Whole Genome Sequencing for Rapid Characterization of Rabies Virus Using Nanopore Technology

The study was approved by the Medical Research Coordinating Committee of the National Institute for Medical Research (NIMR/HQ/R.8a/vol.IX/2788), the Ministry of Regional Administration and Local Government (AB.81/288/01), and Ifakara Health Institute Institutional Review Board (IHI/IRB/No:22-2014) in Tanzania; the University of Nairobi Institute of Tropical and Infectious Diseases (P947/11/2019) and the Kenya Medical Research Institute (KEMRI-SERU; protocol No. 3268) in Kenya; and the Research Institute for Tropical Medicine (RITM), Department of Health (2019-023) in the Philippines. Sequencing of samples originating from Nigeria was undertaken on archived diagnostic material collected as a part of national surveillance.

NOTE: Section 1-4 are prerequisites. Section 5-16 describe the sample-to-sequence-to-interpretation workflow for RABV nanopore sequencing (Figure 1). For subsequent steps in the protocol that need pulse centrifugation, centrifuge at 10-15,000 x g for 5-15 s.

1. Computational environment setup for sequencing and data analysis

1. Open the Oxford Nanopore Technology (ONT) website³⁷ and create an account to access nanopore-specific resources.
   1. Log in and install the ONT sequencing and basecalling software³⁸.
2. Open GitHub³⁹ and create an account.
   1. Go to the artic-rabv⁴⁰ and MADDOG repositories⁴¹ and follow the installation instructions.

2. Design or update the multiplex primer scheme

NOTE: Existing RABV schemes are available in the artic-rabv repository⁴⁰. When targeting a new geographic area, a new scheme should be designed, or an existing scheme modified to incorporate additional diversity.

1. Choose a genome reference set to represent the diversity in the study area; this is typically a set of publicly available sequences (e.g., from NCBI GenBank) or preliminary in-house data. Follow step 2.1.1 to use RABV-GLUE⁴², a RABV sequence data resource, to filter and download NCBI sequences and associated metadata.
   NOTE: Choose reference sequences with complete genomes (i.e., without gaps and masked bases). Choosing up to 10 sequences as a reference set for primer design is recommended. If the available sequence data is incomplete or not representative of the study area, refer to the advice^43,44,45 videodan Supplementary File 1.
   1. Navigate to the NCBI RABV Sequences by Clade page from the Sequence Data drop-down menu in RABV-GLUE. Click the Rabies Virus (RABV) link to access all available data or select a particular clade of interest. Use the filter option to Add filters that fit the desired criteria (e.g., country of origin, sequence length). Download sequences and metadata.
2. Generate a primer scheme for multiplex polymerase chain reaction (PCR) following the instructions provided by Primal Scheme⁴⁶. A 400 bp scheme with a 50 bp overlap is recommended to sequence low-quality samples. Download and save all outputs (do not edit the file or primer names).
   NOTE: The scheme will be indexed to the first sequence in the input fasta, henceforth referred to as the 'index reference' (Figure 2). See Supplementary File 1 for options to optimize primer performance.

3. Set up RAMPART and ARTIC bioinformatics pipelines

1. Refer to Supplementary File 2 to set up a directory structure to manage the input/output files for RAMPART and the ARTIC bioinformatics pipeline.

4. Biosafety and laboratory setup

1. Handle potentially rabies-positive samples in biosafety level (BSL) 2 or 3 conditions.
2. Ensure laboratory staff have completed rabies pre-exposure vaccination and undergo monitoring of immunity according to World Health Organization (WHO) recommendations³.
3. Ensure dedicated standard operating procedures and risk assessments, following national or international guidelines, are in place for the laboratory.
4. Required lab setup: Minimize contamination by maintaining physical separation between pre- and post-PCR areas. In laboratories with limited space or in-field lab settings, use portable glove boxes or makeshift lab stations to minimize contamination.
5. In this protocol, ensure to designate separate areas for:
   1. Sample extraction: Set up a BSL2/3 cabinet/glove box to handle biological material and perform inactivation and RNA extraction.
   2. Template area: Set up a BSL1 cabinet/glove box for the addition of template (RNA/cDNA) to the pre-prepared reaction master mix.
   3. Master mix area: Set up a designated clean area (BSL1 cabinet/glove box) for the preparation of reagent master mixes. There should be no template in this area.
   4. Post-PCR area: Set up a separate area for work on amplicons and sequencing library preparation.
      ​NOTE: All areas should be cleaned with a surface decontaminant and ultraviolet (UV)-sterilized before and after use.

