Jun 07, 2024
Version 4
ARTIC SARS-CoV-2 sequencing protocol v4 (LSK114) V.4
Version 1 is forked from Ebola virus sequencing protocol
- 1University of Birmingham
- ARTIC
Protocol Citation: Josh Quick, Lauren Lansdowne 2024. ARTIC SARS-CoV-2 sequencing protocol v4 (LSK114). protocols.io https://dx.doi.org/10.17504/protocols.io.bp2l6n26rgqe/v4Version created by Josh Quick
Manuscript citation:
Improvements to the ARTIC multiplex PCR method for SARS-CoV-2 genome sequencing using nanopore. John R Tyson, et. al. bioRxiv 2020.09.04.283077; doi:https://doi.org/10.1101/2020.09.04.283077
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: April 09, 2024
Last Modified: June 07, 2024
Protocol Integer ID: 97982
Keywords: sequencing protocol v4, ultra ii ligation module, sequencing protocol, protocol for sar, lunascript rt supermix, ta ligase master mix, artic sar, lsk114, protocol v4, superscript iv, new england biolab
Funders Acknowledgements:
Wellcome Trust
Grant ID: 206298/B/17/Z
Abstract
Amplicon sequencing protocol for SARS-CoV-2 v4 (LSK114)
We thank the ARTIC network, Oxford Nanopore Technologies, New England Biolabs, BCCDC, COG-UK, CanCOGen and protocols.io commenters for their assistance developing this protocol.
Changes in this version:
-Up to 95 samples per run with EXP-NBD196 native barcode kit
-Substitution of SuperScript IV for LunaScript RT SuperMix and reaction volume reduced to 10 uL.
-Substitution of Ultra II Ligation Module for Blunt/TA Ligase Master Mix and reaction volume reduced to 10 µL.
-Native barcode ligation reaction volume reduced to 10 uL.
-SFB wash volume reduced.
Materials
| Component | Supplier | Part number | |
| ARTIC nCoV-2019 V3 panel (100uM) | IDT | See links below | |
| LunaScript RT SuperMix Kit | NEB | E3010 | |
| Q5 Hot Start HF Polymerase or | NEB | M0493 | |
| Q5 Hot Start High-Fidelity 2X Master Mix | NEB | M0494 | |
| dNTP Solution Mix (10 mM ea.) | NEB | N0447 | |
| Nuclease-free water (100 mL) | NEB | B1500 | |
| NEBNext Ultra II End Repair/dA-tailing module | NEB | E7546 | |
| Blunt/TA Ligase Master Mix | NEB | M0367 | |
| Native Barcoding Expansion Kit 1-12 and/or | ONT | EXP-NBD104 | |
| Native Barcoding Expansion Kit 13-24 or | ONT | EXP-NBD114 | |
| Native Barcoding Expansion Kit 96 | ONT | EXP-NBD196 | |
| AMPure XP beads | Beckman | A63881 | |
| NEBNext Quick Ligation Module | NEB | E6056S | |
| Sequencing Auxiliary Vials | ONT | EXP-AUX001 | |
| Short Fragment Buffer Expansion Kit | ONT | EXP-SFB001 | |
| Qubit dsDNA HS Assay Kit | Thermo | Q32854 | |
| Flow Cell Priming Kit | ONT | EXP-FLP002 | |
| Flow Cell Wash Kit (optional) | ONT | EXP-WSH003 | |
| R9.4.1 flow cells | ONT | FLO-MIN106 |
Before start
Prepare between 11 and 95 RNA samples plus 1 negative control using this protocol.
cDNA preparation
30m
Prepare between 11 and 95 RNA samples plus 1 negative control of nuclease-free water per library. If previously frozen, mix by briefly vortexing and pulse spin to collect liquid. Keep samples on ice at all times.
Note
At least 11 samples are required to have sufficient material to load on the sequencer at the end. If you process >23 you will need the EXP-NBD196 expansion kit.
N.B. If you are processing <11 samples, the quantities of DNA used downstream can be increased to compensate (see Section 11, Native Barcoding).
