We acknowledge Leena Patel and Diamond Woodard for helping in the Drosophila medium preparation and Dr. Igor Chesnokov for providing access to the camera-attached stereomicroscope.

1. Preparation of Drosophila (the tools required are depicted in Figure 2A)
<ol>
	<li>For any of the experiments described, collect Drosophila (maintained at room temperature, or 25 &#186;C) 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.</li>
	<li>Sprinkle a small amount of granulated yeast to stimulate Drosophila egg production. Perform experimental manipulation within 2 - 4 days.</li>
</ol>
2. Dissection of Drosophila Ovaries (the tools required are depicted in Figure 2A)
<ol>
	<li>Warm insect dissecting medium (see Materials Table) to room temperature, 25 &#176;C. Fill three wells of an eight-well glass dissecting dish, with 200 &#181;L of medium in each well.</li>
	<li>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.</li>
	<li>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.</li>
	<li>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.</li>
	<li>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).</li>
	<li>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.</li>
</ol>
3. Preparation for Live-tissue Microscopy
NOTE: The tools required are depicted in Figure 2A.
<ol>
	<li>Before Drosophila dissection, prepare polyL-Lysine coated chambers. To this end, place two 20-&#181;L 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 &#186;C 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 &#176;C for a week. If using a previously coated chamber, bring it to room temperature before starting the Drosophila dissection.</li>
	<li>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.</li>
	<li>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-&#181;L drop of insect dissecting medium on top of the ovary, making sure to cover the entire polyL-Lysine-coated area. Cover the petri dish.</li>
	<li>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 &#176;C, which will mimic a heat shock environment for Drosophila tissue.</li>
</ol>
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).
<ol>
	<li>Open the image acquisition software on the confocal microscope and set the appropriate scanning parameters in the &#34;Acquisition&#34; tab (Table 1). Check the &#34;Time series,&#34; &#34;Bleaching,&#34; and &#34;Regions&#34; 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.</li>
	<li>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.</li>
	<li>Click &#34;live&#34; to acquire a live image of the selected field of interest. Click &#34;stop&#34; to stop live scanning.</li>
	<li>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 &#34;range indicator&#34; option is checked), with the defined background as set by adjusting the offset values.</li>
	<li>Draw a small ROI using the Bezier drawing tool from the &#34;Regions&#34; 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.</li>
	<li>Perform the image acquisition by clicking on &#34;start experiment.&#34;</li>
	<li>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.</li>
	<li>Plot the normalized data using any standard plotting software.</li>
</ol>
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).
<ol>
	<li>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 &#956;M.</li>
	<li>After dissection and teasing the ovaries following step 2, place the ovaries into 200 &#181;L 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.</li>
	<li>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).</li>
	<li>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.</li>
	<li>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 &#34;set first&#34; to define the bottommost Z-section. Do the same while moving the focus wheel towards the other direction to define the topmost section.</li>
	<li>Perform the image acquisition by clicking on &#34;start experiment.&#34;</li>
	<li>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.</li>
</ol>
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.
<ol>
	<li>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.</li>
	<li>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.</li>
	<li>Set up a cross by adding the males to the virgin females with a female:male ratio of 2:1.</li>
	<li>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.</li>
	<li>Anesthetize, sort, and collect straight-winged, red-eyed female progeny within 5 days of eclosion.</li>
	<li>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 &#186;C for 1 h, to generate primarily follicle cell clones, and at 37 &#186;C 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.</li>
	<li>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.</li>
	<li>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.</li>
	<li>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.</li>
</ol>
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.
<ol>
	<li>Warm 4% paraformaldehyde (PFA) to room temperature, 25 &#176;C. Caution! Paraformaldehyde is toxic. 
	NOTE: After opening the ampule, PFA should be stored at 4 &#176;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.</li>
	<li>Immediately after the dissection, fix the dissected ovaries by placing them in 200 &#181;L 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 &#181;L of 1x phosphate-buffered saline (PBS). 
	NOTE: The experiment can be stopped here and the samples can be left at 4 &#186;C for 1 day.</li>
	<li>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.</li>
	<li>Permeabilize the teased ovaries by placing them in a microfuge tube with 500 &#181;L 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.</li>
	<li>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.</li>
	<li>Block the ovaries by adding 200 &#181;L 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.</li>
	<li>Add primary antibodies in 200 &#181;L 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.</li>
	<li>Wash the ovaries with PBS-TX 3 times for 15 min each, following step 7.5.</li>
	<li>Add 200 &#181;L 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.</li>
	<li>Wash the ovaries with PBS-TX 3 times for 15 min each, following step 7.5.</li>
	<li>To stain the DNA, add Hoechst (1:1,000 dilution) to the final PBS-TX wash.</li>
	<li>Finally, leave the tissue in 500 &#181;L of 1x PBS.</li>
	<li>Using a 1-mL micropipette, remove the immunostained ovaries from the microfuge tube into a fresh well of a glass dissecting dish containing 200 &#181;L of PBS.</li>
	<li>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.</li>
	<li>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.</li>
	<li>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.</li>
	<li>Air-dry the samples for 15 min and seal the edges of the coverglass carefully with nail polish.</li>
	<li>Perform confocal microscopy as per experimental need (Table 1).</li>
</ol>

