This research was supported by a CAREER award by the National Science Foundation to SH under award number 2047572. We thank Damien Rivera for his assistance in fruit fly rearing and preparation for topical application assay, Dr. Ganetzky at the University of Wisconsin-Madison for sharing his Canton-S fruit fly strain, the Centers for Disease Control and Prevention for sharing the Rockefeller strain, and the United States Department of Agriculture Center for Medical Agricultural and Veterinary Entomology for sharing the IICC isoline strain. Figure 1 was created with BioRender.com.

The authors declare no conflicts of interest.

This paper presents an adapted protocol for the topical application assay for mosquitoes and fruit flies. This procedure could be easily adapted to be used in the field and with other organisms as it requires minimal specialized equipment. Addressed below are this protocol&#39;s critical steps, potential modifications, troubleshooting advice, limitations of the method, and significance of this method.
Critical steps in the protocol: There are three critical steps in the protocol that, if completed incorrectly, can drastically impact the results of the bioassay: insecticide concentration accuracy, specimen knockdown, and mortality assessment.
Insecticide concentration accuracy: 
It is extremely important to have accurate insecticide solutions to obtain replicable dose-response curves and meaningful results. The volumetric approach to insecticide solution preparation is more common within the literature for both CDC bottle bioassay7 and topical applications13,14,43. However, the gravimetric approach described here is inherently more accurate due to the consideration of temperature through the inclusion of (temperature-specific) density, leading to more accurate formulation preparation.
Specimen knockdown: 
Knocking down the specimens is a critical component of this method and allows for the accurate administration of the insecticide and weight measurements. However, knocking down organisms inevitably contains the risk of physical stress and damage, as previously demonstrated30. Therefore, be cautious and mindful when knocking down the specimens to ensure i) each specimen is knocked down for a similar duration, ii) the length of knockdown is kept to a minimum, and iii) the method of knockdown is kept consistent across all specimens. Additionally, it is advised to test the knockdown method separately, prior to insecticide application, to ensure the method is successful and does not induce control mortality greater than 10%. The initial test may take longer for an inexperienced user, leading to longer knockdown times. Therefore, be cautious when interpreting results from the first assays.
Mortality assessment: 
Assessing mortality can be challenging, especially when the insecticide does not completely kill but only knocks down or maims the mosquito or fly. Therefore, it is important to be aware of how the insecticide impacts the target organism and have a clear definition for &#34;dead&#34; (or knocked down) organisms before starting. Additionally, it is recommended to have the same person assess mortality between doses and replicates to reduce variation.
Protocol modifications: Several modifications described below can be applied to this protocol to improve its versatility and accessibility.
Adapting the assay to smaller or larger-sized insects: 
When using smaller or larger specimens, it is advised to apply a smaller or larger dose volume of insecticide, respectively. As an example, we adapted the mosquito protocol to fruit flies by reducing the 0.5 &#181;L dose to a 0.2 &#181;L dose. Ensure the correct syringe size is chosen for the chosen dose volume.
Adapting the assay to field insects: 
When using field insects, there may be more variation in insect size. Therefore, weighing the insects in smaller groups (e.g., per cup) would be recommended instead of as a large group (e.g., all insects used for one experiment). This can help capture the potential variation in insecticide susceptibility associated with the differences in field insect mass.
Equipment modifications: 
Insect handling tent: Dosing of the specimen can be completed under an insect handling tent that is simply constructed with PVC pipe and mosquito netting. This can be an alternative to an enclosed room (e.g., insectary) and help eliminate potential insecticide contamination in areas where insect rearing might occur. This insect handling tent is easy to construct and low-cost (~$70). Alternatively, an insect handling cage could be purchased (~$425).
Chill table: Ice packs or trays of ice can be used for knocking down the specimen and/or keeping the specimen knocked down.
Incubator: Incubators are recommended for rearing the specimen and holding the specimen for 24 h after insecticide treatment. If an incubator is not available, it can be constructed. Equipment needed to build the incubator includes an insulated container, humidifier, heat cables, humidity and temperature controller, and a light, which should add up to a total cost of ~$170, following and expanding upon previous methods44.
Holding cups: Although plastic cups are used to sort and hold the treated specimen, wax-lined paper cups or glass containers would be suitable alternatives.
Organism and life stage modification: 
This method is very adaptable for use with other vectors, insects, and/or arthropods such as Culex quinquefasciatus mosquitoes32, house flies32, and cockroaches45, as well as non-adult life stages, such as mosquito larvae46.
Topical application location modification: 
This method describes applying the insecticide to the ventral thorax and abdomen region for mosquitoes (and the dorsum for fruit flies). However, other application locations can be used as long as the exposure site is consistent. Consistency is important because insecticide sensitivity can vary based on application location32.
Troubleshooting advice: This method has several steps that are initially challenging. Described below are some of the most common issues one might encounter.
Leaking/evaporating insecticide solutions: 
Insecticides are commonly dissolved in acetone, a highly volatile compound. This means acetone evaporates quickly at room temperature, increasing the insecticide concentrations over time. If the insecticide solutions appear to be leaking or evaporating, remake the solutions, ensure the tube&#39;s lid is on tightly, and double-check that the storage protocols are being properly followed (e.g., parafilm is being used, and the tubes are stored upright). If leaking persists, try filling the tubes with a lower volume to allow more room for the change in volume the acetone experiences at different temperatures. Additionally, if using acetone as the solvent, ensure the tubes are rated for acetone storage (e.g., FEP, TFE, and PFA plastics). If using hydrophobic insecticides, store the solutions in glass vials (as hydrophobic insecticides adhere to&#160;glass less than plastic). It is also good practice to mark the meniscus of the solution prior to storing to monitor evaporation.
Weight drifting on microbalance when weighing organisms: 
If the weight reading on the scale is drifting (slowly going up or down), this could be due to static. Drift most often occurs when weighing organisms in plastic items, as plastic can easily hold a static charge. To avoid this, a weighing paper can be placed underneath the plastic container being weighed, or a non-plastic container such as glass can be used.
Abnormal mortality results: 
There are many ways by which the mortality results may seem abnormal, such as observing high mortality in the controls or high/low mortality throughout all insecticide doses. Review the following cases for troubleshooting each scenario.
High control mortality 
If there is high mortality in the control group (10% or greater), evaluate the knockdown method and length of time the specimens are knocked down. If possible, shorten the length of time for which the specimens are knocked down. Other potential factors to consider for high mortality in the controls include i) checking if the incubator settings are correct&#8212;abnormal temperatures and/or humidity could lead to increased mortality. Temperature and humidity should be checked with an independent data logger. ii) Assessing insect handling. Handling insects too much or too roughly could lead to high mortality. iii) Checking if there is no insecticide contamination in the 100% acetone used to treat the control group or on the instrumentation. Replace acetone and clean all instruments with acetone or ethanol. Avoid contamination by frequently replacing gloves, preventing spillage, and cleaning instruments. Note that in Supplemental File 3, a maximum of two mosquitoes died within the control (acetone-only) cups. This level of mortality is not considered high (it is less than 10%), and therefore, there was no cause for concern.
