Jan 22, 2020
Version 1
nCoV-2019 sequencing protocol V.1
Forked from Ebola virus sequencing protocol
- 1University of Birmingham
- ARTIC
- Coronavirus Method Development Community
External link: https://doi.org/10.1016/j.remle.2020.05.007
Protocol Citation: Josh Quick 2020. nCoV-2019 sequencing protocol. protocols.io https://dx.doi.org/10.17504/protocols.io.bbmuik6w
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: January 22, 2020
Last Modified: January 22, 2020
Protocol Integer ID: 32148
Abstract
ARTIC amplicon sequencing protocol for MinION for nCoV-2019
cDNA preparation
cDNA preparation
Mix the following components in an 0.2mL 8-strip tube;
Component Volume
50µM random hexamers 1 µL
10mM dNTPs mix (10mM each) 1 µL
Template RNA 11 µL
Total 13 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
A mastermix should be made up in the mastermix cabinet and aliquoted into PCR strip tubes. Tubes should be wiped down when entering and leaving the mastermix cabinet.
Gently mix by pipetting and pulse spin the tube to collect liquid at the bottom of the tube.
Incubate the reaction as follows:
65 °C for 00:05:00
Place on ice for 00:01:00
Add the following to the annealed template RNA:
Component Volume
SSIV Buffer 4 µL
100mM DTT 1 µL
RNaseOUT RNase Inhibitor 1 µL
SSIV Reverse Transcriptase 1 µL
Total 20 µL
Note
A mastermix should be made up in the mastermix cabinet and added to the denatured RNA in the extraction and sample addition cabinet. Tubes should be wiped down when entering and leaving the mastermix cabinet.
Gently mix by pipetting and pulse spin the tube to collect liquid at the bottom of the tube.
Incubate the reaction as follows:
42 °C 00:50:00
70 °C 00:10:00
Hold at 5 °C
Primer pool preparation
Primer pool preparation
If required resuspend lyophilised primers at a concentration of 100µM each
Note
nCov-2019/V1 primers for this protocol were designed using Primal Scheme and generate overlapping 400nt amplicons. Primer names and dilutions are listed in the table below.
Generate primer pool stocks by adding 5 µL of each primer pair to a 1.5 mL Eppendorf labelled either “Pool 1 (100µM)” or “Pool 2 (100µM)”. Total volume should be 490 µL for Pool 1 (100µM) and 490 µL for Pool 2 (100µM). These are your 100µM 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.
Dilute this primer pool 1:10 in molecular grade water, to generate 10µM primer stocks. It is recommend that multiple aliquots of each primer pool are made to in case of degradation or contamination.
Note
Primers need to be used at a final concentration of 0.015µM per primer. In this case both pools have 98 primers in so the requirement is 3.6µL primer pools (10uM) per 25µL reaction. For other schemes, adjust the volume added appropriately.
Multiplex PCR
Multiplex PCR
In the mastermix hood set up the multiplex PCR reactions as follows in 0.2mL 8-strip PCR tubes:
Component Pool 1 Pool 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
Primer Pool 1 or 2 (10µM) 3.6 µL 3.6 µL
Nuclease-free water 13.15 µL 13.15 µL
Total 22.5 µL 22.5 µL
Note
A PCR mastermix for each pool should be made up in the mastermix cabinet and aliquoted into PCR strip tubes. Tubes should be wiped down when entering and leaving the mastermix cabinet.
In the extraction and sample addition cabinet add 2.5 µL cDNA to each tube and mix well by pipetting.
Note
The extraction and sample addition cabinet should should be cleaned with decontamination wipes and UV sterilised before and after use.
Pulse centrifuge the tubes to collect the contents at the bottom of the tube.
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
PCR clean-up
PCR clean-up
Combine the entire contents of “Pool 1” and “Pool 2” PCR reactions for each biological sample into to a single 1.5 mL Eppendorf tube.
Clean-up the amplicons using the following protocol:
Protocol
NAME
Amplicon clean-up using SPRI beads
CREATED BY
Josh Quick
Note
Amplicon clean-up should be performed in the post-PCR cabinet which should should be cleaned with decontamination wipes and UV sterilised before and after use.
Vortex SPRI beads thoroughly to ensure they are well resuspended, the solution should be a homogenous brown colour.