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The authors have no competing financial interests.

Critical Steps within the Protocol
Photobleaching: Preventing undue photobleaching of fluorescent samples is absolutely necessary to performing efficient confocal microscopy. Therefore, the time used to locate samples through the eyepiece or to set image acquisition parameters through the live scanning mode should be minimized to minimize photobleaching.
Tissue damage: Since mitochondria are considered to be the sensors of cellular health, it...

Mitochondria are classically described as the cellular powerhouse, since they are the main seats of energy production in differentiated cells. Moreover, mitochondria play a critical role in metabolism, heat generation, lipid modification, calcium and redox homeostasis, the orchestration of cell signaling processes, etc1. Mitochondria also play an active role in the induction of cell death2, as well as in cell cycle regulation3. Such multi-functionality raises the following fundamental questions: a) how do mitochondria perform all these functions simultaneously and b) are there specific mitochondrial pools or subzones that are specialized for distinct functions? In this context, it is important to note that the multifunctional mitochondria are dynamic in their shape, size, and structure within individual cells and that the steady-state shape of mitochondria can vary between cell types. Decades of research from various laboratories suggest that the alteration of mitochondrial shape, size, and structure, collectively called mitochondrial dynamics, is crucial for maintaining various mitochondrial functions4,5,6. These findings raise the possibility that mitochondria may accomplish their multi-functionality by virtue of their structural dynamism.
Extensive efforts are underway to understand the mitochondrial structure-function relationship. The dynamism of mitochondrial structure is primarily maintained by their ability to undergo fission and fusion events with each other. Fission of large mitochondria converts them into smaller mitochondrial elements, while fusion between two smaller mitochondria merges them into a larger mitochondrial element7. Moreover, transient fusion of two mitochondria may occur to allow the mixing of their contents. The fission and fusion events of the inner and outer mitochondrial membranes are carefully governed by specific sets of proteins. The core fission machinery is composed of dynamin-related protein 1 (Drp1), which is recruited from the cytosol to the mitochondria by its interaction with certain bona fide mitochondrial proteins (e.g., Fis1 or Mff1), while Drp1 function can also be regulated by other proteins on the mitochondrial surface4. Although Drp1 operates on the outer membrane, its fission abilities impact the inner membrane as well. The orchestration of the fission of outer and inner mitochondrial membranes is not well understood. On the other hand, fusion of the inner membrane is governed at the core by the activities of Opa1, while mitofusins govern the fusion of the outer-membrane5. The balance of the counteracting fission and fusion events of mitochondria dictate the steady-state mitochondrial shape in a cell. For example, repression of mitochondrial fission would result in complete and unopposed fusion, while the over-activity of mitochondrial fission would result in fragmentation of mitochondria3.
The study of the mitochondrial structure-function relationship primarily involves two complimentary approaches: a) analyses of the cellular and organismal phenotypes after genetic manipulation of mitochondrial fission/fusion proteins and b) direct assessments of mitochondrial structure and function. It is noteworthy that genetic analyses may not always reveal the direct functionality of the molecule at hand (in this case, mitochondrial fission/fusion proteins), as the phenotypes may arise due to secondary effects. Therefore, it is of the utmost importance to develop and use tools to study mitochondrial structure and function directly. Any assessment of mitochondrial structure involves various microscopy tools. Use of fluorescence microscopy of live cells has greatly advanced the studies of mitochondrial dynamics, since mitochondrial dynamism can be monitored both qualitatively and quantitatively using the appropriate fluorescence microscopy tools and techniques8. Fluorescence microscopy-based tools have been developed to study mitochondrial structure and function in live and fixed Drosophila melanogaster tissues, elucidating the significance of mitochondrial dynamism in vivo9. These and related methods are described here, with the goal of studying mitochondrial structure and function in the Drosophila ovary.
The Drosophila ovary consists of germline and somatic lineages, which arise from their respective adult stem cells that reside in the germarium10,11. Sixteen syncytial germ cells (GCs) get encapsulated by somatic follicle cells (FCs) to form individual egg chambers that emerge out of the germarium (Figure 1). One of the 16 GCs get committed to become an oocyte, and the remaining 15 GCs develop into nurse cells that support the growth of the oocyte chamber, facilitating the maturation of the egg before it is laid. The majority of the FCs undergo 9 rounds of mitotic divisions before they exit the mitotic cell cycle to terminally differentiate into a patterned epithelial cell layer consisting of anterior follicle cells (AFCs), posterior follicle cells (PFCs), and main body cells (MBCs). The consecutive egg chambers are connected by stalk cells, which are differentiated cells that are also derived from the FCs early in development. Mitochondrial shape regulated by the mitochondrial fission protein Drp1 is actively involved in the process of differentiation during the normal development of the Drosophila ovarian FC layer9,12. The methods used in these studies to identify the involvement of Drp1 in Drosophila follicle cell layer development are described here.

