1. Introduction
Bovine respiratory disease (BRD) is the leading cause of morbidity, mortality and economic loss in cattle of all ages [
1,
2,
3,
4,
5]. The extensive use of vaccines against BRD-associated viral and bacterial pathogens has not reduced the incidence or severity of BRD in cattle. Consequently, large quantities of broad-spectrum antimicrobials are used for the therapeutic treatment of BRD in Europe and the USA [
6,
7]. Easy and economical methodologies that enable the rapid and reliable on-farm detection of viral and bacterial pathogens are required to inform rapid targeted treatment and up-to-date vaccine design.
BRD is a multifactorial disease that is associated with an ever-increasing number of species and strains of viruses and bacteria. Viruses that are commonly associated with BRD cases include bovine herpesvirus type 1 (BoHV-1), bovine parainfluenza 3 virus (BPI-3), bovine respiratory syncytial virus (BRSV), bovine viral diarrhoea virus (BVDV), bovine coronavirus (BCoV) and bovine adenovirus (BAdV) [
8,
9]. BRD-associated bacteria commonly include
Pasteurella multocida,
Mannheimia haemolytica,
Histophilus somni and
Mycoplasma spp. [
8,
9,
10].
New pathogens, e.g., influenza D virus (IDV) [
11] and
Sneathia amnii [
12], are continually added to the list of BRD aetiologies. Viruses are known to initiate the disease by weakening the animal’s defenses, which leads to secondary bacterial infection [
13]. Viruses with RNA and DNA genomes have been associated with BRD [
8].
A diagnosis of BRD is generally based on clinical signs that are assessed via visual inspection of the animal, measurement of rectal temperature and pulmonary auscultation [
2,
3,
14,
15]. If the identification of the causative pathogen(s) is attempted (which is often not the case), a nasal swab from the affected animal is sent to a diagnostic laboratory, where a targeted qPCR diagnostic analysis of only four or five of the most likely (e.g., previously mentioned) bacterial or viral pathogens is conducted [
16,
17]. As there are at least 40 possible bacterial and viral pathogens associated with BRD, it is too expensive and time-consuming to test for all of these using targeted real-time quantitative PCR (qPCR) diagnostics. Consequently, there are considerable delays in receiving the results of the aetiological diagnoses of BRD cases, which are often inconclusive [
18]. For pathogen identification to be of direct practical use in preventing a BRD outbreak, the results would need to be available to a veterinarian within 24 h. As such, qPCR falls far short of what is required for rapid BRD-associated pathogen identification. This has prompted the increased application of viral metagenomic approaches that are based on next-generation sequencing (NGS) (e.g., Illumina platform) [
8,
9] and third-generation sequencing (TGS) (e.g., Oxford Nanopore Technologies platform) [
19] to BRD-associated virus diagnostics. However, the relatively high cost and lack of portability of even the smallest Illumina platform NGS machine (the iSeq 100) restrict its availability and use to mainly large, well-funded laboratories. In addition, the sequencing runs on Illumina NGS machines usually take longer than 24 h and sequences can only be viewed and analysed after the run is complete.
In contrast, the Oxford Nanopore Technologies (ONT) MinION DNA/RNA sequencer is a portable, low-cost sequencing device that allows real-time data analyses [
20] and has the potential to enable untargeted same-day (sample-to-result) viral diagnostics (i.e., identify viruses to species/strain/sequence-variant level and generate a viral sequence for the assembly of new viral genomes) in small static or mobile veterinary surgeries. Sequencing all of the nucleic acid in a sample potentially allows for the detection, in a single assay, of all organisms, including pathogens, that are present in that sample [
20]. On the ONT MinION, DNA or RNA is sequenced on a disposable flowcell that contains a membrane that is covered in active biological nanopores. There are three types of flowcell that can currently be used on a MinION. These are called (1) FLO-MIN106D (R9.4.1), (2) FLO-FLG001 (R9.4.1) and (3) FLO-MIN112 (R10.4). R9.4.1 and R10.4 refer to the types of biological pores in the membrane in each type of flowcell (
https://store.nanoporetech.com/eu/flow-cells.html (accessed on 21 April 2020)). The membrane is flanked on either side by opposing electrical charges, which drive negatively charged individual single-chain strands of DNA or RNA through the nanopores towards the positive charge. A sensor registers the unique change in the current produced by bases as they pass through the nanopore. These changes in current are translated into nucleotide sequence information, in the form of FASTQ files, by neural network basecallers [
21,
22]. As soon as these FASTQ files are generated, they can be uploaded to a cloud-based software platform called Epi2ME, which contains several intuitive point-and-click sequence analysis applications called ‘workflows’.
The objective of the present study was to assess whether the ONT MinION device, Epi2ME cloud-based software and library preparation kits could achieve sufficient sensitivity, accuracy, specificity and speed to correctly detect and generate the genomic sequence of a known BRD virus in nasal swabs, which are the most commonly collected sample in BRD outbreaks, from infected cattle within 24 h. For this, we first tested and optimised ONT procedures using bovine foetal lung cell cultures (bFLC) that were infected with the BRD-associated virus bovine herpes virus-1 (BoHV-1), as, unlike nasal swabs from BoHV-1-infected cattle, the supply of cell cultures was not a limiting factor. BoHV-1 is an enveloped DNA virus with a large genome of 135.3 kb with a high GC content of 72% and is of major economic importance to the cattle industry [
23]. We then applied these procedures to nasal swabs that were collected from Holstein–Friesian calves that had been experimentally challenged with BoHV-1.
