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

NOTE: Insecticides can cause human, animal, and environmental hazards²⁵. Caution, training, and personal protective equipment are highly advised. Be sure to follow the material safety data sheets for all insecticides and solvents used.

1. Rear specimens

1. Rear 3-5-day-old adult mosquitoes.
   NOTE: The protocol below reflects conditions for Aedes aegypti rearing, closely following Food and Agriculture Organization of the United Nations guidelines²⁶.
   1. Rear mosquitoes of all life stages at 27 ± 1 °C and 75 ± 5% relative humidity with 12:12 h light and dark cycling.
   2. Hatch the mosquito eggs by submerging them in deionized water and adding yeast²⁶, or place the submerged eggs inside a vacuum chamber for 30 min.
      NOTE: Both methods decrease the oxygen content within the water and increase hatching²⁷.
   3. Feed the newly hatched larvae fish food (or an equivalent diet such as ground cat kibble) within trays and keep the larval density as similar as possible between trays as larval density impacts development¹² (e.g., 200-250 larvae per tray containing a total of 1.5 L of water).
   4. Feed the larvae every other day until they reach the pupal stage (approximately 7-10 days), increasing the amount of food as needed.
      NOTE: When fed too little, larval growth will be stunted, and the larvae may eat one another. When fed too much, the larvae may die, causing the water to go foul.
   5. Once pupae develop, transfer them daily to a water bowl in adult mosquito cages and provide 10% sucrose solution ad libitum.
   6. Record the first day of adult emergence. Remove the remaining pupae from the cage 2 days after emergence starts.
      NOTE: Male mosquitoes emerge faster. Note the emergence of males and females separately and ensure sufficient males and females are available for each test.
   7. Wait for 3 days after removing the pupae to achieve 3-5-day-old mosquitoes for testing.
2. Rear fruit flies (loosely following protocols of the University of Zurich²⁸).
   1. Rear Drosophila strains in stock bottles at 23 ± 1 °C and 60 ± 5% relative humidity with 12:12 h light and dark cycling.
      NOTE: Drosophila stock bottles should contain 75 mL of a standard fly medium, which is first poured as a liquid into the bottom of the bottles and then allowed to solidify overnight.
   2. Transfer colonies to new stock bottles with fresh food every two weeks to prevent overpopulation and mold growth. To do this, knock down flies using a hand-held carbon dioxide (CO₂) dispenser, transfer the anesthetized flies to a weighing paper on an ice pack or chill table, and brush the flies into a fresh stock bottle using a fine-tipped paintbrush. Be sure to keep the bottles on their sides during this process to prevent flies from falling into the food and drowning.

