A FAIR protocol of the Best-RAD sequencing approach

Laure Benoit, Sabine Nidelet, Emeline Charbonnel, Marie-Pierre Chapuis

Published: 2024-02-16 DOI: 10.17504/protocols.io.rm7vz3ok4gx1/v1

RAD sequencing

BestRAD

Sequence-Based Genotyping

PCR duplicate

multiplexing

adapters

biotinylated tag

SNP

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Abstract

The protocol is based on the Best-RAD sequencing approach developed in Ali et al. (2016) that allows two main improvements of the RAD-sequencing approach developed by Baird et al. (2008) : (1) a lower rate of PCR duplicates generated during the final enrichment and (2) an increase of the multiplexing capacity by a factor equal to the number of well barcode available (for a brief description, see Figure 1). We developed this protocol on individual wild samples of the Oriental fruit fly, Bactrocera dorsalis . We evaluated the library quality not only on a high amount of target DNA fragments but also on low amounts of PCR duplicates, chimeric fragments, and adaptor residues. To this aim, we tested and validated the following critical parameters : the quality and quantity of the DNA input, the ratio of the AMpure purifications and the number of PCR cycles of the final amplification. Consecutive recommendations and expectations (e.g., DNA concentration or fragment size) are indicated in notes throughout the protocol. Furthermore, special care has been taken to ensure the precision of each step (i.e. volumes, quantities, duration, materials with supplier references). Consequently, this protocol follows the FAIR principle and could be useful for an easy implementation of the Best-RAD sequencing approach in other laboratories and biological models.

Figure 1. Schematic overview of the BestRAD method (slightly modified from Ali et al. 2016). Part I: Two wells are depicted in each of two different plates. Genomic DNA is digested with a restriction enzyme and ligated to biotinylated well barcode adapters (yellow and blue bars). DNA from each well is pooled platewise, mechanically sheared. Part II: Pools are incubated with streptavidin beads. Following washing, DNA is cleaved from the beads leaving the well barcodes. Part III: Library preparation is performed where a unique combination of plate barcode is added (red and purple bars). Multiple plate libraries can be pooled.
Figure 1. Schematic overview of the BestRAD method (slightly modified from Ali et al. 2016). Part I: Two wells are depicted in each of two different plates. Genomic DNA is digested with a restriction enzyme and ligated to biotinylated well barcode adapters (yellow and blue bars). DNA from each well is pooled platewise, mechanically sheared. Part II: Pools are incubated with streptavidin beads. Following washing, DNA is cleaved from the beads leaving the well barcodes. Part III: Library preparation is performed where a unique combination of plate barcode is added (red and purple bars). Multiple plate libraries can be pooled.

Before start

DNA quality control Beforehand, DNA quality should be checked with an electrophoresis on an agarose gel. Measurement of DNA quantity Beforehand, the amount of genomic DNA should be determined with a fluorescent method (as Qubit assay) in order to be normalized. Sample metadata Prepare a metadata file containing for each sample, a unique ID and the combination of the three barcodes used. This allows to avoid switching barcodes during the sample pooling and is required for sequence read demultiplexing.

Steps

Part I - Genomic DNA restriction, well barcode ligation, pooling and shearing

1.

Genomic DNA enzymatic restriction

1.1.

In a 96-well microplate, normalize the DNA concentration of each DNA sample in a final volume of 10µL using EB buffer.

Note
Ideally, use a total DNA quantity (i.e. for the batch of samples to be pooled) around 2000ng (e.g. 96 samples x 2ng/µL x 10µL, or 24 samples x 8ng/µL x 10µL). At minimum, use a total DNA quantity of 700ng. We validated this minimal input DNA quantity by sequencing a library constructed from an input of 672ng.It is possible to use a larger input DNA quantity, which allows a more reliable normalization. In this case, it is necessary in step 3 to use a volume that corresponds to around 2000ng. This also allows to keep the remaining volume in step 3 as a back-up.It is possible to apply the protocol in parallel on several plates.

1.2.

