DNA Extraction Protocols


I. HT Homogenizer Extraction Protocol

II. Large-scale DNA Extraction Protocol

III. Extract-N-Amp Protocol

I. HT Homogenizer Extraction Protocol

A. Materials & Reagents

1. Materials

  • a. Three 96-well tube boxes for each set of 96 samples
  • b. Two DNA storage tubes with caps, for each sample
  • c. 1 stainless steel bead per sample, to go in each grinding tube
  • d. Scissors and/or tweezers for collecting tissue
  • e. Plate map worksheet for recording sample ID of each well/tube
  • f. Ice bucket or Styrofoam container, with ice, dry ice, or liquid nitrogen
  • g. Large and small mortars, for weighing down tube boxes in water bath.
  • h. Paper towels
  • i. HT Homogenizer for grinding tissue
  • j. Water bath, set to 65˚C
  • k. Centrifuge, fitted with A-2-DWP swing bucket rotor for 96-well plates/boxes
  • l. Freezer, -20˚C
  • m. Incubator, set to 37˚C

2. Reagents

  • a. Ice, dry ice, or liquid nitrogen
  • b. Extraction Buffer
  • i. 100 mM Tris-HCl pH 8
  • ii. 50 mM EDTA pH 8
  • iii. 500 mM NaCl
  • iv. 1.25% SDS (w/v)
  • v. 8.3 mN NaOH
  • vi. 0.38 g Sodium bisulfate (per 100 mL of buffer)*
  • * add just prior to use
  • c. 24:1: Chloroform:Isoamyl alcohol
  • d. Isopropanol
  • e. 70% Ethanol v/v
  • f. TE buffer, ph 8
  • i. 10 mM Tris-HCl pH 8
  • ii. 1 mM EDTA
  • g. ddH2O

B. Procedure

1. Label a 96-well tube box for orientation purposes.

2. Put 96-well box containing one stainless steel bb in each tube in a box of ice, pushing the ice up around the sides to chill the tubes. (you can put it in a box with a shallow amount of liquid nitrogen or a box with some dry ice in it.)

3. Get a 96-well grid worksheet and orient it the same as your plate. Label as you fill the box.

4. Cut a 3-4cm sample of rice leaf and fold it in thirds than over again and slide it with forceps to about 5mm above the bb. Do no put the tissue in vertically (the bb will simply pass by it while shaking. The tissue must block the path of the bb, but not tightly.)

5. Put strip caps tightly onto the tubes and a paper towel folded in thirds on top of the caps, then cover with a lid. (Tape it tightly shut.)

6. Proceed to stage two or store at –20 until you are ready to grind tissue.

7. Put your plate(s) in a shallow container of liquid nitrogen (do not submerge the plate – there should be about 1” of nitrogen in the box).

8. Use an empty 96 well box and lid to adjust the tightness of the genogrinder while your box chills.

9. Put the box (with very tight caps and lid) onto the homogenizer and lock it in place.

10. Shut the lid of the machine and lock it. Turn the machine on for 2 minutes on setting 8.

11. Turn the machine off and remove the box. Check that grinding is complete. (note: if you try to grind it again, the box may crack!!)

12. If some of your samples did not grind, use a pair of metal forceps to crush the sample (you must re-freeze in nitrogen first). Be sure to clean the forceps between each sample to avoid contamination.

**note: it is rare to have every sample grind. There are usually one or two that you need to do manually. This is most likely a result of improper positioning of the tissue sample.**

13. Store the boxes in a freezer (-20c) until ready to extract.

**note: you must have two boxes at this point – separate your samples if you only have one box. (keep track of the order!) You must do this to be able to centrufuge (you may create a blank balance box if you wish, but it is usually more efficient to separate your samples).**

14. Centrifuge the boxes to bring tissue to the bottom of the tube (just for a minute or two on a moderate speed setting).

15. Mark the upper (or lower) edge of all of your tubes with a permanent marker and note which edge you marked – always do it the same way!!

*samples should be at –20, not liquid n frozen, when you add the buffer!!*


16. Peel the caps off carefully, one row at a time, being careful not to “throw” tissue out and contaminate other tubes. Set your caps aside in a bucket for washing.

