- Materials
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- Native Barcoding Expansion 1-12 (EXP-NBD104) and 13-24 (EXP-NBD114) if multiplexing more than 12 samples
- Consumables
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- Freshly prepared 70% ethanol in nuclease-free water
- 1.5 ml Eppendorf DNA LoBind tubes
- Nuclease-free water (e.g. ThermoFisher, AM9937)
- Agencourt AMPure XP beads (Beckman Coulter, A63881)
- NEB Blunt/TA Ligase Master Mix (NEB, M0367)
- Equipment
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- Magnetic rack, suitable for 1.5 ml Eppendorf tubes
- Hula mixer (gentle rotator mixer)
- Vortex mixer
- Ice bucket with ice
- Microfuge
- P1000 pipette and tips
- P100 pipette and tips
- P10 pipette and tips
- Optional equipment
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- Qubit fluorometer (or equivalent for QC check)
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Prepare the NEB Blunt/TA Ligase Master Mix according to the manufacturer's instructions, and place on ice:
Thaw the reagents at room temperature.
Spin down the reagent tubes for 5 seconds.
Ensure the reagents are fully mixed by performing 10 full volume pipette mixes.
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Thaw the native barcodes at room temperature. Use one barcode per sample. Individually mix the barcodes by pipetting, spin down, and place them on ice.
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Select a unique barcode for every sample to be run together on the same flow cell, from the provided 24 barcodes. Up to 24 samples can be barcoded and combined in one experiment.
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Dilute 100–200 fmol of each end-prepped sample to be barcoded to 22.5 µl in nuclease-free water.
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Add the reagents in the order given below, mixing by flicking the tube between each sequential addition:
Reagent Volume 100–200 fmol end-prepped DNA 22.5 µl Native Barcode 2.5 µl Blunt/TA Ligase Master Mix 25 µl Total 50 µl -
Mix well by pipetting using wide-bore pipette tips. Alternatively, if you are concerned about preserving the integrity of very long DNA fragments, mix gently by flicking the tube, and spin down.
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Incubate the reaction for 10 minutes at room temperature.
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Resuspend the AMPure XP beads by vortexing.
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Add 50 µl of resuspended AMPure XP beads to the reaction and mix by pipetting.
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Incubate on a Hula mixer (rotator mixer) for 5 minutes at room temperature.
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Prepare 500 μl of fresh 70% ethanol in nuclease-free water.
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Spin down the sample and pellet on a magnet. Keep the tube on the magnet, and pipette off the supernatant when clear and colourless.
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Keep the tube on the magnet and wash the beads 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 tube 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 tube from the magnetic rack and resuspend the pellet in 26 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 26 µl of eluate containing the DNA library into a clean 1.5 ml Eppendorf DNA LoBind tube.
Dispose of the pelleted beads
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Quantify 1 µl of eluted sample using a Qubit fluorometer.
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Pool equimolar amounts of each barcoded sample into a 1.5 ml Eppendorf DNA LoBind tube, ensuring that sufficient sample is combined to produce a pooled sample of 100–200 fmol total.
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Quantify 1 µl of pooled and barcoded DNA using a Qubit fluorometer.
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Dilute 100–200 fmol pooled sample to 65 µl in nuclease-free water.
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Optional actionIf 100–200 fmol for R9.4.1 of pooled sample exceeds 65 µl in volume, perform an AMPure clean-up with 2.5x Agencourt AMPure XP beads to pooled sample volume, eluting in 65 µl of nuclease-free water.
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Fragment size and adapter ligation
The amount of adapter has been optimised for fragment sizes greater or equal to 8 kb. If the fragments are generally smaller than 3 kb, adjustments should be made to use 0.1–0.2 pmoles of DNA in the adapter ligation step.