2. Prepare insecticide formulations using the gravimetric approach

1. Make the first stock solution following the gravimetric approach using an analytical scale with 0.1 mg accuracy inside a fume hood.
   NOTE: The gravimetric approach uses mass to measure the amounts of insecticide and solvent added. The standard practice (volumetric approach) will require an analytical scale to measure the amount of (solid) insecticide added when the first stock solution is prepared; however, the amount of solvent added and all following dilutions are measured by volume only. The gravimetric approach has a higher level of accuracy and is therefore preferred.
   1. Determine the target insecticide concentration and target volume (maximum 10 mL is recommended if using 15 mL conical tubes to prevent spillage when storing in a freezer) for the first stock solution and calculate how much insecticide active ingredient (AI) to add using Eq (1):
        (1)
   2. Prepare a storage tube (15 mL conical tube recommended for larger volumes, 1.5 mL microcentrifuge screw cap tubes recommended for volumes of 1 mL or less) and label with insecticide and solvent name, target concentration, and preparation date. Place the tube and lid on the scale within a rack or holder and tare the scale.
   3. Weigh the desired amount of solid or liquid insecticide AI determined from step 2.1.1. (e.g., deltamethrin used for the representative data) into the tube and record the mass.
   4. Tare the scale and add the desired volume of solvent (equivalent to the target volume) to the tube, close the lid immediately, and record the mass. Close the tube's lid immediately after adding the solvent (acetone used here) to avoid evaporation and mix the solution.
   5. Record the room temperature. Some solvents, such as acetone, can have significant changes in volume (and consequently density) depending on temperature.
   6. If storing immediately, wrap the tube's lid in parafilm (to reduce evaporation), place it in a tube rack/holder (to keep upright and prevent leaking), cover in foil (to prevent UV exposure), place it in a resealable plastic bag (to reduce evaporation), and place the bag in a -20 °C freezer. If not stored immediately, make sure the lid is secured and cover in foil or a light-protected container.
   7. Calculate the stock solution's actual concentration (mg/mL) by dividing the mass of insecticide AI added by the volume of solvent added (and the volume of insecticide added if in liquid form). To calculate the volume of added solvent (or liquid insecticide), divide the mass added by the known density that is appropriate for the recorded temperature.
   8. Calculate the density (g/mL) of the stock solution by dividing the total mass added (insecticide and solvent) by the total volume added (solvent and insecticide, if in liquid form). See step 2.1.7 for converting liquid mass to volume.
2. Serially dilute the initial stock solution via 10% dilutions. If needed, use these serial dilutions to create an initial dose-response curve to identify the target range of insecticide concentrations for the bioassay.
   1. Calculate the volume of insecticide stock solution and the solvent to add to each tube (e.g., 1 mL of insecticide stock solution diluted in 9 mL of solvent for a 10 mL dilution of 10% of the previous concentration).
   2. Vortex the stock solution for 10 s. Tare a prelabeled first dilution tube on the scale. Add the required volume of stock solution to the first dilution tube using a pipette. Immediately close the lid of both tubes and record the mass in the first dilution tube.
   3. Tare the first dilution tube again and add the required volume of solvent. Close the lid immediately, record the mass of the added solvent, and vortex the first dilution for 10 s.
   4. Repeat steps 2.2.2 and 2.2.3 for the remaining dilutions.
   5. Store all dilutions as described above in step 2.1.6.
   6. Calculate the actual concentrations of the dilutions by following step 2.1.7.
   7. Calculate the density of each insecticide dilution by dividing the total mass added (insecticide solution and solvent) by the total volume added (insecticide solution and solvent). For each serial dilution, use the previous insecticide stock dilution's density to calculate the new dilution's density following Eq (2):
      (2)
3. Optional: Create insecticide dilutions with smaller increments by serial dilution.
   1. Select the concentrations and volumes of each new solution to make with the aid of a dose-response curve of the initial serial dilutions, previous trials, or published literature.
      NOTE: Chosen concentrations should result in a mortality range of 0-100%, with a minimum of three concentrations from this range to allow for Probit analysis.
   2. Use the serial dilutions as stock solutions to make each new dilution and follow step 2.2 to create the new dilutions between the 10-fold dilutions.
4. Optional: Aliquot the insecticide solution. If larger volumes of the insecticide solutions are made, aliquot the solutions into 1.5 mL screw-cap tubes to avoid contamination, evaporation, and degradation of the stock solutions from frequent handling and exposure to light.
   1. Aliquot the solutions, starting from the lowest concentration and work towards the highest concentration to reduce potential contamination. Mix each stock solution by vortexing for 10 s before opening and pipetting the desired volume (e.g., 0.5 mL) into a prelabeled screw cap tube.
   2. Store the aliquots in a light-resistant container in a -20 °C freezer.
      NOTE: It is recommended to regularly (monthly) replace aliquots with small new aliquots taken directly from the stock pesticide dilutions. This will limit the potential for contamination to be carried over into other experiments or changes due to evaporation or UV degradation while the samples are used on the bench. The protocol can be paused here and restarted even years later, as long as the insecticide solutions are stored properly (see step 2.1.6) and kept in the -20 °C freezer.
5. Use a permanent marker pen to mark the meniscus before storing to monitor solvent evaporation. When removing insecticide solution to make aliquots, mark the meniscus every time the solution is removed.

3. Prepare topical application bioassay workspace

NOTE: It is recommended to work in a benchtop insect handling tent for easier capture of escaping mosquitoes or flies. See Supplemental Figure S1 for images of an insect handling tent.

