Contact: Mary Jo Kirisits

N.B. Acrylamide is a neurotoxin.  Polyacrylamide is never completely polymerized and therefore contains some acrylamide.  Always wear gloves when dealing with (poly)acrylamide.  Dispose of all (poly)acrylamide waste through ORS.


  1. Amplify DNA with one fluorescently labeled primer. I use 8F (labeled; LI-COR: IRD-800 tag) and 926R.
  2. Clean amplicon with Wizard PCR preps or Millipore Microcon (YM-30; Fisher: 42410).
    1. With the Microcon, flush each filter with water. Add 500 μL water and centrifuge at 14,000 x g for 8 minutes.  Flip and centrifuge at 1,000 x g for 2 minutes.
    2. Combine 2 PCR reactions (100 μL total) and add to filter. Add 350 μL of water. Centrifuge at 14,000 x g for 8 minutes.
    3. Put filter in new centrifuge tube. If there is still liquid visible in the microcon, do not add any additional liquid.  Otherwise, add 40 μL water.  Incubate 1 minute.  Flip and centrifuge at 1,000 x g for 2 minutes to collect the DNA.  Put filtrate through a second time (1,000 x g, 2 minutes).
  3. Measure the DNA concentration of the PCR product at A260nm. The measurement will be more accurate when the absorbance is greater than 0.15, but it may be difficult to achieve such a high absorbance.  I usually use 10-20 μL of amplicon in a 150-μL sample. 
  4. Digest with 4-base cutter endonuclease. I use 2-μL HhaI, RsaI, and MspI with a 37 oC  incubation for 12 hours.  I use approximately 300 ng amplicon in a 20-μL reaction.  At the end of the digestion, inactivate HhaI, RsaI, and MspI  by exposing the reactions to 65 oC for 20 min.
  5. During this time, the plates and gel can be prepared. Wash the plates with a brush and NeuRad soap (Fisher: 04-355-10).  Water beads up on one side of the plates but flows evenly off of the other side.  You are interested in the side of the plates where the flow is even.  Otherwise, the gel mixture can get hung up, causing bubbles. Rinse the plates with ethanol.  Put a Kimwipe on the bench and lean the plates against the wall so they can dry. 
  6. Look at the plates for smears of any kind. Wipe with distilled deionized water and a Kimwipe.  Never re-use the same part of the Kimwipe because this can cause acrylamide to get smeared all over everything.
  7. Prepare the bind silane (Sigma: M6514) stock solution: 50 μL bind silane + 10 mL absolute ethanol. Add 100 μL of the stock solution to 100 μL of 10% acetic acid.  Use a Q-tip to apply it to the glass plates in the vicinity of the comb, and let it dry.
  8. Put the 0.4 mm spacers on the plate. If there are chips on the big plate, put them at the top so they don’t interfere with the spacers.  Put the large plate in back and the small plate in front, with the piece cut out of the small plate at the top.  Make sure that the spacers are flush with the sides and bottom of the plates.  Place this sandwich in the clamps.  Tighten the bottom clamps to hold the plates together.
  9. Place the flat piece, with the grooves facing up, onto the gel stand. Place the plate sandwich onto the stand.
  10. Add a plastic plate to the front of the sandwich and tighten with the upper set of clamps. This is to keep the glass plates tightly together.  This is really important to getting sharp bands.
  11. Prepare 25 mL of 5.5% acrylamide gel (LI-COR: 827-05215) and add the APS:
    1. 25 mL Kb+ gel mixture (5.5%, filtered through 0.2 mm filter)
    2. 167 μL 10% APS       
    3. 16.7 μL Temed                       (Do not add Temed now; Temed is added just before pouring.)
  12. Make a test and real acrylamide plug. Using small Falcon tubes, add 3 mL of the acrylamide mixture to 2 tubes.  Use a pipet tip with a filter to transfer the acrylamide mixture.  Add 10% APS to each tube, and Temed to only one of the tubes.  If the test plug (with Temed) polymerizes after a few minutes, add Temed to the real plug. (Note:  Many people make fresh APS every time, but this is not necessary.  It can be used for 1 month.  Keep it in the refrigerator.  Temed is stored at room temperature.)
  13. Test and real plugs:
    1. 3 mL acrylamide mixture
    2. 50 μL 10% APS
    3. 5 μL Temed
  14. Using the same pipet tip that was used for the acrylamide mixture, pipet the plug (acrylamide mixture, APS, TEMED) into the gel apparatus. Pipet into the three grooves on the bottom of the gel stand.  After a few minutes, tip the gel sideways to see if the plug has polymerized. 
  15. Add 16.7 μL Temed to the 25-mL acrylamide mixture. Hold the plate sandwich up at an angle, and use a syringe to pour the gel.  Squirt the gel mixture on one side of the plate.  If there is a bubble, stop pouring the gel; and tap the glass to get rid of the bubble.
  16. Put the comb in at the top. A square tooth comb, 0.4 mm, works well.
  17. Lay the plate sandwich at a slight angle, with the top part resting on the flat end of the gel stand. Put the weight (15 L carboy filled with liquid) on top of the plates.
  18. Let the gel polymerize for 2-3 hours.
