We are grateful to THFC and BDSC for providing the fly strains. Dr. Tong Chao is supported by the National Natural Science Foundation of China (32030027, 91754103, 92157201) and Fundamental research funds for the central universities. We thank the core facility in the Life Sciences Institute (LSI) for providing services.

1. Drosophila crossing and egg laying
<ol>
	<li>Introduce 3 male (genotype hsFLP ubiRFP FRT19A; cgGal4 UAS-GFP-Atg8a) and 15 female (genotype y&#39; w* Mu FRT19A/ FM7, Kr GFP) adult flies (see Table of Materials) into a culture vial (with standard cornmeal/molasses/agar Drosophila media at 25 &#176;C) for mating. 
	NOTE: Multiple culture vials of the same cross must be set up to ensure enough larvae for further experiments. The male fly strain with genotype hsFLP ubiRFP FRT19A; cgGal4 UAS-GFP-Atg8a carries an FRT site at the X chromosome near the centromere (FRT19A). It also expresses FLP upon heat shock at 37 &#176;C. A transgene with ubiquitous RFP expression is inserted on the X chromosome. GFP-Atg8a is expressed under the control of cgGal4 in the larval fat body22. The female fly strain with genotype y&#39; w* Mu FRT19A/ FM7, Kr GFP carries a lethal mutation (Mu) and FRT19A on the X chromosome. The detailed crossing scheme is shown in Figure 1.</li>
	<li>For egg laying, transfer the flies to a new culture vial. At 48 h post introduction, tap the &#34;mating&#34; vial until the flies are stunned and drop down on the media at the base of the vial.
	<ol>
		<li>Unplug the &#34;mating&#34; vial and cover its mouth with an unplugged inverted vial with fresh media (fresh culture vial). Then, transfer the flies to the fresh culture vial by flipping and tapping the vials.</li>
		<li>Plug the fresh culture vial (with the transferred flies) and discard the old culture vial. Place this fresh culture vial in a 25 &#176;C incubator for egg laying on the fresh media. 
		NOTE: Prewarming the fresh culture vials to 25 &#176;C for 15 min before the fly transfer process will help the flies adapt to the new environment quickly and hasten egg laying.</li>
	</ol>
	</li>
	<li>Remove the flies after 6 h of egg laying. If the experiments need to be repeated, transfer these flies into another culture vial (as described in step 1.2). Otherwise, discard the flies by dumping them into a flask containing 75% ethanol).
	<ol>
		<li>Incubate the vial with embryos in a 37 &#176;C water bath for 1 h to induce FLP expression. Subsequently, place them in a 25 &#176;C incubator and allow the embryos to continue to develop. 
		&#8203;NOTE: The successful formation of mutant clones can be confirmed subsequently through imaging. The absence of RFP signals marks the mutant clones.</li>
	</ol>
	</li>
</ol>
2. Amino acid starvation induces autophagy
<ol>
	<li>Using a laboratory spatula, scoop out the media containing the developing larvae into a Petri dish 75 h post egg-laying. Add 3 mL of 1x PBS to the dish and gently separate out the culture media and the larvae using long forceps. Select 10 to 15 early third instar larvae. 
	NOTE: Early third instar larvae need to be chosen carefully. The larvae&#39;s developmental stage is critical for this protocol&#39;s success. Due to the restricted egg laying duration, most larvae in the culture vial are expected to be in the early third instar stage by 75 h of incubation. However, different culture media recipes may lead to variations in the developmental timing of the larvae. The criteria for distinguishing early third instar larvae are the body length, the presence of anterior and posterior spiracles, as well as the mandibular hooks of the mouth apparatus23.</li>
	<li>Fill the wells of a 9-well glass depression spot plate (see Table of Materials) with 1x PBS. Place the separated third instar larvae in the wells using long forceps and wash the larvae thoroughly to remove all the media residues.</li>
	<li>Take 5 mL of 20% sucrose (in 1x PBS) solution in an empty vial and place the clean third instar larvae in this solution using long forceps. Incubate this vial in a 25 &#176;C incubator for 6 h before harvesting them for dissection. 
	&#8203;NOTE: The 20% sucrose (in 1x PBS) solution serves as the amino acid-deficient starvation medium.</li>
</ol>
3. Third instar larvae dissection and sample tissue processing
<ol>
	<li>Sharpen two pairs of #5 forceps (see Table of Materials) evenly on both sides with a sharpening stone.</li>
	<li>Add 400 &#181;L of 1x PBS into each well of the 9-well glass depression spot plate and transfer the larvae into the wells with long forceps. Place one larva in a well with the dorsal side (the side with the trachea) facing up.
	<ol>
		<li>Grip the cuticle of the larva with two #5 forceps in the middle of the larval trunk and gently tear open the cuticles. The exposed fat bodies, along with other larval internal tissues, will still be attached to the carcass of the larva. Pull enough to expose the internal organs as much as possible. Repeat this step for all the larvae. 
		NOTE: Each larva has two large pieces of fat body along the body trunk. Fat body tissue is a white, opaque, and flat monolayer that can be easily distinguished under the dissection microscope24.</li>
	</ol>
	</li>
	<li>Transfer the larval carcass into a 1.5 mL microcentrifuge tube containing 500 &#181;L of 4% paraformaldehyde (PFA). Incubate for 30 min at 25 &#176;C without shaking the tubes. 
	CAUTION: 4% PFA is toxic. Wear gloves and masks for protection. 
	NOTE: Preparing 4% PFA in 1x PBS buffer is recommended. PFA powder does not dissolve in 1x PBS instantly. Hence, the mixture must be incubated at 65 &#176;C in an incubator overnight or shaken intermittently for 45 min at 25 &#176;C.</li>
	<li>After 30 min incubation of the carcass with 4% PFA, pipette out the PFA solution, add 500 &#181;L of 1x PBS into the tube, and gently shake the tube on a flat rotator for 10 min before discarding the 1x PBS solution (repeat 3x).</li>
	<li>Using long forceps, transfer the fixed and washed larval carcass to a well in the 9-depression spot plate filled with 1x PBS. Using #5 forceps, remove all the non-fat body tissues.</li>
	<li>Use #5 forceps to mount the pieces of the fat body on a microscope slide using 80% glycerol as the mounting medium and lay a coverslip on top. 
	NOTE: Before mounting, check the pH of the 80% glycerol (using pH papers, pH = 7 is optimal) to protect the GFP signal. Mounting media (see Table of Materials) with DAPI in 80% glycerol will help in imaging the nuclei of fat body cells. These slides are stable for 1 week when stored in the dark at 4 &#176;C.</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=Mosaic+analysis+in+Drosophila.">Mosaic analysis in Drosophila.</a> Genetics. 208 (2), 473-490 (2018).</li><li>Zappia, M. P., 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=E2F/Dp+inactivation+in+fat+body+cells+triggers+systemic+metabolic+changes.">E2F/Dp inactivation in fat body cells triggers systemic metabolic changes.</a> eLife. 10, 67753 (2021).</li><li>Demerec, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Biology of Drosophila. , (1965).</li><li>Rizki, R. 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The authors declare no conflicts of interest.

