Thanks to members of the Indiana University Mass Spectroscopy Facility and the University of Utah Metabolomics Core Facility for assistance in optimizing this protocol. J.M.T. is supported by the National Institute of General Medical Sciences of the National Institutes of Health under Award Number R35GM119557.

1. Egg Collection
<ol>
	<li>Collect adult males and virgin females of the desired genotypes. Individually age these animals in a food vial with standard Bloomington media for 3&#8211;5 days.</li>
	<li>Set up the appropriate matings by transferring 50 virgin females and 25 males to a new food vial. 
	NOTE: A minimum of six independent matings should be set up for each genotype. Only one sample will be collected from each mating (i.e., six samples collected from six independent matings).</li>
	<li>Prepare molasses egg-laying caps.
	<ol>
		<li>Mix 115 mL of molasses and 29 g of agar with 700 mL of H2O in a 2 L flask.</li>
		<li>Boil the molasses agar mixture on a hot plate.</li>
		<li>Cool to 70 &#176;C.</li>
		<li>Add 25 mL of acid mix (20 mL of 85% phosphoric acid and 209 mL propionic acid in 1 L of H2O; Store in a brown bottle) and 10 mL of 10% p-hydroxy-benzoic acid methyl ester in 95% ethanol.</li>
		<li>Pour the molasses agar into both the lid and bottom portion of a 35 x 10 mm2 tissue culture dish. 
		NOTE: Before pouring molasses agar plates, be sure that the lids and/or base of the 35 x 10 mm2 plate fit snuggly into the mouth of a Drosophila stock bottle (described in Step 1.6). Depending on the brand of plates used, the base might not be usable.</li>
	</ol>
	</li>
	<li>Make fresh yeast paste by adding distilled water to active dry yeast (5 g yeast/7 mL water) until the mixture becomes a paste that can be easily spread on a solid surface (i.e., the consistency of peanut butter).</li>
	<li>Spread approximately 1.5 g of yeast paste on the surface of a molasses egg-laying cap.</li>
	<li>Poke four air holes into opposite sides of a plastic 6 oz. Drosophila stock bottle using a 22 G needle.</li>
	<li>Transfer the newly mated flies from the food vial into culture bottles. 
	NOTE: Flies can be transferred by either using CO2 as a temporary anesthetic or by holding the top of vial in mouth of the bottle and sharply tapping the bottom of the bottle against a benchtop.</li>
	<li>Quickly place a molasses egg-laying cap with yeast paste on the surface into the mouth of a Drosophila stock bottle. Use laboratory tape to secure the egg-laying cap in place. 
	NOTE: If flies were anesthetized prior to transfer, the bottle should be placed horizontally on the benchtop until all flies have recovered. Once flies are completely mobile, place the bottle in an incubator with the molasses egg-laying cap at the bottom of the culture.</li>
	<li>Replace the molasses egg-laying cap at least once a day for the first two days by inverting the stock bottle, tapping the bottom of the bottle against the benchtop, removing the old egg cap, and immediately placing a new egg cap into the mouth of the bottle. Discard the old egg-laying cap. 
	NOTE: This step ensures that females are not holding their eggs and allows for collection of synchronized populations.</li>
	<li>Place a new egg-laying cap into the stock bottle after day 3, replace after 2 h, and discard. Remove and replace the second egg-laying cap after four hours. Label the bottom of this egg-laying cap with the date, time, and genotype. 
	NOTE: The eggs collected during this 4 h window will be used for analysis. This synchronization step is critical for analyzing specific developmental stages. Eggs can be collected in this manner for several days.</li>
	<li>Insert the egg-laying caps into a 60 x 15 mm2 Petri plate and place into an incubator at the desired temperature and humidity.</li>
	<li>Inspect the egg-laying caps every day for problems with starvation, population density or contamination. 
	NOTE: Larvae must have access to adequate food to ensure continuous development. If necessary, fresh yeast paste can be spread on all egg-laying caps that will be used in the experiment. In addition, if the larval population density is too high, larvae will begin to wander out of the culture plate. These samples should not be used for metabolomic analysis because the high population density and intermediate bouts of starvation experienced while wandering will result in inconsistent data. A density of ~80&#8211;100 middle second instar larvae (L2) per plate or 40&#8211;50 middle third instar larvae (L3) per plate is recommended. Samples that are visibly contaminated with bacteria or fungi should be discarded.</li>
</ol>
2. Larval Sample Collection
<ol>
	<li>Age larvae until the desired stage. If collecting L3 larvae, resynchronize samples at the L2&#8211;L3 molt. Resynchronization is commonly achieved using a previously described method that uses the anterior spiracles as a developmental milestone15. Alternatively, mid-L3 larvae can be synchronized using a sgs3 reporter gene16. 