Into a 1.5 ml low bind tube on a cold block, prepare the master mix of enzymatic restriction (vols for 1 rxn) :

0.55µL

1.2µL

0.25µL

  1. Gently vortex and centrifuge the mix.
  2. Using a new plate on a cold block, a Multipette and a 0.5mL Combitip, distribute 2µL of mix into each well.
  3. Using P10 multichannel pipet, add 10µL of normalized DNA.
  4. Close the plate with self-adhesive film.
  5. Vortex and centrifuge.
1.3.

Apply the following restriction program using a thermocycler (heated lid on):

37°C

80°C

10°C

2.

Ligation of well barcodes

2.1.

Into a 1.5 ml low bind tube, prepare the ligation master mix (vols for 1 rxn) :

1.19µL

0.4µL

0.16µL

0.25µL

  1. Gently vortex the mix and centrifuge.
  2. Using a new plate on a cold block, a Multipette and a 0.5mL Combitip, distribute 2µL of mix into each well.
  3. Using P10 multichannel pipet, add 12µL of digested DNA
  4. Using P10 multichannel pipet, add 2µL of well barcodes (0.05µM if SbfI digestion or 1µM if PstI) according to the plate plan.
  5. Close the plate with self-adhesive film
  6. Gently vortex and centrifuge
2.2.

Apply the following ligation program using a thermocycler (heated lid on):

22°C

65°C (decrease the temperature slowly to 65 at 25°C, e.g. 2.7°C/min during 15min using the minimum ramp speed)

25°C

3.

Pooling, purification and concentration

3.1.

Sample pooling

Note
Choose the volume to be pooled according to the amount of DNA input and the number of samples. Total DNA quantity in the pool should be at least 700ng, and ideally 2000ng.
1. Using a multichannel pipet, pool each barcoded sample of a same batch and a same row of a plate in a 0.2mL 8-tubes PCR strip.

  1. Transfer all the samples of a batch, i.e. all the volume of the 8-tubes PCR strip, in a 1.5mL low-bind tube.
3.2.

Purification and concentration on AMpure beads (ratio 1X)

Incubate AMpure beads at room temperature at least 0h 30m 0s.

Prepare 70% ethanol :

For one pool: `954µL` +`2000µL`.
  1. Measure the exact volume of the pool
  2. Add an equivalent volume of AMpure beads and vortex
  3. Incubate0h 5m 0s at room temperature
  4. Place on a magnetic holder for 0h 5m 0s.
  5. Without removing the tube from the magnetic holder, remove supernatant, add 1000µL, incubate 0h 0m 30s and discard the supernatant.
  6. Repeat step 5 one time
  7. Let dry on the magnetic holder for 0h 8m 0s.
  8. Re-suspend in 100µL (or in 102µL to do the optional 4.2 step)
  9. Remove from the magnetic holder and vortex gently
  10. Incubate 0h 2m 0s at room temperature
  11. Centrifuge briefly and place on a magnetic holder for 0h 5m 0s.
  12. Transfer 100µL( or 102µL for the optional 4.2 step) to Diagenode tubes for shearing Vortex and centrifuge.
4.

Shearing

Maintain tubes at 4°C if shearing is planned within 24 hours or freeze.

4.1.
  1. Briefly vortex and centrifuge the tubes (maximum 12 tubes in parallel, i.e. 12 batches of 96 samples)
  2. Check that there are no bubbles and place the tubes in the rotor (12 tubes of 0.5mL).
  3. Place the rotor in the bath.
  4. Run the following shearing program: Four cycles of :

0h 0m 30s

0h 1m 30s

Repeat the four previous steps one more time (for a total of 8 shearing cycles or a different total number of cycles, see note 4.2).

4.2.

Quality and quantity control (optional)

Run 1µL of each pool on an Agilent High Sensitivity (HS) chip (following manufacturer protocol).

Note
The desired fragment size profile depends on the sequencing read length. For a paired-end sequencing of 150 bp, the profile is expected to be centered around 350bp (200-500bp).The size of the fragments obtained may depend on the biological model and the quality of the DNA input. It is cautious to test different numbers of fragmentation cycles in the bioruptor to adapt the shearing program to the experiment. The higher the number of cycles, the shorter the fragments.

