Name: measurement of carbon dioxide production from radiolabeled substrates in drosophila melanogaster
Uploaded: 2016-06-27T13:00:00.000+00:00
Duration: 10 min 1 s
Description: Watch this Scientific Journal Video about Measurement of Carbon Dioxide Production from Radiolabeled Substrates in Drosophila melanogaster at JoVE.com

The author thanks Drs. Mingjian Lu and Morris Birnbaum for helpful advice and funding support from NIDDK (R21DK089391).

1. Preparation of Fly Food Containing Radiolabeled Metabolic Substrates
<ol>
	<li>In a microfuge tube, mix 1 - 2 &#181;Ci radiolabeled substrate (D-[6-14C]-glucose, D-[1-14C]-glucose or [1-14C]-palmitic acid, 40 - 60 mCi/mmol) with 15 &#181;l FD&#38;C No. 1 blue food dye and H2O to equal a total volume of 25 &#181;l. 
	CAUTION: Carbon-14 is a low-energy beta emitter and has a short range in air. However, take care to prevent spilling of radiolabeled molecules or accidental ingestion by the experimenter. Fly food vials tip over easily; be sure to house them in a container that will prevent them from tipping, especially when radiolabel is in liquid form (step 1.2) or in unsolidified food (step 1.4). Flies that are being fed or have been fed radiolabeled substrates will begin to exhale small amounts of radiolabeled CO2. These flies should be kept in acrylic boxes in fume hoods, and a small dish of KOH can be set in the box for use as a CO2 scrubber.</li>
	<li>Pipet all of the radiolabeled substrate and blue dye mix into the bottom of an empty fly food vial (height: 9.4 cm, width: 2.2 cm).</li>
	<li>Heat standard fly food in a microwave oven until food is just liquefied (15 - 20 sec at power level high for one vial containing 10 ml of food).</li>
	<li>Add 975 &#181;l molten food to the radiolabeled substrate/blue dye mix and quickly swirl, monitoring uniformity of blue color to ensure complete mixing.</li>
	<li>Allow food to cool and solidify completely at RT for 20 - 30 min.</li>
</ol>
2. Feeding Radiolabeled Metabolic Substrates to Adult Fruit Flies
<ol>
	<li>Anesthetize adult flies using a CO2 anesthetizing apparatus consisting of porous polyethylene fused to an acrylic base and connected to a CO2 tank with a pressure regulator. Flow CO2 into the anesthetizing apparatus at 5 L/min.
	<ol>
		<li>Transfer anesthetized flies to vials containing radiolabeled, blue-dyed food (n = 15 - 30 flies per vial). Cap vials with a foam stopper, and rest horizontally until flies wake up. Transfer vials to a lidded acrylic container (180 cm x 180 cm x 240 cm high).</li>
	</ol>
	</li>
	<li>For short-term feeding, starve flies for 18 - 24 hr before transferring flies to the radiolabeled food for a 2 - 3 hr feeding step. The starvation step is important because fed flies will not reliably eat a new source of food presented to them. For long-term feeding over the course of 5 - 7 days, flies need not be starved, but the experimenter should monitor the water content of the radiolabeled food in the vials flies are housed in and use a needle and syringe to add water to vials with dry food.</li>
	<li>Transfer flies to unlabeled food by &#34;tapping&#34; them into a new vial. An initial chase period of 2 - 4 hr allows flies to clean radiolabeled food particles from their cuticles and permits digestion of radiolabeled food remaining in the gut. Monitor progress of food through the gut by visual inspection of flies to look for blue abdomens.</li>
	<li>Subsequently, transfer flies to different food types (12 - 24 hr on standard food or 1% agar for measurement of effects of fed versus fasted state on respiration) or ambient temperatures (12 - 96 hr at 18 &#730;C or 30 &#730;C for genetic manipulations using temperature-sensitive GAL80 (GAL80ts), for e.g). 
