Name: dissection and lipid droplet staining of oenocytes in drosophila larvae
Uploaded: 2019-12-28T13:00:00.000+00:00
Duration: 5 min 37 s
Description: Watch this Scientific Journal Video about Dissection and Lipid Droplet Staining of Oenocytes in Drosophila Larvae at JoVE.com

This work was supported by grants from the National Natural Science Foundation of China (31671422, 31529004, and 31601112), the 111 Project (D18010), the Local Innovative and Research Teams Project of Guangdong Perl River Talents Program (2017BT01S155), and the China Postdoctoral Science Foundation (2018M640767).

1. Egg Laying
<ol>
	<li>Prepare the standard cornmeal food for egg laying. 
	NOTE: For the recipe and cooking procedure for standard cornmeal food used here, see the previously published details15.</li>
	<li>Prepare fresh yeast paste by adding 6 mL of distilled water to 4 g of active dried yeast in a 50 mL centrifugal tube. Use a spatula to mix and make a paste.</li>
	<li>Make egg-laying bottles by filling the cornmeal food into the bottles and spread approximately 1 g of yeast paste onto the surface of the cornmeal food with a spatula.</li>
	<li>Place the flies of desired genotypes in an egg-laying bottle and place it in an incubator with a constant temperature of 25 &#176;C and humidity of 60%. 
	NOTE: Ideal crosses usually consist of 150 virgin flies and at least 75 males. Keeping flies in dark by placing a lightproof box over the egg-laying bottles will increase the reproduction rate.</li>
	<li>Before egg collection, let flies lay eggs for 1 h to allow removal of all old eggs that remain stored in the female oviducts.</li>
	<li>Let flies lay eggs in a new egg-laying bottle for 1 h and remove the adults from the bottle. 
	NOTE: Control the egg-laying time to minimize the developmental range in order to obtain larvae within a precisely controlled developmental stage.</li>
	<li>Allow the eggs to develop for 84 h into the third instar larvae in the incubator at 25 &#176;C with a 12 h/12 h light/dark cycle.</li>
</ol>
2. Starvation Treatment for the Larvae
NOTE: As mentioned above, in the third instar larvae, there are few or no detectable lipid droplets in the oenocytes under normal feeding conditions, but numerous large lipid droplets can be induced in the oenocytes under stress conditions, such as starvation. To further verify this method, it is necessary to pretreat these larvae to induce lipid droplet biogenesis in oenocytes. Here, a starvation time course of 12 h, 24 h, and 36 h was chosen as the paradigm. In particular, a short period of starvation (e.g., 3 h) is sufficient to induce detectable lipid droplets in oenocytes. Starvation duration may vary according to specific experimental goals and settings.
<ol>
	<li>Make chambers for starvation and control treatments.
	<ol>
		<li>For starvation treatment chambers: place a filter paper of appropriate size in a 6 cm Petri dish and pipette 1 mL of PBS onto the filter paper.</li>
		<li>For control treatment chambers: place 5 mL of Bloomington standard cornmeal food in the Petri dish.</li>
	</ol>
	</li>
	<li>Use a spatula to gently dig up the top layer of food containing larvae that are still burrowing in the food and transfer them to a Petri dish filled with 5 mL of PBS. Gently stir the larvae in PBS to remove any food contamination from the larvae and make it as clean as possible.</li>
	<li>Use a small paintbrush to collect 40 third instar larvae of the same approximate size. Sort them randomly into a starvation or control chamber, with 20 larvae each.</li>
	<li>Place the chambers in the incubator at 25 &#176;C with 60% humidity and allow development for 12 h, 24 h, and 36 h of treatment. 
	NOTE: For the larvae in the starvation chamber, add 1 mL of PBS every 12 h to avoid larvae dehydration.</li>
</ol>
3. Dissection of Oenocytes
<ol>
	<li>Use a small paintbrush to pick larvae of the appropriate age (12 h, 24 h, or 36 h after treatment), then transfer them into a new Petri dish filled with 5 mL of ice-cold PBS to wash. 
	NOTE: Repeat step 2.2 when dealing with larvae in a control chamber to remove any food contamination.</li>
	<li>Fill a dissection plate with ice-cold PBS and use forceps to gently transfer larvae into the dissection plate. Put the dissection plate under a stereo microscope for the following dissection step. 
	NOTE: The ice-cold temperature will help slow movements of the larvae and facilitate dissection.</li>
	<li>Turn the larvae ventral side up and dorsal side down and gently hold in place using forceps. Secure the larvae to the dissection plate by placing a dissection pin though the pharynx at the anterior end and another pin through the spiracle at the posterior end. 
	NOTE: The dorsal side is most easily identified by presence of the dorsal trunks of the trachea.</li>
	<li>Use Vannas spring scissors to incise (longitudinally) through the epidermis from the anterior to the posterior end.</li>
	<li>Remove the internal tissue of the epidermis using forceps. 
	NOTE: Caution must be taken when removing the tracheal branches to avoid damage to the oenocytes, which are localized in the internal surface of epidermis.</li>
	<li>With forceps, retrieve the dissection pins and transfer the epidermis into a 1.5 mL microcentrifuge tube filled with PBS on ice.</li>
	<li>Continue to dissect other larvae following the procedure described above.</li>
</ol>
4. Lipid Droplet Staining
<ol>
	<li>Incubate the dissected epidermis in the fixation bufferfor 30 min at room temperature (RT) on a rotator. 
	NOTE: The fixation buffer contains 4% paraformaldehyde (PFA) in PBS.</li>
	<li>Remove the fixation buffer, followed by a quick wash. To perform a quick wash, add 1 mL of PBS at RT into the tube after the removal of PFA, gently resuspend the tissues, and discard the PBS. 
	CAUTION: The fixation buffer contains PFA, which is harmful to human health. It is important to properly dispose the fixation buffer as hazardous waste.</li>
	<li>Wash the samples 3x for 5 min each with PBS to wash out all possible PFA residue.</li>
	<li>Incubate the epidermis with BODIPY 493/503 (1 &#181;g/mL; see Table of Materials) for 30 min at RT on a rotator. 
	NOTE: From this step onwards, wrap the microcentrifuge tube with a piece of aluminum foil to protect samples from light and minimize the possible photo-bleaching.</li>
	<li>Remove the BODIPY 493/503 staining solution and wash the samples 3x for 10 min each with PBS to completely remove residual dyes.</li>
</ol>
5. Mounting and Imaging
<ol>
	<li>Place 6 &#181;L of mounting medium on a clean microscope slide. 
	NOTE: Mounting medium is used for longer detection time based on its antifade properties.</li>
	<li>Use forceps to pick up one epidermis and gently remove the residual PBS with a wipe.</li>
	<li>Place the epidermis in the mounting medium and adjust its orientation so that its internal surface containing the oenocytes is on the bottom and its external surface is on the top.</li>
	<li>Gently place a coverslip onto the epidermis. 
	NOTE: Remove any extra mounting medium leaking from under the coverslip with a wipe, if necessary. To facilitate imaging, gently push down on the coverslip with forceps so that when observing the slide through the microscope, the oenocyte cluster can be easily imaged in one single plane. Alternatively, it is practicable to cut the epidermis into two semi-epidermis through the middle line to avoid rolling-up of the whole epidermis when mounting the tissues.</li>
	<li>Apply clear nail polish around the edges of the coverslip to seal.</li>
	<li>Put the slides in a lightproof sample box and allow the nail polish to dry at RT, which may take 5-10 min.</li>
	<li>Proceed with microscopic analysis. Take images using a confocal microscope (magnification of 63x with optimized GFP or FITC filter settings, excitation = 488 nm, emission = 503 nm) to acquire clean and sensitive signals with minimized background.</li>
</ol>

