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.

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		<tr height="21">
			<td height="21" style="height:21px;width:353px;">Unsulfured blackstrap molasses</td>
			<td style="width:141px;">Good Food, INC</td>
			<td style="width:141px;"></td>
			<td style="width:191px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Drosophila Agar Type II</td>
			<td>Genesee Scientific</td>
			<td>66-103</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Pyridine</td>
			<td>EMD Millipore</td>
			<td>PX2012-7</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Methoxyamine hydrocholoride (MOX)</td>
			<td>MP Biomedicals, LLC</td>
			<td>155405</td>
			<td style="width:191px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">MSTFA with 1% trimethylchlorosilane</td>
			<td>Sigma</td>
			<td>69478</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Fleischmann&#8217;s Active dry yeast</td>
			<td>AB Mauri Food Inc</td>
			<td>2192</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">6oz Drosophila stock bottle</td>
			<td>Genesee Scientific</td>
			<td>32-130</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Soft tissue homogenizing mix (2 mL tubes)&#160;</td>
			<td>Omni International</td>
			<td>SKU:19-627</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Vial insert, 250 &#181;L deactivated glass with polymer feet</td>
			<td>Agilent</td>
			<td>5181-8872</td>
			<td></td>
		</tr>
		<tr height="24">
			<td height="24" style="height:24px;">Succinic acid-2,2,3,3-d4</td>
			<td>Sigma</td>
			<td>293075</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">SpeedVac</td>
			<td>Thermo&#160;</td>
			<td>SC210A</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">o-Phosphoric acid</td>
			<td>Fisher Scientific</td>
			<td>A242-1</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Propionic acid</td>
			<td>Sigma</td>
			<td>P5561</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">p-Hydroxy benzoic acid methyl ester</td>
			<td>Genesee Scientific</td>
			<td>20-258</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Bead Ruptor</td>
			<td>Omni International</td>
			<td>SKU:19-040E</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">ThermoMixer F1.5</td>
			<td>Eppendorf</td>
			<td>5384000012</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">MultiTherm Shaker with a 24 X 12 mm block</td>
			<td>Benchmark Scientific</td>
			<td>H5000</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Methanol</td>
			<td>Sigma</td>
			<td>34860</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">1.5 mL centrifuge tube</td>
			<td>Eppendorf</td>
			<td>22364111</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Falcon 35 X 10 mm tissue culture dish</td>
			<td>Corning Incorporated</td>
			<td>353001</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">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

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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 ...

Dissection of Larval Zebrafish Gonadal Tissue

Although wild zebrafish possess a ZZ/ZW sex-determination system, domesticated zebrafish have lost the sex chromosome. They utilize a polygenic sex determination system, where several genes distributed throughout the genome collectively determine the sex identities of individual fish. Currently, the genes involved in regulating gonad development and how they work remain elusive. Normally, isolating gonadal tissue is the first step to examine the sex developmental processes. Here, we present a procedure to isolate gonadal tissue from 17 dpf (days post fertilization) and 25 dpf zebrafish larvae. The isolated gonadal tissue may be subsequently examined by morphology and gene expression profiling.

Here, we present a protocol for isolating gonadal tissue of larval zebrafish, which will facilitate investigations of zebrafish sex differentiation and maintenance.

Although wild zebrafish possess a ZZ/ZW sex-determination system, domesticated zebrafish have lost the sex chromosome. They utilize a polygenic sex ...

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, ...

Establishment of Larval Zebrafish as an Animal Model to Investigate Trypanosoma cruzi Motility In Vivo

Chagas disease is a parasitic infection caused by Trypanosoma cruzi, whose motility is not only important for localization, but also for cellular binding and invasion. Current animal models for the study of T. cruzi allow limited observation of parasites in vivo, representing a challenge for understanding parasite behavior during the initial stages of infection in humans. This protozoan has a flagellar stage in both vector and mammalian hosts, but there are no studies describing its motility in vivo.The objective of this project was to establish a live vertebrate zebrafish model to evaluate T. cruzi motility in the vascular system. Transparent zebrafish larvae were injected with fluorescently labeled trypomastigotes and observed using light sheet fluorescence microscopy (LSFM), a noninvasive method to visualize live organisms with high optical resolution. The parasites could be visualized for extended periods of time due to this technique&#39;s relatively low risk of photodamage compared to confocal or epifluorescence microscopy. T. cruzi parasites were observed traveling in the circulatory system of live zebrafish in different-sized blood vessels and the yolk. They could also be seen attached to the yolk sac wall and to the atrioventricular valve despite the strong forces associated with heart contractions. LSFM of T. cruzi-inoculated zebrafish larvae is a valuable method that can be used to visualize circulating parasites and evaluate their tropism, migration patterns, and motility in the dynamic environment of the cardiovascular system of a live animal.