17. Add 400ul of 65-degree pre-heated extraction buffer to each tube using a multichannel pipette or repeater pipette.

18. Place clean caps on the tubes, put a third-folded paper towel on top of them and replace the lid. Shake on vortex until most of the tissue is resuspended. *hold on to it, pressing the lid down hard*.

19. Put the boxes in the 65degree water bath, and immediately set a large mortar with the small mortar sitting inside of it on top of the plate. (the paper towel should still be on the caps – try not to let it dip into the water.)

*do not have more than about 1.5 inches of water in the bath*

20. Let the plate incubate for about 30 minutes. There is no need to shake the boxes during this time if you successfully resuspended the tissue with the vortex.

21. Take the boxes and mortars out together from the water bath and place on some paper towels. Let them sit for about 2 minutes then remove the small mortar and let them sit another 2 minutes. By then, the pressure in the tubes will have come down and you can easily remove your caps. Pull the caps from the side that is away from you, so they don’t spray you. Be careful of contamination and put the dirty caps in the bucket for washing.

22. In the fume hood, add 400ul of 24:1: chloroform:isoamyl alcohol to each tube.

23. Put new caps on the tubes. (very tightly!) Add a third-folded paper towel to each box and put the lid on. Tape them shut (very tightly!!)

24. Holding the two boxes tightly together, invert gently, in the fume hood, for 5 minutes. If you notice wet paper towels, tighten the caps and replace the towels.

25. Weigh the boxes and balance them by taring the heaviest and then bringing the partner up to zero with bb’s set in the caps (or with paper towels). Put 2 third-folded paper towels on top of each plate and replace the lid. Tape shut (check that this does not affect the balance of the boxes.) Centrifuge the plates at 3500rpm for 10 minutes.

26. In the hood, remove the caps carefully (so the spray will go away from you). Discard the caps into the bucket in the hood – these will not be re-used.

27. Carefully remove 250ul of the upper phase using a multichannel pipette with 300ul tips. Place the sample in a new 96-well box plate.

*be sure the new plate is in the same orientation as the original*

28. Mark the plate with a line, as you did at the beginning and label the box.

29. Add 2/3 volume of isopropanol (room temperature) to each tube to precipiate the dna. Cap the tubes. Balance by weighing as described above and put a paper towel and lid on. Invert gently until the DNA falls out of solution. You may leave them at –20 overnight (or for a while) or you may proceed directly to the next step.

30. Centrifuge the plates for 10 minutes at 3500rpm.

31. In the hood remove the caps carefully (away from you). Dump the liquid into a beaker, one row of tubes at a time, and tap the tubes on a paper towel gently. Watch that the pellet does not slide out. This works best if you do not hesitate when dumping the supernatant out. Replace the row of tubes back into the box in the correct orientation.

32. Add 200ul of room temperature 70% ethanol to each tube and put clean caps on again. Tap gently. Let sit at room temperature overnight (or for a while). You may proceed immediately if your samples look very clean and the purity of your samples is not absolutely critical.

33. Drain the ethanol off as before and tap the tubes to remove as much liquid as possible without dumping the pellet. You may centrifuge before this step, but it is often unnecessary, unless the pellets start to slide down the sides of the tubes while you are dumping. Put the tubes back into the boxes and put in the 37-degree incubator with no shaking to dry. (it is just blowing air.) This will take a few hours. Alternatively, you can leave them in the fume hood and turn the emergency vent button on (please mute it). This will up the air flow and help them dry faster, but this will be slower than the incubater.

34. When samples have no alcohol residue, add 50ul of te buffer for a concentrated stock or 500ul of te buffer for a working dilution. (they will resuspend quicker if you warm your te buffer to 65c.)

35. Store with caps on at –20 or 4 degrees celcius depending on your use.


II. Large-Scale Genomic DNA Extraction

A. Materials & Reagents

1. Materials

  • a. Brown paper envelopes
  • b. Liquid Nitrogen dewar or Styrofoam container
  • c. Mortar and pestle
  • d. 50 ml conical tubes
  • e. Brush (paintbrush style)
  • f. 80-tube storage racks
  • g. Glass Pasteur Pipette with hooked tip
  • h. Centrifuge
  • i. Pipettors, with tips, including wide-bore tips.
  • j. Water Bath
  • k. Refrigerator and Freezer (capable of -20˚C)