5. Field sample collection and diagnosis

NOTE: Samples must be collected by trained and immunized personnel wearing personal protective equipment and following the referenced standard procedures^47,48,49.

1. Collect the sample via the foramen magnum (i.e., the occipital route), as described in detail in Mauti et al.⁵⁰.
2. Diagnose rabies in the field with rapid diagnostic tests and confirm in the laboratory using recommended procedures⁴⁷, such as the direct fluorescent antibody test (DFA), the direct rapid immunohistochemical test (DRIT)^51,52, or real time reverse transcription (RT)-PCR⁵³.
3. Use confirmed positive brain samples for RNA extraction or store in a freezer at -20 °C for 2-3 months or -80 °C for longer periods. Preserve RNA for storage and transport using a suitable DNA/RNA stabilization medium.

6. Sample preparation and RNA extraction (3 h)

NOTE: Use a spin column-based viral RNA extraction kit suitable for the sample type.

1. Prepare two ceramic bead tubes by filling a 2 mL PCR tube with approximately 200 µL tube full of 1.4 mm ceramic beads and label the tube.
2. Add the recommended volume of lysis buffer provided in the RNA extraction kit to the labeled PCR tube.
3. Get approximately a 3 mm cube from the brain sample confirmed with rabies infection using a wooden applicator and put into a labeled tube with sample ID and 100 µL of nuclease-free water into the tube labeled negative control.
   NOTE: Use closed tube bead-based homogenization to limit sample exposure. If not possible, use other suitable mechanical disruptors (e.g., rotor-based) or a manual micro pestle. However, these may be less effective than bead beating on hard surface to disrupt tissue (tissue samples may harden in certain storage media).
4. Disrupt the brain tissue manually using a wooden applicator stick and then vortex at maximum speed until complete tissue homogenization is achieved.
5. Centrifuge the lysate as per the manufacturer's instructions and use a pipette to transfer the supernatant to a new labeled microcentrifuge tube. Only use this supernatant in the subsequent steps.
6. Follow the RNA extraction kit's spin column instructions to obtain purified RNA.
7. Include a negative extraction control (NEC) here and take all the way through to the sequencing stage.

7. cDNA preparation (20 min)

1. In the master mix area, prepare a master mix for first strand cDNA synthesis according to the number of samples and controls to be processed (with an excess volume of 10% to ensure adequate reagent; Table 1). A no-template control (NTC) should be included at this stage.
2. Label 0.2 mL PCR strip tubes and aliquot 5 µL of the master mix into the tubes.
3. Take the prepared tubes to the template area. Add 5 µL of RNA into each labeled tube, including the NEC. Add 5 µL of nuclease-free water (NFW) to the NTC.
4. Incubate in a thermal cycler following the conditions mentioned in Table 1.
   NOTE: Optional pause point: cDNA can be stored at -20 °C for up to 1 month if necessary, but proceeding to PCR is preferred.

8. Primer pool stock preparation (1 h)

NOTE: This step is only necessary if making new stocks from individual primers, after which pre-prepared stock solutions can be used.

