TITLE: Amplicon Sequencing for Genotyping S. Typhi
AUTHORS: Jaspreet Mahindroo, Anton Spadar, Catherine Troman, Zoe Dyson, Kathryn Holt, Nick Grassly

[DESCRIPTION]
The following protocol is for amplifying and sequencing amplicons targeting Salmonella Typhi. It is primarily for use with samples that are already suspected to be positive for S. Typhi and has been designed for use with DNA extracted from environmental surveillance samples. 

The resulting sequences can be analysed and used for genotyping (given that the genotype is targeted by the primer panel) and determining antimicrobial resistance. The genotypes targeted in the panel in this protocol are listed but primers can be designed for other S. typhi genotypes of interest. 

The library preparation steps of this protocol are adapted from the Oxford nanopore protocol "Ligation Sequencing Amplicons - Native Barcoding Kit 24 V14 (SQK-NBD114.24)" which is available on the Nanopore community.

[STEPS]
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12.1. From the ONT kit (SQK-NBD114.24 or .96) thaw AMPure XP beads (AXP), mix by vortexing, then keep at room temperature.

Thaw the NEBnext Ultra II End Repair reagents on ice, flick or invert the tubes to mix, then spin down.
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12.2. To the 12.5μl of prepared 200fmol amplicon DNA, add the following:

 
You may make up a master mix of these reagents for the number of samples (+1 for pipetting error) and aliquot 2.5μl for each sample.
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12.3. Gently mix by pipetting or flicking the tube then spin down.
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13. Native barcode ligation
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12. End-preparation
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12.4. Incubate in a thermal cycler at 20ºC for 5 minutes then 65ºC for 5 minutes
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13.1. From the ONT kit:
Thaw the EDTA at room temperature, mix by vortexing, spin down, then place on ice.
Thaw the required Native barcodes at room temperature (a different barcode for each sample), mix by flicking of pipetting, spin down, then place on ice.

Thaw the Blunt/TA ligase master mix at room temperature, mix by inverting and flicking well, spin down, then place on ice.
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13.2. Select a different barcode for each sample and note this down in your sample spreadsheet (the same spreadsheet used to calculate sample volumes).

Add the following reagents to a clean 0.2ml tube:

 

Mix by gently pipetting or flicking then spin down.
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13.3. Incubate at room temperature for 20 minutes.
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13.4. To each sample add the following volume of EDTA depending on the colour of its cap:

 

Pipette to mix on addition of the EDTA then spin each tube down.
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13.5. Pool all samples into a single 1.5ml tube and note down the final volume.

You can estimate this by multiplying the volume of each sample (22μl if clear cap EDTA, 24μl if blue cap EDTA) by the total number of samples.
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13.6. Add 0.4x AMPure XP beads to the pooled reaction and mix by flicking the tube. 
        For example if the pool volume is 200μl, add 80μl AMPure XP beads.

Incubate at room temperature for 10 minutes.
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13.7. Spin down and pellet the beads on a magnet until the eluate is clear and colourless (3-5 minutes). Whilst still on the magnet remove and discard the supernatant.
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13.8. Wash the pellet with 700μl 80% ethanol, without disturbing the pellet remove the ethanol and repeat this step. 
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13.9. Spin down, place the tube back on the magnet and remove any residual ethanol. Allow the pellet to air dry for 30 seconds then remove from the magnet and resuspend in 35μl nuclease free water.
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13.10. Incubate at 37°C for 10minutes, gently flicking the tube every couple of minutes to encourage elution.
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13.11. Pellet the beads on the magnet until clear and colourless then take 30μl into a clean 1.5ml tube.
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14. Adapter ligation
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14.1. Thaw NEBnext Quick ligation buffer, Short Fragment Buffer (SFB - ONT kit), and Elution Buffer (EB) at room temperature, vortex to mix, spin down and place on ice. 

Flick gently to mix then spin down the NEBNext Quick T4 ligase enzyme and the Native Adapter (NA - from ONT kit) and place on ice. 
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14.2. To the tube with 30μl cleaned barcoded samples, add the following:

 

Gently flick the tube to mix then spin down.
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14.3. Incubate at room temperature for 20 minutes

Note: At this point, you can remove your flow cell (FLO-MIN114) from the fridge to allow it to come to room temperature.
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14.4. Resuspend AMPure beads by vortexing then add 20μl to the reaction and mix by gently flicking the tube. Incubate at room temperature for 10 minutes.

Note: At this point you can start up MinKNOW and run the Flow Cell check.    
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14.5. Spin down and pellet on the magnet. Still on the magnet, remove and discard the supernatant.
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14.6. Wash the pellet with 125μl Short Fragment Buffer, flick the tube to resuspend the beads, spin down, then place back on the magnet to pellet the beads. 
Repeat this step.
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14.7. Spin down and place back on the magnet then remove any residual buffer. Resuspend in 15μl of Elution Buffer.
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14.8. Incubate at 37°C for 10 minutes, flicking every couple of minutes to aid elution.

