Drosophila melanogaster Larva Injection Protocol

1. Fly rearing

NOTE: The D. melanogaster life cycle is divided into four stages: embryo, larva, pupa, and adult. The generation time with optimal rearing conditions in the laboratory (~25 °C, 60% humidity, and sufficient food) is approximately 10 days from fertilized egg to eclosed adult.Females lay ~100 embryos per day, and embryogenesis lasts about 24 h²². The larvae undergo three developmental stages (instars; L1-L3) in ~4 days (L1 and L2: 24 h, and L3: 48 h). The first instar larvae begin to feed immediately on the surface of the medium. Second instar larvae burrow into the medium, whereas third instar larvae leave the medium and wander up the vial walls, looking for a place to pupariate for 24-48 h. The D. melanogaster line used for this protocol is Oregon R (FBsn0000276).

1. Add dry food containing a cornmeal-soy-based diet to a narrow polystyrene vial (25 mm x 95 mm) to aboue 1/5 to 2/5 volume. Then add 9 mL of water and allow the vial to sit for 1 min until the diet is completely hydrated.
2. Add about 10 granules of dry baker's yeast to the vial and place a mixture of at least 20 newly emerged male and female adult flies.
3. Incubate the vial at 25 °C on a 12:12-h light: dark photoperiodic cycle.
4. To ensure life cycle progression, check the fly vials daily and record the initial development stages.

2. Larvae selection for infection

1. Select larvae with a fine paintbrush (camel's hair is best) once they reach the wandering third instar stage on the day the infection will be carried out (Figure 1).
2. Wash the selected larvae with Ringer's solution (100 mM NaCl, 1.8 mM KCl, 2 mM CaCl₂, 1 mM MgCl₂, 5 mM HEPES pH 6.9) in a small Petri dish (60 mm x 15 mm) upon removal from their original vial²³.
3. Place the larvae on filter paper (150 mm diameter) that is slightly moistened with around 5 mL of Ringer's solution in a Petri dish (100 mm x 15 mm) (Figure 2A).
4. Add ~1 mL of Ringer's solution daily to the filter paper as needed to avoid dryness and larval desiccation.

3. Bacterial preparation

1. Culture P. asymbiotica bacteria on Luria Bertani (LB) agar media at 28 °C for 48 h.
2. Use a biosafety level 2 cabinet to transfer a single colony of P. asymbiotica to inoculate a liquid culture in 10 mL of LB media.
3. Incubate the P. asymbiotica liquid culture overnight at 28 °C in a rotary shaker set to 220 rpm.
4. Culture E. coli bacteria in a similar fashion, but carry out the initial growth on agar media and liquid culture incubation at 37 °C overnight.
5. Centrifuge each overnight bacterial culture for 3 min at 13,000 x g and 4 °C.
6. Discard the supernatant and wash the resulting bacterial pellets three times with 10 mL of sterile PBS.
7. Centrifuge the pellets again for 3 min at 13,000 x g and 4 °C.
8. Dilute each pellet with sterile PBS to adjust bacterial concentrations to the optical density (600 nm) of 0.25 for P. asymbiotica and 0.015 for E. coli, which correspond to 100-300 colony forming units per larva of each bacterial preparation, using a spectrophotometer.

4. Injector preparation

1. Prepare capillaries using 3.5″ glass capillary tubes and a micropipette puller. Set the instrument as follows: heat: 580; pull: 143; velocity: 25; delay: 1; pressure: 500.
2. Use straight serrated medium point forceps to open the glass capillary to the degree that will allow the delivery of the experimental treatments while causing minimal damage to the larvae (Figure 2B). To optimize this procedure while practicing injections, use Trypan Blue solution (dilute the stock 0.4% with PBS to 0.2%) to easily track leakage while injecting and the survival of the blue larvae.
3. Fill the capillary with mineral oil using a plastic, disposable 20 mL syringe with a hypodermic needle (22 G, 25 mm length).
4. Set up the nanoliter injector and eject the oil out of the capillary (Figure 2C).
5. Fill the capillary with the desired bacterial preparation for injection. Take the solution from a drop (~20 µL) that is placed on parafilm. In this protocol, 50.2 nL of two bacterial stocks were injected.

5. Larvae injection

1. Anesthetize D. melanogaster larvae using carbon dioxide for ~2 min prior to the procedure.
2. Gently transfer the anesthetized larvae to a filter paper moistened in Ringer's solution in preparation for injection under a stereomicroscope. The larvae will be lethargic or passive at this point.
3. To inject larvae, apply firm pressure on the dorsal side of the posterior end (the tracheal spiracles are apparent at the posterior end versus black mouthparts at the anterior end) using forceps (Figure 2D).
4. Insert the needle horizontally towards the posterior end of the larvae, near the cuticle. A successful injection will not result in a leakage of the injected solution from the larvae (Figure 3).
5. Remove the forceps that were used to apply pressure on the larvae's tail before withdrawing the needle. If not removed first, the forceps can force hemolymph and/or the intestine out of the larvae from the wound site.
6. Infect the appropriate number of larvae for the study being performed. In this particular protocol, 20 larvae for each experimental condition were injected.
7. Using forceps, gently transfer the injected larvae to a separate moist filter paper in a Petri dish (10 larvae per dish) or a food vial (depending on the purpose of the experiment) to allow for recovery.
8. Place the Petri dishes or food vials in an incubator at 25 °C. If larvae are kept in a Petri dish, add Ringer's solution as necessary to prevent desiccation.

6. Recording survival/mortality

1. Record the number of dead and live D. melanogaster larvae according to the set experimental time points. Live larvae will continue to develop into pupae and adults.
2. Use statistical programs, such as Prism, to enter the raw survival/mortality data, analyze them with the log-rank (Mantel-Cox) test, and represent the results in figures.