1. Prepare a primer pool of 100 µM stock in the master mix area.
2. Resuspend the lyophilized primers in 1x tris-EDTA (TE) buffer or NFW at a concentration of 100 µM each. Vortex thoroughly and spin down.
   ​NOTE: In the following steps, individual primers are separated into two primer pools-odd numbered (named Pool A) and even numbered (named Pool B)-to avoid interactions between primers flanking amplicon overlaps. These pools of primers generate overlapping 400 bp amplicons spanning the target genome.
3. Arrange all the odd numbered primers in a tube rack. Generate a primer pool stock by adding 5 µL from each primer to a 1.5 mL microcentrifuge tube labeled "primer scheme name – Pool A (100 µM)".
4. Repeat the process for all the even numbered primers and label as "primer scheme name – Pool B (100 µM)".
5. Dilute each primer pool 1:10 in molecular grade water to generate 10 µM primer stocks.
   ​NOTE: Make multiple aliquots of 10 µM primer dilutions and freeze them in case of degradation or contamination.

9. Multiplex PCR (5 h)

1. Prepare two PCR master mixes, one for each primer pool in the master mix area.
   1. Use a final concentration of 0.015 µM per primer. Calculate the required primer pool volume for the PCR reaction (Table 2) using the following formula:
      Primer pool volume = number of primers x reaction volume x 0.015/concentration (µM) of primer stock
2. Aliquot 10 µL each of Pool A master mix and Pool B master mix to labeled PCR strip tubes in the template area. For every sample, add 2.5 µL of cDNA (from step 3) to each of the corresponding labeled primer Pool A and B reactions. Excess cDNA can be stored at -20 °C.
3. Mix by gently flicking and pulse centrifuge.
4. Incubate the samples with the conditions mentioned in Table 2 on a PCR machine.
   ​NOTE: The program does not include a specific extension step due to the long annealing time of 5 min (required due to the high number of primers) and the short length of the amplicons (400 bp) which is sufficient for the extension.

10. PCR clean up and quantification (3.5 h)

1. Perform all work from this point on in the post-PCR area.
2. Aliquot solid-phase reversible immobilization (SPRI) beads into microcentrifuge tubes from the main bottle. Store at 4 °C.
3. Warm a SPRI bead aliquot to room temperature (RT; ~20 °C) and thoroughly vortex until the beads are fully resuspended in the solution.
4. In 1.5 mL tubes, combine the primer Pool A and primer Pool B PCR products for each sample. If necessary, add water to bring the volume to 25 µL.
5. Add 25 µL of SPRI beads to each sample (1:1 bead:sample ratio). Mix by pipetting up and down or gently tapping the tube.
6. Incubate at RT for 10 min, occasionally inverting or flicking the tubes.
7. Place on a magnetic rack until the beads and solution have fully separated. Remove and discard the supernatant, taking care not to disturb the bead pellet.
8. Wash twice with 80% ethanol (warmed to RT).
   1. Add 200 µL of ethanol to the pellet. Wait for 30 s to ensure the beads are washed properly.
   2. Carefully remove and discard the supernatant, trying not to touch the bead pellet.
   3. Repeat steps 10.8.1-10.8.2 to wash the pellet a second time.
9. Remove all traces of ethanol using a 10 µL tip. Air-dry until trace ethanol has evaporated (with small beads this happens quickly, ~30 s); when this happens, the pellet should go from shiny to matt. Take care not to over-dry (if the pellet is cracking, it is too dry), as this will affect DNA recovery.
10. Resuspend the beads in 15 µL of NFW and incubate at RT (off magnetic rack) for 10 min.
11. Return to the magnetic rack and transfer the supernatant (cleaned product) to a fresh 1.5 mL tube.
12. Prepare a 1:10 dilution of each sample in a separate tube (2 µL of product + 18 µL of NFW).
    NOTE: Be very careful at this stage to avoid cross-contamination. Only have one amplicon tube open at a time. Aliquot 18 µL of water into the tubes first (in a clean master mix area).
13. Measure the DNA concentration of each diluted sample using a highly sensitive and specific fluorometer, as described in protocols.io^54,55.

