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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S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Thiazolidinediones+are+acute,+specific+inhibitors+of+the+mitochondrial+pyruvate+carrier.">Thiazolidinediones are acute, specific inhibitors of the mitochondrial pyruvate carrier.</a> Proceedings of the National Academy of Sciences of the United States of America. 110 (14), 5422-5427 (2013).</li><li>McCommis, K., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=An+ancestral+role+for+the+mitochondrial+pyruvate+carrier+in+glucose-stimulated+insulin+secretion.">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 border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>High-resolution respirometer Oxygraph O2K</td>
			<td>Oroboros Instruments, Innsbruck, Austria</td>
			<td>10022-02</td>
			<td>Startup O2K respirometer kit 
			&#160;</td>
		</tr>
		<tr>
			<td>O2K-Titration Set&#160;</td>
			<td>Oroboros Instruments, Innsbruck, Austria</td>
			<td>20820-03</td>
			<td>Hamilton syringes with different volumes 
			&#160;</td>
		</tr>
		<tr>
			<td>Datlab software</td>
			<td>Oroboros Instruments, Innsbruck, Austria</td>
			<td>20700</td>
			<td>Software for data acquisition and analysis 
			&#160;</td>
		</tr>
		<tr>
			<td>Fine-tipped antimagnetic forceps</td>
			<td>VWR</td>
			<td>82027-400</td>
			<td> 
			&#160;</td>
		</tr>
		<tr>
			<td>Secura225D-1S-DQE</td>
			<td>Sartorius AG, Goettingen, Germany</td>
			<td></td>
			<td>Semi-micro balance (distributed by several companies)&#160; 
			&#160;</td>
		</tr>
		<tr>
			<td>Drosophila melanogaster wild-type w1118</td>
			<td>Bloomington Drosophila stock Center, IN, USA</td>
			<td></td>
			<td> 
			Storage Condition: 24 &#176;C</td>
		</tr>
		<tr>
			<td>Ethylene glycol-bis(2-aminoethylether)-N,N,N&#8242;,N&#8242;-tetraacetic acid</td>
			<td>Sigma-Aldrich</td>
			<td>E4378</td>
			<td>EGTA 
			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. 
			Storage Condition: RT</td>
		</tr>
		<tr>
			<td>CaCO3</td>
			<td>Sigma-Aldrich</td>
			<td>C4830</td>
			<td> 
			Storage Condition: RT</td>
		</tr>
		<tr>
			<td>Na2ATP</td>
			<td>Sigma-Aldrich</td>
			<td>A2383</td>
			<td> 
			Storage Condition: -20 &#176;C</td>
		</tr>
		<tr>
			<td>MgCl2.6H2O</td>
			<td>Sigma-Aldrich</td>
			<td>M9272</td>
			<td> 
			Storage Condition: RT</td>
		</tr>
		<tr>
			<td>Taurine</td>
			<td>Sigma-Aldrich</td>
			<td>T0625</td>
			<td> 
			Storage Condition: RT</td>
		</tr>
		<tr>
			<td>Na2Phosphocreatine</td>
			<td>Sigma-Aldrich</td>
			<td>P7936</td>
			<td> 
			Storage Condition: -20 &#176;C</td>
		</tr>
		<tr>
			<td>Imidazole</td>
			<td>Sigma-Aldrich</td>
			<td>I5513</td>
			<td> 
			Storage Condition: RT</td>
		</tr>
		<tr>
			<td>Dithiothreitol</td>
			<td>Sigma-Aldrich</td>
			<td>D0632</td>
			<td> 
			Storage Condition: 2-8 &#176;C</td>
		</tr>
		<tr>
			<td>MES hydrate</td>
			<td>Sigma-Aldrich</td>
			<td>M8250</td>
			<td> 
			Storage Condition: RT</td>
		</tr>
		<tr>
			<td>Saponin from quillaja bark</td>
			<td>Sigma-Aldrich</td>
			<td>S7900</td>
			<td>Saponin 
			Storage Condition: RT 
			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> 
			Storage Condition: RT</td>
		</tr>
		<tr>
			<td>KH2PO4</td>
			<td>Sigma-Aldrich</td>
			<td>P9791</td>
			<td> 
			Storage Condition: RT</td>
		</tr>
		<tr>
			<td>HEPES</td>
