Wolbachia mosquito releases were undertaken across four local government administrative areas in northern Queensland, Australia: the Cairns Region, the Cassowary Coast Region, the Douglas Shire and the Charters Towers Region ( Table 1, Figure 1– Figure 3). Within each region, the locations for Wolbachia mosquito releases were selected based on historical dengue case reports, human population density, reported presence of Ae. aegypti and logistical considerations. Depending on the specific objective of each Wolbachia release, for example small scale releases to test different release methods (e.g. egg release trials in Stratford 1–3, Cairns North, Bungalow 1–3, Table 1, Figure 2), or an area-wide release across the entire location (e.g. adult releases across 23 Cairns suburbs between Nov 2016 and Jun 2017, Table 1, Figure 1B, Figure 2), each area was mapped and the target release areas (generally the residential areas and some business areas) were identified. Areas deemed unsuitable for Ae. aegypti, such as uninhabited forested and vegetated areas, open or vacant areas, sporting fields, large industrial and commercial areas, agricultural and farming areas, and major transport infrastructure (major roads, highways, railways and airports) were excluded from releases. The bounded size (km ²) of each release area was calculated, along with the residential population and the number of households ( Table 1).

For Wolbachia mosquito releases between 2011–2014, communication and community engagement activities followed the approach described in Hoffmann et al. (2011). This included consultation with key stakeholders and community groups, one-on-one meetings, displays at community events and centers, and door-knocking and mail-outs to householders to assess support for and participation in Wolbachia mosquito releases. Prior to releases residents were asked to provide permission for release of Wolbachia mosquitoes, either as adult stages from the footpath near their house, or as eggs that were placed into containers by program staff in outdoor shaded areas at their properties. During the release period, we continued to undertake close engagement with the local communities to ensure that residents were informed about the study and were comfortable with continuation of release activities. This included random household surveys, meetings with reference groups of residents and community leaders, and ongoing promotion of a free phone number and accessible city project office in Cairns. Periodic result updates were provided to the communities through letterbox leaflet drops, attendance at community events and meetings, paid advertisements in local newspapers, community newsletters and radio and television media outlets. Residences of a small number of people that did not wish to participate were excluded from release activities.

For Wolbachia mosquito releases between 2015–2017, community engagement activities followed the Public Acceptance Model (PAM) as described in O’Neill et al. (2018). The PAM was comprised of the following four components:

In Cairns, the implementation of the PAM commenced in 2015 and included engagement of traditional mass media, establishment of the Cairns sign-up website (where residents registered their interest in participating in the project), leveraging of existing networks to spread information (particularly through educational institutions), direct-to-premise informative mail-outs, information kiosks at community events and public locations, engagement of high-level stakeholders, such as government representatives, indigenous interest groups and environmental advocacy groups, establishment and maintenance of independent community reference groups, maintenance of community feedback channels (telephone survey and online forms), and distribution of quarterly field trial updates to participants and stakeholders. In Charters Towers, Douglas Shire and the Cassowary Coast the PAM was implemented prior to releases.

Release colony maintenance. Two Ae. aegypti wMel-infected lines were used in releases. The Cairns Ae. aegypti wMel-infected line was released across Cairns, Douglas Shire and the Cassowary Coast areas, and a Townsville Ae. aegypti wMel-infected line was released in Charters Towers, based on the proximity of the latter to Townsville where releases were undertaken between 2014–2015 (see description of the Townsville Ae. aegypti wMel-infected line in O’Neill et al., 2018). The Cairns Ae. aegypti wMel-infected line (described in Walker et al., 2011) was backcrossed to the offspring of Cairns field collected mosquitoes for six generations ( Hoffmann et al., 2011). Between 2011 and 2015 the wMel-infected mosquito colony was maintained at James Cook University, Cairns, in large semi-field cages ( Ritchie et al., 2011) using methods described previously ( Hoffmann et al., 2011). Cages contained up to 10,000 wMel-infected Ae. aegypti and were provided access to human volunteer blood-feeders almost daily. To minimize laboratory adaptation, male Ae. aegypti pupae/adults from field collected Ae. aegypti (F1–F2 eggs) were introduced into the colony each generation, so that they constituted around 10–20% of the new male population. Eggs were collected on flannel cloth in plastic buckets acting as oviposition sites that were spread throughout the cage, incubated for 3 days and then stored in the laboratory.

