Journal of Virological Methods 203 (2014) 65–72

Contents lists available at ScienceDirect

Journal of Virological Methods journal homepage: www.elsevier.com/locate/jviromet

Flavivirus detection and differentiation by a microsphere array assay Adam J. Foord, Victoria Boyd, John R. White, David T. Williams, Axel Colling, Hans G. Heine ∗ CSIRO Australian Animal Health Laboratory, Division of Animal, Food and Health Sciences, Geelong, Vic., Australia

a b s t r a c t Article history: Received 10 January 2014 Received in revised form 20 March 2014 Accepted 21 March 2014 Available online 30 March 2014 Keywords: Flavivirus Japanese encephalitis Murray valley encephalitis West Nile virus Kunjin virus Molecular diagnostics

Flaviviruses of the Japanese encephalitis virus (JEV) serocomplex include major human and animal pathogens that have a propensity to spread and emerge in new geographic areas. Different genotypes or genetic lineages have been defined for many of these viruses, and they are distributed worldwide. Tools enabling rapid detection of new or emerging flaviviruses and differentiation of important subgroups have widespread application for arbovirus diagnosis and surveillance, and are crucial for detecting virus incursions, tracking virus emergence and for disease control. A microsphere suspension array assay was developed to identify JEV serocomplex flaviviruses of medical and veterinary importance. Assay performance was evaluated using representative virus strains as well as clinical and surveillance samples. The assay detected all JEV serocomplex viruses tested in this study with an apparent analytical sensitivity equal or better than the reference real-time or conventional RT-PCR assays and was able to identify mixed virus populations. The ability to identify mixed virus populations at a high analytical sensitivity would be pertinent in the Australian context when attempting to detect exotic JEV or West Nile virus (WNV), and differentiate from endemic Murray Valley encephalitis virus and WNV-Kunjin virus. The relatively low cost, the ability to identify mixed virus populations and the multiplex nature makes this assay valuable for a wide range of applications including diagnostic investigations, virus exclusions, and surveillance programs. Crown Copyright © 2014 Published by Elsevier B.V. All rights reserved.

1. Introduction Diseases caused by emerging and re-emerging mosquito-borne flaviviruses are an increasing problem worldwide (Mackenzie et al., 2004; Erlanger et al., 2009; Mackenzie and Williams, 2009; Frost et al., 2012). In an Australasian context, viruses of the Japanese encephalitis virus serocomplex are the most important and can cause viral encephalitis in humans and horses. Japanese encephalitis virus (JEV), Murray valley encephalitis (MVE), West Nile virus (WNV) and the Australian subtype Kunjin virus (KUNV) all belong to this complex. Mosquitoes are the main vector for the transmission of JEV serogroup viruses, with birds considered the major reservoir host for virus maintenance and amplification. For JEV, pigs are also an important amplifying host. Humans and horses are dead-end hosts. The flavivirus genome is approximately 11,000 nucleotides in length and consists of a 5 untranslated region (UTR) of approximately 100 nucleotides, a single open reading frame coding for

∗ Corresponding author at: CSIRO Australian Animal Health Laboratory, Private Bag 24, Geelong, Vic. 3220, Australia. Tel.: +61 3 5227 5278; fax: +61 3 5227 5555. E-mail address: [email protected] (H.G. Heine). http://dx.doi.org/10.1016/j.jviromet.2014.03.018 0166-0934/Crown Copyright © 2014 Published by Elsevier B.V. All rights reserved.

three structural and seven non-structural proteins and a 3 -UTR of approximately 600 nucleotides. In Australia, MVEV is perhaps the most significant member of the JEV serogroup. Although most human infections with MVEV are asymptomatic or cause a non-specific illness, progression to encephalitis can occur (Spencer et al., 2001). Outbreaks of this disease have occurred in mainland Australia in 1951, 1956, 1974, and most recently in 2011 (Marshall, 1988; Smith et al., 2011; Knox et al., 2012). Likewise, KUNV is an endemic Australian flavivirus. It is classified as a sub-species of WNV (lineage 1b; Scherret et al., 2001; Lanciotti et al., 2002) and is the only Australian representative of WNV. KUNV can cause Kunjin encephalitis, a disease that is generally milder in humans than that caused by MVEV (Hall et al., 2002). KUNV and MVEV can also cause neurological disease in horses in Australia (Gard et al., 1977; Frost et al., 2012; Roche et al., 2012). JEV is endemic in many parts of East and Southeast Asia, causing approximately 67,000 cases annually with mortality rate of around 30% (Campbell et al., 2011). The first outbreak in Australia occurred in the Torres Strait islands in 1995 resulting in three human cases, with two being fatal (Hanna et al., 1996). Additional human cases were reported subsequently in Northern Australia in 1998 (Hanna et al., 1999).

66

A.J. Foord et al. / Journal of Virological Methods 203 (2014) 65–72

Table 1 Details of flavivirus strains used for assay validation. Virus species

Strain name

Genotype/lineage

Origin

Year of isolation

JEV JEV JEV JEV JEV MVEV MVEV MVEV MVEV Alfuy virus Alfuy virus WNV WNV-KUNV WNV-KUNV WNV WNV WNV WNV WNV -Koutango Kokobera virus New Mapoon virus

TS00 FU JaOH0566 JKT6468 Muar 08-154300 OR156 NG156 MK6684 MRM3929 CY2269 NY99 MRM61C K16989 Sarafend Rabensburg (97-103) G22886 Sarawak DakAaD5443 NG23516 CY1014

1 2 3 4 5 1 2 3 4 NAa NA 1a 1b 1b 2 3 5 6 7 NA NA

Australia/pig serum Australia/human serum Japan/human Indonesia/mosquito Singapore/brain Australia/horse brain Australia/mosquito Papua New Guinea/human Papua New Guinea/mosquito Australia/swamp pheasant Australia/mosquito USA/human Australia/mosquito Australia/mosquito Unknown/unknown Czech Republic/mosquito India/mosquito Borneo/mosquito Senegal/rodent Australia/mosquito Australia/mosquito

2000 1995 1966 1981 1952 2008 1973 1956 1966 1966 1999 1999 1960 1994 1951 1997 1958 1966 1968 1980 1998

a

NA, not applicable. Virus abbreviations: Japanese encephalitis virus (JEV); Murray Valley encephalitis virus (MVEV); West Nile virus (WNV); WNV -Kunjin (WNV-KUNV).

