SARS-CoV-2 Amplicon-based Illumina Sequencing Protocol

Use either a SuperScript (step 2) or LunaScript kit (step 3) for cDNA synthesis

Option 1: SuperScipt mastermix

In a clean room, mix 1:1 dNTP and random hexamers. Aliquot 2µL per reaction, and add 11µL RNA, as per the table below. Seal plate and gently mix and centrifuge briefly to collect liquid at the bottom of the well.

Component Volume

50 µM random hexamers 1µL

10mM dNTPs mix (10mM each) 1µL

Total Mastermix volume 2µL

(template RNA) 11µL

Total Reaction volume 13µL

The Mastermix should be prepared in a clean room and added to the nucleic acid in a BSC exclusive for RNA work.

Incubate the reaction in a thermocycler as follows:

Step Temperature Time Cycles

1 65°C 0h 5m 0s 1

2 4°C Hold

Prepare the following SuperScipt mastermix:

Component Volume

SSIV Buffer 4µL

100mM DTT 1µL

RNaseOUT RNase Inhibitor 1µL

SSIV Reverse Transcriptase 1µL

Total Mastermix volume 7µL

(denatured RNA) 13µL

Total Reaction volume 20µL

Add 7µL of mastermix to the denatured RNA from the previous step. Cover the plate with seal, mix gently on a plate mixer, and pulse spin the plate to collect liquid at the bottom of the tube.

The Mastermix should be prepared in in a clean room and added to the denatured RNA in a BSC or workbench exclusive for RNA work.

Incubate in a thermocycler as follows:

Step Temperature Time Cycles

1 42°C 0h 50m 0s 1

2 70°C 0h 10m 0s 1

3 4°C Hold

Option 2: LunaScript mastermix

In a clean room, mix the components as per the table below:

Component Volume

Nuclease-free water 5µL

LunaScript RT SuperMix (5x) 4µL

Total Mastermix volume 9µL

(template RNA) 11µL

Total Reaction volume 20µL

The Mastermix should be prepared in a clean room and added to the nucleic acid in a BSC exclusive for RNA work.

Incubate the reaction in a thermocycler as follows:

Step Temperature Time Cycles

1 25°C 0h 2m 0s 1

2 55°C 0h 20m 0s 1

3 95°C 0h 1m 0s 1

4 4°C 0h 2m 0s 1

Prepare two multiplex PCR mastermixes as follows (1 for each pool):

Component Pool 1 Pool 2

5X Q5 Reaction Buffer 5µL 5µL

10 mM dNTPs 0.5µL 0.5µL

Q5 Hot Start DNA Polymerase 0.25µL 0.25µL

ARTIC Primer Pool 1 or 2 (10µM) 3.6µL 3.6µL

Nuclease-free water 13.15µL 13.15µL

Total Mastermix volume 22.5µL 22.5µL

(cDNA) 2.5µL 2.5µL

Total reaction volume 25µL 2.5µL

Prealiquot 22.5µL of each mastermix (pool1 and pool2) to each plate (pool1 and pool2) accordingly. Add 2.5µL cDNA to each well of both plates. Seal the plate, mix gently and centrifuge briefly to collect liquid at the bottom of the well.

Run the 3.5 hours PCR program for each pool:

Step Temperature Time Cycles

Heat Activation 98°C 0h 0m 30s 1

Denaturation 98°C 0h 0m 15s 35

Annealing 65°C 0h 5m 0s 35

Hold 4°C 1

Pool 12.5µL of each pool 1 and 2 together (total 25μl) in an 0.2 ml 96 well PCR plate.

Perform AMPure XP bead cleanup according to directions, as follows.

Add 25µL of AMPure XP (well-vortexed, room temperature) to the sample plate. Cover the plate with seal, mix gently on a plate mixer, and pulse spin the plate to collect liquid at the bottom of the tube. Incubate at Room temperature for 0h 5m 0s.

Place the plate on a magnetic rack for 0h 5m 0s , or until the beads have pelleted and the supernatant is completely clear.

Remove and discard the liquid from each well with a multichannel pippette, being careful not to touch the bead pellet.

Add 200µL of freshly prepared, room temperature 80% ethanol to each well, incubate for 0h 0m 30s , remove the ethanol carefully with a multichannel pipette.

Repeat ethanol wash (step 6.4).Discard all ethanol and carefully remove as much residual ethanol as possible using a multichannel pipette. With the plate uncovered, incubate for 0h 3m 0s to 0h 5m 0s or until the pellet loses its shine (if the pellet dries completely it will crack and become difficult to resuspend).

Remove from magnetic rack, add28µL of EB buffer to wells and mix gently on a plate mixer, ensuring beads are well re-suspended. Briefly centrifuge the plate to collect the liquid at the bottom of the wells. Incubate at room temperature for 0h 5m 0s .

Place the plate on magnetic rack and incubate for 0h 2m 0s to 0h 5m 0sor until the beads have pelleted and the supernatant is completely clear.

Transfer 25µL of the clear supernatant to a new plate, ensuring no beads are transferred.

Optional section; use remaining volumes from Pool 1 and Pool 2 to confirm amplification by gel electrophoresis.

Prepare 1% agarose gels with enough wells to load all samples. Load 2µLof a 100 bp ladder into gel on either side of each row of wells.

Dispense 2µL of 6X loading dye into each sample with a multichannel pipette, mix and load 2µL of this mix into the gel.

Run at 240V for 0h 20m 0s. Visualize PCR products, confirm bands of approximately 300bp size.

