This study was funded by grants from the National Sciences and Engineering Research Council (NSERC, discovery grant) and Universit&#233; de Moncton to NP. LHB would like to acknowledge the funding support from the Canadian Institute of Health Research (CIHR), the New Brunswick Innovation Foundation (NBIF) and Universit&#233; de Moncton. The work of EHC is supported by the Alzheimer Society of Canada, Brain Canada, NSERC, Canadian Breast Cancer Foundation, New Brunswick Innovation Foundation, New Brunswick Health Research Foundation and Universit&#233; de Moncton.

1. Reagents Preparation
<ol>
	<li>Prepare the following solutions for dissection and permeabilization of the tissue.
	<ol>
		<li>Prepare preservation solution: 2.77 mM CaK2EGTA, 7.23 mM K2EGTA, 5.77 mM Na2ATP, 6.56 mM MgCl2, 20 mM taurine, 15 mM Na2phosphocreatine, 20 mM imidazole, 0.5 mM dithiothreitol, and 50 mM K-MES, pH 7.1 (can be stored at -20 &#176;C).</li>
		<li>Prepare saponin solution: 5 mg of saponin in 1 mL of preservation solution (prepare fresh daily).</li>
	</ol>
	</li>
	<li>Prepare the following solutions for the measurement of the respiration.
	<ol>
		<li>Prepare respiration medium: 120 mM KCl, 5 mM KH2PO4, 3 mM HEPES, 1 mM EGTA, 1 mM MgCl2, and 0.2% BSA (w/v), pH 7.2.</li>
		<li>Prepare substrates, uncoupler, inhibitors. Dissolve all substrates in H2O, and dissolve the uncoupler FCCP as well as the inhibitors in ethanol absolute (except malonate which is diluted in H2O). Neutralize most of the substrates as well as malonate (see Table of Materials). All concentrations given in the protocol are final concentrations inside 2 mL chambers.</li>
	</ol>
	</li>
</ol>
2. Preparation and Calibration of the High-Resolution Respirometer.
<ol>
	<li>Wash the chambers, caps, and stoppers of the respirometer three times with 70% ethanol and three times with distilled water.</li>
	<li>Calibrate at zero oxygen concentration: pipet 2.3 mL of respiration medium in the chambers and add 10 mg of sodium dithionite. Although the maximal volume of the chamber is 2 mL, 2.3 mL are added to ensure that no air bubble stays under the stopper when closing the chambers. Close the chambers with the stoppers and measure the oxygen concentration for 10-20 min. Calibrate the electrodes at zero oxygen concentration with the resulting oxygen consumption rate.</li>
	<li>Wash the chambers with distillated water, incubate in absolute ethanol for 10 min, and then rinse with distilled water.</li>
	<li>Calibrate at air saturation: pipet 2.3 mL of respiration medium in the chambers, insert the stoppers, aspirate the excess medium on top of the stoppers, and then lift the stoppers with the spacer. Excess respiration medium is pipetted in the chambers (volume of 2 mL total) to avoid the presence of air bubbles after closing the chambers.</li>
	<li>Monitor the oxygen consumption for 45 min to an hour, until the oxygen concentration (blue trace) is stable around 250 nM at 24 &#176;C.</li>
</ol>
3. Dissection of Flies and Permeabilization of Tissue
<ol>
	<li>Make sure all steps are performed on ice.</li>
	<li>Anaesthetize six male flies of the same strain, same age and raised on the same diet on ice to facilitate dissection of the flies (wild-type w1118, 15 days old, fed on cornmeal medium).</li>
	<li>Using a scalpel and fine-tipped forceps, dissect the flies (remove the heads and abdomens) to keep only the thoraxes containing the flight muscles. This step can be done with the naked eye. Handle the resulting thoraxes with fine-tipped forceps.</li>
	<li>Transfer three dissected thoraxes in a 25 mm Petri dish containing 2 mL of ice-cold preservation solution using the forceps.</li>
	<li>With the fine-tipped forceps, mechanically permeabilize the thoraxes by inserting the tip of the forceps into the thoraxes and repeatedly tear apart the tissue to obtain a loosely connected network.</li>
	<li>In a 24-well plate, fill two wells with 12.5 &#181;L of the saponin solution and 1 mL of preservation solution to obtain a final concentration of 62.5 &#181;g/mL of saponin. Fill the two adjacent wells with 1 mL of respiration medium.</li>
	<li>With the fine-tipped forceps, transfer three permeabilized thoraxes into the diluted saponin solution and incubate with mild agitation on an orbital shaker, on ice, for 20 min.</li>
