Name: dissection and lipid droplet staining of oenocytes in drosophila larvae
Uploaded: 2019-12-28T13:00:00
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 y...

<ol>
	<li>Liu, Z., Huang, X. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Lipid+metabolism+in+Drosophila:+development+and+disease.">Lipid metabolism in Drosophila: development and disease.</a> Acta Biochimica et Biophysica Sinica (Shanghai). 45 (1), 44-50 (2013).</li><li>Cheng, Y., Chen, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Fruit+fly+research+in+China.">Fruit fly research in China.</a> Journal of Genetics and Genomics. 45 (11), 583-592 (2018).</li><li>Warr, C. G., Shaw, K. H., Azim, A., Piper, M. D. W., Parsons, L. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Using+mouse+and+Drosophila+models+to+investigate+the+mechanistic+links+between+diet,+obesity,+type+II+diabetes,+and+cancer.">Using mouse and Drosophila models to investigate the mechanistic links between diet, obesity, type II diabetes, and cancer.</a> International Journal of Molecular Science. 19 (12), (2018).</li><li>Gutierrez, E., Wiggins, D., Fielding, B., Gould, A. P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Specialized+hepatocyte-like+cells+regulate+Drosophila+lipid+metabolism.">Specialized hepatocyte-like cells regulate Drosophila lipid metabolism.</a> Nature. 445 (7125), 275-280 (2007).</li><li>Makki, R., Cinnamon, E., Gould, A. P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+development+and+functions+of+oenocytes.">The development and functions of oenocytes.</a> Annual Review of Entomology. 59, 405-425 (2014).</li><li>Chatterjee, D., 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=Control+of+metabolic+adaptation+to+fasting+by+dILP6-induced+insulin+signaling+in+Drosophila+oenocytes.">Control of metabolic adaptation to fasting by dILP6-induced insulin signaling in Drosophila oenocytes.</a> Proceedings of the National Academy of Sciences of the United States of America. 111 (50), 17959-17964 (2014).</li><li>Yan, Y., 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=HDAC6+suppresses+age-dependent+ectopic+fat+accumulation+by+maintaining+the+proteostasis+of+PLIN2+in+Drosophila.">HDAC6 suppresses age-dependent ectopic fat accumulation by maintaining the proteostasis of PLIN2 in Drosophila.</a> Developmental Cell. 43 (1), 99-111 (2017).</li><li>Yan, Y., 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=HDAC6+regulates+lipid+droplet+turnover+in+response+to+nutrient+deprivation+via+p62-mediated+selective+autophagy.">HDAC6 regulates lipid droplet turnover in response to nutrient deprivation via p62-mediated selective autophagy.</a> Journal of Genetics and Genomics. 46 (4), 221-229 (2019).</li><li>Farese, R. V., Walther, T. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Lipid+droplets+finally+get+a+little+R-E-S-P-E-C-T.">Lipid droplets finally get a little R-E-S-P-E-C-T.</a> Cell. 139 (5), 855-860 (2009).</li><li>Murphy, D. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+dynamic+roles+of+intracellular+lipid+droplets:+from+archaea+to+mammals.">The dynamic roles of intracellular lipid droplets: from archaea to mammals.</a> Protoplasma. 249 (3), 541-585 (2012).</li><li>Tennessen, J. M., Barry, W. E., Cox, J., Thummel, C. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Methods+for+studying+metabolism+in+Drosophila.">Methods for studying metabolism in Drosophila.</a> Methods. 68 (1), 105-115 (2014).</li><li>Fowler, S. D., Greenspan, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Application+of+Nile+red,+a+fluorescent+hydrophobic+probe,+for+the+detection+of+neutral+lipid+deposits+in+tissue+sections:+comparison+with+oil+red+O.">Application of Nile red, a fluorescent hydrophobic probe, for the detection of neutral lipid deposits in tissue sections: comparison with oil red O.</a> Journal of Histochemistry &amp; Cytochemistry. 33 (8), 833-836 (1985).</li><li>Gocze, P. M., Freeman, D. 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>

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 border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<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 &#215; 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 1xPBS</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>1xPBS (137 mM NaCl, 2.7 mM KCl, 10 mM Na2HPO4,1.8 mM KH2PO4, 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 ...

