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.

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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. 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)...

Watch this Scientific Journal Video about Studying Mitochondrial Structure and Function in Drosophila Ovaries at JoVE.com

Badanie struktury i funkcji mitochondriów w jajnikach Drosophila | Biologia | Wideo

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

Dissection Drosophila Jajniki

Studying Aggression in Drosophila (fruit flies)

Aggression is an innate behavior that evolved in the framework of defending or obtaining resources. This complex social behavior is influenced by genetic, hormonal and environmental factors. In many organisms, aggression is critical to survival but controlling and suppressing aggression in distinct contexts also has become increasingly important. In recent years, invertebrates have become increasingly useful as model systems for investigating the genetic and systems biological basis of complex social behavior. This is in part due to the diverse repertoire of behaviors exhibited by these organisms. In the accompanying video, we outline a method for analyzing aggression in Drosophila whose design encompasses important eco-ethological constraints. Details include steps for: making a fighting chamber; isolating and painting flies; adding flies to the fight chamber; and video taping fights. This approach is currently being used to identify candidate genes important in aggression and in elaborating the neuronal circuitry that underlies the output of aggression and other social behaviors.

Studying Aggression in Drosophila fruit flies

Aggression is an innate behavior that evolved in the framework of defending or obtaining resources.  This complex social behavior is influenced by ...

Microinjection Techniques for Studying Mitosis in the Drosophila melanogaster Syncytial Embryo

This protocol describes the use of the Drosophila melanogaster syncytial embryo for studying mitosis1. Drosophila has useful genetics with a sequenced genome, and it can be easily maintained and manipulated. Many mitotic mutants exist, and transgenic flies expressing functional fluorescently (e.g. GFP) - tagged mitotic proteins have been and are being generated. Targeted gene expression is possible using the GAL4/UAS system2. 

The Drosophila early embryo carries out multiple mitoses very rapidly (cell cycle duration, &asymp;10 min). It is well suited for imaging mitosis, because during cycles 10-13, nuclei divide rapidly and synchronously without intervening cytokinesis at the surface of the embryo in a single monolayer just underneath the cortex. These rapidly dividing nuclei probably use the same mitotic machinery as other cells, but they are optimized for speed; the checkpoint is generally believed to not be stringent, allowing the study of mitotic proteins whose absence would cause cell cycle arrest in cells with a strong checkpoint. Embryos expressing GFP labeled proteins or microinjected with fluorescently labeled proteins can be easily imaged to follow live dynamics (Fig. 1). In addition, embryos can be microinjected with function-blocking antibodies or inhibitors of specific proteins to study the effect of the loss or perturbation of their function3. These reagents can diffuse throughout the embryo, reaching many spindles to produce a gradient of concentration of inhibitor, which in turn results in a gradient of defects comparable to an allelic series of mutants. Ideally, if the target protein is fluorescently labeled, the gradient of inhibition can be directly visualized4. It is assumed that the strongest phenotype is comparable to the null phenotype, although it is hard to formally exclude the possibility that the antibodies may have dominant effects in rare instances, so rigorous controls and cautious interpretation must be applied. Further away from the injection site, protein function is only partially lost allowing other functions of the target protein to become evident.

Microinjection Techniques for Studying Mitosis in the Drosophila melanogaster Syncytial Embryo

This protocol describes the use of microinjection and high resolution imaging in the Drosophila melanogaster syncytial embryo to study mitosis.

This protocol describes the use of the Drosophila melanogaster syncytial embryo  for studying mitosis1.  Drosophila has useful genetics with a sequenced ...

Monitoring Heart Function in Larval Drosophila melanogaster for Physiological Studies

We present various methods to record cardiac function in the larval Drosophila. The approaches allow heart rate to be measured in unrestrained and restrained whole larvae. For direct control of the environment around the heart another approach utilizes the dissected larvae and removal of the internal organs in order to bathe the heart in desired compounds. The exposed heart also allows membrane potentials to be monitored which can give insight of the ionic currents generated by the myocytes and for electrical conduction along the heart tube. These approaches have various advantages and disadvantages for future experiments that are discussed. The larval heart preparation provides an additional model besides the Drosophila skeletal NMJ to investigate the role of intracellular calcium regulation on cellular function. Learning more about the underlying ionic currents that shape the action potentials in myocytes in various species, one can hope to get a handle on the known ionic dysfunctions associated to specific genes responsible for various diseases in mammals.