Quantify amplicons using Qubit dsDNA High Sensitivity kit and plate reader according to directions, as follows.

Create Qubit dsDNA HS working solution by mixing 99.5µL X buffer and 0.5µLX dye (X is the total number of samples, including 6 standards). Using a reservoir and multichannel pipette, dispense 98µL into required number of wells of a Costar 3590 flat-bottom plate (or as appropriate for plate reader).

Dilute the clean, pooled amplicons (from step 6.8) 1:10 by mixing3µL of the amplicons in 27µL of nuclease free water.

Make up serial standards using 1:2 dilutions of 10 ng/ul stock (Standard 2) from the Qubit HS. This creates 5 standards in the following concentrations: 10ng/ul 5ng/ul 2.5ng/ul 1.25ng/ul 0.625ng/ulplus Standard 1 0ng/ul .

Mix2µLof diluted amplicons and each of the 6 standards, 98µL of Qubit HS working solution, mix and breifly centrifuge. Use plate reader to obtain concentration reading for each sample and standards. The Qubit standard curve is generated by the Qubit standards.

Based on the amplicon concentration, normalize of all the samples amplicon concentration to 0.2ng/ul.

This can be done by adding 2.5µL of diluted amplicon to a plate with prealiquoted, appropriate amount of nuclease free water.

Prepare sequencing libraries with Nextera XT DNA Library Prep kit at half volume, as follows.

Tagment DNA.

Thaw the following Nextera XT reagents on ice:

Amplicon tagment mix (ATM)

Tagment DNA buffer (TD)

Nextera PCR master mix (NPM)

Invert all reagents 3 - 5 times, followed by pulse spin.

Add the following reagents in order:

5µL of TD buffer

2.5µL of 0.2 ng/ul amplicon (from step 9)

2.5µL of ATM

Cover plate with plate seal, mix gently on plate mixer and centrifuge for 0h 1m 0s.

Incubate in thermocycler with the following steps:

Step Temperature Time Cycles

1 55°C 0h 5m 0s 1

2 10°C Hold

Remove the plate immediately once thermocycler reachs 10°C , and proceed to neutralization.

Add 2.5µL of NT buffer to each well and mix by pipetting up and down for 3 times, briefly spin down the plate and incubate at room temperature for 0h 5m 0s.

PCR Amplification.

Thaw the following reagents on ice:

NPM

Index primers

Resuspension buffer (RSB)

Thaw the index primers, mix by vortexing each vial and spin down the liquid at the bottom of the vials. Option to dispense indexes into 96 well plate for easier pipetting.

Add 7.5µLof Nextera PCR mastermix to each well.

From the pre-aliquoted index plate, add 5µL (2.5µL of each i5 and i7 index) of the corresponding index combination to each well. Cover plate with plate seal, gently mix on plate mixer, and centrifuge for 0h 1m 0s.

Run the PCR program to amplify the libraries:

Step Temperature Time Cycles

1 72°C 0h 3m 0s 1

2 95°C 0h 0m 30s 1

3 95°C 0h 0m 10s 12

3 55°C 0h 0m 30s 12

3 72°C 0h 0m 30s 12

4 72°C 0h 5m 0s 1

5 4°C Hold 1

Repeat the same clean up process as step 6.1-6.8 using 20µL of AMPure XP beads and 28µL of resuspension buffer.

Repeat the same quantification process as Step 8 but do NOT dilute libraries.

Normalize each library to 4nanomolar (nM) by dilution with nuclease free water.

Pool equal volume (e.g. 5µL ) from each of the normalized libraries into a single 1.5 mL microtube.

Verify fragment size and concentration using Agilent D5000 Assay on TapeStation 4200 as follows.

Add 2µL of Sample Buffer and2µL of your pooled libraries in triplicate in a strip tube.

Vortex using the adapter at 2000 rpm for0h 1m 0s .

Load tubes, tapes, and tips into TapeStation. Start run. Using library concentration and fragment size, calculate the molarity of the libraries using the following formula:

Molarity = concentration ng/uL * (1515.1515/fragment size(bp))

Denature and load pooled libraries for MiSeq as follows.

NOTE: Remove sequencing kit components from freezer to thaw at appropriate temperature/time.

Denature the pooled libraries by mixing 5µL of pooled libraries and5µL of freshly made 0.2N NaOH solution.

Incubate for 0h 5m 0s.

Add 990µL of HT1 buffer and mix well with denatured pooled library by pipetting up and down 10 times with P1000.

Load 600µL of the denatured, diluted pooled library into the loading position of the Illumina MiSeq reagent cartridge (V2, 300 cycle kit). Load reagent cartridge, flow cell, and PR2 buffer into Miseq instrument, confirm the metrics and start the run.

For NextSeq loading, combine up to four pools of libraries at equal concentrations. Be careful to use unique index combinations for all pooled samples.

Denature and load pooled libraries for NextSeq as follows.

NOTE: Remove sequencing kit components from freezer to thaw at appropriate temperature/time.

Denature the pooled libraries by mixing 5µL of pooled libraries and5µL of freshly made 0.2N NaOH solution.

Incubate for 0h 5m 0s.

Add 5µL 200 mM TrisHCl pH7.5 and 985µL HT1 buffer,

Pipette 97µL of this denatured library solution into a new tube and add 1203µL of HT1 buffer.

Load 1300µLof the denatured, diluted pooled library into the loading position of the Illumina NextSeq reagent cartridge. Follow the prompts in the instrument to complete loading of flow cell, buffer bottle or cartridge and reagents cartridge