Molecular methods, principally reverse transcription PCR (RTPCR) and quantitative RT-PCR (qRT-PCR), are now well established for the detection of the RNA of these viruses and are in routine use in many laboratories (Lanciotti et al., 2000; Pyke et al., 2004; SanchezSeco et al., 2005; Ayers et al., 2006; Tang et al., 2006; Chao et al., 2007). Generally these methods are highly sensitive and specific to the intended target. However the inability to detect and/or differentiate multiple agents that have divergent sequences is a significant disadvantage with these methods. Microsphere array assays allow for screening of a multitude of agent related nucleotide markers (up to 500 in the Luminex system) in a single reaction and have become a valuable tool for investigation of disease syndromes. Various assay panels for nucleic acid detection have been developed for medical or veterinary applications, including respiratory viral diseases (Jokela et al., 2012), gastroenteritis pathogens (Li et al., 2011), cystic fibrosis (Johnson et al., 2007), biothreat agents (Janse et al., 2012; Yang et al., 2012) and vesicular diseases of livestock (Hindson et al., 2008). PCR amplification of the target regions usually forms the first step of these assays. Proprietary polystyrene microspheres that contain dyes displaying distinct spectral characteristics form the substrate for these assays. Luminex MagPlex-TAG beads (Luminex Corporation, Austin, USA) contain unique 24 base DNA “anti-TAG” sequences covalently coupled to their surface. This facilitates hybridization of specifically amplified and labeled products containing complementary “TAG” sequences and allows identification by association with particular bead sets in a flow cytometry detection system. Here we report the development of a microsphere array assay (microarray assay) for detection and differentiation of genomic RNA from flaviviruses of the JEV group. We assess the relative sensitivity and specificity of this assay in comparison to reference assays, the preliminary diagnostic performance on a small but well defined set of diagnostic samples, and the capability of the assay to differentiate these viruses.

2. Materials and methods 2.1. Viruses and diagnostic samples/specimens Flaviviruses used to evaluate the specificity and analytical sensitivity were derived from stocks held the Australian Animal Health Laboratory (AAHL). Where available, viruses were selected to

represent genotypes or lineages of each of the viral species belonging to the JEV group (Table 1). In addition, the Edge Hill, Dengue, Sindbis, Ross River, Hendra and equine influenza viruses were used to assess assay specificity. Viruses were passaged in PSEK, Vero or C6/36 cells and their titers determined in PSEK cells. WNV Rabensburg was propagated by intracerebral inoculation of 0.02 ml virus in suckling mouse intracerebral, following approved procedures of the AAHL Animal Ethics Committee. Samples utilized in the diagnostic evaluation study included archival clinical specimens from horses and ducks submitted to the laboratory for flavivirus diagnosis in 2011 from the Australian states of Victoria and South Australia. Horse samples were brain and spinal cord specimens, as well as derived tissue culture isolates. Duck samples were brain, lung, liver, kidney, spleen and gastro-intestinal specimens. Virus isolates derived from these tissues in cell culture or embryonated chicken eggs were also tested. Serum samples from ducks experimentally infected with MVEV were also included in this study. Additional tissue samples from infected horses and mosquito pools (Culex annulirostris) were obtained from Dr Mark Fegan (DPI Victoria). Homogenates of mosquito pools collected in Western Australia as part of arbovirus surveillance activities were provided by Dr Cheryl Johansen (University of Western Australia). These comprised pools of 25 mosquitoes comprising a single species of Culex annulirostris, Cx. globocoxitus, Aedes normanensis, Ae. vigilax or Ae. camptorhynchus that were positive for MVEV, KUNV, Sindbis and Ross River viruses, or that tested negative for flaviviruses or alphaviruses using an enzyme immunoassay following cell culture passage of homogenates at the UWA Arbovirus Surveillance Laboratory (Quan et al., 2011). Nucleic acid was isolated from each sample using the MagMAX 96 Viral RNA Extraction Kit (Life Technologies, Carlsbad, USA, Cat. No. AM1836-5).

2.2. Design of the microarray assay An eight-plex microarray assay was developed to identify JEV serocomplex flaviviruses of medical and veterinary importance. The assay is based on a single generic RT-PCR amplification followed by target-specific primer extension (TSPE) in a multiplex reaction (Fig. 1, Table 2). The generic RT-PCR was designed to amplify a highly conserved segment in the 3 -UTR of the flavivirus genome. This RT-PCR generated a 360 bp fragment and a truncated 285 bp fragment due to a second internal binding site for the reverse

A.J. Foord et al. / Journal of Virological Methods 203 (2014) 65–72

67

Fig. 1. Design of JEV serocomplex microarray assay. Virus identification is based on multiplexed target-specific primer extension (TSPE) reactions within a generic JEV serogroup RT-PCR amplicon (primers D-621F and D-622R). Eight TSPE primers (Table 2) binding to target regions 1–4 (shaded gray) facilitated the extension of ssDNA products (dotted line) either generic for all JEV serocomplex strains (Region 1) or specific for only one or several of the virus species/genotypes/lineages (Regions 2–4). Primer specificity is indicated along the dotted line.