2. Materials and Methods
2.1. Bovine Foetal Lung Cells Infected with BoHV-1
Bovine foetal lung cells (bFLCs) were isolated from a bovine foetus. The source, origin and characteristics of these bFLCs are shown in
Table S1. BoHV-1 strain 2011-415 is a virulent field sample that was isolated from a typical fatal clinical case (9-month-old calf) of infectious bovine rhinotracheitis with pulmonary complications. A post-mortem trachea sample from this animal tested positive via qPCR for BoHV-1 and the virus was isolated and stored at −80 °C. A T75 flask (Thermo Fisher Scientific, Agawam, MA, USA) containing bFLC was infected with the BoHV-1 strain 2011-415 isolate with a multiplicity of infection (MOI) of 1. As a negative control, an additional T75 flask containing bFLCs and 5 mL of 2% buffer G-MEM was also used. The two flasks were incubated at 37 °C for 90 min in a CO
2 incubator (Thermo Fisher Scientific, Carlsbad, CA, USA); then, 15 mL of 2% G-MEM buffer was added. The flasks were replaced in the CO
2 incubator at 37 °C. At 48 h post-infection, the two flasks were observed via phase-contrast light microscopy at 40× magnification, with 90% of the cells displaying viral cytopathology. The flasks were then placed in a freezer at −80 °C for 2 h, and subsequently thawed and their contents transferred to sterile 50 mL centrifuge tubes (Thermo Fisher Scientific, Agawam, MA, USA). The tubes were then centrifuged for 5 min at 3660×
g, and 1 mL aliquots of the supernatant were transferred to sterile 1.5 mL microfuge tubes (Eppendorf, Hamburg, Germany) and frozen at −80 °C.
2.2. Experimental Calves
All animal experiments were carried out in accordance with the UK Animals (Scientific Procedures) Act 1986 and with the approval of the Agri-Food and Biosciences Institute Northern Ireland Ethical Review Committee. The study was reported in accordance with ARRIVE guidelines [
24].
As part of a larger study, 12 Holstein–Friesian bull male calves (mean age 21.3 weeks, s.d. ± 3.4) were selected from a larger group of 43 Holstein–Friesian bull calves. The enrolment of calves for the challenge study was based on two criteria: (i) a low level of BoHV-1 antibody and (ii) a negative BoHV-1 qPCR result for a nasal swab collected two weeks before the challenge. The 12 selected calves were assigned to two groups (BoHV-1-challenge and PBS-challenge), with 6 calves per group. For the PBS-challenge group (n = 6), the mean age = 21.4 (s.d. ± 3.3) weeks, mean weight = 173.3 (s.d. ± 23.7) kg and mean BoHV-1 antibody = 18.0 (s.d. ± 4.5%). For the BoHV-1-challenge group (n = 6), the mean age = 21.0 (s.d. ± 4.5) weeks, mean weight = 175.8 (s.d. ± 35.6) kg and BoHV-1 antibody = 20.6 (s.d. ± 13.2%). On day 0, each calf in the BoHV-1-challenge group was experimentally challenged via intranasal atomisation with a 1.35 mL solution of BoHV-1culture. The animals in the PBS-challenge group were mock-challenged (day 0) with intranasal atomisation of 1.35 mL of a sterile PBS solution. The two groups were housed in two separate biocontainment level 3 sheds, each with a 10 m × 5 m floor covered in straw. Daily clinical assessments, nasal swabs, and blood samples were collected from each animal on day −1, day 0, day 1, day 2, day 3, day 4, day 5 and day 6 relative to the day of the challenge. Animals were euthanised on day 6 post-challenge.
2.3. Nasal Swabs from Experimental Calves
For each nasal swab sample, the exterior of the nasal nares of the calf was sterilised with 70% ethanol; then, a sterile swab was removed from its sterile tube and inserted approximately 20 cm into the nostril and rolled on the internal nasal membrane for approximately 5 s. The swab tip was cut with scissors (sterilised with 70% ethanol) into a 2 mL sterile tube and immediately frozen on dry ice. This was repeated once for each animal so that two nasal swabs (nasal swab-1 and nasal swab-2) were taken for each animal on every day of the 8 day trial. Nasal swabs from the BoHV-1 calf challenge model were stored for approximately 6 months at −80 °C prior to nucleic acid extraction qPCR and sequencing.
Immediately prior to nucleic acid extraction, nasal swabs in tubes were removed from storage in a −80 °C freezer to a class 2 biological safety cabinet. A volume of 1.5 mL of molecular grade PBS (Sigma Aldrich, St. Louis, MO, USA) was added to each tube and the tubes were vortexed for 1 min to release the nasal material from the swabs. The resulting nasal swab eluate was then transferred to a sterile 15 mL tube and the swab remained in the 2 mL tube. A further 1.5 mL of PBS was added to each of the swabs in the 2 mL tubes, the tubes were vortexed again, and this second PBS nasal swab eluate was removed and added to the first 1.5 mL PBS nasal swab eluate in the 15 mL tube, resulting in 3 mL of nasal swab eluate for each swab sample.