	<li>After the latter incubation, transfer the fibers into the adjacent wells filled with the respiration medium with mild agitation, on ice, for 5 min to rinse off the saponin.</li>
</ol>
4. Determination of Dry Weight
<ol>
	<li>Dry the permeabilized thoraxes on an absorbent surface and flip 3-4 times using the fine-tipped forceps to make sure all moisture is removed.</li>
	<li>Weigh the three thoraxes together on a microbalance. Note the obtained weight as it will be used to normalize the mitochondrial oxygen consumption rates.</li>
	<li>Transfer the tissues immediately in a drop of respiration medium on ice.</li>
</ol>
5. Oxygen Consumption Rates Determination
<ol>
	<li>Open the chambers of the respirometer (remove the stoppers) and add 10 mM of pyruvate and 2 mM of malate into each chamber.</li>
	<li>Add the permeabilized tissues directly into the chambers filled with the respiration medium.</li>
	<li>Replace the stoppers using the spacer.</li>
	<li>With a 60 mL syringe, collect oxygen directly from an oxygen tank and inject 2-5 mL of oxygen through the stopper capillary of each chamber to make sure oxygen diffusion through the tissue will not limit oxygen consumption.</li>
	<li>Insert a Hamilton syringe into the capillary of each stopper to make sure the thoraxes remain in the chamber.</li>
	<li>Close the chambers completely when the oxygen concentration is around 400 nM, achieving oxygen levels above air saturation (around 160% air saturation) inside the chambers.</li>
	<li>Stop and restart the stirrers to check for air bubbles in the chambers.</li>
	<li>Enter the mass of tissue weighed previously to normalize respiration rates by mass of tissue (mg), displaying mass-specific oxygen consumption (pmol O2 consumed.s-1.mg-1 of tissue).</li>
	<li>Once the oxygen consumption is stabilized (red trace), add 5 mM ADP with a Hamilton syringe.</li>
	<li>After each injection, rinse the syringes with distilled water, 70% ethanol, and distilled water again.</li>
	<li>Once the oxygen consumption is stable again, inject 5 &#181;L of a 4 mM of cytochrome c.</li>
	<li>Proceed to the sequential injections of the following substrates to evaluate the mitochondrial oxygen consumption at different steps of the ETS:
	<ol>
		<li>Add 5 &#181;L of a 2 M Proline.</li>
		<li>Add 10 &#181;L of a 1 M succinate.</li>
		<li>Add 30 &#181;L of a 1 M glycerol-3-phosphate.</li>
	</ol>
	</li>
	<li>Inject the uncoupler FCCP in a titration manner by steps of 0.5-1 &#181;M (2 &#181;L of a 1 mM solution) until the optimal concentration is reached, i.e., the concentration to attain the highest oxygen consumption possible without inhibition (CAUTION: see Table of Materials).</li>
	<li>Inject the following inhibitors sequentially to entirely inhibit the electron flux in the ETS in order to measure the residual oxygen consumption (ROX) i.e. non-respiratory reactions such as oxygenase reactions among others:
	<ol>
		<li>Add 1 &#181;L of a 1 mM rotenone (CAUTION: see Table of Materials).</li>
		<li>Add 5 &#181;L of a 2 M malonate (CAUTION: see Table of Materials).</li>
		<li>Add 1 &#181;L of a 5 mM antimycin A (CAUTION: see Table of Materials).</li>
	</ol>
	</li>
	<li>Add 0.2 mM ascorbate and 0.5 mM TMPD sequentially into the chambers, starting with the ascorbate to measure the oxygen consumption by complex IV.</li>
	<li>Add 20 mM of sodium azide to inhibit complex IV (CAUTION: see Table of Materials).</li>
	<li>Once the signal is stable, open the chambers with the spacer, re-oxygenate the chambers by injecting 2 mL of pure oxygen and close the chambers when the concentration is around 250-300 nmol.mL-1.</li>
	<li>Measure the signal for 10-15 min and calculate the chemical background, i.e., the oxygen consumption due to autoxidation of TMPD.</li>
</ol>
6. Cleaning of the Respirometer
<ol>
	<li>After the measurements, rinse the chambers with distilled water once and rinse the chambers three times in 70% ethanol.</li>
	<li>Incubate in absolute ethanol for 10 min to inactivate the inhibitors.</li>
	<li>Rinse three times with distilled water, fill the chambers with 70% ethanol and replace the stoppers and the caps until next utilization.</li>
</ol>

