Ebola virus sequencing protocol

This document is part of the Ebola virus Nanopore sequencing protocol package:

http://artic.network/ebov/

Related documents:

Ebola primer scheme:

https://github.com/artic-network/primer-schemes/tree/master/ZaireEbola/V3

Ebola virus Nanopore sequencing protocol:

http://artic.network/ebov/ebov-seq-sop.html

Ebola virus Nanopore sequencing kit-list:

http://artic.network/ebov/ebov-seq-kit.html

Funded by the Wellcome Trust

Collaborators Award 206298/Z/17/Z --- ARTIC network

Preparation

Equipment required:

Consumables required:

Safety, containment and contamination recommendations

Protocol

Part 1: cDNA synthesis with Superscript IV reverse transcriptase

NOTE ON HOOD PREPARATION: To prevent cross contamination of both the sample and other reagents, this should be carried out in the SAMPLE PREPARATION HOOD, which is pre-sterilised with UV and treated with MediPal wipes, DNAway and RNAseZap reagent. Wipe down the hood with each sequentially, allowing 5 minutes for drying between each. Pipettes should also be treated in the same way, and UV treated for 30 mins between library preparations.

1. Set up the following reaction:

   50µM random hexamers		1µL
   10mM dNTPs mix (10mM each)	1µL
   Template RNA	11µL
   TOTAL	12µL

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.

1. Gently mix (avoid vortexing) then pulse spin the tube to ensure maximum contact with the thermal cycler.

2. Incubate the reaction as follows:

   Denaturation	65°C	5 mins
   Primer annealing	Ice	1 mins

3. Add the following to the annealed template RNA:

   SSIV Buffer		4µL
   100mM DTT	1µL
   RNaseOUT RNase Inhibitor	1µL
   SSIV Reverse Transcriptase	1µL
   TOTAL	20µL

4. Gently mix (avoid vortexing) then pulse spin the tube to ensure maximum contact with the thermal cycler.

5. Incubate the reaction as follows:

   Extension	42°C	90 mins
   Inactivation	70°C	10 mins

6. cDNA is now ready for amplicon generation.

Part 2: Ebola Amplicon Preparation

NOTE ON HOOD PREPARATION: To prevent cross contamination of both the sample and other reagents, this should be carried out in the MASTERMIX HOOD, which is pre-sterilised with UV and treated with MediPal wipes, DNAway and RNAseZap reagent. Wipe down the hood with each sequentially, allowing 5 minutes for drying between each. Pipettes should also be treated in the same way, and UV treated for 30 mins between library preparations.

Primer dilution and preparation

1. Ebola primers for this protocol were designed using Primal Scheme and generate overlapping 400nt amplicons. Primer names and dilutions are listed in the table below.

2. Primers should be prepped and aliquoted PRIOR TO DEPARTURE in a STERILE PCR CABINET. At NO stage should primers or PCR reagents be anywhere near the template or amplicons until use.

3. Resuspend lyophilised primers at a concentration of 100µM each

4. Generate primer pool stocks by adding 5µL of each primer pair to a 1.5mL Eppendorf labelled “Pool 1, 100µM” or “Pool 2, 100µM”. Total volume should be 505µL of Pool 1 and 530µL of Pool 2. This is a 10x stock of each primer pool.

5. Dilute this primer pool 1:10 in molecular grade water, to generate 10µM primer stocks. Recommend that multiple aliquots of each primer pool are made to account for any risks of degradation of contamination.

NOTE: Primers need to be used at a final concentration of 0.015µM per primer. In this case, Pool 1 has 101 primers in it so the requirement is 3.8µL of 10µM primers Pool 1 per 25µL reaction. Pool 2 has 106 primers so needs 4.0µL of 10µM primers Pool 2 per 25µL reaction. For other schemes, adjust the volume added appropriately.

1. Set up the amplicon PCR reactions as follows in 0.5mL thin-walled PCR or strip-tubes:

   Reagent	Pool 1	Pool 2
   NEB Q5 Polymerase 2X MasterMix	12.5µL	12.5µL
   Primer Pool 1 or 2 (10µM)	3.8µL	4.0µL
   Water	6.2µL	6.0µL
   TOTAL	22.5µL	22.5µL

NOTE: This should be carried out in the mastermix hood and cDNA should not be taken anywhere near the mastermix hood at any stage.