The present protocol describes the methods to (1) generate flies carrying mutant clones in the larval fat bodies, (2) induce autophagy through amino acid starvation, and (3) dissect the larval fat bodies. In order to generate clones successfully in the larval fat bodies, the following critical steps need to be carried out diligently. (1) Timing the heat shock accurately is crucial because mitotic recombination only happens when the tissue is undergoing mitosis, and (2) both heat shock temperature and duration are critica...

Autophagy is a "self-eating" process induced by various stresses, including amino acid starvation1. Macroautophagy (hereafter referred to as autophagy) is the most well-studied type of autophagy and plays an irreplaceable role in maintaining cellular homeostasis2. The malfunction of autophagy is associated with several human diseases3. In addition, some autophagy-related genes are potential targets for treating various diseases4. 
Autophagy is regulated in a highly sophisticated manner5. Upon starvation, the isolation membranes sequester cytoplasmic materials to form double-membraned autophagosomes6. Autophagosomes then fuse with endosomes and lysosomes to form amphisomes and autolysosomes. With the help of lysosomal hydrolytic enzymes, the engulfed cytoplasmic contents are degraded, and the nutrients are recycled7. 
Autophagy is an evolutionarily conserved process8. Drosophila melanogaster is a great model for studying the autophagy process in vivo. Amino acid starvation easily induces autophagy in fly fat body tissue, an analog of human liver and adipose tissue9. Defects in autophagy disrupt the distinct puncta patterns of several autophagy-related proteins, such as Atg8, Atg9, Atg18, Syx17, Rab7, LAMP1, and p62, among others10. Therefore, analyzing the patterns of these autophagy markers will help discern the occurrence of autophagy defects and the defective autophagy step. For example, the ubiquitin-like protein Atg8 is the most commonly used autophagy marker11. In Drosophila melanogaster, transgenic strains with a green fluorescent protein (GFP)-tagged Atg8a have been successfully developed12. GFP-Atg8a is diffused in the cytosol and nuclei in the fed cells. Upon starvation, GFP-Atg8a is processed and modified by phosphatidylethanolamine (PE) and forms puncta, which label the isolation membranes and fully developed autophagosomes13,14. Through direct fluorescence microscopy, the autophagy induction can be easily observed as an increase in GFP-Atg8 puncta formation15. Atg8a puncta would not form in response to starvation in the presence of an autophagy initiation defect. As GFP-Atg8a can be quenched and digested by the low pH in autolysosomes, GFP-Atg8a puncta may increase in numbers if autophagy is blocked at late stages16.
As autophagy is highly sensitive to nutrition availability17, slight differences in culture conditions often lead to variations in phenotypes. Therefore, clonal analysis, a method that analyzes mutant cells versus wild-type control cells in the same tissue, has a major advantage in dissecting autophagy defects18. Taking advantage of flippase/flippase recognition target (FLP/FRT)-mediated site-specific recombination between homologous chromosomes, flies carrying mosaic tissues are readily made19,20. The wild-type cells surrounding the mutant cells form a perfect internal control to avoid individual differences21.