	NOTE: In this regard, we find mid-L2 larvae (~60 h after egg laying) are ideally suited for most analyses because larval development is still relatively synchronous at this stage, carefully collected samples have adequate food to develop to this timepoint, and the number of animals per sample is manageable (see below). An additional synchronization step is necessary for L3 larvae because the numerous ecdysone pulses that occur during L3 development have dramatic effects on metabolism and will generate artifacts in unsynchronized populations.</li>
	<li>Use a dissecting needle to gently lift the yeast paste off the agar. Place the yeast onto a new molasses agar cap. 
	NOTE: Most larvae will eat at the interface between the molasses agar and the yeast paste.</li>
	<li>Use a small paintbrush to collect larvae from the newly exposed surface of the molasses agar. Place the larvae into a 1.5 mL centrifuge tube on ice. 
	NOTE: Appropriate sample sizes for each developmental stage are as follows: 50 first instar larvae (L1); 25 mid-L2 larvae; 20 early L3 larvae; 15 mid-L3 or late L3 larvae.</li>
	<li>Wash and freeze samples (steps 2.3 through 2.8) in small batches (four to six samples) while collecting a large sample set. Samples must be frozen within 20 min of placing larvae in tubes. 
	NOTE: We recommend using Eppendorf 1.5 mL Flex Tubes because other tubes may interfere with the sample transfer at step 3. Samples must be collected in microfuge tubes. Do not collect samples in the bead tubes described in step 3.1. Ceramic beads retain the NaCl wash solution, which results in inaccurate mass measurements and introduces contaminants into the sample.</li>
	<li>Add 1 mL of ice-cold 0.9% NaCl into the tube, close the lid, and vertically flip the tube to thoroughly to wash the larvae.</li>
	<li>Place the tube back on ice for ~30 s. During this time, larvae will sink to the bottom of the tube, but yeast will remain in suspension. Once all larvae have formed a loose pellet, remove the NaCl solution using a 1 mL pipette.</li>
	<li>Repeat step 2.4 and step 2.5 twice, or until the final wash solution is clear. 
	Note: Additional wash steps may be necessary if the collected sample contains an excessive amount of yeast.</li>
	<li>Centrifuge the samples at 2,000 &#215; g for 1 min at 4 &#176;C.</li>
	<li>Remove all residual solution using a 200 &#181;L pipette.</li>
	<li>Immediately freeze the sample in liquid nitrogen. 
	NOTE: The protocol can be paused at this step. Samples can be stored for up to 3 months at -80 &#176;C.</li>
</ol>
3. Transfer of Samples to Bead Tubes
<ol>
	<li>Each larval sample must be transferred from the 1.5 mL microfuge tube to a 2 mL screwcap tubes containing 1.4 mm ceramic beads. In preparation for this transfer step, use an ethanol-proof marker to label the side of a 2 mL screwcap bead tube (markings on the lid will be destroyed during step 4.4).</li>
	<li>Tare the mass of the labeled bead tube using an analytical balance capable of accurately measuring 0.01 mg.</li>
	<li>If larval samples were stored at -80 &#176;C, place sample tubes in liquid nitrogen prior to transfer.</li>
	<li>Use long forceps to remove the 1.5 mL sample tube from the liquid nitrogen dewar.</li>
	<li>Wearing nitrile gloves, grab the frozen tube, invert, and sharply pound the tube lid against the benchtop to dislodge the frozen pellet. Immediately pour the pellet into a pre-tared 2 mL screwcap bead tube. If the pellet fails to dislodge, return the tube to liquid nitrogen and repeat. 
	NOTE: Larval pellets will not dislodge from all microfuge tubes. If using a tube other than the recommended one, determine if larval pellets can be dislodged prior to collecting samples for analysis.</li>
	<li>Quickly measure the combined mass of the larval pellet and bead tube. Immediately place the sample tube into liquid nitrogen. The larval pellet mass will be used to normalize the metabolomic data. 
	NOTE: The protocol can be paused at this step. Samples can be stored for up to 3 months at -80 &#176;C.</li>
</ol>
4. Sample Extraction
<ol>
	<li>Place the sample tubes in a -20 &#176;C benchtop cooler.</li>
	<li>Add 0.8 mL of prechilled (-20 &#176;C) 90% methanol containing 2 &#181;g/mL succinic-d4 acid into each tube. Return the sample to the -20 &#176;C benchtop cooler. 
	NOTE: The succinic-d4 acid serves as an internal standard. Only use HPLC grade H2O and methanol. Since methanol and other organic solvents are volatile and difficult to accurately measure using air displacement pipettes, we recommend using positive-displacement pipettes for this step and all following steps.</li>
	<li>Set up a negative control by adding 0.8 mL of prechilled (-20 &#176;C) 90% methanol containing 2 &#181;g/mL succinic-d4 acid into an empty bead tube.</li>
	<li>Homogenize samples for 30 second at 6.45 m/s using a bead mill homogenizer located in a 4 &#176;C temperature control room. 
	NOTE: Failure to rapidly and completely homogenize the larval pellet is a common source of variability in metabolomics analysis. Only use homogenizers that are capable of destroying frozen larval tissues within 30 s.</li>
	<li>Return the homogenized samples tubes to the -20 &#176;C benchtop cooler and incubate them in a -20 &#176;C freezer for at least 1 h.</li>
	<li>Centrifuge the tubes at 20,000 &#215; g or maximum speed for 5 min at 4 &#176;C to remove the resulting precipitate.</li>