2.4. Non-Viral Nucleic Acid Depletion
For the bead beating step, either 250 µL of BoHV-1 infected bFLC in vitro culture or 1 mL of nasal swab-2 eluate was transferred to a Pathogen Lysis Tubes L (Qiagen, Manchester, UK). For the negative extraction control, 250 µL of molecular grade phosphate-buffered saline (PBS, pH 7.4) (Sigma Aldrich, St. Louis, MO, USA) was added to a Pathogen Lysis Tube L (Qiagen, Manchester, UK). To prevent the escape of aerosols from the tubes during bead beating, tube lids were sealed with Parafilm (Sigma Aldrich, St. Louis, MO, USA). Tubes were placed in a FastPrep-24 disruptor instrument (MP Biomedicals, Irvine, CA, USA) and shaken at high speed (4 ms
−1) for 30 s. The tubes were then removed and centrifuged at 500×
g for 45 s to collect the contents at the bottom of the tube. The supernatant from each Pathogen Lysis Tube was then carefully transferred to 2 mL DNA LoBind Safelock tubes (Eppendorf AG, Hamburg, Germany) and the volume was adjusted to 1 mL with molecular grade PBS. An aliquot of 2.5 µL of RNaseA (4 mg/mL) (Promega, Madison, WI, USA) was added to the tubes, which were then incubated for 15 min at 37 °C in an Eppendorf Thermostat Plus (Eppendorf AG, Hamburg, Germany). Turbo DNase (10 μL) and 10× Turbo DNase buffer (100 μL) were then added (Thermo Fisher Scientific, Carlsbad, CA, USA) [
25] and the tubes were gently mixed by pipetting six times and incubated for 30 min at 37 °C in an Eppendorf Thermostat Plus. A further 10 μL of Turbo DNase was added and the contents were again mixed by gently pipetting six times and incubated for a further 30 min at 37 °C. DNase inactivation reagent (112.5 μL) (Thermo Fisher Scientific, Carlsbad, CA, USA) was then added to the samples and mixed by gentle pipetting six times, and incubated for 5 min at 24 °C. The mixture was centrifuged at 10,000×
g for 90 s; then, the supernatant containing the nucleic acid was transferred to a fresh tube without disturbing the pellet of DNase Inactivation Reagent. Samples were also processed with the omission of certain treatments to test whether these treatments were necessary. Five such treatment regimes were tested. These were: (A) no bead beating or nuclease treatment; (B) bead beating only; (C) no bead beating, 1× RNase and 1× DNase; (D) no bead beating, 1× RNase and 2× DNase; and (E) bead beating, 1× RNase and 2× DNase. A negative extraction control (a tube containing only molecular grade PBS, or a sterile unused swab) was included with each batch of extractions to monitor the contamination of reagents and cross contamination of samples during the extraction process.
2.5. Nucleic Acid Extraction and Purification
For nasal swab-1 eluates, nucleic acid was extracted with a Roche Magnapure using a Roche Total Nucleic Acid Isolation Kit (Roche, Basel, Switzerland). Nucleic acid extraction and purification from nasal swab-2 eluates and bFLCs were performed using the QIAamp UltraSens Virus Kit (Qiagen, Manchester, UK) according to the manufacturer’s instructions [
26] with the exception of the substitution of 5.6 µL of carrier RNA with 5.6 µL of a solution of 5 mg/mL linear acrylamide. Total nucleic acid was immediately extracted and purified from the swab eluates and bFLCs following the nuclease treatment.
2.6. qPCR Analysis
qPCR analysis was performed in triplicate and the mean of the three resulting Cq values was used if the standard deviation was within the default set limits of ABI7500 software (Applied Biosystems, Waltham, MA, USA). The BoHV-1 UL27 gene TaqMan® Custom Gene Expression Assay (Applied Biosystems, Waltham, MA, USA) comprised BoHV-1 F (forward primer) 5′-TGT GGA CCT AAA CCT CAC GGT-3′, BoHV-1 R (reverse primer) 5′-GTA GTC GAG CAG ACC CGT GTC-3′ and a BoHV-1 probe (FAM-MGB) 5′-AGG ACC GCG AGT TCT TGC CGC-3′. The bACTB TaqMan® Custom Gene Expression assay (Applied Biosystems) comprised bACTB F (forward primer) 5′-CCC TGG AGA AGA GCT ACG AG-3′, bACTB R (reverse primer) 5′-CAG GAA GGA AGG CTG GAA GA-3′ and a bACTB probe (FAM-MGB) 5′-CGG TTC CGC TGC CCT GAG GC-3′. For each qPCR reaction, 12.5 µL of 2X RT-PCR buffer (Thermo Fisher Scientific, Carlsbad, CA, USA), 2.5 µL of the TaqMan Custom Gene Expression Assay (Applied Biosystems, Waltham, MA, USA), 1 µL of 25X RT-PCR Enzyme Mix (Thermo Fisher Scientific, Carlsbad, CA, USA) and 1 µL of the sample were combined for a total reaction volume of 25 µL per well of a 96-well qPCR plate. qPCR reactions were run on a 96-well plate on an ABI7500 FAST machine (Applied Biosystems, Waltham, MA, USA) using the following PCR cycling conditions: 50 °C (2 min), 95 °C (10 min), and then 40 cycles of 95 °C (15 s) and 60 °C (1 min). Cq values were converted to relative quantities using 2−(UL27 Cq-bACTB Cq). Cq values were converted to the number of BoHV-1 genome copies per µL and the number of Bos taurus genome copies per µL using two standard curves (qPCR Cq value vs. DNA concentration) generated from a dilution series of extracted Bos taurus genomic DNA and a PCR2.1 TOPO plasmid (Thermo Fisher Scientific, Carlsbad, CA, USA) clone of the BoHV-1 UL27 gene that we prepared (according to the manufacturer’s instructions) and quantified using both spectrophotometry (on a Nanodrop 1000) and DNA specific dye fluorescence (on a Qubit fluorometer). The mean of the Nanodrop DNA concentration value and Qubit DNA concentration value was used for the generation of the standard curves.