11. Normalization (30 min)

1. Use the normalization template (Supplementary File 3) and DNA concentration (ng/µL) of each sample to calculate the volume of diluted (or neat) sample required for 200 fmol of each sample in a total volume of 5 µL.
2. Label new PCR tubes and add computed volumes of NFW and sample to obtain normalized DNA.
3. Use the computed volume for undiluted (neat) samples if over 5 µL of the diluted sample is required to obtain 200 fmol.
   ​NOTE: Optional pause point: At this point, the cleaned-up PCR product can be stored at 4 °C for up to 1 week or placed at -20 °C for longer-term storage if needed.

12. End-prep and barcoding (1.5 h)

NOTE: The next steps assume the use of specific reagents from nanopore-specific barcoding and ligation sequencing kits (see Table of Materials for details). The protocol is transferable across different chemistry versions, but users should take care to use compatible kits, according to manufacturer instructions.

1. End repair and dA-tailing
   1. Set up the end-prep reaction for each sample mentioned in Table 3. Prepare a master mix according to the number of samples (plus 10% excess). Take care when pipetting as reagents are viscous.
   2. Add 5 µL of master mix into each tube of normalized DNA (5 µL). The total reaction mix should be 10 µL. Change the tips each time and only have one tube open at a time.
   3. Incubate in a thermal cycler under the conditions mentioned in Table 3.
2. Barcoding
   1. Aliquot the barcodes from the barcoding kit to PCR strip tubes at 1.25 µL/tube, and record barcode assigned to each sample.
   2. Add 0.75 µL of the end-prepped sample to its assigned barcode aliquot.
   3. Prepare a ligation master mix according to the number of samples (plus 10% excess) (Table 4).
   4. Add 8 µL of ligation master mix to the end-prepped sample + barcodes, giving a total reaction of 10 µL.
   5. Incubate in a thermal cycler using the conditions mentioned in Table 4.
3. SPRI bead clean up and DNA quantification
   1. Thaw short fragment buffer (SFB) at RT, mix by vortexing, pulse centrifuge, and place on ice.
   2. Pool all the barcoded samples together in a 1.5 mL lobind microcentrifuge tube. So as not to make the clean up volume too large to use, pool 12-24 samples (10 µL/sample), up to 48 samples (5 µL/sample), or up to 96 samples (2.5 µL/sample) from each native barcoding reaction.
   3. Add a 0.4x volume of SPRI beads to the barcoded pool. Mix gently (flicking or pipetting) and incubate at RT for 5 min.
   4. Place the samples on a magnet until the beads have pelleted and the supernatant is completely clear (~2 min). Remove and discard the supernatant. Take care not to disturb the beads.
   5. Wash twice with 250 µL of SFB.
      1. Remove the tube from the magnet and completely resuspend the pellet in 250 µL of SFB. Incubate for 30 s, pulse centrifuge, and return to the magnet.
      2. Remove the supernatant and discard.
   6. Repeat step 12.3.5 to perform a second SFB wash.
   7. Pulse centrifuge and remove any residual SFB.
   8. Add 200 µL of 80% (RT) ethanol to bathe the pellet. Remove and discard the ethanol, being careful not to disturb the bead pellet. Air-dry for 30 s or until the pellet has lost its shine.
   9. Resuspend in 22 µL of NFW at RT for 10 min.
   10. Place on the magnet, leave to settle for ~2 min, then carefully remove the solution and transfer to a clean 1.5 mL microcentrifuge tube.
   11. Use 1 µL to obtain the DNA concentration, as described previously (step 10.13).
       ​NOTE: Optional pause point: At this point, the library can be stored at 4 °C for up to 1 week or -20 °C for longer-term storage, but it is preferable to continue with adapter ligation and sequencing.

