Nov 04, 2020
Version 5
COVID-19 ARTIC v3 Illumina library construction and sequencing protocol V.5
- DNA Pipelines R&D1,
- Benjamin Farr1,
- Diana Rajan1,
- Emma Dawson1,
- Lesley Shirley1,
- Michael Quail1,
- Naomi Park1,
- Nicholas Redshaw1,
- Iraad F Bronner1,
- Louise Aigrain1,
- Scott Goodwin1,
- Scott Thurston1,
- Stefanie Lensing1,
- James Bonfield1,
- Keith James1,
- Nicholas Salmon1,
- Charlotte Beaver1,
- Rachel Nelson1,
- David K. Jackson1,
- Alex Alderton1,
- Ian Johnston1
- 1Wellcome Sanger Institute
- Coronavirus Method Development Community
- COG-UK
Protocol Citation: DNA Pipelines R&D, Benjamin Farr, Diana Rajan, Emma Dawson, Lesley Shirley, Michael Quail, Naomi Park, Nicholas Redshaw, Iraad F Bronner, Louise Aigrain, Scott Goodwin, Scott Thurston, Stefanie Lensing, James Bonfield, Keith James, Nicholas Salmon, Charlotte Beaver, Rachel Nelson, David K. Jackson, Alex Alderton, Ian Johnston 2020. COVID-19 ARTIC v3 Illumina library construction and sequencing protocol. protocols.io https://dx.doi.org/10.17504/protocols.io.bibtkann
License: This is an open access protocol distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited
Protocol status: Working
We use this protocol and it's working
Created: July 08, 2020
Last Modified: November 04, 2020
Protocol Integer ID: 38995
Keywords: COVID-19, SARS-Cov-2, amplicon sequencing, ARTIC, Illumina library construction, coronavirus
Abstract
This SOP describes the procedure for generating cDNA from SARS-CoV-2 viral nucleic acid extracts and subsequently producing 400nt amplicons tiling the viral genome using V3 nCov-2019 primers (ARTIC). This is followed by library construction, equivolume pooling of samples and quantitation, prior to sequencing on the Illumina NovaSeq.
It is adapted from the nCov-2019 sequencing protocol which can be found here:
https://www.protocols.io/view/ncov-2019-sequencing-protocol-bbmuik6w
Guidelines
It is vital cDNA setup is performed in a laboratory in which post-PCR COVID-19 amplicons are not present, to minimise any risk of sample contamination.
Note: Throughout the protocol we have indicated the liquid handling automation in use at Sanger for specific parts of the process. However, these steps could be performed on alternative liquid handlers or manually.
Materials
MATERIALS
NEBNext Ultra II DNA Library Prep Kit for Illumina - 96 rxnsNew England BiolabsCatalog #E7645L
2x Kapa HiFi Hotstart Readymix Kapa BiosystemsCatalog #KK2602
LunaScript RT SuperMix KitNew England BiolabsCatalog #
E3010L
Illumina Library Quantitation Complete kit (Universal)Kapa BiosystemsCatalog #KK4824
NEB Q5® Hot Start High-Fidelity 2X Master MixNew England BiolabsCatalog #M0494L
AccuClear® Ultra High Sensitivity dsDNA Quantitation Kit with DNA StandardsBiotiumCatalog ##31028
Primer pool sequences (v3) can be found here:
https://github.com/joshquick/artic-ncov2019/blob/master/primer_schemes/nCoV-2019/V3/nCoV-2019.tsv
Protocol materials
NEBNext Ultra II DNA Library Prep Kit for Illumina - 96 rxnsNew England BiolabsCatalog #E7645L
2x Kapa HiFi Hotstart Readymix Kapa BiosystemsCatalog #KK2602
LunaScript RT SuperMix KitNew England BiolabsCatalog #
E3010L
Illumina Library Quantitation Complete kit (Universal)Kapa BiosystemsCatalog #KK4824
NEB Q5® Hot Start High-Fidelity 2X Master MixNew England BiolabsCatalog #M0494L
AccuClear® Ultra High Sensitivity dsDNA Quantitation Kit with DNA StandardsBiotiumCatalog ##31028
LunaScript RT SuperMix KitNew England BiolabsCatalog #
E3010L
NEB Q5® Hot Start High-Fidelity 2X Master MixNew England BiolabsCatalog #M0494L
AccuClear® Ultra High Sensitivity dsDNA Quantitation Kit with DNA StandardsBiotiumCatalog ##31028
NEBNext Ultra II DNA Library Prep Kit for Illumina - 96 rxnsNew England BiolabsCatalog #E7645L
2x Kapa HiFi Hotstart Readymix Kapa BiosystemsCatalog #KK2602
cDNA generation
cDNA generation
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 10 µL extracted RNA On ice .