During this time, thaw Flow Cell Flush (FCF), Flow Cell Tether (FCT), Sequencing Buffer (SB), and Library Beads (LB) at room temperature, mix by vortexing, then spin down and place on ice.

You can also perform the flow cell check. Insert the MinION device into your laptop, open MinKNOW, insert the room temperature flow cell into the MinION, then select Start, then Flow Cell Check.
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14.9. Spin down then place back on the magnet to pellet the beads. Remove and retain 15μl in a clean 1.5ml tube.
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15. Priming the flow cell and loading the library
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15.1. Prepare the flow cell priming mix by adding the following to a clean 1.5ml tube:

 

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2. Primer reconstitution 
The primers are received in lyophilized form and need to be reconstituted before use. Prepare a stock solution of 100μM primers.
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4. Primer pooling 
The PCR is currently performed with two reactions per sample with a different primer panel used in each reaction. Using the 10µM working stocks prepare the following primer panels:
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5. PCR reaction 
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7. Thermocycler conditions/ program
Set up the thermocycler conditions as follows:
 

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8. Gel electrophoresis (Optional)
You may check the amplification of PCR targets by running the PCR products on a 1% agarose gel or on a TapeStation using a D5000 DNA screen tape. 
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2.1. To reconstitute the lyophilised primers, use the nmole information on the sheet received with the primers.
Convert the nmol to μmol and then divide by the 100 μmol/L.
For example: for a primer with 24 nmoles, to make 100μM stock solution:

24 nmol / 1000 = 0.024 μmol
0.024μmol / 100μmol/L =0.00024 L
0.00024 L x 1000 = 0.24 ml or 240 µl
So add 240 µl to make a 100 μM solution.

In simple words, multiply the nmol value by 10. Example: for 24 nmol x10 = 240 µl to make a 100 μM solution.
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3.1. In a fresh tube add 5 µl of 100µM primer stock to 45 µl nuclease-free water to give 50 µl of 10µM primer.
Store at 4°C for frequent usage or -20°C for long-term storage.
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4.1. Genotyping panel
Pool the primers listed in 1.1 into a single tube as follows:
Add 10 μl of each forward and reverse primer to a single tube, pulse vortex and spin down and use this pool for PCR reactions. Scale up the volume as required.
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4.2. MDR panel
MDR panel has a common reverse primer for nine targets. To set the multiplex reaction pool the primers in 1.2 as follows:
Add 10 µl of each forward primer + 90 µl of common reverse primer + 10 µl of reverse primer for C19241A into a single tube. Pulse vortex, spin down and use this pool for PCR reactions. Scale up the volume as required.
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5.1. Thaw the primer panels and LongAmp taq 2x Mastermix on ice. 

Pulse vortex and spin down the primer panels, then return to ice. 
Do not vortex the LongAmp taq, mix by flicking or pipetting and spinning down before placing back on ice.

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5.2. Prepare the master mix as follows for the number of samples, plus two controls, and one more extra reaction to account for the pipetting error. 
 


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5.3. Dispense 20 µl of master mix per reaction into 0.2 ml PCR tubes. 

Add 5 µl of sample DNA. 

Mix well by pipetting and spin down.
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3. Primer Dilution 
Using the 100µM stock, prepare a 10uM working stock of each primer
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1.2. MDR Panel_version1


 

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1.1. Genotyping Panel_version1

 

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1. The following primers have been designed for identifying some S. Typhi genotypes and markers for AMR both in the chromosome and in a plasmid. These can be ordered lyophilised from your preferred oligo supplier.
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9.3. Incubate on a rotator for 5 minutes at room temperature.
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9.2. Add the required volume of resuspended AMPure XP beads to the reaction and mix by pipetting or flicking the tube.      
The volume is calculated as the 0.8x PCR reaction volume. 
For example: For 25 ul of PCR reaction = 0.8 x 25 = 20 ul of beads
  
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9.1. Prepare the    for use; resuspend by vortexing.

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9. PCR clean-up 
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9.4. Spin down the sample and pellet on a magnet. Keep the tube on the magnet, and pipette off the supernatant.
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9.5. Keep on the magnet, wash beads with 100 µL of freshly prepared 80% ethanol without disturbing the pellet
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9.6. Remove the  80% ethanol using a pipette and discard.
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9.7. Repeat steps 8.5 & 8.6.