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<li>McCommis, K., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((An+ancestral+role+for+the+mitochondrial+pyruvate+carrier+in+glucose-stimulated+insulin+secretion%5BTitle%5D)+AND+%22Molecular+Metabolism%22%5BJournal%5D)">An ancestral role for the mitochondrial pyruvate carrier in glucose-stimulated insulin secretion.</a> Molecular Metabolism. 5 (8), 602-614 (2016).</li>
</ol>

The authors have nothing to disclose.

In this study, a method for sample preparation prior to the measurements of mitochondrial oxygen consumption in Drosophila is described. This method was developed to overcome different problems related to the protocols using mitochondrial isolations, notably in terms of duration and number of individuals required. Instead of working with mitochondrial isolations usually requiring large amount of tissues obtained from several individuals, this experiment is performed on permeabilized muscle fibers from thoraxes of few Dro...

The fruit fly, Drosophila melanogaster, has been used as a model organism for genetic research for over a century1. The study of this organism has not only led to significant fundamental knowledge about sex-linked inheritance2, mutation rate3, the development of neural system and the cell fate determination4, but has also recently emerged as a valuable tool to study the mechanisms inherent to several diseases such as Alzheimer&#39;s and Parkinson&#39;s5,6. Moreover, it is a popular model to study the aging process, as they can be raised in large number over a short period of time and have a short lifespan. They also possess homologous structures to human organs, such as a heart, oenocytes (hepatocyte-like cells), fat bodies (functioning similarly as the liver and white adipose tissue), insulin producing cells (equivalent to &#946;-pancreatic cells), as well as the hemolymph transporting metabolites (analogous to the blood of vertebrates)7. Moreover, the central pathways of intermediary metabolism (including the insulin/insulin-like growth factor-like signaling pathway and Target of Rapamycin-TOR pathways) are also highly conserved7. For these reasons, Drosophila have recently been exploited to describe the fundamental mechanisms that control metabolism, especially in pathological conditions inherent to human metabolic diseases such as diabetes8. A major component of the metabolism is the mitochondrion that integrates multiple pathways and performs one of life&#39;s most important biological functions, ATP production, via the oxidative phosphorylation process (OXPHOS). Considering their central role in organism&#39;s metabolism, it is not surprising that mitochondrial dysfunctions are involved in many diseases like Parkinson&#39;s9 and Alzheimer&#39;s diseases10, as well as in amyotrophic lateral sclerosis11,12. They are also fundamental determinants of the aging process. Indeed, they are the main producers of reactive oxygen species (ROS) in the cell, which can be detrimental to the cell at high concentration through oxidative damages11. Aging has also been associated to the accumulation of damaged or mutated mitochondrial DNA13, mitophagy dysfunctions14,15 as well as impairment of mitochondrial biogenesis16. Mitochondria are also key determinants of the cell&#39;s homeostasis as they can utilize different substrates to adjust several cellular functions according to the abundance or scarcity of macronutrients17,18.
Indeed, the different nutrients in the diet (carbohydrates, lipids and proteins) are digested, absorbed and transported in the cells. They are then transformed in the cytosol, and the derived substrates are transported into the mitochondrial matrix where they produce reducing equivalents, such as NADH and FADH219. These reducing equivalents are then oxidized by different enzymatic complexes of the electron transport system (ETS). These complexes are embedded in the mitochondrial inner membrane, such as Complex I and Complex II. Additionally, other enzymatic complexes such as the mitochondrial glycerol-3-phosphate dehydrogenase and the proline dehydrogenase represent alternate routes for the entry of electrons into the ETS20,21. These &#39;alternative&#39; complexes are particularly important in insects, as according to the species, they can actively participate to increase the respiration20,22,23,21. Electrons from these ETS feeding systems are transferred to the ubiquinone and subsequently to Complex III, and then to Complex IV, until the final acceptor, molecular oxygen. This electron transfer generates a proton-motive force across the inner mitochondrial membrane driving the phosphorylation of ADP to ATP at Complex V (Figure 1). Considering the central role of mitochondria in cell&#39;s homeostasis, studying mitochondrial metabolism using the relevant model D. melanogaster represents a powerful tool to delineate the underlying mechanisms of various pathophysiological conditions or under cellular and environmental stresses. Surprisingly however, only a handful of studies actually measured mitochondrial respiration in Drosophila24,25,26. Indeed, experiments aiming to evaluate mitochondrial oxygen consumption require the isolation of mitochondria. Although advantageous for the measurement of different mitochondrial functions (such as ROS production or P/O ratio as marker of mitochondrial efficiency27,28), these isolations generally require rather large quantities of tissue from several individuals24,29. This requirement for high amounts of tissue and individuals is an important limiting factor, especially considering that all individuals should be the same age and preferably of the same sex for the experiments, making the measure of respiration at different time points laborious at best. Moreover, while mitochondrial isolations can provide significant insight into the fundamental mechanisms governing mitochondrial metabolism, the methods used to isolate mitochondria have several drawbacks such as the difficulty to obtain replicable results, disruption of the mitochondrial network, and alteration of mitochondrial structure and function29,30,31.
The aim of this study is to present a robust protocol to measure mitochondrial oxygen consumption in Drosophila using only a minimal amount of tissue from very few individuals. This protocol consists of measuring mitochondrial oxygen consumption in situ using permeabilized muscle fibers29 from Drosophila thoraxes in combination with high-resolution respirometry32,33,34,35. This method has also additional advantages compared to the classic mitochondrial isolation method since the interactions with the other components of the cell as well as mitochondrial structure and function are more preserved in permeabilized fibers29,31,36, which makes this approach more physiologically relevant. With this protocol, mitochondrial functions can be accurately evaluated using high-resolution respirometry in only three thoraxes of Drosophila, with substrates allowing the determination of oxygen consumption at several different steps of the ETS. Therefore, this protocol could help answer key questions about the fundamental mechanisms that control metabolism in the context of many environmental or pathophysiological conditions by taking advantage of the Drosophila model.
To measure the oxygen consumption at several different steps of the ETS and evaluate how different substrates contribute to the respiration, different substrates (Figure 1), uncoupler, and inhibitors are used30 after permeabilization of the tissue. Specifically, sequential additions of different substrates are performed to stimulate the entry of electrons through different complexes of the ETS. An uncoupler, carbonyl cyanide 4-(trifluoromethoxy)phenylhydrazone (FCCP), is then added at optimal concentration to measure the non-coupled respiration, i.e., the non-phosphorylating respiration stimulated to maximal oxygen consumption. Sequential inhibitions of complexes I, II, and III are then performed to monitor the residual oxygen consumption that is due to non-ETS oxidation reactions. Finally, complex IV maximal respiration capacity can be evaluated by the injection of N,N,N&#39;,N,-Tetramethyl-p-phenylenediamine (TMPD), an artificial electron provider, and ascorbate. It is important to note that the experiments are conducted at 24 &#176;C since it is the temperature at which the flies are raised.