13. Sequencing (48 h maximum)

1. Prepare the computer (refer also to Prerequisites sections 1-4).
   1. Check that there is enough space to store new data (min 150 GB), data from old runs are backed up/moved to a server before deleting, and the latest version of MinKNOW is installed.
2. Remove the stored flow cell from the fridge and allow it to reach RT.
3. Adapter ligation (1 h)
   1. Pulse centrifuge the adapter mix and ligase and place on ice.
   2. Thaw elution buffer (EB), SFB, and ligation buffer at RT. Mix by vortexing, pulse centrifuge, and place on ice.
   3. Prepare the adapter ligation master mix (Table 5), combining reagents in the specified order in a low bind tube.
      NOTE: Alternatives for adapter ligation master mix reagents (Table 5) can be used depending on availability at the lab. See Supplementary File 3 and Table of Materials for a list of alternatives. Use computation in the Supplementary File 3 worksheet to get the volume of DNA library equivalent to 200 fmol. If less than 20 µL is computed, add NFW to make up to 20 µL.
   4. Mix by gentle flicking and pulse centrifuge. Incubate at RT for 20 min.
      NOTE: During incubation, start preparing the flow cell (section 13.5).
4. Clean up using SPRI beads (do not use ethanol as in earlier clean ups).
   1. Add a 0.4x volume of SPRI beads (RT) to the samples. Incubate at RT for 10 min, gently flick intermittently to aid mixing.
   2. Place on the magnet until the beads and solution have fully separated (~5 min). Remove and discard the supernatant; take care not to disturb the bead pellet.
   3. Wash twice with 125 µL of SFB.
   4. Resuspend the pellet completely with 125 µL of SFB by mixing with a pipette. Leave to incubate for 30 s.
   5. Pulse centrifuge to collect liquid at the tube base and place on the magnet. Remove the supernatant and discard.
   6. Repeat steps 13.4.4-13.4.5 to wash the pellet a second time.
   7. Pulse centrifuge and remove excess SFB.
   8. Resuspend in 15 µL of EB and incubate for 10 min at RT.
   9. Return to the magnet for ~2 min and then carefully transfer the solution to a clean 1.5 mL microcentrifuge tube.
   10. Quantify 1 µL of the eluted library, as described previously in step 10.13
       NOTE: For best results, proceed directly to MinION sequencing; however, the final library can be stored in EB at 4 °C for up to 1 week if needed.
5. Run a flow cell quality check.
   1. Connect the sequencing device to a laptop and open the sequencing software.
   2. Select flow cell type, and then click Check Flow Cell and Start Test.
   3. Once complete, the total number of active (i.e., viable) pores will be displayed. A new flow cell should have >800 active pores; if it does not, contact the manufacturer for a replacement.
6. Priming and loading the flow cell (20 min)
   1. Thaw the following reagents at RT and then place sequencing buffer, a flush tether, flush buffer, and loading beads on ice.
   2. Vortex the sequencing buffer and flush buffer, pulse centrifuge, and place on ice.
   3. Pulse centrifuge the flush tether and mix by pipetting; then place on ice.
   4. Prepare the flow cell priming mix by adding 30 µL of flush tether directly to the tube of flush buffer from a flow cell priming kit and mix by pipetting.
   5. Mix the loading beads by pipetting immediately prior to use as they settle quickly.
   6. In a fresh tube, prepare the final library dilution for sequencing, as mentioned in Table 5.
      NOTE: Use computation in the Supplementary File 3 worksheet to get the volume of DNA library equivalent to 50 fmol. If less than 12 µL is computed, add EB to make up to 12 µL.
   7. Flip back the sequencing device lid and slide the priming port cover clockwise so that the priming port is visible (Figure 3)
   8. Remove air bubbles carefully by setting a P1000 pipette to 200 µL, insert the tip into the priming port, and turn the wheel until a small volume entering the pipette tip is seen (max turn to 230 µL).
   9. Load 800 µL of flow cell priming mix into the flow cell via the priming port, taking care to avoid bubbles.
   10. Leave for 5 min.
   11. Lift the sample port cover gently and load 200 µL of priming mix into the flow cell via the priming port using a P1000 pipette.
   12. Pipette the library mix up and down prior to loading, ensuring loading beads in the master mix are resuspended before loading.
   13. Load 75 µL of library mix to the flow cell via the sample port in a dropwise fashion. Ensure that each drip flows into the port before adding the next.
   14. Replace the sample port cover gently, making sure the bung enters sample port.
   15. Close the priming port and replace the sequencing device lid.
7. Sequencing run (48 h maximum)
   1. Connect the sequencing device to the laptop and open the sequencing software.
   2. Click start and then click Start Sequencing.
   3. Click New Experiment and follow the sequencing software graphical user interface (GUI) workflow to set up the parameters for the run.
   4. Type in the experiment name and sample ID (e.g., rabv_run1), and choose the Flow Cell Type from the drop-down menu.
   5. Continue to kit selection and choose the relevant ligation sequencing kit and native barcoding kit(s) used.
   6. Continue to Run options. Keep the defaults, unless it is desired for the run to stop automatically after a certain number of hours (runs can be stopped manually at any time).
   7. Continue to Basecalling. Choose to turn Basecalling On or Off according to the computing resources (see computer setup). Choose Edit Options under barcoding and ensure Barcode Both Ends is turned on. Save and continue to the output section.
   8. Accept the defaults and continue to final review, check the settings, and record the details in worksheet (Supplementary File 3). Click Start.
      NOTE: If the flow cell is being reused, adjust the starting voltage (in the advanced section of the run options), as indicated by the scheme in Supplementary File 3.
   9. Record the initial active channels-if this is significantly lower than the quality control (QC) check, restart the sequencing software. If still lower, then reboot the computer.
   10. Record the initial channels in strand versus single pore to give an approximate pore occupancy. This number will fluctuate, so give an approximation.
   11. Monitor the run as it progresses.

