Here, we present a rapid and cost-effective workflow for characterizing rabies virus (RABV) genomes using nanopore technology. The workflow is intended to support genomics-informed surveillance at a local level, providing information on circulating RABV lineages and their placement within regional phylogenies to guide rabies control measures.
Genomic data can be used to track the transmission and geographic spread of infectious diseases. However, the sequencing capacity required for genomic surveillance remains limited in many low- and middle-income countries (LMICs), where dog-mediated rabies and/or rabies transmitted by wildlife such as vampire bats pose major public health and economic concerns. We present here a rapid and affordable sample-to-sequence-to-interpretation workflow using nanopore technology. Protocols for sample collection and the diagnosis of rabies are briefly described, followed by details of the optimized whole genome sequencing workflow, including primer design and optimization for multiplex polymerase chain reaction (PCR), a modified, low-cost sequencing library preparation, sequencing with live and offline base calling, genetic lineage designation, and phylogenetic analysis. Implementation of the workflow is demonstrated, and critical steps are highlighted for local deployment, such as pipeline validation, primer optimization, inclusion of negative controls, and the use of publicly available data and genomic tools (GLUE, MADDOG) for classification and placement within regional and global phylogenies. The turnaround time for the workflow is 2-3 days, and the cost ranges from $25 per sample for a 96 sample run to $80 per sample for a 12 sample run. We conclude that setting up rabies virus genomic surveillance in LMICs is feasible and can support progress toward the global goal of zero dog-mediated human rabies deaths by 2030, as well as enhanced monitoring of wildlife rabies spread. Moreover, the platform can be adapted for other pathogens, helping to build a versatile genomic capacity that contributes to epidemic and pandemic preparedness.
The rabies virus (RABV) is a lyssavirus in the Rhabdoviridae family that causes a fatal neurological disease in mammals1. Although rabies is 100% preventable by vaccination, it remains a major public health and economic concern in endemic countries. Of the 60,000 human rabies deaths estimated to occur each year, over 95% are in Africa and Asia where dogs are the primary reservoir2. In contrast, dog vaccination has led to the elimination of dog-mediated rabies across Western Europe, North America, and much of Latin America. In these regions, reservoirs of rabies are now restricted to wildlife, such as bats, raccoons, skunks, and wild canids3. Across Latin America, the common vampire bat is a problematic source of rabies due to regular spillover transmission from bats to both humans and livestock during nightly blood feeding4. The annual global economic impact of rabies is estimated to be $8.6 billion, with livestock losses accounting for 6%5.
Sequence data from viral pathogens combined with metadata on the timing and source of infections can provide robust epidemiological insights6. For RABV, sequencing has been used to investigate the origin of outbreaks7,8, identify host associations with wildlife or domestic dogs8,9,10,11,12, and trace sources of human cases13,14. Outbreak investigations using phylogenetic analysis have indicated that rabies emerged in the formerly rabies-free province of Bali, Indonesia, through a single introduction from the nearby endemic areas of Kalimantan or Sulawesi15. Meanwhile, in the Philippines, an outbreak on Tablas Island, Romblon Province was proven to be introduced from the main island of Luzon16. Viral genomic data have also been used to better understand pathogen transmission dynamics required for targeting control measures geographically. For example, genomic characterization of RABV illustrates the geographic clustering of clades17,18,19, co-circulation of lineages20,21,22, human-mediated viral movement17,23,24, and metapopulation dynamics25,26.
Disease monitoring is one important function of genomic surveillance that has been enhanced with the global increase in sequencing capacity in response to the SARS-CoV-2 pandemic. Genomic surveillance has supported the real-time tracking of SARS-COV-2 variants of concern27,28 and associated countermeasures6. Advances in accessible sequencing technology, such as nanopore technology, have led to improved and more affordable protocols for the rapid sequencing of both human29,30,31,32 and animal33,34,35 pathogens. However, in many rabies endemic countries, there are still barriers to operationalizing pathogen genomic surveillance, as shown by global disparities in SARS-CoV-2 sequencing capacity36. Limitations in laboratory infrastructure, supply chains, and technical knowledge make the establishment and routinization of genomic surveillance challenging. In this paper, we demonstrate how an optimized, rapid, and affordable whole genome sequencing workflow can be deployed for RABV surveillance in resource-limited settings.
