Agarose Gel Electrophoresis Protocol

Agarose gel electrophoresis separates DNA (or RNA) fragments by size using an electric field and a porous gel matrix. The negatively charged phosphate backbone of nucleic acids drives them toward the positive electrode, and smaller fragments slip through the gel’s pores faster than larger ones, spreading them into distinct bands you can see and measure.¹ The protocol itself is one of the most routine procedures in molecular biology, but the details you choose at each step, from agarose percentage to voltage to staining method, determine whether you end up with crisp, interpretable bands or a smeared mess.

Why Fragments Separate by Size

DNA has a nearly uniform ratio of mass to charge. Every nucleotide adds roughly the same amount of negative charge relative to its mass, so the electric field pulls on a 500-base-pair fragment and a 5,000-base-pair fragment with proportional force. What differs is how easily each fragment threads through the agarose mesh. Shorter fragments navigate the pores with less resistance and travel farther in a given time, while longer fragments get hung up more often and lag behind. The distance a fragment migrates ends up being inversely proportional to the log of its molecular weight, which is why DNA ladders (size standards) produce evenly spaced bands on a log scale rather than a linear one.²

Picking the Right Agarose Concentration

The percentage of agarose you dissolve into your buffer controls the pore size of the gel and therefore the range of fragment sizes you can resolve well. A 0.7–0.8% gel has large pores and works best for fragments in the range of roughly 5,000 to 10,000 base pairs and above. A 2% gel has much smaller pores and is better suited for separating fragments in the few-hundred-base-pair range. Most routine work lands somewhere around 1–1.5%, which gives decent resolution for fragments between a few hundred and a few thousand base pairs.

Agarose itself is not all the same. Different grades carry different amounts of residual charged groups, a property measured as electroendosmosis. High-electroendosmosis agarose creates a counter-flow of buffer ions that slows DNA migration unevenly and can smear bands. Low-electroendosmosis agarose minimizes this problem, and the difference becomes especially pronounced if you are using a low-ionic-strength buffer.³ For most standard protocols, molecular-biology-grade agarose is low-electroendosmosis by default, but if you are shopping for agarose and see the specification listed, aim low.

Buffer Choice and What It Actually Changes

The two buffers you will see in nearly every protocol are TAE (Tris-acetate-EDTA) and TBE (Tris-borate-EDTA). Both maintain the pH needed for DNA to stay charged and migrate properly, but they are not interchangeable in every situation. TBE gives slightly better resolution of smaller fragments, while TAE tends to resolve larger fragments a bit more cleanly.⁴ For fragments in the middle range of a few hundred to a few thousand base pairs, the two perform similarly and the choice comes down to what else you plan to do with the DNA.

If you intend to cut a band out of the gel and use the DNA for cloning or ligation, TAE is the standard pick. Borate ions from TBE can inhibit certain enzymatic reactions downstream. If you just need to check that a PCR worked and have no plans to recover the DNA, TBE is perfectly fine and arguably gives slightly sharper bands at the small end. Another practical consideration is heat: running gels at higher voltages generates heat, and both buffers can lose their buffering capacity if the run goes long. TBE has a somewhat higher buffering capacity, so it tolerates longer or faster runs better than TAE before the pH drifts.⁵

Casting the Gel

The practical steps are straightforward: weigh out the agarose, add it to a measured volume of buffer in a flask, and heat the mixture (usually by microwaving) until the agarose is fully dissolved and the solution is clear. Swirl the flask a few times during heating to avoid boil-overs. Let the liquid cool until you can comfortably hold the flask, typically around 55–60 °C, then pour it into the casting tray with the comb already in place. Bubbles trapped in the gel create distortions in the lanes, so if you see any, nudge them to the edge with a pipette tip before the gel sets.

Once the gel has solidified, which usually takes 20 to 30 minutes at room temperature, pull the comb out gently and place the gel in the electrophoresis tank. Fill the tank with the same buffer you used to make the gel until the gel is just submerged. Using a different buffer in the tank than in the gel creates ion mismatches that distort migration, so keep them matched.

