- Materials
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- Extracted ASFV DNA
- Consumables
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- ASFV primers
- VeriFi HS Mix (PCRBIO)
- Nuclease-free water
- Agencourt AMPure XP Beads (Beckman Coulter™, A63881)
- Freshly prepared 70% ethanol in nuclease-free water
- 0.2 ml thin-walled PCR tubes
- 1.5 ml Eppendorf DNA LoBind tubes
- Qubit™ Assay Tubes (Invitrogen, Q32856)
- Qubit dsDNA BR Assay Kit (Invitrogen, Q32850)
- Equipment
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- Microfuge
- Thermal cycler and/or heating block
- Hula mixer (gentle rotator mixer)
- Magnetic rack suitable for 0.2 ml PCR tubes
- P1000 pipette and tips
- P200 pipette and tips
- P100 pipette and tips
- P20 pipette and tips
- P2 pipette and tips
- Qubit fluorometer (or equivalent for QC check)
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Prepare the primers according to the manufacturer's instructions.
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Pool the tiled primer pairs in Eppendorf or PCR tubes following these proportions. The final stock concentration should be 100 μM.
Odd primer pool:
Primer pair Concentration 3 0.75X 5 2X 7 1X 9 1X 11 1X 11 alt 1X 13 1X 15 1X 17 1.5X 19 1X 21 1X 23 1X 25 2X 27 1X 29 1X 31 0.5X Even primer pool:
Primer pair Concentration 2 1X 4 2X 6 0.5X 8 1.5X 10 2X 12 2X 14 2.5X 16 1X 18 1.5X 20 1X 22 1.5X 24 1.5X 26 1.5X 28 0.75X 30 1.5X 32 1X Primer one:
Primer number Concentration 1 1X Note: Due to the proximity to the telomeric sequence, primer 1 has a shorter amplicon length. The primer does not perform optimally in when pooled with others, therefore it is prepared separately.
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Optional actionTo conserve the DNA samples, dilute 1:10 using nuclease-free water. Unused sample should be stored at –20°C.
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Immediately before setting up the PCR, dilute each primer pool 1:10 in nuclease-free water to make a 10 μM working stock.
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For each sample, prepare one reaction corresponding to each primer pool in 0.2 ml PCR tubes:
Note: For each sample you will have three reactions – odd primer pool, even primer pool, and primer 1 pool.
Between each addition, pipette mix 10-20 times.
Reagent Volume Nuclease-free water 9 µl ASFV DNA (1:10 dilution) 2 µl Primer pool (1:10 dilution) 1.5 µl VeriFi HS Mix (PCRBIO) 12.5 µl Total 25 µl Note: we recommend to make a PCR master mix, either for individual primer pools for multiple samples or one for the three primers sets.
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Mix well by pipetting and spin down.
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Incubate in a thermocycler using the following program:
Step Temperature Time Cycles Initial denaturation 98°C 1 min 1 Denaturation
Annealing
Extension98°C
60°C
72°C15 sec
15 sec
4 min 40 sec
40Final extension 72°C 5 min 1 Hold 10°C ∞ -
Resuspend the AMPure XP beads by vortexing.
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Add 10 µl of resuspended AMPure XP beads to each tube and mix by gently pipetting.
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Incubate for 10 minutes at room temperature.
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Prepare 50 ml of fresh 70% ethanol in nuclease-free water.
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Spin down the tubes and pellet the beads on a magnet for 5 minutes. Keep the tubes on the magnet until the eluate is clear and colourless, and pipette off the supernatant.
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Keep the tubes on the magnet and wash the beads in each well with 200 µl of freshly prepared 70% ethanol without disturbing the pellet. Remove the ethanol using a pipette and discard.
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Repeat the previous step.
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Spin down and place the tubes back on the magnet. Pipette off any residual ethanol. Allow to dry for ~30 seconds, but do not dry the pellet to the point of cracking.
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Remove the tubes from the magnetic rack and resuspend each pellet in 15 µl nuclease-free water. Incubate for 2 minutes at room temperature.
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Pellet the beads on a magnet until the eluate is clear and colourless.
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Remove and retain 15 µl of eluate containing the DNA for each sample, in its three respective primer pools, into a clean tube.
Note: At this point you should still have three tubes for each DNA sample.
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Quantify 1 μl of each cleaned PCR product using a Qubit ds DNA BR assay.
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Using the Qubit reads, pool each sample by quantity in the following proportions:
Primer pool Proportion Pool 1 48% Pool 2 50% Amplicon 1 2%