Note
A positive control can also be included which may be a synthetic RNA constructs or high-titre clinical sample which can be diluted. This can help monitor run performance.
Mix the following components in a PCR strip-tubes/plate. Gently mix by pipetting and pulse spin the tube to collect liquid.
| Component | Volume | |
| LunaScript RT SuperMix (5X) | 2 µL | |
| Template RNA | 8 µL | |
| Total | 10 µL |
Note
Viral RNA input from a clinical sample should be between Ct 18-35. If Ct is between 12-15, then dilute the sample 100-fold in water, if between 15-18 then dilute 10-fold in water. This will reduce the likelihood of PCR-inhibition.
Note
To prevent pre-PCR contamination the mastermix should be added to the PCR strip-tubes/plate in the mastermix cabinet which should should be cleaned with decontamination wipes and UV sterilised before and after use.
RNA samples should be added in the extraction/sample addition cabinet which should should be cleaned with decontamination wipes and UV sterilised before and after use.
Incubate the reaction as follows:
25 °C for 00:02:00
55 °C for 00:10:00
95 °C for 00:01:00
Hold at 4 °C
Primer pool preparation (optional)
2h
If making up primer pools from individual oligos fully resuspend lyophilised oligos in 1xTE to a concentration of 100 micromolar (µM) , vortex thoroughly and spin down.
Note
If using IDT ARTIC nCoV-2019 V5.3.2 Panel (100 micromolar (µM) ) skip to step 6.
Sort all odd regions primers into one or more tube racks. Add 5 µL of each odd region primer to a 1.5 mL Eppendorf tube labelled "Pool 1 (100 micromolar (µM) )". Repeat the process for all even region primers for Pool 2. These are your 100 micromolar (µM) stocks of each primer pool.
Note
Primers should be diluted and pooled in the mastermix cabinet which should be cleaned with decontamination wipes and UV sterilised before and after use.
Note
For more information see Figure 2 in;
Quick, J. et al. Multiplex PCR method for MinION and Illumina sequencing of Zika and other virus genomes directly from clinical samples. Nat Protoc 12, 1261–1276 (2017). https://doi.org/10.1038/nprot.2017.066
Dilute 100 micromolar (µM) pools 1:10 in molecular grade water, to generate 10 micromolar (µM) primer stocks.
Note
Primers are used at a final concentration of 15 nanomolar (nM) per primer. In this case V3 pools have 110 primers in pool 1 and 108 primers in pool 2. so the requirement is ~4 µL primer pool (10 micromolar (µM) ) per 25 µL reaction.
Note
Make up multiple 100 µL aliquots of 10 micromolar (µM) primer dilutions and freeze them in case of degradation or contamination.
Multiplex PCR
4h
Set up the two PCR reactions per sample as follows in strip-tubes or plates. Gently mix by pipetting and pulse spin the tube to collect liquid at the bottom of the tube.
| Component | Reaction 1 | Reaction 2 | |
| 5X Q5 Reaction Buffer | 5 µL | 5 µL | |
| 10 mM dNTPs | 0.5 µL | 0.5 µL | |
| Q5 Hot Start DNA Polymerase | 0.25 µL | 0.25 µL | |
| V3 Pool 1 (10µM) | 4 µL | 0 µL | |
| V3 Pool 2 (10µM) | 0 µL | 4 µL | |
| Nuclease-free water | 12.75 µL | 12.75 µL | |
| Total | 22.5 µL | 22.5 µL |
For M0493
or
| Component | Reaction 1 | Reaction 2 | |
| Q5 Hot Start High-Fidelity 2X Master Mix | 12.5 µL | 12.5 µL | |
| V3 Pool 1 (10µM) | 4 µL | 0 µL | |
| V3 Pool 2 (10µM) | 0 µL | 4 µL | |
| Nuclease-free water | 6 µL | 6 µL | |
| Total | 22.5 µL | 22.5 µL |
For M0494
Note
Q5 Hot Start High-Fidelity 2X Master Mix can also be used instead of the component kit. Half-scale PCR reactions can also be used to save costs as you will only require 2.5 µL for downstream steps.