2.7. Generation of MinION Nanopore Libraries for Multiplex Rapid Sequencing
MinION nanopore sequencing libraries were generated from nucleic acid extracted from bFLCs and nasal swab-2 eluates with the Rapid PCR Barcoding Kit (SQK-RPB004) [
27] (Oxford Nanopore Technologies, Oxford, UK). For each library, 6 µL of nucleic acid extraction plus 2 µL of fragmentation mix (FRM) were added to a thin-walled 0.5 mL PCR tube (Eppendorf, Hamburg, Germany). The tubes were incubated in a Master Cycler Gradient PCR machine (Eppendorf, Hamburg, Germany) at 30 °C for 1 min and then 80 °C for 1 min, after which they were immediately placed on ice. This resulted in tagmentation of the DNA in the nucleic acid extraction with sequencing adapters.
For PCR amplification of the tagmented DNA, 8 µL of the tagmentation reaction, 16 µL of nuclease-free water, 1 µL of rapid barcode primer (RLB), and either 25 µL of NEB LongAmp Taq 2× Master Mix (New England BioLabs Inc., Ipswich, MA, USA) or 25 µL of NEB Next Ultra II Q5 Master Mix (New England BioLabs Inc.) were added to a 0.5 mL thin-walled PCR tube (Eppendorf, Hamburg, Germany). The tube contents were mixed by gently flicking the tube and then centrifuged for 10 s in a minifuge. The tubes were then placed in a Master Cycler Gradient PCR machine (Eppendorf AG, Hamburg, Germany). Cycle conditions for LongAmp Taq were: 95 °C (3 min); then 10, 20 or 30 cycles of 95 °C (15 s), 56 °C (15 s) and 65 °C (6 min); and then one cycle of 65 °C (6 min) followed by a hold step at 4 °C. Cycle conditions for NEB Next Ultra™ II Q5 were 98 °C (3 min); then 10, 20 or 30 cycles of 98 °C (10 s), 65 °C (30 s) and 72 °C (either 40 s, 120 s, 180 s or 300 s); and then one cycle of 72 °C (2 min) followed by a hold step at 4 °C.
Following PCR, the 12 PCR libraries were combined in a single 1.5 mL Eppendorf DNA Lo-Bind (Eppendorf AG, Hamburg, Germany) tube and a total volume of 360 µL of AMPureXP beads (i.e., 30 µL of beads for each barcoded library) (Beckman Coulter Inc., Brea, CA, USA) was added to the 12 pooled PCR reactions and mixed gently via pipetting. The library pool and beads were incubated in a rotator mixer for 5 min at room temperature. The tube was then removed from the rotator mixer, centrifuged for approximately 20 s in a minifuge, placed on a magnet for 5 min and the supernatant was discarded. Being careful not to dislodge the beads, 1 mL of 70% ethanol was added to the beads, then immediately removed. This ethanol wash step was then repeated once. The tube was then centrifuged for 30 s using a minifuge and placed on the magnet until the beads had bound to the side of the tube, leaving the ethanol at the bottom. Residual ethanol was removed from the bottom of the tube via pipetting without touching the beads. The tube was then left open for 60 s to allow the pellet to air dry. The tube was then removed from the magnet and the pellet was resuspended in 25 µL of a solution of 10 mM Tris-HCl (pH 8.0) and 50 mM NaCl. The solution was mixed by pipetting until the pellet was completely resuspended and the suspension was then incubated for 2 min at room temperature to elute the purified–pooled–barcoded libraries. The tube was placed on the magnet for 5 min and the eluate containing the purified–pooled–barcoded libraries was collected and transferred to a new 1.5 mL DNA LoBind tube (Eppendorf AG, Hamburg, Germany). A volume of 1 µL of the purified–pooled–barcoded libraries was removed to measure its DNA concentration on a Qubit fluorometer using the Qubit 1× dsDNA HS Assay Kit (Thermo Fisher Scientific, Carlsbad, CA, USA), following the manufacturer’s instructions. The volume of the library corresponding to 60 ng dsDNA was calculated and removed to a new 1.5 mL DNA LoBind tube. This volume was then adjusted to 10 µL with 10 mM Tris-HCl (pH 8.0) and 50 mM NaCl. One microliter of rapid adapter (RAP) was then added to the 10 µL of the pooled barcoded library, mixed gently by flicking the tube, centrifuged for approximately 10 s in a minifuge to collect the liquid at the bottom of the tube and incubated at room temperature for 5 min. Subsequently, the sample and flowcell were prepared for priming and loading following the manufacturer’s instructions. The library was loaded into the spot-on port on a spot-on flowcell (FLO-MIN106D R9) (Oxford Nanopore Technologies, Oxford, UK) and sequenced for 48 h on a MinION Mk1b sequencer (Oxford Nanopore Technologies, Oxford, UK) attached to a MinIT compute module (Oxford Nanopore Technologies, Oxford, UK) using MinIT software version 19.05.02 (with Guppy 3.0.3) and the rapid basecalling option. Each flowcell was tested with the ONT configuration program in MinKnow software immediately prior to each run to ensure that the number of active pores was >1000. The output selected for each run was fast basecalling, FASTQ files only.
