**1. Introduction**

The prevention of vector borne diseases is often achieved by controlling the insect population, which currently largely relies on the use of insecticides. Malaria prevalence has halved since 2000, primarily due to vector control interventions, saving 660 million lives, with a large part of the reduction being attributable to the use of insecticides [1]. The primary vector control tools employed against malaria are insecticide treated nets (ITNs) and indoor residual spray (IRS). However, insecticide resistance represents a major threat to human health. Alternative interventions with different active ingredients and/or modes of action, capable of controlling insecticide resistant vectors, as well as vectors which transmit malaria outdoors, are urgently required to ensure the sustainability of malaria control interventions.

A number of Attractive Toxic Sugar Baits (ATSBs) are being evaluated as part of an Integrated Vector Management (IVM) approach. ATSBs deployed outdoors are likely to be particularly effective against outdoor biting mosquitoes, as well as mosquitoes that are resistant to pyrethroid insecticides. An ATSB has been developed by Westham Co. which utilizes the neonicotinoid dinotefuran. The bait station includes a permeable membrane that allows volatile attractive compounds to be emitted and encourage feeding by mosquitoes ye<sup>t</sup> minimizes feeding by non-target organisms (NTOs) and tarsal contact of both mosquitoes and NTOs with the insecticide-treated bait. Because of the inclusion of this permeable membrane, the product is termed an Attractive *Targeted* Sugar Bait (ATSB®) [2]. These bait stations have been shown to be effective in controlling malaria vectors in Mali [3] and are under evaluation in conjunction with the Innovative Vector Control Consortium (IVCC) in trials in Zambia, Kenya, and Mali. Dinotefuran is included as the active ingredient and, since this insecticide is new to public health, it is not expected that target mosquito populations will carry any resistance to it, though another neonicotinoid, clothianidin, is now in use for public health and cross-resistance is a risk. As with any new vector control tool based on insecticides, once dinotefuran-based ATSBs are deployed, susceptibility testing will need to be introduced to allow early detection of possible emerging resistance and enable evidence-based resistance managemen<sup>t</sup> strategies. Conventionally discriminating (or diagnostic) dose bioassays are used to detect resistance to insecticides encountered by mosquitoes through tarsal contact on an ITN or IRS, and so a WHO tube assay [4] or the Centers for Disease Control and Prevention (CDC) bottle bioassay [5] is used. Survival in a discriminating dose (DD) or discriminating concentration (DC) assay is a sign of possible resistance in the target population.

Such diagnostic bioassays are not available for orally ingested insecticides, and nor is a DC recommended by the WHO for susceptibility monitoring of dinotefuran [4] via tarsal contact. Methods for detecting resistance to neurotoxic insecticides via tarsal contact may not be applicable for the orally ingested dinotefuran due to its negative log P, which inhibits tarsal uptake due to epicuticular lipids and barriers. In addition, it is possible that this different method of exposure may be affected by different resistance mechanisms than those responsible for resistance against contact insecticides. It is therefore desirable to establish a suitable method to screen for dinotefuran resistance in ATSB deployment sites.

Ideally, mosquitoes would be fed a discriminating dose of dinotefuran ingested in a sugar solution to most closely match the exposure route in an ATSB. However, to date only a few methods, rather complicated in terms of practical implementation, have been established to feed a spiked sugar meal to mosquitoes with a high enough feeding rate to allow this form of resistance monitoring to be done [6,7]. Assays based on feeding an AI to insects in a sugar meal are vulnerable to huge variability and poor accuracy, due to poor feeding rates, especially with recently colonized or field-caught mosquitoes, and variable volumes taken up by those mosquitoes that do feed. Topical application of insecticide bypasses tarsal barriers by applying insecticide solutions in lipophilic solvent directly to the thorax of the mosquito to be taken in through the cuticle [4]. Although this is not the same entry system as the proposed delivery via ingestion, it is a technique that also bypasses cuticular barriers and therefore may be sufficiently representative of oral uptake whilst being easily applicable for routine susceptibility monitoring in field sites. Indeed, it has been shown in agricultural pests that the response to exposure to neurotoxic insecticides by topical application is a good proxy for the response to ingestion and that resistance monitoring assays for oral insecticides can be based on topical application [8]. Topical application is relatively quick with even large numbers of mosquitoes and can be done with fairly straightforward portable equipment, and as such is a more robust method for susceptibility testing.

