PREPARATION OF HIGHLY PURIFIED PLANT DNA
Collection of Plant Tissue
1. Cut off leaf tissue (with scissors). Avoiding collecting
yellowed tissue or petioles, if possible.
2. Wrap tissue loosely in cheesecloth, tying with string, and
attach cold-resistant label. Tape will crack and fall off at liquid
nitrogen temperatures, a successful method is to punch a hole
in a thick paper label, and pass the string through the hole.
3. Submerge in liquid nitrogen. From this point on, thawing
must be avoided. Store in a styrofoam cooler with liquid nitrogen
until transferred either to an ultracold freezer (-70°C),
or place directly in freeze drier.
4. Freeze dry (usually about 3 days), taking care not to let
tissue thaw. Transfer tissue from cooler or ultracold freezer
directly to freeze drier chamber, don't let tissue touch the warm
metal sides, and start freeze drier immediately. If tissue is
handled correctly, dry tissue is whitish-green colored. If tissue
has thawed in handling, it will be darker green on the thawed
edges. If this occurs, throw the affected tissue away, it is ruined.
DNA Extraction Protocol
1. Crush 2 grams (or more) of freeze-dried plant tissue
with a small amount of sterile silica sand in an acid washed and
sterilized mortar and pestle. Grind as finely as possible.
2. Add crushed samples to 50 ml or 250 ml Oakridge tubes and
add an appropriate volume of sterile extraction buffer (2X CTAB)
up to as much as half the tube volume.
3. Mix tissue in buffer until thoroughly wet. Final product
should not be highly viscous.
4. Incubate at 50°C for 45 minutes, mix gently (so as not
to break DNA) about 10 every minutes.
5. Add equal volume of chloroform:isoamyl alcohol (24:1) and
incubate for 45 minutes at 50°C. Mix gently about every 10
minutes.
6. Balance the tubes to within 1% and centrifuge for 10 minutes
at 8000 rpm (50 ml tubes), or for 10 minutes at 6000
rpm. (250 ml tubes )
7. Using inverted 10 ml pipette (to provide large orifice),
remove supernatant and place in a new acid-washed tube. Remove
only the top phase.
8. Add an equal volume of ice-cold isopropanol, or twice the
volume of 95% ethanol. Invert gently until two phases are no longer
evident. DNA will collect as white stringy mass. Sample may
be stored overnight at - 20°C at this point.
9. Close and hook the end of a pasteur pipette and gently wind
DNA onto end. Let liquid drain off and put into clean sterile
tube containing 76% ethanol-10 mM ammonium acetate. Let stand
for at least 20 minutes. DNA can be left on the pipette.
10. Remove DNA & pipette. Let liquid drain off (until almost
dry & odor of ethanol is no longer evident) and dissolve in
4.0 ml of 0.10X SSC in 15 ml disposable plastic test-tubes. If
suspension is highly viscous add 0.10X SSC in multiples of 4 ml.
It is important to dissolve the DNA completely.
11. For every ml of SSC + DNA, add 1.1 g of CsCl. Add 5 µl
of ethidium bromide (5 mg/ml) for each gram of CsCl.
12. Use a pasteur pipette to slowly draw up the mixture and
place into Beckman ultra-centrifuge tubes. Seal tubes.
13. Balance the tubes to within 1% and run on Beckmann ultra-centrifuge
at 55,000 rpm, 20°C for at least 6 hours.
14. DNA bands will be visible under UV light. Remove the DNA
with an 18 gauge needle on a 1 cc syringe.
15. Place DNA into 15 ml tubes and add 2 to 10 volumes of isoamyl
or isopropyl alcohol saturated with 20X SSC.
16. Mix gently on platform shaker. As ethidium bromide migrates
into top layer discard alcohol and add more. Repeat this procedure
until no more ethidium bromide moves into the alcohol layer (3
changes of 10 volumes over 2 days is usually sufficient for moderate
to large amounts of DNA).
17. To dialyze out the CsCl, remove DNA solution with pasteur
pipette and load into dialysis tubing. Place dialysis tubes in
a beaker and add 2000 ml of TE (pH 8.0). Place in cold room and
change buffer twice during a minimum of 6 hours.
18. Pipette DNA into sterile microfuge tubes. Determine the concentration of DNA solution using UV spectrophotometer.