Reproducible dsRNA Microinjection and Oviposition Bioassay in Mosquitoes and House Flies

1. Insect Preparation

1. Anesthetize mosquitoes by chilling them at 4 °C for several hours or by 2 min exposure to CO₂ and then holding at 4 °C prior to injections. Cold anesthetize flies at 4 °C 2‒3 h prior to injections. Ensure mosquitoes and flies are mated if oviposition is a response variable.
   NOTE: The mosquito anesthetization method depends upon the species and strain, e.g., Culex recover much quicker from cold anesthetization than Aedes aegypti and therefore CO₂ knockdown is preferable.
2. On ice, carefully stage mosquitoes or flies by using soft forceps to lay them ventrally exposed on microscope slides ~0.5 cm apart. Standard slides can hold two rows of 6 mosquitoes each or 6–7 house flies. Leave a 1–1.5 cm space at the left or right end of the slide clear of mosquitoes or flies to aid slide handling.
3. Keep slides of staged insects at 4 °C in large Petri dishes until ready for injection.
   NOTE: Drilling a hole through the Petri dish lid will help prevent insects from being disturbed by air displacement when capping. Placing slides on a couple of glass capillaries secured to the dish bottom will ease removal from Petri dish.

2. Insect Injection

1. Prepare dsRNA as described by Estep et al.⁵ and Sanscrainte et al.⁹. If free of contaminants, dsRNA can be quantified by using a spectrophotometer to measure the absorbance at 260 nm and calculating RNA concentration in μg/mL as A₂₆₀ × dilution factor × 40.
2. Set up dissecting microscope and microinjector over a chill table or shallow ice bucket to perform injections on the cold-anesthetized mosquitoes and flies (see step 1.1–1.3 and Figure 1).
3. Pull glass capillaries to a fine tip with needle puller (settings: heater = 15 units, solenoid = 4 A).
   NOTE: See Table of Materials for manufacturer’s details.
4. Place the capillary needle into a microinjector as per the manufacturer’s instructions and break needle tip by pinching with a pair of forceps to produce a sharp point.
   NOTE: Suggested needle opening sizes are ~150 µm for mosquito and ~250 µm for housefly.
5. Set microinjector for desired delivery volume. Meniscus movement from ~50 nL aliquots is easily observable and recommended for mosquito microinjections. If available, enable fast injection settings. Rinse the needle by filling and expelling nuclease-free water 3 times.
6. Prepare solutions for injection at a concentration such that 100‒150 nL will provide desired dose for mosquitoes and 500 nL will provide desired dose for house flies.
   NOTE: Suggested initial doses: 1 µg for each mosquito and 5 µg for each housefly^5,9.
7. Optionally for mosquitoes, add Rhodamine B (to aid in visualization) to solutions at a final concentration of 3.0 µg/mL.
   NOTE: The dye will be clearly visible by its pink color after injection through the nearly clear cuticle on the ventral surface at the abdomen/thorax intersection.
8. Pipette 3–4 µL of solution to inject onto a clean surface (such as a piece of paraffin film) and draw into the glass needle without sucking in any air. Depress the inject button repeatedly until liquid begins to dispense from the needle and gently wipe droplets off with a delicate task wiper.
   NOTE: While some models of microinjectors require backfilling the syringe, the injector referenced in the Table of Materials does not.
9. Using an ultra-fine point marker, draw hash marks ~1 mm apart starting from the liquid meniscus to the needle shank. This will aid in visualizing the meniscus movement and ensure that the solution has entered the mosquito during injection.
10. Set the slide of mosquitoes or flies (as prepared in steps 1.2–1.3) under the needle on the chill table or ice (Figure 1). Ensure that field of view in the microscope is wide enough to see the solution meniscus in the needle.
11. For mosquitoes, align the needle with the middle one-third of the mesokatepisternum (Figure 2A), and while bracing the mosquito against the needle with forceps placed on the opposite side, gently puncture the cuticle with the needle tip using the microinjector micrometer.
12. For house flies, align the needle with the mesopleuron (Figure 2B) and, while bracing the fly against the needle, gently puncture the cuticle with the needle tip.
13. Gently slide the needle into the mosquito or fly body until the tip has passed through the midline.
    NOTE: Inserting the needle too shallow often results in the injected liquid beading out of the insect (see step 2.15).
14. While watching for movement of the liquid meniscus in the needle, depress the inject button until the desired amount of liquid has been injected. For mosquitoes, if set to 50.6 nL aliquots, press 2X for ~100 nL or 3X for ~150 nL. For house flies, depress the inject button until ~500 nL has been injected (i.e., with 69 nL aliquots press 7X for 483 nL).
15. If the meniscus does not move, slowly slide the mosquito or fly off the needle while watching for meniscus movement. If a portion of the injected solution beads out of the cuticle upon needle removal or if the meniscus fails to move, discard the insect as this injection was not successful.
16. Transfer successfully injected mosquitoes in groups of 10-15 or house flies in groups of 5 to clear 3.5 oz holding cups. Cover with tulle or netting and recover at room temperature.
17. After mosquitoes and flies have recovered (1 – 2 h), invert each holding cup over a cotton ball soaked with 10% sucrose solution.
    NOTE: The inversion of the cup on top of the sucrose cotton aids in survival by providing easier access to a sugar meal¹⁰.
18. Rinse needle as in step 2.5 between injection solutions. When done injecting, discard used needles in an appropriate sharps container.