Run 1µL of each pool on a Qubit High Sensitivity (HS) assay (following manufacturer protocol).

Note
At this step, the amount of DNA should be between 350ng and 1500ng. A DNA loss of about 25-50% is expected and mostly due to AMpure purification. The more degraded the input DNA, the larger the loss.

Part II - Selection of fragments carrying the restriction site

5.

Preparation of Dynabead M-280 magnetic streptavidin beads

Since streptavidin beads adhere to tips, re-suspend by vortexing and not by pipetting.

Incubate the streptavidin beads at room temperature for at least 0h 30m 0s before using it.

  1. For each pool, prepare 1mL 2X Binding and Wash buffer (B&W 2X): 10millimolar (mM)

1millimolar (mM)

2Molarity (M)

  1. For each pool, transfer 30µL of Dynabeads into a new 1.5mL low bind tube.
  2. Place the tube on a magnetic holder and discard the supernatant.
  3. Without removing the tube from the magnetic holder, add 100µL
  4. Remove the tube from the magnetic holder, vortex 0h 0m 30s, centrifuge briefly.
  5. Place the tube on the magnetic holder for 0h 1m 0s and discard the supernatant.
  6. Repeat steps 3 to 5 twice for a total of 3 washes.
  7. Re-suspend beads in 100µL.
6.

Binding DNA to beads

6.1.

Add shared DNA (~100µL) to the prepared beads.1. Briefly vortex and centrifuge the tubes.

  1. Apply the following binding program in a thermomixer:

22°C

900rpm

6.2.

For each pool:

  1. Prepare B&W 1X: 250µL + 250µL. Incubate at Room temperature
  2. Pepare B&W 1X: 175µL + 175µL. Incubate at 56°C in a thermomixer.
  3. Prepare NEB buffer 4 1X: 30µL + 270µL. Incubate at Room temperature.
6.3.

Centrifuge briefly and transfer each pool in a 1.5mL low bind tube. 1. Place tubes on a magnetic holder for 0h 1m 0s and discard the supernatant.

  1. Re-suspend the beads with 150µL at Room temperature.
  2. Vortex gently and centrifuge briefly.
  3. Place the tube on the magnetic holder for 0h 1m 0s and discard the supernatant.
  4. Repeat steps 3 to 5 twice with B&W 1X buffer at Room temperature for a total of 3 washes.
  5. Repeat steps 3 to 5 twice more with buffer B&W 1X at 56°C for a total of 5 washes.
7.

Release DNA from streptavidin beads

  1. Re-suspend the beads in 100µL.
  2. Remove the tube from the magnetic holder, vortex gently and centrifuge briefly
  3. Place the tube on the magnetic holder for 0h 1m 0s and discard the supernatant.
  4. Repeat step 1 to 3 for a total of 2 washes.
  5. Re-suspend the beads in 40µL.
  6. Remove the tube from the magnetic holder, vortex gently and centrifuge briefly.
  7. Add 2µL.
  8. Vortex gently.
  9. Run the following release program in a thermomixer:

37°C

600rpm

  1. Vortex gently and centrifuge briefly.
  2. Place the tube on the magnetic rack for 0h 1m 0s.
  3. Transfer 40µL supernatant in a 1.5mL low bind tube.
8.

Purification on AMpure beads (1X)

Incubate AMpure beads at room temperature at least 0h 30m 0s.

Prepare 70% ethanol :

For one pool: `133µL` +`467µL`.
  1. Add 40µL of AMpure beads to each pool.
  2. Incubate0h 5m 0s at room temperature
  3. Place on a magnetic holder for 0h 5m 0s.
  4. Without removing the tube from the magnetic holder, remove supernatant, add 200µL, incubate 0h 0m 30s and discard the supernatant.
  5. Repeat step 5 one time
  6. Let dry on the magnetic support for 0h 8m 0s.
  7. Re-suspend in 51µL(or 52µL for the optional step 9)
  8. Remove from the magnetic holder and vortex gently
  9. Incubate 0h 2m 0s at room temperature
  10. Centrifuge briefly and place on magnetic holder for 0h 5m 0s.
  11. Transfer 50µL(or 51µL for the optional step 9) to a 0.2mL PCR tube.
  12. Vortex and centrifuge.
9.

