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Rapid Surveillance for Vector Presence (RSVP): Development of a novel system for detecting Aedes aegypti and Aedes albopictus

Rapid Surveillance for Vector Presence (RSVP): Development of a novel system for detecting Aedes aegypti and Aedes albopictus

  • Brian L. Montgomery, 
  • Martin A. Shivas, 
  • Sonja Hall-Mendelin, 
  • Jim Edwards, 
  • Nicholas A. Hamilton, 
  • Cassie C. Jansen, 
  • Jamie L. McMahon, 
  • David Warrilow, 
  • Andrew F. van den Hurk
PLOS
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Abstract

Background

The globally important Zika, dengue and chikungunya viruses are primarily transmitted by the invasive mosquitoes, Aedes aegypti and Aedes albopictus. In Australia, there is an increasing risk that these species may invade highly urbanized regions and trigger outbreaks. We describe the development of a Rapid Surveillance for Vector Presence (RSVP) system to expedite presence- absence surveys for both species.

Methodology/Principal findings

We developed a methodology that uses molecular assays to efficiently screen pooled ovitrap (egg trap) samples for traces of target species ribosomal RNA. Firstly, specific real-time reverse transcription-polymerase chain reaction (RT-PCR) assays were developed which detect a single Ae. aegypti or Ae. albopictus first instar larva in samples containing 4,999 and 999 non-target mosquitoes, respectively. ImageJ software was evaluated as an automated egg counting tool using ovitrap collections obtained from Brisbane, Australia. Qualitative assessment of ovistrips was required prior to automation because ImageJ did not differentiate between Aedes eggs and other objects or contaminants on 44.5% of ovistrips assessed, thus compromising the accuracy of egg counts. As a proof of concept, the RSVP was evaluated in Brisbane, Rockhampton and Goomeri, locations where Ae. aegypti is considered absent, present, and at the margin of its range, respectively. In Brisbane, Ae. aegypti was not detected in 25 pools formed from 477 ovitraps, comprising ≈ 54,300 eggs. In Rockhampton, Ae. aegypti was detected in 4/6 pools derived from 45 ovitraps, comprising ≈ 1,700 eggs. In Goomeri, Ae. aegypti was detected in 5/8 pools derived from 62 ovitraps, comprising ≈ 4,200 eggs.

Conclusions/Significance

RSVP can rapidly detect nucleic acids from low numbers of target species within large samples of endemic species aggregated from multiple ovitraps. This screening capability facilitates deployment of ovitrap configurations of varying spatial scales, from a single residential block to entire suburbs or towns. RSVP is a powerful tool for surveillance of invasive Aedes spp., validation of species eradication and quality assurance for vector control operations implemented during disease outbreaks.

Author summary

Aedes (Stegomyia) vectors of dengue, Zika and chikungunya viruses utilize artificial and natural containers as larval habitats. Adults do not usually disperse far (< 500 m) from these larval habitats in urban and peri-urban environments. Highly heterogeneous distributions raise significant logistic challenges to conduct informative surveillance. Public health imperatives require contemporaneous vector mosquito presence-absence data for highly urbanized regions that are both vulnerable to invasions and have frequent exposure to viremic travellers. We developed a promising tool to expedite presence-absence surveillance of Aedes aegypti and Aedes albopictus by integrating molecular diagnostics with ovitraps and automated egg quantification software. The high sensitivity of the molecular assays enabled samples from multiple ovitraps to be pooled and processed for each diagnostic test. This innovation resolves the considerable logistic constraints inherent in traditional ovitrap surveillance programs. Proof of concept was evaluated in field trials in Queensland geographies where Ae. aegypti is considered either absent, present or at the margin of its range (Brisbane, Rockhampton and Goomeri, respectively). Aedes aegypti was detected in Goomeri and Rockhampton and not detected in Brisbane. Further investigation is required to address the inaccuracy of automated egg counting software whenever contaminants are present. RSVP can accommodate varied ovitrap designs and deployment configurations, improves efficiency in laboratory and labor costs for high volumes of samples, and enables a rapid turnaround of results. The RSVP system can innovate surveillance programs for early-warning of invasion, eradication, and quality assurance for vector control in disease response contexts.

Introduction

Aedes aegypti and Aedes albopictus are invasive mosquito species and global vectors of Zika (ZIKV) [1], dengue (DENVs) [2] and chikungunya (CHIKV) [3] viruses. Both species can coexist in the same ecological niche [4, 5] and share characteristics that are likely to make their detection difficult in the early stages of stochastic invasions, including heterogeneous distributions [68], limited dispersal capability of adults [913], and often low population densities [1416]. Ovipositing females cement eggs inside a wide variety of water-bearing containers and these eggs can resist desiccation for several months [17, 18]. Transport to new regions or countries can occur via the shipment of immature stages in freight, such as used tires [1921] and lucky bamboo [22, 23], or as adults sequestered in aircraft [24].

In Australia, quarantine authorities intercept both species at international seaports and airports [16, 25], with the frequency of detections increasing dramatically since 2012. However, there is a concurrent threat of range expansion from endemic Queensland populations. Aedes aegypti occurs throughout most of Queensland, although dengue outbreaks only occur in north Queensland [2629]. Aedes albopictus has not colonized the Australian mainland following the rapid invasion of island communities of the Torres Strait, north Queensland [16, 30], largely due to suppression programs at transport hubs [31].

Importantly, both species are considered to be absent from southeast (SE) Queensland (population 3.4 million), where ≈ 70% of the Queensland population reside. This status is not based on systematic entomological baseline monitoring, but rather on the lack of local disease transmission following the importation of ZIKV, DENV and CHIKV in viremic travellers, coupled with negative results from ad hoc larval surveys and trapping of peridomestic mosquitoes. Southeast Queensland is considered to be receptive to invasion by both species. Indeed, Ae. aegypti was present in this region until the mid-1950s [29], whilst predictive models indicate the eastern seaboard of Australia could be colonized by Ae. albopictus [25, 32]. Receptivity is considered to be increasing [33, 34], partly due to the recent proliferation of water-storage containers, such as rainwater tanks [35]. In the future, ineffective mosquito-proofing of rainwater tanks or rainwater harvesting structures may increase the number of available larval habitats [3638].

