# Fong:DNA gel electrophoresis

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See the excellent consensus protocol for an overview. This version is intended to provide instructions specific to the equipment and reagents available in the Fong lab.

Agarose gel electrophoresis is used to separate DNA fragments by size. DNA carries a net negative charge, so in the presence of an electrical current it will tend to migrate towards the positive (red) electrode. Smaller DNA fragments can navigate the agarose gel matrix more easily and therefore migrate further than longer fragments.

## Casting agarose gels

Choose buffer and agarose gel concentration according to the expected size range of the DNA to be separated (see table). 1% agarose with TAE is a good choice for general procedures. Choose the casting tray and gel comb depending on the number and volumes of the samples to be loaded. The small casting trays need about 50mL total volume, and the large trays about 100mL.

Final Concentration of Agarose for DNA electrophoresis (%)
Size Range (bp) 1x TAE 1x TBE
1,000-23,000 0.60 0.50
800-10,000 0.80 0.70
400-8,000 1.00 0.85
300-7,000 1.20 1.00
200-4,000 1.50 1.25
100-3,000 2.00 1.75

### Procedure

1. Measure appropriate weight of agarose and add to a sterile 500ml flask.
2. Add the appropriate volume of buffer, mix by swirling the flask, and microwave for about 1.5-2 min. Use this time to set up the casting tray.
3. Allow the mixture to cool enough that you can comfortably touch the flask with your gloved hand, and then add 3.5μL ethidium bromide(for 50mL) and swirl to mix. Dispose of pipette tip in chemical waste.
4. Pour mixture into the casting tray, and use the comb to carefully remove any bubbles. Place the comb in the slot and allow the gel to cool to a solid (~5-10min).
5. Carefully remove comb and place gel into the gel box so that the samples will run towards the red (+) electrode. Gently pour the same buffer used in the gel over the gel until the top of the gel is just covered.
6. Dislodge remaining cooled gel in the flask and dispose in chemical waste (room 409).

### Materials

• Sample
• ddH2O (optional)
• Micropipettes and tips
• Parafilm (optional)

### Procedure

For diagnostic purposes, plan to use enough DNA that clear bands can be obtained, or, for separation purposes, use the entire sample. Use 2μL loading dye for every 10μL sample. If your sample is less than about 3-5μL, it is often a good idea to dilute it with ddH2O so that it is at least 5μL.

1. Load left lane and, space permitting, right lane with ~5μL DNA ladder.
2. On a small piece of parafilm, place a droplet of loading dye for each sample to be run, allowing enough space to prevent any mixing. Add water to each droplet at this step if necessary.
3. With a new tip, add DNA sample to one droplet of loading dye and mix by pipetting.
4. Pipette mixture carefully into a well of the gel, noting carefully which sample corresponds to each lane.
5. Repeat from step 3 until all samples are added.
6. Carefully place lid on gel box and connect electrodes to power supply. Run at 100V for ~1hr. Running voltage and time can be adjusted to improve resolution or lower running speed, depending on your needs. Watch for the blue loading dye to cover about 60% of the gel length.

## Visualizing gel

### Materials

• Gloves! (as always)
• BioRad Universal Hood gel imager, in room 407

### Procedure

1. Remove gel in casting tray from gel box and carry into 407 in some container to prevent dripping (plastic boxes are located by the sink in 409)
2. Open bottom drawer of imager, spray with ethanol and wipe down with Kimwipes. Carefully place gel (without tray) on the center of the imaging glass.
3. Close drawer and press the Trans UV button on the imager.
4. On the computer by the imager, open Quantity One, choose 'Select Scanner', and select 'Gel Doc...' (the first choice).
5. Click on 'Auto Expose' to image the gel. You may need to manually adjust the Low, High, and Gamma bars to get a good image.
6. Save the image in the appropriate folder.
7. Remove the gel from the drawer and dispose in chemical waste (room 409).
8. Spray drawer again with ethanol and wipe dry.