Gel electrophoresis separates DNA fragments by size using an electric field and a porous agarose gel. The process involves dissolving agarose powder in buffer, pouring it into a casting tray, loading your DNA samples, and running current through the gel. The whole procedure takes about 1.5 to 2 hours from start to finish, and the materials are straightforward once you know what concentrations to use.
How the Gel Actually Works
Agarose forms a mesh of tiny pores when it solidifies. DNA carries a negative charge, so when you apply an electric field, every fragment migrates toward the positive electrode. Smaller fragments slip through the pores faster, while larger ones get slowed down. This size-dependent sieving is what produces distinct bands on your gel.
For very large DNA molecules, something interesting happens: the strand is too big to fit through pores normally, so it threads through the gel matrix in a snake-like motion, sliding along its own length. This behavior is why extremely large fragments (above 15-20 kb) become harder to resolve and require lower voltages or specialized techniques like pulsed-field electrophoresis.
Choosing Your Agarose Concentration
The concentration of agarose determines the pore size, which in turn determines which DNA fragments you can separate cleanly. Most gels fall between 0.5% and 2%. Standard agarose gels work best for fragments between 100 base pairs (bp) and 25 kb. Anything smaller than 100 bp is better resolved with polyacrylamide gels instead.
As a general guide:
- 0.5% gel: best for large fragments, roughly 5-25 kb
- 1.0% gel: a good all-purpose choice for fragments in the 500 bp to 10 kb range
- 1.5% gel: resolves smaller fragments well, around 200 bp to 3 kb
- 2.0% gel: best for distinguishing fragments in the 100-1000 bp range
If you’re unsure, a 1% gel is the standard starting point for most applications.
Preparing Your Buffer
You need a running buffer both inside the gel and filling the electrophoresis chamber. The two most common options are TAE (Tris-acetate-EDTA) and TBE (Tris-borate-EDTA). Use the same buffer in both the gel and the tank.
TAE is the more common choice for routine DNA work and is preferred when you plan to cut bands out of the gel for downstream applications like cloning. TBE gives better resolution for smaller fragments (under 1,500 bp) and holds up better during long runs because it has higher buffering capacity. TAE can become depleted during extended electrophoresis, which distorts migration patterns.
Most labs prepare a concentrated stock solution and dilute it fresh before use. For a 50X TAE stock (1 liter): dissolve 242 g Tris base in distilled water, add 57.1 mL glacial acetic acid and 100 mL of 0.5 M EDTA at pH 8.0, then bring the volume to 1 liter. Dilute this 1:50 with ultrapure water to get your working 1X TAE solution. For a 10X TBE stock: dissolve 108 g Tris base and 55 g boric acid in 900 mL distilled water, add 40 mL of 0.5 M EDTA at pH 8.0, and adjust to 1 liter. Dilute 1:10 for your working solution.
Casting the Gel Step by Step
Weigh out the appropriate amount of agarose powder for your desired concentration. For a typical mini-gel requiring 30 mL of buffer, a 1% gel needs 0.3 g of agarose. Add the powder to your 1X buffer in an Erlenmeyer flask.
Here’s where safety matters. Microwave the mixture in short bursts to dissolve the agarose, swirling (not shaking) between pulses. Agarose solutions are prone to superheating, meaning the liquid can appear calm but erupt violently when disturbed. Use a flask that is 2 to 4 times the volume of your solution to leave room for boiling. Cover the flask loosely with a paper towel or vented plastic wrap. Only heat one flask at a time, watch it continuously, and always use heat-resistant gloves when handling the hot flask. Stop and swirl as soon as you see bubbles forming.
Once the solution is completely clear with no visible particles or unmelted solids, let it cool. You want it cool enough to comfortably hold the flask, but still liquid. If you’re adding a DNA stain directly to the gel (the most convenient method), add it at this point. For a concentrated stain like SYBR Safe at 10,000X, add 3 microliters per 30 mL of gel solution. Swirl gently to mix.
Pour the molten agarose into your casting tray with the comb already in place. Avoid creating bubbles. If any appear, push them to the edge with a pipette tip before the gel sets. Let it solidify at room temperature for 15 to 20 minutes until it turns opaque. Carefully remove the comb by pulling straight up to avoid tearing the wells.
Loading and Running Your Samples
Place the gel (still in its tray) into the electrophoresis chamber and fill with 1X buffer until the gel is submerged by a few millimeters. The wells should be positioned near the negative (black) electrode so DNA migrates toward the positive (red) electrode.
Before loading, mix each DNA sample with loading dye. A typical 6X loading dye contains 40% glycerol and tracking dyes like bromophenol blue. The glycerol makes your sample dense enough to sink into the wells instead of floating away into the buffer. The colored dyes let you see the samples as you load them and track how far the gel has run. Pipette your samples slowly into the wells, keeping the pipette tip just above the well opening to avoid puncturing the bottom.
Always load a DNA ladder (molecular weight marker) in at least one lane. This gives you reference bands of known sizes so you can estimate the size of your fragments.
For voltage, the standard recommendation is 4 to 10 volts per centimeter, measured as the distance between the electrodes in the tank (not the gel length). For the sharpest resolution, stay at or below 5 volts per centimeter. Higher voltage runs faster but generates more heat, which can distort bands or even melt the gel. For large fragments above 15 kb, keep voltage below 5 V/cm. A typical mini-gel run takes 30 to 60 minutes depending on fragment size and voltage.
Visualizing Your DNA
If you added a fluorescent stain like SYBR Safe to the gel before casting, your DNA is already stained and ready to image as soon as the run finishes. Place the gel on a transilluminator or imaging system to see the bands. SYBR Safe fluoresces green and works with UV transilluminators (at 254 or 300 nm) or blue-light transilluminators. Blue-light systems are gentler on the DNA and safer for the user, which matters if you plan to cut out bands for further use.
If you didn’t pre-stain the gel, you can soak it in a staining solution after the run, though this adds 20 to 30 minutes and may produce higher background fluorescence.
Common Problems and Fixes
Smeared bands instead of sharp ones are the most frequent complaint. The most common cause is simply overloading the well with too much DNA. Degraded samples also produce smears, so if your DNA sat at room temperature too long or went through too many freeze-thaw cycles, that’s likely the culprit. High salt concentration in your sample can also distort migration, causing lanes to run unevenly.
Bands that curve into a smile pattern across the gel usually indicate the voltage was too high, generating excess heat in the center of the gel. Lower the voltage or run the gel in a cold room. Uneven gels or slanted wells during casting will also cause lanes to migrate at different rates.
If your gel melts during the run, the voltage is far too high relative to the gel’s ability to dissipate heat. Thick gels are also more prone to this problem because heat gets trapped inside. For most applications, a gel thickness of about 5 mm works well.
Bubbles trapped in or near the wells during loading will cause samples to float out or migrate irregularly. Load slowly and steadily, and make sure your wells are fully submerged in buffer before you start pipetting.