TITLE: Modified NEBNext® VarSkip Long SARS-CoV-2 Enrichment and library prep (Native Barcoding Kit V14 Oxford Nanopore Technologies)- adapted for wastewater samples
AUTHORS: Kathryn Judy, Padmini  Ramachandran, Amanda Windsor, Tamara Walsky, Chris  Grim, Maria Hoffmann

[DESCRIPTION]
This protocol details methods for the preparation of SARS-CoV-2 sequencing library using VSL primers from NEB, adapted for wastewater samples. This protocol produces multiplexed amplicon libraries suitable for sequencing on Oxford Nanopore Technologies® (ONT) MinION systems using ONT V14 chemistry (SQK-NBD114).

[BEFORE_START]
Note: We recommend setting up a no template control reaction and all reactions are set-up in a biological safety cabinet.

The presence of carry-over products can interfere with sequencing accuracy, particularly for low copy targets. Therefore, it is important to carry out the appropriate no template control (NTC) reactions to demonstrate that positive reactions are meaningful.

[GUIDELINES]
Overview 
Sequences and information on the NEBNext VarSkip Long primers can be found at https://github.com/nebiolabs/VarSkip. All other enzymes, buffers, beads and oligos required to convert cDNA into targeted, high quality libraries for next-generation sequencing on the Oxford Nanopore platform are available. 

[STEPS]
SECTION: Before you start
1. 
 

 

SECTION: Targeted cDNA Amplification
2. 
 

SECTION: Targeted cDNA Amplification
3. Prepare master mixes fresh immediately before performing cDNA amplification. 

Q5 Hot Start High-Fidelity Polymerase should stay on ice at all times. Do not vortex.
Thaw Q5 Reaction Buffer, MgCl2, dNTPs, and water.
Mix thawed tubes, spin down, and place on ice.
Thaw VarSkip Long Primer Mix 1 and VarSkip Long Primer Mix 2.  
Mix by flicking and spin down both the tubes.
Keep on ice.

Prepare the split pool amplification reactions as described below:

For Pool set A:
Prepare the master mix below in sufficient volume for your samples.
 

For Pool Set B: 
Prepare the master mix below in sufficient volume for your samples.
 
Mix the two master mix tubes by flicking and spin down. Dispense 7.5 µl master mix from each tube into separate PCR tube strips (A and B), two PCR tubes (one for each master mix) per sample to amplify.

SECTION: Targeted cDNA Amplification
6. Gently flick the tube strips to mix and spin down briefly.
SECTION: Targeted cDNA Amplification
7. Incubate Pool A reactions in a thermocycler* with the following steps:
 

Incubate Pool B reactions in a thermocycler* with the following steps:

 

 

SECTION: Cleanup of cDNA Amplicons
8. We highly recommend this clean up step using AMPure® XP beads, though NEBNext sample purification beads can be used as well. 

 

SECTION: Cleanup of cDNA Amplicons
9. For each sample, combine pool A and pool B PCR products (amplicons), measuring the pooled volume.
SECTION: Cleanup of cDNA Amplicons
10. Vortex AMPure® XP beads for 30 s to resuspend.

SECTION: Cleanup of cDNA Amplicons
11. Add 0.6 X to the combined PCR product.  Mix well by flicking the tube and a very short 2-3 seconds quick centrifugation. Be sure to stop the centrifugation before the beads start to settle out.

SECTION: Cleanup of cDNA Amplicons
12. Incubate samples at Room temperature for 5 min.
SECTION: Cleanup of cDNA Amplicons
13. Quickly spin samples to collect the liquid from the sides of the tube before placing on the magnetic stand for 5 min to separate the beads from the supernatant.

SECTION: Cleanup of cDNA Amplicons
14. After 5 minutes (or when the solution is clear), carefully remove and discard the supernatant. Be careful not to disturb the beads that contain DNA targets.
 


SECTION: Cleanup of cDNA Amplicons
15. Add 200 µL to the tube while in the magnetic stand. Incubate at Room temperature for 30 s, and then carefully remove and discard the supernatant. Be careful not to disturb the beads that contain DNA targets.
SECTION: Cleanup of cDNA Amplicons
16. Repeat previous step once for a total of two washes: 
Add 200 µL to the tube while in the magnetic stand. Incubate at Room temperature for 30 s, and then carefully remove and discard the supernatant. Be careful not to disturb the beads that contain DNA targets. 

Be sure to remove all visible liquid after the second wash. If necessary, briefly spin the tube for 1 s, place back on the magnetic stand and remove traces of ethanol with a p10 pipette tip.

SECTION: Cleanup of cDNA Amplicons
17. Air dry the beads for up to 3 min while the tube is on the magnetic stand with the lid open. 
 

SECTION: Cleanup of cDNA Amplicons
18. Remove the tube from the magnetic stand. Elute the DNA target from the beads by adding 18 µL. 
SECTION: Cleanup of cDNA Amplicons
19. Mix well by flicking the tube followed by a very short centrifugation. Incubate for 5 min at Room temperature. If necessary, quickly spin the sample to collect the liquid from the sides of the tube or plate wells before placing back on the magnetic stand. 
SECTION: Cleanup of cDNA Amplicons
20. Place the tube on the magnetic stand. After 2 min (or when the solution is clear), transfer 17 µL to clean PCR tubes.