2.8. Epi2ME Analysis
FASTQ files were downloaded from the MinIT compute module to a 1 TB external hard drive on a laptop by connecting to the MinIT WiFi according to the manufacturer’s instructions. The FASTQ files were then uploaded to ‘Epi2ME Fastq WIMP Workflow’ or ‘Epi2ME Fastq Custom Alignment Workflow’ using the Epi2ME desktop agent installed on a laptop (Dell Latitude Precision 5520), using the default Epi2ME WIMP settings (i.e., no minimum or maximum read length and minimum quality score of 7). Default settings for the Epi2ME Custom Reference Alignment Workflow were also used. Genbank accession number AJ004801.1 was used as the BoHV-1 reference genome in all alignments.
2.9. Sequencing PCR-Free Libraries with the Field Sequencing Kit
Non-barcoded PCR-free libraries were generated using the Field Sequencing Library Preparation Kit (LRK001) (Oxford Nanopore Technologies, UK) according to the manufacturer’s instructions. The libraries were sequenced on a FLO-MIN106D R9.4.1 flowcell (one library per flowcell) on a MinION Mk1b sequencer, attached to a MinIT compute module, for 24 h (using the rapid base calling in ONT Minknow software that was installed on the MinIT) and the resulting FASTQ files were uploaded to the Epi2ME Fastq WIMP Workflow for taxonomic assignment.
2.10. MinION Sequencing of Nasal Swabs Calves Challenged with BoHV-1
Nasal swabs were collected daily from the six BoHV-1 challenged calves and six control (PBS challenged) calves from day −1 to day 6 relative to the challenge. Each swab was diluted in 3 mL of PBS, and a 1 mL aliquot of this was used for depleted nucleic acid extraction for sequencing and library preparation. Nucleic acid extraction and library preparation were performed in batches comprising the 8 nasal swabs that were collected from each animal (one swab per day) plus a clean swab as a negative control. To test the technical replication, in each batch, two libraries with different barcodes were prepared from the same swab for 3 of the 8 nucleic acid swab extracts. Each batch of 12 libraries was run on a separate flowcell. The libraries were sequenced on a FLO-MIN106D R9.4.1 flowcell on a MinION Mk1b sequencer attached to a MinIT compute module for 24 h (using rapid base calling in ONT Minknow software that was installed on the MinIT) and the resulting FASTQ files were uploaded to the Epi2ME Fastq WIMP Workflow for taxonomic assignment.
2.11. Genome Assemblies
Adaptor sequences were removed from sequence reads using Porechop 0.2.4. Reads were aligned to the bovine genome using Minimap2 [
28] (version 2.17-r974) to identify host-derived sequences. These were subsequently removed using SAMtools [
29] (v 1.10). All reads less than 100 bp in length were removed using bbmap [
30] (38.22) and the remaining reads were assembled using Flye [
31] (2.8). Assembled contigs were further polished by aligning the original reads using Medaka (1.0.3) [
32].
4. Discussion
The current work demonstrated that correct same-day detection of a known virus in nasal swabs of experimentally challenged calves was achievable using the ONT MinION and its associated Epi2ME cloud-based software. This involved subjecting the nasal swab eluate to mechanical disruption by bead beating, nuclease depletion of non-viral capsid nucleic acid, simple tagmentation-based library preparation with PCR barcoding (to allow multiplexing of libraries, generated from samples and controls, on a single flowcell), rapid base calling of MinION Nanopore sequenceon a MinIT compute module, and rapid cloud-based Epi2ME Fastq WIMP Workflow pathogen identification software.
Due to the fact it was developed prior to the other two MinION flowcells, most MinION sequencing protocols, including rapid tagmentation library preps, were developed using FLO-MIN106D (R9.4.1), which typically has 1200–1600 active R9.4.1 pores. FLO-FLG001 (R9.4.1) is one-tenth of the cost but has only 80–160 active pores. FLO-MIN112 (R10.4) has higher consensus sequencing accuracy but is currently only optimised for libraries prepared using relatively slow ligation methods [
36]. With the FLO-MIN106D (R9.4.1), using the rapid basecalling default option on ONT MinKnow sequencing software, a file containing 4000 basecalled FASTQ reads is generated approximately every two minutes in the early stages of the sequence run. The rate of sequencing declines as the sequence run progresses.
