The authors acknowledge the Brazilian agency Coordena&#231;&#227;o de Aperfei&#231;oamento de Pesquisa Pessoal de N&#237;vel Superior (CAPES EPIDEMIAS 09 #88887.505377/2020). P.M. (#88887.512821/2020-00) and T.D. (#88887.512883/2020-00) are research fellowship recipients.

We used the strains w1118 (white) and w[*] Pink1[B9]/FM7i, P{w[+mC]=ActGFP}JMR3 (referred to as Pink1B9) (FlyBase ID: FBgn0029891) from the Bloomington Drosophila stock center (ID number: 34749). In this study, male D. melanogaster PINK1B9-null mutants are compared with male D. melanogaster from the w1118&#160;strain, which is used as a control group (genetic background). Other parameters must be analyzed concomitantly with the respirometry experiments to ensure that the flies have the correct genotype (Pink1B9/Y), such as thorax deformities and locomotion problems, which are well described for pink1B9 mutant flies24,25,26.
1. Animals and housing
<ol>
	<li>Keep the flies in glass bottles containing 10 mL of standard diet (yeast 1.73%, soy flour 0.9%, corn flour 7.3%, agar 0.5%, corn syrup 7.6%, and propionic acid 0.48%) at constant temperature and humidity (25 &#176;C and 60%, respectively), with a 12 h:12 h light/dark cycle.</li>
	<li>For each experiment, use two flies aged 1-3 days post-eclosion. 
	&#8203;NOTE: The PINK1B9-null mutant virgin female flies were crossed with w1118 males to obtain the F1 males with genotype Pink1B9/Y.</li>
</ol>
2. Sample preparation
<ol>
	<li>Prepare the MiR05 buffer as described in Table 1.</li>
	<li>Anesthetize the flies on ice and transfer them to microcentrifuge tubes (two flies per tube).</li>
	<li>In each tube, add 200 &#181;L of chilled MiR05 buffer and homogenize the flies. Homogenize manually, applying gentle pressure (approximately 4-6 strokes of the pestle) to prevent the disintegration of mitochondria.</li>
</ol>
3. High-resolution respirometry calibration of polarographic oxygen sensors
NOTE: The OROBOROS chambers have a total volume of approximately 2 mL. Calibration is required to ensure the oxygen flux is close to 0 pmol to start the assay.
<ol>
	<li>Starting the calibration, add 2 mL of MiR05 to the chamber. Cover the chamber with the stopper leaving no air bubbles; then, carefully pull the stopper to form one single air bubble.</li>
	<li>Open the OROBOROS Dat.Lab software. Enter a temperature of 25 &#176;C in the Block temperature field and click on Connect to Oxygraph-2k (Figure 1A).</li>
	<li>Click and select the folder where the calibration is to be saved and click on Save.</li>
	<li>When a new window opens, name the experiment and the samples (if calibrating, simply enter the name calibration) and click on Save.</li>
	<li>Go to the Layout menu and choose the first option: 01 Calibration Exp. Gr.3 - Temp as shown in Figure 1B.</li>
	<li>Wait for the program to redirect to the calibration layout, wait until the red line shows a standard straight line with points around 0 pmol. Then, select two points: one for chamber A and the other for chamber B (Figure 1C) when the red line is exactly at zero. 
	NOTE: The selected points must have oxygen flux (red line) around 0.
	<ol>
		<li>To mark the points of both chambers, select O2 concentration on the right of the screen, select the marked point, and copy both the temperature and barometric pressure values relative to the point. Paste these values in the boxes below and click on Calibrate and Copy to Clipboard in the lower right corner of the screen as shown in Figure 2.</li>
		<li>Carry out this process for both chambers (A and B), then go to the File menu and select Save and Disconnect. 
		&#8203;NOTE: After the calibration process, the program will redirect the user to the home screen so that the software will be ready to start the HRR test. For this test, we will use the SUIT protocol.</li>
	</ol>
	</li>
</ol>
4. SUIT protocol
<ol>
	<li>With the calibration completed, remove the stoppers and open the chambers.</li>
	<li>Wash the stoppers (holding the tip that does not touch the sample) and chambers with 100% ethanol, 70% ethanol, and distilled water, in that order. 
	NOTE: Repeat the process for every sample change.</li>
	<li>Homogenize the flies in 200 &#181;L of buffer, place all sample content in the chamber using a micropipette, and relocate the stopper in the chamber, taking care to not create air bubbles. 
	NOTE: If air bubbles form, remove the stopper gently and add 100 &#181;L of MIR05 buffer.</li>
	<li>Click on the Layout menu and select the option 05 Flow by Corrected Volume as shown in Figure 1B.</li>
	<li>Wait to be redirected to a new window. Click on Save to select the folder where the experiment is to be saved.</li>
	<li>Wait for another new window to open. Name the experiment in the space Experimental Code. Then, name each sample in its respective chamber A and B in the Sample field, and set the field Unit to unit. 
	NOTE: Other units can also be chosen, for example, Millions of cells or mg.</li>
	<li>Considering that this protocol uses two flies (2 units) and the chamber volume is 2 mL, in the Concentration field, define 1 per mL (Figure 1D). 
	NOTE: Here, the sample amount was normalized by the number of flies; however, this normalization may be performed by the protein content of the sample, mitochondrial DNA quantification, or activity of citrate synthase.</li>
	<li>When the oxygen flux signal (red line) is steady on the positive value, start the HRR protocol with the titration of substrates, inhibitors, and uncouplers (Figure 3). All the subsequent reagents and equipment are shown in Table 1.
	<ol>
		<li>Add digitonin (5 &#956;g/mL) to permeabilize the mitochondrial membrane.</li>
		<li>Add pyruvate (5 mM), malate (2 mM), and proline (10 mM) substrates and wait until the oxygen flux increases and stabilizes. To mark the event for each reagent addition, press the F4 key on the keyboard and enter the name of each reagent.</li>
		<li>Add ADP (5mM) to couple the mitochondrial respiratory chain and wait for the increase and stabilization of the oxygen flux.</li>
		<li>Add succinate (10 mM) and wait for the increase and stabilization of the oxygen flux.</li>
		<li>Add Oligomycin (2.5 &#181;M) to inhibit ATP synthase and look for a decrease in oxygen flux.</li>
		<li>Wait until the red line stabilizes and uncouple mitochondrial electron transfer using Carbonyl-4-(trifluoromethoxy) phenylhydrazone cyanide (FCCP) with titers of 0.25 &#181;M until the maximum oxygen consumption is reached, demonstrated by the rise of the red line. After stabilization of the oxygen flux, start adding the inhibitors of each complex, one at a time.</li>
		<li>Add Rotenone (0.5 &#181;M), a complex I inhibitor. Wait for the decrease and stabilization of the oxygen flux to add the next inhibitor.</li>
		<li>Add Malonate (5 mM), a complex II inhibitor. Wait for the decrease and stabilization of the oxygen flux to add the next inhibitor.</li>
		<li>Finally, add Antimycin (2.5 &#181;M), a complex III inhibitor. Wait for the decrease followed by the increase and stabilization of the oxygen flux. 
		NOTE: The oxygen flux should stabilize at a positive value but below the value for the last inhibition of mitochondrial protein complexes.</li>
	</ol>
	</li>
	<li>At the end of the analysis, remove all the content of the chambers using a micropipette. Store at -20 &#176;C for future analysis, when necessary.</li>
</ol>
5. Data analysis
<ol>
	<li>Utilize an appropriate statistical software package for data analysis.</li>
	<li>Perform t-tests to assess differences between the two groups. For situations involving multiple treatments, use Analysis of Variance (ANOVA) as a statistical test 27.</li>
	<li>Extract the O2 flux values from the graph generated by the software. OXPHOS CI refers to the value of O2 flux after adding ADP. OXPHOS CII is obtained by subtracting OXPHOS CI&#38;CII - OXPHOS CI. OXPHOS CI&#38;CII is the value of O2 flux extracted after the addition of succinate. ETS CI&#38;CII is the O2 flux value after adding FCCP. ETS CI = FCCP - Rotenone and ETS CII = Rotenone - Malonate.</li>
	<li>Calculate ATP synthesis as the difference between the OXPHOS (P) and LEAK (L) (P - L)28.</li>
	<li>Determine the Respiratory Control Ratio (RCR) by calculating the ratio OXPHOS/LEAK, where RCR = P/L.11</li>
</ol>