Monitoring Heart Function in Larval Drosophila melanogaster for Physiological Studies

We present various ways to monitor heart function in the larva of Drosophila for assessing questions dealing with the function of gap junctions, ion channel mutations, modulation of pacemaker activity and pharmacological studies.

We present various methods to record cardiac function in the larval Drosophila. The approaches allow heart rate to be measured in unrestrained  and  ...

The Importance of Correct Protein Concentration for Kinetics and Affinity Determination in Structure-function Analysis 

In this study, we explore the interaction between the bovine cysteine protease inhibitor cystatin B and a catalytically inactive form of papain (Fig. 1), a plant cysteine protease, by real-time label-free analysis using Biacore X100. Several cystatin B variants with point mutations in areas of interaction with papain, are produced. For each cystatin B variant we determine its specific binding concentration using calibration-free concentration analysis (CFCA) and compare the values obtained with total protein concentration as determined by A280. After that, the kinetics of each cystatin B variant binding to papain is measured using single-cycle kinetics (SCK). We show that one of the four cystatin B variants we examine is only partially active for binding. This partial activity, revealed by CFCA, translates to a significant difference in the association rate constant (ka) and affinity (KD), compared to the values calculated using total protein concentration. Using CFCA in combination with kinetic analysis in a structure-function study contributes to obtaining reliable results, and helps to make the right interpretation of the interaction mechanism.

The Importance of Correct Protein Concentration for Kinetics and Affinity Determination in Structure-function Analysis

We apply label-free protein interaction analysis using Biacore X100 for structure-function analysis of the binding of several cystatin B mutants to papain through kinetic characterization. Calibration-free concentration analysis (CFCA) measures the concentration of protein with retained binding activity without the need for a standard curve. We show that confirmation of concentrations using CFCA increases the reliability of the kinetic analysis and that kinetic constants can reliably be determined even if the activity of a recombinant protein is reduced.

In this study, we explore the interaction between the bovine cysteine protease inhibitor cystatin B and a catalytically inactive form of papain (Fig. 1), ...

Patch Clamp and Perfusion Techniques for Studying Ion Channels Expressed in Xenopus oocytes

The protocol presented here is designed to study the activation of the large conductance, voltage- and Ca2+-activated K+ (BK) channels. The protocol may also be used to study the structure-function relationship for other ion channels and neurotransmitter receptors1. BK channels are widely expressed in different tissues and have been implicated in many physiological functions, including regulation of smooth muscle contraction, frequency tuning of inner hair cells and regulation of neurotransmitter release2-6. BK channels are activated by membrane depolarization and by intracellular Ca2+ and Mg2+ 6-9. Therefore, the protocol is designed to control both the membrane voltage and the intracellular solution. In this protocol, messenger RNA of BK channels is injected into Xenopus laevis oocytes (stage V-VI) followed by 2-5 days of incubation at 18&deg;C10-13. Membrane patches that contain single or multiple BK channels are excised with the inside-out configuration using patch clamp techniques10-13. The intracellular side of the patch is perfused with desired solutions during recording so that the channel activation under different conditions can be examined. To summarize, the mRNA of BK channels is injected into Xenopus laevis oocytes to express channel proteins on the oocyte membrane; patch clamp techniques are used to record currents flowing through the channels under controlled voltage and intracellular solutions.

Patch Clamp and Perfusion Techniques for Studying Ion Channels Expressed in Xenopus oocytes

Ionic current of BK channels is recorded using patch clamp techniques. BK channels are expressed in Xenopus oocytes by injecting messenger RNA. The intracellular solution during patch clamp recordings is controlled by a perfusion system.

The protocol presented here is designed to study the activation of the large conductance, voltage- and Ca2+-activated K+ (BK) channels. The protocol may ...