Here, an oral application assay has been developed, able to determine dose response curves among two *Anopheles* strains, that prevent tarsal contact while allowing ingestion of a spiked sugar meal. Practical topical application bioassays were also developed by applying dinotefuran to the thorax of adult *Anopheles* mosquitoes with organic carrier to bypass lipid cuticle barriers. The topical application dose response was compared with the oral toxicity bioassays across several *Anopheles* strains.

#### **2. Materials and Methods**

## *2.1. Mosquito Rearing*

Mosquitoes were reared within the insectaries of Liverpool Insect Testing Establishment (LITE) at the Liverpool School of Tropical Medicine as previously described [9], at 26 ± 2 ◦C and 80 ± 10% relative humidity. Four strains were used for experiments, all described by Williams et al. [9]. Kisumu is an insecticide-susceptible strain of *Anopheles gambiae*, colonized from Kenya in 1975. VK7 2014 is a strain of *An. coluzzi* colonized from Valley de Kou, Burkina Faso, in 2014 and resistant to pyrethroids and Dichlorodiphenyltrichloroethane (DDT) as a result of both target site and metabolic resistance mechanisms. Fang is a susceptible colony of *An. funestus* colonized from Calueque, Southern Angola in 2015. FUMOZ-R, also *An. funestus,* was colonized from Mozambique in 2000 before being selected by exposure to lambda-cyhalothrin to produce a strain with a high level of metabolic resistance [10]. Though not of the same species, the VK7 2014 strain was compared to Kisumu, a model laboratory colony of the *Anopheles gambiae* species complex which is susceptible to all classes of insecticide. A direct species comparison of susceptible (Fang) and pyrethroid resistant (FUMOZ-R) strains was possible for *An. Funestus*.

All tests were carried out using 2–5 day old female mosquitoes which had been allowed to sugar feed and mate but not blood feed prior to testing. For mosquito size for each sample, refer to Supplementary Materials.

## *2.2. Ingestion Assay*

Between one hundred and two hundred 2–5 day old female mosquitoes were starved in a standard (30 cm × 30 cm × 30 cm) BugDorm-1 (MegaView Science Co., Ltd., Taichung, Taiwan) cage for approximately 18 h with ad libitum access to purified (Merck Millipore, Darmstadt, Germany) water-soaked cotton on top of the cage mesh. Mosquitoes were then exposed to insecticide mixed into sugar solution by adding 2 feeders, designed to prevent tarsal contact ye<sup>t</sup> allow easy feeding, to each cage for 24 h (Figure 1).

The sugar solution was 10% sucrose (granulated sugar in de-ionized water), 0.8% Uranine (Fluorescein Sodium Salt; Honeywell, Charlotte, NC, USA) fluorescent marker, and treatment-dependent insecticide concentration (0.000001%, 0.00001%, 0.00002%, 0.000025%, 0.00005%, 0.000075%, 0.0001%, and 0.001% *w*/*v*). Between 1 and 5 replications were performed per concentration and between 5 and 9 replications for each control to collect sufficient data to generate LD values using Rstudio (See Supplementary data). Technical grade (98.7%) dinotefuran was sourced from Sigma–Aldrich (Manchester, UK). 10 mL of each insecticide concentration solution was used to soak cotton wool inside sugar feeders less than 5 min prior to adding the feeder to the cage. Two feeders were used per cage to ensure mosquitoes had adequate access to sugar/insecticide solution.

**Figure 1.** Sugar feeder made from a plastic pot of 2.5 cm height, large radius of 4.5 cm, and small radius of 4 cm. A wad of cotton wool was pressed into the pot to be just below the upper lip without touching the netting (judged by eye) and soaked in sugar solution into which the required concentration of insecticide was dissolved. A double layer of netting was stretched across the top and secured using an elastic band.

After the exposure period, all mosquitoes were aspirated out of each cage into holding cups, separated by treatment as well as living or dead, then frozen at −20 ◦C. Once mosquitoes were killed (usually 1–2 h at freezing temperatures), they were scored for fluorescence using a Leica MZ 10 F microscope (Leica Microsystems, Milton Keynes, UK) under a yellow-fluorescent protein (YFP) filter (Figure 2). Only those mosquitoes that were scored as being positive for feeding by fluorescence were included in the mortality calculations. Feeding rate was calculated for each replicate test from the proportion of fluorescent positive mosquitoes relative to all exposed mosquitoes. All raw bioassay data is available in Supplementary Materials.

**Figure 2.** Side by side comparison of two adult female mosquitoes, one fed on 10% sugar solution only (**right**) and one fed on 10% sugar solution with 0.8% Uranine (**left**). Photographs are taken using white light (**A**) and UV light under a YFP filter (**B**).
