1) Do your bench cleanup protocol.
2) Measure the amount of sediment you intend to process.
For small quantities of sediment (1-2 ml), use 10 ml. extraction buffer (recipe
below). For larger samples, use 5 volumes of extraction buffer per volume of
sediment.
3) Make up your extraction buffer.
It’s OK to make this up as a single batch for all of your samples, and
then divide it up later.
For every 10 ml. of extraction buffer you need, combine:
9.75 ml. lysis buffer from sterilized bottles (0.1M Tris-HCl, 0.1M EDTA, pH
8),
0.25 ml. SDS (10% stock: final concentration 0.25%)
10 µl Proteinase K (25 µg/µl stock in water, final conc. =0.25
mg/ml)
4) Defrost your sediment sample on ice.
As soon as the outside thaws a little bit, you should be able to remove the slug of sediment from the tube it was stored in.
5) Wet packed sediment with a minimal amount of extraction buffer. Make sure that the mixture is not too "watery"; this can significantly decrease the amount of cell lysis you get. Grind in sterile mortar and pestle for 1 minute.
Caution! Overgrinding can shear DNA. If the sediment is very coarse—sort of like thin gravel—grind for only about 5-10 seconds. When in doubt, press down hard with the mortar and pestle rather than use a “circular” grinding motion.
6) Add the rest of the extraction buffer; mix. Incubate at 55° C 1 hour,
with occasional shaking.
If you’re doing several tubes, it will make your life easier if you equalize
the amount of material in each tube. You can “top off” a tube with
some extra lysis buffer.
7) Add 0.83 g. sodium chloride to your sample for every 10 ml. of extraction buffer you added. (For example, you would add 20.75 g. to an extract of a 50-ml sediment sample.) Be sure to measure out the sodium chloride without contaminating it. Allow 10 minutes for the NaCl to dissolve in the buffer.
Do not forget this step; failure to add the salt now will ruin the prep.
8) Add 1 ml. CTAB stock for every 10 ml. of buffer and mix. Incubate 65° C 1 hour with occasional shaking.
(Stock: 10% CTAB (cetyltrimethylammonium bromide) in 1 M NaCl. Takes about 3 hours to dissolve; best to make the day before. Stock keeps for months at RT.)
9) Remove sediment and cell debris by centrifuging at 8,000 x g 10” RT. Draw off clear supernatant, bringing along as little of the floating layer of polysaccharides and detergent as possible. A “turkey baster” pipet works great.
10) Pour the supernatant through four layers of cheesecloth into a clean centrifuge tube. Add 1 volume isopropanol; allow to stand at RT 10”. Centrifuge at 12,000 x g 10”. Draw off supernatant and discard.
11) Wash the pellet with 5 ml. 70% ethanol; centrifuge at 12,000 x g 10”. Draw off supernatant and discard. Allow pellet to air dry, and resolubilize in 500 µl sterile TE (10 mM Tris, 10 mM EDTA, pH 8). Place the liquid in a microcentrifuge tube.
12) Mix with 1 volume phenol by vigorous shaking; centrifuge 12,000 x g 2”.
Draw off the top (aqueous) fraction and move it to a fresh tube. Take care to
avoid bringing along any glop that might be present at the interface. Repeat
with 1:1 phenol:chloroform/isoamyl, then with straight 24:1 chloroform:isoamyl
alcohol. Repeat until no solid precipitate is visible at the interface.
During the last extraction, be super-careful not to bring any of the organic
solvent along with the sample.
13) Add 100 µl 10M ammonium acetate to the cleaned aqueous fraction. Mix, then add 1 ml 100% ethanol. Allow to stand at RT 10”. Centrifuge at 12,000 x g 10”. Draw off supernatant and discard.
14)Wash pellet with 500 µl 70% ethanol and centrifuge 12,000 x g 5”. Draw off supernatant and allow pellet to air dry. Solubilize pellet in 50-100 µl sterile water.
Note: The DNA pellet may be difficult to see. These samples are at low concentration; the DNA may look like a glass-clear thin film on the walls of the tube. You may not be able to see it without practice. That’s OK. Have faith and continue.
If the sample is from a cell pellet or from deep-marine sediment, you may dilute your sample 1:10 and use for PCR. If the sample is from temperate sediments or from other sources contaminated with humic acids, or is unusually rich in polysaccharides, following the steps below is highly recommended. You can, if you choose, try PCRing the sample now; if the reaction fails, go ahead and do the rest of this prep. Steps 15-20 need not be done on the same day as steps 1-14.
15). Add appropriate amount of DNA sample buffer. (i.e., 1/5 as much buffer as there is sample.) Run sample on a 1% Tris-acetate agarose gel. (See Gel Protocol for details.)Visually inspect bands; if DNA is unsheared and of good quality, cut the genomic DNA bands out of the gel with a sterile razor blade.
Don’t use the same edge of the blade for more than one sample. If you do, you’ll introduce some DNA from the previous cut into the next sample. You can use one corner of the blade, which will let you excise two samples per blade. If you have run the same DNA sample in more than one lane, it’s OK to use the same blade for those bands.
16. Place gel slices in Eppendorf tubes and weigh. Add 3 volumes of NaI solution per volume of gel slice. Incubate at 70° C for 5 minutes, until gel slice is completely dissolved.
(1 g gel = 1 ml. Example: For a gel slice weighing 150 mg, add 450 µl NaI solution. NaI solution: 6 M NaI, 0.15 M Na2SO3. Don’t forget to correct for the weight of the microcentrifuge tube. They are almost always 1 g. in weight.)
17. Add 5 µl glass slurry. If unmelted bits of agar become visible against the glass, put the sample back into the hot bath; complete melting is essential. Incubate 20 minutes on ice with occasional vortexing.
18. Spin down glass (15 seconds in microfuge). Remove supernatant. Resuspend pellet in glass wash solution (50% EtOH, 0.1M NaCl, 10 mM Tris, pH 7.5, ice cold.) Spin down again and repeat.
19. Carefully remove wash solution from pellet. Add 50-100 µl sterile water. Heat at 70° C for 5 minutes to release DNA. Spin down glass; solution may be stored with glass or removed to a separate tube (preferred).
20. Dilute sample 1:20 for use as a PCR template.