Keywords
cassava, uganda, kenya, tanzania, nanopore, minion, SDG2
This article is included in the Nanopore Analysis gateway.
This article is included in the Agriculture, Food and Nutrition gateway.
cassava, uganda, kenya, tanzania, nanopore, minion, SDG2
The United Nations has listed Zero Hunger as one of the 17 global sustainable development goals to end extreme poverty by 2030. Plant viruses are a major constraint to crop production globally, causing an estimated $30 billion in damage1 and leaving millions of people food-insecure2. In Africa, agriculture employs up to 50% of the workforce, yet only contributes 15% to the GDP on average3, suggesting that there is low productivity and limited value addition. This can be addressed through continued innovation in the fields of science and technology, as suggested in the Science Agenda for Agriculture in Africa (S3A)4. Sustainable management of plant viruses and their associated vectors must include efficient diagnostics for surveillance, detection and identification to inform disease management, including the development and strategic deployment of virus resistant varieties. To date, researchers have been utilizing conventional methods such as; PCR, qPCR, high-throughput sequencing (RNA-Seq, DNA-Seq) and Sanger sequencing for pathogen identification. However, these methods are both costly and time consuming, delaying timely control actions. The emergence of new tools for real-time diagnostics, such as the Oxford Nanopore MinION, have recently proven useful for the early detection of Ebola5 and Zika6,7, even in poorly resourced laboratories. For the first time globally, the MinION portable pocket DNA sequencer was used to sequence whole plant virus genomes. We used this technology to identify the begomoviruses causing the devastating cassava mosaic virus, which is ravaging smallholder farmers’ crops in sub-Saharan Africa. Cassava, a carbohydrate crop from which tapioca originates, is a major source of calories for over 800 million people worldwide. With this technology, farmers struggling with diseased crops can take immediate, restorative action to improve their livelihoods based on information about the health of their plants, generated using a portable, real-time DNA sequencing device.
Portable DNA sequencing technology has great potential to reduce the risk of community crop failure and help improve livelihoods of millions of people, especially in low resourced communities. Plant diseases are a major cause of low crop productivity and viruses such as tobacco mosaic virus, tomato mosaic, tomato spotted wilt, potato leaf roll, Potato virus X and Y in potato, papaya mosaic, citrus tristeza, chilli leaf curl, and banana bunchy top have been implicated. In particular, cassava viruses are among the world’s greatest risk to food insecurity. Losses caused by cassava mosaic disease (CMD) and cassava brown streak disease are estimated at US $2-3 billion annually8.
We visited smallholder farmers in Tanzania, Uganda and Kenya (Table 1) who are suffering yield shortages due to cassava virus infections. We utilized the MinION to test infected material and farmers were informed within 48 hours of the specific strain of the virus that was infecting their cassava, and a resistant cassava variety was deployed. The advantages of adopting this technology far outweigh the challenges (Table 2). Cassava mosaic begomoviruses were in high enough concentration that reads of whole genomes were obtained without an enrichment step (Table 1). As expected, the viral reads increased with the severity of the symptoms observed (Table 1). We detected a dual infection for a leaf sample with the severity score of 5 in Uganda. In addition, one asymptomatic plant in Tanzania had one viral read detected. The shortest time to obtain a viral read was 15 s (severity score 5) and the longest was 4 h 11 m 15 s (severity score 1).
MinION sequencing is superior to traditional methods of PCR identification, given its generation of whole genome sequences which enable the identification of the plant virus strain even if it becomes mutated or divergent, as it is not biased using primers that rely on known virus sequences. With regards to cassava, there are three major advantages of this technology. Firstly, improved diagnostics are required and real-time whole-genome sequencing will help develop diagnostic primers that are up-to-date. Secondly, this technology will assist with the development of resistant cassava varieties and will allow breeders to immediately test the varieties they are developing against different viral strains. Lastly, it ensures the delivery of the correct healthy uninfected planting material to farmers. In addition, we could detect virus in a plant before it showed symptoms (Table 1). Utilizing traditional PCR methods, three samples collected from farmer 1’s field in Tanzania tested positive for EACMVs and none were positive for ACMV. The asymptomatic sample from Mikocheni Agricultural Research Institute (MARI) tested negative for both ACMV and EACMVs. There were eight fresh cassava leaf samples from Uganda found to be dually infected with ACMV and EACMV-UG using conventional PCR primers for ACMV and EACMV-UG. The primers used in this PCR yield products of 1000 bp and 1500 bp for ACMV and EACMV-UG, respectively. In total, 12 Kenyan samples were tested and all but two (barcode 2 and 3) were found to be positive using conventional PCR (Table 1). Further studies are needed to verify our results regarding the sensitivity of the protocol for early detection of CMD in cassava, but these results are very promising for ensuring farmers receive clean planting material through the early detection of viral infection.