14. Live and offline basecalling

NOTE: These instructions assume that the pre-existing directory structure provided in the artic-rabv repository and that Prerequisites sections 1 and 3 of the protocol have been followed.

1. On your local file system, create a new directory called analysis, where you will store all of your analysis outputs. To organize further: create a sub-directory with the name of your project and inside that a new directory for the run, using the sample ID provided to the MinKNOW as the run_name. Do this in one command as follows:
   mkdir -p
   analysis/project_name/run_name
   Then navigate to its location:
   cd
   path/analysis/project_name/run_name
2. Live basecalling
   NOTE: To perform nanopore basecalling in real time, laptops require a NVIDIA CUDA-compatible graphics processing unit (GPU). Ensure instructions for the GPU basecalling setup have been performed using the guppy protocol⁵⁶.
   1. During run setup, turn live basecalling on.
   2. Use RAMPART to monitor the sequencing coverage in real time, as per the instruction below.
   3. In the computer's terminal, activate the artic-rabv conda environment:
      conda activate artic-rabv
   4. Create a new directory for the rampart output inside the run_name directory and navigate into it:
      cd /path/analysis/project_name/run_name
      mkdir rampart_output
      cd rampart_output
   5. Create a barcodes.csv file to pair the barcodes and sample names. It should have one line per barcode and only specify barcodes that are present in the library, with the headings "barcode" and "sample". Follow the example in the artic-rabv directory:
      ​analysis/example_project/example_run/rampart_output/barcodes.csv
   6. Start RAMPART by providing the relevant protocol folder and path to the fastq_pass folder in the MinKNOW output for the run:
      rampart –protocol /path/rampart/scheme_name_V1_protocol – basecalledPath <insertpathTo Fastqpassfolder>
   7. Open a browser window and navigate to localhost:3000 in the URL box. Wait for sufficient data to be basecalled before results appear on the screen.
3. Offline basecalling (performed post-run)
   1. If live basecalling was not set, the output from MinKNOW will be raw signal data (fast5 files). One will not be able to use RAMPART during the run. Convert the fast5 files to basecalled data (fastq files) post-run using guppy (see setup in Prerequisites step 1.1.1.). Run RAMPART post-hoc on the basecalled data.
   2. Run the guppy basecaller:
      guppy_basecaller -c dna_r9.4.1_450bps_fast.cfg -i /path/to/reads/fast5_* -s /path/analysis/project_name/run_name -x auto -r
      -c is the config file to specify the basecalling model, -i is the input path, -s is the save path, -x specifies basecalling by the GPU device (exclude if using the computer version of guppy), and -r specifies to search input files recursively.
      NOTE: The config file (.cfg) can be changed to a high-accuracy basecaller by replacing _fast with _hac, although this will take significantly longer.