Agencourt AMPure XPBeckman CoulterCatalog #A63880
Add an equal volume (1:1) of SPRI beads to the sample tube and mix gently by either flicking or pipetting. For example add 50 µL SPRI beads to a 50 µL reaction.
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 200 µL of room-temperature 70 % volume ethanol to the pellet.
Carefully remove and discard ethanol, being careful not to touch the bead pellet.
go to step #15.7 and repeat 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 Elution Buffer (EB), mix gently by either flicking or pipetting and incubate for 00:02:00 .
Elution Buffer (EB)QiagenCatalog #19086
Place on magnet and transfer sample to a clean 1.5mL Eppendorf tube ensuring no beads are transferred into this tube.
Quantify 1 µL product using the Quantus Fluorometer using the ONE dsDNA assay.
QuantiFluor(R) ONE dsDNA System, 100rxnPromegaCatalog #E4871
Equipment
Quantus
NAME
Fluorometer
TYPE
Promega
BRAND
E6150
SKU
LINK
Quantification and normalisation
Quantification and normalisation
Quantify the amplicon pools using the Quantus Fluorometer using the ONE dsDNA assay.
Protocol
NAME
DNA quantification using the Quantus fluorometer
CREATED BY
Josh Quick
Note
If the concentration is greater than 25 ng/µL dilute the sample by a factor of 10 by adding 270µL 10mM Tris and quantify again using the Quantus fluorometer.
Remove Lambda DNA 400 ng/µL standard from the freezer and leave on ice to thaw. Remove ONE dsDNA dye solution from the fridge and allow to come to room temperature.
QuantiFluor(R) ONE dsDNA System, 500rxnPromegaCatalog #E4870
Set up two 0.5 mL tubes for the calibration and label them 'Blank' and 'Standard'
Add 200 µL ONE dsDNA Dye solution to each tube.
Mix the Lambda DNA standard 400 ng/µL standard by pipetting then add 1 µL to one of the standard tube.
Mix each sample vigorously by vortexing for 00:00:05 and pulse centrifuge to collect the liquid.
Allow both tubes to incubate at room temperature for 00:02:00 before proceeding.
Selection 'Calibrate' then 'ONE DNA' then place the blank sample in the reader then select 'Read Blank'. Now place the standard in the reader and select 'Read Std'.
Set up the required number of 0.5 mL tubes for the number of DNA samples to be quantified.
Note
Use only thin-wall, clear, 0.5mL PCR tubes such as Axygen #PCR-05-C
Label the tubes on the lids, avoid marking the sides of the tube as this could interfere with the sample reading.
Add 199 µL ONE dsDNA dye solution to each tube.
Add 1 µL of each user sample to the appropriate tube.
Note
Use a P2 pipette for highest accuracy.
Mix each sample vigorously by vortexing for 00:00:05 and pulse centrifuge to collect the liquid.
Allow all tubes to incubate at room temperature for00:02:00 before proceeding.
On the Home screen of the Quantus Fluorometer, select `Protocol`, then select `ONE DNA` as the assay type.
Note
If you have already performed a calibration for the selected assay you can continue, there is no need to perform repeat calibrations when using ONE DNA pre diluted dye solution. If you want to use the previous calibration, skip to step 11. Otherwise, continue with step 9.
On the home screen navigate to 'Sample Volume' and set it to 1 µL then 'Units' and set it to ng/µL.
Load the first sample into the reader and close the lid. The sample concentration is automatically read when you close the lid.
Repeat step 16 until all samples have been read.
The value displayed on the screen is the dsDNA concentration in ng/µL, carefully record all results in a spreadsheet or laboratory notebook.
Label a 1.5 mL Eppendorf tube for each sample.
Note
This is a ‘one-pot ligation’ protocol for native barcoded ligation libraries. We have seen no reduction in performance compared to standard libraries, and is made faster by using the Ultra II® ligation module which is compatible with the Ultra II® end repair/dA-tailing module removing a clean-up step.
Normalise the input by diluting each sample to 1 ng/µL . Use 5 µL input for the One-pot native barcoding reaction to give a total of 5 ng per sample.
Note
Input to the one-pot native barcoding reaction will vary depending on the amplicon length but we have determined 5ng is the correct input for efficient barcoding of this amplicon length. Process at least 7 samples plus one negative control per native barcoded library in order to have sufficient material at the end.