High mortality in all exposed groups (but not in control groups) 
Use lower insecticide concentrations or smaller dose volumes for testing. The dosages used might be above the minimum dose that will not induce mortality. Use several 10-fold dilutions to identify the correct dose range, and rule out contamination. To avoid contamination, start dosing with the lowest concentration and work towards the highest concentration. Additionally, make sure all equipment used is regularly cleaned with acetone and/or ethanol, the doses applied to the specimen are very small, and even the slightest cross-contamination could impact the results.
Low mortality in all exposed groups 
Use higher insecticide concentrations. The dosages used might all be too low to cause mortality in the population. To identify the correct dose range, expose specimens to several more 10-fold concentrated dosages. Ensure the insecticide solutions have not expired or degraded (potentially due to high temperature or light exposure). If the solutions have expired or are suspected of having degraded, remake the solutions and ensure proper storage conditions are followed.
Inconsistent mortality between replicates/days 
The time of the day when insects are exposed to the insecticide could affect the level of resistance expressed, especially for metabolic resistance34. Repeat this protocol during the same window of time each day to avoid time-of-day as a potential variable contributing to changes in mortality. Other potential factors contributing to inconsistent mortality between replicates include i) specimens being differentially reared between experiments. Ensure all specimens are of the same age range, reared at the same temperature and similar densities and food availability. ii) insecticide concentrations degrading&#160;over time or becoming&#160;more concentrated due to acetone evaporation. Remake the solutions and ensure proper storage conditions. iii) Inconsistent mortality scoring. Ensure the same person scores mortality or develop a clear protocol to be used consistently across the team. Use blind scoring to reduce bias in mortality scoring.
Insects sticking to the surface of the sorting tray: 
Acetone reacts to plastics used in this protocol, such as Petri dishes. The specimen will likely adhere to the surface if using acetone on Petri dishes or similar plastic surfaces. This adhesion can be avoided by lining the sorting tray with weighing paper or using a non-plastic sorting tray. Additionally, condensation on the surface of plastic in the sorting tray or holding cups can lead to insects adhering to the condensation, or the specimen may be too cold and potentially freeze to the surface. Adjust the knockdown method to reduce condensation while preventing&#160;the specimens becoming too cold/frozen (e.g., place weighing paper between the specimens and the plastic sorting tray).
R analysis errors: 
Once the mortality data are collected, a variety of complications may occur during analysis. The most common reason an R code cannot complete the actions for the data file is that the data format does not match the code (e.g., column headings and/or empty cells). If more serious complications arise, refer to the R help pages built into Rstudio35.
Limitations of the above-described topical application method: 
Insecticide absorption via topical application method does not mimic natural exposure: 
Topical application on the primary body is not the natural way of insecticide absorption. In the field, insects mostly absorb insecticides through their legs over the length of time they are in contact with the insecticide-treated surface or on their wings through small aerosol particles47,48, rather than a rapid exposure on the ventral surface. However, the direct application of a known insecticide dose will accurately establish a phenotypic response to insecticides, needed for genetic and evolutionary studies or comparisons of insecticide susceptibility across space or time. Therefore, this approach is beneficial for testing technical resistance but will not directly measure practical resistance (the efficacy of the actual intervention tool in a field setting15). However, it is important to note that the current standard methods (e.g., WHO tube tests and CDC bottle bioassays) also cannot capture or mimic aerosol (i.e., by fogging) insecticide exposure in the field.
Topical application assays can only assess contact absorption insecticides: 
This method is intended for insecticides that work through contact and absorption of the insecticide and not for use with oral insecticides, such as boric acid commonly used in attractive toxic sugar baits49.
Significance of the method: 
The topical application method expands on well-established standards for insecticide bioassays by calculating the lethal dose (not concentration) and measuring technical (not practical) resistance15. Given below are the advantages and disadvantages of this method over existing insecticide susceptibility assays.
Lethal dose calculation: 
This method determines the lethal dose of the insecticide, rather than the lethal concentration that the CDC and WHO bioassays use to establish the discriminating dose11. The lethal dose is more meaningful because it is a quantified amount of insecticide known to elicit mortality. In contrast, the lethal concentration does not consider how much insecticide the organism actually acquires. When using the lethal dose calculation, differences between sex- or size-dependent susceptibility profiles can be more accurately observed and quantified, making this measurement even more versatile.
Technical resistance: 
This method assesses technical resistance, which is resistance as measured under standardized, controlled environments. Such measurements are suitable for surveillance of the spread of insecticide resistance and linking phenotypic resistance with potential markers15. Because of the decreased variation in mortality resulting from the topical application bioassay, it allows for better identification of new resistance markers. However, due to the unnatural exposure of insecticides to the mosquito, this assay is not suitable for the estimation of efficacy of a specific intervention in a specific population. Other assays are needed for measurements of such practical resistance15.
Specimen adaptability: 
This method can be practiced on other important arthropods such as crop pests (e.g., Colorado potato beetle), house pests (e.g., cockroaches and bed bugs), or pollinators (e.g., bees) with simple changes to the knockdown approach and/or insecticide dose, volume, and/or concentration (as described above). The ease of adaptability can help analogize insecticide resistance research across different research fields. The use of an LD50 value instead of a lethal concentration that kills 50% of the specimens (LC50) allows accurate comparison across species.
Cost: 
Similar to CDC bottle bioassays and WHO tube tests, costs to run the topical application assay are minimal (see the Table of Materials). The essential pieces of equipment are the syringe (approximately $70) and the dispenser (approximately $100), which are reusable across assays.
Number of specimens needed: 
A minimum of 20-25 specimens should be used per topical application assay cup. A minimum of five insecticide concentrations is recommended to be tested per experiment, with a minimum of three replicates recommended for the procedure. Overall, this results in a minimum of 300-375 specimens needed for a complete test, comparable to the number of specimens needed to perform resistance intensity tests using WHO tube tests or CDC bottle bioassays. However, if reduced variability is achieved with the topical application bioassay, the same number of specimens may lead to more statistical power to compare susceptibility data across space or time.