1. Remove the needed insecticide solutions from the freezer, vortex immediately, and place them in a light-resistant container at room temperature to let the insecticides warm to room temperature before using.
   NOTE: Insecticide AIs can separate from the solvent at cooler temperatures. Additionally, acetone volume changes with temperature, which can alter the applied insecticide dose. Mixing the solutions and allowing them to warm to room temperature helps ensure consistency when using the insecticide solutions.
2. Set out all needed tools and materials for the topical application assay in the insect handling tent as referenced in the Table of Materials.
3. Clean the syringe barrel and needle with analytical grade acetone by completing 5 washes per acetone aliquot. Complete this with 5 separate aliquots for a total of 25 washes. See Supplemental Figure S2 for syringe and repeater pipettor parts.
   1. Set out 5 microcentrifuge tubes with 0.5 mL of acetone each.
   2. Fill the syringe barrel with 0.025 mL of acetone from the first tube and then expel the acetone into a waste container by swiftly pushing down on the plunger. Repeat four more times to complete a total of five acetone washes from the same acetone aliquot. Then, fill the syringe barrel completely with air and expel the air and potential acetone remnants into the waste container. Repeat two more times to complete three "washes" with air.
   3. Repeat step 3.3.2 for the remaining 4 tubes of acetone.
   4. Create an air pocket within the barrel between the syringe plunger and the top of the needle by pulling up the plunger slightly into the barrel (~5 mm).
      NOTE: This air pocket protects the plunger from contacting the insecticide solutions and reduces insecticide carryover.
   5. Set the syringe aside until ready to use for topical application.
4. Create a key containing the doses to be applied and assign random IDs following random number or letter generators (see Supplemental File 1).
5. Label the plastic holding cups with the random ID for blind mortality assessment.
   NOTE: If needed, the protocol can be paused here and restarted at a later day and time. If more than a few hours pass while pausing, it is encouraged to repeat step 3.3 to ensure the syringe is clean and to place the insecticide solutions back in the freezer until about an hour before dosing the insects and then repeat step 3.1.

