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1. Preparation of Drosophila (the tools required are depicted in Figure 2A)
- For any of the experiments described, collect Drosophila (maintained at room temperature, or 25 ºC) within 5 days of eclosion and place them in a vial filled with 5 - 7 mL of Drosophila food (see Materials Table), with no more than 25 flies in each vial; maintain a female:male ratio of 2:1.
- Sprinkle a small amount of granulated yeast to stimulate Drosophila egg production. Perform experimental manipulation within 2 - 4 days.
2. Dissection of Drosophila Ovaries (the tools required are depicted in Figure 2A)
- Warm insect dissecting medium (see Materials Table) to room temperature, 25 °C. Fill three wells of an eight-well glass dissecting dish, with 200 µL of medium in each well.
- Anesthetize Drosophila with CO2 by placing the needle of the blow gun under the vial plug. Place them on a fly pad. Using a dissecting microscope, sort out 5 females and place them in the first well of the dissecting dish. Handle one Drosophila at a time when performing live microscopy.
- While looking through the eyepiece of the dissection microscope, sever the thorax from the abdomen using two pairs of forceps. Using the forceps, carefully transfer the abdomens to the second well of the dish.
- Use one pair of forceps to hold the abdomen at the posterior end, and slowly push the ovaries out (along with the other abdominal contents) with the other pair of forceps. Should this attempt fail, carefully remove the abdominal exoskeleton by inserting the forceps into the anterior end to release the ovaries.
- Using the forceps, hold an individual ovary by the opaque posterior end (i.e., the yolk-filled, late-stage eggs) and move it carefully to the third well of the dish for teasing to process it for live microscopy (step 3) or for fixing to perform immunostaining (step 7).
- Carefully tease the protective sheath from around the ovaries by sweeping a teasing needle lightly from the posterior to the anterior end of each ovary while holding it by the posterior end with a pair of forceps.
NOTE: To minimize damage during teasing, bend the needle tip and zoom in on each ovary by increasing the magnification of the microscope (Figure 2A). Teasing should be effective enough to break the sheath, but it should also be carefully done to preserve the integrity of the ovarioles.
3. Preparation for Live-tissue Microscopy
NOTE: The tools required are depicted in Figure 2A.
- Before Drosophila dissection, prepare polyL-Lysine coated chambers. To this end, place two 20-µL drops of polyL-Lysine (0.1 mg/mL) on the coverglass (14 mm, No. 0) of a glass-bottomed petri dish (35 mm) at a reasonable distance from each other in order to prevent the merging of the drops. Air-dry the plates for 1 h at 37 ºC and mark the edges of the white polyL-Lysine film on the underside of the plate with an erasable marker.
NOTE: The polyL-Lysine-coated chambers can be stored at 4 °C for a week. If using a previously coated chamber, bring it to room temperature before starting the Drosophila dissection.
- Set the scanning parameters on the confocal microscope to make sure that the sample can be imaged immediately after the completion of the dissection and mounting, as below.
- Place one dissected and teased ovary (following step 2 and stained, if required, following step 5) on a marked polyL-Lysine-coated region and spread the ovary with the teasing needle to separate the ovarioles. Place a 10-µL drop of insect dissecting medium on top of the ovary, making sure to cover the entire polyL-Lysine-coated area. Cover the petri dish.
- Immediately perform confocal microscopy of the mounted sample at room temperature.
NOTE: Only one Drosophila should be dissected, processed, and imaged at a time. Also, microscopy should not be performed at 37 °C, which will mimic a heat shock environment for Drosophila tissue.
