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    (A) Collection sites within the Australasian region and (B) collection sites in the Torres Strait and southern Papua New Guinea.

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    Sensitivity plots (mean ± SE) of oligonucleotide sets for real-time TaqMan polymerase chain reaction (PCR) detection of DNA isolated from (A) Ae. aegypti, (B) Ae. albopictus, and (C) Ae. scutellaris. Serial 10-fold dilutions of DNA (10 ng/μL initial concentration) were tested in triplicate and amplified simultaneously. Amplification plots for only one specimen of each species are shown.

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    Regression plots used to determine the limit of detection and amplification efficiencies of the (A) Ae. aegypti, (B) Ae. albopictus, and (C) Ae. scutellaris TaqMan assays. Serial 10-fold dilutions of DNA (10 ng/μL initial concentration) were tested in triplicate and amplified simultaneously. Regression plots for only one specimen of each species are shown.

  • 1

    Benedict MQ, Levine RS, Hawley WA, Lounibos LP, 2007. Spread of the tiger: global risk of invasion by the mosquito Aedes albopictus. Vector Borne Zoonotic Dis 7 :76–85.

    • Search Google Scholar
    • Export Citation
  • 2

    Gratz NG, 2004. Critical review of the vector status of Aedes albopictus. Med Vet Entomol 18 :215–227.

  • 3

    Almeida AP, Baptista SS, Sousa CA, Novo MT, Ramos HC, Panella NA, Godsey M, Simões MJ, Anselmo ML, Komar N, Mitchell CJ, Ribeiro H, 2005. Bioecology and vectorial capacity of Aedes albopictus (Diptera: Culicidae) in Macao, China, in relation to dengue virus transmission. J Med Entomol 42 :419–428.

    • Search Google Scholar
    • Export Citation
  • 4

    Effler PV, Pang L, Kitsutani P, Vorndam V, Nakata M, Ayers T, Elm J, Tom T, Reiter P, Rigau-Perez JG, Hayes JM, Mills K, Napier M, Clark GG, Gubler DJ, 2005. Dengue fever, Hawaii, 2001–2002. Emerg Infect Dis 11 :742–749.

    • Search Google Scholar
    • Export Citation
  • 5

    Reiter P, Fontenille D, Paupy C, 2006. Aedes albopictus as an epidemic vector of chikungunya virus: another emerging problem? Lancet Infect Dis 6 :463–464.

    • Search Google Scholar
    • Export Citation
  • 6

    Vazeille M, Moutailler S, Coudrier D, Rousseaux C, Khun H, Huerre M, Thiria J, Dehecq JS, Fontenille D, Schuffenecker I, Despres P, Failloux AB, 2007. Two Chikungunya isolates from the outbreak of La Reunion (Indian Ocean) exhibit different patterns of infection in the mosquito, Aedes albopictus. PLoS ONE 2 :e1168.

    • Search Google Scholar
    • Export Citation
  • 7

    Rezza G, Nicoletti L, Angelini R, Romi R, Finarelli AC, Panning M, Cordioli P, Fortuna C, Boros S, Magurano F, Silvi G, Angelini P, Dottori M, Ciufolini MG, Majori GC, Cassone A, 2007. Infection with chikungunya virus in Italy: an outbreak in a temperate region. Lancet 370 :1840–1846.

    • Search Google Scholar
    • Export Citation
  • 8

    Whelan P, Hapgood G, 2001. A mosquito survey of Dili, East Timor, and implications for disease control. Arbovirus Res Aust 8 :405–416.

  • 9

    Schoenig E, 1972. Distribution of 3 species of Aedes (Stegomyia) carriers of virus diseases on the main island of Papua and New Guinea. Philipp Sci 9 :61–82.

    • Search Google Scholar
    • Export Citation
  • 10

    Kay BH, Prakash G, Andre RG, 1995. Aedes albopictus and Aedes (Stegomyia) species in Fiji. J Am Mosq Control Assoc 11 :230–234.

  • 11

    Elliot SA, 1980. Aedes albopictus in the Solomon and Santa Cruz islands, South Pacific. Trans R Soc Trop Med Hyg 74 :747–748.

  • 12

    Cooper RD, Waterson DGE, Kupo M, Sweeney AW, 1994. Aedes albopictus (Skuse) (Diptera: Culicidae) in the Western Province of Papua New Guinea and the threat of its introduction to Australia. J Aust Entomol Assoc 33 :115–116.

    • Search Google Scholar
    • Export Citation
  • 13

    Russell RC, Williams CR, Sutherst RW, Ritchie SA, 2005. Aedes (Stegomyia) albopictus—a dengue threat for southern Australia. Comm Dis Intell 29 :296–298.

    • Search Google Scholar
    • Export Citation
  • 14

    Mitchell CJ, Gubler DJ, 1987. Vector competence of geographic strains of Aedes albopictus and Aedes polynesiensis and certain other Aedes (Stegomyia) mosquitoes for Ross River virus. J Am Mosq Control Assoc 3 :142–147.

    • Search Google Scholar
    • Export Citation
  • 15

    Ritchie SA, Moore P, Carruthers M, Williams C, Montgomery B, Foley P, Ahboo S, van den Hurk AF, Lindsay MD, Cooper B, Beebe N, Russell RC, 2006. Discovery of a widespread infestation of Aedes albopictus in the Torres Strait, Australia. J Am Mosq Control Assoc 22 :358–365.

    • Search Google Scholar
    • Export Citation
  • 16

    Rai KS, Pashley DP, Munstermann LE, 1982. Genetics of speciation in Aedine mosquitoes. Steiner WM, Tabachnick WJ, Rai KS, Narang S, eds. Recent Developments in the Genetics of Insect Disease Vectors. Champaign, Illinois: Stipes Publishers, 84–129.

  • 17

    Lee DJ, Hicks MM, Griffiths M, Debenham ML, Bryan JH, Russell RC, Geary M, Marks EN, 1987. The Culicidae of the Australasian Region, Vol. 4. Entomology Monograph No. 2. Canberra: Australian Government Publishing Service Press.

  • 18

    Huang YM, 1972. The subgenus Stegomyia of Aedes in Southeast Asia. I—The Scutellaris group of species. Contrib Am Entomol Inst 9 :1–109.

    • Search Google Scholar
    • Export Citation
  • 19

    Lamche GD, Whelan PI, 2003. Variability of larval identification characters of exotic Aedes albopictus (Skuse) intercepted in Darwin, Northern Territory. Comm Dis Intell 27 :105–109.

    • Search Google Scholar
    • Export Citation
  • 20

    Beebe NW, Whelan PI, van den Hurk AF, Ritchie SA, Corcoran S, Cooper RD, 2007. A polymerase chain reaction-based diagnostic to identify larvae and eggs of container mosquito species from the Australian region. J Med Entomol 44 :376–380.