Table 2 Oligonucleotides used in microarray assay. Name

Functiona

Sequence 5 to 3 b

Specificityc

Target amplification D-621F D-622R

RT-PCR fwd (+M13F) RT-PCR rev (+M13R)

TGTAAAACGACGGCCAGTGTAGACGGTGCTGCCTGC CAGGAAACAGCTATGACCGGGTCTCCTCTAACCTCTAGTCC

JEV serocomplex generic JEV serocomplex generic

Virus identification D-649 D-653 D-652 D-674 D-650 D-694 D-654 D-692

TSPE(MTAG-A022) TSPE(MTAG-A039) TSPE(MTAG-A033) TSPE(MTAG-A030) TSPE(MTAG-A025) TSPE(MTAG-A015) TSPE(MTAG-A042) TSPE(MTAG-A048)

CAAACAAACATTCAAATATCAATCAACCCCAGGAGGACTGGGT ACAAATATCTAACTACTATCACAACTCAGAACCGTCTCGGAAG ACTACTTATTCTCAAACTCTAATACTCAGAACCGTCTCGGAAA CTTAACATTTAACTTCTATAACACATGAAGCCCGTGTCAGATCG CTTTCTTAATACATTACAACATACTCAAGGCCCAATGTCAGACCA TACTTCTTTACTACAATTTACAACAGGAGTGCAATCTGTGAG CACTACACATTTATCATAACAAATGGAGAGTGCAGTCTGCGAT AATCAACACACAATAACATTCATAGGAGAGTGCAGTCTGYGAC

JEV serocomplex generic JEV, MVEV, WNV KUNV MVEV WNV, KUNV MVEV WNV KUNV

a

MagPlex-TAG bead numbers (Luminex MTAG#) are indicated for each of the target-specific primer extension (TSPE) primers. Flavivirus-specific sequences are indicated in bold. Extensions incorporating M13 or complement MTAG sequences (Luminex Corporation) are appended to the 5 end of primers. c Japanese encephalitis virus (JEV), Murray valley encephalitis virus (MVEV), West Nile virus [other than Kunjin] (WNV), WNV-Kunjin virus (KUNV). b

primer. M13 sequence extensions were added to the PCR primers (D-621F and D-622R in Table 2) providing an option for convenient sequencing if this was required later. Eight TSPE primers were designed within these PCR fragments to target regions either conserved across all JEV serocomplex strains or only in one or several of the virus species or genotypes/lineages (Fig. 1). Each of the TSPE primers contained a 24 bp anti-tag sequence extension complementary to the unique tag on each of the eight chosen MagPlex-TAG microsphere sets (Luminex, Austin, USA). The reactivity pattern of the 8-plex assay allowed for generic detection of JEV group viruses and differentiation between important virus species and lineages (Fig. 1). Other viruses such as ALFV, KOKV, New Mapoon and Usutu would only be detected by the JEV serocomplex generic TSPE primer (D-649) and require DNA sequence analysis of the primary PCR amplicon for unambiguous virus identification. 2.3. Microarray assay procedure The microarray assay procedure was modified from a previous assay for detection and differentiation of henipaviruses (Foord et al., 2013).

2.3.1. Primary RT-PCR Single-step reverse transcription PCR (RT-PCR) was performed using Superscript III One-Step RT-PCR with Platinum Taq kit (Invitrogen, Carlsbad, USA) with the following conditions: 25 ␮l volume, 200 nM forward (D-621F) and reverse (D-622R) primers, 2.0 mM MgSO4 . Thermal cycling conditions were: 30 min at 48 ◦ C (RT reaction), 2 min at 94 ◦ C (Taq activation), 45 cycles of 30 s at 94 ◦ C, 40 s at 50 ◦ C, and 40 s at 68 ◦ C, followed by 68 ◦ C for 7 min . The unicorporated dNTPs and primers from the initial RT-PCR were removed by treating with ExoSAP-IT (Affymetrix, Santa Clara, USA). Twenty-five microliters of RT-PCR was treated with 10 ␮l ExoSAP-IT and incubated at 37 ◦ C for 30 min, followed by 10 min at 80 ◦ C to inactivate the enzymes.

2.3.2. Target-specific primer extension (TSPE) Linear amplification was undertaken in the presence of biotin labeled cytosine with the eight TSPE primers (Table 2). The 5 ends of the TSPE primers were designed to contain 24 base TAG sequences complementary to the particular microsphere sets, whereas the remainder of the primer sequence was designed to bind to targets within the RT-PCR product.

68

A.J. Foord et al. / Journal of Virological Methods 203 (2014) 65–72

Fig. 2. Distribution of flavivirus positive and negative populations in the microarray assay. Distribution of Median Fluorescence Intensity (MFI) values for flavivirus positive and negative populations are shown for horse (A), duck (B) and mosquitoes (C). Flavivirus status 0 (negative) or 1 (positive) was determined by MVEV and KUNV qRT-PCR (A), MVEV qRT-PCR (B) and flavivirus ELISA and KUNV qRT-PCR (C). Arbitrary cut-off values (>299.5 MFI for horse, >330 MFI for duck and >120 MFI for mosquito pools) were determined by MedCal statistical software (Version 12.3.0.0) to achieve highest combined sensitivity and specificity. Additional data in Supplementary Table 1.

Biotin-dCTP was incorporated in the reaction to allow detection by streptavidin-R-phycoerythrin (SA-PE). Each TSPE reaction contained 5 ␮l of Exo-SAP-treated RT-PCR product, 0.75 U Tsp DNA polymerase (Invitrogen, Carlsbad, USA), 25 nM TSPE primer, 5 ␮M dATP/dTTP/dGTP and biotin-dCTP (Invitrogen, Carlsbad, USA), 1X Tsp DNA polymerase reaction buffer (Invitrogen, Carlsbad, USA) and 4.0 mM MgCl2 . Thermocycling was performed at 95 ◦ C for 2 min, followed by 30 cycles of 94 ◦ C for 30 s, 50 ◦ C for 30 s and 72 ◦ C for 40 s with a final extension at 72 ◦ C for 5 min. 2.3.3. Microsphere hybridization Five microliters of TSPE reaction were hybridized with 500 beads of each microsphere set in a total of 50 ␮L of 1X hybridization buffer (0.2 M NaCl/0.1 M Tris/0.08% Triton X-100, pH 8.0). The hybridization mixture was incubated at 96 ◦ C for 90 s and 37 ◦ C for 30 min. The microsphere mixture was transferred to a blacksided 96 well Bio-plex flat bottom plate (Bio-Rad, Hercules, USA) and magnetic microspheres were washed using an automated plate washer (Bio-Plex pro II wash station; Bio-Rad, Hercules, USA). 2.3.4. Microsphere identification and fluorescence detection Seventy-five microlitres of 1X hybridization buffer containing 2 mg/L streptavidin-R-phycoerythrin (Invitrogen, Carlsbad, USA) was added and the mixture was incubated in the dark at 37 ◦ C for 15 min. Each assay plate was analyzed in a Bio-Plex 200 instrument

