Sep 24, 2024
Double digestion RADseq library
- David Macaya-Sanz1
- 1Institute of Forest Sciences, ICIFOR-INIA-CSIC (RoR: 02nxes898), Madrid, Spain
Protocol Citation: David Macaya-Sanz 2024. Double digestion RADseq library. protocols.io https://dx.doi.org/10.17504/protocols.io.n92ldm66nl5b/v1
License: This is an open access protocol distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited
Protocol status: Working
We use this protocol and it's working
Created: December 20, 2023
Last Modified: September 24, 2024
Protocol Integer ID: 92559
Keywords: RADseq, restriction enzyme, genotyping, GbS, Genotyping-by-Sequencing
Funders Acknowledgement:
DEDGENE: Arquitectura y predicción genómica de la resistencia a la grafiosis del olmo como herramientas para la mejora genética acelerada
Grant ID: PID2021-127347OA-I00
Disclaimer
Protocol has been tested and used to successfully create libraries several times. Thus, it seems to be reliable and transferable. However, we recommend you to set it up and test it in your lab and system, before processing a large amount of samples.
Protocol is inspired in Peterson et al. 2012 (10.1371/journal.pone.0037135) and Parchman, Gompert and Buerkle 2011 rfseq protocol, with multiple modifications.
Abstract
A protocol to create libraries for Illumina to genotype through double digestion sequencing. The goal is to create a protocol that reduces costs and dedicated time at the expense of a slight reduction of quality, meaning that some of your samples may not meet depth minimum, but the specificity is of the tags is remarkable. Thus, you will probably have a low percentage of samples (in our case less than 10%) needed to be re-processed. The amount of library product should be more than enough to be run in an Illumina sequencer. This version of the protocol includes two clean-ups and no Inline tags in the adapters, making it more expensive and laborious than other versions.
Guidelines
The protocol is recommended to be done in three different days:
1- Digestion and first clean-up.
2- Ligation and second clean-up.
3- PCR and last clean-up.
Samples could be conserved for later use after every section, but we recommend to do so after clean-ups or PCR.
Protocol materials
MseI - 500 unitsNew England BiolabsCatalog #R0525S
Step 10
T4 DNA Ligase - 20,000 unitsNew England BiolabsCatalog #M0202S
Step 28
PHUSION PLUS DNA POLYMERASE 500 RXNThermo ScientificCatalog #F630L
Step 43
EcoRI-HF - 10,000 unitsNew England BiolabsCatalog #R3101S
Step 10
Safety warnings
Follow the common molecular biology lab safety procedures.
Ethics statement
Nothing to declare. No animals or personal data was used.
Before start
Before start, make sure you have at hand all the needed reagents, in particular the oligos (adapters and PCR primers). Other important reagents are: digestion enzymes with buffer, T4 ligase, DNA polymerase, Ampure beads, absolute ethanol, TE buffer, NaCl, and molecular grade water. Also a thermocycler, a centrifuge and a magnetic rack for 96-well plate are needed. Other lab equipment is necessary.
Oligo sequences
Oligo sequences
Make sure you have the required adapter oligos.
Note
| A | B | |
| MseI_P2.1_2N | GTCTCGTGGGCTCGGAGATGTGTATAAGAGACAGNN | |
| MseI_P2.2_2N | /5Phos/TANNCTGTCTCTTATACGAGGACAA | |
| EcoRI_P1.1 | TCGTCGGCAGCGTCAGATGTGTATAAGAGACAG | |
| EcoRI_P1.2 | /5Phos/AATTCTGTCTCTTATACACATCTGACGCTGCCGACGA |
Sequences of the adapter oligos. Note that two Ns have been added to MseI to be able to remove PCR duplicates in the bioinformatic pipeline (allowing a maximum of 16 combinations, enough for low-depth sequencing). These Ns can be removed (rendering no deduplication of reads). Also they can be reduced to one (only 4 combinations, thus not recommended). Also they might work if extended to three (allowing 64 combinations, enough for higher sequencing depth), however we have not tested it.
Make sure you have the required PCR primers oligos.