<table><tbody><tr><td>High-resolution respirometer Oxygraph O2K</td><td>Oroboros Instruments, Innsbruck, Austria</td><td>10022-02</td><td>Startup O2K respirometer kit&lt;br/&gt; &amp;nbsp;</td></tr><tr><td>O2K-Titration Set&amp;nbsp;</td><td>Oroboros Instruments, Innsbruck, Austria</td><td>20820-03</td><td>Hamilton syringes with different volumes&lt;br/&gt; &amp;nbsp;</td></tr><tr><td>Datlab software</td><td>Oroboros Instruments, Innsbruck, Austria</td><td>20700</td><td>Software for data acquisition and analysis&lt;br/&gt; &amp;nbsp;</td></tr><tr><td>Fine-tipped antimagnetic forceps</td><td>VWR</td><td>82027-400</td><td>&lt;br/&gt; &amp;nbsp;</td></tr><tr><td>Secura225D-1S-DQE</td><td>Sartorius AG, Goettingen, Germany</td><td></td><td>Semi-micro balance (distributed by several companies)&amp;nbsp;&lt;br/&gt; &amp;nbsp;</td></tr><tr><td>&lt;em&gt;Drosophila melanogaster&lt;/em&gt; wild-type w&lt;sup&gt;1118&lt;/sup&gt;</td><td>Bloomington Drosophila stock Center, IN, USA</td><td></td><td>&lt;br/&gt; Storage Condition: 24 &amp;deg;C</td></tr><tr><td>Ethylene glycol-bis(2-aminoethylether)-&lt;em&gt;N&lt;/em&gt;,&lt;em&gt;N&lt;/em&gt;,&lt;em&gt;N&lt;/em&gt;&amp;prime;,&lt;em&gt;N&lt;/em&gt;&amp;prime;-tetraacetic acid</td><td>Sigma-Aldrich</td><td>E4378</td><td>EGTA&lt;br/&gt; Storage Condition: RT</td></tr><tr><td>KOH</td><td>Sigma-Aldrich</td><td>P1767</td><td>CAUTION: corrosive to metals, acute toxicity, skin corrosion, serious eye damage, acute aquatic toxicity.&lt;br/&gt; Storage Condition: RT</td></tr><tr><td>CaCO&lt;sub&gt;3&lt;/sub&gt;</td><td>Sigma-Aldrich</td><td>C4830</td><td>&lt;br/&gt; Storage Condition: RT</td></tr><tr><td>Na&lt;sub&gt;2&lt;/sub&gt;ATP</td><td>Sigma-Aldrich</td><td>A2383</td><td>&lt;br/&gt; Storage Condition: -20 &amp;deg;C</td></tr><tr><td>MgCl&lt;sub&gt;2&lt;/sub&gt;.6H&lt;sub&gt;2&lt;/sub&gt;O</td><td>Sigma-Aldrich</td><td>M9272</td><td>&lt;br/&gt; Storage Condition: RT</td></tr><tr><td>Taurine</td><td>Sigma-Aldrich</td><td>T0625</td><td>&lt;br/&gt; Storage Condition: RT</td></tr><tr><td>Na&lt;sub&gt;2&lt;/sub&gt;Phosphocreatine</td><td>Sigma-Aldrich</td><td>P7936</td><td>&lt;br/&gt; Storage Condition: -20 &amp;deg;C</td></tr><tr><td>Imidazole</td><td>Sigma-Aldrich</td><td>I5513</td><td>&lt;br/&gt; Storage Condition: RT</td></tr><tr><td>Dithiothreitol</td><td>Sigma-Aldrich</td><td>D0632</td><td>&lt;br/&gt; Storage Condition: 2-8 &amp;deg;C</td></tr><tr><td>MES hydrate</td><td>Sigma-Aldrich</td><td>M8250</td><td>&lt;br/&gt; Storage Condition: RT</td></tr><tr><td>Saponin from quillaja bark</td><td>Sigma-Aldrich</td><td>S7900</td><td>Saponin&lt;br/&gt; Storage Condition: RT&lt;br/&gt; Solution Preparation: 5 mg in 1 mL of preservation solution. Prepare fresh daily.</td></tr><tr><td>KCl</td><td>Sigma-Aldrich</td><td>P9541</td><td>&lt;br/&gt; Storage Condition: RT</td></tr><tr><td>KH&lt;sub&gt;2&lt;/sub&gt;PO&lt;sub&gt;4&lt;/sub&gt;</td><td>Sigma-Aldrich</td><td>P9791</td><td>&lt;br/&gt; Storage Condition: RT</td></tr><tr><td>HEPES</td><td>Sigma-Aldrich</td><td>H3375</td><td>&lt;br/&gt; Storage Condition: RT</td></tr><tr><td>BSA</td><td>Sigma-Aldrich</td><td>05470</td><td>&lt;br/&gt; Storage Condition: 2-8 &amp;deg;C</td></tr><tr><td>Na&lt;sub&gt;2&lt;/sub&gt;S&lt;sub&gt;2&lt;/sub&gt;O&lt;sub&gt;4&lt;/sub&gt;</td><td>Sigma-Aldrich</td><td>157953</td><td>Sodium dithionite. CAUTION: self-heating substances and mixtures, acute toxicity, acute aquatic toxi chronic aquatic toxicity.&lt;br/&gt; Storage Condition: RT</td></tr><tr><td>Sodium pyruvate</td><td>Sigma-Aldrich</td><td>P2256</td><td>Pyruvate&lt;br/&gt; Storage Condition: 2-8 &amp;deg;C&lt;br/&gt; Solution Preparation: In MilliQ water. Prepare fresh daily.