1. In the TEMPLATE HOOD add 2.5µL of cDNA to each Pool1 and Pool2 reaction mix and mix well.

2. Pulse centrifuge the tubes to remove any contents from the lid.

3. Set up the cycling conditions as follows:

   Step	Temperature	Time	Cycles
   Heat Activation	98°C	30 seconds	1
   Denaturation	98°C	15 seconds	25-35
   Annealing	65°C	300 seconds	25-35
   Hold	4°C	Indefinite	1

NOTE: Cycle number should be 25 for Ct18-21 up to a maximum of 35 cycles for Ct 35

1. Clean-up the amplicons using the following protocol in the TEMPLATE HOOD:

   1. Combine the entire contents of “Pool1” and “Pool2” PCR reactions for each biological sample into to a single 1.5mL Eppendorf tube.

   2. Mix sample gently, avoid vortexing.

   3. Ensure Aline beads are well resuspended by thoroughly mixing prior to addition to the sample. Mixture should be a homogenous brown colour.

   4. Add an equal volume of Aline beads to the tube and mix gently by either flicking or pipetting. This should be approximately 50µL, so add 50µL of beads.

   5. Pulse centrifuge the tubes to remove any beads or solution from the lid or side of the tube.

   6. Incubate for 5 mins at RT.

   7. Place on magnetic rack and incubate for 2 mins or until the beads have pelleted against the magnet and the solution is completely clear.

   8. Carefully remove and discard the solution, being careful not to displace the bead pellet.

   9. Add 200µL of room-temperature 70% ethanol to the pellet.

   10. Carefully remove and discard ethanol, being careful not to displace the bead pellet.

   11. Repeat steps i to j to wash the pellet again.

   12. Briefly pulse centrifuge the pellet and carefully remove as much ethanol as possible using a 10µL tip.

   13. Allow the pellet to dry for 1 mins, being careful not to overdry (if the pellet starts to crack then it is too dry).

   14. Resuspend pellet in 30µL of water, and incubate for 2 mins.

   15. Place on magnet and CAREFULLY remove water and transfer to a clean 1.5mL Eppendorf tube. MAKE SURE that no beads are transferred into this tube. In some cases a pulse centrifugation can be used to pellet residual beads.

   16. Quantify the amplicons pools using the Quantus Fluorometer following ONE dsDNA protocol.

Part 3: Quantus Quantification of Amplicon Pools

1. Set up the required number of 0.5mL tubes samples.

NOTE: Use only thin-wall, clear, 0.5mL PCR tubes.

1. Label the tube lids. Do not label the side of the tube as this could interfere with the sample reading.

2. Add 199µL ONE dsDNA dye solution to each tube.

3. Add 1µL of each user sample to the appropriate tube.

NOTE: Use a P2 pipette for highest accuracy.

1. Mix each sample vigorously by vortexing for 3–5 seconds.

2. Allow all tubes to incubate at room temperature for 2 minutes before proceeding.

3. 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.

1. Add 200µL ONE dsDNA Dye solution to two 0.5mL tubes.

2. Add 1µL Lambda DNA standard 400 ng/µL provided in the kit to one of the tube. These two tubes are the blank sample and standard required to perform the single point calibration procedure.

3. 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’.

4. On the home screen navigate to ‘Sample Volume’ and set it to 1 ul then ‘Units’ and set it to ng/µL.

5. Load the first sample into the reader and close the lid. The sample concentration is automatically read when you close the lid.

6. Repeat step 12 until all samples have been read.

7. The value displayed on the screen is the dsDNA concentration, carefully record all results in a spreadsheet or laboratory notebook.

Part 4: Barocoding and adaptor ligation: One-pot protocol.

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. If you have the time I would recommend using the double incubation times in blue, if you are in a hurry the times in red are a good compromise between speed and efficiency.

1. Set up the following end-prep reaction for each biological sample:

   DNA (20 ng)	16.7µL
   Ultra II End Prep Reaction Buffer	2.3µL
   Ultra II End Prep Enzyme Mix	1µL
   Total	20µL

NOTE: Quantity of amplicons can vary from 10-50ng, any more than this and the molarity of DNA ends will be too high for efficient barcoding. You need to have 6 samples per native barcoded library to have sufficient material at the end.

1. Incubate at RT for 20 mins then 65°C for 10 mins

2. Place on ice for 30 secs

3. Add the following directly to the previous reactions:

   NBXX barcode	2.5µL
   Ultra II Ligation Master Mix	22.5µL
   Ligation Enhancer	0.7µL
   Total	45.7µL

NOTE: Use a SINGLE barcode per biological sample.

1. Incubate at RT for 30 mins, 70°C for 10 mins then place on ice.

NOTE: This is to inactivate the DNA ligase to prevent barcode crossover.