Mosquitoes are responsible for over 700,000 deaths each year due to the diseases they transmit to humans, with over half of those deaths due to malaria alone1. The main preventive method against transmission of malaria and other vector-borne diseases is the use of insecticides, often in the form of long-lasting insecticide nets or indoor-residual spraying2. However, insecticide resistance is widespread among mosquitoes and other insect vectors, as well as agricultural pests3,4. To effectively manage resistance, surveillance is of key importance5. For this, highly accurate and high-throughput resistance detection methods are needed. Currently, the most widespread insecticide resistance surveillance tools for mosquitoes are the WHO tube test6 and the CDC bottle bioassay7. For fruit flies the residual contact application method (similar to the CDC bottle bioassay)&#160;is a commonly used insecticide bioassay8,9,10. However, variability in data from these methods is typically high, with measurements of the same laboratory mosquito strain ranging from ~20-70% mortality in CDC bottle assays and 0-50% in WHO tube tests when exposed to sublethal dosages11. Such variation is surprising because the limited genetic variation in most laboratory strains is expected to lead to limited insecticide susceptibility variation in the population. Nevertheless, there is still a high level of variation observed in the bioassay results.
Potential sources of this variation could be a result of heterogeneous insecticide exposure between specimens within the bioassay due to indirect insecticide exposure via the surface, heterogeneous environmental effects, normal biological variation between individuals of the same genotype, and variation in the mass of specimens of the same population12. An infrequently used method with higher replicability is the topical application bioassay. In this assay, the insecticide is directly applied to each insect13,14, removing the factor of heterogeneous exposure of different specimens within the same assay. However, due to the slow-throughput nature of this method, it is not routinely used as an insecticide susceptibility surveillance tool for mosquito populations. This paper presents a modified protocol for the topical application bioassay that allows for higher-throughput exposures while also correcting for variation in insect mass, a parameter that correlates to changes in insecticide susceptibility12. A reduction in noise and mass-associated variation in mortality data from variable insecticide exposure would allow for more accurate technical resistance surveillance11,15. Such data could be used to more accurately associate phenotypic resistance with genetic markers, fitness parameters, and/or vector competence. Additionally, we demonstrate how this assay could easily be adapted to other insect species by using the topical application bioassay on fruit flies, a smaller-bodied insect species.
The main limitation of the aforementioned residual contact applications is that insecticide exposure may vary from specimen to specimen within the same assay. In the case of CDC bottle bioassays and the contact method, insecticide exposure may vary between replicates of the same assay. The insects are exposed to insecticide that is either distributed on the inside of a glass bottle (CDC bottle bioassay and contact method) or on impregnated papers (WHO tube test). The concentration of insecticide on both surfaces (glass and paper) is known and predetermined by screening different species of known genotypes. However, the amount available to potentially be absorbed by the insect can greatly vary depending on the surface used, the insecticide mixture components, and how homogeneously the insecticide is distributed across the surface material16,17. In the CDC bottle bioassay, the insecticide coating on the inside of the bottle is dependent on procedures employed by each laboratory and user. In the WHO tube test, the insecticide-treated papers are centrally produced and thus most likely quite homogeneous across labs. However, in the WHO tube test, the exposure tube allows specimens to land and rest on non-insecticide-exposed metal mesh, leading to potential heterogeneous insecticide exposure among the specimens within each test. The actual amount of insecticide picked up and absorbed by specimens via each method still needs to be explored further18.
Additionally, the CDC bottle bioassay, WHO tube test, and contact method are most commonly used as threshold assays testing only one predetermined insecticide concentration. This approach can accurately detect the presence of resistance and is valuable for resistance surveillance (especially when resistance is spreading). However, threshold assays cannot quantify the strength of the resistance, which might be more predictive of the efficacy of intervention tools. If multiple insecticide concentrations are used with these methods, then they can be used as intensity assays. Intensity assays for the CDC bottle bioassay and the WHO tube test have been introduced by testing 5x and 10x the predetermined discriminating dosages to address this gap in surveillance6,19. While providing greater ability to differentiate between resistant populations, 3-5 (predetermined) dosages provide limited resolution to calculate lethal concentrations. Additionally, mosquitoes of various sizes are used in such assays. Yet, the mass is important to measure as larger specimens might need a higher dose to be killed as the effective dose per unit of mass will be much lower than that of a smaller organism12. Calculating a mass-relativized lethal dose (amount of insecticide per insect mass) would be a more useful metric than the more common lethal concentration (e.g., amount of insecticide per surface area) as it considers the variation of insect mass between sexes, populations, and genotypes. Such data would help fill the gap between genotypic and phenotypic resistance within the laboratory and the field and could also provide an easy way to calculate the needed application concentration to treat a population of insects of a known average mass.
The use of mass-relativized lethal dosages that kill 50% of the specimens (LD50) also incorporates several other benefits. Assessment of the toxicity of a specific compound in mg/kg (= ng/mg) is standard in human and veterinary toxicology14, and LD50 values are found on material safety data sheets. Lethal dosages also allow direct comparison of toxicity between different chemicals toward a particular species or the same chemical toward different species20, as well as high-quality evaluation of novel insecticides and chemicals13. Additionally, the LD50 can provide more meaningful and accurate resistance ratios than those derived from diagnostic dose mortality results, which can result in an overestimation of the resistance level present in a population. Therefore, this assay would be suitable for routine surveillance programs by providing more rigorous resistance monitoring based on mass-relativized lethal doses derived from more specimens than recommended for other bioassays21.
The topical application method has been used in insecticide susceptibility surveillance for mosquitoes and flies as an alternative for the standard insecticide susceptibility bioassays when resistance is already known or suspected22,23, as well as for surveillance in some pest insects24 to more accurately assess resistance profiles and insecticide intrinsic toxicity21. In topical application bioassays, the insecticide is applied to each organism, resulting in minimal variation in insecticide exposure. This paper presents a slightly adapted and improved method that allows for insecticide exposure to be applied to a large number of insects in a short period while also controlling for insect mass22. This higher-throughput method with good levels of replicability could be a useful additional tool for routine insecticide susceptibility surveillance.