An accessible RABV, nanopore-based, whole genome sequencing workflow was developed by Brunker et al.61, using resources from the ARTIC network46. Here, we present an updated workflow, with complete sample-to-sequence-to-interpretation steps. The workflow details the preparation of brain tissue samples for whole genome sequencing, presents a bioinformatics pipeline to process reads and generate consensus sequences, and highlights two rabies-specific tools to automate lineage assignment and determine phylogenetic context. The updated workflow also provides comprehensive instructions for the setup of appropriate computational and laboratory workspaces, with considerations for implementation in different contexts (including low-resource settings). We have demonstrated the successful implementation of the workflow in both academic and research institute settings in four RABV endemic LMICs with no or limited genomic surveillance capacity. The workflow has proven resilient to application across diverse settings, and comprehensible by users with varying expertise.
This workflow for RABV sequencing is the most comprehensive publicly available protocol (covering sample-to-sequence-to-interpretation steps) and specifically adapted to reduce both startup and running costs. The time and cost required for library preparation and sequencing with nanopore technology is greatly reduced relative to other platforms, such as Illumina61, and continual technology developments are improving sequence quality and accuracy to be comparable with Illumina62.
This protocol is designed to be resilient in diverse low-resource contexts. By referring to the troubleshooting and modifications guidance provided alongside the core protocol, users are supported to adapt the workflow to their needs. The addition of user-friendly bioinformatic tools to the workflow constitutes a major development to the original protocol, providing rapid and standardized methods that can be applied by users with minimal prior bioinformatics experience to interpret sequence data in local contexts. The capacity to do this in situ is often limited by the need to have specific programming and phylogenetic skills, which require an intensive and long-term skills training investment. While this skillset is important to thoroughly interpret sequence data, basic and accessible interpretation tools are equally desirable in order to capacitate local "sequencing champions", whose core expertise may be wet lab based, enabling them to interpret and take ownership over their data.
As the protocol has been undertaken for a number of years in several countries, we now can provide guidance on how to optimize multiplex primer schemes to improve coverage and deal with accumulated diversity. Efforts have also been made to help users improve the cost-effectiveness or to allow for ease of procurement in a given region, which is typically a challenge for the sustainability of molecular approaches63. For example, in Africa (Tanzania, Kenya, and Nigeria), we opted for blunt/TA ligase master mix at the adapter ligation step, which was more readily available from local suppliers and a cheaper alternative to other ligation reagents.
From experience, there are several ways of reducing the cost per sample and per run. Reducing the number of samples per run (e.g., from 24 down to 12 samples) can extend the life of flow cells over multiple runs, whereas increasing the number of samples per run maximizes the time and reagents. In our hands, we were able to wash and reuse flow cells for one in every three sequencing runs, enabling an additional 55 samples to be sequenced. Washing the flow cell immediately after use, or if not possible, removing the waste fluid from the waste channel after every run, seemed to preserve the number of pores available for a second run. Taking into consideration the initial number of pores available in a flow cell, one run can also be optimized to plan how many samples to run in a particular flow cell.
Though the workflow aims to be as comprehensive as possible, with the addition of detailed guidance and signposted resources, the procedure is still complex and can be daunting for a new user. The user is encouraged to seek in-person training and support, ideally locally, or alternatively through external collaborators. In the Philippines for example, a project on capacity building within regional laboratories for SARS-CoV-2 genomic surveillance using ONT has developed core competencies among health care diagnosticians that are readily transferrable to RABV sequencing. Important steps, such as SPRI bead clean up, can be difficult to master without hands-on training, and ineffective clean up can damage the flow cell and compromise the run. Sample contamination is always a major concern when amplicons are being processed in the lab and can be difficult to eliminate. In particular, cross-contamination between samples is extremely difficult to detect during post-run bioinformatics. Good laboratory technique and practices, such as maintaining clean work surfaces, separating pre- and post-PCR areas, and incorporating negative controls, are imperative to ensure quality control. The fast pace of nanopore sequencing developments is both an advantage and disadvantage for routine RABV genomic surveillance. Continuing improvements to nanopore's accuracy, accessibility, and protocol repertoire widen and improve the scope for its application. However, the same developments make it challenging to maintain standard operating procedures and bioinformatic pipelines. In this protocol, we provide a document assisting the transition from older to current nanopore library preparation kits (Table of Materials).