Preparing and Loading Samples

Before loading, your DNA samples get mixed with a loading dye. The dye does two things. First, it contains a dense agent like glycerol, sucrose, or Ficoll that makes the sample sink to the bottom of the well instead of floating away into the buffer. Second, it includes colored tracking dyes, commonly bromophenol blue and xylene cyanol, that migrate at known rates through the gel so you can monitor how far the run has progressed without stopping it.⁶ Bromophenol blue runs roughly with 300-base-pair fragments in a 1% gel, while xylene cyanol co-migrates with fragments closer to 4,000 base pairs, so together they bracket a useful range.

Some loading dyes also contain a small amount of SDS (sodium dodecyl sulfate), which halts any enzymatic activity remaining in the sample and can help tighten bands by denaturing proteins that might otherwise interact with the DNA. Whether your loading dye includes SDS depends on the brand; check the label if it matters for your downstream application.⁷

A DNA ladder goes into at least one lane, ideally at each end of the gel if you are running many samples. Pipette slowly when loading to avoid puncturing the bottom of the well or letting sample spill into adjacent lanes. If your wells are small and your sample volume is large, it helps to pre-spin the sample briefly to concentrate it and reduce the volume you need to load.

Running Conditions and Voltage

Voltage is one of the places where people most often go wrong. Running a gel too fast might seem like a time-saver, but higher voltage generates more heat, which can melt the agarose and distort lanes, especially in low-percentage gels. The standard recommendation is around 5 volts per centimeter of distance between the electrodes. For a typical mini-gel apparatus with electrodes about 15 cm apart, that works out to roughly 75 volts.

The relationship between voltage and DNA mobility is not perfectly simple, either. For fragments under about 1,000 base pairs in gels of 1.4% agarose or less, mobility stays constant across a wide range of field strengths. But larger fragments migrate faster as voltage increases, which means that running a gel hot compresses the spacing between bigger bands and can make it hard to tell, say, a 6 kb fragment from an 8 kb fragment.⁸ If you need to resolve large fragments cleanly, slow down and drop the voltage. At higher agarose concentrations (above 2%), the mobilities of all fragment sizes start to depend on field strength, so the “just crank it up” approach distorts everything.⁹

Run time depends on how far apart the bands you care about need to travel. For a quick check of a PCR product, 20 to 30 minutes may be enough. For resolving a restriction digest with multiple fragments of similar size, you might run for an hour or longer at lower voltage. Watch the tracking dyes: once the leading dye is about three-quarters of the way down the gel, it is usually time to stop.

Staining and Visualization

DNA is invisible in the gel until you stain it. The classic stain is ethidium bromide, an intercalating dye that slips between the base pairs and fluoresces bright orange under ultraviolet light. It is sensitive, cheap, and has been the default for decades. The downside is that ethidium bromide is a known mutagen, and its waste solutions require special handling and decontamination.¹⁰ Many labs have shifted to alternative dyes precisely because of that concern.

SYBR Green I is one of the more popular replacements. It is a cyanine dye that binds nucleic acids through a mechanism that is still not fully characterized but likely does not involve classical intercalation the way ethidium bromide does. Testing in the Ames assay, a standard mutagenicity screen, suggests SYBR Green I does not produce point mutations at high frequency, making it a safer option for routine use.¹¹ SYBR Green I is also more sensitive than ethidium bromide for small quantities of DNA, so if you are trying to detect faint bands, it can outperform the old standby. Other options include SYBR Safe, GelRed, and GelGreen, all marketed as less hazardous alternatives with comparable or better sensitivity.

Ethidium bromide can be added directly to the molten agarose before casting, so the gel is pre-stained and you can check band positions during the run. Most of the newer dyes can also be used this way, though some protocols recommend post-staining by soaking the gel in a dilute dye solution after the run, especially if the dye might alter DNA mobility. Post-staining adds 15 to 30 minutes but avoids any interaction between the dye and migration behavior.