4. Prepare specimens for the topical bioassay. See Figure 1 for a procedural overview

1. Sort and weigh the mosquitoes
   1. Using an aspirator powered by suction from inhalation, aspirate the desired number of 3-5-day-old adult mosquitoes needed for the assay, including an excess to account for damaged individuals. Transfer the mosquitoes into a conical tube (up to 100 mosquitoes per tube) by placing the tip of the aspirator into the tube with cotton wrapped around the tip and gently exhale and tap the aspirator. Use the cotton to plug the tube when the aspirator tip is removed and then cap with the lid. Avoid filling the aspirator and tubes with too many mosquitoes at once, as this adds additional stress on the mosquitoes and can cause death.
   2. Briefly knock down the mosquitoes in the tubes by placing them for a minimum of 10 min at 4 °C or burying them under ice in an ice tray.
      NOTE: Mosquitoes can be held at 2 °C for several hours with minimal mortality²⁹; however, it is best to minimize the duration for which the mosquitoes are on ice to reduce potential negative effects.
   3. Transfer the knocked down mosquitoes to the insect handling tent and carefully tip the mosquitoes out onto a plastic tray (e.g., Petri dish) placed on the ice. Pour only about 50 mosquitoes at a time to ensure each touches the cool tray beneath it and stays knocked down.
   4. Sort the mosquitoes by sex by gently picking them up by the leg(s) (or wings) with forceps and place each sex into a separate holding cup. Count the number of mosquitoes of each sex while sorting and stop when the desired number is reached. While sorting, remove any mosquitoes that are injured (e.g., missing legs) or are extra-large (e.g., abnormally enlarged abdomen) or small (easily distinguished with the naked eye as smaller than the average mosquito size of that population).
      NOTE: Handling the mosquitoes by the appendages reduces structural damage to their soft primary bodies (e.g., abdomen).
   5. Record the weight of each cup of mosquitos using an analytical scale with 0.1 mg precision.
      1. Place an empty cup with a Petri dish as a lid on the scale and tare the scale. Pour the mosquitoes into the container, place the lid on top, and place the container on the scale.
      2. Record the combined weight and number of specimens on the score sheet (see Supplemental File 2). Immediately place the cup of specimens back on ice to keep them immobilized.
      3. Repeat steps 4.1.5.1-4.1.5.2 until all cups of specimens are weighed.
   6. Divide the prepared mosquitoes into groups of 20-25 in separate cups placed on ice labeled with the random IDs. When transferring mosquitoes, aim to reduce stress and physical damage caused by the forceps. Ideally, pick the mosquitoes up using forceps 1-2 times only: once for sorting/weighing and a potential second time for transfer to the experimental cups.
      NOTE: An ideal number of mosquitoes per cup is 20-25, which is enough for a replicate, is reasonable to assess the mortality, and should not result in density-induced stress/death in the cup.
2. Sort and weigh the fruit flies
   1. Anesthetize the flies using CO₂ for 7 s.
      NOTE: If flies are exposed to CO₂ for more than 7 s, they may have trouble crawling and flying when they awaken³⁰.
   2. Pour the flies onto an ice pack wrapped in bench paper and use a fine-tipped paintbrush to separate and count the males and females.
   3. Use the paintbrush to gently pick up the chosen flies and place them into a clean, empty stock bottle. Choose equal numbers of male and female fruit flies (e.g., 15 males and 15 females) and label the stock bottles with the strain name and fruit fly total (e.g., Canton-S, 30 flies).
      NOTE: It is important to have equal numbers of female and male fruit flies because male fruit flies can experience heightened aggression towards each other after being removed from the presence of females³¹. Therefore, to avoid non-insecticide mortality or injuries, it is best to have equal numbers of males and females (or omit male fruit flies completely).
   4. Record the weight of each bottle of fruit flies using an analytical scale.
      1. Place an empty vial (labeled with a random ID, refer to step 3.4) with a Petri dish as a lid on the scale and tare the scale.
         NOTE: Glass vials are recommended for use with fruit flies as they significantly reduce the static.
      2. Anesthetize the bottle of fruit flies corresponding to the vial's random ID using CO₂ for 7 s.
      3. Pour the fruit flies onto weighing paper and use the paper as a funnel to introduce the flies into the vial. Place the Petri dish lid on top of the vial of fruit flies and place it on the scale.
      4. Record the combined weight and number of specimens on the score sheet and then immediately place the vial of fruit flies in a tray of ice, with the lid still on top to prevent the flies from escaping.
      5. Repeat steps 4.2.4.1-4.2.4.4 for each bottle of fruit flies.
3. When the above steps are complete, immediately move on to the next section.

5. Dose specimens

1. Load the syringe with the proper insecticide concentration. Start with the least concentrated dose and work towards the most concentrated dose with each group of organisms. To prevent waste, only load the syringe with the needed volume of insecticide plus a recommended extra 2 µL.
2. Tip the specimens onto weighing paper(s) placed atop a tray on the ice. Separate the specimens that are close together using a clean, insecticide-free paintbrush or cotton swab to allow easy access to each specimen for dosing. For mosquitoes, use the paintbrush also to ensure that each specimen is laying on their dorsum and their ventral surface is facing up.
3. Using the syringe, apply one droplet of insecticide solution (or acetone for the control) to the ventral thorax and abdomen area for mosquitoes and the dorsum for fruit flies. Apply a 0.2 µL droplet (which requires a 10 µL syringe) for smaller sized insects such as fruit flies and a 0.5 µL droplet (which requires a 25 µL syringe) for mosquitoes.
   NOTE: Insecticide sensitivity does not significantly differ between primary body parts (such as the head, thorax, and abdomen) compared to appendages (such as wings, legs, or proboscis)³². Therefore, the application site does not have to be exact as long as the dose droplet is applied to the primary body. The ventral thorax and abdomen area are chosen for mosquitoes because they often lay on their dorsal side when knocked down, whereas the dorsum is chosen for fruit flies because they often lay on their ventral side when knocked down. This decreased specificity of the application site helps increase the throughput of this method.
4. Immediately pour the specimens back into the labeled plastic cup and cover the cup with netting and a rubber band. Place the cup into a holding tray and note on the cup any specimens that were killed, damaged, or escaped in this process (to exclude them in the final count of specimens in that cup). For the first cup, record the time when dosing is completed.
5. Replace the weighing paper(s) on which the specimens are placed to avoid insecticide contamination between doses.
6. Repeat dosing for each cup until all specimens have been dosed with the proper insecticide concentrations and record the ending time when all specimens have been dosed.
7. Provide 10% sucrose solution to each cup via a soaked cotton ball and set the cups aside until mortality is assessed the following day. Store the mosquitoes at 27 ± 1 °C with 75 ± 5% relative humidity⁵ and the fruit flies at 23 ± 1 °C with 60 ± 5% relative humidity.
   NOTE: Be careful while squeezing the cotton balls to avoid oversaturation or undersaturation. The cotton balls should be moist but not dripping. Dripping sugar water in the cup can lead to mortality of the specimens and thus impact the mortality assessment of the insecticide.