The present study describes how to induce autophagy by amino acid starvation and generate GFP-Atg8a-expressing mosaic fat body tissues. These protocols can be used for analyzing differences in autophagy among mutant clones.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>1.5 mL microcentrifuge tube</td>
			<td>Axygen</td>
			<td>MCT-150-C</td>
			<td></td>
		</tr>
		<tr>
			<td>#5 Forceps</td>
			<td>Dumont</td>
			<td>RS-5015</td>
			<td></td>
		</tr>
		<tr>
			<td>9 Dressions Spot plate</td>
			<td>PYREX</td>
			<td>7220-85</td>
			<td></td>
		</tr>
		<tr>
			<td>Fluorescence Microscope</td>
			<td>Nikon</td>
			<td>SMZ1500</td>
			<td></td>
		</tr>
		<tr>
			<td>Glycerol</td>
			<td>Sangon Biotech</td>
			<td>A100854-0100</td>
			<td></td>
		</tr>
		<tr>
			<td>KCl</td>
			<td>Sangon Biotech</td>
			<td>A610440-0500</td>
			<td>Composition of 1x PBS solution</td>
		</tr>
		<tr>
			<td>KH2PO4</td>
			<td>Sangon Biotech</td>
			<td>A600445-0500</td>
			<td>Composition of 1x PBS solution</td>
		</tr>
		<tr>
			<td>Laboratory spatula</td>
			<td>Fisher</td>
			<td>14-375-10</td>
			<td></td>
		</tr>
		<tr>
			<td>Long forceps</td>
			<td>R&#39; DEER</td>
			<td>RST-14</td>
			<td></td>
		</tr>
		<tr>
			<td>Microscope cover glass</td>
			<td>CITOTEST</td>
			<td>80340-1130</td>
			<td></td>
		</tr>
		<tr>
			<td>Microscope slides</td>
			<td>CITOTEST</td>
			<td>80302-2104</td>
			<td></td>
		</tr>
		<tr>
			<td>Na2HPO4</td>
			<td>Sangon Biotech</td>
			<td>A501727-0500</td>
			<td>Composition of 1x PBS solution</td>
		</tr>
		<tr>
			<td>NaCl</td>
			<td>Sangon Biotech</td>
			<td>A610476-0005</td>
			<td>Composition of 1x PBS solution</td>
		</tr>
		<tr>
			<td>Paraformaldehyde</td>
			<td>Sigma-Aldrich</td>
			<td>158127</td>
			<td></td>
		</tr>
		<tr>
			<td>Petri dish</td>
			<td>Corning</td>
			<td>430166</td>
			<td></td>
		</tr>
		<tr>
			<td>Standard cornmeal/molasses/agar fly food</td>
			<td></td>
			<td></td>
			<td>Tong Lab-made</td>
		</tr>
		<tr>
			<td>Stereo microscope</td>
			<td>Nikon</td>
			<td>SMZ745</td>
			<td></td>
		</tr>
		<tr>
			<td>Sucrose</td>
			<td>Sinopharm Chemical Reagent Co.,Ltd.</td>
			<td>10021418</td>
			<td></td>
		</tr>
		<tr>
			<td>Vectashield antifade mounting medium with DAPI</td>
			<td>Vectorlabratory</td>
			<td>H-1200-10</td>
			<td>Recommended mounting medium</td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Fly stocks</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>y&#39;w* Iso FRT19A</td>
			<td></td>
			<td></td>
			<td>Tong Lab&#39;s fly stocks</td>
		</tr>
		<tr>
			<td>y&#39;w* Mu1FRT19A/ FM7,Kr GFP</td>
			<td></td>
			<td></td>
			<td>Tong Lab&#39;s fly stocks</td>
		</tr>
		<tr>
			<td>y&#39;w* Mu2 FRT19A/ FM7,Kr GFP</td>
			<td></td>
			<td></td>
			<td>Tong Lab&#39;s fly stocks</td>
		</tr>
		<tr>
			<td>hsFLP ubiRFP FRT19A; cgGal4 UAS-GFP-Atg8a</td>
			<td></td>
			<td></td>
			<td>Tong Lab&#39;s fly stocks</td>
		</tr>
	</tbody>
</table>