Why UV Light Can Be a Problem

Once stained, gels are typically placed on a UV transilluminator to visualize bands. This works well for imaging, but UV light damages DNA. If you plan to excise a band and use the DNA afterward, even brief UV exposure can introduce enough nicks and crosslinks to reduce cloning efficiency. Replacing the UV transilluminator with a blue-light system avoids this damage. One study found that switching to blue light improved the efficiency of ligation-independent cloning enough that even lower-efficiency bacterial cells could be used without extra steps.¹² Blue-light transilluminators require dyes that fluoresce under blue light, like SYBR Safe, rather than ethidium bromide, which needs UV excitation. So your choice of stain and your choice of transilluminator are linked decisions.

Cutting Bands and Extracting DNA

Gel extraction is the step that turns agarose gel electrophoresis from an analytical tool into a preparative one. You identify the band of interest under illumination, cut it out of the gel with a clean scalpel or razor blade, and dissolve the agarose slice to free the DNA. Commercial gel extraction kits use a silica column to bind the DNA after the agarose is melted in a chaotropic buffer, and the purified DNA is eluted in water or a low-salt buffer.

These kits produce clean DNA suitable for cloning, sequencing, and other enzymatic reactions. Testing of regenerated silica columns from gel extraction kits showed that DNA yield and quality remained consistent even after columns were reused multiple times, with successful cloning and correct sequences confirmed by sequencing.¹³ That said, most labs treat columns as single-use to avoid cross-contamination. If you are working on a tight budget, regeneration is an option that does not appear to compromise results.

The buffer you used matters here, too. TAE-based gels are standard for gel extraction because borate ions from TBE co-purify with the DNA and can inhibit downstream enzymes. If you made your gel in TBE and need to extract a band, expect lower yields or interference in ligation reactions.

Adapting the Protocol for RNA

RNA molecules form extensive secondary structures through internal base pairing, which means their migration in a standard native gel does not reliably reflect their true size. A 2 kb RNA that folds into a compact hairpin will run faster than its length would predict. To get accurate size estimates, you need to denature the RNA before and during electrophoresis.

The two common approaches are formaldehyde-denaturing gels and glyoxal-based denaturation. Formaldehyde gels incorporate formaldehyde directly into the agarose and running buffer, maintaining denaturing conditions throughout the run. Glyoxal-based methods treat the RNA with glyoxal and dimethyl sulfoxide before loading, then run the treated sample on a standard agarose gel in a phosphate buffer.¹⁴ Both approaches prevent secondary structures from reforming and allow RNA to migrate according to its actual length.

Visualization of RNA in denaturing gels can use ethidium bromide or SYBR-type dyes, but silver staining is an alternative that provides sharp, well-resolved bands and accurate size estimates without requiring UV excitation at all.¹⁵ Silver staining is more sensitive for low-abundance RNA species and avoids the mutagenicity concerns of ethidium bromide, though the staining protocol itself is more involved and takes longer.

When Standard Gels Cannot Resolve Your Fragments

Standard agarose gel electrophoresis works well for fragments up to roughly 20 or 25 kilobases. Beyond that, the pore size of the gel is simply not large enough to separate very long DNA molecules, and everything above a certain threshold migrates as a single unresolved mass near the top of the gel. This is where pulsed-field gel electrophoresis comes in.

In pulsed-field gel electrophoresis, the electric field alternates between different pairs of electrodes arranged at angles to each other. Each time the field switches direction, the DNA molecules have to reorient before they can start migrating again, and the time needed for reorientation depends on the molecule’s size. Smaller molecules reorient and resume moving faster than larger ones. By tuning the pulse times, you can separate fragments ranging from tens of kilobases up to entire megabase-scale chromosomes.¹⁶ Pulsed-field runs are long, often 12 to 24 hours or more, and the apparatus is more specialized, but for applications like bacterial strain typing or resolving yeast chromosomes, it is the standard tool.