</td></tr><tr><td>L-(-)-Malic acid</td><td>Sigma-Aldrich</td><td>M1000</td><td>Malate&lt;br/&gt; Storage Condition: RT&lt;br/&gt; Solution Preparation: In MilliQ water. Neutralize with KOH and store at -20 &amp;deg;C.</td></tr><tr><td>Adenosine 5&#039;-diphosphate monopotassium salt hydrate</td><td>Sigma-Aldrich</td><td>A5285</td><td>ADP&lt;br/&gt; Storage Condition: -20 &amp;deg;C&lt;br/&gt; Solution Preparation: In MilliQ water. Neutralize with KOH and store at -80 &amp;deg;C.</td></tr><tr><td>Cytochrome c from equine heart</td><td>Sigma-Aldrich</td><td>C7752</td><td>Cytochrome c&lt;br/&gt; Storage Condition: -20 &amp;deg;C&lt;br/&gt; Solution Preparation: In MilliQ water. Store at -20 &amp;deg;C.</td></tr><tr><td>L-Proline</td><td>Sigma-Aldrich</td><td>P0380</td><td>Proline&lt;br/&gt; Storage Condition: RT&lt;br/&gt; Solution Preparation: In MilliQ water. Store at -20 &amp;deg;C.</td></tr><tr><td>Sodium succinate dibasic hexahydrate</td><td>Sigma-Aldrich</td><td>S2378</td><td>Succinate&lt;br/&gt; Storage Condition: RT&lt;br/&gt; Solution Preparation: In MilliQ water. Neutralize with HCl and store at -20 &amp;deg;C.</td></tr><tr><td>sn-Glycerol 3-phosphate bis(cyclohexylammonium) salt</td><td>Sigma-Aldrich</td><td>G7886</td><td>Glycerol-3-phosphate&lt;br/&gt; Storage Condition: -20 &amp;deg;C&lt;br/&gt; Solution Preparation: In MilliQ water. Neutralize with HCl and store at -80 &amp;deg;C.</td></tr><tr><td>Carbonyl cyanide 4-(trifluoromethoxy)phenylhydrazone</td><td>Sigma-Aldrich</td><td>C2920</td><td>FCCP. CAUTION: acute toxicity, skin sensitisation, chronic aquatic toxicity.&lt;br/&gt; Storage Condition: RT&lt;br/&gt; Solution Preparation: In absolute ethanol. Store in glass vials at -20 &amp;deg;C.</td></tr><tr><td>Rotenone</td><td>Sigma-Aldrich</td><td>R8875</td><td>CAUTION: acute toxicity, skin irritation, eye irritation, specific target organ toxicity (respir sytem), acute aquatic toxicity, chronic aquatic toxicity.&lt;br/&gt; Solution Preparation: In absolute ethanol. Store in dark vials at -20 &amp;deg;C.</td></tr><tr><td>Malonic acid</td><td>Sigma-Aldrich</td><td>M1296</td><td>Malonate. CAUTION: acute toxicity, serious eye damage.&lt;br/&gt; Storage Condition: RT&lt;br/&gt; Solution Preparation: In MilliQ water. Neutralize with KOH. Prepare fresh daily.</td></tr><tr><td>Antimycin A from &lt;em&gt;Streptomyces sp&lt;/em&gt;.</td><td>Sigma-Aldrich</td><td>A8674</td><td>Antimycin A. CAUTION: acute toxicity, acute aquatic toxicity, chronic aquatic toxicity.&lt;br/&gt; Storage Condition: -20 &amp;deg;C&lt;br/&gt; Solution Preparation: In absolute ethanol. Store at -20 &amp;deg;C.</td></tr><tr><td>&lt;em&gt;N&lt;/em&gt;,&lt;em&gt;N&lt;/em&gt;,&lt;em&gt;N&lt;/em&gt;&amp;prime;,&lt;em&gt;N&lt;/em&gt;&amp;prime;-Tetramethyl-&lt;em&gt;p&lt;/em&gt;-phenylenediamine</td><td>Sigma-Aldrich</td><td>T7394</td><td>TMPD&lt;br/&gt; Storage Condition: RT&lt;br/&gt; Solution Preparation: In MilliQ water. Store in dark vials at&amp;nbsp; -20 &amp;deg;C.</td></tr><tr><td>(+)-Sodium L-ascorbate</td><td>Sigma-Aldrich</td><td>A4034</td><td>Ascorbate&lt;br/&gt; Storage Condition: RT&lt;br/&gt; Solution Preparation: In MilliQ water. Store in dark vials at&amp;nbsp; -20 &amp;deg;C.</td></tr><tr><td>NaN&lt;sub&gt;3&lt;/sub&gt;</td><td>Sigma-Aldrich</td><td>S2002</td><td>Sodium azide. CAUTION: acute toxicity (oral and dermal), specific target organ toxicity (brain), aquatic toxicity, chronic aquatic toxicity.&amp;nbsp;&lt;br/&gt; Solution Preparation: In MilliQ water. Store at -20 &amp;deg;C.</td></tr></tbody></table>