Note
To prevent pre-PCR contamination the mastermix for each pool should be made up in the mastermix cabinet which should should be cleaned with decontamination wipes and UV sterilised before and after use and aliquoted into PCR strip-tubes/plate
Add 2.5 µL cDNA to each of the PCR reactions, gently mix by pipetting and pulse spin the tube to collect liquid at the bottom of the tube.
Note
Up to 5 µL cDNA can be added to each PCR reaction (in place of nuclease-free water) to improve amplification of low titre samples. Using 5 µL cDNA will require a 20 µL cDNA reaction and may be more likely to cause inhibition so use cautiously.
Note
cDNA should be added in the extraction and sample addition cabinet which should should be cleaned with decontamination wipes and UV sterilised before and after use.
0 µL Set-up the following program on the thermal cycler:
Step Temperature Time Cycles
Heat Activation 98 °C 00:00:30 1
Denaturation 98 °C 00:00:15 25-35
Annealing 65 °C 00:05:00 25-35
Hold 4 °C Indefinite 1
Note
Cycle number should be 25 for Ct 18-21 up to a maximum of 35 cycles for Ct 35.
Note
Thermocycler calibration can vary instrument to instrument. If you see amplicon 64 dropout then decrease the annealing/extension temperature to 63 °C . Denaturation temperature of 95 °C can also be used and may slightly increase PCR yields.
Optional step: If you wish you can check the DNA concentration of your reactions before proceeding, to determine successful amplification (or failure in the case of the negative control). If you are processing many samples, to save time, you can check the concentration of a few samples to check amplification success. In our experience, reactions are typically around 80-100ng/μL. If your samples are <50ng/μL you can double the amount used in the next step. We recommend quantifying DNA with a fluorometer, such as a Qubit or Quantus.
PCR dilution
10m
Label strip-tubes/plate and combine the following volumes of each PCR reaction for 10 µL each sample:
| Component | Volume | |
| Pool 1 PCR reaction | 2.5 µL | |
| Pool 2 PCR reaction | 2.5 µL | |
| Nuclease-free water | 45 µL | |
| Total | 50 µL |
Note
The PCR post-clean up concentration is typically around 100 Mass Percent . This means we can pool them without quantification/normalisation to make a significant time saving. If you require very even barcode representation perform clean-up and normalise to 10 Mass Percent then continue.
Note
Amplicons should be added in the post-PCR cabinet which should should be cleaned with decontamination wipes and UV sterilised before and after use.
Native barocoding
2h
Barcode the amplicon pools using the one-pot native barcoding approach.
In a new PCR strip-tube/plate set up the following reaction for each sample:
| Component | Volume | |
| PCR dilution from previous step | 3.3 µL | |
| Ultra II End Prep Reaction Buffer | 1.2 µL | |
| Ultra II End Prep Enzyme Mix | 0.5 µL | |
| Nuclease-free water | 5 µL | |
| Total | 10 µL |
Note
Make a master mix of end-preparation reagents and nuclease-free water and aliquot into strip-tube/plate to improve reproducability.
Incubate at room temperature for 00:15:00
Incubate at 65 °C f for 00:15:00
Incubate on ice for 00:01:00
In a new PCR strip-tube/plate set up the following reaction for each sample:
| Component | Volume | |
| End-preparation reaction mixture | 0.75 µL | |
| NBXX barcode | 1.25 µL | |
| Blunt/TA Ligase Master Mix | 5 µL | |
| Nuclease-free water | 3 µL | |
| Total | 10 µL |
Note
Use one native barcode from the EXP-NBD104 (1-12), EXP-NBD114 (13-24) or EXP-NBD196 per sample. Use 12 or more barcodes per library or there will be insufficient total material to achieve good yields.
If processing <11 samples, increase quantities in the above reaction to allow for sufficient material
for sequencing. For example: if processing 6 samples, double the component volumes for a final reaction volume of 20µL for each sample.