	Note: The length of this chase period will vary with the experiment being performed. For e.g., a 96 hr chase at 30 &#730;C may be necessary in experiments using GAL80ts in order to fully activate GAL4-dependent transgene expression.</li>
</ol>
3. Preparation of the CO2-Collection Apparatus
<ol>
	<li>Assemble materials to construct &#34;fly pods&#34;, mesh-capped tubes that will contain flies labeled with 14C-glucose or 14C-palmitic acid and that will remain open to the atmosphere. Cut off the top 50 mm of 12 mm x 75 mm polypropylene tubes, leaving 25 mm-long, round-bottomed tubes. Prepare 35 mm x 35 mm squares of 130 &#181;m nylon mesh. Also, obtain transparent tape in a tape dispenser.</li>
	<li>Assemble materials for the CO2-collection apparatuses. For each fly pod, assemble a 20 ml glass scintillation vial, a rubber top stopper with an off-center hole, a center well to be inserted through the hole in the top stopper, grade GF/B glass microfiber filter paper (2.1 cm diameter circle, 1 &#181;m pore). GF/B filter paper is used because of its high wet strength and high loading capacity.</li>
	<li>Prepare 5% KOH fresh on the day of use. Just before step 3.4, fold and place the circular GF/B filter paper into the center well that is threaded through the hole of the rubber top stopper and saturate it with 100 &#181;l of 5% KOH. Refer to Figure 1.</li>
	<li>Following incubation(s) on unlabeled media, anesthetize flies with CO2. Brush flies into the cutoff polypropylene tube, cap with nylon mesh, and use transparent tape to adhere mesh to tube. Work quickly to avoid flies waking up and escaping before the mesh has been securely affixed to the tube. An equal number of flies should be used in each fly pod for a particular experiment; 10 - 20 flies per fly pod produce sufficient radiolabeled-CO2 for measurement by scintillation counting (step 4.1 below).</li>
	<li>Transfer the fly pod to a 20 ml glass scintillation vial. Cap the glass vial with the rubber top stopper holding a center well containing KOH-saturated GF/B filter paper. Fold the wide top of the rubber top stopper over the lip of the glass scintillation vial. Refer to Figure 1.</li>
	<li>Set glass vials containing flies in a lidded, acrylic container, and incubate for varying periods of time (2 to 12 hr works well).</li>
</ol>
4. Analysis of Results
<ol>
	<li>At the end of the incubation, uncap glass vials, and transfer KOH-saturated GF/B filter paper to a 6 ml plastic scintillation vial containing 4 ml scintillation cocktail.</li>
	<li>If needed, freeze flies in fly pods on dry ice and store at -80 &#730;C for later determination of stored substrates. Note that these samples are radioactive and must be stored accordingly.</li>
	<li>Prepare additional scintillation vials: one containing unused GF/B filter paper to serve as background and one to two others containing 0.1 - 0.5 &#181;Ci radiolabeled substrate to determine cpm/&#181;Ci.</li>
	<li>Measure cpm for each sample using a scintillation counter according to manufacturer&#39;s protocol.</li>
	<li>Calculate pmol 14CO2/cpm and pmol 14CO2 per fly per hour. Subtract the background cpm from the cpm value for each sample. From the neat radiolabeled substrate data, calculate the background-corrected cpm/&#181;Ci of sample counted.
	<ol>
		<li>Use the manufacturer-provided specific activity and radiochemical concentration of the radiolabeled substrate (for example, 50 &#181;Ci/&#181;mol and 0.1 &#181;Ci/&#181;l, respectively) to calculate pmol/cpm. Use this factor to convert background-corrected fly pod data from cpm 14CO2/sample to pmol 14CO2/sample. Then divide by the number of flies and the number of hr in the fly pod.</li>
	</ol>
	</li>
</ol>

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The author declares no competing financial interests.