<table><tbody><tr><td>Grace&#039;s Media (Insect Dissecting Medium)</td><td>Fisher Scientific</td><td>30611031-2</td><td></td></tr><tr><td>41 Paraformaldehyde AQ</td><td>Electronic Microscopy Sciences</td><td>50-259-99</td><td></td></tr><tr><td>Mitotracker Green (overall mitochondrial stain)</td><td>Life Technologies</td><td>m7514</td><td>Reconstitute and Aliquot</td></tr><tr><td>Tetramethylrhodamine ethyl ester perchlorate</td><td>Sigma Aldrich</td><td>87917-25MG</td><td>Reconstitute and Aliquot</td></tr><tr><td>MitoSox (Mito-Ros stain)</td><td>Life Technologies</td><td>m36008</td><td>Reconstitute and Aliquot</td></tr><tr><td>PolyLysine</td><td>MP Biomedicals</td><td>ICN15017625</td><td></td></tr><tr><td>Fly Vials</td><td>Fisher Scientific</td><td>AS-515</td><td></td></tr><tr><td>Fly Conicals</td><td>Fisher Scientific</td><td>AS-355</td><td></td></tr><tr><td>Fly Vial Flugs</td><td>Fisher Scientific</td><td>AS273</td><td></td></tr><tr><td>Fly Conical Flugs</td><td>Fisher Scientific</td><td>AS 277</td><td></td></tr><tr><td>Jazzmix Drosophila food (Drosophila food)</td><td>Fisher Scientific</td><td>AS153</td><td></td></tr><tr><td>Bovine Serum Albumin</td><td>Sigma Aldrich</td><td>A9647-50G</td><td></td></tr><tr><td>Cyclin E Antibody (d-300)</td><td>Santa Cruz</td><td>sc- 33748</td><td></td></tr><tr><td>ATPB antibody [3D5] - Mitochondrial Marker</td><td>AbCam</td><td>ab14730</td><td></td></tr><tr><td>Cy3 AffiniPure Goat Anti-Mouse IgG (H+L)</td><td>Jackson ImmunoResearch</td><td>115-165-146</td><td></td></tr><tr><td>Cy5 AffiniPure Goat Anti-Rabbit IgG (H+L)</td><td>Jackson ImmunoResearch</td><td>111-175-144</td><td></td></tr><tr><td>Hoechst</td><td>Fisher Scientific</td><td>H3570</td><td></td></tr><tr><td>VectaShield</td><td>Fisher Scientific</td><td>H100</td><td></td></tr><tr><td>Azer Scientific EverMark Select Microscope Slides</td><td>Fisher Scientific</td><td>22-026-252</td><td></td></tr><tr><td>Microscope Cover Glass</td><td>Fisher Scientific</td><td>12-542-B</td><td></td></tr><tr><td>Mat Tek Corp&amp;nbsp;Glass Bottom Mircrowell Dish</td><td>Fisher Scientific</td><td>P35G-0-14-C</td><td></td></tr><tr><td>Active Dried Yeast</td><td>Fisher Scientific</td><td>ICN10140001</td><td></td></tr><tr><td>Confocal Microscope</td><td>Carl Zeiss</td><td>LSM 700</td><td></td></tr><tr><td>Dumont #5 Forceps</td><td>Fine Science Technologies</td><td>11251-20</td><td></td></tr><tr><td>Moria Nickel Plated Pin Holder</td><td>Fine Science Technologies</td><td>26016-12</td><td></td></tr><tr><td>Minutien Pins</td><td>Fine Science Technologies</td><td>26002-15</td><td></td></tr><tr><td>MYFP ( w[*]; P{w[+mC]=sqh-EYFP-Mito}3 )</td><td>Bloomington Stock Center</td><td>7194</td><td></td></tr><tr><td>Fly Pad</td><td>Fly stuff</td><td>59-118</td><td></td></tr><tr><td>Blowgun</td><td>Fly stuff</td><td>54-104</td><td></td></tr><tr><td>Blowgun needle</td><td>Flystuff</td><td>54-119</td><td></td></tr><tr><td>Dissecting Microscope</td><td>Carl Zeiss</td><td>Stemi 2000</td><td></td></tr><tr><td>Analyses software</td><td>Carl Zeiss</td><td>Zen&amp;nbsp;</td><td></td></tr><tr><td>Analyses software</td><td>Open source</td><td>Image J</td><td></td></tr><tr><td>Research Macro Zoom Microscope</td><td>Olympus</td><td>MVX10</td><td></td></tr><tr><td>QICAM Fast 1394 Cooled Digital Camera, 12-bit, Mono&amp;nbsp;</td><td>QImaging</td><td>QIC-F-M-12-C</td><td></td></tr><tr><td>QCapture Pro 5.1</td><td>QImaging</td><td></td><td></td></tr></tbody></table>