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A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Factors+underlying+the+variability+of+lipid+droplet+fluorescence+in+MA-10+Leydig+tumor+cells.">Factors underlying the variability of lipid droplet fluorescence in MA-10 Leydig tumor cells.</a> Cytometry. 17 (2), 151-158 (1994).</li><li>Fam, T. K., Klymchenko, A. S., Collot, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Recent+advances+in+fluorescent+probes+for+lipid+droplets.">Recent advances in fluorescent probes for lipid droplets.</a> Materials (Basel). 11 (9), (2018).</li><li>BDSC Standard Cornmeal Medium. Bloomington Drosophila Stock Center Available from: <a target="_blank" href="https://bdsc.indiana.edu/information/recipes/bloomfood.html">https://bdsc.indiana.edu/information/recipes/bloomfood.html</a> (2019)</li><li>Milon, A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Do+estrogens+regulate+lipid+status+in+testicular+steroidogenic+Leydig+cell?.">Do estrogens regulate lipid status in testicular steroidogenic Leydig cell?.</a> Acta Histochemica. 121 (5), 611-618 (2019).</li><li>Farmer, B. C., Kluemper, J., Johnson, L. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Apolipoprotein+E4+alters+astrocyte+fatty+acid+metabolism+and+lipid+droplet+formation.">Apolipoprotein E4 alters astrocyte fatty acid metabolism and lipid droplet formation.</a> Cells. 8 (2), (2019).</li></ol>

The authors have no conflicts of interest to disclose.