			<td>Sigma-Aldrich</td>
			<td>H3375</td>
			<td> 
			Storage Condition: RT</td>
		</tr>
		<tr>
			<td>BSA</td>
			<td>Sigma-Aldrich</td>
			<td>05470</td>
			<td> 
			Storage Condition: 2-8 &#176;C</td>
		</tr>
		<tr>
			<td>Na2S2O4</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. 
			Storage Condition: RT</td>
		</tr>
		<tr>
			<td>Sodium pyruvate</td>
			<td>Sigma-Aldrich</td>
			<td>P2256</td>
			<td>Pyruvate 
			Storage Condition: 2-8 &#176;C 
			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 
			Storage Condition: RT 
			Solution Preparation: In MilliQ water. Neutralize with KOH and store at -20 &#176;C.</td>
		</tr>
		<tr>
			<td>Adenosine 5&#39;-diphosphate monopotassium salt hydrate</td>
			<td>Sigma-Aldrich</td>
			<td>A5285</td>
			<td>ADP 
			Storage Condition: -20 &#176;C 
			Solution Preparation: In MilliQ water. Neutralize with KOH and store at -80 &#176;C.</td>
		</tr>
		<tr>
			<td>Cytochrome c from equine heart</td>
			<td>Sigma-Aldrich</td>
			<td>C7752</td>
			<td>Cytochrome c 
			Storage Condition: -20 &#176;C 
			Solution Preparation: In MilliQ water. Store at -20 &#176;C.</td>
		</tr>
		<tr>
			<td>L-Proline</td>
			<td>Sigma-Aldrich</td>
			<td>P0380</td>
			<td>Proline 
			Storage Condition: RT 
			Solution Preparation: In MilliQ water. Store at -20 &#176;C.</td>
		</tr>
		<tr>
			<td>Sodium succinate dibasic hexahydrate</td>
			<td>Sigma-Aldrich</td>
			<td>S2378</td>
			<td>Succinate 
			Storage Condition: RT 
			Solution Preparation: In MilliQ water. Neutralize with HCl and store at -20 &#176;C.</td>
		</tr>
		<tr>
			<td>sn-Glycerol 3-phosphate bis(cyclohexylammonium) salt</td>
			<td>Sigma-Aldrich</td>
			<td>G7886</td>
			<td>Glycerol-3-phosphate 
			Storage Condition: -20 &#176;C 
			Solution Preparation: In MilliQ water. Neutralize with HCl and store at -80 &#176;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. 
			Storage Condition: RT 
			Solution Preparation: In absolute ethanol. Store in glass vials at -20 &#176;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. 
			Solution Preparation: In absolute ethanol. Store in dark vials at -20 &#176;C.</td>
		</tr>
		<tr>
			<td>Malonic acid</td>
			<td>Sigma-Aldrich</td>
			<td>M1296</td>
			<td>Malonate. CAUTION: acute toxicity, serious eye damage. 
			Storage Condition: RT 
			Solution Preparation: In MilliQ water. Neutralize with KOH. Prepare fresh daily.</td>
		</tr>
		<tr>
			<td>Antimycin A from Streptomyces sp.</td>
			<td>Sigma-Aldrich</td>
			<td>A8674</td>
			<td>Antimycin A. CAUTION: acute toxicity, acute aquatic toxicity, chronic aquatic toxicity. 
			Storage Condition: -20 &#176;C 
			Solution Preparation: In absolute ethanol. Store at -20 &#176;C.</td>
		</tr>
		<tr>
			<td>N,N,N&#8242;,N&#8242;-Tetramethyl-p-phenylenediamine</td>
			<td>Sigma-Aldrich</td>
			<td>T7394</td>
			<td>TMPD 
			Storage Condition: RT 
			Solution Preparation: In MilliQ water. Store in dark vials at&#160; -20 &#176;C.</td>
		</tr>
		<tr>
			<td>(+)-Sodium L-ascorbate</td>
			<td>Sigma-Aldrich</td>
			<td>A4034</td>
			<td>Ascorbate 
			Storage Condition: RT 
			Solution Preparation: In MilliQ water. Store in dark vials at&#160; -20 &#176;C.</td>
		</tr>
		<tr>
			<td>NaN3</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.&#160; 
			Solution Preparation: In MilliQ water. Store at -20 &#176;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