LunaScript RT SuperMix KitNew England BiolabsCatalog #
E3010L
Prepare RT mastermix in a dedicated UV treated pre-PCR area to minimise contamination risk.
| RT Master Mix | Vol / RXN (µL) | Vol/96 RXN (µL) inc. excess | |
| LunaScript Super Mix | 4 | 461 | |
| Nuclease-free water | 6 | 691 | |
| Total | 10 | 1152 |
Mix thoroughly by vortexing.
Use the SPT Labtech Dragonfly Discovery to dispense 10 µL of RT mastermix into the PCR plate containing 10 µL extracted RNA.
Seal and briefly centrifuge plate.
Place plate on a thermocycler and run the following program:
| 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).
cDNA amplification
cDNA amplification
Note
Primer pool sequences (v3) can be found here:
https://github.com/joshquick/artic-ncov2019/blob/master/primer_schemes/nCoV-2019/V3/nCoV-2019.tsv
Where an alt primer is available, the non alt version is omitted.
Expected result
Achieving more even genome coverage
A hypothetical 'ideal' multiplex primer pool would generate the same number of reads from each amplicon, so the fraction of reads due to each amplicon would be 1/n, where n is the number of primer pairs in the multiplex pool. In reality this is not achievable, and the fraction of reads observed for each amplicon varies widely.
The ratio [actual observed read fraction/‘ideal’ read fraction] can be calculated for each individual amplicon, as indicated by the differently-coloured dots on the box-and-whisker plots below. This tells us whether a particular amplicon is under-represented (ratio <1x) or over-represented (>1x).
By changing the weights of each primer pair within the primer pool ('rebalancing') the number of reads obtained for each amplicon can be modified, and the effect of the process is illustrated below. The plots show the distribution per amplicon prior to rebalancing primer pair concentrations (above) and after (below). More amplicons cluster around 1x after rebalancing and the distance between the maximum and minimum ratios is also markedly reduced.
Weight to apply per primer pair
A more detailed description of the process is provided in this document: 
Improving the evenness of SARS-CoV-2 genome coverage by titration of primer concentration (final).pdf
Improving the evenness of SARS-CoV-2 genome coverage by titration of primer concentration (final).pdf NEB Q5® Hot Start High-Fidelity 2X Master MixNew England BiolabsCatalog #M0494L
Prepare the following mastermixes:
| Weighted PCR Primer Pool 1 Master Mix | Vol/PCR RXN (µl) | Vol/96 plate (µl) inc. excess | |
| Q5 Hotstart 2X Master Mix | 12.5 | 1440 | |
| Primer Pool 1 (mean 102nM) | 3.6 | 415 | |
| Nuclease-free water | 2.9 | 334 | |
| Total | 19 | 2189 |
| Weighted PCR Primer Pool 2 Master Mix | Vol/PCR RXN (µl) | Vol/96 plate (µl) inc. excess | |
| Q5 Hotstart 2X Master Mix | 12.5 | 1440 | |
| Primer Pool 2 (mean 102nM) | 3.6 | 415 | |
| Nuclease-free water | 2.9 | 334 | |
| Total | 19 | 2189 |
Note
The equivolume primer pools used in the standard protocol are of 10 micromolar (µM) cumulative concentration, therefore each of the 98 primers in each pool is at 102 nanomolar (nM) in the pool and at 15 nanomolar (nM) in the final reaction. With the rebalanced primer pools, for equivalency we dilute them such that the average primer concentration is 102 nanomolar (nM) , and therefore the average concentration of each primer in the final reaction is also 15 nanomolar (nM) .
