TITLE: ddRADseq for animal population genomics/phylogenomics
AUTHORS: Tomasz Suchan, Christophe Dufresnes, Robin Schmidt, Johanna Ambu, Sven Gippner, Miguel Vences

[DESCRIPTION]
Restriction-site Associated DNA sequencing (RADseq) offers a simple, affordable, and versatile approach to swiftly genotype thousands of genetic markers without prior optimization for population genetic and phylogenetic analyses on any kind of organism (Baird et al. 2008). Although RADseq protocols have undergone significant advancements, they remain difficult to implement de novo for researchers inexperienced with the technique in labs that are not a priori equipped for NGS technologies. 

The protocol presented here aims to facilitate such implementation. It consists of an adaptation of the double digest RADseq (ddRADseq) protocol proposed by Peterson et al. (2012) and further modified by Brelsford et al. (2016).  Compared to the latter, it optimizes reagent usages and introduces a facilitated procedure of AMPure purification of an entire 96-well plate at once, by using a custom magnetic 96-pin device.

The procedure goes as follows. Genomic DNA of the samples undergoes enzyme restriction by SbfI, a 'rare'-cutter, and MseI, a more 'frequent'-cutter. The digested DNA fragments are then ligated to adapters on each end, with the SbfI adapter containing individual 4-8 bp barcodes for multiplexing samples in the library. The ligated fragments are then purified and amplified by replicate PCRs using primers that include TruSeq Illumina indices for sequencing as well as indexing. PCR products are then pooled and size selection (400-500bp) is performed either with a PippinPrep Instrument or through ad hoc migration/excision on an agarose gel. The size-selected library is purified, quantified, and controlled before Illumina sequencing.

So far, this protocol, or the previous version it derives from (Brelsford et al., 2016), has been successfully used in phylogeographic studies across various non-model animals, including amphibians, reptiles, mammals, and insects (e.g. Ambu et al., 2023).

 


[BEFORE_START]
Before starting the ddRADseq protocol you should order and prepare all materials including reagents, consumables, and adapters in advance. 

 

Adapter sequences

SbfI adapter p1.1
ACACTCTTTCCCTACACGACGCTCTTCCGATCTNNNNNNTGCA
SbfI adapter p1.2
[5phos]nnnnnnAGATCGGAAGAGCGTCGTGTAGGGAAAGAGTGT

where NNNNNN is a sample-specific 4-7 bp barcode, nnnnnn is a reverse complement; for instance, we use 96 barcoded adapters which allow barcoding a full PCR plate of samples. The barcode appears at the beginning of each read (i.e. "inline" barcode).

MseI adapter p2.1:
GTGACTGGAGTTCAGACGTGTGCTCTTCCGATCT
MseI adapter p2.2:
[5phos]TAAGATCGGAAGAGCGAGAACAA

MseI adapters are universal, the same is used for all the samples.


Primer sequences and preparation

Illumina PCR1 primer:
AATGATACGGCGACCACCGAGATCTACACTCTTTCCCTACACGACG
Indexed PCR2 Primers:
CAAGCAGAAGACGGCATACGAGATnnnnnnGTGACTGGAGTTCAGACGTGTGC

where nnnnnn is an Illumina index. These can be used to uniquely tag pools of samples barcoded with inline barcodes. In our case, we tag 96 samples with inline barcodes (in the P1 adapter) and then index these pools, each pool with a unique indexed primer.

 


[GUIDELINES]
Prevent cross-contamination by using low-retention filter tips for all pipetting procedures. If possible, we recommend working under a laminar flow bench to protect the samples and further avoid contamination. A non-template control on the plate may also be incorporated to stay tuned for possible contamination. 

Handle the SbfI barcoded adapter plate with great care, as cross-contamination on this plate might invalidate your entire study and any future study using that plate. Moreover, barcode cross-contamination may only become apparent through inconsistent results after data analysis.

[STEPS]
SECTION: Preparation
1. Before starting the protocol, it is necessary to prepare the SbfI and MseI adapters as follows:
SECTION: Restriction Reaction
2. In this section of the protocol, the restriction digestion reaction of samples will be carried out.

SECTION: Purification (short fragment removal)
4. 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.
SECTION: Library Amplification
5. The library amplification/PCR step employs Illumina PCR primers to amplify restricted fragments from their ligated adapters.

 

SECTION: Ligation Reaction
3. 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).
SECTION: Illumina Sequencing
10. Proceed to Illumina sequencing or equivalent. DNA quantity and mean fragment length can be used to calculate the molarity of the library.
 

SECTION: Final quantification
9. Quantify the amount of DNA using a fluorimeter (e.g., Qubit).
SECTION: Final quantification
8. Check size selection and library fragment size using gel electrophoresis or a TapeStation/Bioanalyzer/Fragment Analyzer.
SECTION: Size selection
6. 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.
SECTION: Restriction Reaction
2.1. 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 1
0.9 1
0.1 1
0.1 1
SECTION: Restriction Reaction
2.2. Place 6 µL in each well of a 96-well plate.