This protocol describes a method for measuring the oxidation of specific radiolabeled substrates in adult Drosophila melanogaster. This technique is straightforward, quantitative, reproducible and sensitive, and it complements existing methods that measure total CO2 production and O2 consumption because it can be used to identify the nutrient(s) being oxidized for ATP production.
Measurement of CO2 liberated from the oxidation of specific radiolabeled substrates using the method described here is complementary to techniques that measure total CO2 production. Total CO2 production (VCO2) allows estimation of metabolic rate and, when total O2 consumption (VO2) is known, the metabolic rate can be calculated and the fuel source for oxidation can be estimated based on the respiratory exchange ratio (VCO2 / VO2 = RER). When carbohydrates are the exclusive substrate, RER = 1, but when fatty acids are the exclusive source, RER = 0.7, owing to the stoichiometry of the reactions of glucose and lipid oxidation. In the absence of equipment to measure oxygen consumption in Drosophila14, the oxidative fuel source cannot be identified. However, by labeling flies with trace amounts of one radioactive substrate or another and measuring rates of radiolabeled CO2 production, one can draw conclusions about the ability of flies to oxidize a given substrate at different time points during a stimulus or about the effects of a given mutation on fuel oxidation.
There are a number of critical steps in this protocol. First, care should be taken when using radioactive materials to avoid spills and contamination of research areas. Second, when anesthetizing flies during steps 2.1 and 3.4, the experimenter should work quickly to avoid effects of prolonged CO2 exposure on metabolism and behavior. A group of 20 flies can be anesthetized and transferred to a new vial or into a fly pod in 20 sec&#160;or less. When transferred from an anesthesia apparatus to a food vial, the vial should be rested on its side to prevent anesthetized flies from getting stuck on the food surface. When flies are fed radiolabeled food over the course of several days as in step 2.2, the experimenter should insure that the food does not dry out as flies are sensitive to dehydration.
The fed or fasted state prior to labeling, the choice of the label, the length of labeling time, the length of time in the fly pod, and the method of anesthesia are parameters that can be subjected to troubleshooting and modification when using this technique. First, flies subjected to short labeling periods (2 - 3 hr) are unlikely to consume significant quantities of radiolabeled food unless subjected to a fasting period beforehand. Second, the choice of label can affect the conclusions one is able to draw about metabolic phenotypes. For example, radiolabeled CO2 production from D-[1-14C]-glucose can reflect oxidation through the citric acid cycle or the pentose phosphate shunt, whereas radiolabeled CO2 production from D-[6-14C]-glucose reflects oxidation through the citric acid cycle only12,15,16. Feeding flies with a radiolabeled tracer for an essential amino acid17,18 would be a good strategy for assessing oxidation of amino acids derived from proteins. Finally, lengthy labeling periods with radiolabeled glucose also raise the possibility of incorporation of this precursor into other classes of molecules such as triglycerides19 and amino acids such as alanine, which readily interconverts with pyruvate. This can be assessed by measuring radioactivity incorporated into these different classes of molecules6. Indeed, separate cohorts of flies can be fed radiolabeled substrates in the same manner and then used for fly pod experiments or measurement of radiolabel that is stored and remains in glycogen and triglycerides in fed and fasted conditions6, for example. Another strategy is to use short-labeling periods that are likely to reveal oxidation of the radiolabeled dietary nutrients themselves and not storage forms of these nutrients. Third, it is also possible that two groups of flies could have identical rates of 14CO2 production over a given amount of time, but very different rates initially. Therefore, varying the amount of time that flies spend in the fly pod may be an important parameter to alter when assessing phenotypic variation. Finally, this protocol calls for the use of a short period of CO2 anesthesia when putting flies into the fly pod apparatus. It is possible that the CO2 anesthesia may affect metabolic function over the course of the fly pod experiment. An alternate approach is to use nitrogen anesthesia which does not affect metabolic rate 20.
As with any technique that measures such a complex system as fuel oxidation and metabolic rate, the method described here has limitations. First, it is possible that flies in different groups could consume different amounts of radiolabeled food and thus would enter the CO2 collection phase of the assay with different starting amounts of substrate. This can be controlled for to some extent by performing the experiment with large sample sizes and by feeding flies en masse. For short labeling periods, flies with similar blue-colored guts will have consumed the radiolabel and can be carried forward to the chase portion of the experiment. The amount of radiolabel remaining in a group of flies can also be measured at the end of the experiment and in separate cohorts of flies after the end of the feeding and initial chase period. Second, the activity levels between groups of flies in the fly pods may differ. This will have to be determined experimentally and taken into account when interpreting data. Third, this technique does not measure total CO2 production, only production from a specific dietary nutrient or the storage form(s) of that nutrient. This protocol is not a replacement for but instead is complementary to measurements of VCO2 because it provides different and additional information.
The method described here measures exhaled, radiolabeled CO2 that is a product of the oxidation of trace amounts of specific metabolic substrates. By varying the label used - glucose, fatty acid, or amino acid - one can design increasingly sophisticated experiments to assess the contributions of different substrates to metabolism under different physiological conditions such as starvation and on different genetic backgrounds. Future applications of this technique include measurement of metabolism in the larval and pupal stages of development, two stages of the Drosophila life cycle that are characterized by extremes of nutrient storage and breakdown, respectively.