3. Aedes aegypti mortality and oviposition bioassay

1. For three days after injection, continue to provide mosquitoes with 10% sucrose and record the number of visibly dead mosquitoes.
2. Fill a ≤12-inch artificial membrane (such as collagen sausage casing) with fresh blood (i.e., bovine for Aedes aegypti) and heat to 60 °C in hot water bath.
3. Briefly dry the membrane by rolling on a paper towel and lay across the tulle caps of the 3.5 oz clear holding cups. To enhance feeding, spike blood with 1 mM ATP after heating as a phagostimulant or by handle the warmed blood sausage with clean bare hands to leave human volatiles on the casing surface.
4. After blood feeding, replace the cotton ball soaked with 10% sucrose on top of the holding cups and allow mosquitoes to rest for ~24 h before transferring to oviposition cups.
5. Construct oviposition cups by filling clear 3.5 oz bioassay cups with ~30 mL deionized H₂O and placing a piece of seed germination paper (~4 cm wide and 5 cm long with ridges running vertically) at the bottom of the cup and against the side (Figure 3A). Cover with tulle or netting and cut a small slit (~1 cm) in the tulle or netting cap.
6. At 24 h post-blood feeding, cut a small slit (~1 cm) in the tulle cap of the holding cup and transfer only individual females that successfully fed — with a visible blood bolus in abdomen — by gentle mouth aspiration into oviposition cups (1 female/oviposition cup). Seal the small cut in the oviposition cap with a 10% sucrose saturated cotton ball.
7. Hold cups at ~27 °C for 5‒7 days to allow mosquitoes to fully oviposit while tracking daily mortality. Count eggs under a dissecting scope.

4. Musca domestica mortality and oviposition bioassay

1. For three days after injection, continue to provide the flies with 10% sucrose and record the number of visibly dead flies.
2. Three days after injection, anesthetize flies by briefly placing a polyethylene tube dispensing CO₂ over the holding cups until all flies are motionless at bottom of the cups.
3. Transfer flies to a clean cage comprised of a 4 L plastic jar with a stockinette sleeve covering the opening (10 cm, Figure 3B). Provide flies with water and a mixture of granulated sucrose, powdered milk, and dried egg yolk in an 8:8:1 ratio (by volume) for four days, recording and removing dead flies daily.
4. Prepare dry ingredients for fresh fly larval rearing medium by mixing 75% wheat bran with 25% pelleted livestock feed by weight. Add water to reach 62% moisture (until a drop of water can barely be squeezed out).
5. Prepare a 2.5 cm diameter ball from a 1:1 mixture of the above fresh larval media and “used” fly larval medium (medium in which flies have previously pupated). Wrap the ball in a square of black cotton cloth, squeeze lightly until medium liquid seeps through, then use rubber bands to hold the cloth in place around the ball.
6. Put the ball in a 60 mL cup and place it in the fly cage for 5 h (Figure 3B).
7. Rinse eggs off the oviposition ball and ensure that the eggs are removed from under folds in the cloth. Shake eggs to disrupt clusters and transfer them to a graduated 20 mL centrifuge tube with a transfer pipet.
8. Wait until the eggs settle and note the volume of the settled eggs in the graduated tube. Add sufficient water to bring the volume up to 20X the volume of settled eggs. Record the total volume of water plus eggs.
9. While mixing the water and egg suspension with either a magnetic stir bar or vigorous stirring, use a 1 mL pipet tip (with the end of the tip cut off) to dispense 0.5 mL of the water and egg suspension onto a piece of pre-moistened black cloth in a series of lines. Count the eggs under a dissecting microscope.
10. Repeat step 4.9 two more times. If one of the counts is a severe outlier (i.e., differs from the other two counts by >35%), make a fourth count and disregard the outlier.
11. Calculate the mean number of eggs per sample and multiply this value by 2 to get the number of eggs/mL of the suspension. Multiply the eggs/mL value obtained by the volume of the egg suspension from step 4.8. This is the estimate for the total number of eggs laid.