studying mitochondrial structure and function in drosophila ovaries

Analysis of the mitochondrial structure-function relationship is required for a thorough understanding of the regulatory mechanisms of mitochondrial functionality. Fluorescence microscopy is an indispensable tool for the direct assessment of mitochondrial structure and function in live cells and for studying the mitochondrial structure-function relationship, which is primarily modulated by the molecules governing fission and fusion events between mitochondria. This paper describes and demonstrates specific methods for studying mitochondrial structure and function in live as well as in fixed tissue in the model organism Drosophila melanogaster. The tissue of choice here is the Drosophila ovary, which can be isolated and made amenable for ex vivo live confocal microscopy. Furthermore, the paper describes how to genetically manipulate the mitochondrial fission protein, Drp1, in Drosophila ovaries to study the involvement of Drp1-driven mitochondrial fission in modulating the mitochondrial structure-function relationship. The broad use of such methods is demonstrated in already-published as well as in novel data. The described methods can be further extended towards understanding the direct impact of nutrients and/or growth factors on the mitochondrial properties ex vivo. Given that mitochondrial dysregulation underlies the etiology of various diseases, the described innovative methods developed in a genetically tractable model organism, Drosophila, are anticipated to contribute significantly to the understanding of the mechanistic details of the mitochondrial structure-function relationship and to the development of mitochondria-directed therapeutic strategies.

The described methods can be used to study mitochondrial structure and function in live and fixed Drosophila ovaries (Figure 2B). Provided are some examples of anticipated results obtained with the described methods.
Dissection of the Drosophila ovary: When dissected further, the severed abdomens (Figure 3B) from the whole Drosophila (Figure 3A)...

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Studying Mitochondrial Structure and Function in Drosophila Ovaries

Analysis of the mitochondrial structure-function relationship is required for a thorough understanding of the regulatory mechanisms of mitochondrial functionality. Specific methods for studying mitochondrial structure and function in live and fixed Drosophila ovaries are described and demonstrated in this paper.

Studying Mitochondrial Structure and Function in Drosophila Ovaries

Analysis of the mitochondrial structure-function relationship is required for a thorough understanding of the regulatory mechanisms of mitochondrial ...

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23.9K Views.  University of Alabama at Birmingham. Analysis of the mitochondrial structure-function relationship is required for a thorough understanding of the regulatory mechanisms of mitochondrial functionality. Specific methods for studying mitochondrial structure and function in live and fixed Drosophila ovaries are described and demonstrated in this paper. 