    • Search Google Scholar
    • Export Citation
  • 21

    Sinclair D, 1992. The distribution of Aedes aegypti and dengue in Queensland, 1990–June 30, 1992. Arbovirus Res Aust 6 :323.

  • 22

    Beebe NW, Whelan PI, van den Hurk A, Ritchie SA, Cooper RD, 2005. Genetic diversity of the dengue vector Aedes aegypti in Australia and implications for future surveillance and mainland incursion monitoring. Comm Dis Intell 29 :299–304.

    • Search Google Scholar
    • Export Citation
  • 23

    Beebe NW, Cooper RD, Foley DH, Ellis JT, 2000. Populations of the south-west Pacific malaria vector Anopheles farauti s.s. revealed by ribosomal DNA transcribed spacer polymorphisms. Heredity 84 :244–253.

    • Search Google Scholar
    • Export Citation
  • 24

    Heid CA, Stevens J, Livak KJ, Williams PM, 1996. Real time quantitative PCR. Genome Res 6 :986–994.

  • 25

    Livak KJ, Flood SJA, Marmaro J, Giusti W, Deetz K, 1995. Oligonucleotides with fluorescent dyes at opposite ends provide a quenched probe system useful for detecting PCR product and nucleic acid hybridization. PCR Methods Appl 4 :357–362.

    • Search Google Scholar
    • Export Citation
  • 26

    Smith G, Smith I, Harrower B, Warrilow D, Bletchly C, 2006. A simple method for preparing synthetic controls for conventional and real-time PCR for the identification of endemic and exotic disease agents. J Virol Methods 135 :229–234.

    • Search Google Scholar
    • Export Citation
  • 27

    Rutledge RG, Côté C, 2003. Mathematics of quantitative kinetic PCR and the application of standard curves. Nucleic Acids Res 31 :e93.

  • 28

    Bass C, Williamson MS, Wilding CS, Donnelly MJ, Field LM, 2007. Identification of the main malaria vectors in the Anopheles gambiae species complex using a TaqMan real-time PCR assay. Malar J 6 :155.

    • Search Google Scholar
    • Export Citation
  • 29

    Marks EN, 1980. Mosquitoes (Diptera: Culicidae) of Cape York Peninsula, Australia. Stevens NC, Bailey A, eds. Contemporary Cape York Peninsula. Brisbane: The Royal Society of Queensland, 59–76.

  • 30

    Sanogo YO, Kim CH, Lampman R, Novak RJ, 2007. A real-time TaqMan polymerase chain reaction for the identification of Culex vectors of West Nile and Saint Louis encephalitis viruses in North America. Am J Trop Med Hyg 77 :58–66.

    • Search Google Scholar
    • Export Citation

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

Rapid Identification of Aedes albopictus, Aedes scutellaris, and Aedes aegypti Life Stages Using Real-time Polymerase Chain Reaction Assays

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  • 1 School of Molecular and Microbial Sciences, University of Queensland, Brisbane, Queensland, Australia; Tropical Population Health Unit Network, Queensland Health, Cairns, Queensland, Australia; Medical Entomology, Centre for Disease Control, Department of Health and Community Services, Darwin, Northern Territory, Australia; Australian Army Malaria Institute, Gallipoli Barracks, Enoggera, Queensland, Australia; Virology, Forensic and Scientific Services, Queensland Health, Coopers Plains, Queensland, Australia

In 2005, a widespread infestation of Aedes albopictus was discovered in the Torres Strait, the region between northern Australia and New Guinea. To contain this species, an eradication program was implemented in 2006. However, the progress of this program is impeded by the difficulty of morphologically separating Ae. albopictus larvae from the endemic species Aedes scutellaris. In this study, three real-time TaqMan polymerase chain reaction assays that target the ribosomal internal transcribed spacer 1 region were developed to rapidly identify Aedes aegypti, Ae. albopictus, and Ae. scutellaris from northern Australia. Individual eggs, larvae, pupae, and adults, as well as the species composition of mixed pools were accurately identified. The assay method was validated using 703 field-collected specimens from the Torres Strait.

INTRODUCTION

Aedes albopictus is a highly invasive mosquito species that has dramatically expanded from its native range in southeast Asia and become established in North and South America, southern Europe, Africa, and islands of the Pacific.1,2 It is susceptible to infection with at least 22 arboviruses,2 and has been the principle vector of dengue viruses (DENVs) during outbreaks in Hawaii and Macao.3,4 More recently, Ae. albopictus has exhibited enhanced vector competence for the recently emerged strain of chikungunya virus (CHIKV) that caused a major epidemic on La Réunion island in 2005–2006.5,6 It was also the vector during a CHIKV outbreak in the province of Ravenna, Italy in 2007, the first on continental Europe.7

This species is widely distributed in the Australasian region, being reported from Papua New Guinea, the Solomon Islands, Fiji, and East Timor.812 Despite the close proximity of Australia to these countries, and the numerous interceptions by quarantine personnel at international seaports, Ae. albopictus has not become established on the Australian mainland.13 There is concern that if Ae. albopictus were to be introduced onto the Australian mainland, it could rapidly colonize Australian towns and cities, including southern areas where Aedes aegypti does not occur, increasing the possibility of both DENV and CHIKV transmission in these centers. Furthermore, some strains of Ae. albopictus have been shown to be competent laboratory vectors of Ross River virus,14 and the relative role that this species could play in the transmission of other endemic Australian arboviruses is largely unknown.

In 2005, a widespread infestation of Ae. albopictus was discovered on the islands of the Torres Strait, the region that separates mainland Australia from the island of New Guinea.15 The original 2005 surveys found Ae. albopictus on 10 islands, but more recent data has confirmed that its distribution has expanded to 13 islands (J. Davis and G. Hapgood, unpublished data). In response to this discovery and with a view to preventing the spread of this mosquito to the mainland, Queensland Health and the Australian Government Department of Health and Ageing funded the Aedes albopictus Eradication Program (AAEP). This campaign consists of a control team, which undertakes larval habitat reduction, treating containers, adults and larvae with synthetic pyrethroids, and treatment of larger containers with s-methoprene; and a surveillance team, which undertakes pre-and post-treatment assessment of the control operations and the monitoring of uninfested islands.

Aedes albopictus is a member of the Scutellaris group of the Aedes (Stegomyia) subgenus that contains over 40 species.16 Two species have been described from this group in Australia: Aedes katherinensis, which is restricted to northern parts of the mainland, and Aedes scutellaris, which has a more regional distribution encompassing Cape York Peninsula, the Torres Strait, and the Papua and Moluccas regions of Indonesia.17 Although readily identified in the adult stage, larvae of Ae. albopictus are difficult to morphologically separate from Ae. scutellaris, with the distinguishing character being the length and number of branches of hair 1–VII,18 a character that exhibits some variability within and between species.19 Furthermore, eggs, early instar larvae, and pupae of Scutellaris group mosquitoes, as well as other container inhabiting Aedes, are not readily identified using morphologic features.

