DNA extraction | CTAB-chloroform | 96 wells plate

Put the samples to be freeze-dried at -80 and let them sit for at least one hour.

Turn on the freeze dryer and the blue pump attached to the side. The freeze dryer will take 30 minutes to reach the proper conditions.

When the machine is ready, it should say "Load/Unload" in the top left corner, load the samples that have been stored at -80C. Click "Start" and the machine. Let the machine run for 48 hours.

After 48 hours, click "Aerate" to stop the machine and remove the red cap from the tube on the left side to drain the moisture collected. Unload the samples.

Pre-label the tubes

Pre-load cleaned and autoclaved beads into plate of tubes (could use a bead dispenser if available).

Add 10mg of leaves (max. 30 mg) to each tube. Try to be consistent to have a similar yield for each extraction.

Put the samples in the tissue lyser for 2 min at a frequency of 30 (1/s).

Centrifuge the samples for 2 min @2000rpm to get powder off caps and down into bottom of the tubes.

Turn on water bath in fume hood (60°C) and clear workspace in hood.

Make sure you have sufficient CTAB buffer and reagents/supplies for all steps before starting.

Glove up.

In hood, measure out 600 µl of CTAB extraction buffer per sample (+8-10%) into a wide trough (or clean tipbox lid)

Add 1 µl b-mercaptoethanol per 600 µl of CTAB extraction buffer, and stir to mix.

Add 4µl of RNAseA per sample.

Uncap the tubes with the ground samples VERY CAREFULLY/SLOWLY and discard cap strips immediately into waste bin (they are a contamination risk).

Add 600 µl of CTAB buffer (including β-mercaptoethanol and RNAseA) to each sample using multichannel pipettor.

Re-cap tightly with FRESH caps.

Use the vortex or the tissue lyser to homogenize the sample in the buffer. Make sure there is no powder left at the bottom of the tube.

I ncubate tubes for ~20 min. in 60°C water bath.

Meanwhile, label next set(s) of plates/tube-strips to match your collection plates.

Post-incubation, remove plates from the water bath and blot them off. Re-tighten caps.

Centrifuge plates for 2 min to pellet solids @4000 rpm.

Transfer all the supernatant possible to a new labelled 96-well Costar plate using multi-channel pipettor.

Decant chloroform into large dedicated trough (tipbox lid or other HARD plastic).

Add 500 µl of chloroform (equal volume) to each sample tube.

Re-cap samples (tightly!) with fresh caps.

Hand-invert for 5 minutes using a paper towel over the lids to catch leaking from the caps.

Centrifuge for 15 min @ 4000 rpm to separate aqueous (upper) and chloroform (lower) layers.

While you are waiting on the spin:

Fully label a fresh plate of strip tubes -- these will be final home of samples.

Arrange the handy-dandy-cut-down-rack for interface-viewing/pipetting (center front) and your freshly labeled Costar plate of tubes (center-back). Put an empty Costar base for waste tubes (to the left, if right-handed).

Set pipettor to 300 mL and SLOW speed (if automated), and get a fresh box of tips for each plate.

Bring plate of centrifuged samples to hood. Place on right (if right-handed).

One strip at a time, put a sample strip and the corresponding empty strip in the handy-dandy-cut-down-rack.

Un-cap sample (can do before, during, or after moving, depending on preference).

Carefully pipet off the aqueous layer (up to 300mL) of sample strip and transfer it the matched empty strip.

Move chloroform waste to left waste plate and return filled strip to new base.

Repeat until all samples are transfered.

[Out of hood if you like.]

Add 1.5 volumes of cold 100% isopropanol to each aqueous sample, cap well with fresh strip-caps, and mix by hand- inverting plate once or twice.

(e.g. for a sample of 200uL add 300uL of isopropanol)

Place in -20 freezer for at least 30 minutes or as long as overnight (we generally do the latter).

Centrifuge for 15 min. at 4000 rpm.

A greyish gelatinous pellet is often visible in the bottom of the tube (but may not be).

Un-cap. To re-use caps (optional): lay them in order on a trifold towel labelled with plate #, NOT touching.

One strip at a time, gently tilt to pour off isopropanol supernatant (into a dedicated waste tipbox lid).

While each strip is still tilted but mostly empty, blot the open ends with a fresh bit of trifold towel to wick out as much liquid as possible.

Add 200 µl cold 70% ethanol (mol. grade) to each sample.

Flick to rinse pellet. It should float up off the bottom of the tube.

(OR can invert to really get rid of isopropanol on walls. Cap if so)

Centrifuge at 4500rpm for 5 min (no need to re-cap yet, but you can)

Pour off ethanol as with isopropanol, wicking out as much as possible with paper towel at the end of the pour.

Air-dry the pellet for 1-2 hours at room temp. with a Kimwipe placed over the open tubes. If you have to leave for longer, re-cap loosely and dry later. Don't overdry.

Once pellets are dry (no ethanol smell), add 50μL TE buffer to each sample.

Cap with retained or new caps.

Flick tube-strips to mix well and/or leave out at room temperature for at least an hour to fully re-suspend.

Refrigerate if diluting/using wihtin a few weeks, otherwise store in -20 or -80. (Do not freeze before pellet has fully resuspended!)