(Bio-Rad, Hercules, USA) using manufacturer’s protocols. Briefly, 50 ␮L of the microsphere/TSPE/streptavidin-R-phycoerythrin mixture was injected into the instrument at a sample plate temperature of 37 ◦ C and run at high RP1 target setting, with 100 of each microsphere set analyzed per well. Fluorescence was measured as units of Median Fluorescence Intensity (MFI). A positive result was defined as a value greater than three times the MFI obtained from negative controls. Distribution of MFI values for qRT-PCR negative and positive samples is illustrated in Fig. 2. All instrument MFI values in the multiplex reaction were converted to a positive or negative result as determined by cut-off threshold. From these results, flavivirus strains were identified and differentiated by their characteristic reactivity patterns (Table 3). Primer D-649 (bead #22 in target region 1) was designed for generic detection of all JEV serocomplex viruses. Pairwise comparison within region 2 (D-653 bead #39; D-652 bead #33) and region 3 (D-674 bead #30; D-650 bead #25) identified and differentiated between MVEV, WNV species and WNV-KUNV subtype. Reactivity in region 4 (D-694 bead #15; D-654 bead #42; D-692 bead #48) was used to confirm results obtained from regions 2 and 3. Primer D-654 was designed to specifically target WNV lineage 1a viruses. The identity of viruses that are only detected by the JEV serocomplex generic TSPE primer (D-649) or that showed ambiguous reactivity patterns were confirmed using M13 primers for DNA sequencing of the primary PCR amplicon (D-621F and D622R).

A.J. Foord et al. / Journal of Virological Methods 203 (2014) 65–72

69

Table 3 Flavivirus differentiation and limit of detection of the microarray assay. Region 1

Region 2

TSPE primer Specificity

D-649 JEV MVEV WNV KUNV

D-653 JEV MVEV WNV

D-652 KUNV

D-674 MVEV

D-650 WNV KUNV

D-694 MVEV

D-654 WNV

D-692 KUNV

(Reference)

JEV TS00 (genotype 1) FU (genotype 2) JaOH0566 (genotype 3) JKT6468 (genotype 4) JEV Muar (genotype 5)

10e-6 10e-5 10e-5 10e-7 10e-6

10e-6 10e-5 10e-5 10e-7 10e-6

Neg. Neg. Neg. Neg. Neg.

Neg. Neg. Neg. Neg. Neg.

Neg. Neg. Neg. Neg. Neg.

Neg. Neg. Neg. Neg. Neg.

Neg. Neg. Neg. Neg. Neg.

Neg. Neg. Neg. Neg. Neg.

10e-4 (P) Neg. (P) 10e-4 (P) 10e-4 (P) Neg. (P)

MVEV MVE Horse MVE NG156 MVE MK6684 MVE OR156

10e-5 10e-5 10e-6 10e-7

10e-5 10e-5 10e-6 10e-7

Neg. Neg. Neg. Neg.

10e-5 10e-5 10e-6 10e-7

Neg. Neg. Neg. Neg.

10e-5 10e-5 10e-6 10e-7

Neg. Neg. Neg. Neg.

Neg. Neg. Neg. Neg.

10e-7 (P) 10e-4 (P) 10e-7 (P) 10e-6 (P)

WNV (non KUNV) NY99 (lineage 1a) G22886 (lineage 1c/5) Sarafend (lineage 2) Koutango (DakAaD5443) WNV Sarawak (6) Rabensberg

10e-6 10e-6 10e-5 10e-6 10e-5 Pos.

10e-6 10e-6 10e-5 10e-6 10e-5 Neg.

Neg. Neg. Neg. Neg. Neg. Pos.

Neg. Neg. Neg. Neg. Neg. Neg.

10e-6 10e-6 10e-5 10e-6 10e-5 Pos.

Neg. Neg. Neg. Neg. Neg. Neg.

10e-6 Neg. Neg. Neg. 10e-4 Neg.

Neg. Neg. Neg. Neg. Neg. Neg.

10e-6 (L) 10e-6 (T) 10e-5 (T) 10e-6 (T) 10e-6 (T) ND

10e-6 10e-6

Neg. Neg.

10e-6 10e-6

Neg. Neg.

10e-6 10e-6

Neg. Neg.

Neg. Neg.

10e-4 10e-5

10e-6 (P) 10e-7 (P)

10e-4 10e-6

Neg. Neg.

Neg. Neg.

Neg. 10e-4

Neg. Neg.

Neg. Neg.

Neg. Neg.

Neg. Neg.

10e-2 (C) 10e-2 (C)

10e-4 10e-5

Neg. Neg.

Neg. Neg.

Neg. Neg.

Neg. Neg.

Neg. Neg.

Neg. Neg.

Neg. Neg.

10e-2 (C) 10e-5 (C)

WNV-KUNV (lineage 1b) MRM61C K16989 ALFV MRM3929 CY2269 Kokobera virus group Kokobera NG23516 New Mapoon CY1014

Region 3

Comparisona

Target

Region 4

Highest dilutions yielding positive signals using 10 fold serially (10e-4 to 10e-8) diluted RNA extracted from representative strains of JEV, MVEV, WNV ALFV and Kokobera group viruses are indicated in bold. Direct comparison to the relevant diagnostic assay is shown in the last column. Neg. = negative at 10e-4 dilution. ND = not done. a qRT-PCR assay: (L), Lanciotti et al. (2000); (T), Tang et al. (2006); (P), Pyke et al. (2004); RT-PCR assay: (C), Chao et al. (2007).