	<li>Transfer 600 &#181;L of the supernatant into a new 1.5 mL microcentrifuge tube. Do not disturb the precipitate. If the precipitate pellet becomes dislodged while pipetting the supernatant, return all supernatant to the tube and repeat step 4.6.</li>
	<li>Place the sample tubes in a vacuum centrifuge. Be sure that all tubes are open before sealing the centrifuge. Dry the samples at room temperature until all solvent is removed (this step usually takes ~16 h to complete). 
	NOTE: If necessary, the protocol can be paused at this step. Dried sample can be stored at -80 &#176;C.</li>
</ol>
5. Chemical Derivatization
<ol>
	<li>If the dried samples were stored at -80 &#176;C, place unopened sample tubes in a vacuum centrifuge and dry for 30 min. This step must be performed prior to opening the sample tube. 
	NOTE: This step removes the condensation that collects on the outside of the microfuge tube when removed from the freezer. This portion of the protocol is extremely sensitive to H2O and extra precaution must be taken to keep all reagents as dry as possible.</li>
	<li>Prepare a solution of 40 mg/mL methoxylamine hydrochloride (MOX) in anhydrous pyridine. Only use anhydrous pyridine. Store MOX and pyridine in a desiccator and prepare the MOX solution daily. 
	CAUTION: MOX and pyridine are toxic. Prepare this solution in the fume hood.
	<ol>
		<li>Use a heat gun to dry a 1 mL glass syringe.</li>
		<li>Insert the needle of the syringe into the bottle of anhydrous pyridine. Remove 1 mL of pyridine and add to a microfuge tube containing 40 mg of MOX.</li>
		<li>Flush the bottle of anhydrous pyridine with argon. Seal the bottle and return to the desiccator.</li>
		<li>Dissolve the MOX in the pyridine by incubating the tube in a thermal mixer at 35 &#176;C for 10 min at 600 rpm.</li>
	</ol>
	</li>
	<li>Add 40 &#181;L of 40 mg/mL MOX in anhydrous pyridine solution to the dried sample.</li>
	<li>Vortex for 10 s and briefly centrifuge (10,000 x g for 20 s).</li>
	<li>Incubate at 30 &#176;C for 1 h at 600 rpm in a thermal mixer.</li>
	<li>Centrifuge at 20,000 &#215; g or maximum speed for 5 min to remove the particle matter.</li>
	<li>Transfer 25 &#181;L of supernatant into an autosampler vial with a 250 &#181;L deactivated glass microvolume insert.</li>
	<li>Add 40 &#181;L of N-methyl-N-trimethylsilyltrifluoracetamide (MSTFA) containing 1% TMCS. 
	CAUTION: MSTFA is toxic. Perform this step in the fume hood. 
	NOTE: If available, this step can be completed by a Gerstel autosampler.</li>
	<li>Place a cap on the autosampler vial and seal using a crimper tool.</li>
	<li>Incubate the sample at 37 &#176;C for 1 hour with shaking (250 rpm).</li>
	<li>Prepare a fatty acid methyl ester standard (FAMES) solution as previously desceribed3. Add 3 &#181;L of FAMES to the autosampler vial using a robotic autosampler immediately prior to injection. 
	NOTE: FAMEs are used to calibrate the retention time shift and check the instrument performance. If no robotic autosampler is available, FAMEs can be added during step 5.8.</li>
</ol>
6. GC-MS Detection
NOTE: In most cases, the user will conduct this step with the assistance of a mass spectroscopy core facility. This protocol is designed to be used with a 30 m, GC column with a 5 m guard column.
<ol>
	<li>Randomize the sample order.</li>
	<li>Prepare the GC-MS.
	<ol>
		<li>Set the helium carrier gas flow rate to 1 mL/min.</li>
		<li>Set the inlet temperature to 250 &#176;C.</li>
		<li>Program the GC to execute the following temperature gradient:
		<ol>
			<li>Initial temperature to 95 &#176;C with a hold of 1 min.</li>
			<li>Increase the temperature to 110 &#176;C at the rate of 40 &#176;C/min with a hold of 2 min.</li>
			<li>Increase to 250 &#176;C by 5&#176;C/min ramp.</li>
			<li>Increase to 330 &#176;C at the rate of 25 &#176;C/min with a final hold of 4 min.</li>
		</ol>
		</li>
		<li>Set solvent delay to 3.5 min. 
		NOTE: The time can be changed according to the GC-MS system. The purpose of this step is to prevent MSTFA and MOX from damaging the detector.</li>
	</ol>
	</li>
	<li>Inject 1 &#181;L of the derivatized sample into the GC-MS (split ratio of 10:1). 
	NOTE: Inject 2 &#181;L of sample if the intensity of peaks is too low. The injection order of samples should be randomized.</li>
	<li>Operate the mass spectrometer in full scan mode over a mass range of 50&#8211;500 m/z.</li>
</ol>
7. Data Analysis
<ol>
	<li>Analyze metabolomic data using either a targeted or untargeted approach. Targeted analysis is focused on measuring the abundance of a defined set of metabolites, such as the lactate measurements that we describe below. In contrast, untargeted analysis uses an unbiased approach to identify any metabolic feature that is significantly changed between two sample sets. 
	NOTE: Our lab primarily uses the free programs MetAlign17 and MetaboAnalyst18,19 for data analysis. Since an adequate description of both the quality control, normalization, and data processing steps are beyond the scope of this manuscript, we refer the user to more detailed protocols that are devoted to data processing20,21,22. In addition, a flow chart describing the steps required for this analysis can be found elsewhere8.</li>
</ol>