From 2016 the wMel-infected mosquito colony was maintained at Monash University, Melbourne. The Townsville Ae. aegypti wMel-infected line was established by backcrossing the Cairns Ae. aegypti wMel-infected line to the offspring of Townsville collected wildtype mosquitoes for three generations ( O’Neill et al., 2018). Both wMel-infected lines were maintained in controlled laboratory conditions, in 30 cm ² mesh-sided rearing cages (see description of methods in O’Neill et al., 2018). Each cage contained ~600 adults, and was fed using human volunteers once per week for two gonotrophic cycles. These colonies comprised of a broodstock, and a release-production colony. Male Ae. aegypti adults (from F1-F3 field collected material) were introduced into the broodstock cages at a rate of 10% every generation. Material from the broodstock colony was then transferred to the release-production colony where material was amplified through one generation without the addition of field collected males. Eggs were collected on red flannel cotton strips, and were matured for four days before being dried. Once the drying process was complete ( O’Neill et al., 2018), eggs were packed and shipped to field sites.

Adult mosquito rearing for releases. Between 2011–2013, adult mosquitoes for releases were produced using previously described methods ( Hoffmann et al., 2011). Briefly, immature stages were reared in 3 L buckets with 2 L of water and fed a diet of Tetramin Tropical Tablets (Tetra Holding [US] Inc. Germany, Product number 16110) (2011: 150 larvae per bucket) or ground Tetramin Tropical Flakes (Tetra Holding [US] Inc. Germany, Product number 77101) (2012–2013: 500 larvae/bucket). When approximately 90% of larvae had pupated, the larvae/pupae were sieved and the required number of larvae/pupae were then separated into individual 750 mL plastic containers with approximately 200 ml of water. Adults were allowed to emerge and were maintained for 4–6 days on a 50% honey solution. The cups were then stacked into polystyrene boxes for transport to the release site for release.

For the 2016 Charters Towers releases, wMel-infected mosquito eggs were produced at Monash University and were then shipped to Townsville where the eggs were hatched and the immatures stages were reared to adults in cups in a laboratory maintained at 25–28 °C. Egg strips were either hatched in tap water containing Aqua One Vege Wafers (Aqua Pacific UK Ltd, Southampton, UK, Product number 26050), and two days later approximately 100 larvae were aliquoted into individual release cups (paper drinking cups, 550 mL volume, 100 mm width × 180 mm height, C-DC9787, FPA, Australia) each containing 360 mL of tap water, or egg strips were cut into sections based on the required number of eggs to produce a target hatch of 100 larvae and the eggs were added directly to the individual rearing cups. Immatures were fed Aqua One Vege Wafers (1.5 wafers upon setup and 1.5 wafers on day 4). A mesh cover was placed on each cup and adults were maintained for 4-6 days on a 50% honey solution. Release cups were transferred to plastic tubs and transported to Charters Towers for release the following day between 0800–1000 hours. For the 2016–2017 Cairns releases, adult mosquitoes were reared as above, with initial rearing being undertaken at the James Cook University insectary and this transitioned to a laboratory in Cairns, maintained at 25–28 °C, in early 2017.

For the 2017 Douglas Shire releases, wMel-infected mosquito eggs were produced at Monash University and were then shipped to Port Douglas where the eggs were hatched and the immatures stages were reared to adults in a laboratory maintained at 25–28 °C. Rearing followed above procedures except that all eggs were hatched in water containing 1 Tetramin Tropical Tablet, and two days later approximately 100 larvae were aliquoted into individual release cups (Plastic [PET] drinking cup, 425 mL volume, Detpak, Australia) each containing 300 mL of tap water. Plastic (PET) lids (Detpak, Australia) were placed on top of a mesh cover on each cup and adults were maintained for 4–6 days on 50% honey solution. On the morning of release, release cups were transferred to plastic tubs and transported to the field for release between 0800-1000 hours.

