# Goal/Experiment:
Amplicon multiplex PCR sequencing of Rift Valley fever virus (RVFV) on Illumina MiSeq
## Abstract
Amplicon sequencing protocol for Rift Valley fever virus (RVFV).
## RNA Extraction
1. 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
2. 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 | TGAAAATTCCCTGAGACACATGG |
| L | RVFL-2981revAC | ACTTCCTTGCATCACTGATG |
| L | RVFL-probe-2950 | CAATGTAAGGGGCCTGTTGTGGCAGCTGTG |
Table 1. Primers and probe set used for RT-qPCR assay (Bird et al., 2007).
### Mix the following components in PCR strip-tubes/plate
| Component | Volume (µL) |
|-----------------------------------|-------------|
| KiCqStart™ One-Step Probe RT-qPCR ReadyMix™ | 7.5 |
| Nuclease-free water | 4.75 |
| RVFV Oligos (2912FwdGG, 2981revAC, probe-2950) | 0.75 |
| RNA | 2.0 |
| Total | 15 |
Note: Set up the reaction on ice. Incubate the reaction on an Applied Biosystems machine as follows:
- 50 °C for 10:00
- 95 °C for 2:00
- 95 °C for 0:03, 40 cycles
- 60 °C for 0:30
## cDNA Synthesis
3. 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. Mix the following components in PCR strip-tubes/plate. Gently mix by pipetting and performing quick spin to collect the liquid.
| Component | Volume (µL) |
|-----------------|-------------|
| LunaScript RT Supermix (5X) | 2 |
| Template RNA | 8 |
| Total | 10 |
Note: To prevent pre-PCR contamination the mastermix should be added to the PCR strip-tubes/plate in the mastermix cabinet, which should be cleaned with decontamination wipes and UV sterilized before and after use. RNA samples should be added in the extraction/sample addition cabinet which should be cleaned with decontamination wipes and UV sterilized before and after use.
3. Incubate the reaction as follows:
- 25 °C for 2:00
- 55 °C for 10:00
- 95 °C for 1:00
- Hold at 4 °C
## Primer Pool Preparation
4. If making up primer pools from individual oligos, fully resuspend lyophilized oligos in 1xTE to a concentration of 100 micromolar (µM), vortex thoroughly, and spin down.
### Preparing the primer pools:
4.1 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 labeled "Pool 1 (100 µM)". Repeat the process for all even region primers for Pool 2. 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 sterilized before and after use.
4.2 Dilute 100 µM pools 1:10 in molecular grade water, to generate 10 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 are approx. 1.4 µL primer pool (100 µM) per 25 µL reaction.
Note: Make up several 100 µL aliquots of 10 µM primer dilutions and freeze them in case of degradation and/or contamination.
## Multiplex PCR
5. 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 (µL) | Reaction 2 (µL) |
|-------------------------------|-------------------------|-------------------------|
| Q5 Hotstart Mastermix Buffer (5X) | 12.5 | 12.5 |
| V1 Primer Pool 1 | 1.425 | 0 |
| V1 Primer Pool 2 | 0 | 1.35 |
| Nuclease-free water | 6.575 | 6.65 |
| Mastermix Volume | 20.5 | 20.5 |
| (cDNA) | 4.5 | 4.5 |
| **Total reaction Volume** | **25** | **25** |
Note: To prevent pre-PCR contamination the mastermix for each pool should be made up in the mastermix cabinet, which should be cleaned with decontamination wipes and UV sterilized before and after use and aliquoted into PCR strip-tubes/plate.
### 5.2 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 be cleaned with decontamination wipes and UV sterilized before and after use.
### Set up the following program on the thermal cycler:
| Step | Temperature | Time | Cycles |
|----------------|-------------|------------------|---------|
| Heat activation| 98 °C | 0:30 | 1 |
| Denaturation | 95 °C | 0:15 | 35 |
| Annealing | 63 °C | 5:00 | 35 |
| Hold | 4 °C | Indefinite | 1 |
## Amplicon Clean-up
6. 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:
### 6.1 Add 20 µL (0.8X) of AMPure XP beads (thoroughly vortexed and at Room temperature). 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 5 minutes.
### 6.2 Place the tube/plate on a magnetic stand for 5 minutes or until the beads have pelleted and the supernatant is completely clear.
### 6.3 Remove and discard the liquid from each well with a multichannel pipette, being careful not to touch the bead pellet.
### Note: Caution: do not discard the beads.
### 6.4 Add 200 µL of freshly prepared, Room temperature 80% ethanol to each well/tube, incubate for 30 seconds at Room temperature and then carefully remove and discard the supernatant.
### Note: Be careful not to disturb the beads that contain DNA targets.
### 6.5 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.
### 6.6 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.
### 6.7 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).
### 6.8 Mix well by pipetting up and down 10 times, or on a vortex mixer. Incubate for at least 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.