Establishment of Larval Zebrafish as an Animal Model to Investigate Trypanosoma cruzi Motility In Vivo

In this protocol, fluorescently labeled T. cruzi&#160;were injected into transparent zebrafish larvae, and parasite motility was observed in vivo using light sheet fluorescence microscopy.

Chagas disease is a parasitic infection caused by Trypanosoma cruzi, whose motility is not only important for localization, but also for cellular binding ...

Thawing, Culturing, and Cryopreserving Drosophila Cell Lines

There are currently over 160 distinct Drosophila cell lines distributed by the Drosophila Genomics Resource Center (DGRC). With genome engineering, the number of novel cell lines is expected to increase. The DGRC aims to familiarize researchers with using Drosophila cell lines as an experimental tool to complement and drive their research agenda. Procedures for working with a variety of Drosophila cell lines with distinct characteristics are provided, including protocols for thawing, culturing, and cryopreserving cell lines. Importantly, this publication demonstrates the best practices required to work with Drosophila cell lines to minimize the risk of contaminations from adventitious microorganisms or from other cell lines. Researchers who become familiar with these procedures will be able to delve into the many applications that use Drosophila cultured cells including biochemistry, cell biology and functional genomics.

Thawing, Culturing, and Cryopreserving Drosophila Cell Lines

Drosophila cell lines are important reagents for both fundamental and biomedical research. This article provides protocols for thawing, subculturing, and the cryopreservation of commonly used Drosophila cell lines to assist researchers in incorporating the use of these reagents in their research.

There are currently over 160 distinct Drosophila cell lines distributed by the Drosophila Genomics Resource Center (DGRC). With genome engineering, the ...

Assay for Blood-brain Barrier Integrity in Drosophila melanogaster

Proper nervous system development includes the formation of the blood-brain barrier, the diffusion barrier that tightly regulates access to the nervous system and protects neural tissue from toxins and pathogens. Defects in the formation of this barrier have been correlated with neuropathies, and the breakdown of this barrier has been observed in many neurodegenerative diseases. Therefore, it is critical to identify the genes that regulate the formation and maintenance of the blood-brain barrier to identify potential therapeutic targets. In order to understand the exact roles these genes play in neural development, it is necessary to assay the effects of altered gene expression on the integrity of the blood-brain barrier. Many of the molecules that function in the establishment of the blood-brain barrier have been found to be conserved across eukaryotic species, including the fruit fly, Drosophila melanogaster. Fruit flies have proven to be an excellent model system for examining the molecular mechanisms regulating nervous system development and function. This protocol describes a step-by-step procedure to assay for blood-brain barrier integrity during the embryonic and larval stages of D. melanogaster development.

Assay for Blood-brain Barrier Integrity in Drosophila melanogaster

Blood-brain barrier integrity is critical for nervous system function. In Drosophila melanogaster, the blood-brain barrier is formed by glial cells during late embryogenesis. This protocol describes methods to assay for blood-brain barrier formation and maintenance in D. melanogaster embryos and third instar larvae.

Proper nervous system development includes the formation of the blood-brain barrier, the diffusion barrier that tightly regulates access to the nervous ...

Quantifying Liver Size in Larval Zebrafish Using Brightfield Microscopy

In several transgenic zebrafish models of hepatocellular carcinoma (HCC), hepatomegaly can be observed during early larval stages. Quantifying larval liver size in zebrafish HCC models provides a means to rapidly assess the effects of drugs and other manipulations on an oncogene-related phenotype. Here we show how to fix zebrafish larvae, dissect the tissues surrounding the liver, photograph livers using bright-field microscopy, measure liver area, and analyze results. This protocol enables rapid, precise quantification of liver size. As this method involves measuring liver area, it may underestimate differences in liver volume, and complementary methodologies are required to differentiate between changes in cell size and changes in cell number. The dissection technique described herein is an excellent tool to visualize the liver, gut, and pancreas in their natural positions for myriad downstream applications including immunofluorescence staining and in situ hybridization. The described strategy for quantifying larval liver size is applicable to many aspects of liver development and regeneration.

Here we demonstrate a method for quantifying liver size in larval zebrafish, providing a way to assess the effects of genetic and pharmacologic manipulations on liver growth and development.

In several transgenic zebrafish models of hepatocellular carcinoma (HCC), hepatomegaly can be observed during early larval stages. Quantifying larval ...

