COVID-19 ARTIC v4.1 Illumina library construction and sequencing protocol - tailed method

Important! This step must be performed in a RNase free, pre-PCR environment in which post PCR COVID-19 amplicons are not present, to minimise risk of sample contamination.

Decontaminate bench surfaces, pipettes and gloves with RNase ZAP before starting work. Keep reagents and samples chilled throughout the process.

Defrost PCR plate containing 10µL extracted RNA On ice.

Prepare RT mastermix in a dedicated UV treated pre-PCR area to minimise contamination risk.

A	B	C
RT Master Mix	Vol / RXN (µL)	Vol/384 RXN (µL) inc. excess
LunaScript Super Mix	4	1843
Nuclease-free water	6	2765
Total	10	4608

Mix thoroughly by vortexing.

Use the SPT Labtech Dragonfly Discovery to dispense 10µLof RT mastermix into the PCR plate containing 10µL extracted RNA.

Seal plate and place on a BioShake plate shaker for 30 seconds at 1500rpm to mix. Briefly centrifuge plate.

Place plate on a thermocycler and run the following program:

A	B
Temperature	Time
25°C	2 minutes
55°C	20 minutes
95°C	1 minute
4°C	∞
Lid temp: Tracking

PAUSE POINT cDNA can be stored at 4°C (same day) or -20°C (up to a week).

Prepare the following mastermixes:

A	B	C
Weighted PCR Primer Pool 1 Master Mix	Vol/PCR RXN (µl)	Vol/384 plate (µl) inc. excess
Q5 Hotstart 2X Master Mix	12.5	5760
Primer Pool 1 (mean 102nM)	3.6	1659
Nuclease-free water	2.9	1336
Total	19	8755

A	B	C
Weighted PCR Primer Pool 2 Master Mix	Vol/PCR RXN (µl)	Vol/384 plate (µl) inc. excess
Q5 Hotstart 2X Master Mix	12.5	5760
Primer Pool 2 (mean 102nM)	3.6	1659
Nuclease-free water	2.9	1336
Total	19	8755

Note

The equivolume primer pools used in the standard protocol are of 10micromolar (µM)cumulative concentration, therefore each of the 98 primers in each pool is at 102nanomolar (nM)in the pool and at 15nanomolar (nM)in the final reaction. With the rebalanced primer pools, for equivalency we dilute them such that the average primer concentration is 102nanomolar (nM), and therefore the average concentration of each primer in the final reaction is also 15nanomolar (nM).

Mix thoroughly by vortexing.

Use the SPT Labtech Dragonfly Discovery to dispense 19µLmastermix per well into 2x384 well plates.

Use the Agilent Bravo to add 6µL of cDNA template to each primer pool reaction and mix.

Heat seal and place the plates onto a thermocycler and run the following program.

Important! Heat seal to minimise evaporation.

Note: Amplification should ideally be performed in a different lab to minimise the risk of contaminating other samples.

A	B	C
Step	Temperature	Time
1	98°C	30 seconds
2	95°C	15 seconds
3	63°C	5 minutes
4	Repeat steps 2 & 3 for a total of 35 cycles
5	4°C	∞

PAUSE POINT Amplified cDNA can be stored at 4°C (overnight) or -20°C (up to a week).

Defrost two UDI tag plates (one containing each tail primer pool), both of which should contain the same i5 and i7 barcodes per well.

Use the SPT Labtech Mosquito LV to transfer 100nl of amplified pool 1 cDNA into the UDI tag plate containing the pool 1 tailed primers and 100nlof amplified pool 2 cDNA into the UDI tag plate containing the pool 2 tailed primers, maintaining the same well locations throughout. Immediately proceed to the next step.

Use the SPT Labtech Dragonfly Discovery to dispense 6.25µL of Kapa HiFi 2X Mastermix into each well of both UDI tag plates, and place On iceimmediately. The dispense is sufficient to mix all the reagents.

Heat seal and place the two plates onto a thermocycler and run the following program.

Important! Heat seal to minimise evaporation.

A	B	C
Step	Temperature	Time
1	95°C	5 minutes
2	98°C	30 seconds
3	61°C	20 minutes
4	72°C	2 minutes
Repeat steps 2-4 once more
5	98°C	30 seconds
6	65°C	30 seconds
7	72°C	2 minutes
Repeat steps 5-7 six more times
8	72°C	5 minutes
9	4°C	∞

PAUSE POINT Amplified cDNA can be stored at 4°C (overnight) or -20°C (up to a week).