Quantity control (optional)

Run 1µL of each pool on a Qubit HS assay (following manufacturer protocol).

Note
At this step, the amount of DNA should be between 8ng and 30ng. The large loss of DNA is expected and mainly due to step 6, in which the fragments of interest (which carry the restriction site) are bound to the streptavidin beads, while the rest of the genome is washed out.

Part III - Library construction with NEBNext Ultra II kit

10.

Note
The following steps are described in the NEB Next Ultra II kit's manual, except for two modifications concerning the AMpure purification ratios (see steps 12 and 15) and several recommendations (see notes and optional quality controls).
NEBNext End Prep

Directly into the 50µL from step 8, add :

3µL

7µL

  1. Vortex gently and centrifuge briefly (It is important to mix well. The presence of a small amount of bubbles will not interfere with performance)
  2. Place in a thermocycler, with the heated lid set to ≥ 75°C
  3. Run the following program:

20°C

65°C

4°C

Warning: move to the next step immediately.

11.

Plate barcode adapter Ligation

Directly into the 60µL from step 10, on a cold block, add in order:

30µL

1µL

1.25µL 1.5micromolar (µM)

1.25µL 1.5micromolar (µM)

  1. Vortex gently and centrifuge briefly (It is important to mix well. The presence of a small amount of bubbles will not interfere with performance)
  2. Place in a thermocycler, with the heated lid off
  3. Run the following program:

20°C

10°C

Samples can be stored overnight at –20°C.

12.

Cleanup of Adaptor-ligated DNA (ratio 0.65X)

Note
The ratio recommended in the supplier manual is 0.9X. We suggest a more drastic library sizing with 0.65X ratio, which we have found to be the best for removing free adapters.

Incubate AMpure beads at room temperature at least 0h 30m 0s.

Prepare 70% ethanol :

For one pool: `133µL` +`467µL`.
  1. Transfer 93.5µL from step 11 in a 1.5mL low-bind tube.
  2. Add 61µL
  3. Incubate0h 5m 0s at room temperature
  4. Place on magnetic holder for 0h 5m 0s.
  5. Without removing the tube from the magnetic holder, remove supernatant, add 200µL, incubate 0h 0m 30s and discard the supernatant.
  6. Repeat step 5 one time.
  7. Let dry on the magnetic support for 0h 8m 0s.
  8. Re-suspend in 51µL(52µL for the optional step 13).
  9. Remove from the magnetic holder and vortex gently.
  10. Incubate 0h 2m 0s at room temperature.
  11. Centrifuge briefly and place on magnetic holder for 0h 5m 0s.
  12. Transfer 17µL (18µL for the optional step 13) to a 0.2mL PCR tube.
  13. Vortex and centrifuge.
13.

Quantity control (optional)

Run 1µL of each pool on a Qubit HS assay (following manufacturer protocol).

Note
The amount of DNA is expected to be the same or slightly lower as in step 9. If the quantity is significantly lower as in the step 9 (loss of more than 20%), you could go back with the remaining DNA from step 3 if available.

14.

Final PCR enrichment

Note
The number of final enrichment cycles depends on the quality and quantity of the input DNA (i.e. step 10 of this protocol) and also on the final quantity of library desired depending of the sequencing platform and flowcell format used. For a standard application (Illumina sequencing, which may require around 100ng of library), the NEBNext Ultra II manual recommends 6-7 cycles for 10-30ng of input DNA. For applications requiring a larger library quantity (e.g. RAD-capture by probe hybridization, which may require around 300ng of library), it is recommended to increase the number of cycles (otherwise, it comes with the risk of increasing the rate of PCR duplicates).We tested 6, 9 and 12 PCR cycles for 12ng of NEB input DNA and found an optimal amplification with 9 cycles (i.e. 105ng and 11nM of library for an Illumina NovaSeq sequencing). Furthermore, 9 cycles led to good sequencing results for several libraries constructed from a range of 8-30ng of input DNA.

