TITLE: Sub-ARTIC Illumina SARS-CoV-2 Spike sequencing protocol (LoCost) V3.2
AUTHORS: Gavin Horsburgh, Paul Parsons, Kathryn Maher, Steve      Paterson, Terry Burke

[DESCRIPTION]
This protocol describes a procedure for sequencing the Spike region of SARS-CoV-2 using short amplicons (146-208bp). The method has proved to successful with both clinical RNA samples and degraded wastewater samples. The primers are unique to this method. The library prep procedure has been heavily adapted from the ncov-2019 sequencing v3 (ARTIC) protocol by Josh Quick (https://www.protocols.io/view/ncov-2019-sequencing-protocol-v3-locost-bh42j8ye) and the "low cost" method from the NEOF Liverpool Illumina ARTIC protocol.

[BEFORE_START]
Before starting, generate the "odd" and "even" primer pools as follows:

1. Fully resuspend lyophilised oligonucleotides in 1x TE to a concentration of 100 micromolar (µM) , vortex thoroughly and spin down 
2. Sort the odd and even primer sets into separate batches and label two 1.5-ml tubes 
3. Starting with the even primer set add the volume (μl) given overleaf to the pooled 1.5-ml tube (between 7.5 - 15). Repeat with the odds. Vortex and spin down. These are your pooled stocks. 
4. Dilute the pools one in ten across several aliquots with molecular grade water. Vortex and spin down. These are your working primer pools.  


[STEPS]
SECTION: Multiplex PCR
2. Primers are separated into two pools, odd and even, depending on where they sit across the Spike region. See the guidelines section for further details. In a clean pre-PCR hood set up two PCR reactions, one per pool as follows:


 

SECTION: NEBNext Ultra II End Prep
3. Prepare a master mix of the reagents as below by multiplying volumes by the number of samples; add 10% to allow for pipetting error.



 

SECTION: Adapter Ligation
4. Prepare adapter ligation master mix by adding volumes as detailed below for each sample; adding 10% to allow for pipetting error.


 

SECTION: Magnetic Bead clean up
5. Increase volume of sample from 19.7 µL to 25 µL by the addition of nuclease-free water.

SECTION: Addition of NEBNext indexes
6. Add the following components to separate wells of a PCR plate. Make sure to use the indexes in appropriate wells so that each sample has unique i5 and i7 indexes. If using less than 12 samples, the i5 primer can be replaced by the Universal PCR primer.


 
* if using 12 samples or fewer replace index primer i5 with the Universal primer.

SECTION: Pool
7. Pool 5 µL of each sample together in a fresh low-binding microfuge tube and mix. This will give a final volume of 60 µL for twelve samples. If larger numbers of samples are used then pool 5 µL of each and mix. Split into aliquots of about 120 µL.

SECTION: qPCR
8. Perform qPCR by using a Kapa Library Quant kit for Illumina from Roche. Reactions are scaled down to 10 µL. Use the appropriate Kapa kit for the type of qPCR machine used, see https://pim-eservices.roche.com/eLD/api/downloads/ca670ceb-fb38-eb11-0291-005056a71a5d?countryIsoCode=pi

SECTION: Library dilution
9. Take 5 µL of 4 Nanomolar (nM) dilution of library and add 5 µL of 0.1 Molarity (M) NaOH to denature the DNA and incubate at 37 Room temperature for 5 min. 

SECTION: Multiplex PCR
2.1. Add 4.5 µL of cDNA to respective wells to give a total volume of 12.5 µL. Include negative controls. Mix by pipetting, seal plate and centrifuge briefly.

SECTION: Multiplex PCR
2.2. Perform PCR using the following program (with heated lid):


 

 

SECTION: PCR pooling
2.3. In a freshly bleached post-PCR hood that has been subjected to UV for 40 min, combine Pool Even and Pool Odd to give 20 µL in each tube/well. Add 20 µL nuclease-free water to dilute products 1 in 2.
 

SECTION: NEBNext Ultra II End Prep
3.1. Mix well by pipetting and centrifuge briefly.
SECTION: NEBNext Ultra II End Prep
3.2. Combine 2 µL of End Prep master mix with 10 µL of amplified cDNA and mix by pipetting. Spin down, seal, place in a thermocycler and run the following program:


 

SECTION: Adapter Ligation
4.1. Add 6.7 µL ligation mix to each 12 µL amplified cDNA/mastermix and mix by pipetting. Incubate at 20 °C for 15 min.

SECTION: Adapter Ligation
4.2. Add 1 µL of USER enzyme to the ligation mixture. Mix by pipetting, centrifuge briefly and incubate at 37 °C for 15 min. 
SECTION: Magnetic Bead clean up
5.1. Perform a 0.9x Ampure XP bead clean by adding 22.5 µL of Ampure XP beads and mix by pepetting. Incubate for 5 min at 37 Room temperature.

SECTION: Magnetic Bead clean up
5.2. Transfer tube/plate to magnet and allow beads to clump for 5 min. Remove supernatant taking care not to disturb pellet, and discard.

