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Materials

• Agarose
• 1× TAE (Tris-acetate–EDTA buffer)
  • 50× TAE: 40 mM Tris base, 20 mM acetic acid, 2 mM EDTA, pH 8.5
  • Dilute 50× TAE to 1× for use. Add 80 mL empty carboy and fill to 4 L with diH₂O.
• DNA dye (10,000× ethidium bromide)
• 6× DNA loading dye
• DNA samples
• 1× DNA ladder in 1× DNA loading dye
  
  
• Bottle or flask large enough for gel volume
• Microwave oven
• Gel caster
• Gel tray of desired length
• Gel comb of desired size/thickness
• Electrophoresis chamber with clean-enough 1× TAE
• Electrophoresis power supply
• Microwave oven
• Gel viewing equipment (blue-light source and orange glasses, or gel imager)

XC: xylene cyanol. BPB: bromophenol blue. OG: Orange G. 

Procedure

Gel Preparation

1. Weigh and mix agarose powder with 1× TAE to desired agarose concentration in a flask or bottle. An flask for agarose is usually by the gel station. Make enough volume for however many gels at their respective concentrations.
   Alternatively, obtain previously prepared solid agarose gel in bottle/flask.
   • Agarose gels are commonly used in concentrations of 0.7–2%m/V, depending on the expected size of bands needing to be resolved. See table.
2. With a loosened cap, microwave agarose suspension or solid until completely clear and homogeneous liquid (≈1 min per 100 mL on high). Watch as it boils to stop the microwave before it can overflow. Swirl, handling with a silicone mit. Continue microwaving in smaller increments if necessary.
   If needed, dilute molten agarose with 1× TAE to desired percentage, swirling to mix. Make enough for the desired casts and the level you wish to fill. See below.
   • E.g. you can keep a bottle of 2% agarose and melt it when needed, diluting with TAE in a second flask to the desired 0.7–2% concentration. Pouring in a second container and dilution also cools it, so the poured gel solidifies faster.
3. If prestaining the gel, to a final 1× add 10,000× DNA dye (GelGreen or GelRed) to the molten agarose and swirl until color is uniform.
   • Dye can be incorporated into the gel during preparation (prestaining), after gel electrophoresis by incubating in a dye bath (poststaining), or solely added to the DNA sample (sample dying). Prestaining is most robust. Post-staining requires more time and monitoring. Sample staining impacts DNA migration, so is not best for analytical purposes and must be used with a similarly dyed ladder.)
   • Dyes like ethidium bromide are mutagens. Even safe dyes should be handled cautiously.
4. Position tray(s) in center of clamp and tighten clamp moderately. Place desired comb(s) in tray slots, oriented so the prongs are closer to the top of the gel. Ensure tray is level using bubble level, adjusting gel caster's screw feet.
   • Pick gel casting trays appropriate for your use. The short 7 cm tray holds 40 mL when filled to the top of comb prongs. The long , and the 10 cm tray holds 55 mL when filled to the top of the comb prongsmL.
     For gel purification purposes, you may want a thick gel (tray filled to top of comb prongs) to minimize the number of wells needed to hold the samples. For analytical purposes, you may need only a thin gel (casted to half prong height) to hold small volumes of samples such as analytical DNA digests.
   • Pick comb appropriate for your use. Thinner combs are superior for analytical purposes, as they result in thin bands more accurately assessable. Thicker combs are can be superior for purification purposes, as they hold more sample. A full-height narrow well holds ≈30 µL; a full-height wide well holds ≈60 µL.
5. Pour molten agarose in trays to desired level while still hot, but not boiling to prevent tray/clamp warping. Use comb to pull bubbles and impurities away to end of gel.
   • Bubbles can be pushed to the edges of the gel using the comb.
6. Allow the gel to solidify at room temperature or level in the deli fridge  at 4°C, 8–15 min.
   • Pre-chilled caster /clamp and tray speeds solidification.

Sample Preparation, Loading, and Electrophoresis

1. Mix samples with 6× loading dye (loading buffer). If sample-staining, ensure the 6× loading dye contains 1× GelGreen/GelRed.
   • Loading dye does not typically contain any DNA dye, only tracking dyes that allow estimation you to see your sample when loading and visibly estimate DNA migration progression without fluorescence(see chart on left). Loading dye also contains glycerol, sucrose, or PEG to weigh down the sample in the well.
2. Fill electrophoresis chamber with 1× TAE (running buffer) until it's a bit under the gel+tray height.
   • If running buffer is cloudy or has crystals, empty contents into gel liquid waste tank. Gently rinse chamber in sink without pointing stream at deep ends, where it can damage the electrode filaments. Do a final rinse with diH₂O. Refill with 1×TAE.
     