<table><tbody><tr><td>1.5 mL microcentrifuge tubes</td><td>Thomas Scientific</td><td>20A00L068</td><td>Acetone aliquot storage</td></tr><tr><td>1.5 mL screw cap tubes</td><td>Thomas Scientific</td><td>1182K23</td><td>Insecticide dilution storage</td></tr><tr><td>15 mL conical tubes</td><td>VWR</td><td>339651</td><td>Insecticide dilution storage</td></tr><tr><td>20 mL glass scintillation vials</td><td>Fisher Scientific</td><td>0334125D</td><td>Fruit fly weighing</td></tr><tr><td>25 &amp;mu;L syringe</td><td>Fisher Scientific</td><td>14815288</td><td>Topical applicator</td></tr><tr><td>Acetone</td><td>Fisher Scientific</td><td>AC423240040</td><td>ACS 99.6%, 4 L</td></tr><tr><td>&lt;em&gt;Aedes aegypti&lt;/em&gt; (IICC strain)</td><td>USDA CMAVE</td><td>NA</td><td>Insecticide resistant</td></tr><tr><td>&lt;em&gt;Aedes aegypti&lt;/em&gt; (Rockefeller strain)</td><td>CDC</td><td>NA</td><td>Insecticide susceptible</td></tr><tr><td>Analytical scale</td><td>Fisher Scientific</td><td>14-557-409</td><td>Precision up to 0.1 mg</td></tr><tr><td>Aspirator</td><td>Amazon</td><td>6.49986E+11</td><td>Mosquito collection device</td></tr><tr><td>Bench paper</td><td>VWR</td><td>89126-794</td><td>Place under workspace</td></tr><tr><td>Cotton swabs</td><td>Amazon</td><td>B092S8JVQN</td><td>Use for sorting insects</td></tr><tr><td>Cotton wool balls</td><td>Amazon</td><td>B0769MKZWT</td><td>Use for sucrose solution</td></tr><tr><td>Dispenser</td><td>Fisher Scientific</td><td>1482225</td><td>Repeater pipettor</td></tr><tr><td>&lt;em&gt;Drosophila melanogaster&lt;/em&gt; (Canton-S strain)</td><td>University of Wisconsin-Madison</td><td>NA</td><td>Insecticide susceptible</td></tr><tr><td>Fine-tipped paint brushes</td><td>Amazon</td><td>B07KT2X1BK</td><td>Use for sorting insects</td></tr><tr><td>Fruit fly stock bottles</td><td>Fisher Scientific</td><td>AS355</td><td>Use for rearing and sorting fruit flies</td></tr><tr><td>Hand-held CO&lt;sub&gt;2&lt;/sub&gt; dispenser</td><td>Fisher Scientific</td><td>NC1710679</td><td>Use for knocking down insects</td></tr><tr><td>Holding cups</td><td>Amazon</td><td>B08DXG7V1S</td><td>Clear plastic</td></tr><tr><td>Ice pack</td><td>Amazon</td><td>B08QDWMMW5</td><td>Use for knocking down fruit flies</td></tr><tr><td>Ice trays</td><td>Amazon</td><td>9301085269</td><td>Use for knocking down insects</td></tr><tr><td>Insect forceps</td><td>Amazon</td><td>B07B4767WR</td><td>Insect forceps</td></tr><tr><td>Insecticide</td><td>Sigma-Aldrich Inc</td><td>45423-250MG</td><td>Deltamethrin</td></tr><tr><td>Labeling stickers</td><td>Amazon</td><td>B07Q4X9GWX</td><td>3/4&quot; Color dot stickers</td></tr><tr><td>Labeling tape</td><td>Amazon</td><td>B00X6A1GYK</td><td>White tape</td></tr><tr><td>Netting</td><td>Amazon</td><td>B07F2PHHWV</td><td>Use for covering holding cups and insect handling tent</td></tr><tr><td>Petri dishes</td><td>Fisher Scientific</td><td>FB0875712H371</td><td>100 mm x 15 mm</td></tr><tr><td>PVC Pipe</td><td>Lowe&amp;rsquo;s</td><td>23971</td><td>Insect handling tent materials</td></tr><tr><td>Rubber bands</td><td>Amazon</td><td>B00006IBRU</td><td>Use for securing mesh/net on cups</td></tr><tr><td>Sucrose</td><td>Amazon</td><td>B01J78INO0</td><td>Granulated White Sugar</td></tr><tr><td>Weighing paper</td><td>VWR</td><td>12578-165</td><td>4&quot; x 4&quot;</td></tr></tbody></table>

topical application bioassay to quantify insecticide toxicity for mosquitoes and fruit flies