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9.8. Spin down briefly and place the tube back on the magnet.
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9.9. Pipette off any residual 80% ethanol using a small tip (10-20µl).
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9.10. Briefly allow to dry.
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9.11. Remove the tube from the magnetic rack and resuspend the pellet in 20µl of nuclease free water, mix by gently flicking or pipetting.
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9.12. Incubate for 2 minutes at room temperature.
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9.13. Pellet beads on magnet until the eluate is clear and colourless.
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9.14. Still on the magnet, remove 20µl eluate and store it in a clean tube. 

Avoid disturbing the pelleted beads. If you find the beads keep getting drawn up the pipette tip, try removing 18µl instead.
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10. Quantification 
The PCR products are quantified using the      
 

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10.1. Standards
The Qubit dsDNA kit requires 2 standards for calibration-- Standard #1 and Standard #2. 
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10.3. Prepare Qubit working solution for the required number of samples and
standards as follows 

Each sample: 
Qubit dsDNA BR Reagent 1 µL
Qubit dsDNA BR Buffer 199 µL
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10.2. Label the tube lids. Do not label the side of the tube as this could interfere with the sample
reading
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10.4. Aliquot Qubit working solution to each tube:
standard tubes require 190µL of Qubit working solution
sample tubes require 198µL of Qubit working solution

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10.5.  Add 10µL of the standard to the appropriate tube.
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10.6. Add 2µL of each cleaned PCR product to the appropriate tube.

The final volume in each tube must be 200µL once the sample/standard has been added.
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10.7. Mix each tube vigorously by vortexing for 3–5 seconds.
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10.8. Allow all tubes to incubate at room temperature for 2 minutes, then proceed to “Read standards
and samples.
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10.9. On the Home screen of the Qubit Fluorometer, press DNA, then select 1X dsDNA broad range as the assay type. The Read standards screen is displayed. Press Read Standards to proceed.
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10.10. Insert the tube containing Standard #1 into the sample chamber, close the lid, then press Read
standard. When the reading is complete (~3 seconds), remove Standard #1.
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10.11. Insert the tube containing Standard #2 into the sample chamber, close the lid, then press Read
standard. When the reading is complete, remove Standard #2.
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10.12. The instrument displays the results on the Read standard screen. Then press run samples.
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10.13. On the assay screen, select the sample volume and units:

Press the + or – buttons on the wheel, or anywhere on the wheel itself, to select the sample
volume added to the assay tube (2µL).

From the unit dropdown menu, select the units for the output sample concentration (in this
case choose ng/µL).
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10.14. Insert a sample tube into the sample chamber, close the lid, then press Read tube. When the
reading is complete (~3 seconds), remove the sample tube and carefully record the calculated sample concentration.
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10.15. The top value (in large font) is the calculated concentration of the original sample. 

The bottom value is the dilution concentration. For information on interpreting the sample results, refer to the Qubit Fluorometer User Guide.
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10.16. Repeat step 9.14 until all samples have been read.
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10.17. All negative controls should ideally be ‘too low’ to read on the Qubit machine, but MUST be < 1ng
per ul. If your negative controls >1ng per ul, considerable contamination has occurred and you
must redo previous steps.
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11. 200fmol calculation 
The nanopore protocol recommends using 200fmol of your sample DNA in the first step of library preparation (End-preparation)
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11.1. Based on the DNA concentration obtained from Qubit, transfer 200fmol of DNA into a fresh tube and add nuclease free water to total 12.5ul.  

You can use the spreadsheet attached to help calculate the volume required of each sample. We have based the ng required of each sample using the average amplicon length for the primer panel to calculate the ng required for 200fmol product (1170ng for the genotyping panel and 1211ng for the MDR panel)

  

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11.2. Use this 12.5ul for the End-preparation reaction. 
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2.2. Add the required volume of nuclease free water to the primer vial, pulse vortex and spin down. This is the primer stock with 100µM concentration. Store at -20°C for long-term storage.
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15.2. Open the lid of the nanopore sequencing device and slide the flow cell's priming port cover clockwise so that the priming port is visible. After opening the priming port, check for any bubbles under the cover. Draw back a small volume to remove any bubbles (a few µLs). Visually check that there is continuous buffer from the priming port across the sensor array.
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15.3. Using a P1000 pipette, slowly load 800μL of the priming mix into the flow cell via the priming port.

Leave a small amount of liquid in the end of the pipette tip to ensure you do not introduce air into the flowcell.

Leave for 5 minutes.
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15.4. Mix the contents of the LIB tube by pipetting just before adding to the following library mix in a 1.5ml tube:

 

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15.5. Complete the flowcell priming by opening the SpotOn port cover and carefully loading 200μL of the priming mix into the priming port. As before, leave a small amount of liquid in the bottom of the tip to avoid the introduction of air bubbles.

When adding the priming mix, you may see a small amount of liquid come up through the SpotOn port. If you do, pause and allow the liquid to flow back into the flowcell before continuing putting through the priming mix. 
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15.6. Mix the prepared library mix gently by pipetting.