Video: Studying Mitochondrial Structure and Function in Drosophila Ovaries

Dissection of Drosophila Ovaries

In Situ Labeling of Mitochondrial DNA Replication in Drosophila Adult Ovaries by EdU Staining

The mitochondrial genome is inherited exclusively through the maternal line. Understanding of how the mitochondrion and its genome are proliferated and transmitted from one generation to the next through the female oocyte is of fundamental importance. Because of the genetic tractability, and the elegant, ordered simplicity by which oocyte development proceeds, Drosophila oogenesis has become an invaluable system for mitochondrial study. An EdU (5-ethynyl-2&#180;-deoxyuridine) labeling method was utilized to detect mitochondrial DNA (mtDNA) replication in Drosophila ovaries. This method is superior to the BrdU (5-bromo-2'-deoxyuridine) labeling method in that it allows for good structural preservation and efficient fluorescent dye penetration of whole-mount tissues.
Here we describe a detailed protocol for labeling replicating mitochondrial DNA in Drosophila adult ovaries with EdU. Some technical solutions are offered to improve the viability of the ovaries, maintain their health during preparation, and ensure high-quality imaging. Visualization of newly synthesized mtDNA in the ovaries not only reveals the striking temporal and spatial pattern of mtDNA replication through oogenesis, but also allows for simple quantification of mtDNA replication under various genetic and pharmacological perturbations.

In Situ Labeling of Mitochondrial DNA Replication in Drosophila Adult Ovaries by EdU Staining

Drosophila oogenesis continues to be exceptionally useful in the study of mitochondrial proliferation and inheritance. This manuscript describes a detailed protocol used to label the replicating mitochondrial DNA (mtDNA) in Drosophila adult ovaries with 5-ethynyl-2&#180;-deoxyuridine (EdU), which facilitates uncovering mechanisms associated with mitochondrial inheritance that were previously debatable.

The mitochondrial genome is inherited exclusively through the maternal line. Understanding of how the mitochondrion and its genome are proliferated and ...

Genetics

Preparation of Mitochondrial Enriched Fractions for Metabolic Analysis in Drosophila

Since mitochondria play roles in amino acid metabolism, carbohydrate metabolism and fatty acid oxidation, defects in mitochondrial function often compromise the lives of those who suffer from these complex diseases. Detecting mitochondrial metabolic changes is vital to the understanding of mitochondrial disorders and mitochondrial responses to pharmacological agents. Although mitochondrial metabolism is at the core of metabolic regulation, the detection of subtle changes in mitochondrial metabolism may be hindered by the overrepresentation of other cytosolic metabolites obtained using whole organism or whole tissue extractions. 
Here we describe an isolation method that detected pronounced mitochondrial metabolic changes in Drosophila that were distinct between whole-fly and mitochondrial enriched preparations. To illustrate the sensitivity of this method, we used a set of Drosophila harboring genetically diverse mitochondrial DNAs (mtDNA) and exposed them to the drug rapamycin. Using this method we showed that rapamycin modifies mitochondrial metabolism in a mitochondrial-genotype-dependent manner. However, these changes are much more distinct in metabolomics studies when metabolites were extracted from mitochondrial enriched fractions. In contrast, whole tissue extracts only detected metabolic changes mediated by the drug rapamycin independently of mtDNAs.

Preparation of Mitochondrial Enriched Fractions for Metabolic Analysis in Drosophila

Mitochondria play central roles in the regulation of metabolism and homeostasis. Subtle changes in mitochondrial metabolism that affect organismal physiology could be difficult to detect in whole organism metabolomics studies. Here we describe an isolation method that enhances the detection of subtle metabolic shifts in Drosophila melanogaster.

Since mitochondria play roles in amino acid metabolism, carbohydrate metabolism and fatty acid oxidation, defects in mitochondrial function often ...