### 6.9 Place the tube/plate on the magnetic stand. After 5 minutes (or when the solution is clear).
### 6.10 Transfer 25 µL to a new PCR tube, ensuring no beads are transferred.
## Gel Electrophoresis or Tapestation
7. Use remaining volumes from Pool 1 and Pool 2 to confirm amplification (step 5.3).
### 7.1 Make 1% agarose gels with enough wells for all samples.
### 7.2 Load 2 µL of the 100 bp ladder into gel on either side of each row of wells.
### 7.3 Dispense 2 µL of 6X loading dye into each sample with a multichannel pipette, mix and load 2 µL of this mix into the gel.
### 7.4 Run at 240V for 20 minutes. Visualize PCR products, confirm bands of approximately 400bp.
### 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
8. Quantify amplicons using Qubit dsDNA High Sensitivity kit and plate reader according to directions.
## Library Preparation
9. Prepare sequencing libraries with NEBNext Ultra II RNA Library Prep kit at half volume, as follows.
### 9.1 End-Prep
Add the following components to a sterile nuclease-free tube:
| Component | Volume (µL) |
|-----------------------------------|-------------|
| NEBNext Ultra II End Prep Enzyme Mix | 1.5 |
| NEBNext Ultra II Reaction Buffer | 3.5 |
| Targeted cDNA amplicon | 25 |
| **Total volume** | **30** |
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 | 30:00 |
| 65 °C | |
| 4 °C | Indefinite|
### 9.2 Adaptor-ligation
Add the following components directly to the End Prep Reaction Mixture:
| Component | Volume (µL) |
|------------------------------|-------------|
| End Prep Reaction Mixture (step 9.1) | 30 |
| NEBNext Adaptor for Illumina | 1.25 |
| NEBNext Ultra II Ligation Master Mix | 15 |
| **Total volume** | **46.25** |
Note: Mix the NEBNext Ultra II Ligation Master Mix by pipetting up and down several times prior to adding to the reaction. 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.
### 9.3 Set a 100 µL or 200 µL pipette 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.
### 9.4 Incubate at 20 °C for 15 minutes in a thermal cycler with the heated lid off.
### 9.5 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).
### 9.6 Mix well and incubate at 37 °C for 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.
## PCR Enrichment of Adaptor-ligated DNA
10. 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).
### 10.1 Add the following components to a sterile strip tube:
Separate Forward and Reverse Primers
| Component | Volume (µL) |
|---------------------------------------|-------------|
| Adaptor Ligated DNA Fragments (step 9.4 or 9.6) | 7.5 |
| NEBNext Library PCR Master Mix | 12.5 |
| Universal PCR Primer/i5 Primer | 2.5 |
| Index (X) /i7 Primer | 2.5 |
| **Total volume** | **25** |
### 10.2 Add the following components to a sterile strip tube:
Premixed Forward and Reverse Primers
| Component | Volume (µL) |
|---------------------------------------|-------------|
| Adaptor Ligated DNA Fragments (step 9.4 or 9.6) | 7.5 |
| Adaptor Ligated DNA Fragments (step 9.4 or 9.6) | 12.5 |
| Index Primer Mix | 5 |
| **Total volume** | **25** |
### 10.3 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.
### 10.4 Run the PCR program to amplify the libraries:
| Step | Temperature | Time | Cycles |
|-----------------------|-------------|----------------|--------|
| Initial Denaturation | 98 °C | 0:30 | 1 |
| Denaturation | 98 °C | 0:10 | 7 |
| Annealing | 65 °C | 1:15 | 7 |
| Extension | 65 °C | 5:00 | 1 |
| Hold | 4 °C | Indefinite | 1 |
## Library Clean-up
11. Clean Up Libraries:
Repeat the same clean up process as step 6 using 20 µl of AMPure XP beads or NEBNext Sample Purification Beads and 28 µL of Elution Buffer (EB)/ 0.1X TE.
## Library Quantification and Normalization
12.
### 12.1 Analyze 2 µL library using a Qubit dsDNA HS Assay kit.
### 12.2 Calculate the molarity value using the following formula. Use the band size from gel electrophoresis or Tapestation readings (step 7).
Library concentration (µg/µL) / (660 g/mol * average library size (bp)) * 10^6.
### 12.3 Normalize each library by dilution with nuclease free water.
### 12.4 Pool equal volume (e.g., 5 µL) from each of the normalized libraries into a single 1.5 mL Eppendorf tube.
## Sequencing
13. Denature and load pooled libraries as follows:
### 13.1 Denature the pooled libraries by mixing 5 µL of pooled libraries and 5 µL of 0.2N NaOH solution.
### 13.2 Incubate for 5 minutes.
### 13.3 Add 990 µL of HT1 buffer and mix well with denatured pooled library by pipetting up and down 10 times with P1000.
### 13.4 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.
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