Work in the model organism Drosophila melanogaster has contributed greatly to the understanding of genetic principles, developmental processes, growth, aging, behavior, immunity and human disease1,2. A myriad of genetic and cell biological approaches in Drosophila has driven progress in these areas. However, the study of metabolism in the fruit fly has developed more slowly, due in large part to difficulties in measuring metabolic parameters in such a small animal. The interest in using Drosophila as a model for studying human diseases such as diabetes and for understanding the contributions of metabolism to growth and various pathologies has pushed the field to develop and adapt metabolic techniques for this organism3,4.
Reliable methods are now available for the measurement of a number of metabolic parameters in the fruit fly. For example, it is straightforward to assess food intake5, levels of stored triglycerides and glycogen, as well as circulating hemolymph levels of glucose and the major circulating sugar in the fly, trehalose, which is a disaccharide composed of two molecules of glucose4. Use of metabolic tracers has enabled the study of nutrient absorption from the diet and the conversion of absorbed nutrients such as glucose to the storage forms glycogen and triglycerides6. Metabolic rates can be assessed in the fruit fly through the measurement of oxygen consumption7,8 and carbon dioxide (CO2) production. The citric acid cycle oxidizes two-carbon units that can enter the cycle as acetyl coenzyme A (CoA), which is derived from metabolism of dietary and stored carbohydrates and fatty acids. Each turn of the cycle generates one molecule each of GTP (or ATP) and the electron donor FADH2, three molecules of NADH, and two molecules of the waste product CO2. CO2 production can be used to estimate basal metabolic rate. Total CO2 production can be quantified in Drosophila through the use of commercially-available or handmade respirometers9-10,11.
The measurement of total CO2 production, especially when coupled with measurement of O2 consumption, provides valuable insight into whole-body energy metabolism. However, this measure does not identify the nutrient being oxidized for adenosine triphosphate (ATP) production. Three major classes of nutrients can enter the citric acid cycle following conversion to acetyl CoA: carbohydrates, fatty acids, and proteins. Glucose-6-phosphate, derived from dietary glucose or stored glycogen, can be converted to pyruvate that is decarboxylated by pyruvate dehydrogenase to form acetyl CoA. Breakdown of fatty acids derived from the diet or from stored triglycerides by &#223;-oxidation yields acetyl CoA that then enters the citric acid cycle. Finally, a majority of amino acids can enter the citric acid cycle following conversion to pyruvate, acetyl CoA or citric acid cycle intermediates such as alpha-ketoglutarate.
Nutrient-specific contribution to energy metabolism during basal and stimulated conditions can be monitored through the use of radioactive tracers. The protocol presented here is adapted for use in Drosophila from a protocol used to assess oxidation in cells grown in culture12,13. In this approach, fruit flies are fed 14C-radiolabeled metabolic substrates (carbohydrates, fatty acids, or amino acids) in the diet for a short (hours) or long (days) pulse, chased on unlabeled food, and then exposed to a chamber containing potassium hydroxide (KOH)-saturated filter paper for trapping exhaled CO2 as bicarbonate. Radiolabeled&#160;bicarbonate can be quantified by scintillation counting. Differences in radiolabeled-CO2 production between experimental groups of flies may reflect use of different fuels for ATP production over the course of a prolonged fast or intrinsic differences in fuel metabolism between genotypes, for example.