Among those outlined above, there are several critical steps in this protocol, with the egg laying time period being one of these. As a lipid-mobilizing tissue, the oenocyte is highly sensitive to nutrition status6,8. Prolonged egg-laying time periods may result in crowed larvae and increased food competition, leading to inaccurate results. The 1 h egg-laying time period used in this protocol allows the larvae to develop without nutrition competition. A much larg...

Lipids are essential for cell survival. In addition to their traditional role as integral components of cellular membrane systems, lipids also play crucial functions in energy supply and signaling transduction throughout the life cycles of individual animals1. Thus, lipid metabolism must conform to strict regulations to maintain physiological hemostasis in cells. It is known that dysregulation of lipid metabolism results in various diseases, such as diabetes and fatty liver. Despite the great importance of lipid metabolism in animal health, the mechanisms underlying lipid metabolism regulation remain largely unknown.
Drosophila has been extensively used for years since professor Thomas H. Morgan started to use them in studies involving genetics and other basic biological questions2. In the last few decades, emerging evidence has shown that Drosophila is an excellent model organism in the study of many lipid metabolism-associated diseases, such as obesity1,3. In particular, Drosophila shares highly conserved metabolic genes with humans and possess similar relevant tissues/organs and cell types for lipid metabolism.
For example, the fat body of Drosophila, which is responsible for triacylglyceride storage, functions analogous to human adipose tissue. Recently, a cluster of specialized hepatocyte-like cells (i.e., oenocytes), which have been reported to be a functional analog to the human liver, have been shown to be involved in fatty acid and hydrocarbon metabolism in fruit flies4,5. Similar to the case in mammal livers, oenocytes respond to starvation by activating lipid droplet formation in both larval and adult Drosophila, resulting in lipid droplet accumulation in oenocytes4,6,7,8. Anatomically, oenocytes are tightly attached to the basal internal surface of the lateral epidermis in clusters of approximately six cells per abdominal hemisegment, which makes it impracticable to isolate oenocyte clusters from the epidermis. Thus, oenocytes must be attached to the epidermis during dissection and staining.
Lipids are stored in the form of lipid droplets, which are organelles with single layer membranes in cells9. Lipid droplets exist in almost all cell types across different species10. Lipid droplet dynamics, including its size and number, change in response to environmental stressors. This is regarded as a reflection of metabolic status in response to stress, such as aging and starvation7,8. Therefore, it is of great importance to develop a feasible and reliable method to qualitatively determine lipid droplet dynamics in oenocytes during development and under stressful conditions. In particular, in the third instar larvae, oenocytes contain few or no detectable lipid droplets under fed conditions, but they do contain numerous large lipid droplets after nutrition deprivation4. To verify the effectiveness of this method, it is suggested to perform lipid droplet staining in the oenocytes under starved conditions.
Currently, several lipophilic dyes are available for lipid droplets staining, such as the nonfluorescent dyes Sudan Black and Oil Red O and fluorescent dyes Nile Red and BODIPY 493/50311. Sudan Black and Oil Red O are commonly used for tissue cholesteryl esters and triacylglycerols and can be easily detected by light microscopy. However, relatively high background staining and relatively low resolution are two limiting factors for its applications in qualitative analysis of lipid droplet dynamics. To overcome the limitations of nonfluorescent dyes, Nile Red and BODIPY 493/503 are utilized as ideal substitutes for lipid droplet staining. It has been reported that Nile Red can also detect some unesterified cholesterol, which makes BODIPY 493/503 a more specific dye for cellular lipid droplets, to some extent12,13,14.
Above all, to fulfill a need for rapid and sensitive analysis of lipid droplets in oenocytes, this protocol presents a feasible and highly reproducible method of fixative-based lipid droplet-specific staining using BODIPY 493/503 as the stain dye. In this report, oenocytes are dissected, and BODIPY 493/503 is used for lipid droplet staining in the oenocytes, in which lipid droplets are detected by confocal microscopy. The ease and affordability of this procedure make it ideal for modification and further use in other applications, such as flow cytometry.

