ddRADseq for animal population genomics/phylogenomics

Before starting the protocol, it is necessary to prepare the SbfI and MseI adapters as follows:

Barcoded SbfI adapter annealing:

Anneal unique adapter pairs by mixing 2µL and 2µL with 196µL to make 200µL.

Heat to 95°C for 0h 5m 0s and slowly cool to Room temperature. Anneal the set of adapters in a plate format. It will be more convenient for later use in setting up reactions. Perform an additional 20-fold dilution to obtain a working solution at a concentration of 0.1 µM.

Non-barcoded MseI adapter annealing:

Mix 100µL and 100µL with 800µL to make 1000µL.

Heat to 95°C for 0h 5m 0s and slowly cool to Room temperature to anneal the single-stranded adapters into a double-stranded adapter.

In this section of the protocol, the restriction digestion reaction of samples will be carried out.

Prepare the restriction reaction mastermix for all the samples using the amounts of reagents below (3 µL reaction mix per sample), mix by brief vortexing, and shortly centrifuge:

1.9

0.9

0.1

0.1

Place 6µL in each well of a 96-well plate.

Add 3µL to each well of the sample DNA plate.

The total reaction volume should now be 9 µL. Cover and seal the plate, centrifuge, and

incubate at37°C for3h 0m 0s on a thermal cycler with a heated lid. Inactivate the

restriction enzymes fo0h 20m 0s a65°C . Store a4°C .

In this section of the protocol, the prepared double-stranded SbfI and MseI adapters (as described in step 1 - Preparation) will be ligated to the restriction-digested DNA (see step 2 - Restriction Reaction).

Prepare the ligation reaction mastermix for all the samples using the amounts of reagents below (1.6 µL reaction mix per sample), mix by brief vortexing, and shortly centrifuge:

0.26

0.12

1

0.17

0.06

Add 1.6µL to each well of the restriction-digested DNA.

Add 1µL to each well.

The total reaction volume should now be 11.6 µL. Cover and seal the plate, centrifuge, and incubate at 16°C for 3h 0m 0s on a thermal cycler. Store at -20°C if the library is not to be processed immediatly.

Purify the ligation reaction product using an AMPure XP protocol with a 1:1 ratio and elute in 45 uL of nuclease-free water.

We introduced the use of a custom-made magnetic 96-pin plate (see YouTube Tutorial, starting at 11:40) to increase the efficiency of the protocol.

Let the AMPure aliquot at Room temperature for 0h 30m 0s and then shake it gently to resuspend the magnetic beads.

Elute the beads by placing the dried plate/device on the elution plate and remove the 96-pin magnet plate slowly. Then gently shake/move the PCR cover plate to solubilize the beads. Wait 0h 10m 0s to 0h 20m 0s until the solution homogenizes and the DNA fully detaches from the beads.

Introduce the 96-pin magnet plate again on the PCR plate to attract the (now DNA-free) beads. Wait at least one minute, then remove the plate/device and throw the PCR plate with the used AMPure beads away.

Add 8.4µL to each well to have 20µL in each well.

Prepare one microplate with 20µL in each well.

Prepare two microplates with 100µL in each well.

Prepare one microplate with 45µL in each well.

Transfer your 20µL to the AMPure plate and (optionally) mix by pipetting 10 times up and down.

Incubate for 0h 5m 0s at Room temperature.

Equip a 96-pin magnet device (custom-made by Tomasz Suchan) with a clean PCR plate as a cover and apply it to the beads+DNA plate. Wait for 0h 10m 0s to separate the beads from the solution.

Rinse the attached beads twice for 0h 0m 30s each in the ethanol microplates, then hold the plate/device for a couple of minutes to dry the beads. Cracks on the beads indicate drying.

The library amplification/PCR step employs Illumina PCR primers to amplify restricted fragments from their ligated adapters.

Prepare the PCR reaction mastermix for all samples using the amounts of reagents below (7 µL per sample but remember to prepare enough mastermix to run 2 PCR reactions per sample!):

1.48

2.00

0.08

0.67

0.67

0.10

2.00µL

Add 7µL to each well of two PCR plates.