analyzing starvation-induced autophagy in the drosophila melanogaster larval fat body

Autophagy is a cellular self-digestion process. It delivers cargo to the lysosomes for degradation in response to various stresses, including starvation. The malfunction of autophagy is associated with aging and multiple human diseases. The autophagy machinery is highly conserved-from yeast to humans.&#160;The larval fat body of Drosophila melanogaster, an analog for vertebrate liver and adipose tissue, provides a unique model for monitoring autophagy in vivo. Autophagy can be easily induced by nutrient starvation in the larval fat body. Most autophagy-related genes are conserved in Drosophila. Many transgenic fly strains expressing tagged autophagy markers have been developed, which facilitates the monitoring of different steps in the autophagy process. The clonal analysis enables a close comparison of autophagy markers in cells with different genotypes in the same piece of tissue. The current protocol details procedures for (1) generating somatic clones in the larval fat body, (2) inducing autophagy via amino acid starvation, and (3) dissecting the larval fat body, aiming to create a model for analyzing differences in autophagy using an autophagosome marker (GFP-Atg8a) and clonal analysis.

Under fed conditions, the GFP-tagged ubiquitin-like protein, GFP-Atg8a, is diffused inside the cells. Upon starvation, it forms green puncta and labels autophagosomes. Once autophagosomes fuse with lysosomes, GFP is quenched in the acidic autolysosomes, and the green puncta disappear. If autophagy is not induced or the autophagosome maturation is accelerated, the number of GFP puncta is expected to be low. However, if the fusion between autophagosomes and lysosomes is blocked or the pH of the autolysosome becomes basic, ...