<table><tbody><tr><td>50 mL centrifuge tube</td><td>Corning</td><td>430829</td><td>50 mL</td></tr><tr><td>6 cm Petri dish</td><td>Thermo Fisher</td><td>150326</td><td>6 cm</td></tr><tr><td>Agar</td><td></td><td></td><td>For fly food</td></tr><tr><td>Aluminum foil</td><td>N/A</td><td>N/A</td><td>Protect smaple from light</td></tr><tr><td>BODIPY 493/503</td><td>Invitrogen</td><td>D3922</td><td>Lipid droplet staining dye</td></tr><tr><td>Confocal microscope</td><td>Leica</td><td>Leica TSC SP5</td><td>Confocal imaging</td></tr><tr><td>Corn syrup</td><td></td><td></td><td>For fly food</td></tr><tr><td>Cornmeal</td><td></td><td></td><td>For fly food</td></tr><tr><td>Coverslip</td><td>Citoglas</td><td>10212424C</td><td>20 &amp;times; 20 mm, 0.13-0.17 thick</td></tr><tr><td>Dissection pin</td><td>N/A</td><td>N/A</td><td></td></tr><tr><td>Dissection plate</td><td>N/A</td><td>N/A</td><td></td></tr><tr><td>Filter paper</td><td>N/A</td><td>N/A</td><td>Diameter: 11 cm</td></tr><tr><td>Fixation buffer</td><td>N/A</td><td>N/A</td><td>4% Paraformaldehyde (PFA) in 1x PBS</td></tr><tr><td>Forcep</td><td>Dumont</td><td>11252-30</td><td>#5</td></tr><tr><td>Incubator</td><td>Jiangnan</td><td>SPX-380</td><td>For fly culture</td></tr><tr><td>Microcentrifuge tube</td><td>Axygen</td><td>MCT-150-C</td><td>1.5 mL</td></tr><tr><td>Microscopy slide</td><td>Citoglas</td><td>10127105P-G</td><td></td></tr><tr><td>Mounting medium</td><td>VECTASHIELDAntifade Mounting Medium</td><td>H-1000</td><td>Antifade mounting medium</td></tr><tr><td>Nail polish</td><td>PanEra</td><td>AAPR419</td><td>Seal the coverslip</td></tr><tr><td>Paintbrush</td><td>N/A</td><td>N/A</td><td></td></tr><tr><td>PBS</td><td>N/A</td><td>N/A</td><td>1x PBS (137 mM NaCl, 2.7 mM KCl, 10 mM Na&lt;sub&gt;2&lt;/sub&gt;HPO&lt;sub&gt;4&lt;/sub&gt;,1.8 mM KH&lt;sub&gt;2&lt;/sub&gt;PO&lt;sub&gt;4&lt;/sub&gt;, pH 7.4)</td></tr><tr><td>Rotator</td><td>Kylin-Bell Lab Instruments</td><td>WH-986</td><td></td></tr><tr><td>Scissor</td><td>Smartdata Medical</td><td>SR81</td><td>Vannas spring scissor</td></tr><tr><td>Soy flour</td><td></td><td></td><td>For fly food</td></tr><tr><td>Spatula</td><td>N/A</td><td>N/A</td><td></td></tr><tr><td>Standard cornmeal food</td><td>N/A</td><td>N/A</td><td>Accoding to Bloomington standard cornmeal food recipe</td></tr><tr><td>Stereo microscope</td><td>Leica</td><td>Leica S6E</td><td>For tissue dissection</td></tr><tr><td>Wipe paper</td><td>N/A</td><td>N/A</td><td></td></tr><tr><td>Yeast</td><td></td><td></td><td>For fly food</td></tr><tr><td>yw</td><td>Kept as lab stock</td><td>N/A</td><td>Drosophila</td></tr></tbody></table>