The difficulty in separating Ae. albopictus larvae from Ae. scutellaris in the Torres Strait has compromised the surveillance component of the AAEP. In an attempt to resolve this issue, larval specimens are sent to the Queensland Health Forensic and Scientific Services (QHFSS), Brisbane, for identification using a polymerase chain reaction-restriction fragment length polymorphism (PCR-RFLP) procedure developed by Beebe and others.20 This assay involves the amplification of the ITS1 region followed by digestion with RsaI to create restriction fragment-length polymorphism profiles for common endemic Australian container Aedes and Culex species. Although this method has proved successful, the laborious nature of the assay does not make it an ideal system for the rapid throughput of a large number of specimens. Consequently, we report the development of real-time TaqMan PCR assays for the rapid identification of Ae. albopictus and Ae. scutellaris. Furthermore, because Ae. aegypti is endemic in northern Queensland,21 and is occasionally intercepted outside its known geographic distribution,22 assays for this species were also developed. The assays we describe in the current study are able to identify all life stages, including eggs and first instar larvae. Other features of these assays include successful identification of specimens without the need for DNA extraction and in mixed pools of larvae or eggs. Evaluation of the TaqMan assays was then undertaken on specimens collected from the Torres Strait during AAEP surveillance and control operations.

MATERIALS AND METHODS

Mosquito material

Reference specimens of container inhabiting species were collected from various sites in Queensland, the Northern Territory, New South Wales, and Papua New Guinea (Table 1, Figure 1). Additional reference material was obtained from a laboratory colony of Ae. albopictus maintained at QHFSS that was established from eggs collected from Masig Island, Torres Strait, in 2005, and an Ae. aegypti colony housed at the Australian Army Malaria Institute, which was established from larvae collected from Townsville, Queensland in 2001. Mosquito specimens used for assay validation were identified using either morphologic characteristics18 or PCR-RFLP.20 Evaluation of the real-time assays was undertaken on Torres Strait field specimens collected during the surveillance monitoring by the AAEP in 2007. All operational larval specimens identified as belonging to the Scutellaris group, as well as pupae and early instar larvae of other species, were preserved in 70% ethanol and forwarded to QHFSS for molecular identification.

Nucleic acid isolation

Genomic DNA of mosquitoes was recovered using the procedure described by Beebe and others.22 Furthermore, a rapid DNA isolation method was separately developed as follows: individual mosquito specimens were placed in microfuge tubes containing 200 μL of Tris-EDTA (TE) buffer, before homogenization with a sterile pestle and centrifugation for 10 minutes at 14,000 rpm. Supernatants were retained and the pellets were discarded. The DNA was stored at −20°C to await analysis.

ITS1 PCR amplification and sequencing

The ITS1 region of the species listed in Table 1 was amplified and sequenced using the primers ITS1A and ITS1B.23 The PCR amplification reactions were performed in a total volume of 50 μL, including 1.5 mM MgCl2, 0.2 mM dNTPs, 1 × PCR buffer, 1U of AmpliTaq Gold DNA polymerase (Applied Biosystems, Foster City, CA), 800 nM of each primer and 1–10 ng of template DNA. Cycling conditions were as follows: one cycle of 94°C for 10 min, 35 cycles of 94°C for 30 seconds, 51°C for 40 seconds, and 72°C for 30 seconds, after which products were held at 4°C. The PCR amplicons were separated by electrophoresis on a 1% agarose gel and visualized under ultraviolet (UV) light using ethidium bromide staining. Products were then gel purified using the QIAquick Gel Extraction Kit (QIAGEN, Doncaster, VIC, Australia). Nucleotide sequencing of the purified DNA was performed using the Big Dye Terminator version 3.1 Cycle Sequencing Kit (Applied Biosystems, Foster City, CA). With the assistance of the Griffith University Sequencing Facility (GUDSF), the sequence data was then aligned and edited using the computer programs ContigExpress and AlignX (Vector NTI Suite, Invitrogen, Carlsbad, CA).

Real-time PCR primer and probe design

Three sets of TaqMan primers and probes specific for Ae. aegypti, Ae. albopictus, and Ae. scutellaris were designed using Primer Express Software version 2.0 (Applied Biosystems, Foster City, CA), based on the alignment of ITS1 sequences of Ae. aegypti and Ae. albopictus obtained from GenBank (accession nos. M95126 and AB231675, respectively), and sequences obtained in our laboratory (Table 2). Each oligonucleotide was assessed for its specificity using a BlastN search of GenBank sequences (NIH, Bethesda, MD). All probes were dual-labeled with a 6-carboxy-fluorescein (FAM) reporter group at the 5′ end and a 6-carboxy-tetramethyl-rhodamine (TAMRA) quencher group at the 3′ end.24,25 Primers were supplied by GeneWorks (Adelaide, SA, Australia) and probes were synthesized by Sigma-Proligo (Singapore).

Real-time PCR amplifications

Real-time PCR amplifications were performed in 96-well 0.1 mL MicroAmp plates (Applied Biosystems, Foster City, CA) in a 20 μL volume, containing 1 × TaqMan Fast Universal PCR Master Mix (Applied Biosystems, Foster City, CA), 300 nM each primer, 100 nM probe, and 0.1–20 ng of DNA template. Thermal cycling was performed on an ABI 7500 Sequence Detection System (Applied Biosystems, Foster City, CA) using the following conditions: one cycle of 95°C for 20 seconds, followed by 40 cycles of 95°C for 3 seconds, and 60°C for 30 seconds. Data was recorded and analyzed using the 7500 Fast SDS System software (Applied Biosystems, Foster City, CA).

Synthetic primer and probe controls

Synthetic primer and probe controls for each TaqMan assay were produced as described by Smith and others,26 eliminating the need for positive mosquito DNA controls (Table 3). The probe control (GeneWorks, Adelaide, SA, Australia) encompassed the specific TaqMan probe sequence of the target species flanked by forward and reverse primer sequences of rodent glyceraldehyde-3-phosphate dehydrogenase (rGAPDH): 5′-[rGAPDH fwd]-[target species probe]-[rGAPDH rev]-3′. Conversely, the primer control (GeneWorks) consisted of the rGAPDH probe sequence flanked by the primers of the target species: 5′-[target species fwd]-[rGAPDH probe]-[target species rev]-3′. Each synthetic control was titrated individually to obtain an optimal working dilution by testing 2 μL of 10-fold dilutions of each primer or probe in triplicate.

Validation of real-time PCR assays

Two individual fourth instar larvae were used to evaluate the sensitivity of each assay. The concentration of extracted genomic DNA from each Ae. aegypti, Ae. albopictus, and Ae. scutellaris specimen was first quantified using a NanoDrop ND-1000 Spectrophotometer (NanoDrop Technologies, Wilmington, DE) so that samples containing an initial concentration of 10 ng/μL of DNA could be used to prepare serial 10-fold dilutions. Each of the dilutions was then tested in triplicate using the real-time PCR conditions described previously. The limit of detection was determined to be the lowest concentration of DNA with detectable fluorescence above the cycle threshold (Ct) prior to cycle 40.