2. Reagents

  • a. Ice, dry ice, or liquid nitrogen
  • b. Extraction Buffer
  • i. 100 mM Tris-HCl pH 8
  • ii. 50 mM EDTA pH 8
  • iii. 500 mM NaCl
  • iv. 1.25% SDS (w/v)
  • v. 8.3 mN NaOH
  • vi. 0.38 g Sodium bisulfate (per 100 mL of buffer)*
  • * add just prior to use
  • c. 24:1: Chloroform:Isoamyl alcohol
  • d. Isopropanol
  • e. 70% Ethanol v/v
  • f. TE buffer, ph 8
  • i. 10 mM Tris-HCl pH 8
  • ii. 1 mM EDTA
  • g. ddH2O

B. Procedure

1. Harvest the tissue:

  • a. Harvest fresh leaf tissue (enough to make a ball the size of an egg).
  • b. Place it in a brown paper envelope and submerge it in liquid nitrogen.
  • c. This tissue may then be stored at -20°C or -80°C until needed., or you may proceed directly to the next step.

2. Chill a mortar either by placing it in the freezer for several hours prior to use or by setting it in a box of liquid nitrogen for about 5 minutes. (*Use caution when removing it from the box, as it will be extremely cold- you will need to pull it out with a tool of some sort.)

3. Grind the sample:

  • a. Pour some liquid nitrogen into the mortar and put one of the tissue samples into it.
  • McCouch Rice Lab DNA Extraction Protocols June 2, 2010
  • b. Grind the tissue, using a pestle, until it is a fine powdery consistency (all the nitrogen will evaporate and you need to continue grinding at that point to ge it fine enough.)
  • c. Transfer the ground tissue to a 50ml conical tube (that has been labeled and chilled) using a paintbrush.
  • d. Put about 15ml of ground tissue in each tube. Cap the tube and place in the nitrogen or in the freezer.
  • e. Quickly clean off the mortar and pestle with dry paper towels before they begin to thaw. If they do thaw, tissue will stick to them. If this occurs, you can get the tissue off using a piece of sandpaper.
  • f. Do not submerge the frozen mortar and pestle in water – it will break! You can proceed directly to the extraction, or store your ground tissue at -20 or -80C.

4. Add Sodium bisulfite to your extraction buffer and heat to 65°C in a water bath.

5. Add approximately 20 to 25 ml of extraction buffer to each tube of ground tissue.

  • a. This may be added using a re-pipet device or simply by measuring the buffer in a 50ml conical tube and pouring it into the tubes containing tissue. Exact measurement is not essential.
  • b. Cap the tubes and resuspend by tapping gently (or stirring with a clean metal spatual if necessary.) The tissue should be at -20C when you are ready to add the buffer (colder will make it very difficult to resuspend the tissue.)

6. Place the samples in a 65°C water bath for about 30 minutes. Mix occasionally by hand.

7. Remove samples from 65°C. Allow to cool down in a chemical fume hood for 10 to 15 minutes (removing the caps may assist in this).

8. Add about 15 ml chloroform:isoamyl alcohol, 24:1 to each sample. Close each tube making sure that the caps are on properly in order to avoid any leaking.

9. Place the tubes in a rack, then place another rack (or some other solid object) on top of them to secure the tubes. Gently invert the tubes and rock back and forth to mix. Do this for a full 5 minutes. All of this work should be done in the fume hood, as chloroform is a potential carcinogen.

10. Centrifuge samples in a table top centrifuge (about 2500 to 3000 rpm for 10 minutes).

11. Transfer upper aqueous phase to new 50 ml tube.

  • a. If the interface between the upper aqueous layer and the lower organic layer is stable, the upper layer may be transferred by decanting. If not, transfer by pipeting.
  • b. This work should also be done in a chemical fume hood as some chloroform can be detected in the aqueous layer.
  • c. OPTIONAL STEP: Add 10ul od RNase solution to your tubes, mix gently, and let them sit at room temperature for about 15 minutes. Proceed with next step.

12. Add 2/3 volume of isopropanol to each sample (e.g. if 15 ml of aqueous phase is transferred to a new tube, add 10 ml of isopropanol).