As soon as these FASTQ files are generated, they can be uploaded to a cloud-based software platform called Epi2ME, which contains several intuitive point-and-click sequence analysis applications called ‘workflows’. In the present work, the viral, bacterial, fungal and yeast sequences were identified from nanopore FASTQ files using the ‘Epi2ME Fastq WIMP Workflow’, which employed the ‘Centrifuge’ algorithm [
37]. Centrifuge uses an indexing scheme based on the Burrows–Wheeler transform (BWT) and Ferragina–Manzini (FM) index, which were optimised specifically for metagenomic classification. Centrifuge has space-optimised indexing schemes, requires a relatively small index and classifies sequences at a very high speed; therefore, it can process the millions of sequence reads from a typical high-throughput DNA sequencing run within a few minutes on a desktop computer or laptop [
37]. As long as the proportion of pathogen nucleic acid in the samples relative to that of non-pathogen nucleic acid is sufficiently high, Epi2ME Fastq WIMP Workflow enables the identification of a pathogen within approximately 15 to 30 min of loading a sequencing library on a FLO-MIN106D (R9.4.1) flow cell.
The size of most viral genomes is several orders of magnitude lower than those of bacteria and eukaryotes. Consequently, in nasal swabs taken from cattle infected with a BRD-associated virus, the vast majority of the nucleic acid will be prokaryotic and eukaryotic, and just a fraction will be viral. Several methods are commonly employed to enrich the viral sequence relative to the non-viral sequence in a sample to decrease the amount of sequence depth required to obtain a viral genome sequence from complex samples. The ViroCap targeted sequence capture panel was designed to enrich the nucleic acid from DNA and RNA viruses from 34 families that infect vertebrate hosts [
38] but can detect many viruses that are not on the panel [
39]. ViroCap was used to enrich the animal viruses from clinical samples for sequencing on the MinION but the optimal probe hybridistaion time varied for different viruses and a 20 h probe hybridisation time was adopted [
40]. PCR amplification (using overlapping targeted primers spanning an entire viral genome (spiked primer approach) can be used to increase the amount of whole viral genome sequence from a sample if the genome sequence of the virus is known [
41].
Enrichment of viral nucleic acid can also be achieved via the depletion of non-viral material from a sample. Eukaryotic and prokaryote cells can be separated from the much smaller viral capsids via ultracentrifugation [
42]. However, some giant viruses, such as mimiviruses, which are associated with pneumonia in humans, are larger than some bacteria and thus pellet at lower centrifugation speeds than bacteria [
43].
As intact viral capsids are nuclease resistant, RNaseA and DNase1 can be used to selectively digest non-viral capsid nucleic acids. DNase1 and RNaseA are applied following cell disruption so that the eukaryotic and prokaryotic nucleic acids are exposed to the nucleases [
44]. However, in a cell infected with a virus (i.e., a virocell), much of the virus nucleic acid is not protected by a capsid and this unprotected viral sequence can also be lost if cell disruption and nuclease pre-nucleic acid extraction treatments are applied.
Following the depletion of non-viral nucleic acid, there is often insufficient total nucleic acid to generate enough of a sequencing library for NGS and TGS platforms; therefore, following double-stranded cDNA synthesis, whole-genome amplification (WGA) approaches are usually applied to amplify all of the remaining total nucleic acids in a depleted nucleic acid preparation. These approaches include Sequence-Independent, Single-Primer Amplification (SISPA) and Linker Amplified Shotgun Library (LASL) [
32,
33,
42,
45], which both employ PCR, and isothermal multiple displacement amplification (MDA) using podovirus φ29 polymerase [
42]. Not surprisingly, each WGA method was shown to preferentially amplify different families of viruses and MDA is prone to the generation of a chimeric sequence [
45].
The protocol we developed employed the LASL WGA approach. Compared with MDA and SISPA, LASL sequencing requires fewer reagents, thus lower cost, and fewer steps, thus less time from taking the sample to loading the flowcell. With the LASL procedure we developed, the addition of library adapters and WGA simply comprises a 5-min tagmentation of nucleic acid with a sequencing adapter, followed by a 80-min, 30-cycle PCR amplification with barcoded primers.
There is currently a paucity of literature that describes the use of the experimental challenge of cattle with a known BRD virus to assess these relatively new viral metagenomics approaches. Nanopore sequencing has been used and will likely be increasingly used due to its many advantages over other next-generation sequencing platforms to compare the nasal viromes of cattle with and without BRD to attempt to find or confirm the associations of viruses with BRD [
19]. However, nanopore viral metagenomics should be assessed for several BRD-associated viruses using experimental challenges in cattle with known viruses to check for sensitivity and specificity issues caused by extractability from swabs, varying GC content and varying amounts of extractable viral nucleic acid in swabs from the same animal during infection.