2.4. Comparative molecular assays Flavivirus qRT-PCR assays and a conventional RT-PCR were used for comparative assessment of the microarray assay. Specific qRT-PCR assays designed to detect WNV strain NY99 (lineage 1a; Lanciotti et al., 2000), lineage 1 and 2 of WNV (Tang et al., 2006), MVEV, KUNV and JEV (Pyke et al., 2004) were also utilized. Oligonucleotide primers and probes were as described in these manuscripts. Assays were performed using AgPath-ID One-Step RT-PCR Reagents (Life Technologies, Carlsbad, USA) with 250 nM probe and 900 nM forward and reverse primers. Thermal cycling was 45 ◦ C for 10 min, followed by 45 cycles of 95 ◦ C for 15 s and 60 ◦ C for 45 s. Cut-off values were cycle threshold (CT) ≤40 for positive and CT ≥45 for negative. Results with CT values between 40 and 45 were deemed indeterminate, i.e. not conclusively positive or negative. A conventional RT-PCR for detection of all flavivirus based on a qRT-PCR assay (Chao et al., 2007) was utilized for ALFV, Kokobera and New Mapoon viruses where specific qRT-PCR assays were not readily available. Forward and reverse primers were as described in the manuscript and RT-PCR conditions were as described in Section 2.3.1. 2.5. DNA sequencing PCR amplicons containing M13-specific extensions incorporated in forward and reverse primers were analyzed by agarose gel electrophoresis. Fragments of correct size were excised and

extracted using the QIAquick Gel Extraction kit (Qiagen, Hilden, Germany). DNA sequences were determined using Big Dye terminator v1.1 chemistry on an ABI 3130XL Genetic Analyser capillary electrophoresis instrument. Identity of the sequences was determined using alignments of representative strains of flaviviruses and BLAST analyses. 3. Results 3.1. Differentiation of flaviviruses by microarray assay The ability of the microarray assay to differentiate diverse members of the JEV serocomplex of flaviviruses was assessed using representative strains listed in Table 1. Viruses were identified by their reactivity pattern in the microarray assay. All JEV serogroup viruses tested were identified as belonging to this group by their reactivity with the generic assay (Region 1; comprising primer D-649). Region 2 (comprising primers D-653 and D-652) differentiated KUNV and WNV Rabensburg from JEV, MVEV and other WNV strains tested, so that only KUNV and Rabensburg strains were positive in reaction with D-652. Region 3 (comprising primers D674 and D-650) differentiated MVEV from WNV; only MVEV strains were positive in reaction with D-674, while all WNV strains were positive in reaction with D650. Region 4 (comprising primers D694, D-654 and D-692) verified results of the other regions for MVEV and KUNV strains. Primer D-654 detected WNV strains NY99 and Sarawak but not G22886, Sarafend, Koutango and Rabensburg. Although no JEV-specific test was designed, all JEV genotypes

70

A.J. Foord et al. / Journal of Virological Methods 203 (2014) 65–72

Table 4 Ability of the microarray assay to identify WNV and KUNV in a mixed virus population. Virus dilution

KUNV 10e-1

WNV 10e-1 10e-2 10e-3 10e-4 10e-5 10e-6

both both KUNV KUNV KUNV KUNV

10e-2 both both KUNV KUNV KUNV KUNV

10e-3 WNV both both KUNV KUNV KUNV

10e-4 WNV WNV both both both KUNV

10e-5 WNV WNV both both both KUNV

or greater resulted in detection of only one virus. Thus, although both viral genomes can be detected when present equally, there was not consistency in the ability of the assay to detect both viruses when one viral RNA was in excess of the other.

10e-6 WNV WNV WNV both both both

Virus stocks of WNV (NY 99) and KUNV (K16989) were adjusted to 1E + 06 TCID50/ml. RNA was extracted, serially diluted and combined in different ratios. Microarray assay was performed to determine its ability to identify one or both viruses (marked as shaded area) in the mixtures.

3.2.3. DNA sequencing Following DNA sequencing of PCR products on examples of each virus species, alignments to known flaviviruses and blast analysis were performed. The identity of these agents differentiated previously in the microarray assay was confirmed. Where the facility to differentiate clearly was not available, sequence analysis was able to provide this utility. This was evident for JEV, Alfuy and Kokobera viruses which only reacted with region 1 or region 1 and 2 assays, and thus where not definitively classified by the microarray assay. 3.3. Diagnostic evaluation of microarray assay

displayed a characteristic reactivity pattern. Positive results using primers D-649 and D-653 and negative for all other primers indicated a JEV positive sample. The other flaviviruses tested, ALFV, Kokobera and New Mapoon virus were detected by the generic primer D-649 (region 1) but did not display a unique reactivity profile with the TSPE primers. The assay was specific for JEV group viruses as all non-related viruses tested, i.e. Dengue, Edge Hill, Hendra, Nipah and equine influenza were negative in the microarray assay. 3.2. Analytical evaluation of the microarray assay 3.2.1. Comparison of microarray assay to reference assays The analytical sensitivity of the microarray assay was assessed in direct comparison to qRT-PCR assays specific for either WNV (NY99), MVEV, WNV (lineages 1 and 2), KUNV or JEV. A conventional RT-PCR specific for members of the JEV group was also utilized for direct comparison for Alfuy and Kokobera group viruses. The comparative limit of detection of the assays was determined using tenfold serially diluted RNA template from viruses listed in Table 3. Generally the microarray assay had an analytical sensitivity and dynamic range similar to qRT-PCR/conventional RT-PCR. More specifically, the microarray assay performed better than the qRTPCR on the JEV positive samples where 2 of the 5 samples were not detected by the qRT-PCR assay. Of the 3 samples detected by both assays the microarray assay was from 10 to 100X more sensitive. For the MVEV infected samples the microarray assay was 10X more sensitive in 2 of the 4 samples, the qRT-PCR being more sensitive in the remaining 2. For the WNV positive samples (inc. KUNV) the two assays had equivalent sensitivities, apart from Sarawak, and K16989 strains, were qRT-PCR results were 10X more sensitive; however, the microarray assay retained the utility to differentiate KUNV from other strains of WNV at all dilution points (Table 3). For the detection of Alfuy and Kokobera group viruses the microarray assay detected all strains and was as sensitive (New mapoon virus) as qRT-PCR or 100X (Alfuy MRM3929, Kokobera NG23516) to 10,000X (Alfuy CY2269) more sensitive than qRT-PCR (Table 3). 3.2.2. Detection of viruses in a mixed population The capability of the microarray assay to detect and differentiate different flaviviruses in a mixed population was investigated using WNV and KUNV in a checkerboard mixing experiment. Dilutions of KUNV (K16989) and WNV (NY99) RNA were combined in varying ratios, according to Table 4. In the presence of equimolar concentration of viral RNA, the microarray assay was able to detect both RNA species, and this was evident over the entire concentration range (Table 4). In some tests where either KUNV or WNV RNAs were present in 10 to 100X excess, dual detection could be observed. Conversely in other tests, an excess of one viral RNA species by 10-fold

