Plant DNA Barcoding (Rapid DNA Isolation)

Protocol

Note: See expanded protocols at dnabarcoding101.org

Materials

  • Lysis solution (6 M Guanidine Hydrochloride GuHCl) (120 µL)
  • Wash buffer (480 µL)
  • TE buffer (75 µL)
  • Specimen tissue sample(s) (from Part I)
  • Whatman No.1 Chromatography paper discs (2, 3-mm diameter)
  • Microcentrifuge tubes (1.5 mL)
  • Micropipettes and tips (2–1000 µL)
  • Permanent marker
  • Sterile plastic pestles
  • Microcentrifuge tube rack
  • Sterile tweezers

Steps

  1. Obtain plant, fungal, or animal tissue ~10 mg or ⅛- to ¼-inch diameter by removing a piece of the tissue with a razor blade, clean tweezers, scissors, or back of a 10-µL pipette tip to enable efficient lysis. If you are working with more than one sample, be careful not to cross-contaminate specimens. (If you only have one specimen, make a balance tube with the appropriate volume of water for centrifugation steps.) Be sure to preserve the remainder of the organism at -20°C or in 96–100% Ethanol.

    • Tip: Tissue should be no larger than a grain of rice. Using more than the recommended amount can affect amplification.

  2. Place tissue in a clean 1.5-mL tube labeled with a sample identification number.

  3. Add 50 µL of lysis solution to each tube.

    • Tip: Lysis solution dissolves membrane-bound organelles including the
      nucleus, mitochondria, and chloroplast..

  4. Twist a clean plastic pestle against the inner surface of 1.5-mL tube to forcefully grind the tissue for at least 2 minutes. Use a clean pestle for each sample. Ensure the sample is ground into fine particles.

    • Tip: Grinding breaks up cell walls and other tough material. Once ground, the sample should be liquid, but there may be some particulate matter remaining.

  5. For each sample, use a separate sterile tweezer to add one 3-mm diameter disc of Whatman No. 1 Chromatography paper to the lysed extract. Tap or flick the tube gently to ensure the disc is fully submerged in the extract. Allow the disc to soak in the extract for 1 minute.

    • Tip: Whatman chromatography paper binds the DNA, helping separate DNA from contaminants.

  6. While the disc is soaking, add 200 µL of wash buffer to a clean 1.5-mL tube labeled with the sample identification number.

    • Tip: Wash buffer will remove contaminants that can inhibit PCR while the DNA remains bound to the paper.

  7. Remove the disc from the extract using a sterile tweezer or pipette tip and transfer the disc into the fresh tube containing wash buffer. Tap or flick the tube to mix for 5 seconds, then allow the disc to sit in the wash buffer for 1 minute.

    • Tip: Discard/set-aside the tweezer following Step 7. Use of the tweezer to transfer the disc in future steps will contaminate the disc with impurities that may affect PCR.

  8. Use a sterile pipette tip to gently drag the disc out of the wash buffer and up the tube wall to dry at the top of the tube. Ensure that little to no debris is attached to the disk. Allow the disc to air dry for 2 minutes to evaporate the ethanol on the disc.

    • Tip: Ethanol in the wash buffer can inhibit PCR, so drying the paper after the wash step is required.

  9. While the disc is air-drying, add 30 µL of TE to a clean 1.5-mL tube labeled with the sample identification number.

  10. Once dry, carefully transfer the disc using a sterile tweezer or pipette tip into the fresh tube containing 30 µL of TE. Allow the disc to soak for a minimum of 15 minutes at ambient temperature (soaking the disc overnight at 4° C is optimal) to elute the purified DNA.

  11. The disc in TE can be stored at 4° C temporarily or frozen at -20° C for long-term storage until ready to begin Part III; ensure that the disc has incubated at ambient temperature for at least 15 minutes before storage at 4° C or -20° C. In Part III, you will use 2 μL of DNA for each PCR reaction. This is a crude DNA extract and contains nucleases that will eventually fragment the DNA at room temperature. Keep the sample cold to limit this activity.