Mix thoroughly by vortexing.
Use the SPT Labtech Dragonfly Discovery to dispense 19 µL mastermix per well into 2x96 well plates.
Use the Agilent Bravo to add 6 µL of cDNA template to each primer pool reaction and mix.
Note
It is recommended to use filtered tips for this transfer to reduce risk of cross sample contamination via aerosolisation.
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.
Expected result
Critical step: We strongly recommend performing a gradient PCR to determine the optimal annealing temperature for your thermocycler. Subtle differences in thermocycler calibration can result in specific amplicons dropping out. Reducing our annealing temperature from 65°C to 63°C for identical cDNA input recovered amplicon #64 as shown in the image below.
| 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).
Amplified cDNA SPRI
Amplified cDNA SPRI
Allow AMPure XP beads to equilibrate to room temperature (~30 minutes). Ensure solution is homogenous prior to use, mixing gently by inversion.
Centrifuge amplified cDNA plates. 1000 x g, 00:01:00
Use either the Agilent Bravo or Beckman Coulter NX with a 96 well multichannel head to perform the following steps:
Combine the entire volumes of primer 1 and primer 2 PCR reactions per sample into one PCR plate.
Add 0.8X volume of SPRI beads per sample ( 40 µL SPRI : 50 µL amplified cDNA), mix well by pipetting.
Incubate for 00:05:00 at Room temperature
Transfer the plate to the magnet, allow 00:02:00 for the beads to settle.
Carefully remove and discard the supernatant without disturbing the bead pellet.
Wash the beads with 180 µL 75% freshly prepared ethanol for 00:00:30 , then remove ethanol and discard.
(First wash)
Wash the beads with 180 µL 75% freshly prepared ethanol for 00:00:30 , then remove ethanol and discard.
(Second wash)
Allow beads to dry 00:05:00
Remove plate from magnet, add 20 µL nuclease-free water and resuspend by mixing well.
Incubate for 00:03:00 at Room temperature
Transfer the plate to the magnet, allow 00:05:00 for the beads to settle.
Carefully transfer supernatant into a new plate, taking care not to disturb the bead pellet.
PAUSE POINT Purified amplified cDNA can be stored at -20°C for several weeks prior to library preparation.
Amplified cDNA quantification
Amplified cDNA quantification
Note
Purified amplified cDNA is quantified with a fluorescence based assay. We use the AccuClear Ultra High Sensitivity dsDNA Quantitation kit with 7 DNA standards (Biotium) according to manufacturer’s instructions.
To streamline the workflow, we do not normalise sample input for library preparation. Instead we confirm samples are in the range of 50ng-1ug per 20µl sample and take the entire volume into library preparation.
AccuClear® Ultra High Sensitivity dsDNA Quantitation Kit with DNA StandardsBiotiumCatalog ##31028
Pipette 20 µL of each DNA standard into wells A1 – G1 of a PCR plate. Add nuclease-free water to H1.
Dilute the AccuClear dye (100X) to working concentration by mixing 300 µL dye with 30 mL AccuClear buffer in a 50ml Falcon. Mix thoroughly by vortexing and transfer to a 384 well reservoir.
Use the SPT Labtech Mosquito LV to stamp 200nl of amplified cDNA and 1µl of known standards in triplicate into a 384 assay plate. Immediately proceed to the next step.
Use the Agilent Bravo 384ST to add 50 µL 1X AccuClear dye from the reservoir to the assay plate, mix thoroughly by pipetting.
Measure fluorescence values on a BMG FLUOstar Omega plate reader calibrated for use with AccuClear dye.
Confirm known standards are performing as expected.
Dilute any samples >125ng/µl with nuclease free water so they are in the range of 10 - 125ng/µl and repeat quantitation.