SECTION: Cleanup of cDNA Amplicons
21. Assess the concentration of the DNA targets. We recommend using a Qubit fluorometer for concentration assessment. Use 1 µl of sample for the Qubit fluorometer. Amplicons should also be run on Femto or Bioanalyzer® or Tape Station using High Sensitivity (HS) 5000 tape to confirm ~1500-1600 bp size of amplicons.
 



 

SECTION: Targeted cDNA Amplification
4. Add 4.5 µl cDNA into each pre-filled PCR tube, ensuring each sample to be amplified is added into exactly 1 tube in strip A and 1 tube in strip B.
SECTION: Targeted cDNA Amplification
5. While keeping the polymerase on ice, add 0.5 µL Q5 Hot Start High-Fidelity Polymerase to each tube.

SECTION: End-Prep
22. 
Calculate the volume of each sample needed to bring forward at least 200 fmol  per sample using the amplicon size determined after cleanup. We recommend bringing forward approximately the same moles of DNA for each sample.

Aliquot the volume of each sample calculated into fresh PCR tubes and make up each sample to 11.5 µL using nuclease-free. Excess amplicons should be returned to 4 °C
SECTION: End-Prep
23. Make the end repair and dA-tailing master mix by combining the reagents below in the order and amounts listed in the table. Adjust component volumes for your number of samples plus 20% overage.

Thaw DNA Control Sample (DCS) at room temperature, vortex briefly, and place on ice. If this is the first use of the DCS tube, dilute by adding 105 µL, mix gently by pipetting, and spin down. Diluted DCS can be stored at -20 °C after use in Step 24.
Thaw Ultra II End-Prep Buffer at room temperature, then vortex and spin down briefly. 
Thaw Ultra II End Prep Enzyme mix on ice, spin down briefly, and return to ice. Do not vortex.

 
Mix the master mix components by pipetting or gentle flicking and quickly centrifuge. Master mix can remain stable on ice for 4 hours.




SECTION: End-Prep
25. Add 2.5 µL to each sample, mix by gently flicking the tubes, and spin down briefly.

SECTION: End-Prep
24. Add 1 µL to each sample, mix by gentle flicking, and spin down briefly.
SECTION: End-Prep
26. Incubate samples in a thermocycler* with the following settings:

 

SECTION: Native Barcode Ligation and Cleanup
27. 
 

In PCR tubes or a 96-well plate, add reagents below in the order listed in the table.

Thaw Blunt/TA Ligase master mix at room temperature, spin down 5 seconds, then mix with 10 full volume pipette mixes and place on ice. Do not vortex.
Thaw EDTA at room temperature, mix by vortexing, spin down, and place on ice.
Thaw Native Barcodes (ex: NB01-96) required for your number of samples at room temperature, individually mix by pipetting, spin down, and place on ice.
Add a unique barcode to each sample to be run together on a single flowcell (to be pooled).
2-3 barcoding reactions per sample may be desired to increase input DNA. If multiple reactions per sample are performed, be sure to use the same barcode for the same sample in each reaction.

 
Mix components by gently flicking the tubes, then centrifuge briefly.
SECTION: Native Barcode Ligation and Cleanup
28. Incubate samples at Room temperature for 20 min

SECTION: Native Barcode Ligation and Cleanup
29. Add 1 µL to each tube to stop the reaction. Mix well by flicking and spin down briefly.

SECTION: Native Barcode Ligation and Cleanup
30. Pool all barcoded samples to be run on a single flowcell in a 1.5 µl LoBind tube, measuring the volume each sample as it is added. Calculate the final volume of the pooled samples.
SECTION: Native Barcode Ligation and Cleanup
31. Add 0.4 X to each pool and mix by gently flicking followed by a short spin to collect the liquid. Stop the centrifugation before the beads begin to settle.

SECTION: Native Barcode Ligation and Cleanup
32. Incubate at room temperature for 10 min to bind DNA to the beads, agitating the pool(s) every two minutes. If available, pools can be incubated on a Hula mixer (rotator mixer) to agitate the beads instead.

SECTION: Native Barcode Ligation and Cleanup
33. Spin down the pool(s) and place tube(s) in an appropriate magnetic separation rack until the beads have separated, 5 min 

SECTION: Native Barcode Ligation and Cleanup
34. Carefully pipette off the supernatant without disturbing the beads, discarding the supernatant. Do not discard the beads, which contain your DNA target.
SECTION: Native Barcode Ligation and Cleanup
35. Wash the beads with 700 µL. Pipette off the ethanol and discard. If the pellet was disturbed, wait for the beads to pellet again before pipetting off the ethanol. Do not discard the beads.