1. Pool all barcoded fragments together into a clean 1.5 ml Eppendorf tube

2. Add 45.7µL Aline beads per sample.

3. Incubate for 5 mins.

4. Place on a magnet rack for 2 mins or until clear.

5. Remove solution.

6. Add 200µL 70% ethanol to the tube still on the magnetic rack.

7. Remove and discard ethanol without disturbing the pellet.

8. Repeat steps 11 and 12.

9. Spin down and remove residual 70% ethanol and air dry for 1 min.

10. Resuspend in 31µL EB.

11. Incubate off the magnetic rack for 2 mins.

12. Replace on magnetic rack.

13. Wait until clear and then carefully remove solution and transfer to a clean 1.5mL Eppendorf tube.

14. Remove 1µL and assess concentration by Quantus as described in previous section.

15. Set up the following adapter ligation reaction:

    Cleaned-up barcoded amplicon pools (~60ng)	30µL
    NEBNext Quick Ligation Reaction Buffer (5X)	10µL
    AMII adapter mix	5µL
    Quick T4 DNA Ligase	5µL
    Total volume	50µL

16. Incubate at RT for 30 mins.

17. Add 50µL Aline beads

18. Incubate for 5 mins

19. Place on a magnetic rack until clear

20. Remove supernatant

21. Add 200µL SFB and resuspend by flicking

CAUTION: do not use 80% ethanol

1. Place on magnetic rack until clear

2. Remove supernatant

3. Repeat SFB wash

4. Spin down and remove residual SFB

5. Add 15µL EB and resuspend by flicking

6. Incubate at RT for 2 mins.

7. Place on magnetic rack.

8. Carefully transfer solution to a clean 1.5mL Eppendorf tube.

9. Remove 1µL and assess concentration by Quantus (wait until beads have settled before measuring).

NOTE: Library can be now be stored at 4°C if required, but for best results it would be best to proceed immediately to sequencing.

Part 5: Priming and loading the SpotON flow cell

1. Thaw the following at RT before placing on ice:
   • Sequencing buffer (SQB)
   • Loading beads (LB)
   • Flush buffer (FLB)
   • Flush tether (FLT)

2. Add 30 ul FLT to the tube of FLB and mix well.

3. Flip back the MinION lid and slide the priming port cover clockwise so that the priming port is visible.

IMPORTANT: Care must be taken when drawing back buffer from the flow cell. The array of pores must be covered by buffer at all times. Removing more than 20-30µL risks damaging the pores in the array.

1. After opening the priming port, check for small bubble under the cover. Draw back a small volume to remove any bubble (a fewµLs):
   • Set a P1000 pipette to 200µL
   • Insert the tip into the priming port
   • Turn the wheel until the dial shows 220-230µL, or until you can see a small volume of buffer entering the pipette tip.

2. Load 800µL of FLB plus FLT into the flow cell via the priming port, using the dial-down method described in step 5, avoiding the introduction of air bubbles.

3. Wait for 5 minutes.

4. In a new tube prepare the library dilution for sequencing:

   Reagent	Volume
   SQB	37.5µL
   LB	25.5µL
   Library (~30ng)	12µL
   Total	75µL

5. Gently lift the SpotON sample port cover to make the SpotON sample port accessible.

6. Load 200µL of the priming mix into the flow cell via the priming port (NOT the SpotON sample port), avoiding the introduction of air bubbles.

7. Mix the prepared library gently by pipetting up and down just prior to loading.

8. Add 75µL of the prepared library to the flow cell via the SpotON sample port in a dropwise fashion. Ensure each drop siphons into the port before adding the next.

9. Gently replace the SpotON sample port cover, making sure the bung enters the SpotON port, close the priming port and replace the MinION lid.

10. Double–click the MinKNOW icon located on the desktop to open the MinKNOW GUI.

11. If your MinION was disconnected from the computer, plug it back in.

12. Choose the following flow cell type from the selector box:

    • FLO-MIN106 : R9.4.1 flowcell

13. Then mark the flow cell as Selected.

14. Click the New Experiment button at the bottom left of the GUI.

15. 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 LSK109 as there is no option for native barcoding (NBD104)

    • Run Options

      Set the run length, usually 1-2 hours.

    • Basecalling

      Leave basecalling turned on and check the HAC (high accuracy model) is selected

    • Output

      The number of files that MinKNOW will write to a single folder. By default this is set to 4000

16. Click Start run.

17. Allow the script to run to completion.

18. The MinKNOW Experiment page will indicate the progression of the script; this can be accessed through the Experiment tab that will appear at the top right of the screen

19. Monitor the Message panel on the right hand side for errors.