<table><tbody><tr><td>PALMITIC ACID, [1-14C]-, 50 &amp;micro;Ci</td><td>PerkinElmer</td><td>NEC075H050UC</td><td></td></tr><tr><td>GLUCOSE, D-[6-14C]-, 50 &amp;micro;Ci</td><td>PerkinElmer</td><td>NEC045X050UC</td><td></td></tr><tr><td>Glucose, D-[1-14C]-, 50&amp;micro;Ci</td><td>PerkinElmer</td><td>NEC043X050UC</td><td></td></tr><tr><td>Drosophila Vials, Narrow, Polystyrene</td><td>Genesee Scientific</td><td>32-116</td><td></td></tr><tr><td>Round-Bottom Polypropylene Tubes, 12 x 75 mm</td><td>Fisher Scientific</td><td>14-959AA</td><td></td></tr><tr><td>Mesh, nitex nylon, 120 &amp;micro;m</td><td>Genesee Scientific</td><td>57-102</td><td></td></tr><tr><td>20 ml borosilicate glass scintillation vials</td><td>Fisher Scientific</td><td>03-337-15</td><td></td></tr><tr><td>Flask top stopper with off-center hole</td><td>Fisher Scientific</td><td>K882310-0000</td><td></td></tr><tr><td>Polypropylene center well</td><td>Fisher Scientific</td><td>K882320-0000</td><td></td></tr><tr><td>Whatman GF/B glass microfiber filter paper, circle, 2.1cm diameter</td><td>Fisher Scientific</td><td>09-874-20</td><td></td></tr><tr><td>Ecoscint A</td><td>National Diagnostics</td><td>LS-273</td><td></td></tr><tr><td>6 ml Scintillation Vials with Push-On/Twist-Off Caps</td><td>National Diagnostics</td><td>SVC-06</td><td></td></tr></tbody></table>

measurement of carbon dioxide production from radiolabeled substrates in drosophila melanogaster

The power of Drosophila genetics is increasingly being applied to questions of hormone signaling and metabolism and to the development of models of human disease in this organism. Sensitive methods for measurements of parameters such as metabolic rates are needed to drive the understanding of physiology and disease in small animals such as the fruit fly. The method described here assesses fuel oxidation in small numbers of adult flies fed food containing trace amounts of 14C-labeled substrates such as glucose or fatty acid. After the feeding period and any additional experimental manipulations, flies are transferred to short tubes capped with mesh, which are then placed in glass vials containing KOH-saturated filter paper that traps exhaled, radiolabeled CO2 generated from oxidation of radiolabeled substrates as potassium bicarbonate, KHCO3. This radiolabeled bicarbonate is measured by scintillation counting. This is a quantitative, reproducible, and simple approach for the study of fuel oxidation. The use of radiolabeled glucose, fatty acids, or amino acids allows determination of the contribution of these different fuel sources to energy metabolism under different conditions such as feeding and fasting and in different genetic backgrounds. This complements other approaches used to measure in vivo energy metabolism and should further the understanding of metabolic regulation.