SECTION: Magnetic Bead clean up
5.3. Wash the beads with 100 µL 80% ethanol for 30 s then remove ethanol by pipetting. Repeat one more time making sure to remove any residual ethanol.

SECTION: Magnetic Bead clean up
5.4. Allow beads to air dry for around  5 min; ensure that all the ethanol has evaporated. Do not let beads dry to the point of cracking, as this could affect the amount of DNA recovered.

SECTION: Magnetic Bead clean up
5.5. Add 17 µL of nuclease-free water to each well and mix by pipetting. Allow to stand for 5 min then transfer to magnet and allow beads to clump for 5 min. Remove 15 µL into a fresh low-bind tube.

SECTION: Magnetic Bead clean up
5.6.  Perform Qubit assay to determine concentration.
SECTION: Addition of NEBNext indexes
6.1. Mix thoroughly by pipetting and centrifuge briefly.
SECTION: Addition of NEBNext indexes
6.2. Place on thermocycler and perform PCR using the following program:


 
*Refer to NEBNext protocol for number of cycles required. (Examples: 100 ng = ~3 cycles, 50 ng = ~3–4 cycles, 10 ng = ~6–7 cycles)
SECTION: Pool
7.1. Perform a 0.7x bead clean by adding 42 µL or 84 µL magnetic beads, depending on sample quantity. Leave for 5 min to allow beads to bind DNA. 

SECTION: Pool
7.2. Place on a magnet and allow beads to clump for 5 min. Remove supernatant and discard.

SECTION: Pool
7.3. Wash pellet with 100 µL 80% ethanol for 30 s and remove. Repeat one more time. Allow the ethanol to dry off for about 5 min, making sure the pelleted beads do not dry out, which can reduce DNA recovery.

SECTION: Pool
7.4. Add 30 µL of nuclease-free water and mix by pipetting. Leave for 5 min at 37 Room temperature to allow DNA to elute from the beads. 

SECTION: Pool
7.5. Place the tube/plate on the magnet and let the beads clump for 5 min. Elute DNA into a fresh low-binding tube. 

SECTION: Pool
7.6. Perform TapeStation assay to determine if primer dimers are present. If present, perform another 0.7x bead clean and check again.
SECTION: qPCR
8.1. Enter the values obtained from qPCR into the KAPA Library Quantification Data Analysis Template. 



SECTION: Library dilution
9.1. Neutralise with 5 µL of 200 Milimolar (mM) Tris-HCl pH 7.0 for 1 min. 

SECTION: Library dilution
9.3. Take35 µL of 20 Picomolar (pM) sample and add 465 µL of HT1. This will give 500 µL of 1.4 Picomolar (pM)sequencing mix.

SECTION: cDNA prep
1. In a freshly bleached Pre-PCR hood (ideally in an isolated clean room), that has been subjected to UV irradiation for 40 min, add LunaScript and RNA sample to a tube/well as follows (making sure to include negative controls):


 

SECTION: cDNA prep
1.1. Mix thoroughly by pipetting up and down several times, seal plate, and centrifuge briefly.
SECTION: cDNA prep
1.2. Incubate the reaction as follows (with heated lid):


 

SECTION: Library dilution
9.2. Add 985 µL cold HT1 (hybridisation buffer) to give 20 Picomolar (pM) denatured library

SECTION: PhiX control
10. Thaw a tube of 10 Nanomolar (nM) PhiX stock .

SECTION: PhiX control
10.1. Combine the following volumes in a microcentrifuge tube:
10 Nanomolar (nM) PhiX (10 μl) and RSB (Resuspension Buffer) (15 μl). The total volume is 25 μl at 4 Nanomolar (nM). 
Vortex briefly and then pulse centrifuge. The 4 Nanomolar (nM) PhiX can be stored frozen for 3 months.

SECTION: PhiX control
10.2. Combine 5 µL of 4 Nanomolar (nM) PhiX with 5 μl of 0.1 Molarity (M) NaOH. Vortex briefly and pulse centrifuge. Incubate at 37 Room temperaturefor 5 min

SECTION: PhiX control
10.3. Add 5 µL of 200 Nanomolar (nM) Tris-HCl (pH 7.0), vortex briefly, pulse centrifuge and incubate at 37 Room temperature for 1 min

SECTION: PhiX control
10.4. Add 985 µLof HT1 (hybridisation buffer) to give a concentration of 20 Picomolar (pM). Vortex briefly and pulse centrifuge.

SECTION: MiniSeq Sequencing
11. Combine your library with PhiX control to give a final dilution of PhiX of 5%
SECTION: PCR pooling
2.4. Check the quality and concentration of negatives and a selection of samples using a fluorometer and/or Agilent TapeStation. 


 

SECTION: qPCR
8.2. Dilute sample to 4 Nanomolar (nM).
SECTION: PhiX control
10.5. PhiX control at 20 Picomolar (pM) can be frozen at -20°C for up to two weeks. After this time cluster numbers tend to decrease.