The continued use of insecticides for public health and agriculture has led to widespread insecticide resistance and hampering of control methods. Insecticide resistance surveillance of mosquito populations is typically done through Centers for Disease Control and Prevention (CDC) bottle bioassays or World Health Organization (WHO) tube tests. However, these methods can result in a high degree of variability in mortality data due to variable insecticide contact with&#160;the insect, the relatively small numbers of organisms tested, extensive variation in mass between populations, and constantly changing environmental conditions, leading to variable outcomes. This paper presents the topical application bioassay, adapted as a high-throughput phenotypic bioassay for both mosquitoes and fruit flies, to test large numbers of insects along&#160;a range of insecticide concentrations.
This assay 1) ensures consistent treatment and insecticide contact with every organism, 2) produces highly specific dose-response curves that account for differences in average mass between strains and sexes (which is particularly important for field-collected organisms), and 3) allows for the calculation of statistically rigorous median lethal doses (LD50), which are necessary for resistance ratio comparisons-an alternative surveillance approach from diagnostic dose mortality, which is also used for larvicide resistance surveillance. This assay will be a complementary tool for accurately phenotyping mosquito populations and, as illustrated using fruit flies, is easily adaptable for use with other insects. We argue that this assay will help fill the gap between genotypic and phenotypic insecticide resistance in multiple insect species.

These representative results feature two different strains of Ae. aegypti, Rockefeller (ROCK), and an isolated field strain from Florida with known knockdown resistance mutations F1534C and V1016I (IICC genotype). Additionally, Drosophila melanogaster (Canton: S strain) is featured.
Figure 2 and Figure 3 illustrate the dose response of each organism by strain and sex tested following the above protocol. As no differences were observed between the dose-response curves of male and female mosquitoes within each strain (t = 1.70, p = 0.098 for ROCK and t = 0.64, p = 0.527 for IICC), data from both sexes within each mosquito strain were pooled. The mass-relativized LD50 for ROCK and IICC are 0.008 ng/mg (95% CI: 0-0.104) and 0.336 ng/mg (95% CI: 0.235-0.438), respectively. The 95% CIs of these values do not overlap, indicating significantly different dose responses of the strains. The RR of the IICC strain (relative to the ROCK strain) is 41.7, which according to the WHO, is considered highly resistant5. For the Canton-S fruit flies, the mass-relativized LD50 is 0.213 ng/mg (95% CI: 0-0.490).
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/63391/63391fig02.jpg" /> 
Figure 2: Representative data of mosquitoes using topical application bioassay. Representative dose-response data from topical application bioassay following the above protocol using deltamethrin and mosquitoes: (A) female Ae. aegypti ROCK (n = 880) and IICC (n = 550) strains, (B) male Ae. aegypti ROCK (n = 880) and IICC (n = 569) strains. Deltamethrin testing concentrations ranged from 0.00075 ng/&#181;L to 9.68705 ng/&#181;L, and the dose of deltamethrin applied (ng) per average mosquito mass (mg) is reflected on the x-axis. Mortality is shown as a proportion on the y-axis. The black line through each data point cluster represents the strain and sex-specific linear regression. <a href="https://www.jove.com/files/ftp_upload/63391/63391fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/63391/63391fig03.jpg" /> 
Figure 3: Representative data of fruit flies using topical application bioassay. Representative dose-response data from topical application bioassay following the above protocol using deltamethrin and fruit flies: D. melanogaster Canton-S strain (n = 1014). Deltamethrin testing concentrations ranged from 0.00499 to 5.02876 ng/&#181;L, and the dose of deltamethrin applied (ng) per average fruit fly mass (mg) is reflected on the x-axis. Mortality is shown as a proportion on the y-axis. The black line represents the linear regression. <a href="https://www.jove.com/files/ftp_upload/63391/63391fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Supplemental Figure S1: Benchtop insect handling tent. Benchtop insect handling tent is used for easier capture of escaping mosquitoes or flies during the topical application assay. Structure is closed in A and open in B. This structure was built with PVC pipe and fine-mesh fabric. <a href="https://www.jove.com/files/ftp_upload/63391/Supplemental_Figure1.Final.V1.pdf" target="_blank">Please click here to download this File.</a>
Supplemental Figure S2: Syringe and repeater applicator unit. Syringe and repeater applicator unit used for dosing insects. Main parts include 1) needle, 2) syringe barrel, 3) plunger, 4) repeater, and 5) repeater button. <a href="https://www.jove.com/files/ftp_upload/63391/Supplemental_Figure2.Final.V1.pdf" target="_blank">Please click here to download this File.</a>
Supplemental File 1: Randomization script: Randomization script to create non-biased labels for all cups of each experiment. <a href="https://www.jove.com/files/ftp_upload/63391/Supplemental_File1_Randomization.Script.Final.V1.docx" target="_blank">Please click here to download this File.</a>
Supplemental File 2: Mortality score sheet: Mortality score sheet to assist mortality assessment. Sheet also includes places to record all other important information to record, as referenced in the protocol, such as the insecticide application start and end times. <a href="https://www.jove.com/files/ftp_upload/63391/Supplemental_File2_Mortality.Score.Sheet.Final.V1 (1).docx" target="_blank">Please click here to download this File.</a>
Supplemental File 3: Example mortality data: Example data file used to create Figure 2. The column heading descriptions are as follows: &#34;id&#34; = identification code of each data point; &#34;species&#34; = species name (e.g., Aedes aegypti); &#34;insecticide&#34; = name of insecticide topically applied (e.g., Deltamethrin); &#34;strain&#34; = name of mosquito strain (e.g., ROCK); &#34;date&#34; = start date topical application; &#34;sex&#34; = sex of mosquitoes; &#34;age&#34; = age of mosquitoes (young = 3-5-day-old; old = 4 weeks old); &#34;total.mosq&#34; = total number of mosquitoes weighed in batch; &#34;weight&#34; = weight (mg) of all mosquitoes within batch; &#34;concentration&#34; = concentration of insecticide (&#181;g/mL); &#34;syringe&#34; = droplet volume (mL) of syringe; &#34;dose&#34; = amount of insecticide active ingredient applied to each mosquito (ng); &#34;total&#34; = number of mosquitoes in each cup; &#34;dead&#34; = number of dead mosquitoes in each cup. <a href="https://www.jove.com/files/ftp_upload/63391/Supplemental_File3_Example.Data.Final.V1 (1).txt" target="_blank">Please click here to download this File.</a>
Supplemental File 4: R analysis code: Example R code that can be used to complete the Probit analysis (as described in step 8 of the protocol). The representative results (accessible via the supplemental example data file) can be used with this R code. <a href="https://www.jove.com/files/ftp_upload/63391/Supplemental_File4.Final.V3 (1).R" target="_blank">Please click here to download this File.</a>