Add the library mix to the flowcell via the SpotOn port in a dropwise fashion, allowing each drop to flow into the flowcell before adding the next.
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15.7. Replace the SpotOn port cover and close the priming port, then replace the lid of your sequencing device.
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16. Starting the Sequencing Run
Open the ONT MinKNOW software and follow the steps below to set up and start your sequencing run.
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16.1. Click start, then start sequencing.
Create a name for you sequencing run, it is good practise to make this unique and identifiable for if you ever need to revisit the data. The date and an experiment name are recommended. In sample name you can put a number or repeat the experiment name - this is not as important as the run name. Then click continue.
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16.2. Select the kit used - this is SQK-NBD114.24 or SQK-NBD114.96 depending on whether you have purchased the 24 or 96 barcode kit. Click continue.
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16.3. In the run length options, set the run time to 8 hours. Click continue.
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16.4. In the basecalling options, select high accuracy basecalling. In the barcoding options, make sure barcoding is enabled and toggle to use barcode at both ends. Click continue until you reach the run overview, where you can double check the selected options, then click start run.
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17. A nanopore flow cell can be used multiple times, so it must be washed to remove
the library from the previous run.
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17.3. After the sequencing run, close the ONT MinKNOW software. Detach the MinION device from the laptop and leave the flowcell in the device. 
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17.1. Place the tube of Wash Mix (WMX) on ice. Do not vortex the tube and thaw one tube of Wash Diluent (DIL) at room temperature.
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17.2. In a clean 1.5 ml Eppendorf DNA LoBind tube, prepare the following Flow Cell Wash Mix:


 
Mix well by pipetting, and place on ice. Do not vortex the tube.
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17.4. Before removing the waste fluid, ensure that the flow cell priming port cover and SpotON sample port cover are closed.
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17.5. Using a P1000, remove all fluid from the waste channel through waste port 1. As both the flow cell priming port and SpotON sample port are closed, no fluid should leave the sensor array area.
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17.6. Rotate the flow cell priming port cover clockwise so that the priming port is visible.
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17.7. After opening the priming port, check for a small air bubble under the cover. Draw back a small volume to remove any bubbles.
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17.8. Load 400 µl of the prepared Flow Cell Wash Mix into the flow cell priming port, avoiding the introduction of air.
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17.9. Close the flow cell priming port and wait for 60 minutes.
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17.10. Before removing the waste fluid a second time, ensure that the flow cell priming port cover and SpotON sample port cover are closed.
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17.11. Using a P1000, remove all fluid from the waste channel through waste port 1. As both the flow cell priming port and SpotON sample port are closed, no fluid should leave the sensor array area.
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17.13. To run a second library straight away, follow the flow cell priming instructions in step 14.
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17.12. You can now either run a second library on the flow cell straight away or store the flow cell for later use.
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17.14. To store the flowcell for later use, thaw one tube of Storage Buffer (S) at room temperature, mix contents thoroughly by pipetting and spin down briefly.
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17.15. Rotate the flow cell priming port cover clockwise so that the priming port is visible.
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17.16. After opening the priming port, check for a small air bubble under the cover. Draw back a small volume to remove any bubbles.
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17.17. Slowly add 500 μl of Storage Buffer (S) through the flow cell priming port.
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17.18. Close the priming port.
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17.19. Using a P1000, remove all fluid from the waste channel through waste port 1. As both the flow cell priming port and SpotON sample port are closed, no fluid should leave the sensor array area.
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17.20. The flow cell can now be stored at 4-8°C.
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6.1. Positive control: Salmonella Typhi strain H58 DNA (or use appropriate control strain that is available)

Additionally, synthetic gene fragments such as G blocks (IDT) can be used as positive control. Use tviD amplicon sequence and the MDR amplicon sequences as positive control for genotyping and MDR panels respectively. Refer to the attached document for these sequences. 
   

G blocks are received lyophilized and need to be resuspended with Tris-EDTA pH 8.0 or nuclease free water (as recommended by the supplier). Centrifuge the tube briefly to bring down any contents of the tube sticking to the wall or cap. 


For control strain DNA or the G blocks, a concentration of at least 10 ng/μL is recommended for a stock dilution. For example if the stock has 1000ng then resuspend in 100 µl of diluent to get 10 ng/μl concentration. 
A working solution of 1 ng/μL is used in PCR reactions. To make working solution dilute 10 µl  of the 10ng/µl  stock solution in 90 µl  nuclease free water. Quantify using Qubit. 
Make 6 µl aliquots of the working solution to avoid contamination and repeated free-thaw cycles then use 5 µl for each positive control reaction.

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6. PCR Controls
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6.2. Negative control: 5µl nuclease-free water for a no template control