  19. Mix 2 μL of digest and 2 μL of loading dye (LI-COR: 830-05629). Add 3.5 μL of size markers (50-700 bp; LI-COR: 4000-45) to another tube.  Denature them at 95 oC for 2 minutes; this can be done in the thermocycler.  After denaturing, put them immediately on ice to prevent re-annealing of strands as the tubes cool.  Leave them on ice (10 minutes) until you are ready to load them.
  20. Take the plate sandwich from the stand and put it on a clean Kimwipe. This will minimize further cleaning of the plates prior to analysis.
  21. Pull the comb straight out of the sandwich. (If you have a problem seeing the lanes to load them, outline the lanes with a Sharpie marker before removing the comb.)
  22. Remove the small flat plate from the top part of the sandwich, and open the clamps a little more so that the upper buffer tank can be accommodated. The upper buffer tank has something akin to an O ring in the rear (like a piece of tubing), to create a good seal.  When inserting this, make sure that the tubing does not get moved out of its groove.  Use the upper set of clamps to hold the buffer tank in place.
  23. Use DDW and a clean Kimwipe to polish the plates where the laser will read.
  24. Put the lower (gray) buffer tank in place on the sequencer. Put the plate sandwich into the holder in the sequencer.  Put the covers over the upper and lower buffer tank.
  25. Connect the electrodes on the sequencer to that on the buffer tank.
  26. Fill the lower and upper buffer tanks to the indicated line with 0.8 X TBE. (10X TBE from LI-COR: 826-05213.)
  27. Clean out the wells of the gel. Using a syringe and needle and take up a few mLs of TBE buffer. CAREFULLY shoot this along the tops of the wells to dislodge bubbles.  Do this at least 3 times.
  28. Close the sequencer door.
  29. Go to the computer. Press 3 keys to unlock the computer: <>>, <ENTER>, <SHIFT>.
  30. Click on Base ImagIR Data Collection. Then click on Data Collection Dev 8.  Device 8 is the sequencer on the left hand side.
  31. Open a new file on the D drive. Give it a name with no dashes or slashes. The date works well (i.e. 122000 for December 20, 2000).
  32. Go to scanner control, and input the appropriate parameters:
    1. 1500 V
    2. 35 mA
    3. 35 W
    4. 20 frames
    5. Speed 3
    6. Signal channel 3
    7. T = 45 oC
  33. Turn the voltage and scanner ON. There is a column to the left of these toggle buttons, written in light gray, which tells you what the actual conditions are.  Make sure that this column says ON after you turn these things on.
  34. Let this pre-electrophorese for 20-30 minutes. This will allow the gel to be heated and flushed with buffer.  The temperature should reach about 40 oat this time, and the samples can then be loaded.
  35. Turn the voltage and scanner to OFF. Make sure that the light gray left hand lane also says OFF before you open the scanner door.
  36. Clean the wells again to remove bubbles.
  37. If it is difficult to see the wells, it may be good to put a sheet of white paper behind them.
  38. Load size markers (1 μL) on either end of the samples, and load one lane of size markers in the middle. Load the samples (1.5 μL) consisting of the digested amplicon plus loading buffer.  You can use the same flat tip (VWR: 53550-436) for all the loading.
  39. Turn the voltage and scanner back ON.
  40. Focus the scanner. Go to OPTIONS, FOCUS, AUTO.  You want the red line to be in the middle of the peak.  Usually the signal gain is 450-500 and the signal offset is 100-150 after the focusing.
  41. Go to OPTIONS, AUTOGAIN. The settings should be 20, 10.  Then press AUTO.
  42. After doing autogain, you can focus again if needed. (If the red line was not in the middle of the peak the first time.)
  43. Now we will start seeing bands on the gel after about 15-20 minutes. The big blob at the beginning is the primer band.  (The un-used primer will be the smallest band there, and therefore will move to the scanning location first.)
  44. If the sample bands are really dark or seem to run together, the sample can be diluted. The diluted samples can be loaded onto the same gel once the first samples have run off.
  45. At the end of the run, turn the voltage and scanner OFF. Wait until it says OFF in the light gray column on the left before opening the sequencer.  Otherwise, it will give you an error.
  46. Open the sequencer. Unplug the electrode.  Remove the buffer tank covers. 
  47. Loosen the clamps on the upper buffer tank. Let the buffer drain into the lower tank. 
  48. Rinse and dry the buffer tanks and covers.
  49. Remove the plate sandwich. Use a flat spatula to help separate the plates and to remove the gel from the plate.  The gel and all gloves, pipet tips and Kimwipes that touched the acrylamide, can be disposed of in a yellow bag that goes to ORS.
  50. Wash the plates with soap so that no acrylamide is left. Place on a clean Kimwipe and lean them against the wall to dry.

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