Between 2011 and 2015, weekly releases were undertaken on a per household basis, at a density of 1:3 to 1:10 houses. Between 2016 and 2017, releases were undertaken on an area basis, where the target release area was divided into a series of 100 m × 100 m grids, with a single release point located inside each grid. Adult mosquitoes were transported to the field in release cups in vehicles and released on a single day each week, normally at the property line or front yard by removing the container lids and gently shaking. Releases were generally undertaken between 0800–1600 hours each day.

Between 2011 and 2013 the duration of releases was fixed to between 9 and 16 weeks, except for an initial trial of egg releases in Stratford (SF3) where the methodology was being optimized over a longer period (23 weeks). Shorter duration release periods (7- or 8-weeks) were trialed in Charters Towers and Douglas Shire in 2016, and these were extended in Tully and Innisfail (Cassowary Coast) to 12-week releases and 14- to 16-week releases, respectively, in 2017. The 2016–2017 Cairns releases were staged, with releases continuing in a release area until the frequency of Wolbachia in samples of field-caught mosquitoes was above 50% for two consecutive weeks. The duration of releases in these areas varied from 10–12 weeks, except for Cairns North Ext (CNX) where only two weeks of releases were undertaken. Releases in this area were discontinued after this time because the Wolbachia infection frequency in mosquitoes already exceeded 90% following the spread of Wolbachia from nearby release areas. Details of the releases in each area are summarized in Table 1.

Small-scale field trials to develop and optimize the egg release methods were undertaken in parts of Babinda and Machans Beach in 2013, and in Bungalow (BU1–3), Cairns North (CN1) and Stratford (SF1–3) in 2014. For the 2013 trials, 3 L white polypropylene buckets with lids (Piber Plastics, Australia) were used ( Figure 4A). Each bucket had four 6 mm holes drilled 20 mm apart in a square pattern in the side. Tangle-Trap Sticky Coating (Tanglefoot, USA, Product number 300000588) was applied around the perimeter of the emergence holes (1–2 cm away from holes) to prevent the entry of ants into the buckets. The buckets interiors were roughened with sandpaper to provide perching for mosquitoes post-emergence. Containers were filled with 2 L of tap water and 1 lucerne pellet (0.5 g) (LLP, Carole Park, Australia) and 1 TetraMin Tropical Tablet (broken into four pieces) were added, along with an egg strip containing approximately 150 eggs (target emergence rate of 100 adults per container). Containers were placed in a shaded position near the front boundary of selected houses. Containers were serviced every 2 weeks, at which time live/dead larval/pupal counts were made and the general condition of the container (tipped over, dry/empty, fouled water) was recorded for quality assurance. During the cooler months, immature development in some containers was slowed and necessitated leaving containers in place for 3 weeks prior to servicing. Once immature development was complete, the remaining contents of the containers was discarded and the inside of the container was cleaned with a sponge to remove any residual eggs and larvae/pupae. The container was then refilled with clean tap water, and new food and eggs were added as described above.

The 2014 egg release trials were similar to the above, except these involved 2.3 L plastic buckets containing 1 L of tap water. An alternative feeding mixture involving ground red kidney beans (1.25 g per container) was used in these trials. An assessment of egg hatch rate (hatch rate quality assurance) was undertaken in the laboratory prior to each release round and this information was used to calculate the required numbers of eggs to be added to each container. The target emergence rate for these trials was 75 adults per container.

For the 2015 egg releases, the conditions were the same as the 2014 releases except Aqua One Vege Wafers were used as a food source (4 wafers per containers in spring/summer/autumn months, and 5 wafers per containers in winter months), and each container received 100 viable eggs).