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Topical Application Bioassay to Quantify Insecticide Toxicity for Mosquitoes and Fruit Flies

We describe the methodology and importance of the topical application bioassay to measure insecticide susceptibility in mosquitoes and fruit flies. The presented assay is high-throughput, utilizes insect mass-thus allowing for calculating a mass-relativized lethal dose instead of concentration-and likely has lower variability than other similar methods.

The continued use of insecticides for public health and agriculture has led to widespread insecticide resistance and hampering of control methods. ...

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Research

JoVE Journal

Biology

5.7K Views.  Arizona State University. Door insecticide rechtstreeks op het insect aan te brengen in plaats van via oppervlaktecontact, is er minimale variatie in blootstelling aan insecticiden, en dit stelt ons in staat om het gevoeligheidsprofiel van muggenpopulaties nauwkeuriger te meten. Het belangrijkste voordeel van deze techniek is dat het ons in staat stelt om massa-gerelativeerde dodelijke doses van een insecticide te berekenen in plaats van dodelijke concentraties, wat de vergelijkingen van de resistentieverhouding aanzi...

Video: Topical Application Bioassay to Quantify Insecticide Toxicity for Mosquitoes and Fruit Flies

Studying Aggression in Drosophila (fruit flies)

Aggression is an innate behavior that evolved in the framework of defending or obtaining resources. This complex social behavior is influenced by genetic, hormonal and environmental factors. In many organisms, aggression is critical to survival but controlling and suppressing aggression in distinct contexts also has become increasingly important. In recent years, invertebrates have become increasingly useful as model systems for investigating the genetic and systems biological basis of complex social behavior. This is in part due to the diverse repertoire of behaviors exhibited by these organisms. In the accompanying video, we outline a method for analyzing aggression in Drosophila whose design encompasses important eco-ethological constraints. Details include steps for: making a fighting chamber; isolating and painting flies; adding flies to the fight chamber; and video taping fights. This approach is currently being used to identify candidate genes important in aggression and in elaborating the neuronal circuitry that underlies the output of aggression and other social behaviors.

Studying Aggression in Drosophila fruit flies

Aggression is an innate behavior that evolved in the framework of defending or obtaining resources.  This complex social behavior is influenced by ...

Injection of An. stephensi Embryos to Generate Malaria-resistant Mosquitoes

The introduction of exogenous genes into the genomes of mosquitoes requires microinjection techniques tailored to the specific species of interest. This video protocol demonstrates a method used by the James laboratory to microinject DNA constructs into Anopheles stephensi embryos for the generation of transformed mosquitoes. Techniques for preparing microinjection needles, collecting and preparing embryos and performing the microinjection are illustrated.

Anopheles stephensi mosquitoes are vectors for malaria inhabiting India and throughout Asia. This video demonstrates the technique for performing microinjections of this species with transgenes that will confer resistance to the malaria to the mosquito. Much of the methodology demonstrated in this video is applicable to microinjection techniques of other mosquito species.

The introduction of exogenous genes into the genomes of mosquitoes requires microinjection techniques tailored to the specific species of interest. This ...

Visualizing Efficacy of Pesticides Against Disease Vector Mosquitoes in the Field

Efficacy of public health pesticides targeting nuisance and disease-vector insects such as mosquitoes, sand flies, and filth-breeding flies is not uniform across ecological zones. To best protect public and veterinary health from these insects, the environmental limitations of pesticides need to be investigated to inform effective use of the most appropriate pesticide formulations and techniques. We have developed a research program to evaluate combinations of pesticides, pesticide application equipment, and application techniques in hot-arid desert, hot-humid tropical, warm and cool temperate, and urban locations to derive pesticide use guidelines specific to target insect and environment. To these ends we designed a system of protocols to support efficient, cost-effective, portable, and standardized evaluation of a diverse range of pesticides and equipment across multiple environments. At the core of these protocols is the use of an array of small cages with colony-reared sentinel mosquitoes (adults and immatures) and sand flies (adults), strategically arranged in natural habitats and exposed to pesticide spray. Spatial and temporal patterns of pesticide efficacy are derived from percent mortality in sentinel cages, then mapped and visualized in a geographic information system. Maps of sentinel mortality data may be statistically compared to evaluate relative efficacy of a pesticide across multiple environments, or to study multiple pesticides in a single environment. Protocols may be modified to accommodate a variety of scenarios, including, for example, the vertical orientation of sentinels in canopy habitats or simultaneous testing of ground and aerial application methods.