This method of an oenocyte dissection and lipid droplet staining can be used to realize the appearance of lipid droplet in drosophila larvae oenocyte under normal and stressed conditions. This method provide a useful tool for studying lipid metabolism not only in insects but also in mammals with only a few minor modifications. To induce egg laying, place 150 virgin female flies and 75 males of the desired genotypes in an egg-laying bottle and place the bottle under a light-proof box in a 25 degrees Celsius incubator with 60%humidity.After one hour, transfer the flies to a new bottle and let the flies lay eggs in the new bottle for one hour before removing the adults. Then, allow the eggs to develop into the third instar larvae for 84 hours at 25 degrees Celcius with a 12-hour light-dark cycle. Before initiating the starvation treatment, place an appropriately sized piece of filter paper into a six centimeter Petri dish and add one milliliter of PBS onto the paper to create a starvation chamber.To make a control treatment chamber, place five milliliters of Bloomington standard cornmeal food into a six centimeter Petri dish. When the chambers are ready, use a small paint brush to collect instar larvae of the same approximate size from the egg-laying bottle. And 20 larvae randomly into each starvation and the control chamber.Then place the chambers in the incubator for 12, 24 or 36 hours of development and/or starvation. At the end of the treatment period, use forceps to gently transfer larvae from each chamber into a dissection plate containing ice cold PBS and place the plate under a stereo microscope. Orient the first larva ventral side up and use forceps to gently hold the larva in place.Place one pin through the pharynx and another pin through the spiracle to secure the larva to the plate. Use Vannas spring scissors to make a longitudinal incision through the epidermis from the anterior to the posterior end of the animal. Use forceps to remove the internal tissue of the epidermis, taking care to avoid damage to the oenocytes on the internal surface of the epidermis.Then use forceps to retrieve the dissection pins and transfer the epidermis to a 1.5 milliliter microcentrifuge tube of PBS on ice. For lipid droplet staining, incubate the dissected epidermis in fixation buffer for 30 minutes at room temperature on a rotator, followed by a quick resuspension in one milliliter of PBS. Then wash the samples with three five-minute washes in one milliliter of PBS per wash to remove any residual fixative.After the last wash, incubate the epidermal samples with an appropriate lipophilic probe for 30 minutes at room temperature on a rotator. During the incubation, wrap the sample tubes in foil to protect the tissues from light and wash the samples three times in one milliliter of PBS for 10 minutes per wash. To image the oenocytes, transfer each epidermis sample into six microliters of mounting medium on a clean microscope slide, adjust the orientation of the tissue, so that the internal surface containing the oenocytes is touching the bottom of the slide.Gently place a coverslip onto the epidermis and seal the edges with clear nail polish. When the polish has dried, image the samples on a confocal microscope, put a 63 X magnification with the appropriate excitation and emission wavelengths. There are a few detectable lipid droplets within the oenocytes of normal feeding larvae at different developmental stages.As observed in these representative images however, the lipid droplets increase in number within the oenocytes in response to 12, 24, and 36-hour periods of starvation. This method provides and easy and a feasible protocol for researchers in relevant research fields to investigate whether genetic or environmental manipulations cause lipid droplet changes in cells.

dissection-lipid-droplet-staining-oenocytes-drosophila

Research

JoVE Journal

Biology

Dissection of Drosophila Ovaries

Dissection of Larval CNS in Drosophila Melanogaster

The central nervous system (CNS) of Drosophila larvae is complex and poorly understood.  One way to investigate the CNS is to use immunohistochemistry to examine the expression of various novel and marker proteins.  Staining of whole larvae is impractical because the tough cuticle prevents antibodies from penetrating inside the body cavity.  In order to stain these tissues it is necessary to dissect the animal prior to fixing and staining.  In this article we demonstrate how to dissect Drosophila larvae without damaging the CNS.  Begin by tearing the larva in half with a pair of fine forceps, and then turn the cuticle "inside-out" to expose the CNS.  If the dissection is performed carefully the CNS will remain attached to the cuticle.  We usually keep the CNS attached to the cuticle throughout the fixation and staining steps, and only completely remove the CNS from the cuticle just prior to mounting the samples on glass slides.  We also show some representative images of a larval CNS stained with Eve, a transcription factor expressed in a subset of neurons in the CNS.  The article concludes with a discussion of some of the practical uses of this technique and the potential difficulties that may arise.