Another group reported the assessment of nanopore sequencing of nasal swabs and tracheal washes from animals that were identified as infected with a BRD-associated virus [
46]. Rather than using experimentally challenged animals, they screened nasal swabs and tracheal washes from 116 animals using qPCR and MiSeq and found that 19 samples were naturally infected with the influenza D virus (IDV). They performed nuclease depletion (with DNase and RNase) of non-capsid-protected nuclease acid in the IDV-positive nasal swab and tracheal wash samples. However, they did not report cell disruption prior to the nuclease treatment. They used a random primer ‘FR20RV’ for WGS but did not give details of the polymerase or cycling conditions; therefore, it is not possible to know whether their protocol would have been able to sequence large viral genomes or genomes with extreme GC content directly from samples. Unlike BoHV-1, which has a relatively large genome (135.3 kb) with very high GC content (72%), IDV only has a small genome (12.3 kb) with an average GC content (41.5%). They generated libraries with the Ligation 1D Sequencing Kit SQK-LSK108, and thus, library generation would have been considerably slower with many more pipetting steps than our LASL method, although they did not report the time it took from sample to result. They also ran the libraries on a GridION, not a MinION. The GridION uses the same flowcells and similar software to the MinION but, unlike the MinION, it is not portable or low-cost.
By looking at a single ‘known’ BRD virus in infected nasal swabs and cell cultures, we were able to reveal significant technical issues with the standard ONT protocols. One of the major problems we encountered was that LongAmp
Taq failed to amplify the BoHV-1 genome efficiently and, instead, preferentially amplified the non-viral DNA. NEB LongAmp
Taq is suggested by ONT for use in many of their protocols, including the whole-genome amplification of nanopore libraries. The very high GC content of the BoHV-1 genomewas most likely the cause of the failure of the PCR amplification of BoHV-1 DNA with NEB LongAmp
Taq. We showed that NEB Next Ultra II Q5 polymerase gave far higher PCR amplification of BoHV-1 DNA than NEB LongAmp
Taq. Whether NEB Next Ultra II Q5 polymerase gives representative amplification of all viruses in the cattle nasal virome (including dsDNA, ssRNA, dsRNA and ssDNA viruses with high, low or average GC content) will have to be carefully assessed in nasal swabs from experimental challenge models in cattle with a range of viruses. A reverse transcription step will also have to be optimised for RNA viruses, as different reverse transcriptases vary widely in their performance in achieving optimal sequence coverage of RNA viral genomes [
47]. Nevertheless, we demonstrated that, with the introduction of double-stranded cDNA synthesis, a rapid tagmentation-based nanopore viral shotgun metagenomics approach could simultaneously and correctly detect RNA and DNA viruses in control mixtures of cultures of three BRD-associated viruses (BoHV-1, BPI3 and BRSV) [
48,
49]. We also recently showed that the current procedure can detect RNA and DNA viruses (e.g., bovine coronavirus, bovine rhinitis virus and ungulate tetraparvovirus) in nasal swabs from naturally infected animals [
49].
Pooling and mixing swabs from different challenge models would also allow us to test the performance of the MinION sequencing and Epi2ME analysis with a ‘known’ mixture of viruses in infected nasal swabs. Most mock communities for viruses are generated from cell cultures. Cell cultures have far fewer non-viral nucleic acids than nasal swabs; therefore, they are not representative of nasal swabs and are consequently suboptimal for developing nasal swab sequencing protocols. In the current study, we observed a much lower percentage of viral sequence in both the depleted and undepleted libraries derived from nasal swabs than from cell cultures. A mock BRD RNA/DNA virus community from infected nasal swabs from experimentally challenged cattle models would be extremely useful for the assessment and optimisation of nanopore nasal virome sequencing protocols.
Spike-in controls would also allow for the determination of specificity. One or two reads were assigned to BoHV-1 in day −1 and day 0 samples and blank swab/PBS negative extraction controls in the first two of the six batches of swabs we processed from BoHV-1-challenged animals. This could have resulted from cross-contamination during DNA extraction or library preparation, and/or index hopping during sequencing. ONT sequencing was reported to generate 0.02–0.3% index hopping [
50]. In the current work, stringent measures were adopted to avoid cross-contamination. To prevent sample aerosol escape during the high-speed bead beating and centrifugation, the lids of the screw-caps (with o-ring gaskets) pathogen lysis tubes were screwed tight and wrapped with Parafilm for bead beating and high-quality Eppendorf Safelock microfuge tubes were used for all centrifugation steps. Minimisation of manual handling steps of the samples could further reduce possible cross-contamination. Automated sample extraction and library preparation using devices such as the Voltrax [
51] or PDQEX [
52] would eliminate many possible cross-contamination steps, although this would require further optimisation. Index hopping could be reduced or eliminated via improved removal of unligated adapters and improved index sequences in the adapters that are supplied in the rapid PCR-barcoding kit.