dissection and lipid droplet staining of oenocytes in drosophila larvae

Lipids are essential for animal development and physiological homeostasis. Dysregulation of lipid metabolism results in various developmental defects and diseases, such as obesity and fatty liver. Usually, lipids are stored in lipid droplets, which are the multifunctional lipid storage organelles in cells. Lipid droplets vary in size and number in different tissues and under different conditions. It has been reported that lipid droplets are tightly controlled through regulation of its biogenesis and degradation. In Drosophila melanogaster, the oenocyte is an important tissue for lipid metabolism and has been recently identified as a human liver analogue regarding lipid mobilization in response to stress. However, the mechanisms underlying the regulation of lipid droplet metabolism in oenocytes remain elusive. To solve this problem, it is of utmost importance to develop a reliable and sensitive method to directly visualize lipid droplet dynamic changes in oenocytes during development and under stressful conditions. Taking advantage of the lipophilic BODIPY 493/503, a lipid droplet-specific fluorescent dye, described here is a detailed protocol for the dissection and subsequent lipid droplet staining in the oenocytes of Drosophila larvae in response to starvation. This allows for qualitative analysis of lipid droplet dynamics under various conditions by confocal microscopy. Furthermore, this rapid and highly reproducible method can also be used in genetic screens for indentifying novel genetic factors involving lipid droplet metabolism in oenocytes and other tissues.

Successful execution of this procedure should result in clear lipid droplets staining that reveals the number and size of lipid droplets in the oenocytes. Figure 1A,A&#39;,A&#39;&#39; shows that there are few detectable lipid droplets (green dots) in the oenocytes of normal feeding larvae during different developmental stages. Figure 1B,B&#39;,B&#39;&#39; shows increased lipid droplets (green dot...

Watch this Scientific Journal Video about Dissection and Lipid Droplet Staining of Oenocytes in Drosophila Larvae at JoVE.com

Dissection and Lipid Droplet Staining of Oenocytes in Drosophila Larvae

Presented here are detailed methods for the dissection and lipid droplet staining of oenocytes in Drosophila larvae using BODIPY 493/503, a lipid droplet-specific fluorescent dye.

Dissection and Lipid Droplet Staining of Oenocytes in Drosophila Larvae

Lipids are essential for animal development and physiological homeostasis. Dysregulation of lipid metabolism results in various developmental defects and ...

dissection-lipid-droplet-staining-oenocytes-drosophila

Research

JoVE Journal

Biology

11.5K Views.  Guangzhou Medical University. Presented here are detailed methods for the dissection and lipid droplet staining of oenocytes in Drosophila larvae using BODIPY 493/503, a lipid droplet-specific fluorescent dye.

Video: Dissection and Lipid Droplet Staining of Oenocytes in Drosophila Larvae

Single Drosophila Ommatidium Dissection and Imaging

The fruit fly Drosophila melanogaster has made invaluable contributions to neuroscience research and has been used widely as a model for neurodegenerative diseases because of its powerful genetics1. The fly eye in particular has been the organ of choice for neurodegeneration research, being the most accessible and life-dispensable part of the Drosophila nervous system. However the major caveat of intact eyes is the difficulty, because of the intense autofluorescence of the pigment, in imaging intracellular events, such as autophagy dynamics2, which are paramount to understanding of neurodegeneration.
We have recently used the dissection and culture of single ommatidia3 that has been essential for our understanding of autophagic dysfunctions in a fly model of Dentatorubro-Pallidoluysian Atrophy (DRPLA)3, 4.
We now report a comprehensive description of this technique (Fig. 1), adapted from electrophysiological studies5, which is likely to expand dramatically the possibility of fly models for neurodegeneration. This method can be adapted to image live subcellular events and to monitor effective drug administration onto photoreceptor cells (Fig. 2). If used in combination with mosaic techniques6-8, the responses of genetically different cells can be assayed in parallel (Fig. 2).

Single Drosophila Ommatidium Dissection and Imaging

The limiting factor in the use of the adult Drosophila eye to study neurodegeneration and cell biology is the difficult imaging of intracellular processes. We describe the dissection of single ommatidia to generate a bona-fide primary neuronal cell culture, which can be subject to drug treatment and advanced imaging.

The fruit fly Drosophila melanogaster has made invaluable contributions to neuroscience research and has been used widely as a model for neurodegenerative ...