 

 

SECTION: Restriction Reaction
2.3. Add 3 µL to each well of the sample DNA plate.
SECTION: Restriction Reaction
2.4. The total reaction volume should now be 9 µL. Cover and seal the plate, centrifuge, and
incubate at 37 °C for 180 min on a thermal cycler with a heated lid. Inactivate the
restriction enzymes for 20 min at 65 °C. Store at 4 °C.

SECTION: Preparation
1.1. Barcoded SbfI adapter annealing:

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

Heat to 95 °C for 5 min 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.

 

SECTION: Preparation
1.2. Non-barcoded MseI adapter annealing:

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

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

SECTION: Ligation Reaction
3.1. 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 1
0.12 1
1 1
0.17 1
0.06 1
SECTION: Purification (short fragment removal)
4.1. Let the AMPure aliquot at Room temperature for 30 min and then shake it gently to resuspend the magnetic beads.

 

SECTION: Purification (short fragment removal)
4.2. Add 8.4 µL to each well to have 20 µL in each well.

SECTION: Ligation Reaction
3.2. Add 1.6 µL to each well of the restriction-digested DNA.

SECTION: Ligation Reaction
3.3. Add 1 µL to each well.

 

SECTION: Ligation Reaction
3.4. The total reaction volume should now be 11.6 µL. Cover and seal the plate, centrifuge, and incubate at 16 °C for 180 min on a thermal cycler. Store at -20 °C if the library is not to be processed immediatly.

SECTION: Purification (short fragment removal)
4.3. Prepare one microplate with 20 µL in each well.

SECTION: Purification (short fragment removal)
4.4. Prepare two microplates with 100 µL in each well.

SECTION: Purification (short fragment removal)
4.5. Prepare one microplate with 45 µL in each well.

 

SECTION: Purification (short fragment removal)
4.6. Transfer your 20 µL to the AMPure plate and (optionally) mix by pipetting 10 times up and down.

 

SECTION: Purification (short fragment removal)
4.7. Incubate for 5 min at Room temperature.

SECTION: Purification (short fragment removal)
4.8. 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 10 min to separate the beads from the solution. 

 

SECTION: Purification (short fragment removal)
4.9. Rinse the attached beads twice for 30 s 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.

 

SECTION: Purification (short fragment removal)
4.10. 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 10 min to 20 min until the solution homogenizes and the DNA fully detaches from the beads.

SECTION: Purification (short fragment removal)
4.11. 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.

 

SECTION: Library Amplification
5.1. 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 1
2.00 1
0.08 1
0.67 1
0.67 1
0.10 1
2.00 µL

SECTION: Library Amplification
5.2. Add 7 µL to each well of two PCR plates.

SECTION: Library Amplification
5.3. Add 3 µL purified in step 4.

SECTION: Library Amplification
5.4. PCR Reaction:

Denaturation
30 s at 98 °C
20 cycles
20 s at 98 °C,
30 s at 60 °C, 
40 s at 72 °C
Final Elongation
2 min at 72 °C
Hold at 4 °C

SECTION: Library Amplification
5.5. Pool the two replicate plates together to obtain a single plate with 20 uL of amplicons in each well.
SECTION: Library Amplification
5.6. Run 3 µL of each PCR product on a 1.5 % at 120 V for 30 min. 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.

SECTION: Library Amplification
5.7. 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.

SECTION: Purification &amp; Concentration
7. 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.
SECTION: Purification &amp; Concentration
7.1. Let the AMPure aliquot at Room temperature for 30 min and then gently shake it to resuspend the magnetic beads.

 

SECTION: Purification &amp; Concentration
7.2. Add AMPure to your elution following a 1:1 volume ratio. Mix by inverting the tube gently several times and wait 5 min. Put the tube on a magnet rack for tubes.

SECTION: Purification &amp; Concentration
7.3. Wait a couple of minutes until the beads migrate to the tube wall. Then, remove the supernatant by pipetting.
SECTION: Purification &amp; Concentration
7.4. Rince the beads twice using 1000 µL, wait 30 s, and remove the ethanol.
SECTION: Purification &amp; Concentration
7.5. 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.
SECTION: Purification &amp; Concentration
7.6. 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.
SECTION: Purification &amp; Concentration
7.7. 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.

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

 

SECTION: Size selection
6.2. Fill the gel rig with clean gel buffer (e.g., TAE).
SECTION: Size selection
6.3. Prepare a gel template with 400 µL + 100 µL.

SECTION: Size selection
6.4. Load 3 µL in a few wells, leaving a regular interspace.

 

SECTION: Size selection
6.5. Load as much library as possible in the remaining wells, but without overflowing the wells.
SECTION: Size selection
6.6. Migrate the gel at low voltage and for a sufficiently long time to ensure a high-resolution migration. We typically go for 80 V during 180 min.

SECTION: Size selection
6.7. Check for proper migration by very quick UV exposure and prepare tubes for the gel pieces.
SECTION: Size selection
6.8. 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.

 

SECTION: Size selection
6.9. 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.
SECTION: Size selection
6.10. 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).