The quantity of exhaled and trapped radiolabeled CO2 that is produced from the metabolism of a radioactive substrate should correlate with the number of flies in a fly pod as well as the amount of time the animals spend in the CO2 collection apparatus. To test this, flies that were fasted for 18 hr were fed radiolabeled palmitic acid (0.02 &#181;Ci/&#181;l food) for 2 hr and then tested for radiolabeled CO2 production in groups of 12 or 25 animals for 3 or 6 hr incubations. The amount of CO2 produced by 25 flies was nearly double the amount produced by 12 flies, and the amount for each group doubled when the incubation time was increased from 3 hr to 6 hr (Figure 2A). The pmol 14CO2 / fly / hr was nearly equivalent in each group.
Fasting stimulates the breakdown of fatty acids by &#223; oxidation; therefore CO2 production from fatty acid oxidation should be elevated in fasted animals compared with fed animals. To determine whether this was the case, 18-hr fasted flies were fed radiolabeled palmitic acid for 2 hr. Following this short feeding pulse, flies were divided into two groups and transferred to unlabeled fly food or 1% agar for 3 hr and then transferred to fly pods for 2 hr. In two separate experiments, flies chased on agar produced significantly more radiolabeled CO2 compared with flies chased on food (Figure 2B), indicating that &#223; oxidation was increased in the fasted animals.
<img alt="Figure 1" src="/files/ftp_upload/54045/54045fig1.jpg" /> 
Figure 1.The Apparatus for Collecting Exhaled, Radiolabeled CO2. 
Flies that have consumed radiolabeled metabolic substrates are placed into a &#34;fly pod&#34; (FP), a cutoff centrifuge tube capped with mesh (M). The fly pod is placed into a 20 ml glass scintillation vial and capped with a rubber top stopper (S) containing an off-center hole, through which is placed a center well (CW) that contains GF/B filter paper saturated with 100 &#181;l 5% KOH. Exhaled CO2 reacts with the KOH and is trapped on the filter paper as bicarbonate. Trapped, radiolabeled CO2 can be quantified by scintillation counting of the saturated filter paper.&#160; <a href="https://www.jove.com/files/ftp_upload/54045/54045fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" src="/files/ftp_upload/54045/54045fig2.jpg" /> 
Figure 2. 14CO2 Production Correlates with Time and the Number of Flies in a Fly Pod and Responds to Fed and Fasted Conditions. 
(A) The amount of 14CO2 produced in flies fed radiolabeled palmitate correlates with the number of flies tested (n = 12 (open bars) or 25 (closed bars)) and the time spent in a fly pod (3 or 6 hr). On a per-fly basis, flies produced 0.99, 1.13, 1.10 and 1.01 pmol 14CO2 / hr (data listed in the same left to right order as graph data). 
(B) Flies that were chased on agar oxidized more radiolabeled palmitate than flies that were chased on food. Data are from two experiments and are reported as means standard deviation, n = 16 - 17 flies/group (experiment 1); n = 10 flies/group (experiment 2). * p = 0.017, Student&#39;s t-test. <a href="https://www.jove.com/files/ftp_upload/54045/54045fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

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Measurement of Carbon Dioxide Production from Radiolabeled Substrates in Drosophila melanogaster

This paper describes a method for the measurement of fuel oxidation in Drosophila melanogaster in which trace amounts of specific radiolabeled metabolic substrates are fed to flies. The exhaled radiolabeled CO2 that is a produced from fuel oxidation is collected and measured.

Measurement of Carbon Dioxide Production from Radiolabeled Substrates in Drosophila melanogaster

The power of Drosophila genetics is increasingly being applied to questions of hormone signaling and metabolism and to the development of models of human ...

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JoVE Journal

Biology

9.1K Views.  University of Virginia. This paper describes a method for the measurement of fuel oxidation in Drosophila melanogaster in which trace amounts of specific radiolabeled metabolic substrates are fed to flies. The exhaled radiolabeled CO2 that is a produced from fuel oxidation is collected and measured.