Neuroscience

Methods to Examine the Lymph Gland and Hemocytes in Drosophila Larvae

Many parallels exist between the Drosophila and mammalian hematopoietic systems, even though Drosophila lack the lymphoid lineage that characterize mammalian adaptive immunity. Drosophila and mammalian hematopoiesis occur in spatially and temporally distinct phases to produce several blood cell lineages. Both systems maintain reservoirs of blood cell progenitors with which to expand or replace mature lineages. The hematopoietic system allows Drosophila and mammals to respond to and to adapt to immune challenges. Importantly, the transcriptional regulators and signaling pathways that control the generation, maintenance, and function of the hematopoietic system are conserved from flies to mammals. These similarities allow Drosophila to be used to genetically model hematopoietic development and disease.
Here we detail assays to examine the hematopoietic system of Drosophila larvae. In particular, we outline methods to measure blood cell numbers and concentration, visualize a specific mature lineage in vivo, and perform immunohistochemistry on blood cells in circulation and in the hematopoietic organ. These assays can reveal changes in gene expression and cellular processes including signaling, survival, proliferation, and differentiation and can be used to investigate a variety of questions concerning hematopoiesis. Combined with the genetic tools available in Drosophila, these assays can be used to evaluate the hematopoietic system upon defined genetic alterations. While not specifically outlined here, these assays can also be used to examine the effect of environmental alterations, such as infection or diet, on the hematopoietic system.

Methods to Examine the Lymph Gland and Hemocytes in Drosophila Larvae

Drosophila and mammalian hematopoietic systems share many common features, making Drosophila an attractive genetic model to study hematopoiesis. Here we demonstrate dissection and mounting of the major larval hematopoietic organ for immunohistochemistry. We also describe methods to assay various larval hematopoietic compartments including circulating hemocytes and sessile crystal cells.

Many parallels exist between the Drosophila and mammalian hematopoietic systems, even though Drosophila lack the lymphoid lineage that characterize ...

Developmental Biology

Dissection and Immunofluorescent Staining of Mushroom Body and Photoreceptor Neurons in Adult Drosophila melanogaster Brains

Nervous system development involves a sequential series of events that are coordinated by several signaling pathways and regulatory networks. Many of the proteins involved in these pathways are evolutionarily conserved between mammals and other eukaryotes, such as the fruit fly Drosophila melanogaster, suggesting that similar organizing principles exist during the development of these organisms. Importantly, Drosophila has been used extensively to identify cellular and molecular mechanisms regulating processes that are required in mammals including neurogenesis, differentiation, axonal guidance, and synaptogenesis. Flies have also been used successfully to model a variety of human neurodevelopmental diseases. Here we describe a protocol for the step-by-step microdissection, fixation, and immunofluorescent localization of proteins within the adult Drosophila brain. This protocol focuses on two example neuronal populations, mushroom body neurons and retinal photoreceptors, and includes optional steps to trace individual mushroom body neurons using Mosaic Analysis with a Repressible Cell Marker (MARCM) technique. Example data from both wild-type and mutant brains are shown along with a brief description of a scoring criteria for axonal guidance defects. While this protocol highlights two well-established antibodies for investigating the morphology of mushroom body and photoreceptor neurons, other Drosophila brain regions and the localization of proteins within other brain regions can also be investigated using this protocol.

Dissection and Immunofluorescent Staining of Mushroom Body and Photoreceptor Neurons in Adult Drosophila melanogaster Brains

This protocol describes the dissection and immunostaining of adult Drosophila melanogaster brain tissues. Specifically, this protocol highlights the use of Drosophila mushroom body and photoreceptor neurons as example neuronal subsets that can be accurately used to uncover general principles underlying many aspects of neuronal development.

Nervous system development involves a sequential series of events that are coordinated by several signaling pathways and regulatory networks. Many of the ...

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 ...

Visualizing Cytoskeleton-Dependent Trafficking of Lipid-Containing Organelles in Drosophila Embryos

Early Drosophila embryos are large cells containing a vast array of conventional and embryo-specific organelles. During the first three hours of embryogenesis, these organelles undergo dramatic movements powered by actin-based cytoplasmic streaming and motor-driven trafficking along microtubules. The development of a multitude of small, organelle-specific fluorescent probes (FPs) makes it possible to visualize a wide range of different lipid-containing structures in any genotype, allowing live imaging without requiring a genetically encoded fluorophore. This protocol shows how to inject vital dyes and molecular probes into Drosophila embryos to monitor the trafficking of specific organelles by live imaging. This approach is demonstrated by labeling lipid droplets (LDs) and following their bulk movement by particle image velocimetry (PIV). This protocol provides a strategy amenable to the study of other organelles, including lysosomes, mitochondria, yolk vesicles, and the ER, and for tracking the motion of individual LDs along microtubules. Using commercially available dyes brings the benefits of separation into the violet/blue and far-red regions of the spectrum. By multiplex co-labeling of organelles and/or cytoskeletal elements via microinjection, all the genetic resources in Drosophila are available for trafficking studies without the need to introduce fluorescently tagged proteins. Unlike genetically encoded fluorophores, which have low quantum yields and bleach easily, many of the available dyes allow for rapid and simultaneous capture of several channels with high photon yields.

