This protocol is from USU Molecular Ecology Lab created by Jim Walton in Dr. Mock's lab.
1. Prior to starting:
a. Tungsten Carbide Bead Cleaning:
i. Seal the collection microtubes with caps. Place clear cover over rack and knock upside against the bench to free tungsten carbide beads from surrounding material. (May need to loosen bead with paperclip or other sharp object if pellet is allowed to dry).
ii. Empty contents into a sieve and rinse the beads thoroughly with water.
iii. Incubate beads in 0.4 M HCl (I used EtOH) for 1 min. at room temperature to degrade DNA and avoid future cross-contamination.
iv. Rinse with distilled water to remove HCl. Dry beads with paper towel.
b. Add 96-100% EtOH to Buffer AW2 and Buffer AW1 if not already added. See bottle for directions.
c. If precipitates form in Buffer AW1 warm to 65C to redisolve. Do not reheat after adding EtOH.
d. Preheat Buffer AP1 to 65C.
2. Add one tungsten carbide bead to each collection microtube.
3. Remove a piece of tissue a bit smaller than your index fingernail (~ 9mm x 5mm or 5-10mg). Cut into srips perpendicular to veinage (~2mm wide)
in each tube of 2 microtube racks.
4. Packing list: load box with everything listed
2 stuffed extraction plates (verify tight caps and bead is in each tube).
protocol
50 mL centrifuge tube
1 fine point sharpie and 1 regular point sharpie
2 elution microtube racks (dark blue plates)
2 clean S-blocks
2 light blue racks
1 pkg. air pore tape sheets
1 pkg. rubber caps (for elution microtubes)
1 pkg. collection microtube caps
reagent Dx
RNaseA
AP1
P3
Buffer AW1 (with EtOH added)
Buffer AW2 (with EtOH added)
Buffer AE
2 DNeasy plates
5 300 uL rack of tips
2 1500 uL rack of tips
bag of gloves
paper towels
box kimwipes
2 ziploc bags (0.5 gallon for 65C H2O bath)
5 reagent basin (one/each reagent added)
transfer pipette (when you overfill 50mL tube with reagent)
pippetes (1500uL and 300uL multichannel at F&R range can be used)
5. Go to forest and range research lab
a. turn on water bath as soon as you arrive to 65C. Place reagent Dx and Buffer AP1 in bath as soon as warmed.
6. Place plate(s) between adapter plates and place in tissue lyser. Grind 1.5 minutes at 25 Hz. (My samples were at 30 Hz).
7. Reassemble the racks so that the collection microtubes nearest the tissue lyser in step 8 is now furthest from the tissue lyser (flip the cover plates).
8. Grind for another 1.5 minutes at 25 Hz.
9. Remove plate sandwiches and adapter plates from each rack of collection microtubes.
10. After reagent Dx and Buffer AP1 is warmed make Working Lysis Solution: vortex until completely mixed. Make for 1 plate at a time and add to reagent basin, go to step 4 after both lysis solution are made. Flush RNase A and Reagent Dx when mixing.
Table 1. Working Lysis Solution
11. Add 400 uL of working lysis solution into each collection microtube. Seal microtubes with new caps, ensure caps are properly sealed.
12. Shake until all plant material is wet (1 min.).
13. Centrifuge, allowing to reach 3,000 rmp then stop the centrifuge (pulse).
14. Incubate at 65C in a ziploc bag for 10-15 minutes (ensure bags are fully sealed).
15. Centrifuge, allowing to reach 3,000 rpm then stop centrifuge (pulse).
16. Remove and discard caps. Add 130 uL Buffer P3 to each collection microtube.
17. Seal microtubes with new caps. Place cover on rack and shake vigorously for 15 seconds.
18. Centrifuge, allowing to reach 3,000 rpm then stop the centrifuge.
19. Incubate for 10 minutes at -20C. Aids in precipitation of proteins and downstream inhibitors.
20. Centrifuge for 5 minutes at 6,000 rpm. Compact pellets may form.
21. While in centrifuge label the new microcentrifuge collection tubes, DNeasy 96 plates, and elution microtubes RS.
22. Remove and discard caps. Transfer 400 uL of each supernatant to new racks of collection microtubes (careful not to transfer any floating pellets from previous step).
a. Pellets contain carbide bead which can be cleaned and reused.
b. Do not transfer >400 uL of the supernatant or 96 well plates and S-blocks capacity will exceed.
c. If less than 400 uL of supernatant is recovered adjust volume of Buffer AW1 accordingly in next step.
23. Add 1.5 volumes (typically 600 uL) of Buffer AW1 to each sample. White precipitate may form.
24. Close microtubes with new caps. Shake vigorously for 15 seconds, up and down with both hands.
25. Centrifuge, allowing to reach 3,000 rpm then stopping the centrifuge.
26. Place DNeasy 96 plates on top of S-blocks.
27. Remove and discard caps from the collection microtubes. Transfer 1 mL of each sample to each well of the DNeasy 96 plates.
a. Do not get sample on rim of DNeasy 96 plates.
b. Do not lower pipette tip to bottom of well, it may cause overflow.
c. Only remove one row of well caps at a time to avoid contamination of other wells.
28. Seal DNeasy 96 plate with AirPore Tape Sheets. Centrifuge for 4 minutes at 6,000 rpm (fold down edges of AirPore Tape).
a. Check wells in plate, if lysate remains centrifuge for another 4 minutes.
b. Discard flow-through in S-blocks.
29. Remove and discard AirPore Tape. Add 800 uL of Buffer AW2 (400 uL x2) to each sample.
30. Centrifuge for 15 minutes at 6,000 rpm to dry the DNeasy membrane. Reseal during centrifuge, remove tape and centrifuge again to aid in drying is necessary. Residual ethanol in filter will result in downstream inhibition.
31. Place DNeasy 96 plate in correct orientation on a new rack of elution microtubes RS add 100 uL Buffer AE.
32. Seal with AirPore Sealing Tape.
33. Incubate 1 minute at room temperature (12-25C). Centrifuge for 2 minutes at 6,000 rpm.
34. Place caps on the Elution microtubes RS.
35. Clean up.
a. Rinse everything with DI water and soak in water bath with bleach for ~15 minutes.
36. Store samples in -20C freezer.