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The authors have nothing to disclose.

Metabolomics provides an unparalleled opportunity to survey the metabolic reactions that compose intermediary metabolism. The sensitivity of this technology, however, renders data susceptible to genetic background, developmental cues, and a variety of environmental stresses, including temperature, humidity, population density, and nutrient availability. Therefore, a high quality and reproducible metabolomics analysis requires that samples be collected under highly controlled conditions. While several reviews emphasize th...

The fruit fly Drosophila melanogaster has emerged as an ideal system for studying the molecular mechanism that regulate intermediary metabolism. Not only are most metabolic pathways conserved between Drosophila and humans, but key nutrient sensors and growth regulators, such as insulin, Tor, and myc, are also active in the fly1,2. As a result, Drosophila can be used to explore the metabolic basis of human diseases ranging from diabetes and obesity to neurodegeneration and cancer. In this regard, Drosophila larval development provides the ideal framework in which to study a metabolic program known as aerobic glycolysis, or the Warburg effect. Just as many tumors use aerobic glycolysis to generate biomass from carbohydrates, so to do Drosophila larvae activate aerobic glycolysis to promote developmental growth3,4,5. These similarities between larval and tumor metabolism establish Drosophila as a key model for understanding how aerobic glycolysis is regulated in vivo.
Despite the fact that the fly has emerged as a popular model for studying metabolism, most Drosophila studies rely on methods that are designed to measure individual metabolites3, such as trehalose, triglycerides, or ATP. Since a specific protocol is required to measure each metabolite, assay-based studies are labor-intensive, expensive, and biased towards those compounds that can be measured using commercial kits. A solution to these limitations has emerged from the field of metabolomics, which provides a more efficient and unbiased means of studying Drosophila metabolism. Unlike an assay-based study, a single metabolomic analysis can simultaneously measure hundreds of small molecule metabolites and provide a comprehensive understanding of an organism&#39;s metabolic status6,7. This technique has significantly expanded the scope of Drosophila metabolic studies and represents the future of this emerging field8.
Metabolomic studies are primarily conducted using three technologies: (i) nuclear magnetic resonance (NMR), (ii) liquid chromatography-mass spectrometry (LC-MS), and (iii) gas chromatography-mass spectrometry (GC-MS)9. Each approach offers distinct advantages and disadvantages, and all of these technologies have been used to successfully study Drosophila metabolism. Since the research conducted in our lab is focused on small, polar metabolites, we primarily employ a GC-MS-based method. GC-MS provides the user with a number of advantages, including high reproducibility, peak resolution, sensitivity, and the availability of a standard electron impact (EI) spectral library, which allows for the rapid identification of discovered metabolic features10,11. The preparation of samples for GC-MS, however, is somewhat complex and requires a careful attention to detail. Samples must be collected, washed, weighed, and frozen in a manner that quickly quenches metabolic reactions. Furthermore, the fly carcass is resistant to standard homogenization protocols and requires a bead mill to ensure optimal metabolite extraction. Finally, samples analyzed by GC-MS must undergo chemical derivatization prior to detection12. While previously published methods describe all of these steps3,13,14, a visual protocol that would allow the novice user to reproducibly generate high quality data is still needed. Here we demonstrate how to prepare Drosophila larval samples for GC-MS-based metabolomics analysis. This protocol allows the user to reproducibly measure many of the small polar metabolites that compose central carbon metabolism.

