Dec 09, 2021
Version 1
SARS-CoV-2 McGill Nextera Flex sequencing protocol_Lunascript_ARTIC.V3_5uLRT V.1
- 1McGill University
- McGill Genome Centre
Protocol Citation: Sarah J Reiling, Marie-Michelle Simon, Anne-Marie Roy, Shu-Huang Chen, Ioannis Ragoussis 2021. SARS-CoV-2 McGill Nextera Flex sequencing protocol_Lunascript_ARTIC.V3_5uLRT. protocols.io https://dx.doi.org/10.17504/protocols.io.by6xpzfn
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: October 18, 2021
Last Modified: December 09, 2021
Protocol Integer ID: 54199
Abstract
How the Nextera DNA Flex Assay Works
The Nextera DNA Flex library prep kit uses a bead-based transposome complex to tagment genomic DNA, which is a process that fragments DNA and then tags the DNA with adapter sequences in one step. After it is saturated with input DNA, the bead-based transposome complex fragments a set number of DNA molecules. This fragmentation provides flexibility to use a wide DNA input range to generate normalized libraries of consistent tight fragment size distribution. Following tagmentation, a limited-cycle PCR adds Nextera DNA Flex-specific index adapter sequences to the ends of a DNA fragment. This step enables capability across all Illumina sequencing platforms. A subsequent Sample Purification Beads (SPB) cleanup step then purifies libraries for use on an Illumina sequencer. The double-stranded DNA library is denatured before hybridization of the biotin probe oligonucleotide pool.
PCR Amplicons for Nextera Flex
When starting with PCR amplicons, the PCR amplicon must be > 150 bp. The standard clean up protocol depletes libraries < 500 bp. Therefore, Illumina recommends that amplicons < 500 bp undergo a 1.8 x sample purification bead volume ratio to supernatant during Clean Up Libraries on page 11. Shorter amplicons can otherwise be lost during the library cleanup step. Tagmentation cannot add an adapter directly to the distal end of a fragment, so a drop in sequencing coverage of ~50 bp from each distal end is expected. To ensure sufficient coverage of the amplicon target region, design primers to extend beyond the target region by 50 bp per end.
cDNA preparation
cDNA preparation
cDNA PREPARATION
Mix the following components in a 0.2 mL 8-strip tube or in a 96-well plate:
Component Volume
LunaScript RT SuperMix (5x) 4 µL
Nuclease-free water 5 µL
Template RNA 11 µL
Total 20 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:
25 °C for 00:02:00
55 °C for 00:20:00
95 °C for 00:01:00
Place on ice for 00:01:00 or store cDNA at -20C
24m
Primer pool preparation
Primer pool preparation
PRIMER POOL PREPARATION
If required resuspend lyophilised primers at a concentration of 100 µM each
ARTIC nCov-2019 only primers for this protocol were designed using Primal Scheme and generate overlapping 400 nt amplicons. Primer names and dilutions are listed in the table below. https://github.com/sarahreiling/artic-ncov2019/blob/master/primer_schemes/nCoV-2019/V3/nCoV-2019_V3only.scheme.bed
Generate primer pool stocks by adding 5 µL of each primer pair to a 1.5 mL Eppendorf labelled either “Pool 1 (100 µM)” or “Pool 2 (100 µM)”. Total volume should be 490 µL for Pool 1 (100 µM) and 490 µL for Pool 2 (100 µM). These are your 100 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.
Dilute this primer pool 1:10 in molecular grade water, to generate 10 µM 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 0.015 µM per primer. In this case both pools have 98 primers in so the requirement is 3.65 µL primer pools (10 uM) per 25 µL reaction. For other schemes, adjust the volume added appropriately.
Multiplex PCR
Multiplex PCR
MULTIPLEX PCR
In the extraction and sample addition cabinet add 5 µL RT product 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.
In the mastermix hood set up the multiplex PCR reactions as follows in 0.2mL 8-strip PCR tubes:
Component Pool 1 [10 uM primer] Pool 2 [10 uM]
Q5 Hot Start High-Fidelity 2X Master Mix 12.5 µL 12.5 µL
Primer Pool 1 or 2 3.7 µL 3.7 µL
Nuclease-free water 3.8 µL 3.8 µL
Total 20 µL 20 µL
Add 5 ul RT product as mentioned in step 10.
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.
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 36
Annealing 63 °C 00:05:00 36
Hold 4 °C Indefinite 1
Note
Cycle number should be 25 for Ct 18-21 up to a maximum of 36 cycles for Ct 35
PCR clean-up
PCR clean-up
PCR CLEANUP
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:
Add a volume ratio of 0.8x of SPRI beads to the sample tube and mix gently by either flicking or pipetting.
Incubate for 5 min at room temperature.
Pellet on magnet for 5 min. Remove supernatant.
Add 200 ul of 80% ethanol to the pellet and wash twice.
Elute in 30 ul elution buffer.
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.
Amplicon Quantification and normalisation
Amplicon Quantification and normalisation
AMPLICON QUANTIFICATION AND NORMALIZATION
Quantify the amplicon pools using a fluorimetric dsDNA assay.
We expect following concentrations:
Pool 1+2 combined:
100-150 ng/ul for Ct 14-24
30-80 ng/ul for Ct 25-29
10-30 ng/ul for Ct 30-36
Use the 30 ul with 150 ng DNA for Nextera Flex Tagmentation.
Protocol can be found here:
Start on page 7 with the Tagmentation.
Tagment Genomic DNA
Tagment Genomic DNA
Note
For automated protocol : at step 7, 22 ul tagmentation master mix is transfered.
Post Tagmentation Cleanup
Post Tagmentation Cleanup
Amplify Tagmented DNA
Amplify Tagmented DNA
Note
For automated protocol : at step 9, 44 ul tagmentation master mix is transfered.
Note
At step 14, beads can be resuspended in 62 uL RSB , followed by a 60 uL transfer of supernatant at step 18. This is used to decrease the concentration of the final pool in order to facilitate the QC step.
Pool Libraries
Pool Libraries
Check Library Quality
Check Library Quality
Dilute Libraries to the Starting Concentration
Dilute Libraries to the Starting Concentration