Watch this Scientific Journal Video about Analyzing Starvation-Induced Autophagy in the Drosophila melanogaster Larval Fat Body at JoVE.com

Analyzing Starvation-Induced Autophagy in the Drosophila melanogaster Larval Fat Body

The present protocol describes the induction of autophagy in the Drosophila melanogaster larval fat body via nutrient depletion and analyzes changes in autophagy using transgenic fly strains.

Analyzing Starvation-Induced Autophagy in the Drosophila melanogaster Larval Fat Body

Autophagy is a cellular self-digestion process. It delivers cargo to the lysosomes for degradation in response to various stresses, including starvation. ...

analyzing-starvation-induced-autophagy-drosophila-melanogaster-larval

Research

JoVE Journal

Biology

2.2K Views.  Zhejiang University. The present protocol describes the induction of autophagy in the Drosophila melanogaster larval fat body via nutrient depletion and analyzes changes in autophagy using transgenic fly strains.

Video: Analyzing Starvation-Induced Autophagy in the Drosophila melanogaster Larval Fat Body

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.

Dissection z larw CNS na 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 ...

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 ...

Drosophila Larval NMJ Immunohistochemistry

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 31 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. Immunohistochemistry can be used to clearly image the components of the NMJ. In this article, we demonstrate how to use antibody staining to visualize the Drosophila larval NMJ.

Drosophila Larval NMJ Immunohistochemistry

This protocol demonstrates how to perform immunohistochemistry on dissected Drosophila larva.

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  ...

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 ...

Isolation of Drosophila melanogaster Testes

The testes of Drosophila melanogaster provide an important model for the study of stem cell maintenance and differentiation, meiosis, and soma-germline interactions. Testes are typically isolated from adult males 0-3 days after eclosion from the pupal case. The testes of wild-type flies are easily distinguished from other tissues because they are yellow, but the testes of white mutant flies, a common genetic background for laboratory experiments are similar in both shape and color to the fly gut. Performing dissection on a glass microscope slide with a black background makes identifying the testes considerably easier. Testes are removed from the flies using dissecting needles. Compared to protocols that use forceps for testes dissection, our method is far quicker, allowing a well-practiced individual to dissect testes from 200-300 wild-type flies per hour, yielding 400-600 testes. Testes from white flies or from mutants that reduce testes size are harder to dissect and typically yield 200-400 testes per hour.

Isolation of Drosophila melanogaster Testes

Drosophila melanogaster testes can be rapidly and efficiently isolated from adult males using dissecting needles. With practice, one can readily isolate in one or two days an amount of testes sufficient for the analysis of DNA or RNA by high throughput sequencing or more traditional molecular biology methods or of protein for antibody- or enzyme-based assays.

The testes of Drosophila melanogaster provide an important model for the study of stem cell maintenance and differentiation, meiosis, and soma-germline ...

Measurement of Lifespan in Drosophila melanogaster

Aging is a phenomenon that results in steady physiological deterioration in nearly all organisms in which it has been examined, leading to reduced physical performance and increased risk of disease. Individual aging is manifest at the population level as an increase in age-dependent mortality, which is often measured in the laboratory by observing lifespan in large cohorts of age-matched individuals. Experiments that seek to quantify the extent to which genetic or environmental manipulations impact lifespan in simple model organisms have been remarkably successful for understanding the aspects of aging that are conserved across taxa and for inspiring new strategies for extending lifespan and preventing age-associated disease in mammals.
The vinegar fly, Drosophila melanogaster, is an attractive model organism for studying the mechanisms of aging due to its relatively short lifespan, convenient husbandry, and facile genetics. However, demographic measures of aging, including age-specific survival and mortality, are extraordinarily susceptible to even minor variations in experimental design and environment, and the maintenance of strict laboratory practices for the duration of aging experiments is required. These considerations, together with the need to practice careful control of genetic background, are essential for generating robust measurements. Indeed, there are many notable controversies surrounding inference from longevity experiments in yeast, worms, flies and mice that have been traced to environmental or genetic artifacts1-4. In this protocol, we describe a set of procedures that have been optimized over many years of measuring longevity in Drosophila using laboratory vials. We also describe the use of the dLife software, which was developed by our laboratory and is available for download (<a href="http://sitemaker.umich.edu/pletcherlab/software">http://sitemaker.umich.edu/pletcherlab/software</a>). dLife accelerates throughput and promotes good practices by incorporating optimal experimental design, simplifying fly handling and data collection, and standardizing data analysis. We will also discuss the many potential pitfalls in the design, collection, and interpretation of lifespan data, and we provide steps to avoid these dangers.

