DGGE Protocol

Preparation of Acrylamide Solutions

0% Solution

40% acrylamide/bis (37.5:1, BioRad) 22.5mL
50xTAE buffer 3mL
Milli-Q Water 124.5mL
    • Combine first two components
    • Bring volume up to 150mL with Milli-Q
    • Aliquot into three 50 mL centrifuge tubes
    • Store at 4oC

80% Solution

40% acrylamide/bis (37.5:1, BioRad) 22.5mL
50xTAE buffer 3mL
Formamide (freshly deionized) 48mL
Urea 50.4g
Milli-Q Water ~75mL
    • Combine first four components (urea dissolves slowly)
    • Bring volume to 150mL with Milli-Q
    • Aliquot into three 50 mL centrifuge tubes
    • Store at 4oC

Deionization of Formamide

Formamide 70mL
Mixed Bead Resin 1-2 spoonfulls
  • NOTE: Formamide must be kept wrapped in foil throughout its use due to its sensitivity to light
  • Combine formamide and resin
  • Mix on stir plate for 60 minutes
  • Use Whatman filter paper to remove resin from formamide

Cleaning the Glass Plates

  • Remove one set of glass plates (on tall and one short) and two spacers from the soap solution
  • Wash both sides thoroughly with dish soap and rinse with tap water followed by distillied water
  • Wash the gel-side of each plate with the following reagents in specified order:
    • Windex
    • Acetone
    • 70% Ethanol

Plate Assembly

  • Place the spacers along the edges of the tall glass plate (gel-side up), with the notches facing towards the center of the plate
  • Place the short glass plate (gel-side down) onto the spacers. Align the assemblage with all sides flush, making sure spacers extend slightly past the glass plate edges
  • Lightly tighten the supports to either side of the assemblage, ensuring that the arrows on the supports are facing up and towards each other
  • Place the glass plate assemblage onto the positioning area of the already leveled base
  • Release the tension of the supports and position the plates and the spacers so that they are flush against each other. Proper alignment of the plates is essential for the formation of a good seal
  • Tighten the supports to lock the assemblage into place
  • Pull out the knobs on either side of the base and insert the plates into the pouring area, ensuring that the assemblage is flush against the grey strip
  • Push in the knobs and rotate the screws in a counterclockwise direction (as looking straight down) to secure the plates
  • Verify that there is some bowing in the grey strip to ensure a proper seal

Pouring the Gel

  • Make three solutions from the following chart in 15 mL centrifuge tubes (one of these must be the 0% plug)
0% 10% 20% 30% 40% 50% 60% 80%
 0% solution 6 mL 9.625 mL 8.25 mL 6.875 mL 5.5 mL 4.125 mL 2.75 mL 0 mL
80% solution 0 mL 1.375 mL 2.75 mL 4.125 mL 5.5 mL 6.875 mL 8.25 mL 11 mL
APS (10%) 44 µL 52 µL 52 µL 52 µL 52 µL 52 µL 52 µL 52 µL
TEMED 6.0 µL 6.9 µL 6.9 µL 6.9 µL 6.9 µL 6.9 µL 6.9 µL 6.9 µL
  • Set up the puring apparatus, making sure that the valve connecting the two wells of the gradient maker is closed (~30o from horizontal)
  • Turn on stir bar to 6
  • Center the comb in between the plates and place the syringe between the plates and against one of the spacers
  • Thoroughly mix the acrylamide solutions (20% with either 40% or 60% or 80%) with the APS and TEMED
  • As quickly as possible, pour the higher concentration solution into the front chamber (the one closest to the outlet valve) and pour the lower concentration solution into the back chamber
  • Open the valve between the wells on the gradient chamber (bring to horizontal position) and begin the mixing and the pouring of the gel
  • To promote even mixing, reduce the speed of the stir bar as the gel pours (one should just barely be able to see the mixing in the front chamber). If the stirring speed is too great, the higher concentration denaturant will flow back into the other chamber
  • When the wells are approximately 75% empty, thoroughly combine APS, TEMED and the 0% solution. Add this mixture to the rear well as the front one empties
  • Once both wells are empty, clean the pouring apparatus by running Milli-Q water through the system multiple times
  • Allow the gel to sit for at least two hours. If leaving overnight (recommended), cover assemblage with a large ziploc bag and store in a large tupperware container with a beaker full of water (to reduce dessication)

Assembling the D-Code (Gel Running) System

  • Clean off a set of glass plates (dish soap only)
  • Line the space between the plates with Parafilm
  • Lock plates into place with supports, ensuring they are flush
  • Carefully remove the comb from the gel, along with any excess gel from around the wells
  • With the tall glass facing out and short glass facing in, snap both assemblages into the holder
  • Fill D-Code system with approximately 7 L of 1xTAE (140 mL 50xTAE and 6860 mL Milli-Q water) until fill-line is reached
  • Place the holder down into the buffer, ensuring that it is properly in place
  • Place the top of the D-Code system on top of the apparatus. Verify that all electrical connections are created
  • Plug in electrical chord and turn power switch on
  • Set temperature to 65oC and turn on the heater and pump. Make sure that the buffer level rises and remains above the negative electrode wire
  • Allow D-Code to reach 65oC (90 to 120 minutes)

Loading the Gel

  • Turn off power and remove top of D-Code system
  • Prepare samples by combining equal amounts of template (at least 300 ng) with 5x loading buffer in 500 µL PCR tubes
  • Using a P-100 and corresponding gel loading tips, load samples by placing the pipettor tip half-way into the well and expunging sample with extreme care (loading the wells is made easier by avoiding air bubbles at the pipette tip and expunging slowly)
  • Load the standard into every fourth lane
  • Replace top to the D-Cody system, ensuring all electrical connections are made
  • Plug in D-Code and plug it into both the electrical outlet and the BioRad PowerPack300 (with DCODE chip)
  • Turn on D-Code system and set temperature to 60oC
  • Turn on heater
  • Set PowerPack300 (with DCODE chip) to 200V
  • Turn on the pump only after samples are pulled into the gel (approximately 15 minutes)
  • Let gel run for 3.5 to 4 hours at 200V (or 12 hours at 100V)

Staining and Imaging the Gel

  • Drain the excess buffer from the top of the gel and dismantle the apparatus
  • With the upmost of caution, remove the supports and spacers
  • With extreme care, use one of the spacers as a lever to pry off the short glass plate, leaving the gel on the tall glass plate
  • Soak the tall plate (with gel) in an ethidium bromide stain, and place on rocker (slow mode) for 30 minutes
  • Destain in Milli-Q water for 15-20 minutes
  • Thoroughly wet the imager’s glass plate with water
  • Tilt the glass plate onto the wet surface of the imager, allowing the gel to slowly slide off (continually wetting the edges of the gel is helpful)
  • Once imaging completed and bands to be cut are identified, remove the gel by re-wetting the glass plate and carefully sliding the gel onto it
  • Transport gel to transilluminator for band cutting

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