Dissection and Staining of Drosophila Larval Ovaries

Many organs depend on stem cells for their development during embryogenesis and for maintenance or repair during adult life. Understanding how stem cells form, and how they interact with their environment is therefore crucial for understanding development, homeostasis and disease. The ovary of the fruit fly Drosophila melanogaster has served as an influential model for the interaction of germ line stem cells (GSCs) with their somatic support cells (niche) 1, 2. The known location of the niche and the GSCs, coupled to the ability to genetically manipulate them, has allowed researchers to elucidate a variety of interactions between stem cells and their niches 3-12.
Despite the wealth of information about mechanisms controlling GSC maintenance and differentiation, relatively little is known about how GSCs and their somatic niches form during development. About 18 somatic niches, whose cellular components include terminal filament and cap cells (Figure 1), form during the third larval instar 13-17. GSCs originate from primordial germ cells (PGCs). PGCs proliferate at early larval stages, but following the formation of the niche a subgroup of PGCs becomes GSCs 7, 16, 18, 19. Together, the somatic niche cells and the GSCs make a functional unit that produces eggs throughout the lifetime of the organism.
Many questions regarding the formation of the GSC unit remain unanswered. Processes such as coordination between precursor cells for niches and stem cell precursors, or the generation of asymmetry within PGCs as they become GSCs, can best be studied in the larva. However, a methodical study of larval ovary development is physically challenging. First, larval ovaries are small. Even at late larval stages they are only 100&mu;m across. In addition, the ovaries are transparent and are embedded in a white fat body. Here we describe a step-by-step protocol for isolating ovaries from late third instar (LL3) Drosophila larvae, followed by staining with fluorescent antibodies. We offer some technical solutions to problems such as locating the ovaries, staining and washing tissues that do not sink, and making sure that antibodies penetrate into the tissue. This protocol can be applied to earlier larval stages and to larval testes as well.

Dissection and Staining of Drosophila Larval Ovaries

How niches and stem cells form during development is an important question with practical implications. In the Drosophila ovary, germ line stem cells and their somatic niches form during larval development. This video demonstrates how to dissect, stain and mount female gonads from late third instar (LL3) Drosophila larvae.

Many organs depend on stem cells for their development during embryogenesis and for maintenance or repair during adult life. Understanding how stem cells ...

A Protocol for Computer-Based Protein Structure and Function Prediction

Genome sequencing projects have ciphered millions of protein sequence, which require knowledge of their structure and function to improve the understanding of their biological role. Although experimental methods can provide detailed information for a small fraction of these proteins, computational modeling is needed for the majority of protein molecules which are experimentally uncharacterized. The I-TASSER server is an on-line workbench for high-resolution modeling of protein structure and function. Given a protein sequence, a typical output from the I-TASSER server includes secondary structure prediction, predicted solvent accessibility of each residue, homologous template proteins detected by threading and structure alignments, up to five full-length tertiary structural models, and structure-based functional annotations for enzyme classification, Gene Ontology terms and protein-ligand binding sites. All the predictions are tagged with a confidence score which tells how accurate the predictions are without knowing the experimental data. To facilitate the special requests of end users, the server provides channels to accept user-specified inter-residue distance and contact maps to interactively change the I-TASSER modeling; it also allows users to specify any proteins as template, or to exclude any template proteins during the structure assembly simulations. The structural information could be collected by the users based on experimental evidences or biological insights with the purpose of improving the quality of I-TASSER predictions. The server was evaluated as the best programs for protein structure and function predictions in the recent community-wide CASP experiments. There are currently &gt;20,000 registered scientists from over 100 countries who are using the on-line I-TASSER server.

Guidelines for computer based structural and functional characterization of protein using the I-TASSER pipeline is described. Starting from query protein sequence, 3D models are generated using multiple threading alignments and iterative structural assembly simulations. Functional inferences are thereafter drawn based on matches to proteins with known structure and functions.

Genome sequencing projects have ciphered millions of protein sequence, which require knowledge of their structure and function to improve the ...