Nanopore sequencing technology has wide applications globally, but in East Africa these include: (a) crop improvement by screening for virus resistant germplasm and genetic diversity during breeding; (b) indexing of cassava planting materials for virus presence or absence to ensure that only clean materials in multiplication fields are distributed to farmers; (c) detection and identification of alternative plant species for cassava-infecting begomoviruses, so that farmers are advised to remove and/or grow crops away from such plants as a management strategy; and (d) virus and biodiversity studies.
In Tanzania, three cassava mosaic disease (CMD) symptomatic cassava leaf samples (Figure 1, Table 1) were collected from the smallholder cassava farmer 1’s field in Bagamoyo. Disease severity was assessed as described by Legg et al.2, where 1 is healthy and 5 shows severe symptoms of the disease, including leaf distortion and stunting of the plant. One more asymptomatic leaf sample was collected at MARI, Dar es Salaam. In total, seven CMD symptomatic plants were collected from farmer 2’s farm in Wakiso district in Uganda. Both Tanzanian and Ugandan samples were collected in September 2017. A total of 12 samples from Kenya were collected in February 2018 from various sources (Table 1). High-quality DNA was isolated using the cetyl trimethylammonium bromide method9. Each DNA sample (Table 1) was quantified and the purity checked using a NanoDrop 2000c UV–vis Spectrophotometer (Thermo Fisher Scientific, Wilmington, DE, USA) was used to check the purity and quantity of DNA for each sample, and results were recorded in Table 1.
In Tanzania and Uganda, the Rapid Barcoding kit SQK-RBK001 and MinION 9.4.1 flow cells (Oxford Nanopore) were used to process genomic DNA extracted using a standard CTAB method9. We utilized the Rapid Barcoding kit SQK-RBK004 with 9.4.1 flow cells in Kenya. DNA was diluted to 700 ng as specified in the library protocol. The SQK-RBK001 (Sept 2017) and/or the SQK-RBK004 (Feb 2018) protocols were performed as described by the manufacturer. In Tanzania and Uganda, the MinION was run for 24 h instead of the recommended 48 h, and in Kenya we had a total run time of approximately 17 h due to power interruptions.
In Tanzania and Uganda, Albacore 2.0.2 was used for base calling. In Kenya, Albacore 2.1.10 was used and the scripts were modified to reflect the newest rapid barcoding kit RBK004. Fastq files were imported into Geneious10 and a local blast database of all known cassava mosaic begomovirus whole genomes were downloaded from GenBank (program set to megablast, maximum hits=1, scoring (match mismatch 1-2, max e-value 1e-1, word size 28, gap cost=linear) and a local blast was performed on each of the reads generated using the Nanopore device. There are other bioinformatics pipelines that are used for Nanopore data, but the focus of our study was to use a simple, efficient analyses pipeline, which was a local blast database and a well-curated reference dataset11. To ensure we did not miss other begomoviruses or pathogens, we confirmed these in-field results by performing a post diagnostic blast of reads on the Nimbus Cloud at the Pawsey Supercomputing Center (Kensington, Australia) with blast 2.2.31 against the full NCBI nucleotide database to confirm results. For reference, the specific database used was {$ blastcmd –db nt/nt –info} Database: Nucleotide collection (nt) 46,853,753 sequences; 170,830,796,758 total bases Date: Feb 20, 2018 5:04 PM. The data were processed into a blast archive using a blast script with the following parameters (Script attached) {$blastn -query "$file" -db /mnt/nucdb/nt/nt -outfmt 11 -culling_limit 10 -out "out.$file.asn" -num_threads 17 } then converted into XML (for loading into Geneious) and HTML for viewing where possible.