Contemporaneous baseline monitoring is essential to increase certainty that vector species are absent in geographies that are vulnerable to invasion [39] to minimize the risk of cryptic disease outbreaks. However, existing surveillance options for peridomestic mosquito species have operational weaknesses. Larval and pupal surveys are labor-intensive [4042] and can be compromised by inaccessible premises or larval habitats, cryptic containers [43, 44], or timing a survey when mosquito abundance is low [16]. In terms of adult traps, some designs (e.g., Biogents BG-Sentinel trap) are highly sensitive [45] but require electricity and incur significant procurement, servicing, and maintenance costs [46, 47]. Novel adult traps (BG-Gravid Aedes Trap) are cheaper but less sensitive [4850] and may be expensive to deploy in extensive arrays over a large spatial scale [6, 47, 51, 52]. Ovitraps provide a cheap and sensitive tool for presence-absence surveys of peridomestic species [17, 53, 54] but require an investment in time and laboratory resources to count eggs, rear and identify larvae [54, 55].

We report the development of a sensitive, easy-to-use, and cost-effective system, which we call Rapid Surveillance for Vector Presence (RSVP), to expedite presence-absence surveys of invasive Aedes species. The RSVP provides a powerful validation tool to confirm a species is absent at various spatial scales (property, suburban block, town or region), within an eradication program, and potentially as a quality assurance measure for vector control strategies implemented in response to ZIKV, DENV or CHIKV outbreaks.

Methods

Development and optimization of molecular assays to detect Ae. aegypti and Ae. albopictus in ovitrap samples

Eggs were hatched overnight by submerging ovistrips in 200 mL of Milli-Q water in rectangular 750 mL polyethylene containers, to which ≈ 10 mg of brain-heart powder was added to stimulate hatching. Ovistrips were removed and larvae extracted from the water via a vacuum filtration protocol. Specifically, the membrane and support pad of a MicroFunnel 300 filter funnel (Pall Life Sciences, Ann Arbor, MI, USA) was replaced with an FTA card (GE Healthcare Life Sciences, PA, USA) trimmed to fit the base of the funnel. The filter funnel was then placed on top of a vacuum flask. The water containing the larvae was gently agitated before it was poured into the cylinder of the filter funnel and a vacuum applied. The filter funnel and hatching container were rinsed with Milli-Q water until there were no larvae visible. The cards were then removed from the base of the funnel and air-dried overnight at room temperature.

The protocol of Hall-Mendelin et al. [56] was used to prepare the FTA cards and dried larvae for nucleic acid extraction. Briefly, the cards were cut into 4–5 strips and placed in a 5 mL vial containing 1 mL of Milli-Q water. Vials were placed on wet ice and vortexed every 5 min for 15 sec for a total of 20 min. The cards were then placed in a 5 mL syringe from which the plunger had been removed. The plunger was used to squeeze the liquid from the cards into a 2 mL vial. Nucleic acids were extracted from 140 μL of the eluate using the Bio Robot Universal System (Qiagen, Hilden, Germany) and the QIAamp Virus BioRobot MDx Kit (Qiagen, Clifton Hill, Australia) according to the manufacturer’s instructions.

Real-time TaqMan reverse transcription-polymerase chain reaction (RT-PCR) assays were used to detect target Ae. aegypti and Ae. albopictus against a background of endemic species. The Ae. aegypti ribosomal RNA was detected using real-time TaqMan RT-PCR with forward GCAGTCAGATGTTTGATACCGC and reverse GGTTGACGTATTATCAGGTCACACTA primers at 500 nM, and probe FAM-TGGGCGCCTCGGTGTCCCG-TAMRA at 300 nM. The Ae. albopictus ribosomal RNA was detected using forward CCGACAAGGCAATATGTC and reverse ACGCGTACGGACATTG primers at 300 nM, and probe FAM-TTCCCTCCGATCAGCGAACTC-TAMRA at 200 nM. The cycling conditions consisted of one cycle at 50°C for 5 min, one cycle at 95°C for 2 min, and 40 cycles at 95°C for 3 sec and 60°C for 30 sec. A positive result, indicating the presence of target species RNA, corresponded to cycle threshold values of ≤ 40 cycles. Reaction controls always included a no-template and synthetically generated template samples. The specificity of the 2 TaqMan RT-PCR assays was tested using 4th instar larval samples of Ae. aegypti and Ae. albopictus, as well as other peridomestic species, including Aedes katherinensis, Aedes notoscriptus, Aedes palmarum, Aedes scutellaris, Aedes tremulus and Culex quinquefasciatus.

Pools of varying sizes were produced to test the sensitivity of the TaqMan RT-PCR assays in detecting Ae. aegypti and Ae. albopictus. The Ae. aegypti eggs used to optimize the sensitivity of the assay were obtained from the F2-4 generations of a colony originating from Townsville, Australia, while Fo Ae. notoscriptus were collected in ovitraps deployed in Brisbane, Australia. The Ae. albopictus were from a colony established from eggs collected from Hammond Island (Torres Strait, Australia) and had been in colony for < 10 generations. Following hatching as described above, single 1st instar Ae. aegypti or Ae. albopictus larvae were added to batches of 1st instar Ae. notoscriptus or Ae. aegypti larvae, respectively, to produce pool sizes of 10, 100, 1,000 and 5,000 larvae. In addition, a pool comprised of a single Ae. aegypti or Ae. albopictus 1st instar larvae, and pools containing only Ae. notoscriptus or Ae. aegypti 1st instar larvae were produced. The pools were processed using the vacuum filtration method described above and Ae. aegypti and Ae. albopictus detected using the species-specific TaqMan RT-PCR assays.

Field validation of the RSVP as a tool for detection of Ae. aegypti

Study sites.