Add 3µL purified in step 4.

PCR Reaction:

Denaturation

0h 0m 30s at 98°C

20 cycles

0h 0m 20s at 98°C,

0h 0m 30s at 60°C,

0h 0m 40s at 72°C

Final Elongation

0h 2m 0s at 72°C

Hold at 4°C

Pool the two replicate plates together to obtain a single plate with 20 uL of amplicons in each well.

Run 3 µL of each PCR product on a 1.5% at 120V for 0h 30m 0s. You should see a smear of PCR product from 200 bp to 1000 bp, sometimes with a bright band of adapter dimer at 120 bp.

Pool the PCR products of all samples into one tube. Samples that failed to amplify, or amplified only the adapter dimer, as revealed by the gel, can be excluded from the pool.

Conduct size selection within the range of 400 to 500 base pairs. You can achieve this using a PippinPrep instrument, following the provided guidelines from the manufacturer. Another method involves a prolonged migration on an agarose gel, excising the desired region, and purifying it using a gel extraction kit, as outlined below.

Prepare a 2.0% using clean TAE buffer and a large comb, with the exact volume adjusted as needed.

Accurately measure the total elution volume (in µL, with a pipet) and its concentration (in ng/µL, with a fluorometer, e.g., Qubit, using 1 µL of sample). Multiply both numbers to obtain the amount of DNA in your library (in ng). You can then divide this number by 25 to calculate the elution volume needed to concentrate the library to an optimal 25 ng/µL in the next step. Keep in mind that you might lose 10 - 15 % of your DNA yield during this concentration step and that it is best to have at least 10 µL of library (down to 3-4 ng/µL is still ok).

Fill the gel rig with clean gel buffer (e.g., TAE).

Prepare a gel template with 400µL + 100µL.

Load 3µL in a few wells, leaving a regular interspace.

Load as much library as possible in the remaining wells, but without overflowing the wells.

Migrate the gel at low voltage and for a sufficiently long time to ensure a high-resolution migration. We typically go for 80V during 3h 0m 0s.

Check for proper migration by very quick UV exposure and prepare tubes for the gel pieces.

Cut the 400 - 500 bp region on a UV table using sterile scalpel/razor blades. Minimize UV exposure by only cutting a few gel pieces at a time and individualize each piece in the prepared tubes. Try to remove the empty agarose to limit the amount of gel to be purified and be careful not to include ladder fragments in your excisions.

Purify each gel slice separately using a gel extraction kit (e.g., Monarch DNA Gel Extraction Kit from New England Biolabs). Elute in ~15 - 20 µL of elution buffer and pool elution products into a single tube.

Purify the ligation reaction product using the AMPure XP protocol with a 1:1 ratio and elute in the calculated volume of nuclease-free water.

Let the AMPure aliquot at Room temperature for 0h 30m 0s and then gently shake it to resuspend the magnetic beads.

Add AMPure to your elution following a 1:1 volume ratio. Mix by inverting the tube gently several times and wait 0h 5m 0s. Put the tube on a magnet rack for tubes.

Wait a couple of minutes until the beads migrate to the tube wall. Then, remove the supernatant by pipetting.

Rince the beads twice using 1000µL, wait 0h 0m 30s, and remove the ethanol.

After the second rinsing step, let the beads dry. This may take up to 30-40 minutes, especially in 1.5 mL tubes and without air conditioning. The beads will progressively lose their shine, which will indicate drying.

Elute the DNA by pipetting the desired volume of water on the beads, to detach them from the tube wall. You should remove the tube from the rack and re-pipet the elution liquid on the beads until all of them are detached.

Wait a few minutes and put the tube back on the magnet rack. Then wait a few more minutes until the beads get stuck again. Carefully pipet the elution volume (avoiding the beads) in a clean final and properly labeled 1.5 ml tube.

Check size selection and library fragment size using gel electrophoresis or a TapeStation/Bioanalyzer/Fragment Analyzer.

Quantify the amount of DNA using a fluorimeter (e.g., Qubit).

Proceed to Illumina sequencing or equivalent. DNA quantity and mean fragment length can be used to calculate the molarity of the library.