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JoVE Journal

Biochemistry

11.8K Views.  Université de Moncton. 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. 

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

Using Scaffold Liposomes to Reconstitute Lipid-proximal Protein-protein Interactions In Vitro

Studies of integral membrane proteins in vitro are frequently complicated by the presence of a hydrophobic transmembrane domain. Further complicating these studies, reincorporation of detergent-solubilized membrane proteins into liposomes is a stochastic process where protein topology is impossible to enforce. This paper offers an alternative method to these challenging techniques that utilizes a liposome-based scaffold. Protein solubility is enhanced by deletion of the transmembrane domain, and these amino acids are replaced with a tethering moiety, such as a His-tag. This tether interacts with an anchoring group (Ni2+ coordinated by nitrilotriacetic acid (NTA(Ni2+)) for His-tagged proteins), which enforces a uniform protein topology at the surface of the liposome. An example is presented wherein the interaction between Dynamin-related protein 1 (Drp1) with an integral membrane protein, Mitochondrial Fission Factor (Mff), was investigated using this scaffold liposome method. In this work, we have demonstrated the ability of Mff to efficiently recruit soluble Drp1 to the surface of liposomes, which stimulated its GTPase activity. Moreover, Drp1 was able to tubulate the Mff-decorated lipid template in the presence of specific lipids. This example demonstrates the effectiveness of scaffold liposomes using structural and functional assays and highlights the role of Mff in regulating Drp1 activity.

Using Scaffold Liposomes to Reconstitute Lipid-proximal Protein-protein Interactions In Vitro

This paper describes a method for assessing the interactions and assemblies of integral membrane proteins in vitro with various partner factors in a lipid-proximal environment.

Studies of integral membrane proteins in vitro are frequently complicated by the presence of a hydrophobic transmembrane domain. Further complicating ...

Simultaneous Measurement of Superoxide/Hydrogen Peroxide and NADH Production by Flavin-containing Mitochondrial Dehydrogenases

It has been reported that mitochondria can contain up to 12 enzymatic sources of reactive oxygen species (ROS). A majority of these sites include flavin-dependent respiratory complexes and dehydrogenases that produce a mixture of superoxide (O2&#9679;-) and hydrogen peroxide (H2O2). Accurate quantification of the ROS-producing potential of individual sites in isolated mitochondria can be challenging due to the presence of antioxidant defense systems&#160;and side reactions that also form O2&#9679;-/H2O2. Use&#160;of nonspecific inhibitors that can disrupt mitochondrial bioenergetics can also compromise measurements by altering&#160;ROS release from other sites of production. Here, we present an easy method for the simultaneous measurement of H2O2 release and nicotinamide adenine dinucleotide (NADH) production by purified flavin-linked dehydrogenases. For our purposes here, we have used purified pyruvate dehydrogenase complex (PDHC) and &#945;-ketoglutarate dehydrogenase complex (KGDHC) of porcine heart origin as examples. This method allows for an accurate measure of native H2O2 release rates by individual sites of production by eliminating other potential sources of ROS and antioxidant systems. In addition, this method allows for a direct comparison of the relationship between H2O2 release and enzyme activity and the screening of the effectiveness and selectivity of inhibitors for ROS production. Overall, this approach can allow for the in-depth assessment of native rates of ROS release for individual enzymes prior to conducting more sophisticated experiments with isolated mitochondria or permeabilized muscle fiber.