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.

A representative trace of mitochondrial oxygen consumption using the protocol described above is provided in Figure 2. The pyruvate and malate injected in the chambers along with the permeabilized muscle fibers are referred to the CI-LEAK respiration, i.e., when the complex I of the ETS is stimulated by the NADH produced through oxidation of pyruvate and malate via the tricarboxylic acid cycle (CI). During this respiration rate, the mitochondrial oxy...

Watch this Scientific Journal Video about Measurement of Mitochondrial Oxygen Consumption in Permeabilized Fibers of Drosophila Using Minimal Amounts of Tissue at JoVE.com

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

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 ...

measurement-mitochondrial-oxygen-consumption-permeabilized-fibers

Research

JoVE Journal

Biochemistry

11.9K Views.  Université de Moncton. In dit document wordt een methode voor het meten van zuurstofverbruik met behulp van hoge-resolutie respirometrie in permeabel thoraxes van Drosophila beschreven. Deze techniek vereist een minimale hoeveelheid weefsel in vergelijking met de klassieke mitochondriale isolatie-techniek en de verkregen resultaten meer fysiologisch relevant zijn.

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

Using Isolated Mitochondria from Minimal Quantities of Mouse Skeletal Muscle for High throughput Microplate Respiratory Measurements

Skeletal muscle mitochondria play a specific role in many disease pathologies. As such, the measurement of oxygen consumption as an indicator of mitochondrial function in this tissue has become more prevalent. Although many technologies and assays exist that measure mitochondrial respiratory pathways in a variety of cells, tissue and species, there is currently a void in the literature in regards to the compilation of these assays using isolated mitochondria from mouse skeletal muscle for use in microplate based technologies. Importantly, the use of microplate based respirometric assays is growing among mitochondrial biologists as it allows for high throughput measurements using minimal quantities of isolated mitochondria. Therefore, a collection of microplate based respirometric assays were developed that are able to assess mechanistic changes/adaptations in oxygen consumption in a commonly used animal model. The methods presented herein provide step-by-step instructions to perform these assays with an optimal amount of mitochondrial protein and reagents, and high precision as evidenced by the minimal variance across the dynamic range of each assay.

The methods presented provide step-by-step instructions for the performance of a collection of microplate based respirometric assays using isolated mitochondria from minimal quantities of mouse skeletal muscle. These assays are able to measure mechanistic changes/adaptations in mitochondrial oxygen consumption in a commonly used animal model.

Skeletal muscle mitochondria play a specific role in many disease pathologies. As such, the measurement of oxygen consumption as an indicator of ...

Developmental Biology

Measuring and Interpreting Oxygen Consumption Rates in Whole Fly Head Segments

Regulated metabolic activity is essential for the normal functioning of living cells. Indeed, altered metabolic activity is causally linked with the progression of cancer, diabetes, neurodegeneration, and aging to name a few. For instance, changes in mitochondrial activity, the cell&#39;s metabolic powerhouse, have been characterized in many such diseases. Generally, the oxygen consumption rates of mitochondria were considered a reliable readout of mitochondrial activity and measurements in some of these studies were based on isolated mitochondria or cells. However, such conditions may not represent the complexity of a whole tissue. Recently, we have developed a novel method that enables the dynamic measurement of oxygen consumption rates from whole isolated fly heads. By utilizing this method, we have recorded lower oxygen consumption rates of the whole head segment in young versus aged flies. Secondly, we have discovered that lysine deacetylase inhibitors rapidly alter the oxygen consumption in the whole head. Our novel technique may therefore aid in uncovering new properties of various drugs, which may impact metabolic rates. Furthermore, our method may give a better understanding of metabolic behavior in an experimental setup that more closely resembles physiological states.

Measuring alterations in metabolic rates is central to understanding the progression of various diseases and aging. Here, we present a novel technique to measure whole head oxygen consumption that more closely resembles the physiological state and may aid in revealing novel drugs that modify mitochondrial activity.

Regulated metabolic activity is essential for the normal functioning of living cells. Indeed, altered metabolic activity is causally linked with the ...

An Affordable and Efficient "Homemade" Platform for Drosophila Behavioral Studies, and an Accompanying Protocol for Larval Mitochondrial Respirometry

The usefulness of Drosophila as a model organism for the study of human diseases, behaviors and basic biology is unquestionable. Although practical, Drosophila research lacks popularity in developing countries, possibly due to the misinformed idea that establishing a lab and performing relevant experiments with such tiny insects is difficult and requires expensive, specialized apparatuses. Here, we describe how to build an affordable flylab to quantitatively analyze a myriad of behavioral parameters in D. melanogaster, by 3D-printing many of the necessary pieces of equipment. We provide protocols to build in-house vial racks, courtship arenas, apparatuses for locomotor assays, etc., to be used for general fly maintenance and to perform behavioral experiments using adult flies and larvae. We also provide protocols on how to use more sophisticated systems, such as a high resolution oxygraph, to measure mitochondrial oxygen consumption in larval samples, and show its association with behavioral changes in the larvae upon the xenotopic expression of the mitochondrial alternative oxidase (AOX). AOX increases larval activity and mitochondrial leak respiration, and accelerates development at low temperatures, which is consistent with a thermogenic role for the enzyme. We hope these protocols will inspire researchers, especially from developing countries, to use Drosophila to easily combine behavior and mitochondrial metabolism data, which may lead to information on genes and/or environmental conditions that may also regulate human physiology and disease states.