Incubate at room temperature for 00:20:00
Incubate at 65 °C for 00:10:00
Incubate on ice for 00:01:00
Note
The 65°C incubation is to inactivate the DNA ligase to prevent barcode cross-ligation when reactions are pooled in the next step.
In a new 1.5 mL Eppendorf tube pool all one-pot barcoding reactions together.
Note
If processing <24 samples pool the total volume from all barcodes.
if processing 48 samples pool 5 µL from each native barcoding reaction.
If processing 96 samples pool 2.5 µL from each native barcoding reaction so as not to exceed a pool volume of 240 µL which would make the clean-up volume too large.
Add 0.4x volume of SPRI beads to the sample tube and mix gently by either flicking or pipetting. For example add 96 µL SPRI beads to 240 µL pooled one-pot barcoding reactions.
Note
0.4x volume of SPRI is sufficient to bind 400 bp amplicons in the presence of ligation buffer, do not use 1x as this will result in an excessive large bead pellet.
Mix by vortexing and pulse centrifuge to collect all liquid at the bottom of the tube. Incubate for 00:05:00 at room temperature.
Place on magnetic rack and incubate for 00:02:00 or until the beads have pelleted and the supernatant is completely clear. Carefully remove and discard the supernatant, being careful not to touch the bead pellet.
Add 250 µL SFB and resuspend beads completely by pipette mixing. Pulse centrifuge to collect all liquid at the bottom of the tube and place on the magnet. Remove supernatant and discard.
Note
SFB will remove excess adapter without damaging the adapter-protein complexes. Do not use 70% ethanol as in early clean-ups.
Repeat steps 11.9 to perform a second SFB wash. Pulse centrifuge and remove any residual SFB.
Note
You do not need to allow to air dry with SFB washes.
Add 200 µL of room-temperature 70 % volume ethanol to bathe the pellet. Carefully remove and discard ethanol, being careful not to touch the bead pellet.
Note
Only perform 1x 70% ethanol wash
Pulse centrifuge to collect all liquid at the bottom of the tube and carefully remove as much residual ethanol as possible using a P10 pipette.
With the tube lid open incubate for 00:01:00 or until the pellet loses it's shine (if the pellet dries completely it will crack and become difficult to resuspend).
Resuspend pellet in 30 µL 10 millimolar (mM) Tris pH 8.0, mix gently by either flicking or pipetting and incubate for 00:02:00 .
Place on magnet and transfer sample to a clean 1.5 mL Eppendorf tube ensuring no beads are transferred into this tube.
Quantify the barcoded amplicons using a fluorometer such as a Qubit or Quatus. Concentration will vary depending on number and Ct of samples and but you need about 30 ng total at this stage to achieve maximum run yield.
Set up the Native Adapter (NA) ligation, using short fragment buffer (SFB) for clean-up, following the current ONT protocol for the kit you have purchased.
Quantify the barcoded amplicons using a fluorometer such as a Qubit or Quatus. Concentration will vary depending on number and Ct of samples but 15 ng final library is usually required to acheive maximum run yield.
Note
Final library can be now be stored in 10 millimolar (mM) Tris 8 at 4 °C for up to a week if needed otherwise proceed directly to MinION sequencing.
MinION sequencing
1d
Prime and load the flowcell with 15 ng of sequencing library, following the current ONT protocol for the kit you have purchased.
Note
From experience we know 15 ng is optimum loading input for short amplicons. Speed drop during the run indicates excessive library was loaded. Low run yield <20M reads indicates insufficient library.
Start the sequencing run using MinKNOW.
Note
If using Live basecalling ensure to turn on double-ended barcoding in the basecalling settings.
Plug the MinION into the computer with the flowcell loaded, and wait for it to be detected.
Follow the sequential screens on MinKNOW to name the run, and select the appropriate kit(s). You should select ‘barcode both ends’, and we also recommend using the ‘trim barcodes’ function. If your computer has sufficient processing you can select ‘live basecalling’ (recommended), otherwise basecalling can be performed afterwards. We recommend using the High Accuracy basecalling mode.