A common roadblock to sequencing in LMICs is accessibility, including not only the cost but also the ability to procure consumables in a timely manner (in particular sequencing reagents, which are relatively new to procurement teams and suppliers) and computational resources, as well as simply having access to stable power and the internet. Using portable nanopore sequencing technology as the foundation of this workflow helps with many of these accessibility issues, and we have demonstrated the use of our protocol across a range of settings, conducting the full protocol and analysis in-country. Admittedly, procuring equipment and sequencing consumables in a timely manner remains a challenge and, in many instances, we were forced to carry or ship reagents from the UK. However, in some areas, we were able to rely entirely on local supply routes for reagents, benefiting from investment in SARS-CoV-2 sequencing (e.g., the Philippines) that has streamlined procurement processes and begun to normalize the application of pathogen genomics.
The need for a stable internet connection is minimized by one-time-only installs; for instance, GitHub repositories, software download, and nanopore sequencing itself only require internet access to start the run (not throughout) or can be performed completely offline with agreement from the company. If mobile data is available, a phone can be used as a hotspot to the laptop to begin the sequencing run, before disconnecting for the run duration. When routinely processing samples, data storage requirements can grow rapidly, and ideally data would be stored on a server. Otherwise, solid state drive (SSD) hard drives are relatively cheap to source.
While we recognize that there are still barriers to genomic surveillance in LMICs, increasing investment in building genomics accessibility and expertise (e.g., Africa Pathogen Genomics Initiative [Africa PGI])64 suggests that this situation will improve. Genomic surveillance is critical for pandemic preparedness6, and capacity can be established through routinizing the genomic surveillance of endemic pathogens such as RABV. Global disparities in sequencing capacities highlighted during the SARS-CoV-2 pandemic should be a driver of catalytic change to address these structural inequities.
This sample-to-sequence-to-interpretation workflow for RABV, including accessible bioinformatics tools, has the potential to be used to guide control measures targeting the goal of zero human deaths from dog-mediated rabies by 2030, and ultimately for the elimination of RABV variants. Combined with relevant metadata, genomic data generated from this protocol facilitates rapid RABV characterization during outbreak investigations and in the identification of circulating lineages in a country or region60,61,65. We illustrate our pipeline mostly using examples from dog-mediated rabies; however, the workflow is directly applicable to wildlife rabies. This transferability and low cost minimize the challenges in making routine sequencing easily available, not only for rabies but also for other pathogens46,66,67, to improve disease management and control.
The authors have nothing to disclose.
This work was supported by Wellcome [207569/Z/17/Z, 224670/Z/21/Z], Newton funding from the Medical Research Council [MR/R025649/1] and the Philippines Department of Science and Technology (DOST), the UK Research and Innovation Global effort on COVID-19 [MR/V035444/1], the University of Glasgow Institutional Strategic Support Fund [204820], Medical Research Council New Investigator Award (KB) [MR/X002047/1], and International Partnership Development Fund, a DOST British Council-Philippines studentship (CB), a National Institute for Health Research [17/63/82] GemVi scholarship (GJ), and University of Glasgow studentships from the MVLS DTP (KC) [125638-06], the EPSRC DTP (RD) [EP/T517896/1], and the Wellcome IIB DTP (HF) [218518/Z/19/Z]. We are grateful to colleagues and collaborators who have supported this work: Daniel Streicker, Alice Broos, Elizabeth Miranda, DVM, Daria Manalo, DVM, Thumbi Mwangi, Kennedy Lushasi, Charles Kayuki, Jude Karlo Bolivar, Jeromir Bondoc, Esteven Balbin, Ronnel Tongohan, Agatha Ukande, Davis Kuchaka, Mumbua Mutunga, Lwitiko Sikana, and Anna Czupryna.