Three sites were chosen to represent the trichotomy of Ae. aegypti distribution in Queensland. Brisbane (27° 28’15.6”S, 153°1’24.4”E) is the state capital (> 1 million population; > 380,000 private dwellings) [57] within SE Queensland with no recent detections of Ae. aegypti. The city provided a large geographical area (> 1,367 km2) to test RSVP in a species absence context. Rockhampton (-23°22’49”S, 150°30’21”E) is a small regional city 636 km NW of Brisbane (80,000 population; > 32,000 private dwellings). Aedes aegypti is locally abundant and therefore provided a suitable site for field validation of species presence. Goomeri is a small town (< 500 population, > 200 private dwellings), 235 km NW of Brisbane and represents the ill-defined margin of Ae. aegypti distribution nearest SE Queensland. Aedes aegypti has previously been detected during infrequent larval surveys.

Trapping strategy.

Large ovitraps were assembled to maximize visual and olfactory cues [58, 59]. Black plastic buckets (9.3L) were provided with an overflow hole and exposed to the outdoor environment for 2 weeks prior to deployment, to remove volatile chemicals that may repel ovipositing females. An ‘ovistrip’ [60] (380 x 50 mm) made from white Tork Cleaning Cloth (SCA, Stockholm, Sweden) was affixed to the inside wall of each bucket with a fold-back clip. A pellet of compacted alfalfa was added to 3-L of tap water to produce an organic infusion [61]. Black plastic mesh covers (20 mm aperture) were fitted to prevent access by children or domestic animals.

Ovitrap sites included well-shaded urban yards, premises near imported dengue cases, commercial precincts and transport hubs (Fig 1). Ovitraps were deployed with occupier consent in outdoor sites that were adjacent to premises and protected from rain and wind [62]. Brisbane ovitraps were primarily deployed in a routine monitoring context from January to July 2015 and were collected after 2 weeks. Rockhampton ovitraps were deployed in May and June 2015, and were collected after 7 days. Goomeri ovitraps were deployed in February and March 2016 for periods ranging from 11–15 days. Ovistrips were transferred to the local office in individual sealable bags and dried overnight at room temperature. Brisbane ovistrips were pooled each fortnight, whilst additional ad hoc pools were submitted from suburbs where an imported dengue case was notified. Rockhampton ovistrips were pooled according to geographical clusters, or the suburb where an imported case of dengue was notified. Goomeri ovistrips were pooled per each trapping event. All ovistrips were forwarded to Forensic and Scientific Services, Department of Health, Queensland Government, Brisbane, for molecular identification.

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Fig 1. Spatial distribution of RSVP ovitraps (n = 477) in Brisbane deployed at commercial and residential dwellings (29 January- 9 July 2015).

The map layer was created using ArcGIS Online (https://www.arcgis.com/features/index.html).

https://doi.org/10.1371/journal.pntd.0005505.g001

Egg quantification using image analyses

We assessed ImageJ software [63] as a platform to count egg numbers on ovistrips in situ from Brisbane ovitraps deployed during the 2013–14 summer. Using a stereo microscope, each ovistrip was inspected, eggs counted and relative contamination assessed to ascertain whether the ovistrip was of sufficient quality to be analyzed by ImageJ (Fig 2). Quality of the ovistrips was especially important, as ImageJ cannot differentiate Aedes eggs from other objects or contaminants, such as debris, leaves, insect cadavers and/or fungal growth, on the ovistrip. This could potentially produce an inaccurate count of the eggs on the ovistrips. Consequently, an arbitrary grade for each ovistrip was assigned to determine suitability for ImageJ analysis and comprised suitable (Fig 2A), suitable with image manipulation (Fig 2B) and unsuitable (Fig 2C).

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Fig 2. Examples of ovistrip condition (n = 193) for submission to quantify the number of eggs by ImageJ analyses.

(A) Image suitable for immediate analysis; (B) image requires manipulation for image analysis; and (C) image unsuitable for analysis.

https://doi.org/10.1371/journal.pntd.0005505.g002

An image of each ovistrip was then captured using a standard flatbed scanner (Epson Perfection V370 Photo, Suwa, Nagano, Japan). Images were saved in .jpg format under standardized conditions of ovistrip presentation, background, debris removal and a fixed resolution (1200 dpi). We created a macro to analyze each image individually within a directory of imagery. The image was initially split into red, green and blue colour channels. In the red channel, eggs appeared as dark regions on a light background. Accurate selection and measurement of dark regions required the image to be smoothed with a median filter (radius 2 pixels) and inverted (eggs appear as light regions on a dark background). A threshold was then applied (intensity 140) to binarize the image so that egg regions were rendered white (intensity 255) on a black background (intensity 0).

ImageJ produces two estimates of egg number; the number of distinct/disjoint eggs or egg regions (the unit method), and the total area (in pixels) of egg regions (the area method). For the latter method, an average value of 168 pixels per Ae. notoscriptus egg (n = 25) was calculated.

The script was automated upon selection of a directory and dual results (a spread sheet containing quantification for each image and binary images showing the regions selected by the macro as being egg-covered) placed in a subdirectory. The binary images provided a quality control mechanism to check that the macro functions correctly and egg regions are located accurately.

Results

Sensitivity of the novel real-time TaqMan RT-PCR assays for detection of Ae. and Ae. albopictus

The Ae. aegypti TaqMan RT-PCR assay was able to detect single 1st instar larvae in all pools that contained increasing numbers of Ae. notoscriptus up to a pool size of at least 5,000 (Table 1). The Ae. albopictus assay was not as sensitive, with the limit of detection being a single 1st instar larva in a pool of 1,000 Ae. aegypti.

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Table 1. Identification of single 1st instar Ae. aegypti or Ae. albopictus larvae in pools of 1st instar Ae. notoscriptus or Ae. aegypti larvae, respectively using the TaqMan RT-PCR assays.

https://doi.org/10.1371/journal.pntd.0005505.t001

Field validation of the RSVP system

A total of 477 ovitraps were deployed in Brisbane of which 400 (83.9%) collected >75,000 Aedes spp. eggs. The processing of 25 pools, representing an estimated ≈ 54,400 eggs, using the Ae. aegypti TaqMan assay did not detect Ae. aegypti.

