Agarose gel electrophoresis is the standard lab method for separating DNA or RNA fragments by size, using an electric current to pull nucleic acids through a porous gel matrix. Smaller fragments move faster and travel further than larger ones in a fixed run time, so after staining, the gel shows a ladder of bands that reveals fragment sizes directly. It is one of the most frequently run techniques in a molecular biology lab — used to check a PCR product, verify a restriction digest, confirm plasmid identity, or assess whether extracted DNA or RNA is intact — and it is usually the first checkpoint before a sample moves on to sequencing, cloning, or a more sensitive downstream assay.
How It Works: Separating Nucleic Acids by Size
DNA and RNA both carry a negatively charged phosphate backbone, so when a gel is placed in an electric field, nucleic acid fragments migrate toward the positive electrode (the anode). The agarose itself forms a mesh of pores when it sets, and that mesh acts as a molecular sieve: small fragments thread through the pores relatively easily and move quickly, while large fragments have to work harder to squeeze through and move more slowly. Over a fixed run time, this produces size-based separation — fragments spread out into distinct bands ordered by length, with the smallest fragments furthest from the wells and the largest fragments closest to them.
This is different from separation by charge or shape, which is roughly how proteins would behave without help — nucleic acids don’t need an added coating like SDS the way proteins do in SDS-PAGE, because the phosphate backbone already gives every fragment a charge proportional to its length, independent of base sequence.
When to Use Agarose Gel Electrophoresis (and When Not To)
Agarose gel electrophoresis is the right tool for resolving nucleic acid fragments roughly 100 base pairs to 25 kilobases in size — the range covers most PCR products, restriction digests, and plasmid preparations. For separating very small fragments (under ~100 bp, such as short oligonucleotides or certain forensic/STR fragments) or for resolving fragments that differ by only a few base pairs, polyacrylamide gel electrophoresis (PAGE) has finer resolving power because its pore size is smaller and more tunable. For proteins rather than nucleic acids, SDS-PAGE (not agarose) is the standard method, since the separation mechanism and staining/detection chemistry are different — see CASRAI’s western blot protocol guide for how SDS-PAGE fits into that workflow. Agarose is preferred over PAGE for routine nucleic acid work specifically because it is faster and simpler to cast and run, at the cost of somewhat coarser resolution.
What You Need Before You Start
A standard run requires: agarose powder, a running buffer (most commonly TAE — Tris-acetate-EDTA — or TBE, Tris-borate-EDTA), a casting tray and comb to form sample wells, a horizontal electrophoresis chamber with a power supply, a nucleic acid stain (see the staining section below for the ethidium bromide vs. safer-alternative choice), loading dye to track migration and weigh down the sample in the well, a DNA or RNA size ladder run alongside samples as a reference, and a gel imaging system (UV or blue-light transilluminator with a camera, or a dedicated gel-doc imager). TAE and TBE are not interchangeable mid-protocol: TBE has higher buffering capacity and is generally preferred for longer runs or when fragments need sharper resolution, while TAE is more common for everyday work and for fragments that will be excised from the gel afterward (TBE’s borate can interfere with some downstream enzymatic steps). For calculating buffer molarity from a stock concentrate or a solid reagent, see CASRAI’s guide to molarity and solution calculations for the lab.
Step 1: Choosing the Right Agarose Percentage
Agarose percentage controls pore size, and pore size determines which fragment sizes the gel resolves well. As a general guide:
- 0.7% agarose resolves large fragments, roughly 0.8–12 kb — useful for large plasmids or genomic DNA fragments.
- 1.0% agarose is the standard, general-purpose percentage, resolving roughly 0.5–10 kb — the default choice for most PCR product and restriction digest checks.
- 2.0% agarose resolves small fragments well, roughly 0.1–3 kb — useful for small PCR products or distinguishing fragments that are close in size.
Higher percentages give finer resolution of small fragments but run more slowly and produce a gel that is more difficult to handle without tearing; lower percentages run faster and handle large fragments better but blur the distinction between similarly sized bands. When in doubt, 1% is a reasonable default for a first look at an unfamiliar sample.