6. Assess mortality

1. Record specimen mortality at 24 h after the start of insecticide exposure. Classify mosquitoes as alive if they can fly and hold themselves upright; as dead if they are immobile or ataxic (unable to stand or take off for flight), as described by the WHO⁶. Follow the same mortality assessment for fruit flies^8,33.
   NOTE: To assess delayed mortality, mortality can additionally be assessed after 48 and 72 h with daily sugar water changes.
2. After mortality is recorded, place all the cups of specimens in a contained bag in a freezer for at least 1 h to ensure all specimens are dead before disposal or subsequent use (e.g., molecular or chemical analysis).

7. Perform replicates

1. Repeat steps 3-6 on a new set of specimens, taking care to perform replicates at the same time each day, as insecticide susceptibility can change depending on the time of day³⁴.
2. Ensure a minimum of 3 replicates for each concentration for accurate estimation of the lethal dose that kills 50% of the specimens (LD₅₀). Include more replicates if a high level of variability is observed.
3. Complete the analysis after all data are collected.

8. Analyze the results

1. Record data in a spreadsheet program and use the random ID key to unmask the data (reference step 3.4). Save the data as a text file (see example data in Supplemental File 3) for analysis in the statistical program R³⁵ (see example R code in Supplemental File 4) or other software of choice³⁶.
2. Within the software program, complete the following analysis. See Supplemental File 4 for an example R code.
   1. Calculate the dose of insecticide (ng) per specimen mass (mg) following Eq (3) below:
      (3)
   2. Calculate mortality and apply Abbott's formula³⁷ to correct mortality relative to the mortality observed in each control³⁷. Alternatively, use the Schneider-Orelli (1947) formula to correct mortality³⁸. With either formula, apply the correction to all data regardless of mortality in each control, as previously described³⁷ and implemented³⁹, unless the control data are unusually high (see discussion below).
      NOTE: Abbott's formula and equivalent alternatives, such as the Schneider-Orelli formula, adjust mortality values proportionately to the extent of mortality not observed in the controls and will not cause a decrease in mortality for cups that had 100% mortality. For more information, see the cited references for these formulas.
   3. Transform corrected mortality data into probit (probability unit) values⁴⁰ and perform linear regression between the insecticide dose and transformed mortality data. Use a chi-square test to assess the fit of the linear model(s).
      NOTE: Mortality values of 0 (0% mortality) or 1 (100% mortality) are removed from the data before completing the probit transformation. This is necessary due to the nature of the probit transformation. As such, the graphed data will not include positive or negative controls or any other data that resulted in 0% or 100% mortality (after Abbott's correction has been applied).
   4. Calculate the LD₅₀ and 95% confidence intervals (CIs) per specimen strain, population, and/or sex following previously published methods^39,41,42.
   5. NOTE: If the 95% CIs of two strains do not overlap, the strains have significantly different dose responses.
   6. If applicable, calculate resistance ratios (RRs) by dividing the LD₅₀ of the strain of interest by the LD₅₀ of the reference/control strain.

Figure 1: Topical application assay protocol diagram. Topical application assay protocol begins with (A) sorting specimens on ice, followed by (B) weighing specimens on an analytical scale, (C) dosing specimens with insecticide solution(s), and (D) 24 h waiting period post insecticide exposure with access to 10% sucrose solution ad libitum (via a soaked cotton ball), followed by mortality assessment. Red arrows indicate target insecticide application location for mosquitoes (left) and fruit flies (right). Note that the image is not to scale. Please click here to view a larger version of this figure.