Note
| Step2_NXTi7_N701 | CAAGCAGAAGACGGCATACGAGATTCGCCTTAGTCTCGTGGGCTCGG | |
| Step2_NXTi7_N702 | CAAGCAGAAGACGGCATACGAGATCTAGTACGGTCTCGTGGGCTCGG | |
| Step2_NXTi7_N703 | CAAGCAGAAGACGGCATACGAGATTTCTGCCTGTCTCGTGGGCTCGG | |
| Step2_NXTi7_N704 | CAAGCAGAAGACGGCATACGAGATGCTCAGGAGTCTCGTGGGCTCGG | |
| Step2_NXTi7_N705 | CAAGCAGAAGACGGCATACGAGATAGGAGTCCGTCTCGTGGGCTCGG | |
| Step2_NXTi7_N706 | CAAGCAGAAGACGGCATACGAGATCATGCCTAGTCTCGTGGGCTCGG | |
| Step2_NXTi7_N707 | CAAGCAGAAGACGGCATACGAGATGTAGAGAGGTCTCGTGGGCTCGG | |
| Step2_NXTi7_N708 | CAAGCAGAAGACGGCATACGAGATCCTCTCTGGTCTCGTGGGCTCGG | |
| Step2_NXTi7_N709 | CAAGCAGAAGACGGCATACGAGATAGCGTAGCGTCTCGTGGGCTCGG | |
| Step2_NXTi7_N710 | CAAGCAGAAGACGGCATACGAGATCAGCCTCGGTCTCGTGGGCTCGG | |
| Step2_NXTi7_N711 | CAAGCAGAAGACGGCATACGAGATTGCCTCTTGTCTCGTGGGCTCGG | |
| Step2_NXTi7_N712 | CAAGCAGAAGACGGCATACGAGATTCCTCTACGTCTCGTGGGCTCGG | |
| Step2_NXTi5_S502 | AATGATACGGCGACCACCGAGATCTACACCTCTCTATTCGTCGGCAGCGTC | |
| Step2_NXTi5_S503 | AATGATACGGCGACCACCGAGATCTACACTATCCTCTTCGTCGGCAGCGTC | |
| Step2_NXTi5_S505 | AATGATACGGCGACCACCGAGATCTACACGTAAGGAGTCGTCGGCAGCGTC | |
| Step2_NXTi5_S506 | AATGATACGGCGACCACCGAGATCTACACACTGCATATCGTCGGCAGCGTC | |
| Step2_NXTi5_S507 | AATGATACGGCGACCACCGAGATCTACACAAGGAGTATCGTCGGCAGCGTC | |
| Step2_NXTi5_S508 | AATGATACGGCGACCACCGAGATCTACACCTAAGCCTTCGTCGGCAGCGTC | |
| Step2_NXTi5_S510 | AATGATACGGCGACCACCGAGATCTACACCGTCTAATTCGTCGGCAGCGTC | |
| Step2_NXTi5_S511 | AATGATACGGCGACCACCGAGATCTACACTCTCTCCGTCGTCGGCAGCGTC |
Sequences of PCR primers, to allow pooling 96 samples. Those primers produce Nextera-like Illumina libraries. More primers can be created following Illumina scheme. These primers can be used to generate Metabarcoding libraries as well.
DNA dilution
DNA dilution
3h
3h
Measure your DNA stock concentration. Recommended Nanodrop One.
Equipment
NanoDrop™ One/OneC Microvolume UV-Vis Spectrophotometer
NAME
UV-Vis Spectrophotometer
TYPE
Thermo Scientific
BRAND
ND-ONE-W
SKU
1h
Dilute DNA stock to 20 ng/µl at minimum volume of 7.5 µL
2h
Annealing of adapters
Annealing of adapters
1h 50m
1h 50m
Dilute oligos EcoRI_P1.1, EcoRI_P1.2, MseI_P2.1_2N, and MseI_P2.2_2N to 100 micromolar (µM)
In case you have Tris EDTA (TE) ready, you can dilute the oligos in the buffer.
10m
Create Annealing buffer stock (10x) (it is TE buffer (10x) plus the sodium chloride):
100 millimolar (mM) Tris HCl, pH 8
10 millimolar (mM) EDTA
500 millimolar (mM) NaCl
In a 1L flask:
800 mL water
15.759 g Tris-HCl
2.922 g EDTA
29.22 g NaCL
Adjust pH to 8 and bring volume to 1L.