Brand name | |||
Software | |||
Sequencing software (MinKnow) |
Oxford Nanopore Technologies | https://community.nanoporetech.com/downloads | |
Bioinformatics tool kit (Guppy) |
Oxford Nanopore Technologies | https://community.nanoporetech.com/docs/prepare/library_prep_protocols/Guppy-protocol/v/gpb_2003_v1_revao_14dec2018 | |
Equipment | |||
Thermal cycler (miniPCR™ mini16 thermal cycler) |
Cambio | MP-QP-1016-01 | |
Homogenizer (Precellys Evolution Touch Homogenizer) |
Bertin Instruments | EQ02520-300 | |
Cold Racks (0.2-0.5mL) (PCR Mini-cooler with transparent lid) |
BRAND | 781260 | |
Pipettor | |||
(Pipetman L Fixed F1000L, 1000 uL) | Gilson | SKU: FA10030 | |
(Pipetman L Fixed F100L, 100 uL) | Gilson | SKU: FA10024 | |
(Pipetman L Fixed F10L, 10 uL) | Gilson | SKU: FA10020 | |
(Pipetman L Fixed F1L, 1 uL) | Gilson | SKU: FA10025 | |
(Pipetman L Fixed F20L, 20 uL) | Gilson | SKU: FA10021 | |
(Pipetman L Fixed F250L, 250 uL) | Gilson | SKU: FA10026 | |
Fluorometer (Qubit 4 Fluorometer) |
Thermofisher scientific/Fisher scientific | Q33238 | |
Laptop (Any brand with ~2 GB of drive space, minimum of 512 GB storage space, msi installer [GPU]) |
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Microcentrifuge (Refrigerated centrifuge) |
Thermofisher scientific/Fisher scientific | 75004081 | |
Vortex mixer (Basic vortex mixer) |
Thermofisher scientific/Fisher scientific | 88882011 | |
Magnetic rack (DynaMag -2 Magnet) |
Thermofisher scientific/Fisher scientific | 12321D | |
Sequencing device (MinION) |
Oxford Nanopore Technologies | MinION Mk1B | |
RNA Extraction | |||
RNA extraction kit (Qiagen RNEasy Mini Kit 250) |
Qiagen | 74106 | |
RNA stabilizing reagents | |||
(RNA later) | Invitrogen | AM7020 | |
(DNA/RNA Shield) | Zymo Research | R1100-50 | |
PCR | |||
Nuclease-free Water (Nuclease-free Water [not DEPC-treated]) |
Thermofisher scientific/Fisher scientific | AM9937 | |
Master mix for first strand cDNA synthesis (LunaScript RT SuperMix Kit) |
New England Biolabs | E3010S | |
DNA amplification master mix (Q5® Hot Start High-Fidelity 2X Master Mix [NEB]) |
New England Biolabs | M0494L | |
Primer (Scheme) (Custom DNA oligos) |
Invitrogen | ||
SPRI Bead Clean-up | |||
SPRI beads (Aline Biosciences PCR Clean DX ) |
Cambio | AL-AC1003-50 | |
Ethanol, Pure Absolute, >99.8% (GC) [Riedel-De Haen] | Merck | 818760 | |
Short Fragment buffer (SFB expansion pack) |
Oxford Nanopore Technologies | EXP-SFB001 | |
DNA Quantification | |||
DNA quantification kit (Qubit® dsDNA HS Assay Kit) |
Thermofisher scientific/Fisher scientific | Q32854 | |
DNA quantification assay tubes (Qubit™ Assay Tubes) |
Thermofisher scientific/Fisher scientific | Q32856 | |
End Prep and barcoding (Qubit™ Assay Tubes) |
|||
End Prep master mix (NEBNext Ultra End Repair/dA-Tailing Module) |
New England Biolabs | E7546L | |
Barcoding kit | |||
*Chemistry 9 | Oxford Nanopore Technologies | *Chemistry 9 | |
(Native Barcoding Expansion 1-12) | EXP-NBD104 | ||
(Native Barcoding Expansion 13-24) | EXP-NBD114 | ||
(Native Barcoding Expansion 96) | EXP-NBD196 | ||
*Chemistry 14 | Oxford Nanopore Technologies | *Chemistry 14 | |
(not compatible) | (not compatible) | ||
(Native Barcoding Kit 24 V14) | SQK-NBD114.24 | ||
(Native Barcoding Kit 96 V14) | SQK-NBD114.96 | ||
Ligation mastermix (Blunt/TA Ligase Master Mix) |
New England Biolabs | M0367S | |
Adapter Ligation | |||
Adapter ligation master mix | |||
(NEBNext Quick Ligation Module) | New England Biolabs | E6056S | |