Developmental Biology

Measurement of Mitochondrial Oxygen Consumption in Permeabilized Fibers of Drosophila Using Minimal Amounts of Tissue

The fruit fly, Drosophila melanogaster, represents an emerging model for the study of metabolism. Indeed, drosophila have structures homologous to human organs, possess highly conserved metabolic pathways and have a relatively short lifespan that allows the study of different fundamental mechanisms in a short period of time. It is, however, surprising that one of the mechanisms essential for cellular metabolism, the mitochondrial respiration, has not been thoroughly investigated in this model. It is likely because the measure of the mitochondrial respiration in Drosophila usually requires a very large number of individuals and the results obtained are not highly reproducible. Here, a method allowing the precise measurement of mitochondrial oxygen consumption using minimal amounts of tissue from Drosophila is described. In this method, the thoraxes are dissected and permeabilized both mechanically with sharp forceps and chemically with saponin, allowing different compounds to cross the cell membrane and modulate the mitochondrial respiration. After permeabilization, a protocol is performed to evaluate the capacity of the different complexes of the electron transport system (ETS) to oxidize different substrates, as well as their response to an uncoupler and to several inhibitors. This method presents many advantages compared to methods using mitochondrial isolations, as it is more physiologically relevant because the mitochondria are still interacting with the other cellular components and the mitochondrial morphology is conserved. Moreover, sample preparations are faster, and the results obtained are highly reproducible. By combining the advantages of Drosophila as a model for the study of metabolism with the evaluation of mitochondrial respiration, important new insights can be unveiled, especially when the flies are experiencing different environmental or pathophysiological conditions.

In this paper, a method to measure oxygen consumption using high-resolution respirometry in permeabilized thoraxes of Drosophila is described. This technique requires a minimal amount of tissue compared to the classic mitochondrial isolation technique and the results obtained are more physiologically relevant.

The fruit fly, Drosophila melanogaster, represents an emerging model for the study of metabolism. Indeed, drosophila have structures homologous to human ...

Biochemistry

Dissection and Staining of Drosophila Pupal Ovaries

Unlike adult Drosophila ovaries, pupal ovaries are relatively difficult to access and examine due to their small size, translucence, and encasing within a pupal case. The challenge of dissecting pupal ovaries also lies in their physical location within the pupa: the ovaries are surrounded by fat body cells inside the pupal abdomen, and these fat cells must be removed to allow for proper antibody staining. To overcome these challenges, this protocol utilizes customized Pasteur pipets to extract fat body cells from the pupal abdomen. Moreover, a chambered coverglass is used in place of a microcentrifuge tube during the staining process to improve visibility of the pupae. However, despite these and other advantages of the tools used in this protocol, successful execution of these techniques may still involve several days of practice due to the small size of pupal ovaries. The techniques outlined in this protocol could be applied to time course experiments in which ovaries are analyzed at various stages of pupal development.

Dissection and Staining of Drosophila Pupal Ovaries

The Drosophila ovary is an excellent model system for studying stem cell niche development. Though methods for dissecting larval and adult ovaries have been published, pupal ovary dissections require different techniques that have not been published in detail. Here we outline a protocol for dissecting, staining, and mounting pupal ovaries.

Unlike adult Drosophila ovaries, pupal ovaries are relatively difficult to access and examine due to their small size, translucence, and encasing within a ...

Confocal and Super-Resolution Imaging of Polarized Intracellular Trafficking and Secretion of Basement Membrane Proteins During Drosophila Oogenesis

The basement membrane (BM) - a specialized sheet of extracellular matrix present at the basal side of epithelial cells - is critical for the establishment and maintenance of epithelial tissue morphology and organ morphogenesis. Moreover, the BM is essential for tissue modeling, serving as a signaling platform, and providing external forces to shape tissues and organs. Despite the many important roles that the BM plays during normal development and pathological conditions, the biological pathways controlling the intracellular trafficking of BM-containing vesicles and how basal secretion leads to the polarized deposition of BM proteins are poorly understood. The follicular epithelium of the Drosophila ovary is an excellent model system to study the basal deposition of BM membrane proteins, as it produces and secretes all major components of the BM. Confocal and super-resolution imaging combined with image processing in fixed tissues allows for the identification and characterization of cellular factors specifically involved in the intracellular trafficking and deposition of BM proteins. This article presents a detailed protocol for staining and imaging BM-containing vesicles and deposited BM using endogenously tagged proteins in the follicular epithelium of the Drosophila ovary. This protocol can be applied to address both qualitative and quantitative questions and it was developed to accommodate high-throughput screening, allowing for the rapid and efficient identification of factors involved in the polarized intracellular trafficking and secretion of vesicles during epithelial tissue development.