In case of TE buffer ready, you can just prepare a NaCl 0.5 M dilution on the TE.
30m
Combine oligos EcoRI_P1.1 and EcoRI_P1.2 into the same stock to a final 20 micromolar (µM) total dimer concetration
Therefore, add 5 µL EcoRI_P1.1 , 5 µL EcoRI_P1.2 , 2.5 µL 10x annealing buffer , and 12.5 µL water
In case of TE buffer ready: add 5 µL EcoRI_P1.1 , 5 µL EcoRI_P1.2 , 2.5 µL NaCl 0.5M , and 12.5 µL TE buffer
5m
Combine oligos MseI_P2.1_2N and MseI_P2.2_2N, in the same fashion as with EcoRI_P1.
5m
Incubate each oligo combination in thermal cycler: 97.5 °C 00:02:30 followed by a touchdown of 1 °C 00:00:20 until reaching 21 °C . Hold afterwards at 4 °C
1h
Double digest
Double digest
2h 15m
2h 15m
Prepare Digestion Mix adding 0.9 units EcoRI enzyme , 0.45 units MseI enzyme and 1 µL rCutSmart Buffer per sample. Then, add water to get a final volume of 2.5 µL Digestion Mix per sample.
For example, for 48 samples, prepare mix in excess, so factor would be 110x:
5 µL EcoRI-HF at 20000 units/ml total 100 units
5 µL MseI at 10000 units/ml total 50 units
110 µL rCutSmart Buffer at 10 X
155 µL water (you can add some extra water: such as 5 µL
EcoRI-HF - 10,000 unitsNew England BiolabsCatalog #R3101S
MseI - 500 unitsNew England BiolabsCatalog #R0525S
15m
On a 96-well PCR plate, dispense 2.5 µL Digestion Mix in empty PCR tubes or plate wells, keeping it On ice
Spin the plate, to bring to the bottom the digestion mix.
10m
Transfer 7.5 µL genomic DNA at 20 ng/µl to the wells and mix DNA and mix pipetting up and down a few times. Total volume should be 10 µL . It is fundamental to keep track of the layout of the DNA samples transferred into the plate.
20m
Incubate at 37 °C for 01:30:00 . Hold at 4 °C afterwards. Do not heat kill enzymes.
Reactions can be stored at 4 °C Overnight
1h 30m
Clean-up
Clean-up
2h 45m
2h 45m
Potentially size selection could be done here, but In this version we are proceeding with a simple clean-up.
Take an aliquot of magnetics beads, out of the fridge to let them reach Room temperature before initiating the protocol (at least 00:30:00 at Room temperature )
For 96 samples, at least 1 mL magnetic beads
30m
Add 10 µL magnetic beads (1X) to each digestion reaction. Mix well by pipetting up and down at least 10 times. If centrifuging samples after mixing, be sure to stop the centrifugation before the beads start to settle down (no more than 1000 rpm).
20m
Incubate samples on bench top for at least 00:05:00 at room temperature.
5m
Place the plate on a magnetic stand, and wait 00:05:00 or until the solution is clear.
While waiting, prepare 35 mL Ethanol 80% (28 mL ethanol absolute + 7 mL water ). It has to be freshly prepared.
5m
Carefully remove and discard the supernatant (approximately 17 µL ), not disturbing the beads.
20m
Wash with 160 µL freshly prepared ethanol 80% , not removing the plate from the magnetic stand. Incubate beads in the ethanol for 00:00:30 . Remove and discard the ethanol, but do not disturbing the beads. Some ethanol can remain if so needed not to disturb the beads.
Recommendation: remove 75 µL ethanol in two pipetting moves.
15m
Wash AGAIN with 160 µL freshly prepared ethanol 80% , not removing the plate from the magnetic stand. Incubate beads in the ethanol for 00:00:30 . Remove and discard the ethanol, but do not disturbing the beads. In this step, ethanol remaining should be as little as possible.
Recommendation: remove 80 µL ethanol with 200 µL tips , and afterwards use 10 µL tips to remove the last drops of ethanol.