Measurement of Lifespan in Drosophila melanogaster

Drosophila melanogaster is a powerful model organism for exploring the molecular basis of longevity regulation. This protocol will discuss the steps involved in generating a reproducible, population-based measurement of longevity as well as potential pitfalls and how to avoid them.

Aging is a phenomenon that results in steady physiological deterioration in nearly all organisms in which it has been examined, leading to reduced ...

Imaging Through the Pupal Case of Drosophila melanogaster

The longstanding use of Drosophila as a model for cell and developmental biology has yielded an array of tools. Together, these techniques have enabled analysis of cell and developmental biology from a variety of methodological angles. Live imaging is an emerging method for observing dynamic cell processes, such as cell division or cell motility. Having isolated mutations in uncharacterized putative cell cycle proteins it became essential to observe mitosis in situ using live imaging. Most live imaging studies in Drosophila have focused on the embryonic stages that are accessible to manipulation and observation because of their small size and optical clarity. However, in these stages the cell cycle is unusual in that it lacks one or both of the gap phases. By contrast, cells of the pupal wing of Drosophila have a typical cell cycle and undergo a period of rapid mitosis spanning about 20 hr of pupal development. It is easy to identify and isolate pupae of the appropriate stage to catch mitosis in situ. Mounting intact pupae provided the best combination of tractability and durability during imaging, allowing experiments to run for several hours with minimal impact on cell and animal viability. The method allows observation of features as small as, or smaller than, fly chromosomes. Adjustment of microscope settings and the details of mounting, allowed extension of the preparation to visualize membrane dynamics of adjacent cells and fluorescently labeled proteins such as tubulin. This method works for all tested fluorescent proteins and can capture submicron scale features over a variety of time scales. While limited to the outer 20 &#181;m of the pupa with a conventional confocal microscope, this approach to observing protein and cellular dynamics in pupal tissues in vivo may be generally useful in the study of cell and developmental biology in these tissues.

Imaging Through the Pupal Case of Drosophila melanogaster

This paper demonstrates the use of a fast scanning confocal microscope to image cell behavior directly through the puparium. By leaving the pupal case intact, this method allows observation and measurement of dynamic cell processes at a stage of Drosophila development that is difficult to study directly.

The longstanding use of Drosophila as a model for cell and developmental biology has yielded an array of tools. Together, these techniques have enabled ...

Measurement of Larval Activity in the Drosophila Activity Monitor

Drosophila larvae are used in many behavioral studies, yet a simple device for measuring basic parameters of larval activity has not been available. This protocol repurposes an instrument often used to measure adult activity, the TriKinetics Drosophila activity monitor (MB5 Multi-Beam Activity Monitor) to study larval activity. The instrument can monitor the movements of animals in 16 individual 8 cm glass assay tubes, using 17 infrared detection beams per tube. Logging software automatically saves data to a computer, recording parameters such as number of moves, times sensors were triggered, and animals&#8217; positions within the tubes. The data can then be analyzed to represent overall locomotion and/or position preference as well as other measurements. All data are easily accessible and compatible with basic graphing and data manipulation software. This protocol will discuss how to use the apparatus, how to operate the software and how to run a larval activity assay from start to finish.

Measurement of Larval Activity in the Drosophila Activity Monitor

This report describes a method for measuring Drosophila larval activity using the TriKinetics Drosophila Activity Monitor. The device employs infrared beams to detect movements of up to 16 individual animals. Data can be analyzed to represent motion parameters including rates and the positions of the animals within the assay chambers.

Drosophila larvae are used in many behavioral studies, yet a simple device for measuring basic parameters of larval activity has not been available. This ...