Lipid Droplet Staining of Larval Drosophila Oenocytes: A Technique for Assessment of Lipid Droplet Accumulation Under Normal and Stressed Condition Using BODIPY-Based Dye

1. Starvation Treatment for the Larvae

NOTE: In the third instar larvae, there are few or no detectable lipid droplets in the oenocytes under normal feeding conditions, but numerous large lipid droplets can be induced in the oenocytes under stress conditions, such as starvation. To further verify this method, it is necessary to pretreat these larvae to induce lipid droplet biogenesis in oenocytes. Here, a starvation time course of 12 h, 24 h, and 36 h was chosen as the paradigm. In particular, a short period of starvation (e.g., 3 h) is sufficient to induce detectable lipid droplets in oenocytes. Starvation duration may vary according to specific experimental goals and settings.

1. Make chambers for starvation and control treatments.
   1. For starvation treatment chambers: place a filter paper of appropriate size in a 6 cm Petri dish and pipette 1 mL of PBS onto the filter paper.
   2. For control treatment chambers: place 5 mL of Bloomington standard cornmeal food in the Petri dish.
2. Use a spatula to gently dig up the top layer of food containing larvae that are still burrowing in the food and transfer them to a Petri dish filled with 5 mL of PBS. Gently stir the larvae in PBS to remove any food contamination from the larvae and make it as clean as possible.
3. Use a small paintbrush to collect 40 third instar larvae of the same approximate size. Sort them randomly into a starvation or control chamber, with 20 larvae each.
4. Place the chambers in the incubator at 25 °C with 60% humidity and allow development for 12 h, 24 h, and 36 h of treatment.
   NOTE: For the larvae in the starvation chamber, add 1 mL of PBS every 12 h to avoid larvae dehydration.

2. Dissection of Oenocytes

1. Use a small paintbrush to pick larvae of the appropriate age (12 h, 24 h, or 36 h after treatment), then transfer them into a new Petri dish filled with 5 mL of ice-cold PBS to wash.
   NOTE: Repeat step 2.2 when dealing with larvae in a control chamber to remove any food contamination.
2. Fill a dissection plate with ice-cold PBS and use forceps to gently transfer larvae into the dissection plate. Put the dissection plate under a stereo microscope for the following dissection step.
   NOTE: The ice-cold temperature will help slow movements of the larvae and facilitate dissection.
3. Turn the larvae ventral side up and dorsal side down and gently hold in place using forceps. Secure the larvae to the dissection plate by placing a dissection pin though the pharynx at the anterior end and another pin through the spiracle at the posterior end.
   NOTE: The dorsal side is most easily identified by presence of the dorsal trunks of the trachea.
4. Use Vannas spring scissors to incise (longitudinally) through the epidermis from the anterior to the posterior end.
5. Remove the internal tissue of the epidermis using forceps.
   NOTE: Caution must be taken when removing the tracheal branches to avoid damage to the oenocytes, which are localized in the internal surface of epidermis.
6. With forceps, retrieve the dissection pins and transfer the epidermis into a 1.5 mL microcentrifuge tube filled with PBS on ice.
7. Continue to dissect other larvae following the procedure described above.

3. Lipid Droplet Staining

1. Incubate the dissected epidermis in the fixation bufferfor 30 min at room temperature (RT) on a rotator.
   NOTE: The fixation buffer contains 4% paraformaldehyde (PFA) in PBS.
2. Remove the fixation buffer, followed by a quick wash. To perform a quick wash, add 1 mL of PBS at RT into the tube after the removal of PFA, gently resuspend the tissues, and discard the PBS.
   CAUTION: The fixation buffer contains PFA, which is harmful to human health. It is important to properly dispose the fixation buffer as hazardous waste.
3. Wash the samples 3x for 5 min each with PBS to wash out all possible PFA residue.
4. Incubate the epidermis with BODIPY 493/503 (1 µg/mL; see table of materials) for 30 min at RT on a rotator.
   NOTE: From this step onwards, wrap the microcentrifuge tube with a piece of aluminum foil to protect samples from light and minimize the possible photo-bleaching.
5. Remove the BODIPY 493/503 staining solution and wash the samples 3x for 10 min each with PBS to completely remove residual dyes.

4. Mounting and Imaging

1. Place 6 µL of mounting medium on a clean microscope slide.
   NOTE: Mounting medium is used for longer detection time based on its antifade properties.
2. Use forceps to pick up one epidermis and gently remove the residual PBS with a wipe.
3. Place the epidermis in the mounting medium and adjust its orientation so that its internal surface containing the oenocytes is on the bottom and its external surface is on the top.
4. Gently place a coverslip onto the epidermis.
   NOTE: Remove any extra mounting medium leaking from under the coverslip with a wipe, if necessary. To facilitate imaging, gently push down on the coverslip with forceps so that when observing the slide through the microscope, the oenocyte cluster can be easily imaged in one single plane. Alternatively, it is practicable to cut the epidermis into two semi-epidermis through the middle line to avoid rolling-up of the whole epidermis when mounting the tissues.
5. Apply clear nail polish around the edges of the coverslip to seal.
6. Put the slides in a lightproof sample box and allow the nail polish to dry at RT, which may take 5-10 min.
7. Proceed with microscopic analysis. Take images using a confocal microscope (magnification of 63x with optimized GFP or FITC filter settings, excitation = 488 nm, emission = 503 nm) to acquire clean and sensitive signals with minimized background.