Mitochondria contain several flavin-dependent enzymes that can produce reactive oxygen species (ROS). Monitoring ROS release from individual sites in mitochondria is challenging due to unwanted side reactions. We present an easy, inexpensive method for direct assessment of native rates for ROS release using purified flavoenzymes and microplate fluorometry.

It has been reported that mitochondria can contain up to 12 enzymatic sources of reactive oxygen species (ROS). A majority of these sites include ...

Analyses of Mitochondrial Calcium Influx in Isolated Mitochondria and Cultured Cells

Ca2+ handling by mitochondria is a critical function regulating both physiological and pathophysiological processes in a broad spectrum of cells. The ability to accurately measure the influx and efflux of Ca2+ from mitochondria is important for determining the role of mitochondrial Ca2+ handling in these processes. In this report, we present two methods for the measurement of mitochondrial Ca2+ handling in both isolated mitochondria and cultured cells. We first detail a plate reader-based platform for measuring mitochondrial Ca2+ uptake using the Ca2+ sensitive dye calcium green-5N. The plate reader-based format circumvents the need for specialized equipment, and the calcium green-5N dye is ideally suited for measuring Ca2+ from isolated tissue mitochondria. For our application, we describe the measurement of mitochondrial Ca2+ uptake in mitochondria isolated from mouse heart tissue; however, this procedure can be applied to measure mitochondrial Ca2+ uptake in mitochondria isolated from other tissues such as liver, skeletal muscle, and brain. Secondly, we describe a confocal microscopy-based assay for measurement of mitochondrial Ca2+ in permeabilized cells using the Ca2+ sensitive dye Rhod-2/AM and imaging using 2-dimensional laser-scanning microscopy. This permeabilization protocol eliminates cytosolic dye contamination, allowing for specific recording of changes in mitochondrial Ca2+. Moreover, laser-scanning microscopy allows for high frame rates to capture rapid changes in mitochondrial Ca2+ in response to various drugs or reagents applied in the external solution. This protocol can be applied to measure mitochondrial Ca2+ uptake in many cell types including primary cells such as cardiac myocytes and neurons, and immortalized cell lines.

Here, we present two protocols for the measurement of mitochondrial Ca2+ influx in isolated mitochondria and cultured cells. For isolated mitochondria, we detail a plate reader-based Ca2+ import assay using the Ca2+ sensitive dye calcium green-5N. For cultured cells, we describe a confocal microscopy method using the Ca2+ dye Rhod-2/AM.

Ca2+ handling by mitochondria is a critical function regulating both physiological and pathophysiological processes in a broad spectrum of cells. The ...

Measuring Liver Mitochondrial Oxygen Consumption and Proton Leak Kinetics to Estimate Mitochondrial Respiration in Holstein Dairy Cattle

Oxygen consumption, proton motive force (PMF) and proton leak are measurements of mitochondrial respiration, or how well mitochondria are able to convert NADH and FADH into ATP. Since mitochondria are also the primary site for oxygen use and nutrient oxidation to carbon dioxide and water, how efficiently they use oxygen and produce ATP directly relates to the efficiency of nutrient metabolism, nutrient requirements of the animal, and health of the animal. The purpose of this method is to examine mitochondrial respiration, which can be used to examine the effects of different drugs, diets and environmental effects on mitochondrial metabolism. Results include oxygen consumption measured as proton dependent respiration (State 3) and proton leak dependent respiration (State 4). The ratio of State 3 / State 4 respiration is defined as respiratory control ratio (RCR) and can represent mitochondrial energetic efficiency. Mitochondrial proton leak is a process that allows dissipation of mitochondrial membrane potential (MMP) by uncoupling oxidative phosphorylation from ADP decreasing the efficiency of ATP synthesis. Oxygen and TRMP+ sensitive electrodes with mitochondrial substrates and electron transport chain inhibitors are used to measure State 3 and State 4 respiration, mitochondrial membrane PMF (or the potential to produce ATP) and proton leak. Limitations to this method are that liver tissue must be as fresh as possible and all biopsies and assays must be performed in less than 10 h. This limits the number of samples that can be collected and processed by a single person in a day to approximately 5. However, only 1 g of liver tissue is needed, so in large animals, such as dairy cattle, the amount of sample needed is small relative to liver size and there is little recovery time needed.

