TITLE: Double digestion RADseq library
AUTHORS: David Macaya-Sanz

[DESCRIPTION]
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. 

[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. 

[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.

[STEPS]
SECTION: DNA dilution
3. Measure your DNA stock concentration. Recommended Nanodrop One.
 

SECTION: DNA dilution
4. Dilute DNA stock to 20 ng/µl at minimum volume of 7.5 µL 

4. 
SECTION: Annealing of adapters
5. 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. 
SECTION: Pooling and size selection
46. Combine 4 µL of each successful reaction, into a single 1.5 mL. 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.

SECTION: Ligation
25. Estimate proper amounts of T4 ligase.

 

SECTION: Clean-up
14. 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 30 min at Room temperature )
For 96 samples, at least 1 mL 
SECTION: Double digest
13. Incubate at 37 °C for 90 min . Hold at 4 °C afterwards. Do not heat kill enzymes.
Reactions can be stored at 4 °C   

SECTION: Double digest
11. On a 96-well PCR plate, dispense 2.5 µL in empty PCR tubes or plate wells, keeping it on ice 
Spin the plate, to bring to the bottom the digestion mix.
SECTION: Double digest
10. Prepare Digestion Mix adding 0.9 units , 0.45 units and 1 µL per sample. Then, add water to get a final volume of 2.5 µL per sample. 

For example, for 48 samples, prepare mix in excess, so factor would be 110x:
5 µL at 20000 units/ml total 100 units 
5 µL at 10000 units/ml total 50 units 
110 µL at 10 X 
155 µL (you can add some extra water: such as 5 µL 

  
  
SECTION: Double digest
12. Transfer 7.5 µL 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.
 
SECTION: Annealing of adapters
9. Incubate each oligo combination in thermal cycler: 97.5 °C 150 s followed by a touchdown of 1 °C 20 s until reaching 21 °C . Hold afterwards at 4 °C 

SECTION: Annealing of adapters
8. Combine oligos MseI_P2.1_2N and MseI_P2.2_2N, in the same fashion as with EcoRI_P1.
SECTION: Annealing of adapters
7. Combine oligos EcoRI_P1.1 and EcoRI_P1.2 into the same stock to a final 20 micromolar (µM) 
Therefore, add 5 µL, 5 µL, 2.5 µL, and 12.5 µL 

In case of TE buffer ready: add 5 µL, 5 µL, 2.5 µL, and 12.5 µL 
SECTION: Annealing of adapters
6. Create Annealing buffer stock (10x) (it is TE buffer (10x) plus the sodium chloride):
100 millimolar (mM) 
10 millimolar (mM) 
500 millimolar (mM) 

In a 1L flask:
800 mL 
15.759 g 
2.922 g 
29.22 g 
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.
SECTION: Clean-up
15. Add 10 µL  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). 
SECTION: Clean-up
16. Incubate samples on bench top for at least 5 min  at room temperature.
SECTION: Clean-up
17. Place the plate on a magnetic stand, and wait 5 min or until the solution is clear. 
While waiting, prepare 35 mL (28 mL + 7 mL). It has to be freshly prepared.

SECTION: Clean-up
18. Carefully remove and discard the supernatant (approximately 17 µL ), not disturbing the beads.
SECTION: Clean-up
19. Wash with 160 µL , not removing the plate from the magnetic stand. Incubate beads in the ethanol for 30 s . 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  in two pipetting moves.
SECTION: Clean-up
21. After removing all visible liquid, let the beads air dry approximately 3 min , 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.  
SECTION: Clean-up
20. Wash AGAIN with 160 µL , not removing the plate from the magnetic stand. Incubate beads in the ethanol for 30 s . 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 with 200 µL , and afterwards use 10 µL to remove the last drops of ethanol.
SECTION: Clean-up
22. Remove from stand, add 13 µL 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  2 min atRoom temperature . Elution buffer could be water, Tris or TE buffers. 
SECTION: Clean-up
23. Place back on the magnetic stand and, after 5 min or when the solution is clear, transfer 10 µL to a new plate or tube. Only if possible, use a pipette that can transfer the whole volume in a single maneuver.
SECTION: Clean-up
24. 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.
SECTION: Ligation
30. Incubate the total volume (15 µL ) at Room temperature or 23 °C  for 60 min, then heat-kill at 65 °C for 10 minAfter the heat-kill, cool the solution at 2 °C per 90 s until it reaches room temperature.
SECTION: Ligation
26. Prepare Adapter Mix.
For EcoRI dilution, mix:
0.9 µL  of 20 micromolar (µM) 
4.91 µL of 10 X 
44.19 µL

For MseI dilution, mix:
30.55 µL  of 20 micromolar (µM) 
11.95 µL of 10 X 
107.5 µL

Finally, mix:
50 µL 
150 µL 
SECTION: Ligation
27. In the plate with the clean-up digested DNA, dispense 2 µL into each well. You can use same pipette tip, leaving the drop in the upper portion of the well. Spin down the liquids.
SECTION: Ligation
28. Prepare Ligation Mix. Per sample:
1.5 µL of 10 X 
0.03 µL of 400000 units/ml 
1.47 µL 

For 96 samples (approx. 110x,or even 105x):
165 µL of 10 X 
161.7 µL of 400000 units/ml 
52.8 µL 

  
SECTION: Ligation
29. With the plate on ice , quickly dispense 3 µL 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.
SECTION: Clean-up
31. Take an aliquot of magnetics beads, out of the fridge to let them reach Room temperature  before initiating the protocol (at least 30 min at Room temperature )
For 96 samples, at least 1.5 mL 
SECTION: Clean-up
32. Add 15 µL  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).
SECTION: Clean-up
33. Incubate samples on bench top for at least 5 min  at room temperature.
SECTION: Clean-up
34. Place the plate on a magnetic stand, and wait 5 min or until the solution is clear. 
While waiting, prepare 35 mL (28 mL + 7 mL). It has to be freshly prepared.