Confocal and Super-Resolution Imaging of Polarized Intracellular Trafficking and Secretion of Basement Membrane Proteins During Drosophila Oogenesis

The basement membrane is essential for tissue and organ morphogenesis during development. To better understand the mechanisms leading to proper placement of this structure, the protocol presented describes methods to visualize and characterize the intracellular trafficking and secretion of basement membrane proteins in epithelial cells using confocal and super-resolution microscopy.

The basement membrane (BM) - a specialized sheet of extracellular matrix present at the basal side of epithelial cells - is critical for the establishment ...

Analyzing Mitochondrial Physiology and Bioenergetics in Fruit Flies

Analyzing Mitochondrial Function in a Drosophila melanogaster PINK1B9-Null Mutant Using High-resolution Respirometry

Neurodegenerative diseases, including Parkinson&#39;s Disease (PD), and cellular disturbances such as cancer are some of the disorders that disrupt energy metabolism with impairment of mitochondrial functions. Mitochondria are organelles that control both energy metabolism and cellular processes involved in cell survival and death. For this reason, approaches to evaluate mitochondrial function can offer important insights into cellular conditions in pathological and physiological processes. In this regard, high-resolution respirometry (HRR) protocols allow evaluation of the whole mitochondrial respiratory chain function or the activity of specific mitochondrial complexes. Furthermore, studying mitochondrial physiology and bioenergetics requires genetically and experimentally tractable models such as Drosophila melanogaster.
This model presents several advantages, such as its similarity to human physiology, its rapid life cycle, easy maintenance, cost-effectiveness, high throughput capabilities, and a minimized number of ethical concerns. These attributes collectively establish it as an invaluable tool for dissecting complex cellular processes. The present work explains how to analyze mitochondrial function using the Drosophila melanogaster PINK1B9-null mutant. The pink1 gene is responsible for encoding PTEN-induced putative kinase 1, through a process recognized as mitophagy, which is crucial for the removal of dysfunctional mitochondria from the mitochondrial network. Mutations in this gene have been associated with an autosomal recessive early-onset familial form of PD. This model can be used to study mitochondrial dysfunction involved in the pathophysiology of PD.

Author Spotlight: Exploring Mitochondrial Function and Chemical Toxicity Using Drosophila melanogaster 

Here we present a high-resolution respirometry protocol to analyze bioenergetics in PINK1B9-null mutant fruit flies. The method uses the Substrate-Uncoupler-Inhibitor-Titration (SUIT) protocol.

Neurodegenerative diseases, including Parkinson&#39;s Disease (PD), and cellular disturbances such as cancer are some of the disorders that disrupt energy ...

Studying Mitotic Checkpoint by Illustrating Dynamic Kinetochore Protein Behavior and Chromosome Motion in Living Drosophila Syncytial Embryos

The spindle assembly checkpoint (SAC) mechanism is an active signal, which monitors the interaction between chromosome kinetochores and spindle microtubules to prevent anaphase onset until the chromosomes are properly connected. Cells use this mechanism to prevent aneuploidy or genomic instability, and hence cancers and other human diseases like birth defects and Alzheimer's1. A number of the SAC components such as Mad1, Mad2, Bub1, BubR1, Bub3, Mps1, Zw10, Rod and Aurora B kinase have been identified and they are all kinetochore dynamic proteins2. Evidence suggests that the kinetochore is where the SAC signal is initiated. The SAC prime regulatory target is Cdc20. Cdc20 is one of the essential APC/C (Anaphase Promoting Complex or Cyclosome) activators3 and is also a kinetochore dynamic protein4-6. When activated, the SAC inhibits the activity of the APC/C to prevent the destruction of two key substrates, cyclin B and securin, thereby preventing the metaphase to anaphase transition7,8. Exactly how the SAC signal is initiated and assembled on the kinetochores and relayed onto the APC/C to inhibit its function still remains elusive.
Drosophila is an extremely tractable experimental system; a much simpler and better-understood organism compared to the human but one that shares fundamental processes in common. It is, perhaps, one of the best organisms to use for bio-imaging studies in living cells, especially for visualization of the mitotic events in space and time, as the early embryo goes through 13 rapid nuclear division cycles synchronously (8-10 minutes for each cycle at 25 &deg;C) and gradually organizes the nuclei in a single monolayer just underneath the cortex9.
Here, I present a bio-imaging method using transgenic Drosophila expressing GFP (Green Fluorescent Protein) or its variant-targeted proteins of interest and a Leica TCS SP2 confocal laser scanning microscope system to study the SAC function in flies, by showing images of GFP fusion proteins of some of the SAC components, Cdc20 and Mad2, as the example.