In this article we demonstrate how to dissect the central nervous system from third instar Drosophila larvae.

The central nervous system (CNS) of Drosophila larvae is complex and poorly understood.  One way to investigate the CNS is to use immunohistochemistry to ...

Dissection of Imaginal Discs from 3rd Instar Drosophila Larvae

This protocol demonstrates the dissection technique used for isolating imaginal discs from drosophila larvae at the 3rd instar stage. Methods for fixing the tissue after saying and removing the wing, leg, and eye discs are demonstrated directly on a microscope slide for subsequent visualization.

High-resolution Measurement of Odor-Driven Behavior in Drosophila Larvae

Olfactory responses in Drosophila larvae have been traditionally studied in Petri dishes comprising a single peripheral odor source. In this behavioral paradigm, the experimenter usually assumes that the rapid diffusion of odorant molecules from the source leads to the creation of a stable gradient in the dish. To establish a quantitative correlation between sensory inputs and behavioral responses, it is necessary to achieve a more thorough characterization of the odorant stimulus conditions. In this video article, we describe a new method allowing the construction of odorant gradients with stable and controllable geometries. We briefly illustrate how these gradients can be used to screen for olfactory defects (full and partial anosmia) and to study more subtle features of chemotaxis behavior.

In this video article, we describe a new method allowing the construction of odorant gradients with stable and controllable geometries. We briefly illustrate how these gradients can be used to screen for olfactory defects (full and partial anosmia) and to study more subtle features of chemotaxis behavior.

Olfactory responses in Drosophila larvae have been traditionally studied in Petri dishes comprising a single peripheral odor source. In this behavioral ...

Drosophila Larval NMJ Dissection

The Drosophila neuromuscular junction (NMJ) is an established model system used for the study of synaptic development and plasticity. The widespread use of the Drosophila motor system is due to its high accessibility. It can be analyzed with single-cell resolution. There are 30 muscles per hemisegment whose arrangement within the peripheral body wall are known. A total of 35 motor neurons attach to these muscles in a pattern that has high fidelity. Using molecular biology and genetics, one can create transgenic animals or mutants. Then, one can study the developmental consequences on the morphology and function of the NMJ. In order to access the NMJ for study, it is necessary to carefully dissect each larva. In this article we demonstrate how to properly dissect Drosophila larvae for study of the NMJ by removing all internal organs while leaving the body wall intact. This technique is suitable to prepare larvae for imaging, immunohistochemistry, or electrophysiology.

This protocol demonstrates how to dissect Drosophila larvae in preparation for immunohistochemistry and/or imaging of the neuromuscular junction.

The Drosophila neuromuscular junction (NMJ) is an established model system used for the study of synaptic development and plasticity. The widespread use ...

Live Dissection of Drosophila Embryos: Streamlined Methods for Screening Mutant Collections by Antibody Staining

Drosophila embryos between stages 14 and 17 of embryonic development can be readily dissected to generate "fillet" preparations. In these preparations, the central nervous system runs down the middle, and is flanked by the body walls. Many different phenotypes have been examined using such preparations. In most cases, the fillets were generated by dissection of antibody-stained fixed whole-mount embryos. These "fixed dissections" have some disadvantages, however. They are time-consuming to execute, and it is difficult to sort mutant (GFP-negative) embryos from stocks in which mutations are maintained over GFP balancer chromosomes. Since 2002, our group has been conducting deficiency and ectopic expression screens to identify ligands for orphan receptors. In order to do this, we developed streamlined protocols for live embryo dissection and antibody staining of collections containing hundreds of balanced lines. We have concluded that it is considerably more efficient to examine phenotypes in large collections of stocks by live dissection than by fixed dissection. Using the protocol described here, a single trained individual can screen up to 10 lines per day for phenotypes, examining 4-7 mutant embryos from each line under a compound microscope. This allows the identification of mutations conferring subtle, low-penetrance phenotypes, since up to 70 hemisegments per line are scored at high magnification with a 40X water-immersion lens.

