Anolis

Isolating Anolis DNA (Phenol-Chlorofom-Isoamyl-Alcohol Based Protocol)

Based off of the original protocol for extraction of Stickleback DNA. Modified by Tim for use with Anolis tail tips- April 2016

Day 1: Tissue collection and digestion

Materials

• 1.5 ml Eppendorf tubes (1 per specimen)

• razor blades (1 per specimen)

• small tweezers (for handling tail tips)

• large tweezers (for handling razor blades)

• aluminum foil

• Kimwipes

• Bunsen burner

Reagents

• 600 μl of tissue digestion buffer*1 per sample

• 10 μl proteinase K2 per sample

1. Preparation:

   1. Turn on water bath, make sure it is set to 55ºC.

   2. Aliquot 600 μl of tissue digestion buffer*1 into appropriate number of 1.5 ml tubes (1 tube per specimen). Label both the top and side of the tube with the tissue sample #.

   3. Ignite Bunsen burner to a low flame.

   Warning

   REMOVE GLOVES when igniting Bunsen burner. GLOVES ARE FLAMMABLE.

   4. Flame small tweezers with ethanol (be careful not to use too much), hold razor blade in large tweezers and heat briefly in flame of Bunsen burner.

2. Use tweezers to remove tail tip from storage solution (usually 100% ethanol), place onto a fresh Kimwipe. Blot away all excess ethanol, ethanol may interfere with the digestion process.

3. Place tail tip on a small square of aluminum foil. Record total size and width of tail tip (estimate, no need to actually measure). Use razor blade to cut a piece of tissue from the proximal (thicker) end of the tail tip. We’d like to ensure that tail tips can be extracted multiple times, in case one extraction fails. So, for smaller tail tips, cut away ~2-2.5 mm. On larger tail tips, take ~4.5-5mm. Return the rest of the tail tip to the sample tube.

4. Use the razor blade to cut the sample into smaller pieces, about ~1mm in length (there’s no need to chop them super-small, and in doing so you may lose bits of tissue). Place the chopped-up sample into appropriate tube with 600 μl of tissue digestion buffer.

5. Repeat steps 2 through # for all samples. Use a new razor blade, kimwipe, and square of aluminum foil for every sample. Flame small tweezers with ethanol between samples to avoid DNA cross-contamination.

6. Add 10 μl of proteinase K2 to each sample tube.

7. Vortex briefly, place tubes in 55ºC water bath (record time), digest overnight.

Day 2: DNA Isolation – Step 1

Materials

• 1.5 ml Eppendorf tubes (1 per specimen)

• timer

Reagents

• 600 μl of 25:24:1 phenol:chloroform:isoamyl alcohol solution3 per sample

• 900 μl of COLD (-20°C) 100% ethanol per sample.

1. Remove samples from water bath (record time). Vortex briefly. Remember to turn off water bath.

2. Take samples to hood. Turn on hood fan and light. Add 600 µl of 25:24:1 phenol:chloroform:isoamyl alcohol solution3 to each tube (this solution is highly toxic and tends to drip from the pipette, so handle with care).

3. Shake in hood for ~ 3 minutes until completely mixed (hold tops of tubes, otherwise they may pop open!). Solution should look milky white (it is important to mix completely).

4. 1. Spin 10 minutes at room temperature at maximum speed 15,000 rpm.

   2. While samples are spinning, label (top and side) a new 1.5 ml Eppendorf tube for each sample, and fill each tube with 900µl of COLD (-20°c) 100% ETOH. Places these tubes in the -20 freezer while centrifuge finishes.

   3. When spinning is complete, gently remove tubes from centrifuge and take them to hood.

5. 1. In hood, carefully remove the top layer into the new tube filled with 900µl of COLD (-20°c) 100% ETOH. Use P200 to remove top layer. You should be able to get ~ 550µl max. Discard the lower layer in the hazardous waste receptacle in the hood.

   2. This is a good time to change gloves and tube rack.

6. Mix by inverting several times.

7. Place at -20°c overnight to let DNA precipitate, ideally for 24 hours. Note time samples were placed in the freezer.

Day 3: DNA Isolation – Step 2

Materials

• timer

Reagents

• 500 μl of freshly prepared 70% ethanol per sample, cold (place in freezer before starting)

• 100 μl of TE6 per sample

1. Prepare 70% ethanol, place in freezer (-20). Wait about 15-20 minute before proceeding to step 2, so that ethanol has time to cool down.

2. Remove samples from freezer, noting time.

3. Spin 10 minutes at room temperature at 12,100 rpm. Place tubes into centrifuge in such a way that you know where the pellet will form (e.g., with the cap hinge pointed up).

4. Carefully remove the tubes, check each for a gray/white pellet at the bottom of the tube (you may not see a pellet). Remove ethanol without disturbing the pellet, using a P200. If you placed your tubes in the centrifuge properly, you’ll know where the pellet is, even if you can’t see it. For this step, it is better to err on the side of leaving a little ethanol, rather than disturbing the pellet.

5. Add 500 μl of freshly prepared 70% (at -20) ethanol5.

6. Spin 5 minutes at room temperature at 12,100 rpm. Carefully remove the tubes, checking for pellets (more likely to see them on this step).

7. Remove 70% ethanol carefully with a pipette. Use a P200 at first, and switch to a P10 for the final little bit. Once again, don’t disturb the pellet.

8. Spin briefly (hold the “short” button on the centrifuge until the RPMs reach 12100, about 20 seconds). Then, use a P10 to remove as much of the residual ethanol as possible (otherwise ethanol may interfere with PCR).

9. Dry pellet for 15 minutes at room temperature. To do this, invert tube on the white racks. Can dry for longer, if there’s still residual ethanol after 15 minutes.

10. Resuspend the DNA (pipette up and down several times to dissolve the pellet) in 100 μl of TE6, this is concentrated stock (store at -80ºC if needed). Make sure the pellet dissolves. May help to vortex briefly.

11. Quantify with Nanoquant or PicoGreen for dsDNA ratio.

12. Working stock for PCR reactions is 1:25 dilution- 5 μl concentrated stock + 120 μl H2O. Use PCR tube racks and store at -20ºC. Concentrated stock is ideal for ddRAD library prep.

Solutions

1. 1. Tissue digestion Buffer:

      • 10 mM Tris, pH 8.0

      • 100 mM NaCl

      • 10 mM EDTA

      • 0.5% SDS

   2. Combine the following to make 500 ml:

      • dH2O | 450 ml

      • 1 M Tris, pH 8.0 | 5 ml

      • 5 M NaCl | 10 ml

      • 0.5 M EDTA | 10 ml

      • 10% SDS | 25 ml

   3. Notes on making stock solutions:

   Reagent	MW/FW	M	For 250 mL	Comments
   SDS				(Comes in 20% Solution)
   Tris	121.1	1	30.275 g	Adjust pH with HCl
   NaCl	58.44	5	73.05 g
   EDTA*	292.25	0.5	36.53 g	*Will not dissolve if pH is below 8. Adjust pH with Sodium hydroxide pellets as you mix on stirring plate

2. Proteinase K (20 mg/ml, comes in solution)

3. 25:24:1 Phenol:chloroform:isoamyl:alcohol solution (or 1:1 phenol chloroform solution).

4. 100% ethanol

5. 70% ethanol

6. TE: Tris-EDTA (10 mM Tris pH 8, 1 mM EDTA pH8). To make 250 ml combine 2.5 ml 1 M Tris, 0.5 ml of 0.5M EDTA, and 247 ml H2O.