A total of 45 ovitraps were deployed in Rockhampton, of which 18 (40.0%) collected an estimated total of ≈ 1,700 Aedes spp. eggs. Aedes aegypti was detected in 4 of 6 pools.

A total of 62 ovitraps were deployed in Goomeri, of which 32 (51.6%) collected an estimated ≈ 4,200 eggs. Aedes aegypti was detected in 5 of 8 pools.

Utility of image analysis for quantifying eggs on ovistrips

Due to an absence of visual contaminants, the eggs on 107 of 193 (55.4%) of ovistrips were able to be accurately quantified using ImageJ. Of the remainder, accretion of contaminants rendered 41 of 193 (21.2%) of ovistrips unsuitable for analysis. The eggs on 45 of 193 (23.3%) ovistrips could be quantified with manipulation, such as removing debris or digitally enhancing the image.

Discussion

The RSVP system is a powerful and flexible tool for presence-absence surveillance of invasive Aedes (Stegomyia) mosquito species. RSVP utilizes sensitive molecular diagnostics to resolve logistical and time constraints typically associated with ovitrap surveillance. Molecular diagnoses take approx. 48 h, including hatching of eggs, nucleic acid extraction and real-time RT-PCR. This represents an appreciable time saving, considering that 1,000s of larvae from multiple ovitraps can be processed concurrently. In comparison, ovitraps require approx. 7–10 days to rear cohorts to 4th instar larvae or adults before each specimen is identified by the microscopic examination of key morphological characters. Molecular analyses mitigates the risk of human error in detecting a small number of specimens of an invasive species that are obscured by many 1,000s of larvae from endemic species [64]. Hatching eggs to 1st instar larvae prior to RT-PCR analysis removes requirements for a specific ovitrap format (design, size, material), ovistrip size [65] or substrate (e.g., cloth, paper, germination paper, wood). Shipment of large numbers of egg samples by postal service is also potentially more logistically viable than couriering water samples for newly developed environmental DNA analysis [66].

When viewed in isolation, this application of molecular diagnostics may seem expensive, with a single test costing approx. AU$50.00. However, compared to morphological identification, the saving in time and labor to perform one molecular diagnosis for 10 pooled samples (or AU$5.00 per ovitrap) more than offsets the cost of performing the molecular assays. Furthermore, the cost to perform each test will decrease as the number of cards tested increases. Molecular diagnostics can detect multiple target species (e.g. Ae. albopictus and Ae. aegypti) once the nucleic acids have been extracted, with only a marginal cost increase to include an additional species. Other real-time TaqMan assays can be developed to identify other invasive species, such as Aedes japonicus and Aedes koreicus, which have recently colonized North America and Europe, respectively [67].

Estimates of egg abundance on each ovistrip will guide the aggregation of ovistrips into pool sizes that are below the TaqMan RT-PCR assay’s threshold of detection, and enable comparisons between trap sites or to generate spatial ‘heat maps’ [7]. Egg counting software has the potential to remove the need for manual counting and improve the accuracy and consistency of subjective rapid visual estimates [68]. We demonstrated that systems developed to count eggs harvested under laboratory conditions [69, 70] may not be suitable for rapid analyses of field collections due to greater exposure of ovistrips to contaminants. It should be noted that eggs on any contaminated ovistrips would remain viable for molecular analyses. Pre-trial testing in Brisbane determined that a 2 week ovitrap deployment minimized the accretion of contaminants, such as plant debris and/or bacterial and fungal growth. Whilst the relatively high number of contaminated ovistrips limited its accuracy for automated egg counting in our trials, ImageJ or other image processing software warrants further investigation.

The spatial scale of an RSVP ovitrap network can be adjusted to service early-warning, eradication or quality assurance programs. RSVP was trialled by inexperienced field staff in locations where Ae. aegypti is present (Rockhampton), absent (Brisbane) and near the margin of its geographic range (Goomeri). The simplicity, operational flexibility and the rapidity of diagnosis that RSVP provided was well received by these staff. The RSVP system is currently being translated into an expanded local government program in southeast Queensland. Furthermore, the RSVP is also providing a diagnostic platform for a pilot citizen-science project in southeast Queensland. This pilot will emulate citizen-science programs that monitor invasive mosquitoes in Europe [71] and North America (Invasive Mosquito Project), and may complement community-release programs for novel biological control strategies, such as Wolbachia-based programs [7274].

RSVP may provide a powerful quality assurance tool [75] for ZIKV, DENV or CHIKV responses, either through rapid analyses of eggs from lethal ovitraps [60, 76] or non-lethal ovitraps. Vector presence-absence data within small spatial scales (e.g. residential block) [27, 77, 78] may facilitate risk-based assessment for persistence of local transmission and direct subsequent control operations toward blocks that are infested. The RSVP has potential to be integrated with the roll-out of Wolbachia-based programs. For example, ovitraps were used to assess the rate of invasion by Ae. aegypti infected with the endosymbiont Wolbachia pipientis in open field releases in north Queensland [78]. RSVP may also provide a tool to facilitate the measurement of the heterogeneity of Ae. aegypti in the assessment of candidate sites before releases [79], or aid the assessment of Wolbachia invasion during and after releases for interventions aimed either at the reduction of vector populations [8082] or for control of ZIKV [83, 84], DENV [77, 85, 86] and CHIKV [87, 88] outbreaks.

Acknowledgments

We thank Ian Myles and Michael Onn (Brisbane City Council), Dallas Einsiedel and Melinda Piispanen (Queensland Health) for deploying ovitraps in Brisbane; Nadia Bannerman, Tracey Pottle (Gympie Regional Council) and Robert Mrozowski (Queensland Health) for deploying ovitraps in Goomeri and all the residents and businesses that provided permission. We also thank Scott Ritchie, Gregor Devine and Jonathon Darbro for providing Ae. aegypti and Ae. albopictus for assay development. Ian Mackay and Glen Hewitson assisted with optimization of the molecular assays. Finally, we are indebted to the molecular diagnostic team within Public Health Virology, Forensic and Scientific Services, for conducting the molecular analysis of the field samples.