Specificity of the TaqMan primer and probe sets was assessed by testing a minimum of 5 individual specimens of each of the species listed in Table 1. In addition, intraspecific and interspecific variation between geographic populations was also analyzed when samples from different locations were available. To account for variability in detection rates and potential false negatives due to unknown DNA concentration in undiluted samples, the majority of these reference samples were also tested at 10−1 or 10−2 dilutions.

Furthermore, the real-time PCR assays designed to detect Ae. aegypti and Ae. albopictus were tested using ≥ 5 specimens from each of the following life stages: egg, first instar larva, fourth instar larva, pupa, and adult. For Ae. scutellaris, only fourth instar larvae and adults were available for testing. Finally, DNA from different ratios of Ae. aegypti and Ae. albopictus fourth instar larvae or eggs in pools of 5 individuals was isolated using the rapid isolation method and the method of Beebe and others,22 respectively, and tested at the 10−1 dilution using the respective TaqMan assays.

Assessment of the real-time PCR assays using field collected samples

Larvae and pupae stored in 100% ethanol were forwarded to QHFSS for identification using the TaqMan assays. The DNA was isolated using the rapid method described previously and ≥ 3 specimens per larval container were analyzed, depending on the number of larvae or pupae submitted per container.

Evaluation of FTA cards and filter paper for sample preservation

Because the carriage of ethanol is restricted on aircraft in the Torres Strait, we tested larvae that were stored dry on FTA cards or filter paper as alternative methods of sample preservation. Whole live fourth instar larvae of Ae. aegypti and Ae. albopictus were smeared onto FTA Classic Cards (Whatman, Kent, United Kingdom) and Whatman No. 1 filter paper (Whatman, Kent, United Kingdom) using a wooden applicator stick. Samples were allowed to air dry before being stored in an envelope at 23°C. On days 7, 14, and 21, the piece of card or paper containing the smear was placed in 0.5 mL TE buffer and incubated at 4°C for 30 minutes. Samples of 4 μL of the undiluted eluate and additional 10−1 and 10−2 dilutions were added to the reaction mix and analyzed using the TaqMan assay as described previously.

RESULTS

PCR amplification and sequencing of ITS1

Amplification of the ITS1 region using the universal primers ITS1A and ITS1B generated PCR products of ~600 bp from Ae. scutellaris and Ae. katherinensis; 700 bp from Ae. albopictus, Ae. aegypti, Ae. tremulus, Ae. notoscriptus, and Ae. palmarum; and 850 bp from Culex quinquefasciatus. A BlastN search using the ITS1 sequences of Ae. albopictus (accession nos. EU359684, EU359685, EU359686, EU359687, EU359688) and Ae. aegypti (EU359689) confirmed that they were homologous with published sequences of the ribosomal DNA genes of Ae. albopictus (accession no. AB213675) and Ae. aegypti (accession no. M95126), respectively. These sequences, as well as Ae. scutellaris (EU359690, EU359691), Ae. katherinensis (EU359692, EU359693), Ae. palmarum (EU359694, EU359695), Ae. notoscriptus (EU359696), and Cx. quinquefasciatus (EU359697) ITS1 sequences were deposited on GenBank. Unfortunately, the sequencing trace of all Ae. tremulus (N = 13) displayed secondary peaks, so reliable sequence data were not obtainable for this species.

Synthetic control development

Final working dilutions for the synthetic primer and probe controls for each real-time assay were determined from the dilutions in which fluorescence was detected between 24 and 28 cycles. From the initial 200 nM stock solution, a dilution of 10−9, which was equivalent to 2.0 pM, was selected for both the Ae. aegypti and Ae. scutellaris TaqMan primer and probe controls. For the Ae. albopictus TaqMan primer and probe controls, dilutions of 10−9 (equivalent to 2.0 pM) and 10−10 (equivalent to 200 fM) of stock solution were selected, respectively.

Sensitivity of real-time PCR assays

The sensitivity of each of the three real-time assays was assessed using 10-fold serial dilutions ranging from 10 ng/μL to 10 fg/μL of target DNA. From the amplification plots, the lowest concentration of DNA that was detected above the threshold for all replicates was 0.1 pg for all three species (Figure 2A–C). Regression analysis of the standard curves (Figure 3A–C) determined the theoretical minimum concentration of DNA able to be detected by cycle 40 to be 2.18 × 10−3 pg for Ae. aegypti, 7.62 × 10−3 pg for Ae. albopictus, and 7.60 × 10−3 pg for Ae. scutellaris. The regression slopes provided estimates of the amplification efficiencies for each assay. A slope of −3.33 is the number of cycles required for a 10-fold increase in amplicon concentration if the amplification efficiency of the reaction is 100%.27 Each assay demonstrated high real-time PCR efficiency rates: 95.9% for the Ae. aegypti assay, 101.6% for the Ae. albopictus assay, and 90.8% for the Ae. scutellaris assay, as well as high linearity, with a correlation coefficient (r2) > 0.99 for all three assays.

Specificity of real-time PCR assays

Because preliminary experiments demonstrated that the rapid isolation method in TE buffer provided sufficient template for analysis in the TaqMan assays, DNA from the majority of specimens was isolated using this procedure. Each TaqMan primer and probe set designed in this study was shown to be specific for its target species (Table 1). However, for < 1% of reactions using template DNA from non-target species, fluorescence was detected above the threshold, but at much later cycles (Ct > 31) compared with the range of detection of positive target samples (Ct < 28). Nearly all of the non-target species tested in each assay had at least one specimen exhibit these late curves. Subsequently, the real-time reactions were shortened to 30 cycles to eliminate these ambiguous results, and positive samples were defined as those that produced fluorescence at a Ct value of ≤ 28, which was one standard deviation above the mean Ct value. Samples with a Ct value > 28 were reanalyzed using a different dilution, different species–specific TaqMan assay, or the PCR-RFLP method, until their identification could be resolved.

Identification of different life stages and mixed pools

The TaqMan assays were able to identify all life stages tested, using either undiluted template or template at the 10−1 and 10−2 dilution (Table 4). Furthermore, mixed pools of Ae. aegypti and Ae. albopictus fourth instar larvae and eggs were detected using their respective TaqMan assays (Table 5). Some variability in the detection rate of Ae. aegypti eggs was observed, with some individual eggs not being detected until after 30 cycles, and eggs within mixed pools not being detected at all.