Community egg releases undertaken by school children (Bentley Park 2016, Charters Towers 2016, Mission Beach 2016) and local council staff and Rotary group members (Douglas Shire 2016) involved single use “Mozzie Box” containers which consisted of a 775 ml food container (Detpak, Australia) without handle, and with measurements of 104 × 92 mm (top), 79 × 61 mm (base), 104 mm (height) ( Figure 4B, O’Neill et al., 2018). Four holes were punched into each container and 400ml of water was added to each container. Each Mozzie Box container received food and eggs as described above for 2015 egg releases.

Mosquito collections were undertaken during and after releases using BG Sentinel (BGS) traps (Biogents AG, Regensburg, Germany, Product number NR10030). Mosquitoes were collected and returned to the laboratory for sorting, morphological identification and counting. Aedes aegypti samples were stored in 70% ethanol prior to screening for Wolbachia infection status. BGS trap samples were collected each week during releases, and every one to two after releases. Routine BGS trap collections were maintained for between 2–18 months post release, except for Charters Towers where BGS traps were withdrawn three weeks after completion of releases. After this time BGS traps were reinstalled periodically every 6-12 months and samples were collected after 1-2 weeks.

The number and density of BGS traps in each area varied across the different release periods. Monitoring of releases up until 2014 involved relatively high numbers of BGS traps per area, ranging from 11–80 traps per area and equivalent to 27–160 traps per km ². From 2015, as release areas increased in size, BGS monitoring was undertaken on a per area basis, with densities of 8 BGS traps per km ² during 2015 in Cairns, and this was further reduced to 4 BGS traps per km ² from 2016 onwards in Cairns, Charters Towers and the Cassowary Coast. Ae. aegypti counts and Wolbachia screening results, aggregated by release area and collection period are available as Underlying data ( Ryan, 2019).

Adult Ae. aegypti were screened for Wolbachia using Taqman qPCR on a Roche LightCycler 480 using an internally controlled qualitative assay for the presence or absence of Wolbachia as previously described ( Dar et al., 2008; O’Neill et al., 2018; Yeap et al., 2014). The qPCR cycling program consisted of a denaturation at 95°C for 5 min followed by 45 cycles of PCR (denaturation at 95 °C for 10 sec, annealing at 60 °C for 15 sec, and extension at 72 °C for 1 sec with single acquisition) followed by a cooling down step at 40°C for 10 sec. From September 2018 Wolbachia diagnostics were performed by LAMP. LAMP reactions were performed in a Bio-Rad C1000 96-well PCR thermocycler with a 30min incubation at 65°C as previously described ( O’Neill et al., 2018). Individual reactions consisted of 2X WarmStart® Colorimetric LAMP Master Mix (New England BioLabs, Cat# M1800S), primers and 1 μL of target DNA in a total reaction volume of 17 μL. Reactions for individual samples were performed in 96-well PCR plates (LabAdvantage 96-well PCR plates, full skirt, clear). Plates were incubated in a thermocycler (BioRad C1000) at 65°C for 30 minutes then held at 12°C until scoring. Within one hour of incubation, colour changes of individual samples were recorded. Primers were as follows FIP 5’ TGTATGCGCCTGCATCAGCTTCGGTTCTTATGGTGCTAA, BIP 5’ GCAGAAGCTGGAGTAGCGTTGTGTCATGCCACTTAGATGG, F3 5’ TGATGTAACTCCAGAAGTCA, B3 5’ CTTATTGGACCAACAGGATCG, LpF 5’ AGCCTGTCCGGTTGAATT, LpB 5’ CAGTCTTGTTATCCCAGTGAGT.

