Studying Mitochondrial Structure and Function in Drosophila Ovaries

The overall goal of this procedure, is to demonstrate methods for studying mitochondrial structure and function in live as well as in fixed tissue in the model organism drosophila melanogaster. This method can help answer key questions in the field of Mitochondrial Biology. Such as the regulatory mechanisms of mitochondrial function.

The main advantage of this technique is that it performs detailed analysis of the mitochondrial structure/function relationship. After sorting out five anesthetized female flies according to the text protocol, under the dissection microscope, use two pairs of forceps to sever the thorax from the abdomen. Then, carefully transfer the abdomens to the second well of the dish.

Using one pair of forceps to hold the abdomen at the posterior end, use the other pair of forceps to slowly push the ovaries out. Then use the forceps to hold an individual ovary by the opaque posterior end and carefully move it to the third well of the dish for teasing or fixing. Carefully tease the protective sheath from around the ovaries, by using a pair of forceps to hold the posterior end and lightly sweeping a teasing needle from the posterior to the anterior end.

To prepare polylysene coated chambers, place two separated 20 microliter drops of polylysene on the cover glass of a glass-bottom petri dish. Air dry the plates at 37 degrees celsius for one hour, and outline the polylysine on the underside of the plate. Set the scanning parameters on the confocal microscope by opening the image acquisition software, and in the acquisition tab, set the appropriate scanning parameters.

Check the time series, bleaching and regions boxes to open the individual tabs. Then, enter the appropriate acquisition parameter values into each tab. Next, place one dissected and teased ovary onto a marked polylysine coated region and use a teasing needle to spread the ovary and separate the ovarials.

Place a 10 micro liter drop of insect dissecting medium on top of the ovary, making sure to cover the entire polylysine coated area. Then, cover the petri dish. Immediately after mounting the ovary, place the sample on the microscope stage and use the eye piece to quickly locate the field of interest in the live tissue.

Click live to acquire a live image of the selected field of interest, then click stop to stop live scanning. If necessary, adjust the acquisition parameters so that the detected fluorescent signal is below the saturation levels with the defined background as set by adjusting the offset values. From the graphic tab, use the bezier drawing tool to draw a small ROI to demarcate the photo bleaching zone on the image acquired by scanning.

Then perform image acquisition, by clicking on start experiment. To carry out live-staining with fluorescent mitochondrial dyes, dilute the stock of stains in warm dissecting medium to final working concentrations. After dissection and teasing is demonstrated earlier, place the ovaries into 200 microliters of staining solution in a well of a dissection dish.

Cover the dish with aluminum foil wrapped box to protect it from light and incubate the tissue for 10 minutes. Next, wash the stained ovaries by using forceps to move them carefully into three consecutive wells containing medium without stain. For co-staining with TMRE, in the compatible overall mitrochondrial stain, carry-out staining with TMRE first, then without washing the tissue, immediately transfer the sample into the overall mitochondrial stain.

After washing and mounting the ovaries as demonstrated earlier in this video, to carry out confocal imaging on the sample, 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 bottom-most z-section. Then move the focus wheel in the opposite direction to define the top-most section.

Perform image acquisition by clicking on start experiment. In the case of live xvivamicroscopy, the data should be obtained within 15 minutes of tissue mounting, otherwise, the experimental results may be affected by alterations of physiology due to insuing death of the tissue. Immediately after dissection under a fume hood place ovaries into 200 microliters of fresh room temperature PFA in the well of a glass dissecting dish.

Incubate the tissue for 15 minutes to fix it. After washing the ovaries in PBS according to the text protocol, while still in PBS, tease the ovaries to carefully remove the protective fiber sheath that may hinder antigen access from the antibody. Permeabilize the teased ovaries by placing them in a microfuse tube with 500 microliters of 0.5 percent tritan x 100 in PBS.

Next, place the tube in a covered box and set to rock at 25 RPM’s for 30 minutes. Remove the PBS/TX according to the text protocol block the tissue by adding 200 microliters of BSA dissolved in 0.5 percent PBS/TX. Incubate the samples on the rocker at room temperature for one hour.

After removing the blocking solution, add anti-rabid desycline E and anti-mouse ATPB antibodies. Incubate the tissue on the rocker at room temperature for two hours. Following the incubation, use PBS-TX to wash the ovaries three times for 15 minutes each.

Then, add 200 microliters of the appropriate secondary antibodies and fresh PBS-TX and incubate at room temperature for one hour. After washing the tissue two times, add hooks to the third wash. Then with a one milliliter micropipet, remove the immuno-stained ovaries from the microfuse tube into a fresh well of a glass dissecting dish containing 200 microliters of PBS.

Add one drop of glycerol based mounting medium to a glass slide and add the immuno-stained ovaries one-by-one to the mounting medium. Under a dissection microscope, while holding the opaque portion of the ovary with forceps, use a teasing needle to gently pluck the transparent ovarials from the opaque mature egg chambers. Then, remove the mature egg chambers from the mounting medium.

Place a cover glass on the slide and lightly press to ensure the mounting medium spreads uniformly. Air dry the samples for 15 minutes before using nail polish to seal the edges. Finally, perform confocal microscopy as needed.

As shown here, flip targeted to one of the mitochondrial clouds associated with the nerse cells leads to greater than 50 percent loss of the fluorescent signal from the blue and white ROI’s surrounding the red flip ROI within 200 seconds while the yellow control ROI’s minimally bleached. The drosophilla ovaries exhibited a decrease in signal with a depth of tissue, and it is important to discern the maximal depth of tissue within which the phlorophores such as TMRE and mitostain, can be detected during live imaging. A qualitative analysis from co-staining with TMRE and the overall mitochondrial stain, shows that the drisophilla germarium exhibits higher mitochondrial potential per unit mass in stage 2B.

In this figure, semi-qualitative analysis demonstrates the difference in mitochondrial potential per unit mass between AFC’s and the MBC’s as analyzed from a stage 9 egg chamber. These images reveal that mitosox uptake occurs in the mitotic and post-mitotic drisophilla follicle cells. In this experiment, co-immuno staining identifies elevated decycline signal overlapping with the mitochondrial ATPB signal and the GFP negative DRP1 nole clones in the Germarium and in a stage 9 egg chamber.

The described methods can be modified and employed to study mitochondrial structure function in other drisophilla tissues. After watching this video, you should have a good understanding of how to dissect, stain and image fixed and live drisophilla ovaries to study mitochonrial structure and function.