The efficacy of public health pesticides targeting nuisance and disease-vector insects is not uniform across different ecological zones. Here we present a system of techniques using captive vector insects as sentinels for pesticide efficacy to derive electronic maps supporting the standard evaluation of pesticides across multiple environments.

Efficacy of public health pesticides targeting nuisance and disease-vector insects such as mosquitoes, sand flies, and filth-breeding flies is not uniform ...

Environment

Bioassays for Monitoring Insecticide Resistance

Pest resistance to pesticides is an increasing problem because pesticides are an integral part of high-yielding production agriculture. When few products are labeled for an individual pest within a particular crop system, chemical control options are limited. Therefore, the same product(s) are used repeatedly and continual selection pressure is placed on the target pest. There are both financial and environmental costs associated with the development of resistant populations. The cost of pesticide resistance has been estimated at approximately $ 1.5 billion annually in the United States. This paper will describe protocols, currently used to monitor arthropod (specifically insects) populations for the development of resistance. The adult vial test is used to measure the toxicity to contact insecticides and a modification of this test is used for plant-systemic insecticides. In these bioassays, insects are exposed to technical grade insecticide and responses (mortality) recorded at a specific post-exposure interval. The mortality data are subjected to Log Dose probit analysis to generate estimates of a lethal concentration that provides mortality to 50% (LC50) of the target populations and a series of confidence limits (CL's) as estimates of data variability. When these data are collected for a range of insecticide-susceptible populations, the LC50 can be used as baseline data for future monitoring purposes. After populations have been exposed to products, the results can be compared to a previously determined LC50 using the same methodology.

This manuscript demonstrates and discusses techniques used to survey pesticide susceptibility and detect resistance to contact and systemic pesticides in arthropod pests.

Pest resistance to pesticides is an increasing problem because pesticides are an integral part of high-yielding production agriculture. When few products ...

An Optimized Protocol for Rearing Fopius arisanus, a Parasitoid of Tephritid Fruit Flies

Fopius arisanus (Sonan) is an important parasitoid of Tephritid fruit flies for at least two reasons. First, it is the one of only three opiine parasitoids known to infect the host during the egg stage1. Second, it has a wide range of potential fruit fly hosts. Perhaps due to its life history, F. arisanus has been a successfully used for biological control of fruit flies in multiple tropical regions2-4. One impediment to the wide use of F. arisanus for fruit fly control is that it is difficult to establish a stable laboratory colony5-9. Despite this difficulty, in the 1990s USDA researchers developed a reliable method to maintain laboratory populations of F. arisanus10-12. There is significant interest in F. arisanus biology13,14, especially regarding its ability to colonize a wide variety of Tephritid hosts14-17; interest is especially driven by the alarming spread of Bactrocera fruit fly pests to new continents in the last decade18. Further research on F. arisanus and additional deployments of this species as a biological control agent will benefit from optimizations and improvements of rearing methods. In this protocol and associated video article we describe an optimized method for rearing F. arisanus based on a previously described approach12. The method we describe here allows rearing of F. arisanus in a small scale without the use of fruit, using materials available in tropical regions around the world and with relatively low manual labor requirements.

An Optimized Protocol for Rearing Fopius arisanus, a Parasitoid of Tephritid Fruit Flies

Fopius arisanus is an egg-larval parasitoid of Tephritid fruit flies that is successfully used in biological control of these important tropical pests. We describe here an optimized protocol for rearing F. arisanus on a small scale using readily available materials.

Fopius arisanus (Sonan) is an important parasitoid of Tephritid fruit flies for at least two reasons. First, it is the one of only three opiine ...

Isolation and Biophysical Study of Fruit Cuticles

The cuticle, a hydrophobic protective layer on the aerial parts of terrestrial plants, functions as a versatile defensive barrier to various biotic and abiotic stresses and also regulates water flow from the external environment.1 A biopolyester (cutin) and long-chain fatty acids (waxes) form the principal structural framework of the cuticle; the functional integrity of the cuticular layer depends on the outer 'epicuticular' layer as well as the blend consisting of the cutin biopolymer and 'intracuticular' waxes.2 Herein, we describe a comprehensive protocol to extract waxes exhaustively from commercial tomato (Solanum lycopersicum) fruit cuticles or to remove epicuticular and intracuticular waxes sequentially and selectively from the cuticle composite. The method of Jetter and Sch&auml;ffer (2001) was adapted for the stepwise extraction of epicuticular and intracuticular waxes from the fruit cuticle.3,4 To monitor the process of sequential wax removal, solid-state cross-polarization magic-angle-spinning (CPMAS) 13C NMR spectroscopy was used in parallel with atomic force microscopy (AFM), providing molecular-level structural profiles of the bulk materials complemented by information on the microscale topography and roughness of the cuticular surfaces. To evaluate the cross-linking capabilities of dewaxed cuticles from cultivated wild-type and single-gene mutant tomato fruits, MAS 13C NMR was used to compare the relative proportions of oxygenated aliphatic (CHO and CH2O) chemical moieties.
Exhaustive dewaxing by stepwise Soxhlet extraction with a panel of solvents of varying polarity provides an effective means to isolate wax moieties based on the hydrophobic characteristics of their aliphatic and aromatic constituents, while preserving the chemical structure of the cutin biopolyester. The mechanical extraction of epicuticular waxes and selective removal of intracuticular waxes, when monitored by complementary physical methodologies, provides an unprecedented means to investigate the cuticle assembly: this approach reveals the supramolecular organization and structural integration of various types of waxes, the architecture of the cutin-wax matrix, and the chemical composition of each constituent. In addition, solid-state 13C NMR reveals differences in the relative numbers of CHO and CH2O chemical moieties for wild-type and mutant red ripe tomato fruits. The NMR techniques offer exceptional tools to fingerprint the molecular structure of cuticular materials that are insoluble, amorphous, and chemically heterogeneous. As a noninvasive surface-selective imaging technique, AFM furnishes an effective and direct means to probe the structural organization of the cuticular assembly on the nm-&mu;m length scale.