Visualizing Cytoskeleton-Dependent Trafficking of Lipid-Containing Organelles in Drosophila Embryos

In the early Drosophila embryo, many organelles are motile. In principle, they can be imaged live via specific fluorescent probes, but the eggshell prevents direct application to the embryo. This protocol describes how to introduce such probes via microinjection, and then analyze bulk organelle motion via particle image velocimetry.

Early Drosophila embryos are large cells containing a vast array of conventional and embryo-specific organelles. During the first three hours of ...

Dissection of Oenocytes from Adult Drosophila melanogaster

In Drosophila melanogaster, as in other insects, a waxy layer on the outer surface of the cuticle, composed primarily of hydrocarbon compounds, provides protection against desiccation and other environmental challenges. Several of these cuticular hydrocarbon (CHC) compounds also function as semiochemical signals, and as such mediate pheromonal communications between members of the same species, or in some instances between different species, and influence behavior. Specialized cells referred to as oenocytes are regarded as the primary site for CHC synthesis. However, relatively little is known regarding the involvement of the oenocytes in the regulation of the biosynthetic, transport, and deposition pathways contributing to CHC output. Given the significant role that CHCs play in several aspects of insect biology, including chemical communication, desiccation resistance, and immunity, it is important to gain a greater understanding of the molecular and genetic regulation of CHC production within these specialized cells. The adult oenocytes of D. melanogaster are located within the abdominal integument, and are metamerically arrayed in ribbon-like clusters radiating along the inner cuticular surface of each abdominal segment. In this video article we demonstrate a dissection technique used for the preparation of oenocytes from adult D. melanogaster. Specifically, we provide a detailed step-by-step demonstration of (1) how to fillet prepare an adult Drosophila abdomen, (2) how to identify the oenocytes and discern them from other tissues, and (3) how to remove intact oenocyte clusters from the abdominal integument. A brief experimental illustration of how this preparation can be used to examine the expression of genes involved in hydrocarbon synthesis is included. The dissected preparation demonstrated herein will allow for the detailed molecular and genetic analysis of oenocyte function in the adult fruit fly.

Dissection of Oenocytes from Adult Drosophila melanogaster

In insects, the oenocytes produce cuticular hydrocarbon compounds. These compounds protect against desiccation and facilitate chemical communication. Here we demonstrate a dissection technique used to isolate the oenocytes from adult Drosophila melanogaster, and illustrate how this preparation can be utilized to study genes involved in hydrocarbon synthesis.

In Drosophila melanogaster, as in other insects, a waxy layer on the outer surface of the cuticle, composed primarily of hydrocarbon compounds, provides ...

Paraffin-Embedded and Frozen Sections of Drosophila Adult Muscles

The molecular characterization of muscular dystrophies and myopathies in humans has revealed the complexity of muscle disease and genetic analysis of muscle specification, formation and function in model systems has provided valuable insight into muscle physiology. Therefore, identifying and characterizing molecular mechanisms that underlie muscle damage is critical. The structure of adult Drosophila multi-fiber muscles resemble vertebrate striated muscles 1 and the genetic tractability of Drosophila has made it a great system to analyze dystrophic muscle morphology and characterize the processes affecting muscular function in ageing adult flies 2. Here we present the histological technique for preparing paraffin-embedded and frozen sections of Drosophila thoracic muscles. These preparations allow for the tissue to be stained with classical histological stains and labeled with protein detecting dyes, and specifically cryosections are ideal for immunohistochemical detection of proteins in intact muscles. This allows for analysis of muscle tissue structure, identification of morphological defects, and detection of the expression pattern for muscle/neuron-specific proteins in Drosophila adult muscles. These techniques can also be slightly modified for sectioning of other body parts.