<table><tbody><tr><td>Unsulfured blackstrap molasses</td><td>Good Food, INC</td><td></td><td></td></tr><tr><td>Drosophila Agar Type II</td><td>Genesee Scientific</td><td>66-103</td><td></td></tr><tr><td>Pyridine</td><td>EMD Millipore</td><td>PX2012-7</td><td></td></tr><tr><td>Methoxyamine hydrocholoride (MOX)</td><td>MP Biomedicals, LLC</td><td>155405</td><td></td></tr><tr><td>MSTFA with 1% trimethylchlorosilane</td><td>Sigma</td><td>69478</td><td></td></tr><tr><td>Fleischmann&amp;rsquo;s Active dry yeast</td><td>AB Mauri Food Inc</td><td>2192</td><td></td></tr><tr><td>6oz Drosophila stock bottle</td><td>Genesee Scientific</td><td>32-130</td><td></td></tr><tr><td>Soft tissue homogenizing mix (2 mL tubes)&amp;nbsp;</td><td>Omni International</td><td>SKU:19-627</td><td></td></tr><tr><td>Vial insert, 250 &amp;micro;L deactivated glass with polymer feet</td><td>Agilent</td><td>5181-8872</td><td></td></tr><tr><td>Succinic acid-2,2,3,3-d&lt;sub&gt;4&lt;/sub&gt;</td><td>Sigma</td><td>293075</td><td></td></tr><tr><td>SpeedVac</td><td>Thermo&amp;nbsp;</td><td>SC210A</td><td></td></tr><tr><td>o-Phosphoric acid</td><td>Fisher Scientific</td><td>A242-1</td><td></td></tr><tr><td>Propionic acid</td><td>Sigma</td><td>P5561</td><td></td></tr><tr><td>p-Hydroxy benzoic acid methyl ester</td><td>Genesee Scientific</td><td>20-258</td><td></td></tr><tr><td>Bead Ruptor</td><td>Omni International</td><td>SKU:19-040E</td><td></td></tr><tr><td>ThermoMixer F1.5</td><td>Eppendorf</td><td>5384000012</td><td></td></tr><tr><td>MultiTherm Shaker with a 24 X 12 mm block</td><td>Benchmark Scientific</td><td>H5000</td><td></td></tr><tr><td>Methanol</td><td>Sigma</td><td>34860</td><td></td></tr><tr><td>1.5 mL centrifuge tube</td><td>Eppendorf</td><td>22364111</td><td></td></tr><tr><td>Falcon 35 X 10 mm tissue culture dish</td><td>Corning Incorporated</td><td>353001</td><td></td></tr><tr><td>GC column</td><td>Phenomex</td><td>ZB-5MSi</td><td></td></tr></tbody></table>

preparation of drosophila larval samples for gas chromatography-mass spectrometry (gc-ms)-based metabolomics

Recent advances in the field of metabolomics have established the fruit fly Drosophila melanogaster as a powerful genetic model for studying animal metabolism. By combining the vast array of Drosophila genetic tools with the ability to survey large swaths of intermediary metabolism, a metabolomics approach can reveal complex interactions between diet, genotype, life-history events, and environmental cues. In addition, metabolomics studies can discover novel enzymatic mechanisms and uncover previously unknown connections between seemingly disparate metabolic pathways. In order to facilitate more widespread use of this technology among the Drosophila community, here we provide a detailed protocol that describes how to prepare Drosophila larval samples for gas chromatography-mass spectrometry (GC-MS)-based metabolomic analysis. Our protocol includes descriptions of larval sample collection, metabolite extraction, chemical derivatization, and GC-MS analysis. Successful completion of this protocol will allow users to measure the relative abundance of small polar metabolites, including amino acids, sugars, and organic acids involved in glycolysis and the TCA cycles.

Lactate dehydrogenase (dLDH) mutants, which lack dLDH activity4, and genetically-matched controls were collected as mid-L2 larvae and processed according to protocol described above. When compared with controls, mutant larvae exhibit significant changes in lactate, pyruvate, and L-2-hydroxyglutarate4. Spectra were acquired with an Agilent GC6890-5973i MS system. An example of the GC-MS spectra generate with our protocol is shown in ...

Watch this Scientific Journal Video about Preparation of Drosophila Larval Samples for Gas Chromatography-Mass Spectrometry GC-MS-based Metabolomics at JoVE.com

Preparation of Drosophila Larval Samples for Gas Chromatography-Mass Spectrometry GC-MS-based Metabolomics

This protocol describes how to prepare Drosophila larvae for GC-MS-based metabolomic analysis.

Preparation of Drosophila Larval Samples for Gas Chromatography-Mass Spectrometry (GC-MS)-based Metabolomics

Recent advances in the field of metabolomics have established the fruit fly Drosophila melanogaster as a powerful genetic model for studying animal ...

preparation-drosophila-larval-samples-for-gas-chromatography-mass

Research

JoVE Journal

Developmental Biology

11.0K Views.  Indiana University. This protocol describes how to prepare Drosophila larvae for GC-MS-based metabolomic analysis.

Video: Preparation of Drosophila Larval Samples for Gas Chromatography-Mass Spectrometry GC-MS-based Metabolomics

Preparation of Mitochondrial Enriched Fractions for Metabolic Analysis in Drosophila

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

Preparation of Mitochondrial Enriched Fractions for Metabolic Analysis in Drosophila

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

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

Genome-wide Surveillance of Transcription Errors in Eukaryotic Organisms

Accurate transcription is required for the faithful expression of genetic information. Surprisingly though, little is known about the mechanisms that control the fidelity of transcription. To fill this gap in scientific knowledge, we recently optimized the circle-sequencing assay to detect transcription errors throughout the transcriptome of Saccharomyces cerevisiae, Drosophila melanogaster, and Caenorhabditis elegans. This protocol will provide researchers with a powerful new tool to map the landscape of transcription errors in eukaryotic cells so that the mechanisms that control the fidelity of transcription can be elucidated in unprecedented detail.

This protocol provides researchers with a new tool to monitor the fidelity of transcription in multiple model organisms.