Dengue is a notifiable disease in Australia, which mandates clinicians and laboratories to report confirmed and suspected cases to local health authorities. De-identified data was provided by the Queensland Health Communicable Diseases Branch on all laboratory-confirmed and clinically diagnosed (probable) dengue cases notified from Townsville and Cairns and Hinterland Hospital and Health Services (THHS and CHHHS) to the Notifiable Conditions System (NoCS) between 1 January 2000 and 31 March 2019. The NoCS case records include a variable indicating whether the case was classified as imported, on the basis of a history of overseas travel during the 3-12 days prior to illness onset, or locally-acquired. This information is routinely captured in case notifications based on interview by local public health units.

The Townsville Public Health Unit (PHU) provided line-listed data on addresses of all dengue cases notified from THHS and CHHHS, from the operational databases of the Cairns Tropical Public Health Service and the Townsville PHU. This data included one or more addresses per case, with an indication of the primary residential address, and of any address that had been classified as the probable location of dengue acquisition or a possible site of exposure, during the course of public health follow up. We linked the PHU dataset to the NoCS dataset by unique case notification identifier. All NoCS records were retained, with or without a linked PHU record. PHU records without a linked NoCS record were excluded, as NoCS is considered the master source of case notifications data, and holds the travel history variable to distinguish locally-acquired from imported cases.

Approval to access anonymized spatially identifiable dengue case notification data, collected as part of routine disease surveillance, was obtained from the Townsville Hospital and Health Service human research ethics committee (HREC/16/QTHS/108) and research governance office (SSA/16/QTHS/238), and from the Office of the Director-General, Queensland Department of Health, under the Public Health Act 2005.

A case’s location, for the purpose of classifying Wolbachia exposure status, was determined using geolocatable address information from the PHU operational database. The address indicated in the PHU dataset as the probable location of dengue acquisition was used where available (53%); if unavailable then the primary residential address was used (22%). For 20% of cases a geolocatable address was available in the operational database but was not designated as either ‘acquired’ or ‘residential’, and for the remaining 4% no geolocatable address was available in the PHU database, and the suburb of residence from the NOCS dataset was used to define the case’s designated location. Case geolocations were overlaid with Wolbachia release area boundaries in ArcMap (version 10.5, ESRI, Redlands, USA) (QGIS is an open-access alternative) to classify the Wolbachia exposure status of each case at the time of case onset. An area was considered exposed (post-intervention) from the date of last release. Five ‘non-release areas’ in central Cairns, where BGS monitoring indicated that Wolbachia had established by dispersal from neighboring release areas, were considered exposed from the inferred date that the Ae. aegypti infection curve crossed 80%. The local government area (LGA) of each case was determined from its location, as classified above, and any cases located outside of Cairns, the Cassowary Coast, Charters Towers or Douglas LGAs were excluded from the analysis.

The populations of each release area and Wolbachia-exposed non-release area ( Figure 1– Figure 3) were estimated by aggregating from mesh blocks (ABS, 2016) to the boundaries of each intervention area ( Table 1), in ArcMap. In all but three release areas, the boundaries of the release area and monitored area were aligned, and defined the Wolbachia-exposed population for the purpose of epidemiological analyses. The exceptions were Gordonvale, Yorkeys Knob, and Babinda, where monitoring extended beyond the boundaries of the release area and demonstrated Wolbachia establishment throughout this extended area. The population denominator for these three areas was therefore calculated for the larger monitored area ( Table 1 footnote). A further exception was Stratford, where concurrent releases were conducted in three small non-contiguous areas, which represented ~75% of the area and 90% of the population of the suburb of Stratford. For the purpose of epidemiological analysis, the whole suburb population was considered Wolbachia-exposed from the completion of the last releases in the three release zones. For the interrupted time series analysis, monthly aggregate treated and untreated areas (and their resident populations) were calculated, dynamic over time, with the treated area in any given month defined as the total area where Wolbachia deployments had been completed to date (or where, for the five central Cairns non-release areas where Wolbachia established, the inferred local Wolbachia frequency had reached 80%, as above).