Diagnostic sensitivity and specificity was evaluated using a limited but well defined set of samples. This consisted of equine and avian (duck) samples from diagnostic submissions and duck serum samples taken over the course of an experimental infection study. There was a clear differentiation of MFI values for the positive and negative samples from horses (Fig. 2A), ducks (Fig. 2B) and mosquito pools (Fig. 2C). There was strong concordance between the microarray assay data with retrospective qRT-PCR results. Correlation between microarray assay results (MFI values) and qRT-PCR (Ct values) and virus identification are available in the Supplementary Table 1. Of 53 equine diagnostic samples both microarray assay and qRT-PCR assay identified 17 samples as flavivirus positive. Ten were identified as MVEV by both assays. One of the samples identified as MVEV positive by qRT-PCR reacted positive for both MVEV and KUNV by microarray assay, indicative of a mixed infection. All seven KUNV positive samples by qRT-PCR were also positive in the microarray assay. Ninety-one duck samples were available for retrospective analysis consisting of 11 diagnostic submissions and 80 samples from an MVEV experimental infection study. For the diagnostic submissions there was 100% concordance of results between qRT-PCR and microarray assay (MVEV positive n = 9, negative n = 2). Of the 80 experimental infection samples, 69 were positive and 11 negative for MVEV in the microarray. In the MVEV qRT-PCR assay 68 samples were positive seven negative and 5 indeterminate. One sample negative by qRT-PCR was positive by microarray assay. The true status of this sample remains unknown as additional sample was not available. As a consequence it could have been either a false positive in the array or a false negative in qRT-PCR. Of the five samples that returned qRT-PCR indeterminate results two were positive and three negative by microarray assay. Twenty-nine pools of mosquito samples from Western Australia (n = 27) and Victoria (n = 2) were analyzed by microarray assay. All pools of known flavivirus positives (n = 11) were correctly identified by the microarray assay. Of these, three pools of Cx. annulirostris mosquitoes and one pool of Ae. vigilax were identified as KUNV, six pools of Cx. annulirostris as MVEV and one as a mixed population of KUNV and MVEV. All flavivirus negative pools (n = 9; Cx. annulirostris) and those previously identified as infected with SINV (n = 3; Cx. annulirsotris) or RRV (n = 6; Ae. camptorhynchus, Cx. globocoxitus) were negative in the microarray assay. 4. Discussion Flaviviruses of the JEV serocomplex include major human and animal pathogens that have a propensity to spread and emerge in new geographic areas. Different genotypes or genetic lineages have been defined for many of these viruses, and they are distributed worldwide. JEV group viruses have a complex life cycle, involving birds or pigs as primary hosts, mosquitoes as primary vectors,

A.J. Foord et al. / Journal of Virological Methods 203 (2014) 65–72

and humans, horses, and other mammals as incidental hosts. Tools enabling rapid detection of new or emerging flaviviruses and differentiation of important subgroups have widespread application for arbovirus diagnosis and surveillance, and are crucial for detecting virus incursions, tracking virus emergence and for disease control. The 8-plex microarray assay described in the study detected viruses representative of the JEV serocomplex, with an apparent analytical sensitivity equal or better than the reference qRT-PCR or RT-PCR assays. To achieve the range of viruses detected in the microarray assay three separate virus-specific qRT-PCR (Pyke et al., 2004) and one conventional RT-PCR assay (Chao et al., 2007) were required. The qRT-PCR assays have the advantage of less potential for contamination due to the closed system employed (tube or sealed plate), less hands on time and a faster result (approx. 2 h). Microarray on the other hand contains multiple tube opening and pipetting steps that carry the risk of cross contamination and require careful PCR procedures and workflow. In addition to more manipulations, the time to result is longer (approx. 8 h). A significant advantage of the microarray assay was the ability for simultaneous detection of a range of viruses and the differentiation of closely related viruses. A single TSPE reaction comprising of region 1 detected all strains of the JEV serocomplex. The differentiation of JEV from other flaviviruses was not pursued. This was principally due to the complexities of designing additional multiple primers encompassing the sequence variations associated with the different genotypes. Viruses positive in region 1, but not identifiable using regions 2 to 4 can be characterized by DNA sequencing of the RT-PCR product. In this study, DNA sequencing was facilitated by the insertion of M13 primer extensions in the forward and reverse primers, and was used to identify ALFUY and Kokobera viruses. Although Usutu virus (Savini et al., 2011) samples were not available for testing in this study, in silico analysis of target primer sequences indicated positive reaction is expected with generic JEV group TSPE primer D-649. The results obtained from regions 2 and 3 provided a unique pattern for each of the JEV, MVEV, WNV and KUNV species, allowing unambiguous differentiation. The identification of JEV relied on a single reaction with TSPE primer D-653 in region 2 and may require confirmation by DNA sequencing of the initial RT-PCR product for unambiguous identification. MVEV and KUNV, which co-circulate in Australia, could be differentiated from each other and from exotic WNV lineages and JEV. Importantly, KUNV (WNV lineage 1b) was clearly distinguishable from closely related WNV NY strain (lineage 1a) and other WNV strains. The differentiation of these closely related strains has been a challenge for other diagnostic assays, including qRT-PCR. This differentiation was based on the reactivity pattern of virus-specific TSPE reactions for the major virus groups in four different genomic regions. The utility of the assay is enhanced by the ability to detect mixed virus populations. An interesting finding was the identification of an apparent double infection of MVEV and KUNV in one of the equine diagnostic samples and the presence of both viruses in one of the mosquito pools. This was not readily apparent in the original diagnosis using qRT-PCR alone. The ability to detect mixed virus populations was demonstrated over a wide concentration range and various ratios of exotic WNV in the presence of endemic KUNV. However, the ability to detect the minority virus may be limited by the greater amplification and reaction saturation by the predominant virus. The capability to detect multiple viruses would be pertinent in the Australian context when attempting to detect exotic JEV or WNV, and differentiate from endemic MVEV and KUNV. The diagnostic utility of the microarray assay was demonstrated by the strong agreement with qRT-PCR results from samples from the Australian 2011 flavivirus outbreak (Mann et al., 2013) and from ducks experimentally infected with MVEV. The microarray assay reported here offers unique advantages for the detection and differentiation of medically or economically