Note
Note: We use 5X the volume of standard vs sample in our assay setup, which should allow a quantitative range of 0.15ng/µl – 125ng/µl. This deviates from the standard kit SOP which has a stated range of 0.03ng/µl – 25ng/µl.
Ensure all samples (20µl total volume) are in the range of 2.5-50ng/µl prior to proceeding with library preparation.
Library preparation for Illumina sequencing
Library preparation for Illumina sequencing
Note
We use the NEB NEBNext® Ultra™ II DNA Library Prep Kit for Illumina, which we have automated on the Agilent Bravo platform with some modifications. 200ng is our standard input for library preparation, an acceptable range is 50ng – 1ug per sample. We use a custom adapter set, however any TruSeq adapters are suitable.
NEBNext Ultra II DNA Library Prep Kit for Illumina - 96 rxnsNew England BiolabsCatalog #E7645L
Prepare end prep mastermix On ice :
| NEBNext End Prep | Vol/PCR RXN (µl) | Vol/96 plate (µl) inc. excess | |
| NEBNext Ultra II End Prep Enzyme Mix | 1.2 | 144 | |
| NEBNext Ultra II End Prep Reaction Buffer | 2.8 | 336 | |
| Total | 4 | 480 |
Mix well by pipetting.
The Bravo will combine 4 µL of end prep mastermix with 20 µL amplified cDNA and mix by pipetting.
Seal and transfer the plate to a thermocycler and run the following program:
| Temperature | Time | |
| 20°C | 30 minutes | |
| 65°C | 30 minutes | |
| 4°C | ∞ |
Prepare adapter ligation mastermix On ice :
| Adapter Ligation | Vol/PCR RXN (µl) | Vol/96 plate (µl) inc. excess | |
| NEBNext Ultra II Ligation Master Mix | 12 | 1440 | |
| NEBNext Ligation Enhancer | 0.4 | 48 | |
| TruSeq adapter (10µM) | 1 | 120 | |
| Total | 13.4 | 1608 |
Mix well by pipetting.
The Bravo will add 13.4 µL adapter ligation mastermix to each sample and mix by pipetting.
The plate is incubated on deck at 20 °C for 00:15:00 , however this step may also be performed on a thermocycler.
Note
Note: We use alternative TruSeq compatible adapters, which do not require the USER enzyme incubation step. If using NEBNext adapters, follow the steps in the NEB protocol to add USER enzyme to the ligation reaction.
A 0.8X SPRI is performed post-ligation.
Ensure AMPure XP beads have been equilibrated to room temperature (~30 minutes) and the solution is homogenous prior to use.
The Bravo will perform a 0.8X SPRI clean-up and elute sample in 25 µl nuclease-free water as follows:
Add 0.8X volume of SPRI beads per sample, mix well by pipetting.
Incubate for 00:05:00 at Room temperature .
Transfer the plate to the magnet, allow 00:02:00 for the beads to settle.
Carefully remove and discard the supernatant without disturbing the bead pellet.
Wash the beads with 180 µL 75% freshly prepared ethanol for 00:00:30 , then remove ethanol and discard.
(First wash)
Wash the beads with 180 µL 75% freshly prepared ethanol for 00:00:30 , then remove ethanol and discard.
(Second wash)
Allow beads to dry for 00:05:00
Remove plate from magnet, add 25 µL nuclease free water and resuspend by mixing well.
Incubate for 00:03:00 atRoom temperature .
Transfer the plate to the magnet, allow 00:05:00 for the beads to settle.
Carefully transfer supernatant into a new plate, taking care not to disturb the bead pellet.
Half of this eluate (12.5µl) is used as input for library PCR.
Library PCR
Library PCR
Note
We use KAPA HiFi HotStart ReadyMix and unique dual indexed (UDI) tag plates for library PCR.
Note: this deviates from the standard NEB protocol which uses NEBNext Ultra II Q5 Master Mix and different cycling conditions.