Aerial plant organs are protected by the cuticle, a supramolecular biopolyester-wax assembly. We present protocols to monitor selective removal of epi- and intracuticular waxes from tomato fruit cuticles on molecular and micro scales by solid-state NMR and atomic force microscopy, respectively, and to assess the cross-linking capacity of engineered cuticular biopolyesters.

The cuticle, a hydrophobic protective layer on the aerial parts of terrestrial plants, functions as a versatile defensive barrier to various biotic and ...

A Multi-detection Assay for Malaria Transmitting Mosquitoes

The Anopheles gambiae species complex includes the major malaria transmitting mosquitoes in Africa. Because these species are of such medical importance, several traits are typically characterized using molecular assays to aid in epidemiological studies. These traits include species identification, insecticide resistance, parasite infection status, and host preference. Since populations of the Anopheles gambiae complex are morphologically indistinguishable, a polymerase chain reaction (PCR) is traditionally used to identify species. Once the species is known, several downstream assays are routinely performed to elucidate further characteristics. For instance, mutations known as KDR in a para gene confer resistance against DDT and pyrethroid insecticides. Additionally, enzyme-linked immunosorbent assays (ELISAs) or Plasmodium parasite DNA detection PCR assays are used to detect parasites present in mosquito tissues. Lastly, a combination of PCR and restriction enzyme digests can be used to elucidate host preference (e.g., human vs. animal blood) by screening the mosquito bloodmeal for host-specific DNA. We have developed a multi-detection assay (MDA) that combines all of the aforementioned assays into a single multiplex reaction genotyping 33SNPs for 96 or 384 samples at a time. Because the MDA includes multiple markers for species, Plasmodium detection, and host blood identification, the likelihood of generating false positives or negatives is greatly reduced from previous assays that include only one marker per trait. This robust and simple assay can detect these key mosquito traits cost-effectively and in a fraction of the time of existing assays.

Malaria transmitting mosquitoes have a number of epidemiologically important characteristics that can only be detected using molecular techniques. Utilizing a MALDI-TOF based SNP genotyping platform, we developed an assay for simultaneously detecting multiple key traits (species, insecticide resistance, parasite infection and host choice) of malaria vectors.

The Anopheles gambiae species complex includes the major malaria transmitting mosquitoes in Africa. Because these species are of such medical importance, ...

Protocols for Testing the Toxicity of Novel Insecticidal Chemistries to Mosquitoes

New classes of insecticides with novel modes of action are needed to control insecticide resistant populations of mosquitoes that transmit diseases such as Zika, dengue and malaria. Assays for rapid, high-throughput analyses of unformulated novel chemistries against mosquito larvae and adults are presented. We describe protocols for single point-dose and dose response assays to evaluate the toxicity of small molecule chemistries to the Aedes aegypti vector of Zika, dengue and yellow fever,&#160;the malaria vector, Anopheles gambiae and the northern house mosquito, Culex quinquefasciatus, on contact and via ingestion. As an example, we evaluated the toxicity of amitriptyline, a small molecule antagonist of G protein-coupled receptors, via larval, adult topical and adult blood-feeding assay. The protocols provide a starting point to investigate insecticide potential. Results are discussed in the context of additional experiments to explore product applications and mechanisms for delivery.

Protocols are described for assessing the toxicity of chemistries to immature and adult mosquitoes for development as new classes of larvicides, adulticides and endectocides. The protocols enable high-throughput testing of multiple chemistries at single-point dose and subsequent evaluation via dose response assay to determine toxicity on contact or via ingestion.

New classes of insecticides with novel modes of action are needed to control insecticide resistant populations of mosquitoes that transmit diseases such ...

Topical Testing Assay: A Technique to Evaluate the Insecticidal Potential of Dopamine Receptor Antagonists on Adult Female Mosquitoes by Topical Application

Source: Brito-Sierra, C.A. et al., <a href="https://www.jove.com/v/57768">Protocols for Testing the Toxicity of Novel Insecticidal Chemistries to Mosquitoes.</a> J. Vis. Exp. (2019).
In this video, we demonstrate the topical testing assay to determine the toxicity of a potential insecticide on adult female mosquitoes. Using a dopamine receptor antagonist that causes lethal inhibition of neurotransmission in mosquitoes, this method describes a way to evaluate the toxic effects of insecticides upon topical application.

Source: Brito-Sierra, C.A. et al., Protocols for Testing the Toxicity of Novel Insecticidal Chemistries to Mosquitoes. J. Vis. Exp. (2019).
In this video, ...

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Mosquito Feeding Assay: A Method to Evaluate Test Compound Effects on Mortality and Fecundity of Adult Female Mosquitoes

Source: Brito-Sierra, C. A. et al., <a href="https://www.jove.com/v/57768">Protocols for Testing the Toxicity of Novel Insecticidal Chemistries to Mosquitoes.</a> J. Vis. Exp. (2019).
This video demonstrates a blood-feeding assay to evaluate the mortality and fecundity of test compounds with novel modes of action on female mosquitoes. This technique helps in screening new classes of insecticides to kill the resistant population of mosquitoes and control disease spread.

Source: Brito-Sierra, C. A. et al., Protocols for Testing the Toxicity of Novel Insecticidal Chemistries to Mosquitoes. J. Vis. Exp. (2019).
This video ...

Topical Application Bioassay to Quantify Insecticide Toxicity for Mosquitoes and Fruit Flies

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