AFLP
Restriction Digestion of Genomic DNA 1. Make a master mix as follows:
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| 5 Reaction buffer |
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| EcoR I + Mse I (1.25 u each/ ml) |
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| AFLP-grade H2O |
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| Adapter ligation solution |
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| T4 DNA Ligase |
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2. To the digestion (6.25 ml), add 6.25 ml of master mix. Incubate at 20°C ± 2° for 2 hours.
3. Dilute reaction 10_: take 2.5 ml of reaction, and add 22.5
ml of TE. Store diluted ligation mix, and remaining undiluted
ligation mix at -20°C.
Preamplification
1. Make a master mix as follows:
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| Preamplification primer |
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| 10_ PCR buffer |
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| Taq polymerase (5 u/ml) |
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| Water |
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2. Place 0.5 ml of diluted ligation mixture into a PCR tube. Add 4.6 ml of master mix. Run a PCR program of 20 cycles of [94°C for 30 seconds, 56°C for 1 minute, 72°C for 1 minute].
3. Dilute entire reaction by 50_ by adding 249.9 ml of TE,
then transfer to a 0.5 ml tube for storage at -20°C.
Primer Labelling
1. Make a primer labelling reaction as follows:
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| EcoR I primer |
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| Water |
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| Kinase buffer |
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| 33P |
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| PNKinase |
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2. Incubate at 37°C for 1 hour, then heat inactivate at
70°C for 10 minutes.
Selective Amplifications
1. Make mix "A" as follows:
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| Water |
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| PCR buffer |
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| Taq polymerase (5 u/ml) |
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2. Using the tube of 33P-labelled primer, make mix "B" as follows:
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| Labelled EcoRI Primer |
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| 10_ PCR buffer |
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| Total volume |
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3. Into each PCR tube, add 2.5 ml of mix "A". Next add 1.25 ml of diluted pre-amplified DNA. Finally add 1.25 ml of mix "B". Cover, hand-centrifuge, and wipe caps with damp towel to reduce static electricity. Run with PCR program #27.
4. Add an equal volume (5 ml) of formamide buffer.
Formamide buffer To make 200 ml:
deionized formamide 10 ml xylene cyanol FF 10 mg bromophenol blue 10 mg 0.5 M EDTA 200 ml pH 8.0
Preparing an Acrylamide gel 1. Set up work area: White absorbent paper, spill catch tray, plate support (hunk of styrofoam).
2. About every 10 runs, the long plate is cleaned with 2N NaOH, and the short plate is treated with Rain-X.
3. In sink, wash the long glass plate with Alconox and water, using just a gloved hand to rub. Rinse with tap distilled water. Dry the non-gel side with a paper towel, and set on styrofoam support. Dry the gel side with Kimwipes. Rinse with some 95% ethanol (pure grade from glass bottles) and some water, and dry with Kimwipes. Rinse again with just 95% ethanol, and dry again.
4. Repeat procedure with short plate.
5. Place spacers on long plate, and place short plate on top. Leave no gap between the thick spacer part and the top plate.
6. Place 7 binder clips on each side, centering the clip over the spacer. Binder clips are right next to each other.
7. Place a 3rd spacer at the bottom, and add 6 more binder clips.
8. For a 60 ml gel, mix 12 ml 5X TBE and 13 ml dH2O. Place on hot plate, warm the solution, and add 27 g urea and a stir bar. Stir until dissolved.
9. Bring to 51 ml volume with dH2O.
10. Filter through a 0.45 mm 25 mm diameter membrane filter using the 60 ml "Urea" syringe. Filter directly into the acrylamide squirt bottle.
11. Add 9 ml of 40% acrylamide stock.
12. When everything is ready, add 320 ml of 10% APS and 30 ml TEMED. Swirl to mix, and immediately pour gel. (APS amount is increased gradually as it gets older)
13. Insert combs until orange part is even with top of plate. Cover with extra acrylamide solution.
14. Add 2 binder clips, one over each half of comb. Tilt gel to 15° angle, and allow to polymerize for 1 to 1.5 hours.
15. Pour excess acrylamide solution into marked waste bottle, and clean acrylamide squirt bottle immediately.
16. Gel may be left overnight. Add moistened Kim-Wipes and cover with plastic film to prevent evaporation.
17. Remove combs and excess acrylamide. Wipe off any residue.
18. Close drain.
19. Clamp gel in place in running apparatus. Tighten clamps "until your fingers fall off."
20. Mix 800 ml 1x TBE. (160 ml of 5X TBE, bring to 800 ml with water) Keep the volume to this minimum to reduce the amout of radioactive waste. Add buffer to top reservoir to just cover the short plate.
21. Use the 60 ml "Acrylamide" syringe to rinse out well area. Pull up some buffer, and squirt it into well area.
22. Pour remaining TBE into bottom reservoir. Fill from very right end to avoid bubbles between glass plates. If bubbles are caught, force out somehow with syringe.
23. Pre-run at 60 watts for about 30 minutes.
24. Disconnect power cables. Clean out well area again with 60 ml syringe.
25. Carefully insert combs so that teeth touch but do not puncture top of the gel.
26. Load samples. Samples are denatured for 3 minutes at 90°C (Program #90) and then kept on ice.
27. Run for 2 hours at 60 watts.
While gel is running
28. Turn on cold trap at least 1 hour before use.
29. Put ice in pre-trap.
30. Cut piece of Whatman 3MM paper about 13.5 X 16.5 inches. Cut off corner to line up with lower left side of gel.
Disassembling the gel
31. Turn off power and remove cables.
32. Drain upper reservoir by opening valve.
33. Remove glass plates, blot off any surface liquid, and set on ice to cool. Let cool for 2 to 3 minutes.
34. Remove combs, and slide spacers out from the sides. Pry plates apart. Gel should stick only to the long plate.
35. Place a piece of Whatman 3MM paper (about 13.5 X 16.5 inches) on top of the gel. Smooth it with your fingers. Carefully peel the gel off, which will stick to the paper.
36. Place gel+paper on the drying machine, gel side up. Cover with plastic wrap, and dry under vacuum with heat set to 80°C for 50 minutes.
37. Dump liquid waste into appropriate radioactive waste bottle. Wash plates, spacers, gel apparatus, etc.
38. When gel is dry, place gel side in contact with X-ray film for 36 to 72 hours in a room temperature cassette.