Video: Measurement of Carbon Dioxide Production from Radiolabeled Substrates in Drosophila melanogaster

Dissection of Larval CNS in Drosophila Melanogaster

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

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

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

Measurement of Mitochondrial Oxygen Consumption in Permeabilized Fibers of Drosophila Using Minimal Amounts of Tissue

The fruit fly, Drosophila melanogaster, represents an emerging model for the study of metabolism. Indeed, drosophila have structures homologous to human organs, possess highly conserved metabolic pathways and have a relatively short lifespan that allows the study of different fundamental mechanisms in a short period of time. It is, however, surprising that one of the mechanisms essential for cellular metabolism, the mitochondrial respiration, has not been thoroughly investigated in this model. It is likely because the measure of the mitochondrial respiration in Drosophila usually requires a very large number of individuals and the results obtained are not highly reproducible. Here, a method allowing the precise measurement of mitochondrial oxygen consumption using minimal amounts of tissue from Drosophila is described. In this method, the thoraxes are dissected and permeabilized both mechanically with sharp forceps and chemically with saponin, allowing different compounds to cross the cell membrane and modulate the mitochondrial respiration. After permeabilization, a protocol is performed to evaluate the capacity of the different complexes of the electron transport system (ETS) to oxidize different substrates, as well as their response to an uncoupler and to several inhibitors. This method presents many advantages compared to methods using mitochondrial isolations, as it is more physiologically relevant because the mitochondria are still interacting with the other cellular components and the mitochondrial morphology is conserved. Moreover, sample preparations are faster, and the results obtained are highly reproducible. By combining the advantages of Drosophila as a model for the study of metabolism with the evaluation of mitochondrial respiration, important new insights can be unveiled, especially when the flies are experiencing different environmental or pathophysiological conditions.

In this paper, a method to measure oxygen consumption using high-resolution respirometry in permeabilized thoraxes of Drosophila is described. This technique requires a minimal amount of tissue compared to the classic mitochondrial isolation technique and the results obtained are more physiologically relevant.

The fruit fly, Drosophila melanogaster, represents an emerging model for the study of metabolism. Indeed, drosophila have structures homologous to human ...

Biochemistry

High-Resolution Video Tracking of Locomotion in Adult Drosophila Melanogaster

Flies provide an important model for studying complex behavior due to the plethora of genetic tools available to researchers in this field. Studying locomotor behavior in Drosophila melanogaster relies on the ability to be able to quantify changes in motion during or in response to a given task. For this reason, a high-resolution video tracking system, such as the one we describe in this paper, is a valuable tool for measuring locomotion in real-time. Our protocol involves the use of an initial air pulse to break the flies momentum, followed by a thirty second filming period in a square chamber. A tracking program is then used to calculate the instantaneous speed of each fly within the chamber in 10 msec increments. Analysis software then compiles this data, and outputs a variety of parameters such as average speed, max speed, time spent in motion, acceleration, etc. This protocol will discuss proper feeding and management of flies for behavioral tasks, handling flies without anesthetization or immobilization, setting up a controlled environment, and running the assay from start to finish.

The study of complex locomotor behavior in Drosophila melanogaster is dependent upon the ability to quantify changes in a given fly's movement. This article demonstrates how to do this using a high-resolution tracking system.

Flies provide an important model for studying complex behavior due to the plethora of genetic tools available to researchers in this field.  Studying ...

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  ...

Live Imaging Of Drosophila melanogaster Embryonic Hemocyte Migrations

Many studies address cell migration using in vitro methods, whereas the physiologically relevant environment is that of the organism itself. Here we present a protocol for the mounting of Drosophila melanogaster embryos and subsequent live imaging of fluorescently labeled hemocytes, the embryonic macrophages of this organism. Using the Gal4-uas system1 we drive the expression of a variety of genetically encoded, fluorescently tagged markers in hemocytes to follow their developmental dispersal throughout the embryo. Following collection of embryos at the desired stage of development, the outer chorion is removed and the embryos are then mounted in halocarbon oil between a hydrophobic, gas-permeable membrane and a glass coverslip for live imaging. In addition to gross migratory parameters such as speed and directionality, higher resolution imaging coupled with the use of fluorescent reporters of F-actin and microtubules can provide more detailed information concerning the dynamics of these cytoskeletal components.

Live Imaging Of Drosophila melanogaster Embryonic Hemocyte Migrations

Drosophila hemocytes disperse over the entirety of the developing embryo. This protocol demonstrates how to mount and image these migrations using embryos with fluorescently labelled hemocytes.