Assessment of real-time PCR assays using field collected samples

A total of 703 specimens, comprising 671 larvae, 29 pupae, 1 adult, and 2 damaged life stages, collected from 204 containers on 11 Torres Strait islands were analyzed using the Ae. albopictus and Ae. scutellaris TaqMan assays (Table 6). The TaqMan assays identified 629 (89.5%) of the larvae and or pupae, consisting of 65% and 35% Ae. albopictus and Ae. scutellaris, respectively. Importantly, no additional Ae. albopictus or Ae. scutellaris were detected using the PCR-RFLP. Of the specimens identified using the PCR-RFLP, 5 were Ae. notoscriptus and 1 was Ae. aegypti. The remaining 68 specimens could not be amplified using the PCR-RFLP.

Evaluation of FTA cards for sample preservation

With the exception of samples tested undiluted from FTA cards, all Ae. aegypti and Ae. albopictus fourth instar larvae could be detected using the respective TaqMan assays stored on FTA cards and filter paper for ≤ 21 days (Table 7).

DISCUSSION

On the basis of its role in arbovirus transmission cycles and its pestiferous nature, the discovery of Ae. albopictus in the Torres Strait is of particular concern to Australian quarantine and health authorities alike. The implementation of the eradication program in the Torres Strait has highlighted the difficulty in accurately separating fourth instar larvae of Ae. albopictus and Ae. scutellaris, the life stage most often collected during container surveys. Furthermore, it is also difficult to morphologically distinguish early instar larvae, pupae, and eggs of container-inhabiting Aedes, all of which can only be identified by expert taxonomists. Although used for routine identification of samples from the Torres Strait, molecular assays can also be used to identify Ae. albopictus and Ae. aegypti intercepted during quarantine operations at other Australian seaports and airports or during routine surveillance undertaken by regional health authorities.22 The development of molecular methods of identification means that eggs removed from ovitraps do not have to be reared to adult or fourth instar larva for identification, which can take ≤ 10 days. The PCR-RFLP assay, while being able to identify common container inhabiting species, is laborious and has not proven ideal when used for large scale operational identification of specimens collected by the AAEP.

The real-time TaqMan PCR assays we describe in the current study have facilitated the rapid identification of larval specimens submitted to QHFSS. Importantly, we have calculated that following rapid DNA isolation, 22 specimens can be analyzed in ~70 minutes with the TaqMan PCR assays, which compares favorably against the 7 hours it takes to complete the PCR-RFLP for the same number of samples. For the TaqMan assays, this comprises 15 minutes to load the samples, 40 minutes for the PCR reaction using the fast chemistry, and 15 minutes for analysis. In contrast, the PCR-RFLP requires 3 hours for amplification, followed by ≥ 1 hour for restriction digest, ≥ 2 hours for size separation of digested products and analysis, and > 1 hour for all sample loading. Processing time for the TaqMan assays could be reduced further by analyzing specimens in pools, especially when only information on the presence or absence of a species is required. An alternative way to reduce processing time could be to develop a multiplexed TaqMan assay, as has been described for members of the Anopheles gambiae species complex.28

The inclusion of synthetic primer and probe controls eliminates the need for positive mosquito DNA controls and together with the fact that the assay is performed using a closed tube system, reduces the risk of sample contamination. Nonetheless, the detection of fluorescence during later cycles (Ct > 30 cycles) observed in some reactions with non-target species may have been caused by carry-over contamination from the high concentrations of DNA present in each extract and the high sensitivities of the assays. However, for a given specimen, late curves were not reproducible at a particular dilution, did not occur over the range of dilutions tested, or when the non-target specimens were tested on a run without specimens of the target species. Consequently, to eliminate these ambiguous results during routine identification, a cut-off Ct value of 28 was established for all three assays.

The Ae. albopictus and Ae. scutellaris assays used for the operational trial identified almost 90% of specimens. The PCR-RFLP was only able to identify six additional larvae, none of which were Ae. albopictus or Ae. scutellaris. It is possible that the DNA had degraded in the remaining samples or that inhibitors to the PCR reaction were present. Indeed, when the nucleic acids from a sub-sample of the negative samples were electrophorized, there was either no DNA present, or there was evidence of DNA degradation. Both these issues could have been the result of inadequate storage of samples in the field or the rapid procedure used to isolate DNA from these specimens. Improved quality DNA could be obtained by routinely undertaking the DNA isolation method of Beebe and others,22 although this method is more time consuming, taking ~4 hours to process 22 specimens. Therefore, each laboratory may have to determine an optimal compromise between detectability and labor. The inability to detect DNA in eggs using the Ae. aegypti assay could have been the result of incomplete homogenization of the eggs while isolating the DNA. Another method, involving boiling of the sample, may be required for isolation of DNA from eggs.20

We have demonstrated that larval specimens can be squashed straight onto filter paper or FTA cards. Once air-dried, the samples can be stored for at least 21 days, which is longer than the period of time that specimens are kept prior to being analyzed. This alternative method of preservation and storage of mosquito samples circumvents the necessity to store samples in 70% ethanol. This is especially important in the Torres Strait, where air travel is the only government-sanctioned means for the AAEP staff to travel between islands and where dangerous goods legislation limits the amount of ethanol that can be taken on flights. The inability to detect specimens processed undiluted from the FTA cards may be due to the inhibition of the PCR reaction caused by the presence of proprietary chemicals impregnated in the card.

We were unable to find suitable sequence variation in the ITS1 region to produce TaqMan primers and probes that would differentiate Ae. scutellaris from Ae. katherinensis. Beebe and others also observed that both species produced identical banding profiles after digestion with restriction enzymes, so separation of these two species using PCR-RFLP was not possible.20 As Ae. katherinensis does not occur in the Torres Strait, the inability to separate these two species using current molecular methods is not critical to the AAEP. In locations where Ae. scutellaris and Ae. katherinensis occur sympatrically, such as on the tip of Cape York Peninsula,29 separation of these species may be resolved by targeting a different gene sequence, such as the acetylcholinesterase gene (Ace2) used to identify North American Culex spp.30

Table 1

Mosquito specimens (larvae or adults) used for specificity testing of the TaqMan assays