Accurate transcription is required for the faithful expression of genetic information. Surprisingly though, little is known about the mechanisms that ...

Genetics

Medium-scale Preparation of Drosophila Embryo Extracts for Proteomic Experiments

Analysis of protein-protein interactions (PPIs) has become an indispensable approach to study biological processes and mechanisms, such as cell signaling, organism development, and disease. It is often desirable to obtain PPI information using in vivo material, to gain the most natural and unbiased view of the interaction networks. The fruit fly Drosophila melanogaster is an excellent platform to study PPIs in vivo, and lends itself to straightforward approaches to isolating material for biochemical experiments. In particular, fruit fly embryos represent a convenient type of tissue to study PPIs, due to the ease of collecting animals at this developmental stage and the fact that the majority of proteins are expressed in embryogenesis, thus providing a relevant environment to reveal most PPIs. Here we present a protocol for collection of Drosophila embryos at medium scale (0.5-1 g), which is an ideal amount for a wide range of proteomic applications, including analysis of PPIs by affinity purification-mass spectrometry (AP-MS). We describe our designs for 1 L and 5 L cages for embryo collections that can be easily and inexpensively set up in any laboratory. We also provide a general protocol for embryo collection and protein extraction to generate lysates that can be directly used in downstream applications such as AP-MS. Our goal is to provide an accessible means for all researchers to carry out the analyses of PPIs in vivo.

Medium-scale Preparation of Drosophila Embryo Extracts for Proteomic Experiments

The goal of this protocol is to provide a straightforward and inexpensive approach to collecting Drosophila embryos at medium scale (0.5-1 g) and preparing protein extracts that can be used in downstream proteomic applications, such as affinity purification-mass spectrometry (AP-MS).

Analysis of protein-protein interactions (PPIs) has become an indispensable approach to study biological processes and mechanisms, such as cell signaling, ...

Analysis of Lipid Signaling in Drosophila Photoreceptors using Mass Spectrometry

The activation of phospholipase C&#946; (PLC&#946;) is an essential step during sensory transduction in Drosophila photoreceptors. PLC&#946; activity results in the hydrolysis of the membrane lipid phosphatidylinositol 4,5 bisphosphate [PI(4,5)P2] leading ultimately to the activation of transient receptor potential (TRP) and TRP like (TRPL) channels. The activity of PLC&#946; also leads subsequently to the generation of many lipid species several of which have been proposed to play a role in TRP and TRPL activation. In addition, several classes of lipids have been proposed to play key roles in organizing the cell biology of photoreceptors to optimize signaling reactions for optimal sensory transduction. Historically, these discoveries have been driven by the ability to isolate Drosophila mutants for enzymes that control the levels of specific lipids and perform analysis of photoreceptor physiology in these mutants. More recently, powerful mass spectrometry methods for isolation and quantitative analysis of lipids with high sensitivity and specificity have been developed. These are particularly suited for use in Drosophila where lipid analysis is now possible from photoreceptors without the need for radionuclide labeling. In this article, the conceptual and practical considerations in the use of lipid mass spectrometry for the robust, sensitive, and accurate quantitative assessment of various signaling lipids in Drosophila photoreceptors are covered. Along with existing methods in molecular genetics and physiological analysis such lipid is likely to enhance the power of photoreceptors as a model system for discoveries in biology.

Analysis of Lipid Signaling in Drosophila Photoreceptors using Mass Spectrometry

The manuscript presents versatile, robust, and sensitive mass spectrometry protocols to identify and quantify several classes of lipids from Drosophila photoreceptors.

The activation of phospholipase C&#946; (PLC&#946;) is an essential step during sensory transduction in Drosophila photoreceptors. PLC&#946; activity ...

Biochemistry

Metabolic Analysis of Drosophila melanogaster Larval and Adult Brains

This protocol describes a method for measuring the metabolism in Drosophila melanogaster larval and adult brains. Quantifying metabolism in whole organs provides a tissue-level understanding of energy utilization that cannot be captured when analyzing primary cells and cell lines. While this analysis is ex vivo, it allows for the measurement from a number of specialized cells working together to perform a function in one tissue and more closely models the in vivo organ. Metabolic reprogramming has been observed in many neurological diseases, including neoplasia, and neurodegenerative diseases. This protocol was developed to assist the D. melanogaster community&#39;s investigation of metabolism in neurological disease models using a commercially available metabolic analyzer. Measuring metabolism of whole brains in the metabolic analyzer is challenging due to the geometry of the brain. This analyzer requires samples to remain at the bottom of a 96-well plate. Cell samples and tissue punches can adhere to the surface of the cell plate or utilize spheroid plates, respectively. However, the spherical, three-dimensional shape of D. melanogaster brains prevents the tissue from adhering to the plate. This protocol requires a specially designed and manufactured micro-tissue restraint that circumvents this problem by preventing any movement of the brain while still allowing metabolic measurements from the analyzer&#39;s two solid-state sensor probes. Oxygen consumption and extracellular acidification rates are reproducible and sensitive to a treatment with metabolic inhibitors. With a minor optimization, this protocol can be adapted for use with any whole tissue and/or model system, provided that the sample size does not exceed the chamber generated by the restraint. While basal metabolic measurements and an analysis after a treatment with mitochondrial inhibitors are described within this protocol, countless experimental conditions, such as energy source preference and rearing environment, could be interrogated.