  • a. Mix well by inverting the tube several times.
  • McCouch Rice Lab DNA Extraction Protocols June 2, 2010
  • b. A whitish, stringy precipitate consisting of DNA and RNA should be visible at this point.
  • c. Samples may be stored either at 4°C or -20°C for a few hours or overnight in order to assist in the precipitation or as a stopping point in the protocol. Alternately, you may proceed directly to the next step.

13. Remove the precipitate.

  • a. If the precipitate can be hooked out by using a glass Pasteur pipette that has been shaped into a hook, proceed to Step 14.
  • b. If the precipitate does not float or form a cohesive mass, it may be collected by centrifugation as in step 10. Then proceed to Step 14.b.

14. Resuspend the DNA pellet.

  • a. Dip the DNA on on the hook into some cold 70% ethanol. It will stay on the hook. Remove it and blot it gently on a kimwipe. (Do this very carefully so the nucleic acid does not stick to the kimwipe!)
  • b. Place the nearly dry pellet into a 1.5ml tube containing 200-1000ul of TE buffer. (The amount of TE necessary depends on the size of your nucleic acid pellet.)
  • c. *NOTE: There cannot be ANY ethanol in the pellet or PCR reactions will be adversely affected. If necessary, let the pellet sit in the tube a while to dry before adding TE buffer. This is not typically necessary when using this method, as enough ethanol seems to be drawn out onto the kimwipe. Proceed to Step 15.
  • d. (from Step 13.b) Decant the liquid from the centrifuged tube and add several ml of cold 70% ethanol.
  • e. Tap gently and let this sit at room temp for a few minutes to overnight depending on your needs and time constraints. (If the DNA is really “dirty-looking” you may want to let it sit overnight, but this is not typically what we do.) Centrifuge the tube again at 2500 – 3000 rpm, but for only 5 minutes.
  • f. Decant the liquid and let all the ethanol evaporate. (You can let the tubes sit in the fume hood.) This will take a while! When the pellet is dry, add 200-1000ul of TE buffer (depending on the size of your pellet). Using a wide-bore tip, transfer the solution to a 1.5ml tube. Proceed to Step 15.

15. Resuspension may take several hours and you can simply leave the tubes in the refrigerator overnight if you have the time.

  • a. To help with reuspension of DNA, TE buffer can be heated to 65C before adding to the pellet.
  • b. Alternatively, the 1.5ml tubes containing the DNA can be warmed for a little while in a 65C water bath (do not submerge the tubes in the water,but set them above the water so the warm air is surrounding them) for 15-30 minutes (use as little time as possible to maintain integrity of DNA).
  • Note: If you have added more TE and no more of your pellet seems to be going into solution:
  • Some samples may contain large amounts of polysaccharides that will not dissolve and may give the appearance that the DNA pellet has not gone into solution. In these cases, the samples should be centrifuged at maximum speed (13,000rpm) in a microcentrifuge for 10 minutes and the supernatant which contains the DNA should be transferred to a clean tube.

III. Extract-N-Amp Protocol

A. Materials & Reagents

1. Materials

  • a. 96-well microtitre plate and seals
  • b. Tweezers and scissors for tissue collection
  • c. Ice bucket or Styrofoam box
  • d. Thermal cycler
  • e. Centrifuge with microtitre plate rotor and adaptor
  • f. -20 ˚C Freezer
  • g. Solution troughs

2. Reagents

  • a. Ice
  • b. Deionized, distilled water (ddH2O)
  • c. Extract-N-Amp Extraction Buffer
  • d. Extract-N-Amp Dilution Buffer

B. Procedure

1. Collect ~ 1 mm2-sized tissue samples into a 96-well microtitre plate on ice.

2. Store sealed plate of tissue samples at -20 ˚C until extraction.

3. Add 10 μL of Extract-N-Amp Extraction Buffer to each well of tissue plate, ensuring each sample is at least partially submerged in this buffer. Vortex briefly, centrifuge for 10 sec at 3000 rpm to submerge tissue sample in buffer, if necessary.

4. Run sealed plate through a brief incubation program on a thermal cycler, at 94˚C, for 10 minutes.

5. Add 10 μL of Extract-N-Amp Dilution Buffer. Vortex briefly, then centrifuge

6. Add 80 μL of ddH2O (making sample DNA concentration 1:4 relative to initially extracted sample DNA).