Author Contributions

  1. Conceptualization: BLM SHM AFvdH.
  2. Data curation: MAS NAH.
  3. Formal analysis: BLM AFvdH.
  4. Investigation: MAS SHM JE CCJ JLM DW.
  5. Methodology: BLM MAS SHM NAH DW AFvdH.
  6. Project administration: BLM AFvdH.
  7. Writing – original draft: BLM AFvdH.
  8. Writing – review & editing: BLM MAS SHM NAH CCJ DW AFvdH.

References

  1. 1. Musso D, Gubler DJ. Zika Virus. Clin Microbiol Rev. 2016;29:487–524. pmid:27029595
  2. 2. Carrington LB, Simmons CP. Human to mosquito transmission of dengue viruses. Front Immunol. 2014;5:290. pmid:24987394
  3. 3. Weaver SC, Forrester NL. Chikungunya: Evolutionary history and recent epidemic spread. Antiviral Res. 2015;120:32–39. pmid:25979669
  4. 4. Lourenco-de-Oliveira R, Castro MG, Braks MA, Lounibos LP. The invasion of urban forest by dengue vectors in Rio de Janeiro. J Vector Ecol. 2004;29:94–100. pmid:15266746
  5. 5. Juliano SA, Lounibos LP. Ecology of invasive mosquitoes: effects on resident species and on human health. Ecol Lett. 2005;8:558–574. pmid:17637849
  6. 6. Williams CR, Long SA, Webb CE, Bitzhenner M, Geier M, Russell RC, et al. Aedes aegypti population sampling using BG-Sentinel traps in north Queensland Australia: statistical considerations for trap deployment and sampling strategy. J Med Entomol. 2007;44:345–350. pmid:17427707
  7. 7. Unlu I, Klingler K, Indelicato N, Faraji A, Strickman D. Suppression of Aedes albopictus, the Asian tiger mosquito, using a "hot spot" approach. Pest Manag Sci. 2015;72:1427–1432. pmid:26482455
  8. 8. Tantowijoyo W, Arguni E, Johnson P, Budiwati N, Nurhayati PI, Fitriana I, et al. Spatial and temporal variation in Aedes aegypti and Aedes albopictus (Diptera: Culicidae) numbers in the Yogyakarta area of Java, Indonesia, with implications for Wolbachia releases. J Med Entomol. 2016;53:188–198. pmid:26576934
  9. 9. Fonseca DM, Unlu I, Crepeau T, Farajollahi A, Healy SP, Bartlett-Healy K, et al. Area-wide management of Aedes albopictus. Part 2: gauging the efficacy of traditional integrated pest control measures against urban container mosquitoes. Pest Manag Sci. 2013;69:1351–1361. pmid:23649950
  10. 10. Maciel-de-Freitas R, Codeco CT, Lourenco-de-Oliveira R. Daily survival rates and dispersal of Aedes aegypti females in Rio de Janeiro, Brazil. Am J Trop Med Hyg. 2007;76:659–665. pmid:17426166
  11. 11. Marini F, Caputo B, Pombi M, Tarsitani G, della Torre A. Study of Aedes albopictus dispersal in Rome, Italy, using sticky traps in mark-release-recapture experiments. Med Vet Entomol. 2010;24:361–368. pmid:20666995
  12. 12. Niebylski ML, Craig GB Jr. Dispersal and survival of Aedes albopictus at a scrap tire yard in Missouri. J Am Mosq Control Assoc. 1994;10:339–343. pmid:7807074
  13. 13. Russell RC, Webb CE, Williams CR, Ritchie SA. Mark-release-recapture study to measure dispersal of the mosquito Aedes aegypti in Cairns, Queensland, Australia. Med Vet Entomol. 2005;19:451–457. pmid:16336310
  14. 14. Ritchie SA, Montgomery BL, Hoffmann AA. Novel estimates of Aedes aegypti (Diptera: Culicidae) population size and adult survival based on Wolbachia releases. J Med Entomol. 2013;50:624–631. pmid:23802459
  15. 15. Lounibos LP, O'Meara GF, Juliano SA, Nishimura N, Escher RL, Reiskind MH, et al. Differential survivorship of invasive mosquito species in south Florida cemeteries: do site-specific microclimates explain patterns of coexistence and exclusion? Ann Entomol Soc Am. 2010;103:757–770. pmid:20852732
  16. 16. Ritchie SA, Moore P, Carruthers M, Williams C, Montgomery B, Foley P, et al. Discovery of a widespread infestation of Aedes albopictus in the Torres Strait, Australia. J Am Mosq Control Assoc. 2006;22:358–365. pmid:17067032
  17. 17. Fonseca DM, Kaplan LR, Heiry RA, Strickman D. Density-dependent oviposition by female Aedes albopictus (Diptera: Culicidae) spreads eggs among containers during the summer but accumulates them in the fall. J Med Entomol. 2015;52:705–712. pmid:26335478
  18. 18. Reiskind MH, Lounibos LP. Spatial and temporal patterns of abundance of Aedes aegypti L. (Stegomyia aegypti) and Aedes albopictus (Skuse) [Stegomyia albopictus (Skuse)] in southern Florida. Med Vet Entomol. 2013;27:421–429. pmid:23278304
  19. 19. Kay BH, Ives WA, Whelan PI, Barker-Hudson P, Fanning ID, Marks EN. Is Aedes albopictus in Australia? Med J Aust. 1990;153:31–34. pmid:2381357
  20. 20. Reiter P. Aedes albopictus and the world trade in used tires, 1988–1995: the shape of things to come? J Am Mosq Control Assoc. 1998;14:83–94. pmid:9599329
  21. 21. Brown JE, Scholte EJ, Dik M, Den Hartog W, Beeuwkes J, Powell JR. Aedes aegypti mosquitoes imported into the Netherlands, 2010. Emerg Infect Dis. 2011;17:2335–2337. pmid:22172498
  22. 22. Madon MB, Hazelrigg JE, Shaw MW, Kluh S, Mulla MS. Has Aedes albopictus established in California? J Am Mosq Control Assoc. 2003;19:297–300. pmid:14710729