An Affordable and Efficient "Homemade" Platform for Drosophila Behavioral Studies, and an Accompanying Protocol for Larval Mitochondrial Respirometry

We provide protocols for anyone with a "maker culture" mind to start building a flylab for quantitative analysis of a myriad of behavioral parameters in Drosophila melanogaster, by 3D-printing many of the necessary pieces of equipment. We also describe a high resolution respirometry protocol using larvae to combine behavioral and mitochondrial metabolism data.

The usefulness of Drosophila as a model organism for the study of human diseases, behaviors and basic biology is unquestionable. Although practical, ...

Behavior

Assessing Mitochondrial Function in Sciatic Nerve by High-Resolution Respirometry

Mitochondrial dysfunction in peripheral nerves accompanies several diseases associated with peripheral neuropathy, which can be triggered by multiple causes, including autoimmune diseases, diabetes, infections, inherited disorders, and tumors. Assessing mitochondrial function in mouse peripheral nerves can be challenging due to the small sample size, a limited number of mitochondria present in the tissue, and the presence of a myelin sheath. The technique described in this work minimizes these challenges by using a unique permeabilization protocol adapted from one used for muscle fibers, to assess sciatic nerve mitochondrial function instead of isolating the mitochondria from the tissue. By measuring fluorimetric reactive species production with Amplex Red/Peroxidase and comparing different mitochondrial substrates and inhibitors in saponin-permeabilized nerves, it was possible to detect mitochondrial respiratory states, reactive oxygen species (ROS), and the activity of mitochondrial complexes simultaneously. Therefore, the method presented here offers advantages compared to the assessment of mitochondrial function by other techniques.

High-resolution respirometry coupled to fluorescence sensors determines mitochondrial oxygen consumption and reactive oxygen species (ROS) generation. The present protocol describes a technique to assess mitochondrial respiratory rates and ROS production in the permeabilized sciatic nerve.

Mitochondrial dysfunction in peripheral nerves accompanies several diseases associated with peripheral neuropathy, which can be triggered by multiple ...

Analyzing Mitochondrial Physiology and Bioenergetics in Fruit Flies

Analyzing Mitochondrial Function in a Drosophila melanogaster PINK1B9-Null Mutant Using High-resolution Respirometry

Neurodegenerative diseases, including Parkinson&#39;s Disease (PD), and cellular disturbances such as cancer are some of the disorders that disrupt energy metabolism with impairment of mitochondrial functions. Mitochondria are organelles that control both energy metabolism and cellular processes involved in cell survival and death. For this reason, approaches to evaluate mitochondrial function can offer important insights into cellular conditions in pathological and physiological processes. In this regard, high-resolution respirometry (HRR) protocols allow evaluation of the whole mitochondrial respiratory chain function or the activity of specific mitochondrial complexes. Furthermore, studying mitochondrial physiology and bioenergetics requires genetically and experimentally tractable models such as Drosophila melanogaster.
This model presents several advantages, such as its similarity to human physiology, its rapid life cycle, easy maintenance, cost-effectiveness, high throughput capabilities, and a minimized number of ethical concerns. These attributes collectively establish it as an invaluable tool for dissecting complex cellular processes. The present work explains how to analyze mitochondrial function using the Drosophila melanogaster PINK1B9-null mutant. The pink1 gene is responsible for encoding PTEN-induced putative kinase 1, through a process recognized as mitophagy, which is crucial for the removal of dysfunctional mitochondria from the mitochondrial network. Mutations in this gene have been associated with an autosomal recessive early-onset familial form of PD. This model can be used to study mitochondrial dysfunction involved in the pathophysiology of PD.

Author Spotlight: Exploring Mitochondrial Function and Chemical Toxicity Using Drosophila melanogaster 

Here we present a high-resolution respirometry protocol to analyze bioenergetics in PINK1B9-null mutant fruit flies. The method uses the Substrate-Uncoupler-Inhibitor-Titration (SUIT) protocol.

Neurodegenerative diseases, including Parkinson&#39;s Disease (PD), and cellular disturbances such as cancer are some of the disorders that disrupt energy ...

Mitochondrial Functional Assessment in Brush Lizard Liver Mitochondria

Assessment of Mitochondrial Oxygen Consumption Using a Plate Reader-based Fluorescent Assay

Mitochondria serve many important functions, including cellular respiration, ATP production, controlling apoptosis, and acting as a central hub of metabolic pathways. Therefore, experimentally assessing mitochondrial functionality can provide insight into variations among different populations or disease states. Additionally, it is valuable to assess whether isolated mitochondria are healthy enough to proceed with experiments. One characteristic often used to compare mitochondrial function in different samples is the rate of oxygen consumption. Oxygen consumption and subsequent calculation of the respiratory control ratio in either intact cells or mitochondria isolated from tissue can serve all three purposes. Using mitochondria isolated from the livers of brush lizards in conjunction with a phosphorescent probe that is sensitive to the fluctuations in oxygen concentration of a solution, we measured oxygen consumption using a fluorescent plate reader. This method is not only quick and efficient but also can be conducted with a small amount of mitochondria and without the need for specialized equipment. The step-by-step protocol described here increases the accessibility of mitochondrial functional assessment to researchers.