15. Washing the flow cells

1. The flow cells can be washed and reused to sequence new libraries if pores are still viable. See instructions for washing at the ONT flow cell wash protocol⁵⁷.

16. Analysis and interpretation

1. Consensus sequence generation with ARTIC bioinformatics pipeline
   1. Follow the instructions detailed in the artic-rabv GitHub repository⁴⁰ in the rabv_protocols folder to generate consensus sequences from raw fast5 or basecalled fastq files.
      NOTE: Refer to Artic pipeline – Core pipeline⁵⁸ for further guidance.
2. Optional: Analyze the average read depth per amplicon.
   1. Adapt the scripts available from the artic-rabv repository, referring to Supplementary File 1. Briefly, in-depth statistics are generated using SAMtools⁵⁹ and coverage per nucleotide plotted in R.
3. Phylogenetic analysis using GLUE
   1. From RABV_GLUE⁴², select Analysis > Genotyping and Interpretation tab > Add Files, selecting the fasta file of consensus sequences.
   2. Click Gönder and wait. Once the analyses are complete, the Show Analysis button will be available to click, showing clade and subclade assignments, coverage per gene, variation from reference sequences, and closest relative.
   3. Relevant contextual sequences can also be identified in the Sequence Data > NCBI Sequences by Clade section.
   4. Select the clade identified or click Rabies Virus (RABV) to see all the available sequences.
   5. Filter for relevant sequences (e.g., country of origin).
   6. Download these sequences and corresponding metadata for analysis and comparison.
4. Lineage assignment using MADDOG⁴¹
   1. Pull the MADDOG repository from GitHub to ensure you are working with the most up-to-date version.
   2. Create an assignment folder within the local MADDOG repository (previously created in the Prerequisites section) called the run name.
   3. Inside the folder, add the fasta file containing the consensus sequences.
   4. Add a metadata file to the folder.
      ​NOTE: This file must be a csv with 4 columns called 'ID,' 'country,' 'year,' and 'assignment,' detailing the sequence IDs, the country of sampling, and the year of sample collection, while the 'assignment' column should be blank.
   5. In the command line interface, activate the conda environment: conda activate MADDOG.
   6. In the command line interface, navigate to the MADDOG repository folder.
   7. Initially, undertake lineage assignment on sequences to check for any potential abnormalities and to identify if running the longer lineage designation step would be appropriate. For this, type this in the command line: sh assignment.sh.
   8. When prompted, enter Y to indicate you have pulled the repository and are working with the most up-to-date version of MADDOG.
   9. When prompted, enter the name of the folder within the MADDOG repository folder that contains the fasta file.
   10. When the lineage assignment is complete, check the output file in your folder. If the output is as expected and there are multiple sequences assigned to the same lineage, run the lineage designation.
   11. If running lineage designation, delete the assignment output file just created.
   12. In the terminal, inside the MADDOG repository folder, run the command sh designation.sh.
   13. When prompted enter Y to indicate you have pulled the repository and are working with the most up-to-date version of MADDOG.
   14. When prompted, enter the folder name within the MADDOG repository folder containing the fasta file and metadata. This outputs lineage information about each sequence, a phylogeny of the new and relevant previous sequences (from 16.3.6), hierarchical information about the lineages, and details of potentially emerging lineages and areas of undersampling.
       NOTE: Full details of the protocol, usage, and outputs can be found in Campbell et al.⁶⁰.
   15. When the initial analysis has been completed, if prompted to also test for emerging and undersampled lineages, enter Y, if this is required. Otherwise, enter N.
   16. If prompted to confirm newly found lineages, enter Y and follow instructions in the resultant NEXT_STEPS.eml file. Otherwise, enter N.