Studying Mitotic Checkpoint by Illustrating Dynamic Kinetochore Protein Behavior and Chromosome Motion in Living Drosophila Syncytial Embryos

The kinetochore is where the SAC initiates its signal monitoring the mitotic segregation of the sister chromatids. A method is described to visualize the recruitment and turnover of one of the kinetochore proteins and its coordination with the chromosome motion in Drosophila embryos using a Leica laser scanning confocal system.

The spindle assembly checkpoint (SAC) mechanism is an active signal, which monitors the interaction between chromosome kinetochores and spindle ...

Techniques for Imaging Prometaphase and Metaphase of Meiosis I in Fixed Drosophila Oocytes

Chromosome segregation in human oocytes is error prone, resulting in aneuploidy, which is the leading genetic cause of miscarriage and birth defects. The study of chromosome behavior in oocytes from model organisms holds much promise to uncover the molecular basis of the susceptibility of human oocytes to aneuploidy. Drosophila melanogaster is amenable to genetic manipulation, with over 100 years of research, community, and technique development. Visualizing chromosome behavior and spindle assembly in Drosophila oocytes has particular challenges, however, due primarily to the presence of membranes surrounding the oocyte that are impenetrable to antibodies. We describe here protocols for the collection, preparation, and imaging of meiosis I spindle assembly and chromosome behavior in Drosophila oocytes, which allow the molecular dissection of chromosome segregation in this important model organism.

Techniques for Imaging Prometaphase and Metaphase of Meiosis I in Fixed Drosophila Oocytes

We present protocols for the collection, preparation, and imaging of mature Drosophila oocytes. These methods allow the visualization of chromosome behavior and spindle assembly and function during meiosis.

Chromosome segregation in human oocytes is error prone, resulting in aneuploidy, which is the leading genetic cause of miscarriage and birth defects. The ...

3D Mitochondrial Ultrastructure of Drosophila Indirect Flight Muscle Revealed by Serial-section Electron Tomography

Mitochondria are cellular powerhouses that produce ATP, lipids, and metabolites, as well as regulate calcium homeostasis and cell death. The unique cristae-rich double membrane ultrastructure of this organelle is elegantly arranged to carry out multiple functions by partitioning biomolecules. Mitochondrial ultrastructure is intimately linked with various functions; however, the fine details of these structure-function relationships are only beginning to be described. Here, we demonstrate the application of serial-section electron tomography to elucidate mitochondrial structure in Drosophila indirect flight muscle. Serial-section electron tomography may be adapted to study any cellular structure in three-dimensions.

3D Mitochondrial Ultrastructure of Drosophila Indirect Flight Muscle Revealed by Serial-section Electron Tomography

In this protocol, we demonstrate the application of serial-section electron tomography to elucidate mitochondrial structure in Drosophila indirect flight muscle.

Mitochondria are cellular powerhouses that produce ATP, lipids, and metabolites, as well as regulate calcium homeostasis and cell death. The unique ...

Studying Mitochondrial Structure and Function in Drosophila Ovaries

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Chapters in this video

Dissection of Drosophila Ovaries

In Situ Labeling of Mitochondrial DNA Replication in Drosophila Adult Ovaries by EdU Staining

Preparation of Mitochondrial Enriched Fractions for Metabolic Analysis in Drosophila

Measurement of Mitochondrial Oxygen Consumption in Permeabilized Fibers of Drosophila Using Minimal Amounts of Tissue

Dissection and Staining of Drosophila Pupal Ovaries

Confocal and Super-Resolution Imaging of Polarized Intracellular Trafficking and Secretion of Basement Membrane Proteins During Drosophila Oogenesis

Analyzing Mitochondrial Function in a Drosophila melanogaster PINK1^B9-Null Mutant Using High-resolution Respirometry

Studying Mitotic Checkpoint by Illustrating Dynamic Kinetochore Protein Behavior and Chromosome Motion in Living Drosophila Syncytial Embryos

Techniques for Imaging Prometaphase and Metaphase of Meiosis I in Fixed Drosophila Oocytes

3D Mitochondrial Ultrastructure of Drosophila Indirect Flight Muscle Revealed by Serial-section Electron Tomography