Traditional PCR was used to verify our DNA sequencing results. In Tanzania and Kenya, two primer pairs: EAB 555F/EAB 555F12 and JSP001/JSP00213, which amplify 556 bp and 774 bp, respectively, were used to detect East African CMVs (EACMVs) and African CMVs (ACMVs), respectively. Specifically, high quality DNA were isolated using CTAB method as described by Lodhi et al.9. Electrophoresis and spectrophotometric measurements were used to check the quality and quantity of DNA. PCR reactions were carried out in a 25 µl volume, with 12.5 µl (1X final concentration) of One Taq PCR 2X Master Mix (New England Biolabs), One Taq DNA Polymerase in an optimized buffer with 1.5 mM Mg2+ and 0.2 mM each dNTPs in 1X final concentration. A total of 0.2 µM (0.5 µl of each) of primer sets were used in the reaction and nuclease-free water were added to 25µl final reaction volume. The template used was less than 500 ng per reaction. PCR was carried out in 2720 Thermal Cycler (Applied Biosystems) with the following programme: 1 cycle of 3 min at 94°C, then 30 cycles at 94°C for 45 s, 56°C for 45 s, 72°C for 1 min, and a final cycle at 72°C for 7 min. Electrophoresis of the PCR product was run on 1% agarose gel stained in ethidium bromide in 1X TAE buffer in a submarine gel electrophoresis unit and visualized using a BioDoc-It 210 Imaging System m-20V Transilluminator (Thomas Scientific).
In Uganda, the presence of ACMV and EACMV in each sample was detected using a pair of specific primers for ACMV, ACMV-AL1/F and ACMV-ARO/R, and primers specific for EACMV-UG2, UV-AL1/F and ACMV-CP/R314. Specifically, the presence of ACMV and EACMV in each sample was detected using a pair of specific primers for ACMV, ACMV-AL1/F (5’-GCC GGA ATC CCT AAC ATT ATC -3’) and ACMV-ARO/R (5’-GCT CGT ATG TAT CCT CTA AGG CCT G -3’) and specific for EACMV-UG2, UV-AL1/F (5’-TGT CTT CTG GGA CTT GTG TG -3’) and ACMV-CP/R3 (5’- TGC CTC CTG ATG ATT ATA TGT C-3’) described by Zhou et al.14. The primers amplify about 1000 bp for ACMV and 1500 bp for EACMV-UG2 of the Coat Protein and AV2 gene sequences of the ACMV and EACMV genomes, respectively. The PCR reaction was set up using GoTaq® Green Master Mix (Promega, Madison USA). Each of the 25 µl PCR mix contained 12.0 µl of 2X GoTaq® Green Master Mix [containing GoTaq® DNA polymerase, 2X Green GoTaq® Reaction Buffer (pH 8.5), 400 µM dATP, 400 µM dGTP, 400 µM dCTP, 400 µM dTTP and 3 mM MgCl2], 1.0 µl of forward primer, 1.0 µl of reverse primer, 10.0 µl of nuclease-free water (Amressco, Ohio, USA) and 1.0 µl of DNA. PCR was performed using a Biometra professional thermocycler (Biometra, Gottingen, Germany) programmed as follows: 94 °C for 2 min for initial denaturation followed by 30 cycles of 94 °C for 1 min, 60 °C for 1.5 min, 72°C for 2 min and 72°C for 10 min for denaturation, annealing, extension and final extension, respectively. PCR amplicons were separated by electrophoresis in a 1× Tris-acetate-EDTA (TAE) buffer in a 1.2% agarose gel, stained with ethidium bromide (0.1 mg/ml) and visualized using a U:Genius3 (Syngene, Cambridge, UK) gel documentation system.
The image in Figure 1 was captured in Kiromo-Kitonga Bagamoyo, Tanzania by J.N. using a Samsung s8 smartphone.
An earlier version of this article can be found on bioRxiv (DOI: https://doi.org/10.1101/314526).
Raw data for this study are available on figshare: DOI: https://doi.org/10.6084/m9.figshare.666740911.
This work was part-funded by the Crawford Fund, Australia (grant number WA-803-2017).
The funders had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript.
Funding for the Kenyan trip was provided by the Crawford Fund. We also thank the participants from the University of Eldoret who assisted in the preparation of libraries for the Kenyan samples.
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Competing Interests: No competing interests were disclosed.
Reviewer Expertise: My expertise areas are the microbial genomics, nanopore DNA sequencing technology, molecular biology, and the massive DNA data analysis.
Competing Interests: No competing interests were disclosed.
Reviewer Expertise: I'm a Research Microbiologist for the US Food and Drug Administration. My research focuses on the validation of phylogenetic tools for molecular surveillance. I also manage a genome surveillance network for foodborne pathogens called the GenomeTrakr, that collects and publishes WGS data from food and environmental isolates.
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