Metabolic Analysis of Drosophila melanogaster Larval and Adult Brains

We present a protocol for measuring oxygen consumption and extracellular acidification in Drosophila melanogaster larval and adult brains. A metabolic analyzer is utilized with an adapted and optimized protocol. Micro-tissue restraints are a critical component of this protocol and were designed and created specifically for their use in this analysis.

This protocol describes a method for measuring the metabolism in Drosophila melanogaster larval and adult brains. Quantifying metabolism in whole organs ...

Neuroscience

Purification of Transcripts and Metabolites from Drosophila Heads

For the last decade, we have tried to understand the molecular and cellular mechanisms of neuronal degeneration using Drosophila as a model organism. Although fruit flies provide obvious experimental advantages, research on neurodegenerative diseases has mostly relied on traditional techniques, including genetic interaction, histology, immunofluorescence, and protein biochemistry. These techniques are effective for mechanistic, hypothesis-driven studies, which lead to a detailed understanding of the role of single genes in well-defined biological problems. However, neurodegenerative diseases are highly complex and affect multiple cellular organelles and processes over time. The advent of new technologies and the omics age provides a unique opportunity to understand the global cellular perturbations underlying complex diseases. Flexible model organisms such as Drosophila are ideal for adapting these new technologies because of their strong annotation and high tractability. One challenge with these small animals, though, is the purification of enough informational molecules (DNA, mRNA, protein, metabolites) from highly relevant tissues such as fly brains. Other challenges consist of collecting large numbers of flies for experimental replicates (critical for statistical robustness) and developing consistent procedures for the purification of high-quality biological material. Here, we describe the procedures for collecting thousands of fly heads and the extraction of transcripts and metabolites to understand how global changes in gene expression and metabolism contribute to neurodegenerative diseases. These procedures are easily scalable and can be applied to the study of proteomic and epigenomic contributions to disease.

Purification of Transcripts and Metabolites from Drosophila Heads

We describe here the procedures for the extraction and purification of mRNA and metabolites from Drosophila heads. We are applying these techniques to better understand the cellular perturbations underlying neuronal degeneration. These methodologies can be easily scaled and adapted for other "omic" projects.

For the last decade, we have tried to understand the molecular and cellular mechanisms of neuronal degeneration using Drosophila as a model organism. ...

Biology

Optimizing Sample Preparation Process for Transmission Electron Microscopy of Neuromuscular Junctions in Drosophila Larvae

The Drosophila neuromuscular junction (NMJ) has emerged as a valuable model system in the field of neuroscience. The application of confocal microscopy at the Drosophila NMJ enables researchers to acquire synaptic information, encompassing both quantitative data on synapse abundance and detailed insights into their morphology. However, the diffuse distribution and limited visual range of the TEM present challenges for the ultrastructural analysis. This study introduces an innovative and efficient sample preparation method that surpasses the conventional approach. The procedure begins by placing a metal mesh at the base of a flat-bottomed bottle or test tube, followed by positioning fixed larvae samples onto the mesh. An additional mesh is placed over the samples, ensuring that they are positioned between the two meshes. The fixed samples are thoroughly dehydrated and infiltrated before proceeding with the embedding procedure. Then embedding of the samples in epoxy resin is performed in a flat sheet manner, which allows for the preparation of muscles for positioning and sectioning. Benefiting from these steps, all the muscles of Drosophila larvae can be visualized under light microscopy, therey facilitating subsequent positioning and sectioning. Excess resin is removed after locating the 6th and 7th muscles of body segments A2 and A3. Serial ultra-thin sectioning of the 6th or 7th muscle is performed.

Optimizing Sample Preparation Process for Transmission Electron Microscopy of Neuromuscular Junctions in Drosophila Larvae

This report provides a new sample preparation procedure for visualizing neuromuscular junctions in Drosophila Larvae. This method is more effective in preventing the curling of the samples compared to the traditional method and is particularly useful for Drosophila neuromuscular junction ultrastructural analysis.

The Drosophila neuromuscular junction (NMJ) has emerged as a valuable model system in the field of neuroscience. The application of confocal microscopy at ...

Monitoring Brain Metabolite Transport in Drosophila Using FRET Sensors

Visualizing Monocarboxylates and Other Relevant Metabolites in the Ex Vivo Drosophila Larval Brain Using Genetically Encoded Sensors

The high energy requirements of brains due to electrical activity are one of their most distinguishing features. These requirements are met by the production of ATP from glucose and its metabolites, such as the monocarboxylates lactate and pyruvate. It is still unclear how this process is regulated or who the key players are, particularly in Drosophila.
Using genetically encoded F&#246;rster resonance energy transfer-based sensors, we present a simple method for measuring the transport of monocarboxylates and glucose in glial cells and neurons in an ex-vivo Drosophila larval brain preparation. The protocol describes how to dissect and adhere a larval brain expressing one of the sensors to a glass coverslip. 
We present the results of an entire experiment in which lactate transport was measured in larval brains by knocking down previously identified monocarboxylate transporters in glial cells. Furthermore, we demonstrate how to rapidly increase neuronal activity and track metabolite changes in the active brain. The described method, which provides all necessary information, can be used to analyze other Drosophila living tissues.