4. Fluorescence Loss In Photobleaching (FLIP) Assay to Assess Mitochondrial Matrix Continuity
NOTE: Mitochondrial matrix continuity in a fused mitochondrial structure is established after the complete fusion of the mitochondrial inner and outer membranes following a progression through the intermediate steps. Fission of mitochondria may follow the same steps but in the reverse direction (Figure 3A). FLIP is a time-lapse microscopy-based semi-quantitative method that can be used to assess mitochondrial matrix continuity in the final fused state of ex vivo mitochondria (steps 3 and 4 in Figure 3A) in live Drosophila ovaries9. The FLIP assay is performed as a small region of interest (ROI) of the mitochondria expressing a fluorescent molecule in the mitochondrial matrix that is photobleached at regular intervals (FLIP ROI in Figure 3A). As a result, any surrounding mitochondrial region that is continuous with the FLIP ROI (experimental ROI in Figure 3A) will lose signal due to the exchange of molecules in the continuous mitochondrial matrix. The FLIP experiments demonstrated here are performed on transgenic Drosophila expressing mitoYFP, which contains the mitochondrial targeting sequence of the human cytochrome oxidase VIII subunit tagged with YFP to target it to the mitochondrial matrix in a freely diffusible form. A similar experiment can also be performed with the mito pUASP-mito-GFP transgene, as reported previously9. A similar FLIP protocol may be used with a probe targeted to the mitochondrial inter-membrane space to be able to detect the continuity resulting from the fusion of the outer but not the inner mitochondrial membranes (step 2 in Figure 3A).
- Open the image acquisition software on the confocal microscope and set the appropriate scanning parameters in the "Acquisition" tab (Table 1). Check the "Time series," "Bleaching," and "Regions" boxes to open the individual tabs. Put the appropriate acquisition parameter values in each tab (Table 1).
NOTE: The pinhole should be left open, as this experiment is designed to monitor overall signal from the whole mitochondrial population in individual cells.
- Use the eyepiece to quickly locate the field of interest in the mounted live tissue.
NOTE: Select the ovarioles that are well spread on the glass-bottomed dish, since confocal microscopy cannot be performed on floating ovarioles.
- Click "live" to acquire a live image of the selected field of interest. Click "stop" to stop live scanning.
- If necessary, adjust the acquisition parameters such that the detected fluorescent signal is below the saturation levels (indicated by the absence of red pixels when the "range indicator" option is checked), with the defined background as set by adjusting the offset values.
- Draw a small ROI using the Bezier drawing tool from the "Regions" tab to demarcate the photobleaching zone on the image acquired by live scanning.
NOTE: The size of the ROI should be around 20 - 50% of the total fluorescent mitochondrial signal within the cell.
- Perform the image acquisition by clicking on "start experiment."
- Quantify the fluorescence intensity using the proprietary or the open-source software (see Materials Table). Record the mean signal from the ROI where the repetitive bleaching is targeted (FLIP); the ROIs where bleaching has not been performed in the same cell (Experimental); the ROI from another unbleached cell in the same field of view, for assessing overall bleaching during the experimental period (Bleaching); and the ROI on the background area (Background). Subtract the mean background signal obtained from the mean signal in the other ROIs. Normalize the fluorescent signal with the initial pre-bleach signal for the respective cell.
- Plot the normalized data using any standard plotting software.
5. Live Staining with Fluorescent Mitochondrial Dyes
NOTE: Steady-state mitochondrial structure and potential can be assessed using dyes that specifically incorporate into mitochondria in live cells and tissues. Live Drosophila ovaries can be stained ex vivo with fluorescent mitochondrial stains to visualize the mitochondria, to assess mitochondrial reactive oxygen species (mito-ROS) production, and to assess mitochondrial potential per unit mass. This can be accomplished by co-staining with the mitochondrial potentiometric dye tetramethylrhodamine ethyl ester (TMRE) and a compatible live mitochondrial stain representing the mitochondrial mass (see Materials Table for the specific dyes).
- Dilute the stock of the stains in warm insect dissecting medium to the final working concentrations: mitochondrial stain, 250 nM; TMRE, 50 nM; and mito-ROS stain, 5 μM.
- After dissection and teasing the ovaries following step 2, place the ovaries into 200 µL of any particular staining solution in a well of a dissection dish. Incubate them for 10 min with the dish covered by a suitable box wrapped with aluminum foil to protect it from light. Wash the stained ovaries by moving them carefully with forceps into 3 consecutive wells containing medium without stain.