Into a PCR tube, prepare PCR master mix (vols for 1 rxn) :

25µL

5µL 10micromolar (µM)

5µL 10micromolar (µM)

15µL

  1. Vortex gently and centrifuge briefly
  2. Place in a thermocycler, with the heated lid on, and run the following program:

98°C

6-12 cycles of :

98°C

65°C

65°C

10°C

15.

Cleanup of amplified library (ratio 0.65X)

Note
The ratio recommended in the supplier manual is 0.9X. We suggest a more drastic library sizing with 0.65X ratio, which we have found to be the best for removing free adapters.
Incubate AMpure beads at room temperature at least 0h 30m 0s.

Prepare 70% ethanol :

For one pool: `133µL` +`467µL`.
  1. Verify the 50µL from step 13 and transfert in a new low bind tube.
  2. Add 32.5µL
  3. Place on a magnetic holder for 0h 5m 0s.
  4. Without removing the tube from the magnetic holder, remove supernatant, add 200µL, incubate 0h 0m 30s and discard the supernatant.
  5. Repeat step 4 one time
  6. Let dry on the magnetic support for 0h 8m 0s.
  7. Re-suspend in 22µL
  8. Remove from the magnetic holder and vortex gently
  9. Incubate 0h 2m 0s at room temperature
  10. Centrifuge briefly and place on magnetic holder for 0h 5m 0s.
  11. Transfer 21µL to a 0.2mL PCR tube.
  12. Vortex and centrifuge.
16.

Final quality controls

Run 1µL of each library on a Qubit HS assay (following manufacturer protocol).

Note
At this step, the amount of DNA should be 3-10 times higher than in step 13.

Run 1µL of each library on an Agilent HS chip (following manufacturer protocol).

Note
Calculate the average size of the fragments. It should be around 120bp more than the average of the sheared DNA fragments (see Step 4.2).
Profile of a high-quality library on an Agilent HS chip (NEB input of 12ng, 0.65X AMpure purifications, 9 PCR cycles, final library of 180ng). Fragment sizes are tightly centered around the mean (534 bp), without unexpectedly large fragments.
Profile of a high-quality library on an Agilent HS chip (NEB input of 12ng, 0.65X AMpure purifications, 9 PCR cycles, final library of 180ng). Fragment sizes are tightly centered around the mean (534 bp), without unexpectedly large fragments.
Profile of two over-amplified libraries with free adapters on an Agilent HS chip (NEB input 50ng, 0.9X AMpure purifications, 12 PCR cycles). Low molecular weight peaks form a scale pattern. Over-amplification is visible around 6000bp.
Profile of two over-amplified libraries with free adapters on an Agilent HS chip (NEB input 50ng, 0.9X AMpure purifications, 12 PCR cycles). Low molecular weight peaks form a scale pattern. Over-amplification is visible around 6000bp.
Unexpectedly low molecular weight peaks (about 120bp, 240bp) are likely free adapters and adapter doublets. If peak heights are relatively elevated, the amount of adapters used was too large and/or the AMpure purification from step 12 and 15 was not efficient enough. You can apply once more cleanup of step 15 in order to attempt to remove this unexpected peaks. Unexpectedly high molecular weight peaks (e.g., above 6000bp) are likely artificial concatenation, i.e. artifact sequences formed by two or more biological sequences. These fragments are caused by over-amplification in step 14, because of a too large number of cycles. Over-amplification will also produce more PCR duplicates. See the note of step 14 to optimize the number of PCR cycles. Small fragments are favored by Illumina sequencing. Thus, unexpected low molecular weight peaks are more problematic than high ones.

Run 2µL of each library on a Kapa assay (following manufacturer protocol).

Note
At this step, the concentration should ideally reach a minimum of 10nM. However, when input DNA is of low quality or quantity, 2nM may be sufficient (depending on the sequencing facility).KAPA library concentration can be used to normalize several batches of samples aimed to be pooled altogether before Illumina sequencing. Don't use concentration values obtained by a Qubit assay, because it represents all DNA fragments and not only those with both Illumina adapters which will be sequenced.

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