Native barocoding
Native barocoding
Barcode the amplicon pools using native barcodes.
Protocol
NAME
One-pot native barcoding of amplicons
CREATED BY
Josh Quick
Set up the following reaction for each sample:
Component Volume
DNA amplicons 5 µL
Nuclease-free water 7.5 µL
Ultra II End Prep Reaction Buffer 1.75 µL
Ultra II End Prep Enzyme Mix 0.75 µL
Total 15 µL
Incubate at room temperature for 00:10:00
Incubate at 65 °C f for 00:05:00
Incubate on ice for 00:01:00
Add the following directly to the previous reactions:
Component Volume
NBXX barcode 2.5 µL
Ultra II Ligation Master Mix 17.5 µL
Ligation Enhancer 0.5 µL
Total 35.5 µL
Note
Use one native barcode from the EXP-NBD104 (1-12) or EXP-NBD114 (13-24) per sample. Use from 6 to 24 barcodes in a library, any fewer and there will be insufficient total material to achieve good yields.
Incubate at room temperature for 00:15:00
Incubate at 70 °C for 00:10:00
Incubate on ice for 00:01:00
Note
The 70°C incubation is to inactivate the DNA ligase to prevent barcode cross-ligation when reactions are pooled in the next step.
Pool all barcoded fragments together into a new 1.5 ml Eppendorf tube.
Protocol
NAME
Amplicon clean-up using SPRI beads
CREATED BY
Josh Quick
Quantify the barcoded amplicon pools using the Quantus Fluorometer using the ONE dsDNA assay.
Protocol
NAME
DNA quantification using the Quantus fluorometer
CREATED BY
Josh Quick
Set up the following AMII adapter ligation reaction:
Component Volume
Barcoded amplicon pools 30 µL
NEBNext Quick Ligation Reaction Buffer (5X) 10 µL
AMII adapter mix 5 µL
Quick T4 DNA Ligase 5 µL
Total 50 µL
Note
The input of barcoded amplicon pools will depend on the number of barcoded pools and should be between 40 ng (8 barcodes) and 160 ng (24 barcodes).
Incubate at room temperature for 00:15:00
Add 50 µL (1:1) of SPRI beads to the sample tube and mix gently by either flicking or pipetting.
Note
Vortex SPRI beads thoroughly before use to ensure they are well resuspended, the solution should be a homogenous brown colour.
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 200 µL SFB and resuspend beads completely by pipette mixing.
Note
SFB will remove excess adapter without damaging the adapter-protein complexes. Do not use 70% ethanol as in early clean-ups.
Pulse centrifuge to collect all liquid at the bottom of the tube.
Remove supernatant and discard.
Repeat steps 14-16 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 15 µL EB and resuspend beads by pipette mixing.
Incubate at room temperature for 00:02:00 .
Place on magnetic rack.
Transfer final library to a new 1.5mL Eppendorf tube.
Quantify the final library using the Quantus Fluorometer using the ONE dsDNA assay.
Protocol
NAME
DNA quantification using the Quantus fluorometer
CREATED BY
Josh Quick
Note
Final library can be now be stored in 10 mM Tris pH 8 at 4°C for up to a week if needed otherwise proceed directly to MinION sequencing.
Remove Lambda DNA 400 ng/µL standard from the freezer and leave on ice to thaw. Remove ONE dsDNA dye solution from the fridge and allow to come to room temperature.
QuantiFluor(R) ONE dsDNA System, 500rxnPromegaCatalog #E4870
Set up two 0.5 mL tubes for the calibration and label them 'Blank' and 'Standard'
Add 200 µL ONE dsDNA Dye solution to each tube.
Mix the Lambda DNA standard 400 ng/µL standard by pipetting then add 1 µL to one of the standard tube.
Mix each sample vigorously by vortexing for 00:00:05 and pulse centrifuge to collect the liquid.
Allow both tubes to incubate at room temperature for 00:02:00 before proceeding.
Selection 'Calibrate' then 'ONE DNA' then place the blank sample in the reader then select 'Read Blank'. Now place the standard in the reader and select 'Read Std'.
Set up the required number of 0.5 mL tubes for the number of DNA samples to be quantified.