It would be useful if the quantity of virus in the nasal swab could be estimated from the viral shotgun metagenomics sequence data. However, in the current work, the qPCR analysis showed that BoHV-1 was greatly reduced in depleted, compared with undepleted, nasal swab eluates. Adding an accurately quantified spike-in control of a cocktail of different viruses immediately prior to extraction could allow for an estimation of the loss of viral nucleic acids due to depletion, thus enabling an estimation of the quantity of the virus in the sample prior to depletion. The addition of a second nucleic acid spike-in control immediately prior to adapter ligation and WGA library generation would also be necessary to control for the effects of library preparation and would allow for the estimation of viral quantities in nasal swab samples [
53]. Adding spike-ins introduces the risk of cross-contamination of the viral sequence in the sample being analysed with the spike-in sequence [
54]. Therefore, again, this would have to be carefully tested to ensure that the spike-ins are sufficiently different from the virus sequences in the sample being analysed. As we were still assessing which virus sequences were common to BRD nasal swabs in Ireland, we decided not to use spike-ins in the current work.
Although spike-ins would be necessary to allow for an estimation of the absolute amounts of virus from sequencing, the relative abundance between samples can be estimated without spike-ins. The only quantitative output from Epi2Me Fastq WIMP analysis is read counts. We found that the Epi2Me Fastq WIMP BoHV-1 read counts were very different when we ran the same extracted nucleic acid sample with two different barcodes on the same flowcell. Therefore, read counts (assigned by Epi2ME Fastq WIMP) for a particular virus cannot be relied on for comparing the relative abundance of that virus between samples. However, there was reasonable consistency between the same nucleic acid extract run with different barcodes when the Epi2ME Fastq Custom Alignment Workflow was used to calculate the percentage of bases that aligned with the BoHV-1 reference genome. The percentage of bases that aligned with the BoHV-1 reference genome also showed a stronger relationship with the qPCR analysis of BoHV-1 in the nasal swab eluates. Therefore, for quantification purposes, the Epi2ME Fastq Custom Alignment Workflow is more useful than the Epi2ME Fastq WIMP workflow.
Incorrect assignments of bovine nanopore sequence reads to viral and bacterial taxa by Epi2ME Fastq WIMP is a serious issue. The assignment of bovine sequence to Clostridium botulinum taxa could lead to the application of inappropriate, unnecessary and costly treatment and prevention measures on a farm. The incorrect assignment of a bovine sequence to Proteus phage VB_PmiS-Isfahan is also highly misleading in a research environment and the required time and effort to investigate and discount this as being present in the cattle upper respiratory tract was substantial. These instances were both due to the low-quality assemblies being released in RefSeq. Epi2ME Fastq WIMP cannot currently be relied on in a real-world situation and requires either a database with better curation than NCBI RefSeq or for NCBI to scrutinise submissions more carefully before releasing them onto the RefSeq database. There are likely many more low-quality assemblies in the 56,044 RefSeq sequences used by Epi2ME Fastq WIMP, as we observed that many unexpected, i.e., non-herpes viruses, were detected by this software and not by STAT. However, we did not investigate all of these to determine whether they were incorrectly assigned bovine sequences.
The cultivation of viruses is slow, biased and challenging, and the vast majority of viruses remain uncultivated to date [
55]. Therefore, another objective of using viral shotgun metagenomics on the BoHV-1 nasal swabs was to assess whether the relatively large viral genome of BoHV-1 could be assembled directly from nanopore sequence from swabs from an infected animal. In a previous report, IDV genomes were assembled directly from BRD nasal swab nanopore sequences [
46]. The largest IDV contigs assembled for each sample from Nanopore data ranged from 626 bases to 2308 bases [
46]. The length of our BoHV-1 contigs ranged from 9547 bases to 23,959 bases and we were able to assemble 60% of the BoHV-1 genome directly from nasal swab nanopore sequence taken from a single BRD case. Therefore, depending on the viral load, it should possible to obtain a sequence of a new/unknown virus directly from a nasal swab sample within 24–48 h. However, obtaining swabs with high enough levels of virus to allow a full genome assembly of a DNA virus with a genome >100 kb presents technical difficulties if swabs are taken from animals only after they show symptoms, such as high temperature. In four of the six BoHV-1 challenged calves, the highest number of BoHV-1 nanopore sequence reads were obtained in the swabs taken on days 1 and 2 post-BoHV-1 challenge. This was before the increases in rectal temperatures, which were observed on days 3, 4, 5 and 6 post-BoHV-1 challenge. In a BRD outbreak, it would be necessary to take swabs from a group of symptomatic and close-contact symptom-free animals over several days in order to acquire pre-symptomatic swabs with high enough viral loads to generate high-quality viral genome assemblies directly from nasal swab nanopore sequences.
For the viral genome assembly, we had to use third-party software on a local server, as Epi2ME so far only has an assembly workflow for SARS-CoV-2. While the Epi2ME software platform has huge potential for rapid user-friendly pathogen diagnostics, a major shortfall is the lack of an automated workflow for whole viral genome assembly from FAST5 or FASTQ files generated on the MinION from viral shotgun metagenomic sequencing. The addition of a rapid ‘Epi2ME Fastq viral shotgun metagenomics de novo genome assembly workflow’ to the Epi2ME suite of nanopore sequence analysis software would be extremely valuable for improving the diagnostics of BRD and ultimately the reduction of the incidence of this costly disease and its knock-on effects, particularly the spread of antibiotic resistance caused by the BRD-associated high-level use of broad-spectrum antibiotics.