TaqMan assay tested
Ae. aegyptiAe. albopictusAe. scutellaris
SpeciesCollection location*nResult‡nResultnResult
* Abbreviations for collection locations include: QLD, Queensland; TS, Torres Strait; PNG, Papua New Guinea; WP, West Papua; NT, Northern Territory; and NSW, New South Wales.
n = number of individual specimens tested by each assay.
‡ DET denotes the samples were detected; ND denotes that the samples were not detected.
Ae. aegyptiQLD, Cairns1DET1ND1ND
QLD, Daintree village3DET1ND1ND
QLD, Port Douglas3DET1ND1ND
QLD, Yorkey’s Knob3DET1ND1ND
TS, Mer Island3DET1ND1ND
TS, Poruma Island2DET2ND2ND
TS, Thursday Island3DET3ND3ND
Ae. albopictusPNG, Kulalae1ND4DET1ND
PNG, Lihir2ND6DET2ND
PNG, Mabudauan2ND5DET2ND
TS, Badu Island1ND1DET1ND
TS, Erub Island1ND1DET1ND
TS, Masig Island3ND8DET4ND
TS, Moa Island1ND1DET1ND
TS, Prince of Wales Island2ND2DET2ND
TS, Ugar Island1ND1DET1ND
TS, Warraber Island1ND1DET1ND
WP, Timika1ND3DET1ND
Ae. scutellarisTS, Badu Island1ND1ND1DET
TS, Boigu Island1ND1ND1DET
TS, Dauan Island1ND2ND1DET
TS, Erub Island1ND3ND3DET
TS, Moa Island1ND2ND2DET
TS, Saibai Island1ND2ND2DET
TS, Thursday Island3ND3ND1DET
TS, Ugar Island1ND2ND2DET
Ae. katherinensisNT, Alyangula2ND3ND3DET
NT, Kakadu5ND5ND5DET
Ae. tremulusNT, Darwin3ND4ND5ND
NT, Tennant Creek3ND3ND3ND
TS, Badu Island1ND1ND1ND
Ae. notoscriptusQLD, Brisbane2ND2ND3ND
TS, Moa Island2ND2ND2ND
TS, Prince of Wales Island2ND2ND2ND
Ae. palmarumQLD, Cairns5ND5ND5ND
Cx. quinquefasciatusQLD, Brisbane2ND2ND2ND
NSW, Sydney3ND3ND3ND
Table 2

Oligonucleotide primers and probes for real-time PCR assays

Real-time assayDescriptionName5′–3′ SequenceLength (bp)Tm (°C)*
* Tm = melting temperature (°C).
Ae. aegyptiForward primerITS1_F338CGCTCGGACGCTCGTAC1757.6
Reverse primerITS1_R427CTTCGAGCTTCGACGACACA2058.8
ProbeAegyITS1PFAM-CAGAACACGCCAGACACGTTCGTACG-TAMRA2669.2
Ae. albopictusForward primerITS1_F440GTCAGCAGGGCCGAACC1758.7
Reverse primerITS1_R510GACGACCCGCCACTTAGCT1958.9
ProbeAlboITS1PFAM-CAGGGCACATACGTCCGCTTTGGTT-TAMRA2569.3
Ae. scutellarisForward primerITS1_F330GACACGGCTTGCCACCA1758.7
Reverse primerITS1_R448TAAAACACTTCGCTTTCGCAGTT2359.1
ProbeScutITS1PFAM-CGTCCGGGCCAAGTCAATCCAG-TAMRA2268.5
Table 3

Synthetic primer and probe control DNA templates

Synthetic controls5′–3′ Sequence*Length (bp)
* rGAPDH primer and probe sequences are in bold font.
Aedes albopictus
    Primer controlGTCAGCAGGGCCGAACCACAGAAGACTGTGGATGGCCCCTCAAGCTAAGTGGCGGGTCGTC61
    Probe controlTGCACCACCAACTGCTTAG ACAGGGCACATACGTCCGCTTTGGTTAGAACATCATCCCTGCATCC65
Aedes aegypti
    Primer controlCGCTCGGACGCTCGTACACAGAAGACTGTGGATGGCCCCTC ATGTGTCGTCGAAGCTCGAAG62
    Probe controlTGCACCACCAACTGCTTAG ACAGAACACGCCAGACACGTTCGTACGAGAACATCATCCCTGCATCC66
Aedes scutellaris
    Primer controlGACACGGCTTGCCACCAACAGAAGACTGTGGATGGCCCCTCAAACTGCGAAAGCGAAGTGTTTTA65
    Probe controlTGCACCACCAACTGCTTAG ACGTCCGGGCCAAGTCAATCCAGAGAACATCATCCCTGCATCC62
Table 4

Mean ± SD Ct values for different life stages of reference specimens tested using the TaqMan assays at the 100 to 10−2 dilutions

Mean ± SD Ct value
AssayLife stagen10010−110− 2
* The mean ± SD Ct value for Ae. aegypti eggs at the 10−2 dilution was calculated for 4 from 5 specimens, as one egg was not detected.
Ae. aegyptiEgg527.5 ± 4.331.0 ± 4.732.5 ± 3.2*
First instar larva516.3 ± 0.620.1 ± 0.723.7 ± 0.6
Fourth instar larva520.9 ± 9.121.6 ± 2.424.9 ± 2.3
Pupa515.9 ± 3.918.5 ± 2.221.6 ± 1.1
Adult520.5 ± 4.822.6 ± 2.623.0 ± 1.2
Ae. albopictusEgg519.1 ± 4.422.4 ± 4.225.6 ± 4.1
First instar larva518.8 ± 0.422.0 ± 0.425.5 ± 0.1
Fourth instar larva818.4 ± 0.721.7 ± 0.724.7 ± 0.7
Pupa515.8 ± 0.619.2 ± 0.622.5 ± 0.7
Adult516.9 ± 1.520.0 ± 0.823.1 ± 0.6
Ae. scutellarisFourth instar larva715.6 ± 5.618.1 ± 2.322.1 ± 1.9
Adult614.9 ± 5.318.6 ± 2.022.7 ± 2.1
Table 5

Identification of mixed pools of Aedes aegypti and Aedes albopictus fourth instar larvae and eggs using the TaqMan assays

Ae. aegypti TaqManAe. albopictus TaqMan
Life stage*Species ratio†nMean + SD Ct valuenMean + SD Ct value
*DNA from larvae was extracted using the rapid isolation method; Egg DNA was isolated using the method of Beebe and others.22
†Ratio of individual Ae. aegypti to Ae. albopictus in mixed pools (pool size = 5).
n = number of pools detected/number tested.
§The mean ± SD Ct value for Ae. aegypti eggs was calculated only for pools that were detected.
Larvae4:13/311.7 ± 1.33/314.6 ± 0.7
3:23/312.0 ± 0.93/313.7 ± 0.5
2:33/312.5 ± 1.13/312.6 ± 0.1
1:43/313.7 ± 0.73/312.7 ± 0.5
Eggs§4:12/321.2 ± 0.33/322.7 ± 2.9
3:22/319.3 ± 1.73/319.9 ± 0.5
2:32/322.8 ± 0.53/321.3 ± 2.3
1:41/319.13/318.0 ± 0.3
Table 6

Operational assessment of the TaqMan assays using material (larvae and pupae) collected from the Torres Strait in 2007

Islandn containersn specimensn (%) identified by TaqManAe. albopictus TaqMan positive (%)Ae. scutellaris TaqMan positive (%)ITS1 PCR-RFLP identification*
* Number of real-time polymerase chain reaction (PCR) negative specimens identified by the ITS1 PCR-restriction fragment length polymorphism procedure.
Badu52317(73.9)17(100.0)0(0.0)2 Ae. notoscriptus
Erub54147124(84.4)70(56.5)54(43.5)0
Iama125251(98.1)0(0.0)51(100.0)0
Mabuiag156551(78.5)51(100.0)0(0.0)1 Ae. notoscriptus
Masig28109107(98.2)105(98.1)2(1.9)0
Mer168874(84.1)7(9.5)67(90.5)0
Moa (St. Pauls)82321(91.3)16(76.2)5(23.8)0
Poruma175149(96.1)49(100.0)0(0.0)2 Ae. notoscriptus
Waiben110(0.0)0(0.0)0(0.0)1 Ae. aegypti
Ugar266963(91.3)33(52.4)30(47.6)0
Warraber227572(96.0)61(84.7)11(15.3)0
Table 7