Here, we share methods for measuring mitochondrial oxygen consumption, a defining concept of nutritional energetics, and proton leak, the primary cause of inefficiency in mitochondrial generation of ATP. These results can account for 30% of the energy lost in nutrient utilization to help evaluate mitochondrial function.

Oxygen consumption, proton motive force (PMF) and proton leak are measurements of mitochondrial respiration, or how well mitochondria are able to convert ...

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 ...

Direct Measurement of KDM1A Target Engagement Using Chemoprobe-based Immunoassays

The assessment of the target engagement, defined as the interaction of a drug with the protein it was designed for, is a basic requirement for the interpretation of the biological activity of any compound in drug development or in basic research projects. In epigenetics, target engagement is most often assessed by the analysis of proxy markers instead of measuring the union of the compound to the target. Downstream biological readouts that have been analyzed include the histone mark modulation or gene expression changes. KDM1A is a lysine demethylase that removes methyl groups from mono- and dimethylated H3K4, a modification associated with the silencing of gene expression. Modulation of the proxy markers is dependent on the cell type and function of the genetic make-up of the cells investigated, which can make interpretation and cross-case comparison quite difficult. To circumvent these problems, a versatile protocol is presented to assess the dose effects and dynamics of the direct KDM1A target engagement. The assay described makes use of a KDM1A chemoprobe to capture and quantify uninhibited enzyme, can be broadly applied to cells or tissue samples without the need for genetic modification, has an excellent window of detection, and can be used both for basic research and analysis of clinical samples.

Here, we present a protocol to measure KDM1A target engagement in a human or animal cell, tissue or blood samples treated with KDM1A inhibitors. The protocol employs chemoprobe tagging of the free KDM1A enzyme and direct quantification of the target occupation using chemoprobe-based immunoassays and can be used in preclinical and clinical studies.

The assessment of the target engagement, defined as the interaction of a drug with the protein it was designed for, is a basic requirement for the ...

Rapid and Cost-Effective RNA Extraction of Rat Pancreatic Tissue

Regardless of the extraction method, optimized RNA extraction of tissues and cell lines are carried out in four stages: 1) homogenization, 2) effective denaturation of proteins from RNA, 3) ribonuclease inactivation, and 4) removal of contamination from DNA, proteins, and carbohydrates. However, it is very laborious to maintain the integrity of RNA when there are high levels of RNase in the tissue. Spontaneous autolysis makes it very difficult to extract RNA from pancreatic tissue without damaging it. Thus, a practical RNA extraction method is needed to maintain the integrity of pancreatic tissues during the extraction process. An experimental and comparative study of existing protocols was carried out by obtaining 20-30 mg of rat pancreatic tissues in less than 2 minutes and extracting the RNA. The results were assessed by electrophoresis. The experiments were carried out three times for generalization of the results. Immersing pancreatic tissue in RNA stabilization reagent at -80 &#176;C for 24 h yielded high integrity RNA, when the RNA extraction reagent was used as the reagent. The results obtained were comparable to the results obtained from commercial kits with spin column bindings.

The purity and integrity of the isolated RNA is a vital step in RNA dependent assays. Here, we present a practical, rapid, and inexpensive method to extract RNA from a small quantity of undamaged pancreatic tissue.

Regardless of the extraction method, optimized RNA extraction of tissues and cell lines are carried out in four stages: 1) homogenization, 2) effective ...

Measuring Mitochondrial Electron Transfer Complexes in Previously Frozen Cardiac Tissue from the Offspring of Sow: A Model to Assess Exercise-Induced Mitochondrial Bioenergetics Changes

The mitochondrial electron transfer complex (ETC) profile is modified in the heart tissue of the offspring born to an exercised sow. The hypothesis proposed and tested was that a regular maternal exercise of a sow during pregnancy would increase the mitochondrial efficiency of offspring heart bioenergetics. This hypothesis was tested by isolating mitochondria using a mild-isolation procedure to assess mitochondrial ETC and supercomplex profiles. The procedure described here allowed for the processing of previously frozen archived heart tissues and eliminated the necessity of fresh mitochondria preparation for the assessment of mitochondrial ETC complexes, supercomplexes, and ETC complex activity profiles. This protocol describes the optimal ETC protein complex measurement in multiplexed antibody-based immunoblotting and super complex assessment using blue-native gel electrophoresis.