Author Spotlight: Advances in Brain Energy Metabolism Research Using the Drosophila Model

Here we present a protocol to visualize the transport of monocarboxylates, glucose, and ATP in glial cells and neurons using genetically encoded F&#246;rster resonance energy transfer-based sensors in an ex-vivo Drosophila larval brain preparation.

The high energy requirements of brains due to electrical activity are one of their most distinguishing features. These requirements are met by the ...

Sample Preparation for Rapid Lipid Analysis in Drosophila Brain Using Matrix-Assisted Laser Desorption/Ionization Mass Spectrometry Imaging

Lipid profiling, or lipidomics, is a well-established technique used to study the entire lipid content of a cell or tissue. Information acquired from&#160;lipidomics is valuable in studying the pathways involved in development, disease, and cellular metabolism. Many tools and instrumentations have aided lipidomics projects, most notably various combinations of mass spectrometry and liquid chromatography techniques. Matrix-assisted laser desorption/ionization mass spectrometry imaging (MALDI MSI) has recently emerged as a powerful imaging technique that complements conventional approaches. This novel technique provides unique information on the spatial distribution of lipids within tissue compartments, which was previously unattainable without the use of excessive modifications. The sample preparation of the MALDI MSI approach is critical and, therefore, is the focus of this paper. This paper presents a rapid lipid analysis of a large number of Drosophila brains embedded in optimal cutting temperature compound (OCT) to provide a detailed protocol for the preparation of small tissues for lipid analysis or metabolite and small molecule analysis through MALDI MSI.

Sample Preparation for Rapid Lipid Analysis in Drosophila Brain Using Matrix-Assisted Laser Desorption/Ionization Mass Spectrometry Imaging

The aim of this protocol is to provide detailed guidance on the proper sample preparation for lipid and metabolite analysis in small tissues, such as the Drosophila brain, using matrix-assisted laser desorption/ionization (MALDI) mass spectrometry imaging.

Lipid profiling, or lipidomics, is a well-established technique used to study the entire lipid content of a cell or tissue. Information acquired ...

High-Resolution Imaging and Analysis of Drosophila Eye Defects

A Rapid, Simple Workflow for Quantification of External Adult Drosophila Structures

The Drosophila compound eye is a precisely patterned tissue that has revealed molecular mechanisms and biological processes that drive morphogenesis. It is a simple structure of repeating unit eyes, termed ommatidia, that is used to characterize genetic interactions and gene functions. Mutations that affect eye architecture can be easily detected and analyzed; hence, this system is frequently used in under-resourced institutions. Further phenotypic analysis often includes a Scanning Electron Microscope (SEM) to generate high-magnification images suitable for quantitative analysis. However, SEMs are expensive and require costly reagents; sample preparation spans days; and, often, they need full-time staff for sample preparation and instrument maintenance. This limits their utility at under-resourced institutions or during budgetary austerity. In entomology, the use of high-resolution digital imaging technology is a common practice for the identification and characterization of species. This paper describes a method that combines strategies and allows for high-resolution digital imaging of adult Drosophila structures and quantitative analysis using the open software ImageJ. The workflow is a rapid and student-friendly alternative that remedies the limitations of underfunded and under-resourced research facilities with a cost-effective and rapid approach to quantitative phenotypic analysis.

Here, we present a rapid, low-cost, workflow for high-resolution imaging of adult Drosophila eyes to quantify patterning and growth defects. We describe our protocol for sample preparation by point-mounting, high-resolution image acquisition, and image analysis.

The Drosophila compound eye is a precisely patterned tissue that has revealed molecular mechanisms and biological processes that drive morphogenesis. It ...

Preparation of Drosophila Larval Samples for Gas Chromatography-Mass Spectrometry (GC-MS)-based Metabolomics

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Preparation of Mitochondrial Enriched Fractions for Metabolic Analysis in Drosophila

Genome-wide Surveillance of Transcription Errors in Eukaryotic Organisms

Medium-scale Preparation of Drosophila Embryo Extracts for Proteomic Experiments

Analysis of Lipid Signaling in Drosophila Photoreceptors using Mass Spectrometry

Metabolic Analysis of Drosophila melanogaster Larval and Adult Brains

Purification of Transcripts and Metabolites from Drosophila Heads

Optimizing Sample Preparation Process for Transmission Electron Microscopy of Neuromuscular Junctions in Drosophila Larvae

Visualizing Monocarboxylates and Other Relevant Metabolites in the Ex Vivo Drosophila Larval Brain Using Genetically Encoded Sensors

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A Rapid, Simple Workflow for Quantification of External Adult Drosophila Structures