25m
After removing all visible liquid, let the beads air dry approximately 00:03:00 , but DO NOT over-dry (losing the glossy shine, turning lighter brown and starting to crack). Beads may overdry really quickly with these volumes. If necessary, do not remove ethanol from all wells at once. Do it in a manner that the first columns of the plate do not overdry, so removing ethanol and adding elution buffer (next step) might intercalate.
3m
Remove from stand, add 13 µL elution buffer and completely dissolve beads pipetting up and down. Use a pipette that can dispense the complete amount in a single maneuver. Make a very short spin (no more than 1000 rpm) to bring down the elution and let it rest at least 00:02:00 atRoom temperature . Elution buffer could be water, Tris or TE buffers.
20m
Place back on the magnetic stand and, after 00:05:00 or when the solution is clear, transfer 10 µL DNA elution to a new plate or tube. Only if possible, use a pipette that can transfer the whole volume in a single maneuver.
20m
Quantify all or a random subset of samples. Fluorometric (such as Qubit) is recommended, but not essential. Nanodrop can be used as well. If so, you will have to scale up the volumes in the overall protocol to allow for the extra amount needed for quantification, or dilute and recover DNA at two steps above with more volume.
Using amounts of the current protocol, concentration after elution is down to 5 ng/µl , having lost approximately half of the DNA in the digestion and clean-up.
Ligation
Ligation
2h
2h
Estimate proper amounts of T4 ligase.
Note
Example for elm:
Based on estimations on simRAD R package, there are 17.5 million cut sites per elm genome 2.1 Gbp.
Considering that 1 Gbp is approximately 1 pg, then we have 8.3 billion cut sites per ng.
For our digestion output we have 420 billion cut sites per sample (50 ng in 10 ul), that translates in 0.66 pmol (in 15 ul will be 0.044 uM).
One unit is defined as the amount of enzyme required to give 50% ligation of HindIII fragments of λ DNA (5´ DNA termini concentration of 0.12 µM, 300- µg/ml) in a total reaction volume of 20 μl in 30 minutes at 16°C in 1X T4 DNA Ligase Reaction Buffer.
Given that T4 Ligase is 400 units/ul, 12 units is 0.03 ul per sample should suffice.
Prepare Adapter Mix.
For EcoRI dilution, mix:
0.9 µL of 20 micromolar (µM) annealed adapters
4.91 µL of 10 X annealing buffer or NaCl
44.19 µL water or TE buffer
For MseI dilution, mix:
30.55 µL of 20 micromolar (µM) annealed adapters
11.95 µL of 10 X annealing buffer or NaCl
107.5 µL water or TE buffer
Finally, mix:
50 µL EcoRI dilution
150 µL MseI dilution
10m
In the plate with the clean-up digested DNA, dispense 2 µL Adapter Mix into each well. You can use same pipette tip, leaving the drop in the upper portion of the well. Spin down the liquids.
10m
Prepare Ligation Mix. Per sample:
1.5 µL of 10 X T4 ligation buffer
0.03 µL of 400000 units/ml T4 ligase
1.47 µL water
For 96 samples (approx. 110x,or even 105x):
165 µL of 10 X T4 ligation buffer
161.7 µL of 400000 units/ml T4 ligase
52.8 µL water
T4 DNA Ligase - 20,000 unitsNew England BiolabsCatalog #M0202S
10m
With the plate On ice , quickly dispense 3 µL Ligation Mix into each well. Mix by pipetting up and down a few times and spinning down.
Alternatively, you can use same pipette tip, leaving the drop in the upper portion of the well. Spin down the liquids. Perhaps, vortexing the covered plate in a plate vortexer. Spin down again. However, we have not try this approach.
20m
Incubate the total volume (15 µL ) at Room temperature or 23 °C for 01:00:00 , then heat-kill at 65 °C for 00:10:00 After the heat-kill, cool the solution at 2 °C per 00:01:30 until it reaches room temperature.
