.

Matching lengths constrain an arrangement without determining every nucleotide sequence. For example, any change of letters preserving all three cut positions leaves this predicted pattern unchanged. Even the order of fragments can be ambiguous when only their unordered lengths are observed. DNA sequencing across both joins and through the plasmid insert tests additional claims that this length model cannot decide.

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]