Development of a Larval Zebrafish Infection Model for Clostridioides difficile

Clostridioides difficile infection (CDI) is considered to be one of the most common healthcare-associated gastrointestinal infections in the United States. The innate immune response against C. difficile has been described, but the exact roles of neutrophils and macrophages in CDI are less understood. In the current study, Danio rerio (zebrafish) larvae are used to establish a C. difficile infection model for imaging the behavior and cooperation of these innate immune cells in vivo. To monitor C. difficile, a labeling protocol using a fluorescent dye has been established. A localized infection is achieved by microinjecting labeled C. difficile, which actively grows in the zebrafish intestinal tract and mimics the intestinal epithelial damage in CDI. However, this direct infection protocol is invasive and causes microscopic wounds, which can affect experimental results. Hence, a more noninvasive microgavage protocol is described here. The method involves delivery of C. difficile cells directly into the intestine of zebrafish larvae by intubation through the open mouth. This infection method closely mimics the natural infection route of C. difficile.

Development of a Larval Zebrafish Infection Model for Clostridioides difficile

Presented here is a safe and effective method to infect zebrafish larvae with fluorescently labeled anaerobic C. difficile by microinjection and noninvasive microgavage.

Clostridioides difficile infection (CDI) is considered to be one of the most common healthcare-associated gastrointestinal infections in the United ...

Preparation of Drosophila Larval and Pupal Testes for Analysis of Cell Division in Live, Intact Tissue

Experimental analysis of cells dividing in living, intact tissues and organs is essential to our understanding of how cell division integrates with development, tissue homeostasis, and disease processes. Drosophila spermatocytes undergoing meiosis are ideal for this analysis because (1) whole Drosophila testes containing spermatocytes are relatively easy to prepare for microscopy, (2) the spermatocytes&#8217; large size makes them well suited for high resolution imaging, and (3) powerful Drosophila genetic tools can be integrated with in vivo analysis. Here, we present a readily accessible protocol for the preparation of whole testes from Drosophila third instar larvae and early pupae. We describe how to identify meiotic spermatocytes in prepared whole testes and how to image them live by time-lapse microscopy. Protocols for fixation and immunostaining whole testes are also provided. The use of larval testes has several advantages over available protocols that use adult testes for spermatocyte analysis. Most importantly, larval testes are smaller and less crowded with cells than adult testes, and this greatly facilitates high resolution imaging of spermatocytes. To demonstrate these advantages and the applications of the protocols, we present results showing the redistribution of the endoplasmic reticulum with respect to spindle microtubules during cell division in a single spermatocyte imaged by time-lapse confocal microscopy. The protocols can be combined with expression of any number of fluorescently tagged proteins or organelle markers, as well as gene mutations and other genetic tools, making this approach especially powerful for analysis of cell division mechanisms in the physiological context of whole tissues and organs.

Preparation of Drosophila Larval and Pupal Testes for Analysis of Cell Division in Live, Intact Tissue

The goal of this protocol is to analyze cell division in intact tissue by live and fixed cell microscopy using Drosophila meiotic spermatocytes. The protocol demonstrates how to isolate whole, intact testes from Drosophila larvae and early pupae, and how to process and mount them for microscopy.

Experimental analysis of cells dividing in living, intact tissues and organs is essential to our understanding of how cell division integrates with ...

Dissection, Immunohistochemistry and Mounting of Larval and Adult Drosophila Brains for Optic Lobe Visualization

The Drosophila optic lobe, comprised of four neuropils: the lamina, medulla, lobula and lobula plate, is an excellent model system for exploring the developmental mechanisms that generate neural diversity and drive circuit assembly. Given its complex three-dimensional organization, analysis of the optic lobe requires that one understand how its adult neuropils and larval progenitors are positioned relative to each other and the central brain. Here, we describe a protocol for the dissection, immunostaining and mounting of larval and adult brains for optic lobe imaging. Special emphasis is placed on the relationship between mounting orientation and the spatial organization of the optic lobe. We describe three mounting strategies in the larva (anterior, posterior and lateral) and three in the adult (anterior, posterior and horizontal), each of which provide an ideal imaging angle for a distinct optic lobe structure.

Dissection, Immunohistochemistry and Mounting of Larval and Adult Drosophila Brains for Optic Lobe Visualization

This protocol describes three steps to prepare larval and adult Drosophila optic lobes for imaging: 1) brain dissections, 2) immunohistochemistry and 3) mounting. Emphasis is placed on step 3, as distinct mounting orientations are required to visualize specific optic lobe structures.&#160;

The Drosophila optic lobe, comprised of four neuropils: the lamina, medulla, lobula and lobula plate, is an excellent model system for exploring the ...