1h 11m 30s
Clean-up
Clean-up
2h 45m
2h 45m
Take an aliquot of magnetics beads, out of the fridge to let them reach Room temperature before initiating the protocol (at least 00:30:00 at Room temperature )
For 96 samples, at least 1.5 mL magnetic beads
30m
Add 15 µL magnetic beads (1X) to each digestion reaction. Mix well by pipetting up and down at least 10 times. If centrifuging samples after mixing, be sure to stop the centrifugation before the beads start to settle down (no more than 1000 rpm).
20m
Incubate samples on bench top for at least 00:05:00 at room temperature.
5m
Place the plate on a magnetic stand, and wait 00:05:00 or until the solution is clear.
While waiting, prepare 35 mL Ethanol 80% (28 mL Ethanol absolute + 7 mL Water ). It has to be freshly prepared.
5m
Carefully remove and discard the supernatant (approximately 27 µL ), not disturbing the beads.
20m
Wash with 160 µL freshly prepared ethanol 80% , not removing the plate from the magnetic stand. Incubate beads in the ethanol for 00:00:30 . Remove and discard the ethanol, but do not disturbing the beads. Some ethanol can remain if so needed not to disturb the beads.
So removing 150 µL ethanol would be enough.
15m
Wash AGAIN with 160 µL freshly prepared ethanol 80% , not removing the plate from the magnetic stand. Incubate beads in the ethanol for 00:00:30 . Remove and discard the ethanol, but do not disturbing the beads. In this step, ethanol remaining should be as little as possible.
Recommendation: remove 160 µL ethanol with 200 µL tips , and afterwards use 10 µL tips to remove the last drops of ethanol.
25m
After removing all visible liquid, let the beads air dry approximately 00:03:00 , but DO NOT over-dry (losing the glossy shine, turning lighter brown and start to crack). Beads may overdry really quick with these volumes. If necessary, do not remove ethanol from all wells at once. Do it in a manner that the first columns of the plate do not overdry, so removing ethanol and adding elution buffer (next step) might intercalate. Normally, half plate can be done with no need of intercalation.
3m
Remove from stand, add 20 µL elution buffer and completely dissolve beads pipetting up and down. Use a pipette that can dispense the complete amount in a single maneuver. Make a very short spin (no more than 1000 rpm) to bring down the elution and let it rest at least 00:02:00 atRoom temperature . Elution buffer could be water, Tris or TE buffers.
20m
Place back on the magnetic stand and, after 00:05:00 or when the solution is clear, transfer 16 µL DNA elution to a new plate or tube. Only if possible, use a pipette that can transfer the whole volume in a single maneuver. In this step, after transferring 8 µL DNA elution to a plate for storage, the remaining 8 µL DNA elution can be transferred straight to the plate for PCR reaction, with Primers already dispensed (see next section).
20m
Amplification
Amplification
3h
3h
In an empty half-plate, dispense to the bottom of each well, avoiding touching the walls, or touching consistently the same side wall:
1.5 µL of 5 micromolar (µM) Primer R 5xx
All the wells of ROW A must have the same Primer, same for B, and the rest. In this way, you will have deployed 8 different primers, each one 12 times.
20m
Now transfer 8 µL cleaned-up ligation product , to the bottom of the wells. That would be no more than 10 ng of non-size-selected DNA.
10m
For a total volume 15 µL , prepare PCR mix:
Per sample, the final volumes :
3 µL of 5 X Buffer
0.3 µL of 10 millimolar (mM) each dNTPs
0.55 µL water
0.15 µL polymerase
1.5 µL of 5 micromolar (µM) Primer F 7xx (to dispense in aliquoted mix)
1.5 µL of 5 micromolar (µM) Primer R 5xx (already dispensed)
PHUSION PLUS DNA POLYMERASE 500 RXNThermo ScientificCatalog #F630L
For half a plate (102x), first mix:
306 µL of 5 X Buffer
30.6 µL of 10 millimolar (mM) dNTPs
56.1 µL water
15.3 µL polymerase
Aliquot the mix in 12 microtubes 34 µL each and add to each 12.75 µL of a different Primer F 7xx. Dispense at the well rims 5.5 µL specific PCR Mix to each well, in the following manner:
Dispense aliquot number 1 to all the wells of the COLUMN 1; aliquot number 2 to COLUMN 2; and so on. Spin down.
If you really want to avoid possible traces of cross-contamination, you should use different pipette tips per well.