Many studies address cell migration using in vitro methods, whereas the physiologically relevant environment is that of the organism itself. Here we ...

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 ...

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

Measurement of Metabolic Rate in Drosophila using Respirometry

Metabolic disorders are a frequent problem affecting human health. Therefore, understanding the mechanisms that regulate metabolism is a crucial scientific task. Many disease causing genes in humans have a fly homologue, making Drosophila a good model to study signaling pathways involved in the development of different disorders. Additionally, the tractability of Drosophila simplifies genetic screens to aid in identifying novel therapeutic targets that may regulate metabolism. In order to perform such a screen a simple and fast method to identify changes in the metabolic state of flies is necessary. In general, carbon dioxide production is a good indicator of substrate oxidation and energy expenditure providing information about metabolic state. In this protocol we introduce a simple method to measure CO2 output from flies. This technique can potentially aid in the identification of genetic perturbations affecting metabolic rate.

Measurement of Metabolic Rate in Drosophila using Respirometry

Metabolic disorders are among one of the most common diseases in humans. The genetically tractable model organism D. melanogaster can be used to identify novel genes that regulate metabolism. This paper describes a relatively simple method which allows studying the metabolic rate in flies by measuring their CO2 production.

Metabolic disorders are a frequent problem affecting human health. Therefore, understanding the mechanisms that regulate metabolism is a crucial ...

Adapting Microrespirometry for Drosophila - Insights into CASK Mutant Metabolism

Measuring O2 Consumption in Drosophila melanogaster Using Coulometric Microrespirometry

Coulometric microrespirometry is a straightforward, inexpensive method for measuring the O2 consumption of small organisms while maintaining a stable environment. A coulometric microrespirometer consists of an airtight chamber in which O2 is consumed and the CO2 produced by the organism is removed by an absorbent medium. The resulting pressure decrease triggers electrolytic O2 production, and the amount of O2 produced is measured by recording the amount of charge used to generate it. In the present study, the method has been adapted to Drosophila melanogaster tested in small groups, with the sensitivity of the apparatus and the environmental conditions optimized for high stability. The amount of O2 consumed by wildtype flies in this apparatus is consistent with that measured by previous studies. Mass-specific O2 consumption by CASK mutants, which are smaller and known to be less active, was not different from congenic controls. However, the small size of CASK mutants resulted in a significant reduction in O2 consumption on a per-fly basis. Therefore, the microrespirometer is capable of measuring O2 consumption in D. melanogaster, can distinguish modest differences between genotypes, and adds a versatile tool for measuring metabolic rates.

Author Spotlight: Influence of Temperature on Drosophila melanogaster and Desert-Adapted Beetles 

Coulometric respirometry is ideal for measuring the metabolic rate of small organisms. When adapted for Drosophila melanogaster in the present study, measured O2 consumption was within the range reported for wildtype D. melanogaster by previous studies. Per-fly O2 consumption by CASK mutants, which are smaller and less active, was significantly lower than the wildtype.

Coulometric microrespirometry is a straightforward, inexpensive method for measuring the O2 consumption of small organisms while maintaining a stable ...

Measurement of Carbon Dioxide Production from Radiolabeled Substrates in Drosophila melanogaster

Summary

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Chapters in this video

Dissection of Larval CNS in Drosophila Melanogaster

Measurement of Mitochondrial Oxygen Consumption in Permeabilized Fibers of Drosophila Using Minimal Amounts of Tissue

High-Resolution Video Tracking of Locomotion in Adult Drosophila Melanogaster

Monitoring Heart Function in Larval Drosophila melanogaster for Physiological Studies

Live Imaging Of Drosophila melanogaster Embryonic Hemocyte Migrations

Dissection of Oenocytes from Adult Drosophila melanogaster

Isolation of Drosophila melanogaster Testes

Metabolic Analysis of Drosophila melanogaster Larval and Adult Brains

Measurement of Metabolic Rate in Drosophila using Respirometry

Measuring O₂ Consumption in Drosophila melanogaster Using Coulometric Microrespirometry