In a post-PCR lab, use the Agilent Bravo to combine and mix 5µL of pool 1 and pool 2 PCR2 reactions per sample into one plate.

Use the Hamilton STAR to combine 3µL of each sample to form an equivolume pool of 384 samples.

Allow AMPure XP beads to equilibrate to room temperature (~30 minutes). Ensure solution is homogenous prior to use, mixing gently by inversion.

The Hamilton STAR will perform a 0.8X SPRI clean-up and elute the final pool in 1ml elution buffer as follows:

Add 0.8X volume of SPRI beads per pool tube, mix well by pipetting.

Incubate for 0h 3m 0s at 20Room temperature

Transfer tube to magnet, allow 0h 5m 0sfor the beads to form a pellet.

Carefully transfer supernatant into a new tube, taking care not to disturb the bead pellet.

Incubate for 0h 6m 0s at 20Room temperature.

Transfer the tube to a magnet, allow 0h 4m 0sfor the beads to form a pellet.

Carefully remove and discard the supernatant, taking care not to disturb the bead pellet.

Wash the beads with 500µL 75% ethanol for 0h 0m 15s then carefully remove ethanol and discard.

(First wash)

Wash the beads with 500µL 75% ethanol for 0h 0m 15s then carefully remove ethanol and discard.

(Second wash)

Wash the beads with 500µL 75% ethanol for 0h 0m 15s then carefully remove ethanol and discard.

(Third wash)

Allow beads to dry for 0h 5m 0s.

Remove tube from magnet and resuspend beads in 200µL elution buffer, mix well by pipetting.

qPCR

Quantify samples in triplicate using the KAPA Complete kit (Universal) for Illumina (KK4824) plus the KAPA Library Quantification Dilution Control (KK4906).

We use the SPT Labtech Mosquito LV to stamp library pools in triplicate into a 384 assay plate, and the Agilent Bravo to setup the qPCR reactions (1:1600 dilution).

qPCR is performed on the Roche LightCycler 480.

Agilent Bioanalyzer

Prepare 3 dilutions of the equivolume pool (1:10, 1:100, 1:1000). Run 1µl of each dilution in triplicate using the High Sensitivity DNA assay kit.

Confirm size distribution is as expected, check there is no primer-dimer present.

Agilent TapeStation

Run 1µl of each pool in triplicate using the Agilent D5000 ScreenTape System.

Confirm size distribution is as expected, check there is no primer-dimer present.

Set the region size range to 250-1500bp to quantify the pool.

The following protocol is for loading a NovaSeq. We currently plex up to 384 samples per NovaSeq SP lane.

Steps must be performed within a given timeframe or data quality may be affected. Therefore, ensure the instrument is washed, waste containers emptied and ready for use prior to beginning step 46.

Defrost Illumina NovaSeq SP SBS and cluster reagent cartridges for 2-4 hours in a 20Room temperature water bath. Use a lint free tissue to blot any water present on the foil seal. Gently mix cartridges 10X by inversion. Gently tap the bottom of the cartridges on the bench to reduce air bubbles.

Defrost components DPX1, DPX2 and DPX3 from a NovaSeq XP-2 lane kit, then keep 20On ice

Bring flow cell to 20Room temperature (~10 minutes) prior to use.

18µL of each 1nanomolar (nM) pool is required per SP lane.

Denature pools by adding4µL 0.2N NaOH per 18µl. Vortex briefly to mix.

Incubate at 20Room temperature for 0h 8m 0s

Add 5µL 400mM Tris-HCl, pH8.0 to each tube to neutralise the reaction. Vortex briefly to mix, then keep 20On ice.

Important! Use mastermix within 1h 0m 0sof preparation for optimal sequencing performance.

Prepare ExAmp mastermix on ice:

A	B
ExAmp Master Mix	Volume per SP flow cell (µl)
DPX1	126
DPX2	18
DPX3	66
Total	210

Vortex0h 0m 30sto mix, then centrifuge briefly up to 280x g,0h 0m 0s

Add 63µL ExAmp mastermix to each denatured pool, mix well by pipetting.

Prepare the flowcell for sample loading by placing into the flow cell dock with the 2-lane manifold clamped in place.

Pipette 80µL of library + ExAmp pool mix per manifold well. Wait for approximately 2 minutes to allow the solution to fill the lane.

Important! The sequencing run must be started within 0h 30m 0s of libraries being loaded onto the flow cell.

Unclamp the flow cell dock and discard the manifold. Load the flow cell onto the NovaSeq flow cell stage.

Load the SBS and cluster reagent cartridges.

Start sequencing run (250PE).