30m
Run a PCR amplification following the polymerase recommended program:
98 °C for 00:00:30
12 cycles of: 98 °C for 00:00:10 ; 55 °C for 00:00:10 ; 72 °C for 00:00:20
72 °C for 00:05:00
30m
It is highly recommended to quantify in Qubit and/or agarose gel. Empirically, we obtain 10 ng/µl with 12 cycles .
1h 30m
Pooling and size selection
Pooling and size selection
1h 55m
1h 55m
Combine 4 µL of each successful reaction, into a single 1.5 mL microtube . You may use same pipette tip. Different volumes per sample can be pooled, if yields have been uneven. Let's say total volume of the pool is 384 µL . Transfer 320 µL to a new 1.5 mL microtube .
10m
Choosing fragments between 330bp and 410bp. Adding adapter lengths (140bp total) that would be 470bp to 550bp. In our experience, size selection with Ampure beads should follow then these ratios: 0.5X in the first step and 0.2X in the second; to achieve these sizes. You can try different ratios to evaluate selection ranges.
Take an aliquot of magnetics beads, out of the fride to let them reach Room temperature before initiating the protocol (at least 00:30:00 at Room temperature )
For 96 samples, at least 0.24 mL magnetic beads
30m
Add 160 µL magnetic beads (0.5X) to the PCR pool . Mix well by pipetting up and down at least 10 times. If centrifuging samples after mixing, be sure to stop the centrifugation before the beads start to settle down (no more than 1000 rpm).
3m
Incubate samples on bench top for at least 00:05:00 at room temperature.
5m
Place the plate on a magnetic stand, and wait 00:05:00 or until the solution is clear.
5m
Transfer all the supernatant to a new 1.5 mL microtube , discarding the beads. Tried not to transfer beads since they are attached to large fragments of DNA.
3m
Add 64 µL magnetic beads (0.2X) to the PCR pool supernatant, just transfered in a new tube. Mix well by pipetting up and down at least 10 times. If centrifuging samples after mixing, be sure to stop the centrifugation before the beads start to settle down (no more than 1000 rpm).
3m
Incubate samples on bench top for at least 00:05:00 at room temperature.
5m
Place the plate on a magnetic stand, and wait 00:05:00 or until the solution is clear.
While waiting, prepare 2 mL Ethanol 80% (1.6 mL Ethanol absolute + 0.8 mL Water ). It has to be freshly prepared.
5m
Carefully remove and discard the supernatant, not disturbing the beads.
3m
Wash with 800 µL freshly prepared ethanol 80% , not removing the plate from the magnetic stand. Incubate beads in the ethanol for 00:00:30 . Remove and discard the ethanol, but do not disturb the beads. Some ethanol can remain if so needed not to disturb the beads.
3m
Wash AGAIN with 800 µL freshly prepared ethanol 80% , not removing the plate from the magnetic stand. Incubate beads in the ethanol for 00:00:30 . Remove and discard the ethanol, but do not disturb the beads. In this step, ethanol remaining should be as little as possible. Use 10 µL tips to remove the last drops of ethanol.
3m
After removing all visible liquid, let the beads air dry approximately 00:05:00 , but DO NOT over-dry (losing the glossy shine, turning lighter brown and start to crack).
5m
Remove from stand, add 28 µL elution buffer and completely dissolve beads pipetting up and down. Use a pipette that can dispense the complete amount in a single maneuver. Make a very short spin (no more than 1000 rpm) to bring down the elution and let it rest at least 00:02:00 atRoom temperature . Elution buffer could be water, Tris or TE buffers.
5m
Place back on the magnetic stand and, after 00:05:00 or when the solution is clear, transfer 24 µL DNA elution to a tube.
5m
Quantify in Qubit. Following the protocol, we obtained a concentration higher than 15 ng/µl .
Normally, Illumina services require 20 µL at 10 nanomolar (nM) .
Following the conversion formula at:
For a mean library size of 510bp, it is needed less than 3.5 ng/µl .
20m
Protocol references
Protocol is inspired in Peterson et al. 2012 (10.1371/journal.pone.0037135) and Parchman, Gompert and Buerkle 2011 rfseq protocol, with multiple modifications.