SECTION: Native Barcode Ligation and Cleanup
36. Repeat the previous step once for a total of two washes:
Wash the beads with 700 µL. Pipette off the ethanol and discard. If the pellet was disturbed, wait for the beads to pellet again before pipetting off the ethanol. Do not discard the beads.
SECTION: Native Barcode Ligation and Cleanup
37. To remove residual ethanol, quickly spin pool(s) and return the tube(s) to the magnetic rack, allowing beads to separate fully. Pipette off residual ethanol with a P20 pipette and discard. Do not discard the beads.
SECTION: Native Barcode Ligation and Cleanup
38. Remove samples from the magnetic rack and immediately add 35 µL. Resuspend beads by flicking, then quickly spin to collect liquid.

SECTION: Native Barcode Ligation and Cleanup
39. Incubate pool(s) at 37 °C for 10 min to elute DNA. Every two minutes, agitate the sample by gentle flicking for 10 seconds to encourage elution.

SECTION: Native Barcode Ligation and Cleanup
40. Place samples on the magnetic rack until beads separate fully from the solution, 5 min 

SECTION: Native Barcode Ligation and Cleanup
41. Slowly pipette 35 µL of clear eluate without disturbing the beads and transfer to new 1.5 µl LoBind tube(s). Discard the old sample tube(s) with beads. Do not discard the supernatant.


 



SECTION: Adapter Ligation and Cleanup
42. 
 
Add the following components to a 1.5 µl LoBind tube.

Thaw NEBNext Quick Ligation Reaction Buffer at room temperature, pipette up and down several times to break up precipitate, vortex for several seconds, then spin down briefly.
Spin down Quick T4 DNA Ligase, pipette mix, and place on ice. Do not vortex.
Spin down Native Adapter, pipette mix, and place on ice.
Thaw Short Fragment Buffer (SFB) at room temperature, mix by vortexing, spin down, and place on ice.

 
Thoroughly mix components by pipetting or gently flicking the tube, then quickly centrifuge to mix.
SECTION: Adapter Ligation and Cleanup
43. Incubate the reaction for 20 min at room temperature.
SECTION: Adapter Ligation and Cleanup
44. Thaw elution buffer at room temperature. 

Add 20 µL (0.4X) to each nuclease-treated sample and mix by gently flicking followed by a short spin to collect the liquid. Stop the centrifugation before the beads begin to settle.
SECTION: Adapter Ligation and Cleanup
45. Incubate at room temperature for 10 min to bind DNA to the beads, agitating every two minutes. If available, the library can be incubated on a Hula mixer (rotator mixer) to agitate the beads instead.

SECTION: Adapter Ligation and Cleanup
46. Spin down and place tube in an appropriate magnetic separation rack until the beads have separated, 5 min 


SECTION: Adapter Ligation and Cleanup
47. Carefully pipette off the supernatant without disturbing the beads, discarding the supernatant. Do not discard the beads, which contain your DNA target.
SECTION: Adapter Ligation and Cleanup
48. Add 125 µL to each sample tube. Flick the tube to resuspend beads, collect liquid with a quick spin, and return the tube to the magnetic rack.

When the beads have pelleted, remove the supernatant and discard without disturbing the beads. Do not discard the beads.

SECTION: Adapter Ligation and Cleanup
49. Repeat the previous step once for a total of two washes:
Add 125 µL to each sample tube. Flick the tube to resuspend beads, collect liquid with a quick spin, and return the tube to the magnetic rack.

When the beads have pelleted, remove the supernatant and discard without disturbing the beads. Do not discard the beads.
SECTION: Adapter Ligation and Cleanup
51. Remove the tube from the magnetic rack and immediately add 15 µL. Resuspend beads by flicking, then quickly spin to collect liquid.

SECTION: Adapter Ligation and Cleanup
52. Incubate at 37 °C for 10 min to elute DNA. Every two minutes, agitate the sample by gentle flicking for 10 seconds to encourage elution.

SECTION: Adapter Ligation and Cleanup
53. Place tube on the magnetic rack until beads separate fully from the eluate, usually less than 5 min 

SECTION: Adapter Ligation and Cleanup
54. Slowly pipette 15 µL of clear supernatant without disturbing the beads and transfer to a new LoBind tube. Discard the old tube with beads. Do not discard the supernatant.
SECTION: Adapter Ligation and Cleanup
55. Measure DNA concentration with a Qubit Fluorometer using the 1x dsDNA HS kit.  The final library should also be run on Femto or Bioanalyzer® or Tape Station using High Sensitivity (HS) 5000 tape to confirm the size of the final library.

 

SECTION: End Protocol
56. Based on the concentration and library size determined in step 55, aliquot 10-20 fmol and make up to 12 µL using nuclease-free water in a new 0.5 µl LoBind tube. This will be used to load the flowcell.

Prime and load the R10.4.1 (FLO-MIN114) SpotON flowcell following the Oxford Nanopore SQK-LSK114 protocol. We recommend using High Accuracy Basecalling (HAC) at 260bps ("Accurate") speed.
SECTION: Adapter Ligation and Cleanup
50. To remove residual Short Fragment Buffer, quickly spin the tube and return to the magnetic rack, allowing beads to separate fully. Pipette off residual supernatant with a P20 pipette and discard. Do not discard the beads.