Note
Use only thin-wall, clear, 0.5mL PCR tubes such as Axygen #PCR-05-C
Label the tubes on the lids, avoid marking the sides of the tube as this could interfere with the sample reading.
Add 199 µL ONE dsDNA dye solution to each tube.
Add 1 µL of each user sample to the appropriate tube.
Note
Use a P2 pipette for highest accuracy.
Mix each sample vigorously by vortexing for 00:00:05 and pulse centrifuge to collect the liquid.
Allow all tubes to incubate at room temperature for00:02:00 before proceeding.
On the Home screen of the Quantus Fluorometer, select `Protocol`, then select `ONE DNA` as the assay type.
Note
If you have already performed a calibration for the selected assay you can continue, there is no need to perform repeat calibrations when using ONE DNA pre diluted dye solution. If you want to use the previous calibration, skip to step 11. Otherwise, continue with step 9.
On the home screen navigate to 'Sample Volume' and set it to 1 µL then 'Units' and set it to ng/µL.
Load the first sample into the reader and close the lid. The sample concentration is automatically read when you close the lid.
Repeat step 16 until all samples have been read.
The value displayed on the screen is the dsDNA concentration in ng/µL, carefully record all results in a spreadsheet or laboratory notebook.
MinION sequencing
MinION sequencing
Prime the flowcell and load 20 ng sequencing library onto the flowcell.
Protocol
NAME
Priming and loading a MinION flowcell
CREATED BY
Josh Quick
Note
From experience we know 20 ng is optimum loading input for short amplicons.
Thaw the following reagents at room temperature before placing on ice:
Sequencing buffer (SQB)
Loading beads (LB)
Flush buffer (FLB)
Flush tether (FLT)
Add 30 µL FLT to the FLB tube and mix well by vortexing.
If required place a new MinION flowcell onto the MinION by flipping open the lip and pushing one end of the flowcell under the clip and pushing down gently.
Rotate the inlet port cover clockwise by 90° so that the priming port is visible.
Take a P1000 pipette and tip and set the volume to 800 µL . Place the tip in the inlet port and holding perpendicularly to the plane of the flowell remove any air from the inlet port by turning the volume dial anti-clockwise.
Note
Be careful not to remove so much volume that air is introduced onto the rectangular array via the outlet.
Load 800 µL of FLB (plus FLT) into the flow cell via the inlet port, dispense slowly and smoothly trying to avoid the introduction of any air bubbles.
Wait for00:05:00 .
Gently lift the SpotON cover to open the SpotON port.
Load another 200 µL of FLB (plus FLT) into the flow cell via the inlet port, this will initiate a siphon at the SpotON port to allow you to load the library dilution.
In a new tube prepare the library dilution for sequencing:
Component Volume
SQB 37.5 µL
LB 25.5 µL
Final library 12 µL
Total 75 µL
Note
Mix LB immediately before use as they settle quickly.
Dilute library in EB if required.
Mix the prepared library gently by pipetting up and down just prior to loading.
Add the 75 µL library dilution to the flow cell via the SpotON sample port in a dropwise fashion. Ensure each drop siphons into the port before adding the next.
Gently replace the SpotON sample port cover, making sure the bung enters the SpotON port, close the inlet port and close the MinION lid.
Start the sequencing run using MinKNOW.
Protocol
NAME
Starting a MinION sequencing run using MinKNOWCREATED BY
Josh Quick
If required plug the MinION into the computer and wait for the MinION and flowcell to ben detected.
Choose flow cell 'FLO-MIN106' from the drop-down menu.
Then select the flowcell so a tick appears.
Click the 'New Experiment' button in the bottom left of the screen.
On the New experiment popup screen, select the running parameters for your experiment from the individual tabs:
Experiment: Name the run in the experiment field, leave the sample field blank.
Kit: Selection: Select LSK109 as there is no option for native barcoding (NBD104).
Run Options: Set the run length to 6 hours (you can stop the run once sufficient data has been collected as determined using RAMPART).
Basecalling: Leave basecalling turned but select 'fast basecalling'.
Output: The number of files that MinKNOW will write to a single folder. By default this is set to 4000 but can be reduced to make RAMPART update more frequently.
Click 'Start run'.
Monitor the progress of the run using the MinKNOW interface.