Mean ± SD TaqMan assay Ct values of 4th instar larvae of Aedes aegypti (N = 5) and Ae. albopictus (N = 4) stored on FTA cards and filter paper tested at the 100 to 10−2 dilutions

Ae. aegypti TaqManAe. albopictus TaqMan
Storage substrateDay10010−110−210010−110− 2
* ND denotes that the samples were not detected (Ct value > 40 cycles).
FTA card7ND*22.0 ± 0.923.4 ± 0.6ND21.9 ± 1.123.1 ± 0.5
14ND22.6 ± 1.022.5 ± 0.6ND22.3 ± 1.322.3 ± 1.0
21ND21.7 ± 0.723.2 ± 0.6ND22.3 ± 0.824.1 ± 0.3
Filter paper718.9 ± 1.520.9 ± 1.523.7 ± 1.717.8 ± 1.120.1 ± 0.322.7 ± 0.3
1419.6 ± 0.721.2 ± 0.723.8 ± 0.618.2 ± 0.919.9 ± 0.622.3 ± 0.6
2118.3 ± 1.020.6 ± 0.323.6 ± 0.319.1 ± 1.421.5 ± 1.524.4 ± 1.5
Figure 1.
Figure 1.

(A) Collection sites within the Australasian region and (B) collection sites in the Torres Strait and southern Papua New Guinea.

Citation: The American Journal of Tropical Medicine and Hygiene Am J Trop Med Hyg 79, 6; 10.4269/ajtmh.2008.79.866

Figure 2.
Figure 2.

Sensitivity plots (mean ± SE) of oligonucleotide sets for real-time TaqMan polymerase chain reaction (PCR) detection of DNA isolated from (A) Ae. aegypti, (B) Ae. albopictus, and (C) Ae. scutellaris. Serial 10-fold dilutions of DNA (10 ng/μL initial concentration) were tested in triplicate and amplified simultaneously. Amplification plots for only one specimen of each species are shown.

Citation: The American Journal of Tropical Medicine and Hygiene Am J Trop Med Hyg 79, 6; 10.4269/ajtmh.2008.79.866

Figure 3.
Figure 3.

Regression plots used to determine the limit of detection and amplification efficiencies of the (A) Ae. aegypti, (B) Ae. albopictus, and (C) Ae. scutellaris TaqMan assays. Serial 10-fold dilutions of DNA (10 ng/μL initial concentration) were tested in triplicate and amplified simultaneously. Regression plots for only one specimen of each species are shown.

Citation: The American Journal of Tropical Medicine and Hygiene Am J Trop Med Hyg 79, 6; 10.4269/ajtmh.2008.79.866

*

Address correspondence to Andrew F. van den Hurk, Virology, Queensland Health Forensic and Scientific Services, 39 Kessels Road, Coopers Plains, Queensland, 4108, Australia. E-mail: andrew_hurk@health.qld.gov.au

Authors’ addresses: Lydia A. Hill, Department of Microbiology and Parasitology, School of Molecular and Microbial Sciences, University of Queensland, Brisbane, Queensland, 4072, Australia. Joseph B. Davis, George Hapgood, and Scott A. Ritchie, Tropical Population Health Unit Network, Queensland Health, PO Box 1103, Cairns, Queensland, 4870, Australia, E-mails: joe_davis@health.qld.gov.au, george_hapgood@health.qld.gov.au, and scott_ritchie@health.qld.gov.au. Peter I. Whelan, Medical Entomology, Centre for Disease Control, Department of Health and Community Services, Darwin, Northern Territory, 0810 Australia, E-mail: Peter.Whelan@ nt.gov.au. Greg A. Smith and Andrew F. van den Hurk, Virology, Queensland Health Forensic and Scientific Services, 39 Kessels Rd., Coopers Plains, Queensland, 4108, Australia, E-mails: greg_smith@health.qld.gov.au and andrew_hurk@health.qld.gov.au. R. D. Cooper, Australian Army Malaria Institute, Gallipoli Barracks, Enoggera, Queensland, 4051, Australia, E-mail: Bob.Cooper@defence.gov.au.

Acknowledgments: The authors thank Nigel Beebe, Donna Mackenzie, Sonja Hall-Mendelin, Petrina Johnson, Russell Simmons, Judy Northill, and Ina Smith for advice and assistance with aspects of this study. We also thank Peter Ebbsworth and Esonke Waraung for collecting the larval samples from New Guinea. Finally, the authors thank Alyssa Pyke for reading a draft of the manuscript.

Financial Support: Funding for this study was provided by the Australian Biosecurity Cooperative Research Centre for Emerging Infectious Disease.

REFERENCES

  • 1

    Benedict MQ, Levine RS, Hawley WA, Lounibos LP, 2007. Spread of the tiger: global risk of invasion by the mosquito Aedes albopictus. Vector Borne Zoonotic Dis 7 :76–85.

    • Search Google Scholar
    • Export Citation
  • 2

    Gratz NG, 2004. Critical review of the vector status of Aedes albopictus. Med Vet Entomol 18 :215–227.

  • 3

    Almeida AP, Baptista SS, Sousa CA, Novo MT, Ramos HC, Panella NA, Godsey M, Simões MJ, Anselmo ML, Komar N, Mitchell CJ, Ribeiro H, 2005. Bioecology and vectorial capacity of Aedes albopictus (Diptera: Culicidae) in Macao, China, in relation to dengue virus transmission. J Med Entomol 42 :419–428.

    • Search Google Scholar
    • Export Citation
  • 4

    Effler PV, Pang L, Kitsutani P, Vorndam V, Nakata M, Ayers T, Elm J, Tom T, Reiter P, Rigau-Perez JG, Hayes JM, Mills K, Napier M, Clark GG, Gubler DJ, 2005. Dengue fever, Hawaii, 2001–2002. Emerg Infect Dis 11 :742–749.

    • Search Google Scholar
    • Export Citation
  • 5

    Reiter P, Fontenille D, Paupy C, 2006. Aedes albopictus as an epidemic vector of chikungunya virus: another emerging problem? Lancet Infect Dis 6 :463–464.

    • Search Google Scholar
    • Export Citation
  • 6

    Vazeille M, Moutailler S, Coudrier D, Rousseaux C, Khun H, Huerre M, Thiria J, Dehecq JS, Fontenille D, Schuffenecker I, Despres P, Failloux AB, 2007. Two Chikungunya isolates from the outbreak of La Reunion (Indian Ocean) exhibit different patterns of infection in the mosquito, Aedes albopictus. PLoS ONE 2 :e1168.