Assessment of oxygen consumption provides integral information about mitochondrial function. Using a phosphorescent probe with a fluorescent plate reader, accurate and reproducible data can be obtained easily without specialized equipment. This assay enables any lab to measure the oxygen consumption of isolated mitochondria and calculate respiratory control ratios.

Mitochondria serve many important functions, including cellular respiration, ATP production, controlling apoptosis, and acting as a central hub of ...

Multi-parameter Measurement of the Permeability Transition Pore Opening in Isolated Mouse Heart Mitochondria

The mitochondrial permeability transition pore (mtPTP) is a non specific channel that forms in the inner mitochondrial membrane to transport solutes with a molecular mass smaller than 1.5 kDa. Although the definitive molecular identity of the pore is still under debate, proteins such as cyclophilin D, VDAC and ANT contribute to mtPTP formation. While the involvement of mtPTP opening in cell death is well established1, accumulating evidence indicates that the mtPTP serves a physiologic role during mitochondrial Ca2+ homeostasis2, bioenergetics and redox signaling 3.
mtPTP opening is triggered by matrix Ca2+ but its activity can be modulated by several other factors such as oxidative stress, adenine nucleotide depletion, high concentrations of Pi, mitochondrial membrane depolarization or uncoupling, and long chain fatty acids4. In vitro, mtPTP opening can be achieved by increasing Ca2+ concentration inside the mitochondrial matrix through exogenous additions of Ca2+ (calcium retention capacity). When Ca2+ levels inside mitochondria reach a certain threshold, the mtPTP opens and facilitates Ca2+ release, dissipation of the proton motive force, membrane potential collapse and an increase in mitochondrial matrix volume (swelling) that ultimately leads to the rupture of the outer mitochondrial membrane and irreversible loss of organelle function.
Here we describe a fluorometric assay that allows for a comprehensive characterization of mtPTP opening in isolated mouse heart mitochondria. The assay involves the simultaneous measurement of 3 mitochondrial parameters that are altered when mtPTP opening occurs: mitochondrial Ca2+ handling (uptake and release, as measured by Ca2+ concentration in the assay medium), mitochondrial membrane potential, and mitochondrial volume. The dyes employed for Ca2+ measurement in the assay medium and mitochondrial membrane potential are Fura FF, a membrane impermeant, ratiometric indicator which undergoes a shift in the excitation wavelength in the presence of Ca2+, and JC-1, a cationic, ratiometric indicator which forms green monomers or red aggregates at low and high membrane potential, respectively. Changes in mitochondrial volume are measured by recording light scattering by the mitochondrial suspension. Since high-quality, functional mitochondria are required for the mtPTP opening assay, we also describe the steps necessary to obtain intact, highly coupled and functional isolated heart mitochondria.

A spectrofluorometric protocol for the measurement of the mitochondrial permeability transition pore opening in isolated mouse heart mitochondria is presented here. The assay involves the simultaneous measurement of mitochondria Ca2+ handling, mitochondrial membrane potential and mitochondrial volume. The procedure for obtaining high-quality and functional heart mitochondria is also described.

The mitochondrial permeability transition pore (mtPTP) is a non specific channel that forms in the inner mitochondrial membrane to transport solutes with ...

Biology

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

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 ...

Studying Mitochondrial Structure and Function in Drosophila Ovaries

Analysis of the mitochondrial structure-function relationship is required for a thorough understanding of the regulatory mechanisms of mitochondrial functionality. Fluorescence microscopy is an indispensable tool for the direct assessment of mitochondrial structure and function in live cells and for studying the mitochondrial structure-function relationship, which is primarily modulated by the molecules governing fission and fusion events between mitochondria. This paper describes and demonstrates specific methods for studying mitochondrial structure and function in live as well as in fixed tissue in the model organism Drosophila melanogaster. The tissue of choice here is the Drosophila ovary, which can be isolated and made amenable for ex vivo live confocal microscopy. Furthermore, the paper describes how to genetically manipulate the mitochondrial fission protein, Drp1, in Drosophila ovaries to study the involvement of Drp1-driven mitochondrial fission in modulating the mitochondrial structure-function relationship. The broad use of such methods is demonstrated in already-published as well as in novel data. The described methods can be further extended towards understanding the direct impact of nutrients and/or growth factors on the mitochondrial properties ex vivo. Given that mitochondrial dysregulation underlies the etiology of various diseases, the described innovative methods developed in a genetically tractable model organism, Drosophila, are anticipated to contribute significantly to the understanding of the mechanistic details of the mitochondrial structure-function relationship and to the development of mitochondria-directed therapeutic strategies.

Studying Mitochondrial Structure and Function in Drosophila Ovaries

Analysis of the mitochondrial structure-function relationship is required for a thorough understanding of the regulatory mechanisms of mitochondrial functionality. Specific methods for studying mitochondrial structure and function in live and fixed Drosophila ovaries are described and demonstrated in this paper.

Analysis of the mitochondrial structure-function relationship is required for a thorough understanding of the regulatory mechanisms of mitochondrial ...

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

Samenvatting

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