  23. 23. Medlock JM, Hansford KM, Schaffner F, Versteirt V, Hendrickx G, Zeller H, et al. A review of the invasive mosquitoes in Europe: ecology, public health risks, and control options. Vector Borne Zoonotic Dis. 2012;12:435–447. pmid:22448724
  24. 24. Whelan P, Nguyen H, Hajkowicz K, Davis J, Smith D, Pyke A, et al. Evidence in Australia for a case of airport dengue. PLoS Negl Trop Dis. 2012;6:e1619. pmid:23029566
  25. 25. Russell RC, Williams CR, Sutherst RW, Ritchie SA. Aedes (Stegomyia) albopictus—a dengue threat for southern Australia? Commun Dis Intell. 2005;29:296–298.
  26. 26. Hanna JN, Ritchie SA. Outbreaks of dengue in north Queensland, 1990–2008. Commun Dis Intell. 2009;33:32–33.
  27. 27. Ritchie SA, Pyke AT, Hall-Mendelin S, Day A, Mores CN, Christofferson RC, et al. An explosive epidemic of DENV-3 in Cairns, Australia. PLoS One. 2013;8:e68137. pmid:23874522
  28. 28. Viennet E, Ritchie SA, Williams CR, Faddy HM, Harley D. Public health responses to and challenges for the control of dengue transmission in high-income countries: four case studies. PLoS Negl Trop Dis. 2016;10:e0004943. pmid:27643596
  29. 29. Russell RC, Currie BJ, Lindsay MD, Mackenzie JS, Ritchie SA, Whelan PI. Dengue and climate change in Australia: predictions for the future should incorporate knowledge from the past. Med J Aust. 2009;190:265–268. pmid:19296793
  30. 30. Beebe NW, Ambrose L, Hill LA, Davis JB, Hapgood G, Cooper RD, et al. Tracing the tiger: population genetics provides valuable insights into the Aedes (Stegomyia) albopictus invasion of the Australasian Region. PLoS Negl Trop Dis. 2013;7:e2361. pmid:23951380
  31. 31. van den Hurk AF, Nicholson J, Beebe NW, Davis J, Muzari OM, Russell RC, et al. Ten years of the Tiger: Aedes albopictus presence in Australia since its discovery in the Torres Strait in 2005. One Health. 2016;2:19–24.
  32. 32. Hill MP, Axford JK, Hoffmann AA. Predicting the spread of Aedes albopictus in Australia under current and future climates: multiple approaches and datasets to incorporate potential evolutionary divergence. Austral Ecology. 2014;39:469–478.
  33. 33. Nicholson J, Ritchie SA, Russell RC, Zalucki MP, van den Hurk AF. Ability for Aedes albopictus (Diptera: Culicidae) to survive at the climatic limits of its potential range in eastern Australia. J Med Entomol. 2014;51:948–957. pmid:25276922
  34. 34. van den Hurk AF, Craig SB, Tulsiani SM, Jansen CC. Emerging tropical diseases in Australia. Part 4. Mosquito-borne diseases. Ann Trop Med Parasitol. 2010;104:623–640. pmid:21144182
  35. 35. Trewin BJ, Kay BH, Darbro JM, Hurst TP. Increased container-breeding mosquito risk owing to drought-induced changes in water harvesting and storage in Brisbane, Australia. Int Health. 2013;5:251–258. pmid:24225151
  36. 36. Mariappan T, Srinivasan R, Jambulingam P. Defective rainwater harvesting structure and dengue vector productivity compared with peridomestic habitats in a coastal town in southern India. J Med Entomol. 2008;45:148–156. pmid:18283956
  37. 37. Unlu I, Faraji A, Indelicato N, Fonseca DM. The hidden world of Asian tiger mosquitoes: immature Aedes albopictus (Skuse) dominate in rainwater corrugated extension spouts. Trans R Soc Trop Med Hyg. 2014;108:699–705. pmid:25193027
  38. 38. Ritchie S, Montgomery B, Walsh I. Production of mosquitoes in rainwater tanks and wells on Yorke Island, Torres Strait: preliminary study. Environmental Health (Australia). 2002;2:13–18.
  39. 39. Kraemer MU, Sinka ME, Duda KA, Mylne AQ, Shearer FM, Barker CM, et al. The global distribution of the arbovirus vectors Aedes aegypti and Ae. albopictus. Elife. 2015;4:e08347. pmid:26126267
  40. 40. Focks DA, Chadee DD. Pupal survey: an epidemiologically significant surveillance method for Aedes aegypti: an example using data from Trinidad. Am J Trop Med Hyg. 1997;56:159–167. pmid:9080874
  41. 41. Wai KT, Arunachalam N, Tana S, Espino F, Kittayapong P, Abeyewickreme W, et al. Estimating dengue vector abundance in the wet and dry season: implications for targeted vector control in urban and peri-urban Asia. Pathog Glob Health. 2012;106:436–445. pmid:23318235
  42. 42. Wu HH, Wang CY, Teng HJ, Lin C, Lu LC, Jian SW, et al. A dengue vector surveillance by human population-stratified ovitrap survey for Aedes (Diptera: Culicidae) adult and egg collections in high dengue-risk areas of Taiwan. J Med Entomol. 2013;50:261–269. pmid:23540112
  43. 43. Montgomery BL, Ritchie SA. Roof gutters: a key container for Aedes aegypti and Ochlerotatus notoscriptus (Diptera: Culicidae) in Australia. Am J Trop Med Hyg. 2002;67:244–246. pmid:12408662
  44. 44. Montgomery BL, Ritchie SA, Hart AJ, Long SA, Walsh ID. Subsoil drain sumps are a key container for Aedes aegypti in Cairns, Australia. J Am Mosq Control Assoc. 2004;20:365–369. pmid:15669376
  45. 45. Krockel U, Rose A, Eiras AE, Geier M. New tools for surveillance of adult yellow fever mosquitoes: comparison of trap catches with human landing rates in an urban environment. J Am Mosq Control Assoc. 2006;22:229–238. pmid:17019768