- For co-staining with TMRE and the compatible overall mitochondrial stain, follow the above protocol to stain first with TMRE and then immediately with the overall mitochondrial stain (without any wash steps in between).
- Mount the ovaries on a polyL-Lysine coated glass-bottomed dish following step 3 and prepare for confocal microscopy with the appropriate scanning parameters (Table 1), following steps 4.2-4.4.
NOTE: The signal from the incorporated dyes did not last when attempting to mount the stained samples in mounting medium.
- Check the Z-sectioning box to open the tab. Turn the focus wheel towards the bottom of the sample while it is being live-scanned and click on "set first" to define the bottommost Z-section. Do the same while moving the focus wheel towards the other direction to define the topmost section.
- Perform the image acquisition by clicking on "start experiment."
- Quantify the background-corrected fluorescent intensity from the ROIs for the background signal and the individual cells (as in step 4.7) and plot the data using any plotting software.
6. Generation of Drp1 Null Mosaics
NOTE: The clonal strategy used here introduces green fluorescent protein (GFP)-negative Drp1 null clones in the background of a GFP-positive, phenotypically wild-type background that is genotypically heterozygous for the Drp1 null mutation9. Heat shock-induced flippase-flippase recognition target (FLP-FRT)-mediated site-specific mitotic recombination creates homozygous clones of the functionally null drpKG03815 allele. The genotype of Drosophila carrying the Drp1 mutant is drpKG03815 FRT40A/CYO, whereas the genotype carrying the heat shock-induced FLP (hsFLP) and UbiGFP clonal marker is hsflp; ubiquitin nls-GFP (UbiGFP) FRT 40A/CyO. The genotype of the selected offspring of the cross between the above genotypes is hsFLP/+; drpKG03815FRT40A/UbiGFPFRT40A.
- Synchronize the Drosophila for virgin collection by moving them into new vials of fresh food every 2 to 3 days. Monitor the pupariating vials daily in order to collect emerging virgin females.
- Collect red-eyed, curly-wing virgin females from the Drp1 mutant genotype every day, once in the morning and once in the evening, and place them into a separate vial. In parallel, collect male Drosophila with dark red eyes and straight wings carrying hsflp and UbiGFP within 5 days of eclosion.
- Set up a cross by adding the males to the virgin females with a female:male ratio of 2:1.
- Sprinkle a small amount of granulated yeast to stimulate egg production.
NOTE: Carefully move the Drosophila to a fresh vial of media every 2 to 3 days to increase the amount of progeny and to reduce vial crowding.
- Anesthetize, sort, and collect straight-winged, red-eyed female progeny within 5 days of eclosion.
- For the heat shock, place the collected Drosophila into empty vials with a small amount of granulated yeast and a small, soft wipe (to absorb the moisture during the heat shock). Place the vial with the Drosophila in a water bath at 38 ºC for 1 h, to generate primarily follicle cell clones, and at 37 ºC for 1 h twice a day (allowing at least 5 h between the 2 heat shocks) for 2 consecutive days, to generate both germline and follicle cell clones.
NOTE: Make sure that the vial is completely submerged in the water up to the level of the plug.
- The heat shock may make the Drosophila immobile. Allow the heat-shocked Drosophila to recover for 1 h at room temperature when they become mobile again.
- Add the males back to the females in the same proportion as in the cross, and move them to fresh vials with medium sprinkled with a small amount of granulated yeast. Maintain the Drosophila for at least 5 days.
- Dissect the ovaries as necessary for a live or fixed experiment.
NOTE: Ovaries isolated from the parental Drosophila expressing UbiGFP should be used as negative controls to confirm the efficient induction of GFP-negative clones by the heat shock.