Paraffin-Embedded and Frozen Sections of Drosophila Adult Muscles

Identification of mechanisms underlying muscle damage is crucial. Here we present the histological technique for preparing paraffin-embedded and frozen sections of Drosophila thoracic muscles. This allows analysis of muscle morphology and localization of protein and other muscle cell components.

The molecular characterization of muscular dystrophies and myopathies in humans has revealed the complexity of muscle disease and genetic analysis of ...

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 ...

Preparation of Adult Drosophila Eyes for Thin Sectioning and Microscopic Analysis

Drosophila has long been used as model system to study development, mainly due to the ease with which it is genetically tractable. Over the years, a plethora of mutant strains and technical tricks have been developed to allow sophisticated questions to be asked and answered in a reasonable amount of time. Fundamental insight into the interplay of components of all known major signaling pathways has been obtained in forward and reverse genetic Drosophila studies. The fly eye has proven to be exceptionally well suited for mutational analysis, since, under laboratory conditions, flies can survive without functional eyes. Furthermore, the surface of the insect eye is composed of some 800 individual unit eyes (facets or ommatidia) that form a regular, smooth surface when looked at under a dissecting microscope. Thus, it is easy to see whether a mutation might affect eye development or growth by externally looking for the loss of the smooth surface ('rough eye' phenotype; Fig. 1) or overall eye size, respectively (for examples of screens based on external eye morphology see e.g.1). Subsequent detailed analyses of eye phenotypes require fixation, plastic embedding and thin-sectioning of adult eyes.
The Drosophila eye develops from the so-called eye imaginal disc, a bag of epithelial cells that proliferate and differentiate during larval and pupal stages (for review see e.g. 2). Each ommatidium consists of 20 cells, including eight photoreceptors (PR or R-cells; Fig. 2), four lens-secreting cone cells, pigment cells ('hexagon' around R-cell cluster) and a bristle. The photoreceptors of each ommatidium, most easily identified by their light sensitive organelles, the rhabdomeres, are organized in a trapezoid made up of the six "outer" (R1-6) and two "inner" photoreceptors (R7/8; R8 [Fig. 2] is underneath R7 and thus only seen in sections from deeper areas of the eye). The trapezoid of each facet is precisely aligned with those of its neighbors and the overall anteroposterior and dorsoventral axes of the eye (Fig. 3A). In particular, the ommatidia of the dorsal and ventral (black and red arrows, respectively) halves of the eye are mirror images of each other and correspond to two chiral forms established during planar cell polarity signaling (for review see e.g. 3).
The method to generate semi-thin eye sections (such as those presented in Fig. 3) described here is slightly modified from the one originally described by Tomlinson and Ready4. It allows the morphological analysis of all cells except for the transparent cone cells. In addition, the pigment of R-cells (blue arrowheads in Fig. 2 and 3) can be used as a cell-autonomous marker for the genotype of a R-cell, thus genetic requirements of genes in a subset of R-cells can readily be determined5,6.

Preparation of Adult Drosophila Eyes for Thin Sectioning and Microscopic Analysis

A standard approach to prepare adult Drosophila eyes for semi-thin sectioning and light microscopic analysis is presented here. The protocol can be used for gross morphological analysis of eye defects, or with the indicated adjustments can be used to determine genetic requirements of genes in specific cell types of the eye (e.g. clonal analysis of photoreceptors) or for electron microscopic analysis.

Drosophila has long been used as model system to study development, mainly due to the ease with which it is genetically tractable. Over the years, a ...

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.

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.

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

Dissection and Lipid Droplet Staining of Oenocytes in Drosophila Larvae

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Single Drosophila Ommatidium Dissection and Imaging

Methods to Examine the Lymph Gland and Hemocytes in Drosophila Larvae

Dissection and Immunofluorescent Staining of Mushroom Body and Photoreceptor Neurons in Adult Drosophila melanogaster Brains

Dissection and Staining of Drosophila Pupal Ovaries

Visualizing Cytoskeleton-Dependent Trafficking of Lipid-Containing Organelles in Drosophila Embryos

Dissection of Oenocytes from Adult Drosophila melanogaster

Paraffin-Embedded and Frozen Sections of Drosophila Adult Muscles

Dissection and Staining of Drosophila Larval Ovaries

Preparation of Adult Drosophila Eyes for Thin Sectioning and Microscopic Analysis

Studying Mitochondrial Structure and Function in Drosophila Ovaries