Nov 14, 2023

Amplicon multiplex PCR sequencing of Rift Valley fever virus (RVFV) on Illumina MiSeq  V.1

This protocol is a draft, published without a DOI.
Amplicon multiplex PCR sequencing of Rift Valley fever virus (RVFV) on Illumina MiSeq
  • 1International Livestock Research Institute (ILRI);
  • 2South Africa National Bioinformatics Institute (SANBI);
  • 3Kenya Medical Research Institute (KEMRI);
  • 4Rwanda Inspectorate, Competition and Consumer Protection Authority;
  • 5Division of Immunology, Department of Human Pathology, University of Cape Town;
  • 6South African National Bioinformatics Institute (SANBI), University of the Western Cape
John Juma
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Protocol Citation: John Juma, Samuel Oyola, Samson Konongoi, Isidore Nsengimana [Rwanda Inspectorate, Competition and Consumer Protection Authority], Reuben Mwangi [International Livestock Research Institute], James Akoko, Richard Nyamota, Collins Muli, Edward Kiritu, Paul Dobi, Shebbar Osiany, Amos Onwong'a, Rachael Gachogo, Rosemary Sang, Alan Christoffels, Kristina Roesel, Bernard Bett, Samuel Oyola 2023. Amplicon multiplex PCR sequencing of Rift Valley fever virus (RVFV) on Illumina MiSeq . protocols.io https://dx.doi.org/
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: December 10, 2022
Last Modified: November 14, 2023
Protocol Integer ID: 73818
Keywords: amplicon, Rift Valley fever, sequencing, protocol for rift valley fever virus, rift valley fever virus, amplicon multiplex pcr, sequencing protocol, virus, rvfv, pcr
Funders Acknowledgements:
DTRA
Grant ID: HDTRA11910031
Abstract
Amplicon sequencing protocol for Rift Valley fever virus (RVFV)
RNA extraction
Extract viral RNA from serum or cell-culture supernatants using QIAamp Viral RNA kit (QIAGEN, Hilden, Germany), according to the manufacturer’s instructions. Begin with a volume of 140 µL

RT-qPCR
Determine cycle threshold (Ct) values on RNA samples using probe-based reverse transcription quantitative real-time PCR against the highly conserved domain on the L-segment of the virus (using 5' Fam reporter dye and 3' BHQ1 quencher dye).

RVFV segment Primer name Sequence 5’-3’
L RVFL-2912fwdGG TGAAAATTCCTGAGACACATGG
L RVFL-2981revAC ACTTCCTTGCATCATCTGATG
L RVFL-probe-2950 CAATGTAAGGGGCCTGTGTGGACTTGTG
Table 1. Primers and probe set used for RT-qPCR assay (Bird et al., 2007).

Mix the following components in PCR strip-tubes/plate

AB
ComponentVolume (uL)
KiCqStart™ One-Step Probe RT-qPCR ReadyMix™7.5
Nuclease-free water4.75
RVFV Oligos (2912fwdGG, 2981revAC, probe-2950)0.75
RNA2.0
Total15


Note
Set up the reaction on ice.
Incubate the reaction on a Applied Biosystems machine as follows:

50 °C for 00:10:00
95 °C for 00:02:00
95 °C for 00:00:03 for 40 cycles
60 °C for 00:00:30

12m 33s
cDNA synthesis
30m
  1. Prepare RNA samples and include a negative control (nuclease-free water) per library. If previously frozen, mix by vortexing briefly and quick spin to collect the liquid. At all times, keep the samples on ice.
  2. Mix the following components in PCR strip-tubes/plate. Gently mix by pipetting and performing quick spin to collect the liquid.

AB
ComponentVolume
LunaScript RT Supermix (5X)2 uL
Template RNA8 uL
Total10 uL


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.


3. 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

13m
Primer pool preparation
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.


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.

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, V1 pools have 38 primers in pool 1 and 36 primers in pool 2, so the requirements is approx. 1.4 µL primer pool (100 micromolar (µM) ) per 25 µL reaction.


Note
Make up several 100 µL aliquots of 10 micromolar (µM) primer dilutions and freeze them in case of degradation and/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.


ABC
ComponentReaction 1Reaction 2
Q5 Hotstart Mastermix Buffer (5X)12.5 uL12.5 uL
V1 Primer Pool 11.425 uL0 uL
V1 Primer Pool 20 uL1.35 uL
Nuclease-free water6.575 uL6.65 uL
Mastermix Volume20.5 uL20.5 uL
(cDNA)4.5 uL4.5 uL
Total reaction Volume25 uL25 uL

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 4.5 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
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.