7. Co-immunostaining for Cyclin E and Mitochondria
NOTE: To detect Drosophila Cyclin E (dCyclinE), we have used a commercially obtained antibody raised specifically against dCyclinE9 (see Materials Table). As a mitochondrial marker, we used an antibody against ATP-B (a subunit of the mitochondrial ATP synthase complex)9.
- Warm 4% paraformaldehyde (PFA) to room temperature, 25 °C. Caution! Paraformaldehyde is toxic.
NOTE: After opening the ampule, PFA should be stored at 4 °C and used within 7 days. This is because storage of PFA may allow oxidation to methanol, which would dramatically alter mitochondrial membranes during fixation, even if present in trace amounts.
- Immediately after the dissection, fix the dissected ovaries by placing them in 200 µL of fresh PFA in a well of a glass dissecting dish (without teasing). Keep the dish in a fume hood for 15 min. Wash the fixed ovaries by moving them carefully with the forceps into 3 consecutive wells, each containing 200 µL of 1x phosphate-buffered saline (PBS).
NOTE: The experiment can be stopped here and the samples can be left at 4 ºC for 1 day.
- Tease the ovaries more thoroughly in PBS (similar to step 2.6) to carefully remove the protective fibrous sheath that may hinder antigen access by the antibody.
- Permeabilize the teased ovaries by placing them in a microfuge tube with 500 µL of freshly made 0.5% PBS-Triton-X100 (PBS-TX) and incubate it for 30 min while rocking at 25 rpm.
NOTE: During the rocking, place the tubes parallel to the rocking movement; placing them perpendicularly may allow the tissue to stick to the cap of the tubes, thus resulting in tissue loss.
- To remove the PBS-TX, place the microfuge tubes in a rack to allow the tissue to settle down at the bottom of the tubes. Inspect them visually and tap the tubes, if necessary, to bring any floating tissues down to the bottom. Aspirate the solution carefully from the top, making sure that the tissue at the bottom of the tubes remains undisturbed.
- Block the ovaries by adding 200 µL of 2% bovine serum albumin (BSA) dissolved in 0.5% PBS-TX, and incubate it on the rocker at room temperature for 1 h. Remove the blocking agent by following step 7.5.
- Add primary antibodies in 200 µL of fresh blocking agent: anti-rabbit dCyclinE antibody (1:100) and anti-mouse ATP-B antibody (1:100). Incubate the tissues on the rocker for 2 h at room temperature.
- Wash the ovaries with PBS-TX 3 times for 15 min each, following step 7.5.
- Add 200 µL of the appropriate secondary antibodies in fresh PBS-TX: anti-mouse-CY3 (1:1,000) and anti-rabbit-CY5 (1:500). Incubate on the rocker for 1 h at room temperature.
- Wash the ovaries with PBS-TX 3 times for 15 min each, following step 7.5.
- To stain the DNA, add Hoechst (1:1,000 dilution) to the final PBS-TX wash.
- Finally, leave the tissue in 500 µL of 1x PBS.
- Using a 1-mL micropipette, remove the immunostained ovaries from the microfuge tube into a fresh well of a glass dissecting dish containing 200 µL of PBS.
- Add one drop of glycerol-based mounting medium to a glass slide and add the immunostained ovaries one by one to the mounting medium.
NOTE: Make sure that the ovaries are indeed transferred to the mounting medium. Failing to do so will lead to tissue loss.
- While looking through the dissection microscope, gently pluck the transparent ovarioles (younger stages) from the opaque mature egg chambers using the teasing needle while holding the opaque portion of the ovary with the forceps. Remove the mature egg chambers from the mounting medium.
- Place a coverglass (22 mm, No. 1) on the slide and press lightly to ensure that the mounting medium is spread uniformly underneath.
NOTE: Pressing on the coverglass also ensures the proper alignment of the tissue along the coverglass. Failing to do so optimally may allow the smaller stages to float in the mounting medium, preventing optimal microscopy of those tissues.
- Air-dry the samples for 15 min and seal the edges of the coverglass carefully with nail polish.
- Perform confocal microscopy as per experimental need (Table 1).