  46. 46. Williams CR, Long SA, Russell RC, Ritchie SA. Field efficacy of the BG-Sentinel compared with CDC Backpack Aspirators and CO2-baited EVS traps for collection of adult Aedes aegypti in Cairns, Queensland, Australia. J Am Mosq Control Assoc. 2006;22:296–300. pmid:17019776
  47. 47. Crepeau TN, Unlu I, Healy SP, Farajollahi A, Fonseca DM. Experiences with the large-scale operation of the Biogents Sentinel trap. J Am Mosq Control Assoc. 2013;29:177–180. pmid:23923335
  48. 48. Eiras AE, Buhagiar TS, Ritchie SA. Development of the gravid Aedes trap for the capture of adult female container-exploiting mosquitoes (Diptera: Culicidae). J Med Entomol. 2014;51:200–209. pmid:24605470
  49. 49. Ritchie SA, Buhagiar TS, Townsend M, Hoffmann A, van den Hurk AF, McMahon JL, et al. Field validation of the gravid Aedes trap (GAT) for collection of Aedes aegypti (Diptera: Culicidae). J Med Entomol. 2014;51:210–219. pmid:24605471
  50. 50. Johnson BJ, Hurst T, Quoc HL, Unlu I, Freebairn C, Faraji A, et al. Field comparisons of the Gravid Aedes Trap (GAT) and BG-Sentinel trap for monitoring Aedes albopictus (Diptera: Culicidae) populations and notes on indoor GAT collections in Vietnam. J Med Entomol. 2016;In press.
  51. 51. Azil AH, Bruce D, Williams CR. Determining the spatial autocorrelation of dengue vector populations: influences of mosquito sampling method, covariables, and vector control. J Vector Ecol. 2014;39:153–163. pmid:24820568
  52. 52. Degener CM, Eiras AE, Azara TM, Roque RA, Rosner S, Codeco CT, et al. Evaluation of the effectiveness of mass trapping with BG-sentinel traps for dengue vector control: a cluster randomized controlled trial in Manaus, Brazil. J Med Entomol. 2014;51:408–420. pmid:24724291
  53. 53. Dibo MR, Chierotti AP, Ferrari MS, Mendonca AL, Chiaravalloti Neto F. Study of the relationship between Aedes (Stegomyia) aegypti egg and adult densities, dengue fever and climate in Mirassol, state of Sao Paulo, Brazil. Mem Inst Oswaldo Cruz. 2008;103:554–560. pmid:18949325
  54. 54. Codeco CT, Lima AW, Araujo SC, Lima JB, Maciel-de-Freitas R, Honorio NA, et al. Surveillance of Aedes aegypti: comparison of house index with four alternative traps. PLoS Negl Trop Dis. 2015;9:e0003475. pmid:25668559
  55. 55. Regis L, Monteiro AM, Melo-Santos MA, SilveiraJr JC, Furtado AF, Acioli RV, et al. Developing new approaches for detecting and preventing Aedes aegypti population outbreaks: basis for surveillance, alert and control system. Mem Inst Oswaldo Cruz. 2008;103:50–59. pmid:18368236
  56. 56. Hall-Mendelin S, Ritchie SA, Johansen CA, Zborowski P, Cortis G, Dandridge S, et al. Exploiting mosquito sugar feeding to detect mosquito-borne pathogens. Proc Natl Acad Sci U S A. 2010;107:11255–11259. pmid:20534559
  57. 57. Australian Bureau of Statistics. Data by Region: Local Government Areas [Internet]. stat.data.abs.gov.au. 2011 [accessed 21 November 2016].
  58. 58. Harrington LC, Ponlawat A, Edman JD, Scott TW, Vermeylen F. Influence of container size, location, and time of day on oviposition patterns of the dengue vector, Aedes aegypti, in Thailand. Vector Borne Zoonotic Dis. 2008;8:415–423. pmid:18279006
  59. 59. Reiskind MH, Zarrabi AA. Water surface area and depth determine oviposition choice in Aedes albopictus (Diptera: Culicidae). J Med Entomol. 2012;49:71–76. pmid:22308773
  60. 60. Williams CR, Ritchie SA, Long SA, Dennison N, Russell RC. Impact of a bifenthrin-treated lethal ovitrap on Aedes aegypti oviposition and mortality in north Queensland, Australia. J Med Entomol. 2007;44:256–262. pmid:17427694
  61. 61. Ritchie SA. Effect of some animal feeds and oviposition substrates on Aedes oviposition in ovitraps in Cairns, Australia. J Am Mosq Control Assoc. 2001;17:206–208. pmid:14529089
  62. 62. Williams CR, Long SA, Russell RC, Ritchie SA. Optimizing ovitrap use for Aedes aegypti in Cairns, Queensland, Australia: effects of some abiotic factors on field efficacy. J Am Mosq Control Assoc. 2006;22:635–640. pmid:17304930
  63. 63. Schneider CA, Rasband WS, Eliceiri KW. NIH Image to ImageJ: 25 years of image analysis. Nat Methods. 2012;9:671–675. pmid:22930834
  64. 64. Lamche GD, Whelan PI. Variability of larval identification characters of exotic Aedes albopictus (Skuse) intercepted in Darwin, Northern Territory. Commun Dis Intell. 2003;27:105–109.
  65. 65. Falsone L, Brianti E, Severini F, Giannetto S, Romi R. Oviposition substrate in Asian tiger mosquito surveillance: do the sizes matter? J Vector Ecol. 2015;40:256–261. pmid:26611959
  66. 66. Schneider J, Valentini A, Dejean T, Montarsi F, Taberlet P, Glaizot O, et al. Detection of invasive mosquito vectors using environmental DNA (eDNA) from water samples. PLoS One. 2016;11:e0162493. pmid:27626642