Set-up the following program on the thermal cycler:

Step Temperature Time Cycles

Heat activation 98 °C 00:00:30 1
Denaturation 95 °C 00:00:15 35
Annealing 63 °C 00:05:00 35
Hold 4 °C Indefinite 1

5m 45s
Amplicon clean-up
1h
Combine the two pools of amplicons:
Add 12.5 µL of each primer pool (Pool 1 and Pool 2, total of 25 µL ) in new PCR strip-tubes/plate.

Perform NEBNext Sample Purification Beads/AMPure XP bead cleanup as follows:
Add 20 µL (0.8X) of AMPure XP beads (thoroughly vortexed and at Room temperature ) to the combined amplicons plate. Cover the plate with seal, gently mix on a plate mixer and pulse spin to bring down the components at the bottom of the tube. Incubate at Room temperature for 00:05:00 (5 minutes).

5m
Place the tube/plate on a magnetic stand for 00:05:00 or until the beads have pelleted and the supernatant is completely clear.

5m
Remove and discard the liquid from each well with a multichannel pippette, being careful not to touch the bead pellet.


Note
Caution: do not discard the beads

Add 200 µL of freshly prepared, Room temperature 80% ethanol to each well/tube, incubate for 00:00:30 at Room temperature and then carefully remove and discard the supernatant.


Note
Be careful not to disturb the beads that contain DNA targets.

30s
Repeat ethanol wash (step 6.3 and 6.4).
Be sure to remove all visible liquid after the second wash. If necessary, briefly spin the tube/plate, place back on the magnet and remove traces of ethanol with a p10 pipette tip.
Air dry the beads for up to 5 minutes while the tube/plate is on the magnetic stand with the lid open.


Note
Caution: Do not over-dry the beads. This may result in lower recovery of DNA. Elute the samples when the beads are still dark brown and glossy looking, but when all visible liquid has evaporated. When the beads turn lighter brown and start to crack, they are too dry.

Remove the tube/plate from the magnetic stand. Elute the DNA target from the beads by adding 28 µL 0.1X TE or Elution Buffer (EB).
Mix well by pipetting up and down 10 times, or on a vortex mixer. Incubate for at least 00:02:00 (2 minutes) at room temperature. If necessary, quickly spin the sample to collect the liquid from the sides of the tube or plate wells before placing back on the magnetic stand.

2m
Place the tube/plate on the magnetic stand. After 00:05:00 (5 minutes) (or when the solution is clear).

5m
Transfer 25 µL to a new PCR tube, ensuring no beads are transferred.

Gel electrophoresis or Tapestation
20m
Use remaining volumes from Pool 1 and Pool 2 to confirm amplification (step 5.3).
Make 1% agarose gels with enough wells for all samples.
Load 2 µL of the 100 bp ladder into gel on either side of each row of wells.

Dispense 2 µL of 6X loading dye into each sample with a multichannel pipette, mix and load 2 µL 2of this mix into the gel.

Run at 240V for 00:20:00 . Visualize PCR products, confirm bands of approximately 400bp size.

20m
Run pooled cDNA amplicons on a TapeStation® without cleanup. To run on a TapeStation, dilute an aliquot of the pooled amplicons 10-fold with 0.1X TE Buffer and run 2 µL on a DNA High Sensitivity ScreenTape.

Amplicon quantification
Quantify amplicons using Qubit dsDNA High Sensitivity kit and plate reader according to directions.
Library preparation
1h 30m
Prepare sequencing libraries with NEBNext Ultra II RNA Library Prep kit at half volume, as follows.
End-Prep

Add the following components to a sterile nuclease-free tube:


AB
ComponentVolume
NEBNext Ultra II End Prep Enzyme Mix1.5 uL
NEBNext Ultra II Reaction Buffer3.5 uL
Targeted cDNA amplicon25 uL
Total volume30 uL

Set a 100 µL or 200 µL pipette to 25 µL and then pipette the entire volume up and down at least 10 times to mix thoroughly. Perform a quick spin to collect all liquid from the sides of the tube.

In a thermal cycler with lid heated to 75 °C , run the following program:

Temperature Time
20 °C 00:30:00
65 °C 00:30:00
4 °C Indefinite

1h
Adaptor-ligation


Add the following components directly to the End Prep Reaction Mixture


AB
ComponentVolume
End Prep Reaction Mixture (step 9.1)30 uL
NEBNext Adaptor for Illumina1.25 uL
NEBNext Ultra II Ligation Master Mix15 uL
Total volume46.25



Note
  1. Mix the NEBNext Ultra II Ligation Master Mix by pipetting up and down several times prior to adding to the reaction
  2. The NEBNext adaptor is provided in NEBNext Oligo kits. NEB has several oligo options which are supplied separately from the library prep kit. Please see www.neb.com/oligos for additional information

Do not premix adaptor with the Ligation Master Mix.