Step 2: Casting the Gel
Weigh out agarose powder and dissolve it in the chosen running buffer at the target percentage (for example, 1 g of agarose in 100 mL of buffer for a 1% gel), then heat the mixture — typically in a microwave, swirling at intervals — until the agarose fully dissolves and the solution turns clear. Let the molten agarose cool to roughly 50–60°C (cool enough not to warp a plastic casting tray or melt a comb, but well before it starts to set) before adding stain, if the stain is being cast directly into the gel rather than applied after the run. Pour the gel into a casting tray with a comb positioned to form sample wells, and let it solidify fully at room temperature, typically 20–30 minutes, before removing the comb carefully to avoid tearing the wells.
Step 3: Preparing and Loading Samples
Mix each sample with loading dye before loading — loading dye serves two purposes: it adds density so the sample sinks into the well instead of diffusing into the buffer, and it contains a visible tracking dye that migrates through the gel at a roughly predictable rate, giving a rough visual cue for how far the run has progressed without needing to stop and check under UV or blue light repeatedly. Load a DNA or RNA ladder into at least one well on the gel — without a ladder run alongside the samples, there is no way to estimate fragment size from the finished gel, only relative comparisons between sample lanes. Load samples carefully and consistently; uneven loading volumes across lanes make band-intensity comparisons between samples unreliable.
Step 4: Running the Gel
Submerge the cast gel in an electrophoresis chamber filled with the same running buffer used to cast it — mixing TAE-cast gels with TBE running buffer, or vice versa, changes the effective ionic strength and produces inconsistent migration. Apply a constant voltage, commonly in the range of about 5–10 volts per centimeter of gel length, and run until the tracking dye has migrated an appropriate distance for the fragment sizes being resolved — this is typically 30 minutes to a few hours depending on gel percentage, voltage, and fragment size. Running too fast (high voltage) generates heat that can distort bands, especially on longer runs; running too slow wastes time without improving resolution meaningfully once the dye has migrated far enough.
Step 5: Staining and Visualization
Nucleic acids in an unstained gel are invisible to the eye, so a fluorescent dye that intercalates into (or otherwise binds) DNA or RNA is needed to visualize bands under UV or blue light. Ethidium bromide has historically been the most common stain because it is inexpensive and highly sensitive, but it is a mutagen and requires careful handling and disposal as hazardous waste under most institutional environmental health and safety programs. Safer alternatives — such as SYBR Safe, GelRed, and similar dyes — are now widely used in place of ethidium bromide because they are designed to be less mutagenic and, in some cases, compatible with standard (non-hazardous) waste disposal, though institutional EHS policy should always be confirmed rather than assumed, since disposal rules vary by dye, concentration, and institution. Stain can be added directly into the molten agarose before casting, into the running buffer, or applied to the gel as a post-run soak — each has trade-offs in sensitivity, background, and how much extra handling time it adds.
Step 6: Reading and Interpreting Results
Image the gel under UV or blue-light transillumination and compare each sample band’s migration distance to the ladder to estimate fragment size — the closer a sample band lines up with a specific ladder band, the closer its size is to that reference. A single sharp band at the expected size is the typical sign of a clean, successful reaction (a specific PCR product, a completed restriction digest). Multiple bands can mean multiple real products (partial digestion, alternative splice products, non-specific PCR amplification) or contamination, and distinguishing between those explanations usually requires additional evidence beyond the gel itself, such as sequencing or a repeat reaction with adjusted conditions. A diffuse smear rather than a distinct band is a common sign of degraded or sheared nucleic acid, especially for RNA, which is far more prone to degradation than DNA due to the ubiquity of RNases; a smear in an RNA gel is often the first indicator that a sample needs to be re-extracted rather than run forward into a downstream application.
Troubleshooting Common Problems
- No bands at all: check that the gel was oriented correctly in the buffer chamber (DNA migrates toward the positive electrode — a gel run backward shows nothing in the expected lanes), that stain was actually present in or applied to the gel, and that the sample itself contains enough nucleic acid to detect.