71

important JEV serocomplex viruses. The microarray assay was able to detect a wide range of viruses and simultaneously differentiate closely related viruses, in particular KUNV and WNV (NY99). This was achieved at sensitivities equal to published molecular assays. The relatively low cost and the ability to identify mixed virus populations makes this assay valuable for a wide range of diagnostic and epidemiological investigations, such as virus exclusions and surveillance programs. Acknowledgements We thank Susie Daglas for assistance with the preparation and titration of viruses, Tyrone McDonald for sample identification and help with qPCR, Vicky Stevens and Kelly Davies for DNA sequencing. We thank Dr Mark Fegan (DPI Victoria) and Dr Cheryl Johansen (University of Western Australia) for providing diagnostic samples. We also thank Prof. Roy Hall (University of Queensland), Prof. Norbert Nowotny (University of Veterinary Medicine, Vienna) and Dr Ken Morita (Nagasaki University) for generously providing flavivirus stocks. Appendix A. Supplementary data Supplementary material related to this article can be found, in the online version, at http://dx.doi.org/10.1016/j.jviromet. 2014.03.018. References Ayers, M., Adachi, D., Johnson, G., Andonova, M., Drebot, M., Tellier, R., 2006. A single tube RT-PCR assay for the detection of mosquito-borne flaviviruses. J. Virol. Methods 135, 235–239. Campbell, G.L., Hills, S.L., Fischer, M., Jacobson, J.A., Hoke, C.H., Hombach, J.M., Marfin, A.A., Solomon, T., Tsai, T.F., Tsu, V.D., Ginsburg, A.S., 2011. Estimated global incidence of Japanese encephalitis: a systematic review. Bull. World Health Organ 89, 766–774. Chao, D.Y., Davis, B.S., Chang, G.J., 2007. Development of multiplex real-time reverse transcriptase PCR assays for detecting eight medically important flaviviruses in mosquitoes. J. Clin. Microbiol. 45, 584–589. Erlanger, T.E., Weiss, S., Keiser, J., Utzinger, J., Wiedenmayer, K., 2009. Past, present, and future of Japanese encephalitis. Emerg. Infect. Dis. 15, 1–7. Foord, A.J., White, J.R., Colling, A., Heine, H.G., 2013. Microsphere suspension array assays for detection and differentiation of Hendra and Nipah Viruses. Biomed. Res. Int. 2013, 1–8. Frost, M.J., Zhang, J., Edmonds, J.H., Prow, N.A., Gu, X., Davis, R., Hornitzky, C., Arzey, K.E., Finlaison, D., Hick, P., Read, A., Hobson-Peters, J., May, F.J., Doggett, S.L., Haniotis, J., Russell, R.C., Hall, R.A., Khromykh, A.A., Kirkland, P.D., 2012. Characterization of virulent West Nile virus Kunjin strain, Australia, 2011. Emerg. Infect. Dis. 18, 792–800. Gard, G.P., Marshall, I.D., Walker, K.H., Acland, H.M., Saren, W.G., 1977. Association of Australian arboviruses with nervous disease in horses. Aust. Vet. J. 53, 61–66. Hall, R.A., Broom, A.K., Smith, D.W., Mackenzie, J.S., 2002. The ecology and epidemiology of Kunjin virus. Curr. Top. Microbiol. Immunol. 267, 253–269. Hanna, J.N., Ritchie, S.A., Phillips, D.A., Shield, J., Bailey, M.C., Mackenzie, J.S., Poidinger, M., McCall, B.J., Mills, P.J., 1996. An outbreak of Japanese encephalitis in the Torres Strait, Australia, 1995. Med. J. Aust. 165, 256–260. Hanna, J.N., Ritchie, S.A., Phillips, D.A., Lee, J.M., Hills, S.L., van den Hurk, A.F., Pyke, A.T., Johansen, C.A., Mackenzie, J.S., 1999. Japanese encephalitis in north Queensland, Australia, 1998. Med. J. Aust. 170, 533–536. Hindson, B.J., Reid, S.M., Baker, B.R., Ebert, K., Ferris, N.P., Tammero, L.F., Lenhoff, R.J., Naraghi-Arani, P., Vitalis, E.A., Slezak, T.R., Hullinger, P.J., King, D.P., 2008. Diagnostic evaluation of multiplexed reverse transcription-PCR microsphere array assay for detection of foot-and-mouth and look-alike disease viruses. J. Clin. Microbiol. 46, 1081–1089. Janse, I., Bok, J.M., Hamidjaja, R.A., Hodemaekers, H.M., van Rotterdam, B.J., 2012. Development and comparison of two assay formats for parallel detection of four biothreat pathogens by using suspension microarrays. PLoS One 7 (2), e31958. Johnson, M.A., Yoshitomi, M.J., Richards, C.S., 2007. A comparative study of five technologically diverse CFTR testing platforms. J. Mol. Diagn. 9, 401–407. Jokela, P., Piiparinen, H., Mannonen, L., Auvinen, E., Lappalainen, M., 2012. Performance of the Luminex xTAG Respiratory Viral Panel Fast in a clinical laboratory setting. J. Virol. Methods 182, 82–86. Knox, J., Cowan, R.U., Doyle, J.S., Ligtermoet, M.K., Archer, J.S., Burrow, J.N., Tong, S.Y., Currie, B.J., Mackenzie, J.S., Smith, D.W., Catton, M., Moran, R.J., Aboltins, C.A., Richards, J.S., 2012. Murray Valley encephalitis: a review of clinical features, diagnosis and treatment. Med. J. Aust. 196, 322–326.