SECTION: Clean-up
35. Carefully remove and discard the supernatant (approximately 27 µL ), not disturbing the beads.
SECTION: Clean-up
36. Wash with 160 µL , not removing the plate from the magnetic stand. Incubate beads in the ethanol for 30 s . 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 would be enough.
SECTION: Clean-up
37. Wash AGAIN with 160 µL , not removing the plate from the magnetic stand. Incubate beads in the ethanol for 30 s . 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 with 200 µL , and afterwards use 10 µL to remove the last drops of ethanol.
SECTION: Clean-up
38. After removing all visible liquid, let the beads air dry approximately 3 min , 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.
SECTION: Clean-up
39. Remove from stand, add 20 µL 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  2 min atRoom temperature . Elution buffer could be water, Tris or TE buffers. 
SECTION: Clean-up
40. Place back on the magnetic stand and, after 5 min or when the solution is clear, transfer 16 µL 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 to a plate for storage, the remaining  8 µL can be transferred straight to the plate for PCR reaction, with Primers already dispensed (see next section).
SECTION: Amplification
43. For a total volume 15 µL, prepare PCR mix:
Per sample, the final volumes : 
3 µL of 5 X 
0.3 µL of 10 millimolar (mM) 
0.55 µL 
0.15 µL 
1.5 µL of 5 micromolar (µM)  (to dispense in aliquoted mix)
1.5 µL of 5 micromolar (µM)  (already dispensed)

  

For half a plate (102x), first mix:
306 µL of 5 X 
30.6 µL of 10 millimolar (mM) 
56.1 µL 
15.3 µL 

Aliquot the mix in 12 microtubes 34 µL  and add to each 12.75 µL of a different Primer F 7xx. Dispense at the well rims 5.5 µL 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.
SECTION: Amplification
44. Run a PCR amplification following the polymerase recommended program:
98 °C for 30 s 
12 cycles of: 98 °C for 10 s ; 55 °C for 10 s ; 72 °C for 20 s 
72 °C for 5 min 
SECTION: Amplification
41. 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) 
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. 
SECTION: Amplification
42. Now transfer 8 µL , to the bottom of the wells. That would be no more than 10 ng of non-size-selected DNA.
SECTION: Amplification
45. It is highly recommended to quantify in Qubit and/or agarose gel. Empirically, we obtain 10 ng/µl with 12 cycles .
SECTION: Pooling and size selection
47. 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.
SECTION: Pooling and size selection
48. Take an aliquot of magnetics beads, out of the fride to let them reach Room temperature  before initiating the protocol (at least 30 min at Room temperature )
For 96 samples, at least 0.24 mL 
SECTION: Pooling and size selection
49. Add 160 µL 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).
SECTION: Pooling and size selection
50. Incubate samples on bench top for at least 5 min  at room temperature.
SECTION: Pooling and size selection
55. Place the plate on a magnetic stand, and wait 5 min or until the solution is clear. 
While waiting, prepare 2 mL (1.6 mL + 0.8 mL). It has to be freshly prepared.
SECTION: Pooling and size selection
56. Carefully remove and discard the supernatant, not disturbing the beads.
SECTION: Pooling and size selection
57. Wash with 800 µL , not removing the plate from the magnetic stand. Incubate beads in the ethanol for 30 s . Remove and discard the ethanol, but do not disturb the beads. Some ethanol can remain if so needed not to disturb the beads. 

SECTION: Pooling and size selection
58. Wash AGAIN with 800 µL , not removing the plate from the magnetic stand. Incubate beads in the ethanol for 30 s . 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 to remove the last drops of ethanol.
SECTION: Pooling and size selection
59. After removing all visible liquid, let the beads air dry approximately 5 min , but DO NOT over-dry (losing the glossy shine, turning lighter brown and start to crack). 
SECTION: Pooling and size selection
60. Remove from stand, add 28 µL 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  2 min atRoom temperature . Elution buffer could be water, Tris or TE buffers. 
SECTION: Pooling and size selection
61. Place back on the magnetic stand and, after 5 min or when the solution is clear, transfer 24 µL to a tube. 
SECTION: Pooling and size selection
53. Add 64 µL 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).
SECTION: Pooling and size selection
54. Incubate samples on bench top for at least 5 min  at room temperature.
SECTION: Pooling and size selection
51. Place the plate on a magnetic stand, and wait 5 min or until the solution is clear. 
SECTION: Pooling and size selection
52. Transfer all the supernatant to a new 1.5 mL, discarding the beads. Tried not to transfer beads since they are attached to large fragments of DNA.
SECTION: Pooling and size selection
62. 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:
 https://knowledge.illumina.com/library-preparation/dna-library-prep/library-preparation-dna-library-prep-reference_material-list/000001240
For a mean library size of 510bp, it is needed less than  3.5 ng/µl .


SECTION: Oligo sequences
1. Make sure you have the required adapter oligos.

 

SECTION: Oligo sequences
2. Make sure you have the required PCR primers oligos.