- Smeared bands: most often degraded sample (especially likely for RNA), overloaded wells, or a gel run at too high a voltage for too long, generating heat that distorts the matrix.
- Bands that don’t line up evenly (a “smiling” gel): usually uneven heat distribution during the run, often from running at too high a voltage or an inconsistent buffer level across the gel.
- Faint or missing ladder: check ladder concentration and loading volume against the manufacturer’s recommendation, and confirm the stain is actually reaching the ladder lane if stain was applied post-run rather than cast into the gel.
- Bands at an unexpected size: can indicate non-specific PCR amplification, incomplete restriction digestion, primer-dimer formation (very small unexpected bands), or genuine biological variation (an unexpected splice variant or insertion/deletion) — treat it as a real result to investigate rather than an error to explain away.
Agarose Gel Electrophoresis vs. Related Techniques
Agarose gel electrophoresis is sometimes confused with related separation methods that serve different purposes. Polyacrylamide gel electrophoresis (PAGE) uses a synthetic polymer matrix instead of agarose and offers finer resolution, making it the better choice for very small nucleic acid fragments or for protein separation (as SDS-PAGE, the first stage of a western blot). Capillary electrophoresis runs the separation inside a thin capillary tube rather than a flat gel slab, giving higher resolution and enabling automated, quantitative sizing (used in DNA sequencing and fragment-length analysis such as microsatellite genotyping), at the cost of specialized instrumentation that a standard gel rig doesn’t require. Choosing among these comes down to what the experiment needs: agarose for routine, fast, low-cost checks of nucleic acid fragments in the common size range; PAGE for finer resolution or protein separation; capillary electrophoresis for automated, high-precision sizing.
Frequently Asked Questions
What is agarose gel electrophoresis?
It is a laboratory technique that separates DNA or RNA fragments by size by pulling them through a porous agarose gel matrix using an electric current — smaller fragments move faster and travel further than larger fragments in a given run time, producing a pattern of bands that reveals fragment sizes when compared against a size ladder.
Why does DNA migrate toward the positive electrode?
DNA’s phosphate backbone carries a negative charge along its entire length, so in an electric field the molecule is pulled toward the positive electrode (the anode). This charge is roughly proportional to fragment length, which is part of why migration through the gel’s pores — not raw charge alone — ends up being the dominant factor in size-based separation.
What’s the difference between TAE and TBE buffer?
Both are Tris-based buffers used to run and sometimes cast agarose gels. TBE (Tris-borate-EDTA) has higher buffering capacity, making it better suited to longer runs and sharper resolution, while TAE (Tris-acetate-EDTA) is more common for everyday, shorter runs and for samples that will be excised from the gel afterward, since TBE’s borate can interfere with some downstream enzymatic reactions.
How long does an agarose gel run take?
Typically 30 minutes to a couple of hours, depending on gel percentage, applied voltage, and how far the fragments need to migrate to separate clearly — a quick check of a PCR product on a 1% gel at a moderate voltage often takes 30–45 minutes, while resolving closely sized fragments on a higher-percentage gel can take longer.
Why is my DNA or RNA smearing instead of forming a clean band?
A smear instead of a sharp band most often indicates degraded or sheared nucleic acid — RNA is especially prone to this because RNases are extremely common and hard to fully eliminate from a lab environment. Overloading the well or running the gel too hot (too high a voltage for too long) can also produce smearing even with intact sample.
Is ethidium bromide still used for staining agarose gels?
It is still used in some labs because it is inexpensive and sensitive, but many labs have shifted to alternatives such as SYBR Safe or GelRed, which are designed to be less mutagenic and, depending on institutional policy, may allow simpler waste disposal. Whichever stain is used, always follow the specific institutional EHS guidance for handling and disposal rather than assuming a given dye is exempt from hazardous-waste rules.
For the concentration and dilution math behind preparing running buffer or diluting a stock reagent to a working molarity, see CASRAI’s guide to molarity and solution calculations for the lab, and for the serial-dilution technique often used to prepare a dilution series of a sample or standard, see CASRAI’s serial dilution guide.