    • Search Google Scholar
    • Export Citation
  • 7

    Rezza G, Nicoletti L, Angelini R, Romi R, Finarelli AC, Panning M, Cordioli P, Fortuna C, Boros S, Magurano F, Silvi G, Angelini P, Dottori M, Ciufolini MG, Majori GC, Cassone A, 2007. Infection with chikungunya virus in Italy: an outbreak in a temperate region. Lancet 370 :1840–1846.

    • Search Google Scholar
    • Export Citation
  • 8

    Whelan P, Hapgood G, 2001. A mosquito survey of Dili, East Timor, and implications for disease control. Arbovirus Res Aust 8 :405–416.

  • 9

    Schoenig E, 1972. Distribution of 3 species of Aedes (Stegomyia) carriers of virus diseases on the main island of Papua and New Guinea. Philipp Sci 9 :61–82.

    • Search Google Scholar
    • Export Citation
  • 10

    Kay BH, Prakash G, Andre RG, 1995. Aedes albopictus and Aedes (Stegomyia) species in Fiji. J Am Mosq Control Assoc 11 :230–234.

  • 11

    Elliot SA, 1980. Aedes albopictus in the Solomon and Santa Cruz islands, South Pacific. Trans R Soc Trop Med Hyg 74 :747–748.

  • 12

    Cooper RD, Waterson DGE, Kupo M, Sweeney AW, 1994. Aedes albopictus (Skuse) (Diptera: Culicidae) in the Western Province of Papua New Guinea and the threat of its introduction to Australia. J Aust Entomol Assoc 33 :115–116.

    • Search Google Scholar
    • Export Citation
  • 13

    Russell RC, Williams CR, Sutherst RW, Ritchie SA, 2005. Aedes (Stegomyia) albopictus—a dengue threat for southern Australia. Comm Dis Intell 29 :296–298.

    • Search Google Scholar
    • Export Citation
  • 14

    Mitchell CJ, Gubler DJ, 1987. Vector competence of geographic strains of Aedes albopictus and Aedes polynesiensis and certain other Aedes (Stegomyia) mosquitoes for Ross River virus. J Am Mosq Control Assoc 3 :142–147.

    • Search Google Scholar
    • Export Citation
  • 15

    Ritchie SA, Moore P, Carruthers M, Williams C, Montgomery B, Foley P, Ahboo S, van den Hurk AF, Lindsay MD, Cooper B, Beebe N, Russell RC, 2006. Discovery of a widespread infestation of Aedes albopictus in the Torres Strait, Australia. J Am Mosq Control Assoc 22 :358–365.

    • Search Google Scholar
    • Export Citation
  • 16

    Rai KS, Pashley DP, Munstermann LE, 1982. Genetics of speciation in Aedine mosquitoes. Steiner WM, Tabachnick WJ, Rai KS, Narang S, eds. Recent Developments in the Genetics of Insect Disease Vectors. Champaign, Illinois: Stipes Publishers, 84–129.

  • 17

    Lee DJ, Hicks MM, Griffiths M, Debenham ML, Bryan JH, Russell RC, Geary M, Marks EN, 1987. The Culicidae of the Australasian Region, Vol. 4. Entomology Monograph No. 2. Canberra: Australian Government Publishing Service Press.

  • 18

    Huang YM, 1972. The subgenus Stegomyia of Aedes in Southeast Asia. I—The Scutellaris group of species. Contrib Am Entomol Inst 9 :1–109.

    • Search Google Scholar
    • Export Citation
  • 19

    Lamche GD, Whelan PI, 2003. Variability of larval identification characters of exotic Aedes albopictus (Skuse) intercepted in Darwin, Northern Territory. Comm Dis Intell 27 :105–109.

    • Search Google Scholar
    • Export Citation
  • 20

    Beebe NW, Whelan PI, van den Hurk AF, Ritchie SA, Corcoran S, Cooper RD, 2007. A polymerase chain reaction-based diagnostic to identify larvae and eggs of container mosquito species from the Australian region. J Med Entomol 44 :376–380.

    • Search Google Scholar
    • Export Citation
  • 21

    Sinclair D, 1992. The distribution of Aedes aegypti and dengue in Queensland, 1990–June 30, 1992. Arbovirus Res Aust 6 :323.

  • 22

    Beebe NW, Whelan PI, van den Hurk A, Ritchie SA, Cooper RD, 2005. Genetic diversity of the dengue vector Aedes aegypti in Australia and implications for future surveillance and mainland incursion monitoring. Comm Dis Intell 29 :299–304.

    • Search Google Scholar
    • Export Citation
  • 23

    Beebe NW, Cooper RD, Foley DH, Ellis JT, 2000. Populations of the south-west Pacific malaria vector Anopheles farauti s.s. revealed by ribosomal DNA transcribed spacer polymorphisms. Heredity 84 :244–253.

    • Search Google Scholar
    • Export Citation
  • 24

    Heid CA, Stevens J, Livak KJ, Williams PM, 1996. Real time quantitative PCR. Genome Res 6 :986–994.

  • 25

    Livak KJ, Flood SJA, Marmaro J, Giusti W, Deetz K, 1995. Oligonucleotides with fluorescent dyes at opposite ends provide a quenched probe system useful for detecting PCR product and nucleic acid hybridization. PCR Methods Appl 4 :357–362.

    • Search Google Scholar
    • Export Citation
  • 26

    Smith G, Smith I, Harrower B, Warrilow D, Bletchly C, 2006. A simple method for preparing synthetic controls for conventional and real-time PCR for the identification of endemic and exotic disease agents. J Virol Methods 135 :229–234.

    • Search Google Scholar
    • Export Citation
  • 27

    Rutledge RG, Côté C, 2003. Mathematics of quantitative kinetic PCR and the application of standard curves. Nucleic Acids Res 31 :e93.

  • 28

    Bass C, Williamson MS, Wilding CS, Donnelly MJ, Field LM, 2007. Identification of the main malaria vectors in the Anopheles gambiae species complex using a TaqMan real-time PCR assay. Malar J 6 :155.

    • Search Google Scholar
    • Export Citation
  • 29

    Marks EN, 1980. Mosquitoes (Diptera: Culicidae) of Cape York Peninsula, Australia. Stevens NC, Bailey A, eds. Contemporary Cape York Peninsula. Brisbane: The Royal Society of Queensland, 59–76.

  • 30

    Sanogo YO, Kim CH, Lampman R, Novak RJ, 2007. A real-time TaqMan polymerase chain reaction for the identification of Culex vectors of West Nile and Saint Louis encephalitis viruses in North America. Am J Trop Med Hyg 77 :58–66.

    • Search Google Scholar
    • Export Citation
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