DNA EXTRACTION The “mini-extraction” procedure detailed below takes ca. 4 hours TOTAL from start to finish for 24 samples if the major steps of the extraction process are not interrupted. It is best to do the first 2 hrs, steps 1-13 under "Procedure", then leave tubes in freezer overnight or longer before completing the extraction protocol. SETUP Before you begin, do the following. 1. Turn on the hot-water bath to 37-40C 30-45 min beforehand. 2. Replace the lab benchtop “diaper” paper to help prevent contamination. 3. Label 2 sets of 1.5 ml plastic tubes for your samples; put one set in front of you and the second set in a separate rack away from your immediate work area. 4. Set out the 40-200µl pipettor with unfiltered 200µl (yellow) tips, the 200-1000µl pipettor with unfiltered 1000µl (blue) tips, and the 0.5-10l pipettor with filtered 10l tips. 5. Set an empty rack in front of you. 6. Get your leaf tissue samples for extraction ready and arrange them in precisely the sequential order of the labeled tubes. 7. Fill a 500ml beaker with water and a little Alconox to make a soapy solution, and place this in the back near your work area, to put used pestles. 8. Set the wastebasket next to you, to easily discard pipette tips as you work. 9. Immediately before you begin, get a small glass watch-glass to use as a “dipper”, and a small amount of liquid nitrogen in a styrofoam box or insulated thermos with sides easily allowing you to pull out the “fluid”. PROCEDURE The DNA extraction procedure follows; points at which you can leave samples in the refrigerator-freezer for 1 or more days before returning to finish up the extraction procedure are marked with an asterisk (*). 1. Remove a 1.5 labeled tube to the empty “working” rack and place ca. 1 lid-worth (approximately 1-2 paper punch-sized pieces) of leaf tissue into it with forceps. 2. Immediately pour a partially filled watch-glass “dipper”-ful of liquid nitrogen into the tube; when the hissing begins to subside, gently insert a clean pestle and firmly but quickly grind the tissue to small bits 3. Immediately pipette 400l of SDS buffer into the tube and grind again briefly to mix tissue with buffer; set tube (with pestle still stuck in it) onto the rack of leaf samples. 4. Repeat steps 1-3 until all samples have been frozen and ground in liquid nitrogen, and ground and mixed thoroughly in SDS buffer, and are now back in the original rack(s). 5. Gently twist pestles once more in each tube, and remove pestles to beaker with soapy water; pipette 200l more SDS buffer into each tube. 6. Momentarily vortex at medium speed to mix tissue with SDS buffer solution. 7. Soak tubes in floating rack in the hot-water bath for 15-30min; set aside styrofoam box with unused liquid nitrogen, to allow the latter to evaporate spontaneously. 8. Remove the tubes to a plastic rack and take them to the fume hood. 9. Under the hood, open the tubes and pipette 400µl 24:1 chloroform-isoamyl alcohol mixture into each one.

10. Still under the hood, invert each tube twice carefully, lift the cap (holding the tube away from the other samples in case contents spill out!) to remove initial pressure, then shake tube vigorously to form milky emulsion, finally lifting the cap again (slowly!) to expel pressure. 11. Close the lids on the tubes, and centrifuge in the Marathon 24-well microcentrifuge at 13,000 rpm for 5 min (leave other tubes not yet centrifuged under the hood); immediately and gently remove samples from microcentrifuge to their rack, to avoid remixing the aqueous upper layer with the particulate layer below it. 12. Bring 2nd set of labeled 1.5ml tubes to hood and set beside the rack of centrifuged tubes; transfer 400µl of clear upper aqueous phase (avoid particles!) to second set of tubes, pouring off the (hazardous!) halogenated solution into the brown toxic waste bottle marked “halogenated materials” in the hood and putting the used pipette tips and old 1.5ml tubes into the bag next to the bottle. 13. Pipette 400µl ice-cold isopropanol from freezer into each tube, close the lids and invert gently 10-20 times to mix [you may observe clouds or “threads” of DNA precipitating as you do this]; freeze tubes for 20min to 1hr in the refrigerator-freezer to precipitate DNA. This is a good time to end the process; leave tubes in freezer overnight to greatly increase yield! 14. Centrifuge at 13,000 rpm for 5min, open the lids, and pour off supernatant into sink, taking care not to lose the pellet. 15. Pipette 100µl TE buffer from refrigerator into each tube, close lids and invert tubes gently to dissolve DNA pellet (soaking for 5min in hot-water bath will facilitate this). Use clean spatula to break up pellet gently if necessary. 16. Open lids and pipette 50µl 7.5M ammonium acetate, 10l 3M sodium acetate, and 300µl 95% ethanol into each tube; close lids and invert gently several times to mix; freeze for 20min to 1hr in refrigerator-freezer [can leave the process at this point for several hours or days if necessary--but label your tube rack with tape]. 17. Centrifuge at 13,000 rpm for 5min, open the lids, and pour off supernatant into sink. 18. Pipette 100µl 70% ethanol into each tube; let tubes sit undisturbed for 5 min, then gently pour off ethanol into the sink without losing pellet in bottom of tubes; tap mouths of tubes gently upside down on paper towel to remove excess alcohol. 19. Vacuum-dry tubes with lids open for 20min. 20. Pipette 100µl TE buffer from refrigerator into each tube, close lids and invert tubes gently to dissolve DNA pellet (soaking for 5min in hot-water bath will facilitate this). Use clean spatula to break up pellet gently if necessary (it may not go much into solution, but don't worry; freeze-thaw cycles will do this eventually). NOTES  For older herbarium material, storing tubes in a refrigerator-freezer for 3-5 days in isopropanol at step 16 improves DNA precipitation and, consequently, final DNA yield.  You should try wherever possible to restrict the leaf or other tissue for each extraction to tissue from a single individual plant, such as a piece of one leaf, so that the extract may be useful in the future as representative of a single individual rather than as a bulk population sample  For fibrous leaf tissue, gymnosperm needles and graminoid/cyperoid tissue, some folks add a pinch of clean white sand to the tube at the very first step and grind the leaf tissue with that, immediately followed by liquid nitrogen or buffer.

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