Preparation of mitochondria-enriched samples from previously frozen archived solid tissues allowed the investigators to perform both functional and analytical assessments of mitochondria in various experimental modalities. This study demonstrates how to prepare mitochondria-enriched preparations from frozen heart tissue and perform analytical assessments of mitochondria.

The mitochondrial electron transfer complex (ETC) profile is modified in the heart tissue of the offspring born to an exercised sow. The hypothesis ...

Quantification of Subcellular Glycogen Distribution in Skeletal Muscle Fibers using Transmission Electron Microscopy

With the use of transmission electron microscopy, high-resolution images of fixed samples containing individual muscle fibers can be obtained. This enables quantifications of ultrastructural aspects such as volume fractions, surface area to volume ratios, morphometry, and physical contact sites of different subcellular structures. In the 1970s, a protocol for enhanced staining of glycogen in cells was developed and paved the way for a string of studies on the subcellular localization of glycogen and glycogen particle size using transmission electron microscopy. While most analyses interpret glycogen as if it is homogeneously distributed within the muscle fibers, providing only a single value (e.g., an average concentration), transmission electron microscopy has revealed that glycogen is stored as discrete glycogen particles located in distinct subcellular compartments. Here, the step-by-step protocol from tissue collection to the quantitative determination of the volume fraction and particle diameter of glycogen in the distinct subcellular compartments of individual skeletal muscle fibers is described. Considerations on how to 1) collect and stain tissue specimens, 2) perform image analyses and data handling, 3) evaluate the precision of estimates, 4) discriminate between muscle fiber types, and 5) methodological pitfalls and limitations are included.

A modified post-fixation procedure increases the contrast of glycogen particles in tissue. This paper provides a step-by-step protocol describing how to handle the tissue, conduct the imaging, and use stereological methods to obtain unbiased and quantitative data on fiber type-specific subcellular glycogen distribution in skeletal muscle.

With the use of transmission electron microscopy, high-resolution images of fixed samples containing individual muscle fibers can be obtained. This ...

Generation of Tissue Spheroids via a 3D Printed Stamp-Like Device

Advances in 3D cell culture have developed more physiologically relevant in vitro models, such as tissue spheroids. Cells cultivated as spheroids have more realistic biological responses that resemble the in vivo environment. Due to their advantages, tissue spheroids represent an emerging trend toward superior, more reliable, and more predictive study models with a broad range of biotechnological applicability. However, reproducible platforms that can achieve large-scale production of tissue spheroids have become an unmet need in fully exploring and boosting their potential. Herein, the large-scale production of homogeneous tissue spheroids is reported using a low-cost and time-effective methodology. A 3D printed stamp-like device is developed to generate up to 4,716 spheroids per 6-well plate. The device is fabricated by the stereolithography method using a photocurable resin. The final device is composed of cylindrical micropins, with a height of 1.3 mm and a width of 650 &#181;m. This approach allows the fast generation of homogeneous spheroids and co-cultured spheroids with uniform shape and size and &gt;95% cell viability. Moreover, the stamp-like device is tunable for different sizes of well plates and Petri dishes. It is easily sterilized and can be reused for long periods. The efficient large-scale production of homogeneous tissue spheroids is essential to leverage their translation for multiple areas of industry, such as tissue engineering, drug development, disease modeling, and on-demand personalized medicine.

Generation of Tissue Spheroids via a 3D Printed Stamp-Like Device

The present protocol describes a technique to produce tissue spheroids on a large scale cost-effectively using a 3D printed stamp-like device.

Advances in 3D cell culture have developed more physiologically relevant in vitro models, such as tissue spheroids. Cells cultivated as spheroids have ...