2x Kapa HiFi Hotstart Readymix Kapa BiosystemsCatalog #KK2602
Prepare PCR mastermix On ice :
| KAPA PCR Mastermix | Vol/PCR RXN (µl) | Vol/96 plate (µl) inc. excess | |
| KAPA HiFi HotStart ReadyMix | 25 | 3000 | |
| Water | 12.5 | 1500 | |
| Total | 37.5 | 4500 |
Mix well by pipetting.
The Bravo will add 37.5 µL PCR mastermix and 12.5 µL sample into a lyophilised plate of UDIs and mix thoroughly by pipetting. The final concentration of each UDI in the PCR reaction is 2μM.
Seal and transfer the plate to a thermocycler and run the following program:
| Temperature | Time | |
| 95°C | 5 minutes | |
| 98°C | 30 seconds | |
| 65°C | 30 seconds | |
| 72°C | 2 minutes | |
| Repeat 4 times | ||
| 72°C | 5 minutes | |
| 4°C | ∞ |
Construct equivolume pool
Construct equivolume pool
In a post-PCR lab, combine 5µl of each sample per plate to form an equivolume pool of 96 samples.
Using a multichannel pipette, transfer 5 µL of each sample in the PCR plate into a low volume reservoir.
Transfer the contents of the reservoir into an Eppendorf tube and mix well.
Equivolume pool SPRI
Equivolume pool SPRI
Allow AMPure XP beads to equilibrate to room temperature (~30 minutes). Ensure solution is homogenous prior to use, mixing gently by inversion.
Add 0.8X volume of SPRI beads per pool tube, mix well by pipetting.
Incubate for 00:05:00 at Room temperature .
Transfer the tube to a magnet, allow 00:05:00 for the beads to form a pellet.
Carefully remove and discard the supernatant, taking care not to disturb the bead pellet.
Wash the beads with 1 mL 75% ethanol for 00:00:30 then carefully remove ethanol and discard.
(First wash)
Wash the beads with 1 mL 75% ethanol for 00:00:30 then carefully remove ethanol and discard.
(Second wash)
Allow beads to dry for 00:05:00 .
Remove tube from magnet and resuspend beads in 1 mL elution buffer, mix well by pipetting.
Incubate for 00:03:00 at Room temperature
Transfer tube to magnet, allow 00:05:00 for the beads to form a pellet.
Carefully transfer supernatant into a new tube, taking care not to disturb the bead pellet.
Equivolume pool quantification
Equivolume pool quantification
Note
Equivolume pools may be quantified either by qPCR or on an Agilent Bioanalyzer. Pools are then diluted to 1nM for sequencing.
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 or adapter-dimer present.
Sequencing
Sequencing
Note
We currently sequence samples on an Illumina NovaSeq SP flow cell, using the XP workflow.
Alternatively, samples may be sequenced on an Illumina MiSeq using either v2 (500 cycle) or v3 (600 cycle) reagent kits. We have plexed up to 96 samples per run, this could be increased further depending on coverage requirements. Loading concentration will need to be optimised for MiSeq.
MiSeq run parameters: Read length 212 paired end + 16bp.
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 Room 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 On ice
Bring flow cell to Room temperature (~10 minutes) prior to use.
18 µL of each 1 nanomolar (nM) pool is required per SP lane.
Denature pools by adding4 µL 0.2N NaOH per 18µl. Vortex briefly to mix.
Incubate at Room temperature for 00:08:00
Add 5 µL 400mM Tris-HCl, pH8.0 to each tube to neutralise the reaction. Vortex briefly to mix, then keep On ice .
Note
For the following steps, keep samples and mastermix On ice until ready for loading onto the flow cell.
Important! Use mastermix within 01:00:00 of preparation for optimal sequencing performance.
Prepare ExAmp mastermix on ice:
| ExAmp Master Mix | Volume per SP flow cell (µl) | |
| DPX1 | 126 | |
| DPX2 | 18 | |
| DPX3 | 66 | |
| Total | 210 |
Vortex00:00:30 to mix, then centrifuge briefly up to 280 x g
Add 63 µL 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 80 µL 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 00:30:00 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).