   • When using ethidium bromide, some people add it to the running buffer as well as the gel.
3. Remove comb from cast gel and submerge gel with +tray in an the electrophoresis chamber, oriented with wells at the top and the negative (black) terminals of the chamber at the top.
   If TAE doesn't cover the gel, add sufficient TAE. 
   • Gels can be cut to save lanes for later use. The tray isn't always necessary. Gels must be stored in TAE + DNA dye (if prestained), TAE (if unstained), or wrapped ≈airtight in plastic wrap.
   • If chamber liquid is cloudy or has crystals, empty contents into gel liquid waste tank. Gently rinse chamber in sink without pointing stream at deep ends, where it can damage the filament. Do a final rinse with diH2O. Refill with 1×TAE.
4. Load samples and, finally, ladder into wells. Pre-dyed ladder is generally at the gel station.
   • While loading, maintain positive pressure on the sample to prevent bubbles or buffer from entering the tip. Release any air at the tip of the tip so no bubbles are released when loading.
     Hover the tip of the tip above the well. Slowly and steadily, push the sample out and watch as the sample fills the well. After all of the sample is unloaded, push the pipettor to the second stop as you raise the pipette out of the buffer.
   • Pipetting into the bottom of the well (a more difficult method) allows more sample to fit in a well. Moving the gel or pipette tip while it's in a well containing sample may expel part of the sample from the well. Keep everything steady when ejecting inside a well.
5. Secure the lid onto the chamber. Plug , and plug in lid into power supply, matching .
   
   • Match terminal colors. Ensure negative
   ,
   • black terminals are at the top of the chamber and lid and positive red at the bottom. Thus anionic nucleic acids run to the red anode. Run to red. 
     
   Program in voltage (80–150 5–10 V/cm electrode distance) and run time (15–45 min) and runstart. Confirm running by spotting bubbles coming from the terminalsemanating from the electrodes.
   • Common settings are 80–120–150 V, 60–30–15 min. Run time is inversely correlated with voltage.
6. Run until bromophenol blue has run ≈75% the gel length or Orange G is ≈90% the gel length. Try not to run DNA off the gel or into a lower row of lanes.
   • 120 V for ≈25 min is sufficient for most applications. Higher voltage shortens run time, but can sometimes result in band smearing or other problems due to heating.
   • The lower, blue/purple band migrates as ≈300 bp DNA, and should typically migrate ≈75% of the lane length before ending.
   Stop the program, remove the lid, and remove the gel with tray.
   • Don't leave the gel in the buffer; NAs will slowly diffuse out.
7. (opt.) If When the run is over, if poststaining, stain the gel in TAE + 1× 1–3× DNA dye solution, rocking for 20–30 15–30 min, followed by destaining in water if necessary for sensitivity.
8. Inspect/cut the gel under blue light with orange glasses or UV with UV shield. Image the gel with the blue light (or UV) imager: Open Canon EOS utility on the computer to awaken the connected camera and use Remote Live View to adjust the gel position and autofocus before taking a picture. The camera should typically be in the No Flash setting (auto-set exposure, ISO, and aperture). Exit EOS Utility when done.
   • Blue light is safer to work with and less damaging to DNA than UV light. Requires GelGreen, SYBR green, SYBR Safe, etc.

GelGreen and GelRed Dyes

Stability

GelGreen is known to be stable for years in aqueous solution at room temperature, including dilutions. GelGreen and GelRed do degrade slowly over light exposure time, so it is best to keep the dyes in amber tubes and dyed gels in closed containers.

Dyed gels can be stored for later use by wrapping in plastic wrap and placing in a box at 4°. Low temperature further slows evaporation, allowing storage for several weeks.

Toxicity

So long as we don't get the DMSO-based one, GelGreen/Red aren't glove-permeable or cell-permeable. We currently own a water-based GelGreen.

SYBR® Safe is one of the original ethidium bromide alternatives to be marketed as safe. However, it has been reported to show mutagenicity after metabolic activation in the Ames test (9). While SYBR® Safe is reported to be non-mutagenic in Syrian hamster embryo (SHE) cells and L5178YTK +/- mouse lymphoma cells (10), it was tested at concentrations well below its 1× working concentration of 0.66 µg/mL (11) due to its excessive cytotoxicity (9). These results are consistent with the fact that SYBR® Safe rapidly penetrates cell membranes and stains the nucleus of live cells. Nancy-520, a SYBR derivative, also readily penetrates living cells to interact with DNA, suggesting that it has similar potential for cytotoxicity. Moreover, these dyes also showed no advantage in sensitivity compared to GelRed™ or GelGreen™.

SafeView™ FireRed was found to contain propidium iodide, which is commonly used to stain dead cells. Consistent with this, SafeView™ FireRed did not stain live cells. However, it has been reported that propidium iodide is cell permeable and cytotoxic to J774 cells with prolonged exposure (1 day) (5).

In the same study, GelRed™ and GelGreen™ were found to be non-permeable and non-toxic for up to three days of incubation (5). EZ-Vision® In-Gel also was found to be cell membrane impermeable, consistent with the finding that it contains DAPI. However, EZ-Vision® has very low sensitivity, and the added disadvantage of being incompatible with visible blue gel imagers, unlike GelGreen™.