An In Vivo Assay to Quantify Bacterial Phagocytosis in Adult Drosophila Flies

1. Prepare Fluorescein particles for injection

1. Reconstitute 10 mg of commercially available, heat-killed bacteria particles labeled with fluorescein (see Table of Materials) to a stock concentration of 10 mg/mL by adding 990 µL sterile 1x PBS and 10 µL 50 mM sodium azide. Vortex to mix.
   1. Divide into single-use 8 µL aliquots in 0.2 mL tubes and store in a dark box at 4 °C to minimize light-associated sensitivity.
      NOTE: Sodium azide preservative is optional and can be omitted if 10 mg/mL stocks are made with 1 mL sterile 1x PBS, aliquoted, and stored at -20 °C.
2. Make a 10 mL solution of 5% food coloring in 1x PBS by mixing 500 µL syringe-filtered green food coloring and 9.5 mL sterile 1x PBS.
3. Wash particles before injection to remove sodium azide. Mix 42 µL sterile 1x PBS and 8 µL of 10 mg/mL in a 1.7 mL tube. Centrifuge at max speed for 2.5 min at room temperature.
   1. Remove the supernatant, add 50 µL 1x PBS, and centrifuge at max speed for 2.5 min at room temperature.
   2. Repeat steps 1.3 and 1.3.1 2x, for a total of 3 washes.
   3. After the final wash, discard the supernatant and re-suspend particles to 1.6 mg/mL in 50 µL of 5% food coloring in 1x PBS.
   4. Wrap the tube in aluminum foil to protect it from light. Store at 4 °C, discard after 1 week.

2. Prepare the injection station and flies

1. Prepare the injection pad. To inject up to 4 genotypes of flies at the same time, use laboratory tape to divide a rectangular CO₂ fly pad into 4 sections. On the bench near the microscope, designate areas to place the vials once flies have been lined up on the pad (one for each corner of the pad).
2. Prepare vials of age-matched, 4-7 days-old, flies for injection. For each strain to be tested, transfer 5 males and 5 females into a fresh, labeled vial of prepared fly food and keep it at 25 °C.
3. Prepare the pneumatic injector (see Table of Materials) by setting the instrument to 100 ms (short bursts of gas pressure to expel the liquid – allowing the delivery of sub-nanoliter volumes) TIMED mode.
4. Prepare the microscope slides. Cut 1.5-inch strips of electrical tape, fold them into a loop with the adhesive side out, and place them onto a labeled microscope slide.

3. Prepare glass capillary needles

1. Pull glass needles (thin wall glass capillaries) using a needle puller.
   1. Hold the needle under the microscope with a micrometer and break the tip using #5 fine-point stainless steel tweezers. 100 µm tips are sufficient to pierce the fly's cuticle while minimizing wounding.
   2. Measure the volume of liquid that will be injected into each fly. Load the needle with sterile 5% food coloring in 1x PBS and expel the liquid onto a drop of mineral oil on a 0.01 mm stage micrometer.
      NOTE: If the liquid droplet is spherical, the volume in picoliters is calculated as (size)³/1910. A needle with a 100 µm diameter will eject ~2 nL in 100 ms.

4. Inject flies

1. Pipette 10 µL of 1.6 mg/mL particles onto a small square of parafilm.
   1. Pull the liquid into the needle and mount it in the injector nozzle (see Table of Materials).
   2. Anesthetize flies with CO₂ and line them up in their designated area on the flypad, with the ventral side up and the heads oriented towards the front of the pad. Place vials in corresponding areas on the bench.
   3. Inject flies at the upper corner of the abdomen with 5, 100 ms pumps of liquid (~10 nL total).
   4. Transfer the injected flies to the appropriate vials, note the time on the vial. Keep at 25 °C.
2. Load a new needle with 0.4% Trypan Blue Solution.
3. Set the pneumatic injector to GATED, which allows a constant flow of air to push the liquid out of the needle.
4. Anesthetize flies after they have rested for 30 min and inject with Trypan Blue until the abdomen is full and distended.
   NOTE: When examining phagosome maturation with particles labeled with a pH-sensitive dye, allow flies to rest for 1 h and do not inject with trypan blue before mounting flies.
5. Mount flies on microscope slides with electrical tape, ventral side down. Push the wings to the side of the fly and secure them to the tape. Also, gently push the head into the tape to ensure that the fly will not move.
6. Immediately move to step 5.

5. Imaging flies

1. Image flies, one at a time, at 25x or 32x magnification using an inverted fluorescence microscope attached to a digital camera and computer (see Table of Materials). Focus on the dorsal vessel of the fly using computer software for the digital camera.
2. Record the exposure time and magnification between experiments.
   NOTE: Losing track of genotypes is a potential source of error when photographing multiple strains in a single sitting. To avoid mislabeling flies, make a note of the image number of the first and last fly photograph for each genotype.

6. Quantifying and normalizing the fluorescence

1. Open the software and open one image at a time.
   1. Measure the fluorescence intensity of the dorsal vessel. Draw a polygon around the dorsal vessel. Select Measure and record the fluorescence intensity inside the polygon.
   2. Determine the background fluorescence intensity. Copy the first polygon and move it to an area adjacent to the dorsal vessel of the fly. Select Measure and record fluorescence intensity of the background area.
   3. Normalize the dorsal vessel fluorescence by the background fluorescence:
      Dorsal vessel ÷ background.
   4. Calculate the average normalized dorsal vessel fluorescence intensity of all flies in a strain.
   5. Repeat the experiment 2 more times.
   6. Use a Student's unpaired t-test to compare the mean relative fluorescence intensities of control flies and test flies from the three experiments. Calculate the effect size using the formula: Cohen's d = (M₁-M₂) ÷ SD_pooled, where M₁ is the mean of genotype 1 and M₂ is the mean of genotype 2 &
      SD_pooled =
      where SD1 is the standard deviation of genotype 1 and SD2 is the standard deviation of genotype 2.