(NEBNext Ultra II Ligation Module) | New England Biolabs | E7595S | |
(Blunt/TA Ligase Master Mix) | New England Biolabs | M0367S | |
Adapter mix | |||
*Chemistry 9 | Oxford Nanopore Technologies | *Chemistry 9 EXP-AMII001 |
|
(Adapter Mix II [AMII]) | |||
*Chemistry 14 | Oxford Nanopore Technologies | *Chemistry 14 EXP-NBA114 |
|
(Native adapter [NA]) | |||
Sequencing | |||
Flowcell priming kit | |||
*Chemistry 9 | Oxford Nanopore Technologies | *Chemistry 9 EXP-FLP002 |
|
(Flush Buffer [FB]) | |||
(Flush Tether [FT]) | |||
*Chemistry 14 | Oxford Nanopore Technologies | *Chemistry 14 EXP-FLP004 |
|
(Flow Cell Flush [FCF]) | |||
(Flow Cell Tether [FCT]) | |||
Ligation Sequencing Kit | |||
*Chemistry 9 | Oxford Nanopore Technologies | *Chemistry 9 SQK-LSK109 |
|
Adapter Mix (Adapter Mix [AMX]) |
|||
Ligation Buffer (Ligation buffer [LNB]) |
|||
Short Fragment Buffer (Short Fragment buffer [SFB]) |
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Sequencing Buffer (Sequencing Buffer [SQB]) |
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Elution Buffer (Elution buffer [EB]) |
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Loading Beads (Loading Beads [LB]) |
|||
Sequencing Tether (Sequencing Tether [SQT]) |
|||
*Chemistry 14 | Oxford Nanopore Technologies | *Chemistry 14 SQK-LSK114 |
|
Adapter Mix (Ligation Adapter [LA]) |
|||
Ligation Buffer (Ligation buffer [LNB]) |
|||
Short Fragment Buffer (Short Fragment buffer [SFB]) |
|||
Sequencing Buffer (Sequencing Buffer [SB]) |
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Elution Buffer (Elution buffer [EB]) |
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Loading Beads (Loading Beads [LIB]) |
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Sequencing Tether (Flow Cell Tether [FCT]) |
|||
Library solution (Library solution [LIS]) |
|||
Flush buffer (Flow Cell Flush [FCF]) |
|||
Flow Cell | |||
*Chemistry 9 | Oxford Nanopore Technologies | *Chemistry 9 FLO-MIN106D |
|
(Flow Cell [R9.4.1]) | |||
*Chemistry 14 | Oxford Nanopore Technologies | *Chemistry 14 FLO-MIN114 |
|
(Flow Cell [R10.4.1]) | |||
Flow Cell wash | |||
Flowcell wash kit (Flow cell wash kit) |
Oxford Nanopore Technologies | EXP-WSH004 | |
Consummables | |||
Surface decontaminant | |||
(DNA Away Surface Decontaminant, Squeeze Bottle [Molecular Bio]) | Thermofisher scientific/Fisher scientific | 7010PK | |
(RNase Away Surface Decontaminant, Bottle [Molecular Bio]) | Thermofisher scientific/Fisher scientific | 7002PK | |
PCR 8-Tube Strip 0.2ml, individual cap (PCR 8-Tube Strip 0.2ml, with Individual attached Flat Caps, Sterile, DNAse/RNAse, Pyrogen Free,Natural [Greiner]) |
Greiner | 608281 | |
PCR Tube 0.2ml (PCR Tube 0.2ml, Natural [Domed Cap] Bagged in 500s, Non-Sterile [Greiner]) |
Greiner | 671201 | |
1000µL Filter Tips (500) (Stacked 1000µL Filter Tips [500]) |
Thermofisher scientific/Fisher scientific | 11977724 | |
100µL Filter Tips (1000) | Thermofisher scientific/Fisher scientific | 11947724 | |
10µL Filter Tips (1000) (Stacked 100µL Filter Tips [1000]) |
Thermofisher scientific/Fisher scientific | 11907724 | |
Reinforced tubes tubes (2ml) with screw caps and o-rings (Fisherbrand™ Bulk tubes) |
Thermofisher scientific/Fisher scientific | 15545809 | |
Microcentrifuge tube (1.5ml) (1.5 ml Eppendorf Tubes [500]) |
Eppendorf | 1229888 | |
DNA LoBind Tubes (1.5ml) (DNA LoBind Tubes) |
Thermofisher scientific/Fisher scientific | 10051232 | |
Cryobabies labels | |||
Gloves (S/M/L) | |||
Paper towel |