72

A.J. Foord et al. / Journal of Virological Methods 203 (2014) 65–72

Lanciotti, R.S., Kerst, A.J., Nasci, R.S., Godsey, M.S., Mitchell, C.J., Savage, H.M., Komar, N., Panella, N.A., Allen, B.C., Volpe, K.E., Davis, B.S., Roehrig, J.T., 2000. Rapid detection of West Nile virus from human clinical specimens, field-collected mosquitoes, and avian samples by a TaqMan reverse transcriptase-PCR assay. J. Clin. Microbiol. 38, 4066–4071. Lanciotti, R.S., Ebel, G.D., Deubel, V., Kerst, A.J., Murri, S., Meyer, R., Bowen, M., McKinney, N., Morrill, W.E., Crabtree, M.B., Kramer, L.D., Roehrig, J.T., 2002. Complete genome sequences and phylogenetic analysis of West Nile virus strains isolated from the United States, Europe, and the Middle East. Virology 298, 96–105. Li, S., Fang, M., Zhou, B., Ni, H., Shen, Q., Zhang, H., Han, Y., Yin, J., Chang, W., Xu, G., Cao, G., 2011. Simultaneous detection and differentiation of dengue virus serotypes 1-4, Japanese encephalitis virus, and West Nile virus by a combined reverse-transcription loop-mediated isothermal amplification assay. Virol. J. 8, 360. Mackenzie, J.S., Gubler, D.J., Petersen, L.R., 2004. Emerging flaviviruses: the spread and resurgence of Japanese encephalitis, West Nile and dengue viruses. Nat. Med. 10 (12 Suppl.), S98–S109. Mackenzie, J.S., Williams, D.T., 2009. The zoonotic flaviviruses of Southern, SouthEastern and Eastern Asia, and Australasia: the potential for emergent viruses. Zoonoses Public Hlth. 56, 338–356. Mann, R.A., Fegan, M., O‘Riley, K., Motha, J., Warner, S., 2013. Molecular characterization and phylogenetic analysis of Murray Valley encephalitis virus and West Nile virus (Kunjin subtype) from an arbovirus disease outbreak in horses in Victoria, Australia, in 2011. J. Vet. Diagn. Invest. 25, 35–44. Marshall, I.D., 1988. Murray Valley and Kunjin encephalitis. In: Monath, T.P. (Ed.), The Arboviruses: Epidemiology and Ecology. CRC Press, Boca Raton, FL, p. 151. Pyke, A.T., Smith, I.L., van den Hurk, A.F., Northill, J.A., Chuan, T.F., Westacott, A.J., Smith, G.A., 2004. Detection of Australasian Flavivirus encephalitic viruses using rapid fluorogenic TaqMan RT-PCR assays. J. Virol. Methods 117, 161–167. Quan, P.L., Williams, D.T., Johansen, C.A., Jain, K., Petrosov, A., Diviney, S.M., Tashmukhamedova, A., Hutchison, S.K., Tesh, R.B., Mackenzie, J.S., Briese, T., Lipkin,

W.I., 2011. Genetic characterization of K13965, a strain of Oak Vale virus from Western Australia. Virus Res. 160, 206–213. Roche, S.E., Wicks, R., Garner, M.G., East, I.J., Paskin, R., Moloney, B.J., Carr, M., Kirkland, P., 2012. Descriptive overview of the 2011 epidemic of arboviral disease in horses in Australia. Aust. Vet. J. 91, 5–13. Sanchez-Seco, M.P., Rosario, D., Domingo, C., Hernandez, L., Valdes, K., Guzman, M.G., Tenorio, A., 2005. Generic RT-nested-PCR for detection of flaviviruses using degenerated primers and internal control followed by sequencing for specific identification. J. Virol. Methods 126, 101–109. Savini, G., Monaco, F., Terregino, C., Di Gennaro, A., Bano, L., Pinoni, C., De Nardi, R., Bonilauri, P., Pecorari, M., Di Gialleonardo, L., Bonfanti, L., Polci, A., Calistri, P., Lelli, R., 2011. Usutu virus in Italy: an emergence or a silent infection? Vet. Microbiol. 151, 264–274. Scherret, J.H., Poidinger, M., Mackenzie, J.S., Broom, A.K., Deubel, V., Lipkin, W.I., Briese, T., Gould, E.A., Hall, R.A., 2001. The relationships between West Nile and Kunjin viruses. Emerg. Infect. Dis. 7, 697–705. Smith, D.W., Speers, D.J., Mackenzie, J.S., 2011. The viruses of Australia and the risk to tourists. Travel Med. Infect. Dis. 9, 113–125. Spencer, J.D., Azoulas, J., Broom, A.K., Buick, T.D., Currie, B., Daniels, P.W., Doggett, S.L., Hapgood, G.D., Jarrett, P.J., Lindsay, M.D., Lloyd, G., Mackenzie, J.S., Merianos, A., Moran, R.J., Ritchie, S.A., Russell, R.C., Smith, D.W., Stenhouse, F.O., Whelan, P.I., 2001. Murray Valley encephalitis virus surveillance and control initiatives in Australia. National Arbovirus Advisory Committee of the Communicable Diseases Network Australia. Commun. Dis. Intell. Q Rep. 25, 33–47. Tang, Y.L., Hapip, C.A., Liu, B., Fang, C.T., 2006. Highly sensitive TaqMan RT-PCR assay for detection and quantification of both lineages of West Nile virus RNA. J. Clin. Virol. 36, 177–182. Yang, Y., Wang, J., Wen, H., Liu, H., 2012. Comparison of two suspension arrays for simultaneous detection of five biothreat bacterial in powder samples. J. Biomed. Biotechnol. 2012, 831052, http://dx.doi.org/10.1155/2012/831052.