Gel Electrophoresis

What gel electrophoresis actually does

Hand someone a tube of DNA and ask "how big are these fragments?" and you have posed a surprisingly hard question. The fragments are invisible, weigh femtograms, and a single microlitre holds billions of them in a tangle of every possible length. Gel electrophoresis answers that question with nothing more than a slab of gelled seaweed extract, a tank of salt water, and a 9-volt-to-150-volt battery's worth of electricity. Within an hour it sorts the tangle into a tidy ladder of bands, each band a population of molecules of one size, ranked from largest at the top to smallest at the bottom.

The technique exploits two physical facts at once. First, DNA is an electrically charged molecule: every nucleotide carries a phosphate group that is negatively charged at normal buffer pH, so an electric field exerts a force on it. Second, the field is applied not in open solution but through a gel — a microscopic three-dimensional mesh that acts as a molecular sieve. The combination of a uniform pulling force and a size-dependent obstacle course is what produces size separation. Take away the gel and every fragment, large or small, would move at nearly the same speed, because in free solution DNA's charge and drag scale together and cancel out. The gel breaks that symmetry.

Why everything runs to the anode

DNA's backbone is a chain of alternating sugars and phosphates. At a typical running pH of around 8 (set by a TAE or TBE buffer), each phosphate has lost its proton and sits as a negatively charged group. There is exactly one phosphate per nucleotide, so a 1,000-base-pair fragment carries about 2,000 negative charges and a 100-base-pair fragment about 200 — the charge-to-mass ratio is constant no matter the sequence or length. That uniformity is the quiet genius of the method: because charge per unit length is fixed, the electric force per unit length is fixed too, so the only thing left to distinguish fragments is how the gel impedes them.

When you switch on the power supply, the electrode where electrons leave the solution becomes positive (the anode) and the other negative (the cathode). Negative charges are pulled toward positive, so DNA marches steadily toward the anode. Bench convention colors the anode red, which gives students the mnemonic "DNA runs to red." A loading dye mixed into the sample carries a visible tracking front — bromophenol blue migrates roughly with a 300-bp fragment, xylene cyanol with about 4 kb — so you can watch the run progress and stop it before your bands run off the bottom edge.

The sieve: how the gel sorts by length

Pour molten agarose, let it cool, and it sets into a gel riddled with water-filled pores a few hundred nanometres across — comparable in scale to the molecules being separated. A short DNA fragment is compact and finds an open path through the mesh almost immediately, so it migrates fast. A long fragment must reptate: it threads through head-first like a snake worming through a chain-link fence, constantly snagging on the polymer strands. The longer the molecule, the more it snags, and the slower it moves.

The result is a clean inverse relationship. Over the useful range of a gel, migration distance is proportional to the logarithm of fragment length (not the length itself), which is why a size ladder's bands crowd together at the bottom and spread out at the top. Below a lower size cutoff every small fragment runs at nearly the same maximum speed and the bands merge; above an upper cutoff the largest fragments barely move and pile up near the well. You tune that resolving window with the gel's percentage: a loose 0.5–0.8% agarose gel resolves large fragments (1–25 kb) by giving them big pores, while a dense 2–3% gel resolves tiny fragments (50–500 bp) by tightening the mesh.

The numbers that govern a run

Electrophoresis is governed by a few interacting knobs. Voltage sets the driving force; higher voltage runs faster but generates more heat, and too much heat warps the gel and smears bands. The practical sweet spot is about 5–10 volts per centimetre of electrode-to-electrode distance — roughly 80–150 V for a standard 10–15 cm tank — for 30 to 90 minutes. The buffer carries the current; without ions in the water the circuit is open and nothing migrates, which is why a gel made with pure water simply does nothing.

The table below compares the two dominant gel systems and the molecules each is built to resolve.

Property	Agarose gel	Polyacrylamide gel (PAGE)
Typical concentration	0.5–3% (w/v)	4–20% (acrylamide)
Pore size	Large, loosely tunable	Small, finely tunable
Resolving range	~100 bp – 25 kb DNA	~5 bp – 1 kb DNA; proteins
Resolution	Tens of base pairs	Single base / single amino acid
Format	Horizontal, poured on bench	Vertical, cast between plates
Main use	Routine DNA & RNA sizing	Sequencing-grade DNA; SDS-PAGE proteins
Hazard	Stain (intercalating dye)	Unpolymerized acrylamide neurotoxin

Making invisible bands visible

DNA is colorless, so after the run you stain it. The classic stain is ethidium bromide, a flat molecule that slides (intercalates) between the stacked base pairs of the double helix; once wedged in, it fluoresces bright orange under ultraviolet light, roughly 20 times brighter than in free solution. Because it binds in proportion to the amount of DNA, band brightness reports quantity as well as position. Ethidium bromide is a suspected mutagen — it intercalates your DNA just as happily as the sample's — so many labs now use safer dyes such as SYBR Safe or GelRed that fluoresce under less hazardous blue light.

To convert band position into an actual size you run a ladder — a commercial mixture of fragments of precisely known lengths (for example, bands at 100, 200, 300… up to 1,000 bp) — in an adjacent lane. Plotting log(size) of the ladder bands against their migration distance gives a near-straight standard curve, and reading an unknown band off that curve typically pins its size to within 5–10%. This is how a single photograph of a glowing gel becomes a quantitative measurement.

Sorting proteins: the SDS trick

Proteins seem like they should defeat the method, because unlike DNA they have wildly different intrinsic charges and fold into compact globular shapes rather than uniform rods. The fix is the detergent sodium dodecyl sulfate (SDS). SDS unfolds each protein into a linear chain and coats it with negative charge in fixed proportion — about one SDS molecule per two amino acid residues — which swamps the protein's own charge and shape. Now every protein behaves like DNA: a uniformly charged rod whose only distinguishing feature is length. A reducing agent such as β-mercaptoethanol snaps disulfide bonds so multi-subunit proteins split into separate chains. The result, SDS-PAGE, separates proteins purely by molecular weight and is the workhorse behind Western blots and protein purity checks.

Why it is in every molecular biology lab

Gel electrophoresis is the universal quality-control step. After a PCR amplification, a quick gel confirms you made a product of the right size and only that product. After cutting DNA with restriction enzymes, the pattern of band sizes tells you the digest worked and which fragments you have. When building a recombinant plasmid, you screen clones by digesting and running them. The technique underlies DNA fingerprinting and paternity testing (everyone's restriction-fragment pattern differs), the read-out of classic Sanger DNA sequencing, and the Southern, Northern, and Western blots that probe specific sequences or proteins after separation.

Its enduring appeal is economic and practical: a single gel costs only cents in reagents, runs in under an hour, needs no expensive instrument beyond a power supply and a UV box, and gives an instantly interpretable visual answer. Newer technologies — capillary electrophoresis, microfluidic "lab-on-a-chip" systems, and next-generation sequencing — automate or miniaturise the same physics, but the slab gel poured on a bench remains the first thing most biologists reach for when they need to know, quickly and cheaply, how big their molecules are.

Where runs go wrong

• Smeared bands. Voltage too high (overheating), gel too old, or degraded sample. Run cooler and slower.
• No bands at all. Forgot the stain, ran the gel backwards (toward the cathode), or used buffer with no ions. DNA runs to red.
• Bands ran off the gel. Run too long, or gel percentage wrong for the size range. Watch the dye front.
• Fuzzy small fragments. Gel percentage too low; switch to a denser gel to resolve short fragments.
• One giant band stuck in the well. Fragments too large for the pore size, or DNA not fully cut.