Functional Reconstitution and Channel Activity Measurements of Purified Wildtype and Mutant CFTR Protein

The Cystic Fibrosis Transmembrane Conductance Regulator (CFTR) is a unique channel-forming member of the ATP Binding Cassette (ABC) superfamily of transporters. The phosphorylation and nucleotide dependent chloride channel activity of CFTR has been frequently studied in whole cell systems and as single channels in excised membrane patches. Many Cystic Fibrosis-causing mutations have been shown to alter this activity. While a small number of purification protocols have been published, a fast reconstitution method that retains channel activity and a suitable method for studying population channel activity in a purified system have been lacking. Here rapid methods are described for purification and functional reconstitution of the full-length CFTR protein into proteoliposomes of defined lipid composition that retains activity as a regulated halide channel. This reconstitution method together with a novel flux-based assay of channel activity is a suitable system for studying the population channel properties of wild type CFTR and the disease-causing mutants F508del- and G551D-CFTR. Specifically, the method has utility in studying the direct effects of phosphorylation, nucleotides and small molecules such as potentiators and inhibitors on CFTR channel activity. The methods are also amenable to the study of other membrane channels/transporters for anionic substrates.

Described here is a rapid and effective procedure for functional reconstitution of purified wild-type and mutant CFTR protein that preserves activity for this chloride channel, which is defective in Cystic Fibrosis. Iodide efflux from reconstituted proteoliposomes mediated by CFTR allows studies of channel activity and the effects of small molecules.

The Cystic Fibrosis Transmembrane Conductance Regulator (CFTR) is a unique channel-forming member of the ATP Binding Cassette (ABC) superfamily of ...

Experimental Approaches to Study Mitochondrial Localization and Function of a Nuclear Cell Cycle Kinase, Cdk1

Although mitochondria possess their own transcriptional machinery, merely 1% of mitochondrial proteins are synthesized inside the organelle. The nuclear-encoded proteins are transported into mitochondria guided by their mitochondria targeting sequences (MTS); however, a majority of mitochondrial localized proteins lack an identifiable MTS. Nevertheless, the fact that MTS can instruct proteins to go into the mitochondria provides a valuable tool for studying mitochondrial functions of normally nuclear and/or cytoplasmic proteins. We have recently identified the cell cycle kinase CyclinB1/Cdk1 complex in the mitochondria. To specifically study the mitochondrial functions of this complex, mitochondrial overexpression and knock-down of this complex without interfering with its nuclear or cytoplasmic functions were essential. By tagging CyclinB1/Cdk1 with MTS, we were able to achieve mitochondrial overexpression of this complex to study its mitochondrial targets as well as functions. Via tagging dominant-negative Cdk1 with MTS, inhibition of Cdk1 activity was accomplished particularly in the mitochondria. Potential mitochondrial targets of CyclinB1/Cdk1 complex were identified using a gel-based proteomics approach. Unlike traditional 2D gel analysis, we employed 2-dimensional difference gel electrophoresis (2D-DIGE) technology followed by phosphoprotein staining to fluorescently label differentially phosphorylated proteins in mitochondrial Cdk1 expressing cells. Identification of phosphoprotein spots that were altered in wild type versus dominant negative Cdk1 bearing mitochondria revealed the identity of mitochondrial targets of Cdk1. Finally, to determine the effect of CyclinB1/Cdk1 mitochondrial localization in cell cycle progression, a cell proliferation assay using a synthetic thymidine analogue EdU (5-ethynyl-2&#8242;-deoxyuridine) was used to monitor the cells as they go through the cell cycle and replicate their DNA. Altogether, we demonstrated a variety of approaches available to study mitochondrial localization and activity of a cell cycle kinase. These are advanced, yet easy to follow methods that will be beneficial to many cell biology researchers.

Here, we outline how to study mitochondrial localization of a (cell cycle) kinase, and how to determine its sub-mitochondrial location as well as potential mitochondrial substrates/targets. Forced expression of proteins into the mitochondria provides a useful tool for studying the functional consequences of mitochondrial localization of a protein of interest.

Studying Mitochondrial Structure and Function in Drosophila Ovaries

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A Protocol for Computer-Based Protein Structure and Function Prediction

Functional Reconstitution and Channel Activity Measurements of Purified Wildtype and Mutant CFTR Protein

Experimental Approaches to Study Mitochondrial Localization and Function of a Nuclear Cell Cycle Kinase, Cdk1