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Oligo Purification using PolyAcrylamide Gel

The quality of an oligonucleotide (ie. how much is full-length product versus incomplete products) may be assessed by loading approximately 5 ug on a 19 or 20% polyacrylamide/urea minigel and staining with ethidium bromide. This same gel system may also be used in a preparative manner to purify the full-length from the incomplete oligonucleotide products.

  1. Recipe for 19% acrylamide/urea gel

    This recipe is for 30ml, which is enough for a standard size gel. You need only 5-10ml for a typical minigel (for quality assessment or for purifying only a portion of the oligonucleotide). There is no stacking gel in this gel system.

    • 15 ml of 40% acrylamide (1:22)*
    • 14.4 g urea
      • >
    • 3 ml 10X TBE


    1. warm to 37o to dissolve urea

    2. Add water to fill to 30ml. Just before pouring gel, add:
      • 120 ul of 15% ammonium persulfate
      • 20 ul TEMED
      3.  
    3. Pour the gel. Put in the appropriate comb (either a "welled" comb or preparative style), and allow the gel to polymerize at a 20º angle. Add more acrylamide solution to the top of the gel around the comb during polymerization if required.
       
      4.  
    4. Pre-run the gel in 1X TBE for 1 hour at 200V (minigel) to 400 V (regular size gel).


    *(1:22) 40% acrylamide
    22.9g acrylamide
    1.09g bis-acrylamide
    Fill to 60ml with water. This may be stored dark, 4ºC for several weeks after filtering and degassing.


  2. Running the Sample

    1. Prepare the sample by dissolving the desired amount in water and adding an equal volume of formamide. Load one lane with xylene cyanol/bromophenol blue, with an equal volume of formamide, as marker dyes (these run approximately equivalent to an 8 mer and 28 mer oligo in this gel system). Heat samples and markers to 65o for 10 minutes. Clean out the wells by gently squirting with a pasteur pipet until unpolymerized material is gone, and load the samples. If running a quality assessment gel to be stained with ethidium bromide, use approximately 5-10ug of material, with the final volume of sample (with the formamide) being appropriate for the well size of the gel you are using. If running a preparative gel for purification, load about one third to one half of the sample into the preparative well, keeping the final volume with the formamide appropriate for the well volume. Use a separate well to load the marker dyes.


    2. Run the gel under the same conditions as pre-running.


    3. When dyes have migrated the desired distance, remove the gel and either stain in ethidium bromide or wrap in saran wrap for ultraviolet shadowing (for the purification gel). Lay the wrapped gel on a treated silica plate (fluoresces at 254nm) and illuminate with a handheld UV illuminator at 254nm (short wave). The DNA will absorb the light and appear dark against the fluorescing silica. Work quickly, and using a clean razor blade cut out the largest (full-length) dark band. Cut the acrylamide strip into small bits and transfer to a tube(s).


    4. Add 1 ml of 0.3M Na0Ac per 2 cm length of gel slice. Agitate, if possible (improves elution of oligo), at 37ºC overnight. Spin down acrylamide and carefully transfer supernatant and respin to ensure all acrylamide bits are removed. Extract the supernatant with an equal volume of phenol:chloroform (1:1). Precipitate the oligonucleotide with 2 volumes of ethanol at -20ºC. (If sample will not precipitate, use 6 volumes of acetone or a mix of ethanol and acetone.) Wash the pellet with1 ml of 95% ethanol, dry, resuspend, and determine the concentration of oligonucleotide spectrophotometrically. You should be able to recover approximately 100-400ug of oligonucleotide per 200nmol synthesis with this method.