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oligo_purity_by_denaturing_PAGE

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					                    Checking oligo purity by denaturing PAGE

For short oligos (<18nt), use 20% gel acrylamide stock. I have not run longer oligos, but I
have recipes for 18% and 12% gel stock also.

20% acrylamide gel stock (1L):
    420 g urea
    200 mL 5x TBE
    500 mL 40% acrylamide (19:1 acryl:bis)
    Mix in a clean glass beaker with a stir bar over low heat.
    Allow to cool, then sterile filter using Nalgene filter/bottle.
    This stock solution must be stored in the dark!

18% Acrylamide Gel Stock:
    420 g Urea
    200 mL 5x TBE
    450 mL 40% Acrylamide (19:1)
    Fill to 1L with sterile ddH2O in a graduated cylinder.
    Mix all components in a beaker with a stir bar over low heat, once the urea is
      dissolved, allow to cool.
    Sterile filter using Nalgene filter/bottle.
    This stock solution must be stored in the dark!

12% Acrylamide Gel Stock:
    420 g Urea
    200 mL 5X TBE
    300 mL 40% Acrylamide (19:1)
    Fill to 1L with sterile ddH2O in a graduated cylinder.
    Mix all components in a beaker with a stir bar over low heat, once the urea is
      dissolved, allow to cool.
    Sterile filter using Nalgene filter/bottle.
    This stock solution must be stored in the dark!

5x TBE (1L):
     54 g Tris-base
     27.5 g boric acid
     20 mL 0.5 M EDTA
     H2O to 1L

Polymerization ratio:
    ~30 mL acrylamide gel stock
    300 L 10% APS
      12 L TEMED
      This volume is good for one 20cm x 20cm x 0.75mm gel.

To make and run a gel:
   1. Wash the two gel plates, then, using a Kim wipe, wash the faces that will be
      touching the gel with Windex first, then with ethanol. Wash the spacers and
      wipe with ethanol. Assemble on top of some tip boxes, Tupperware, or some
      other raised surface. Sandwich all three spacers between the two plates so that
      the clean plate faces are aimed towards each other; the bottom spacer should
      be flush with the bottom of the plates (sticking out on the sides) and the side
      spacers should seat flush against the bottom spacer, with the excess sticking out
      of the top of the plates (see picture below). Clamp the bottom spacer with two
      clips. Adjust the side spacers so that they are flush with the edge of the bottom
      spacer and the sides of the plates. Insure a tight connection by placing a clip at
      the bottom of each side spacer and then pushing the bottom and side spacers
      closer together. Once a tight connection is achieved, add two more clips to each
      side of the plates.
   2. Pour ~ 30 mL of the Acrylamide Gel stock of choice into a 200 mL beaker. Pipette
      in the APS and the TEMED in the amounts shown above and gently swirl, taking
      care to get no bubbles.
   3. Tilt the plates with one hand and use the beaker to pour the gel solution in one
      corner (see red arrow in image below) into the plates in a continuous stream.
      Make sure the gel is free of bubbles (this may not be possible, but pouring in one
      continuous, steady stream will minimize bubbles, and you will usually only get
      only one or two at the side where you are pouring – this does not matter) and
      then immediately put the comb in the top by rocking it into place. The comb
      slots are rounded; seat the comb in between the plates such that an even and
      small amount (1-2 mm) of rounded slot protruded above the front plate. Allow
      to sit ~15 minutes or until the gel is visibly solidified (tilt the beaker with the
      excess gel solution to observe polymerization).
   4. Remove the bottom spacer and the comb. Label your wells and center your
      samples in the wells (e.g. if you have 4 samples, load as shown below). Clamp
      the plates against the gasket with the wells facing the upper buffer reservoir
      (two clamps on each side of the plates), so you can read the labeled wells on the
      back. Place one of the 100 mL bottles of water (rad gel bench) in the upper
      buffer reservoir. Fill with 1x TBE to ~ 1 cm above the wells. Fill the lower buffer
      reservoir with 1x TBE to 5-10 mm above the bottom spacer line. Remove bubbles
      from the space between the plates where the bottom spacer used to be with the
      bent needle syringe on the rad gel bench. Plug into a power source and pre-run
      ~45 mins at 20 watts (~500V) the volts is not exact, but the 20 watts is important
      (the voltage will go up as the gel runs.
   5. Put your samples in loading buffer (usually 80% Formamide, 25 mM EDTA, and a
      dye or two – bromophenol blue and/or xylene cyanol). Heat to ~90C for about 3
      mins. Turn off the power to the gel and squirt out the settled Urea from each
   lane (including any you are not using) using a syringe. Then load samples and run
   at 20 watts for ~ 1.5 hrs or until the loading dies are almost off the gel.
6. To stain the gel, carefully pull the plates apart and transfer the gel to a bin
   containing enough Stains-All working solution (Pardi Lab) to cover the gel. The
   stain is light sensitive, so be sure to cover with foil or something similar. Put on
   the gel shaker at slow revolutions for ~15 mins or until staining has occurred.
   Destain to desired amount in water. Sandwich between 2 transparencies and
   image on scanner.




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