Locally-acquired and imported dengue case notifications were cross-tabulated by month of illness onset and LGA. Scaled ‘time-since-release’ (TSR) was calculated for each case located within a Wolbachia exposed area, as (date of case onset – date of last release in the local release area) rounded to the nearest month. TSR is equal to zero for cases with onset in the same month as releases were completed, and is positive for cases with onset post-intervention. Cases located in Wolbachia non-exposed areas were excluded from the TSR analysis. The staggered deployment of Wolbachia across release areas from January 2011 to May 2017 means the distribution of pre-intervention and post-intervention time within the dengue case time series (Jan 2000 – Mar 2019) is variable between release areas. For each release area, the pre-intervention period was calculated as months from January 2000 to end of releases, and post-intervention period as months from end of releases to March 2019. Locally-acquired and imported dengue case notifications were tabulated by month of TSR, and summary statistics for the distribution of pre- and post-intervention time were calculated.

To better visualize the temporal distribution of locally-acquired and imported dengue case notifications with respect to Wolbachia deployments, release areas were grouped by calendar quarter of last release, and cases were plotted by release area group against date of illness onset, stratified by locally-acquired vs imported cases.

Negative binomial regression was used to model monthly counts of locally-acquired dengue cases (January 2000 – March 2019) in aggregate Wolbachia-treated and not-yet-treated areas. Cases located in Wolbachia non-exposed areas were excluded from the analysis. The regression model was fitted in Stata (SE version 14.2, StataCorp, TX) using generalized estimating equations, with epidemic year (September – August) as the cluster variable to account for temporal autocorrelation in the monthly case counts, adjusting for monthly imported dengue cases (any vs none) and season (dry: June – November vs wet: December – May), with a population size offset. A binary intervention variable was included in the regression model to distinguish the pre- or post-intervention status of each area in any given month, the coefficient of which provided the estimate of intervention effect (incidence rate ratio).

Regulatory approval for the release of Aedes aegypti containing Wolbachia was provided by the Australian Pesticides and Veterinary Medicines Authority (Permit numbers 12311, 13183, 13718, 13810, 14266, 14762, 14530, 82947, 14762).

Ethics approval for human blood feeding mosquito colonies in Melbourne was issued from Monash University (CF11/0766 a 2011000387). All volunteers (no children involved) provided written consent.

In Cairns, Human Ethics approval for bloodfeeding was provided by Human Research Ethics Committee, James Cook University (H4907). All adult subjects provided informed oral consent (no children were involved). Names of subjects providing oral consent were recorded in writing.

Verbal and/or written consent from participants was obtained by phone, online or face-to-face to set BG traps, set MRCs, or participate in Community Mosquito Releases.

Surveys were undertaken under Monash ethics: CF13/2407 – 2013001272 Community knowledge of dengue and Wolbachia based dengue control and CF13/2805 – 2013001515 Community knowledge of dengue and Wolbachia based dengue control – Townsville.

Approval to access anonymized spatially identifiable dengue case notification data, collected as part of routine disease surveillance, was obtained from the Townsville Hospital and Health Service human research ethics committee (HREC/16/QTHS/108) and research governance office (SSA/16/QTHS/238), and from the Office of the Director-General, Queensland Department of Health, under the Public Health Act 2005.

Figshare: CNS_Monitoring_Results.xlsx. https://doi.org/10.6084/m9.figshare.9831113 ( Ryan, 2019).

This project contains all data underlying results presented in Figure 6– Figure 15.

Human dengue case notification data was provided to us by Queensland Health. The conditions of the release of the raw dengue case notifications data to us by the Communicable Disease Branch of Queensland Health do not permit further sharing to a third party. This data (locally acquired and imported dengue case notifications) can be acquired by application to Queensland Health: https://www.health.qld.gov.au/clinical-practice/guidelines-procedures/diseases-infection/surveillance/reports/notifiable/data-request. Access to data will be considered following completion of the Data request form (PDF, 237 kB).

Data deposited with Figshare are available under the terms of the Creative Commons Attribution 4.0 International license (CC-BY 4.0).