Handling Ethidium Bromide Waste

If your lab still uses ethidium bromide, disposing of it properly matters. Ethidium bromide solutions are toxic and mutagenic, and dumping them down the drain is both a safety violation and an environmental hazard.¹⁷ Most institutions require that ethidium bromide waste be collected separately and either chemically decontaminated or sent out for hazardous waste disposal. Common decontamination methods include charcoal filtration, bleach treatment, or commercial destaining bags that adsorb the dye from solution. Gels containing ethidium bromide are generally treated as solid hazardous waste and bagged separately from regular lab trash.

Switching to SYBR-type dyes eliminates much of this burden. The newer dyes can typically be disposed of through normal chemical waste channels or even down the drain in many jurisdictions, depending on local regulations. Given that SYBR dyes also tend to be more sensitive, the main reason labs still cling to ethidium bromide is inertia and familiarity, not performance.

Capillary Electrophoresis as an Alternative

Automated capillary electrophoresis systems have become common in clinical and high-throughput research labs. Instead of a slab gel, capillary electrophoresis runs samples through a narrow capillary tube filled with a polymer matrix, and a detector reads the separated components in real time. Compared with agarose gel electrophoresis, capillary systems produce narrower, more efficient peaks and can process samples faster with less hands-on time.¹⁸

For protein electrophoresis in clinical diagnostics, such as detecting abnormal immunoglobulin bands, the two methods agree closely on major findings but diverge on subtle ones. Agarose gels tend to offer slightly better resolution in certain protein zones, while capillary electrophoresis picks up a marginally higher number of faint oligoclonal bands.¹⁹ For nucleic acid work, instruments like the Agilent Bioanalyzer and TapeStation use capillary or microfluidic electrophoresis to size and quantify DNA and RNA fragments from very small sample volumes, eliminating the gel-casting and staining steps entirely.

That said, capillary instruments are expensive and overkill for many routine applications. If you are checking a handful of PCR reactions or running a restriction digest, a $10 agarose gel that takes 30 minutes to pour and run is hard to beat on cost and simplicity. Agarose gels also give you the option to recover specific bands by gel extraction, which is not straightforward with capillary systems. The two approaches are complementary rather than competitive: agarose gels for everyday preparative and analytical work, and capillary electrophoresis for high-throughput, quantitative, or low-sample-volume applications.

Common Mistakes and How to Avoid Them

A few issues come up again and again, especially for people new to the technique:

Smiling bands: Bands that curve upward at the edges of the gel typically indicate the gel was run too fast and the center overheated. Lower the voltage or run in a cold room.
Fuzzy or smeared lanes: This can mean degraded DNA, overloaded wells, or that the gel melted during the run. Check your DNA quality on a spectrophotometer first, load less sample, or reduce voltage.
No visible bands: Confirm your stain is working by running a positive control (the ladder should always be visible). If the ladder shows up but your sample does not, the problem is your sample, not your gel.
Bands in the wrong position: If your fragment runs at an unexpected size, double-check that you used the right ladder for your gel percentage. A supercoiled plasmid, for example, migrates much faster than its linearized form and will appear to be a smaller fragment than it actually is.
Uneven wells: If the comb was not perfectly horizontal when the gel set, wells will have different depths and volumes. This produces uneven loading and lanes that are hard to compare. Make sure the casting tray is level before pouring.

Supercoiled DNA deserves a special mention because it trips people up frequently. Uncut plasmid DNA migrates as multiple bands representing the supercoiled, relaxed circular, and linear forms, each of which moves at a different rate through the gel. If you are trying to assess plasmid integrity, you need to recognize these forms rather than assuming multiple bands mean contamination. Linearizing the plasmid with a single-cut restriction enzyme gives you one clean band at the true plasmid size.