Live Dissection of Drosophila Embryos: Streamlined Methods for Screening Mutant Collections by Antibody Staining

We describe a streamlined protocol for generating "fillet" preparations of Drosophila embryos of specific genotypes. This protocol allows efficient execution of a variety of genetic screens. It also allows excellent visualization of structures in the late embryo.

Drosophila embryos between stages 14 and 17 of embryonic development can be readily dissected to generate "fillet" preparations.  In these preparations, ...

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 ...

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 ...

Quantification of Lipid Abundance and Evaluation of Lipid Distribution in Caenorhabditis elegans by Nile Red and Oil Red O Staining

Caenorhabditis elegans is an exceptional model organism in which to study lipid metabolism and energy homeostasis. Many of its lipid genes are conserved in humans and are associated with metabolic syndrome or other diseases. Examination of lipid accumulation in this organism can be carried out by fixative dyes or label-free methods. Fixative stains like Nile red and oil red O are inexpensive, reliable ways to quantitatively measure lipid levels and to qualitatively observe lipid distribution across tissues, respectively. Moreover, these stains allow for high-throughput screening of various lipid metabolism genes and pathways. Additionally, their hydrophobic nature facilitates lipid solubility, reduces interaction with surrounding tissues, and prevents dissociation into the solvent. Though these methods are effective at examining general lipid content, they do not provide detailed information about the chemical composition and diversity of lipid deposits. For these purposes, label-free methods such as GC-MS and CARS microscopy are better suited, their costs notwithstanding.

Quantification of Lipid Abundance and Evaluation of Lipid Distribution in Caenorhabditis elegans by Nile Red and Oil Red O Staining

Nile red staining of fixed Caenorhabditis elegans is a method for quantitative measurement of neutral lipid deposits, while oil red O staining facilitates qualitative assessment of lipid distribution among tissues.

Caenorhabditis elegans is an exceptional model organism in which to study lipid metabolism and energy homeostasis. Many of its lipid genes are conserved ...

Plastoglobule Lipid Droplet Isolation from Plant Leaf Tissue and Cyanobacteria

Plastoglobule lipid droplets are a dynamic sub-compartment of plant chloroplasts and cyanobacteria. Found ubiquitously among photosynthetic species, they are believed to serve a central role in the adaptation and remodeling of the thylakoid membrane under rapidly changing environmental conditions. The capacity to isolate plastoglobules of high purity has greatly facilitated their study through proteomic, lipidomic, and other methodologies. With plastoglobules of high purity and yield, it is possible to investigate their lipid and protein composition, enzymatic activity, and protein topology, among other possible molecular characteristics. This article presents a rapid and effective protocol for the isolation of plastoglobules from chloroplasts of plant leaf tissue and presents methodological variations for the isolation of plastoglobules and related lipid droplet structures from maize leaves, the desiccated leaf tissue of the resurrection plant, Eragrostis nindensis, and the cyanobacterium, Synechocystis sp. PCC 6803. Isolation relies on the low density of these lipid-rich particles, which facilitates their purification by sucrose density flotation. This methodology will prove valuable in the study of plastoglobules from diverse species.

A fast and efficient protocol is presented for the isolation of plastoglobule lipid droplets associated with various photosynthetic organisms. The successful preparation of isolated plastoglobules is a crucial first step that precedes detailed molecular investigations such as proteomic and lipidomic analyses.

Plastoglobule lipid droplets are a dynamic sub-compartment of plant chloroplasts and cyanobacteria. Found ubiquitously among photosynthetic species, they ...