Set a 100 µL or 200 µL 2pipette to 40 µL and then pipette the entire volume up and down at least 10 times to mix thoroughly. Perform a quick spin to collect all liquid from the sides of the tube.


Note
Caution: The NEBNext Ultra II Ligation Master Mix is very viscous. Care should be taken to ensure adequate mixing of the ligation reaction, as incomplete mixing will result in reduced ligation efficiency. The presence of a small amount of bubbles will not interfere with performance)

Incubate at 20 °C for 00:15:00 (15 minutes) in a thermal cycler with the heated lid off.

15m
Add 1.5 µL of USER® Enzyme to the ligation mixture from Step 9.4.


Note
Steps 9.5. and 9.6. are only required for use with NEBNext Adaptors. USER enzyme can be found in the NEBNext Multiplex Oligos (www.neb.com/oligos).

Mix well and incubate at 37 °C for 00:15:00 (15 minutes) with the heated lid set to ≥ 47 °C .


Note
Samples can be stored overnight at –20°C.
Note: Only a portion of the ligation reaction (7.5 µl) will move forward to PCR enrichment.

15m
PCR Enrichment of Adaptor-ligated DNA
6m 55s
Follow Section 10.1. if you are using the following oligos: Use option A for any NEBNext Oligo kit where index primers are supplied in tubes. These kits have the forward and reverse primers supplied in separate tubes. Primers are supplied at 10 micromolar (µM) .

Follow Section 10.2. if you are using the following oligos: Use Option B for any NEBNext Oligo kit where index primers are supplied in a 96-well plate format. These kits have the forward and reverse (i7 and i5) primers combined. Primers are supplied at 10 micromolar (µM) .



Add the following components to a sterile strip tube:

Separate Forward and Reverse Primers


AB
ComponentVolume
Adaptor Ligated DNA Fragments (step 9.4 or 9.6)7.5 uL
NEBNext Library PCR Master Mix12.5 uL
Universal PCR Primer/i5 Primer2.5 uL
Index (X) /i7 Primer2.5 uL
Total volume25 uL


Add the following components to a sterile strip tube:

Premixed Forward and Reverse Primers


AB
ComponentVolume
Adaptor Ligated DNA Fragments (step 9.4 or 9.6)7.5 uL
Adaptor Ligated DNA Fragments (step 9.4 or 9.6)12.5 uL
Index Primer Mix5 uL
Total volume25 uL


Set a 100 µL pipette to 20 µL and then pipette the entire volume up and down at least 10 times to mix thoroughly. Perform a quick spin to collect all liquid from the sides of the tube.

Run the PCR program to amplify the libraries:

Step Temperature Time Cycles
Initial Denaturation 98 °C 00:00:30 1
Denaturation 98 °C 00:00:10 7
Annealing 65 °C 00:01:15 7
Extension 65 °C 00:05:00 1
Hold 4 °C Indefinite



6m 55s
Library Clean-up
Clean Up Libraries
Repeat the same clean up process as step 6 using 20 µL of AMPure beads or NEBNext Sample Purification Beads and 28 µL of Elution Buffer (EB)/ 0.1X TE.

Library quantification and normalization

Analyze 2 µL library using a Qubit dsDNA HS Assay kit

Calculate the molarity value using the following formula. Use the band size from gel electrophoresis or Tapestation readings (step 7).

Library concentration (0 µg/µL ) / (660 g/mol * average library size (bp)) * 10^6



Normalize each library by dilution with nuclease free water.
Pool equal volume (e.g. 5 µL ) from each of the normalized libraries into a single 1.5 mL Eppendorf tube.

Sequencing
5m
Denature and load pooled libraries as follows:
Denature the pooled libraries by mixing 5 µL of pooled libraries and 5 µL of 0.2N NaOH solution.

Incubate for 00:05:00 (5 minutes)

5m
Add 990 µL of HT1 buffer and mix well with denatured pooled library by pipetting up and down 10 times with P1000.

Load 600 µL of the denatured, diluted pooled library into the loading position of the Illumina reagent cartridge (V2, 300 cycle kit). Load reagent cartridge, flow cell, and PR2 buffer into Miseq instrument, confirm the metrics and start the run.