  67. 67. Deblauwe I, Sohier C, Schaffner F, Rakotoarivony LM, Coosemans M. Implementation of surveillance of invasive mosquitoes in Belgium according to the ECDC guidelines. Parasit Vectors. 2014;7:201. pmid:24766783
  68. 68. Brasil LM, Gomes MM, Miosso CJ, da Silva MM, Amvame-Nze GD. Web platform using digital image processing and geographic information system tools: a Brazilian case study on dengue. Biomed Eng Online. 2015;14:69. pmid:26178732
  69. 69. Mains JW, Mercer DR, Dobson SL. Digital image analysis to estimate numbers of Aedes eggs oviposited in containers. J Am Mosq Control Assoc. 2008;24:496–501. pmid:19181055
  70. 70. Mello CB, dos Santos WP, Rodrigues MAB, Candeias ALB, Gusmao CMG, Portela NM. Automatic counting of Aedes aegypti eggs in images of ovitraps. In: Naik GR, editor. Recent Advances in Biomedical Engineering. Open Access: InTech; 2009.
  71. 71. Kampen H, Medlock JM, Vaux AG, Koenraadt CJ, van Vliet AJ, Bartumeus F, et al. Approaches to passive mosquito surveillance in the EU. Parasit Vectors. 2015;8:9. pmid:25567671
  72. 72. McNaughton D. The importance of long-term social research in enabling participation and developing engagement strategies for new dengue control technologies. PLoS Negl Trop Dis. 2012;6:e1785. pmid:22953011
  73. 73. McNaughton D, Clough A, Johnson P, Ritchie S, O'Neill S. Beyond the 'back yard': Lay knowledge about Aedes aegypti in northern Australia and its implications for policy and practice. Acta Trop. 2010;116:74–80. pmid:20540930
  74. 74. McNaughton D, Duong TT. Designing a community engagement framework for a new dengue control method: a case study from central Vietnam. PLoS Negl Trop Dis. 2014;8:e2794. pmid:24853391
  75. 75. Morrison AC, Zielinski-Gutierrez E, Scott TW, Rosenberg R. Defining challenges and proposing solutions for control of the virus vector Aedes aegypti. PLoS Med. 2008;5:e68. pmid:18351798
  76. 76. Rapley LP, Johnson PH, Williams CR, Silcock RM, Larkman M, Long SA, et al. A lethal ovitrap-based mass trapping scheme for dengue control in Australia: II. Impact on populations of the mosquito Aedes aegypti. Med Vet Entomol. 2009;23:303–316. pmid:19941596
  77. 77. Hoffmann AA, Goundar AA, Long SA, Johnson PH, Ritchie SA. Invasion of Wolbachia at the residential block level is associated with local abundance of Stegomyia aegypti, yellow fever mosquito, populations and property attributes. Med Vet Entomol. 2014;28 Suppl 1:90–97.
  78. 78. Hoffmann AA, Montgomery BL, Popovici J, Iturbe-Ormaetxe I, Johnson PH, Muzzi F, et al. Successful establishment of Wolbachia in Aedes populations to suppress dengue transmission. Nature. 2011;476:454–457. pmid:21866160
  79. 79. Dutra HL, Dos Santos LM, Caragata EP, Silva JB, Villela DA, Maciel-de-Freitas R, et al. From lab to field: the influence of urban landscapes on the invasive potential of Wolbachia in Brazilian Aedes aegypti mosquitoes. PLoS Negl Trop Dis. 2015;9:e0003689. pmid:25905888
  80. 80. Rasic G, Endersby NM, Williams C, Hoffmann AA. Using Wolbachia-based release for suppression of Aedes mosquitoes: insights from genetic data and population simulations. Ecol Appl. 2014;24:1226–34. pmid:25154109
  81. 81. Ritchie SA, Townsend M, Paton CJ, Callahan AG, Hoffmann AA. Application of wMelPop Wolbachia strain to crash local populations of Aedes aegypti. PLoS Negl Trop Dis. 2015;9:e0003930. pmid:26204449
  82. 82. Zhang D, Zheng X, Xi Z, Bourtzis K, Gilles JR. Combining the sterile insect technique with the incompatible insect technique: I-Impact of Wolbachia infection on the fitness of triple- and double-infected strains of Aedes albopictus. PLoS One. 2015;10(4):e0121126. pmid:25849812
  83. 83. Aliota MT, Peinado SA, Velez ID, Osorio JE. The wMel strain of Wolbachia reduces transmission of Zika virus by Aedes aegypti. Sci Rep. 2016;6:28792. pmid:27364935
  84. 84. Dutra HL, Rocha MN, Dias FB, Mansur SB, Caragata EP, Moreira LA. Wolbachia blocks currently circulating Zika virus isolates in Brazilian Aedes aegypti mosquitoes. Cell Host Microbe. 2016;19:771–774. pmid:27156023
  85. 85. Hoffmann AA, Iturbe-Ormaetxe I, Callahan AG, Phillips BL, Billington K, Axford JK, et al. Stability of the wMel Wolbachia Infection following invasion into Aedes aegypti populations. PLoS Negl Trop Dis. 2014;8:e3115. pmid:25211492
  86. 86. Lambrechts L, Ferguson NM, Harris E, Holmes EC, McGraw EA, O'Neill SL, et al. Assessing the epidemiological effect of Wolbachia for dengue control. Lancet Infect Dis. 2015;15:862–866. pmid:26051887
  87. 87. Aliota MT, Walker EC, Uribe Yepes A, Velez ID, Christensen BM, Osorio JE. The wMel strain of Wolbachia reduces transmission of chikungunya virus in Aedes aegypti. PLoS Negl Trop Dis. 2016;10:e0004677. pmid:27124663
  88. 88. van den Hurk AF, Hall-Mendelin S, Pyke AT, Frentiu FD, McElroy K, Day A, et al. Impact of Wolbachia on infection with chikungunya and yellow fever viruses in the mosquito vector Aedes aegypti. PLoS Negl Trop Dis. 2012;6:e1892. pmid:23133693