Analyzing Mitochondrial Physiology and Bioenergetics in Fruit Flies

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Cell Physiol. 292 (1), C125-C136 (2007).</li><li>Divakaruni, A. S., Jastroch, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+practical+guide+for+the+analysis,+standardization+and+interpretation+of+oxygen+consumption+measurements.">A practical guide for the analysis, standardization and interpretation of oxygen consumption measurements.</a> Nat Metab. 4 (8), 978-994 (2022).</li><li>Bingol, B., Sheng, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mechanisms+of+mitophagy:+PINK1,+Parkin,+USP30+and+beyond.">Mechanisms of mitophagy: PINK1, Parkin, USP30 and beyond.</a> Free Radic Biol Med. 100, 210-222 (2016).</li></ol>

The authors declare no competing interests.

HRR is a powerful technique for studying mitochondrial respiration and energy metabolism in D. melanogaster and other organisms. It provides a detailed and quantitative assessment of mitochondrial function, allowing researchers to gain insights into the bioenergetics of the cells. The protocol presented here describes the evaluation of mitochondrial respiratory chain function and the activity of specific mitochondrial complexes using the SUIT protocol in D. melanogaster. The SUIT protocol involves systematically manipulating various substrates, uncouplers, and inhibitors to examine different aspects of mitochondrial respiration.
The technique described allows the assessment of respiratory inhibition resulting from effects on the OXPHOS or the ETS, dehydrogenase activity (CI and CII), and membrane integrity (coupling of OXPHOS). Here, we performed the experiments using PINK1B9-null flies to study mitochondrial dysfunction since it is associated with PD34,35. However, this protocol can be useful for different disease models, drug treatments, and toxicology studies.
In addition, the sample preparation may be adapted to fit experimental requirements36. A critical step in the respirometry protocol is sample preparation. It is critical that the correct protocol be established for the type of sample (cell, isolated mitochondria, homogenate), and it is important to consider the appropriate method for sample normalization (protein amount, DNA content, citrate synthase activity). It is also important to use the same buffer for sample preparation and the respirometry assay.
As the sample preparation protocol described here is simple, it does not usually pose problems for the technique. If another type of sample is chosen or its preparation changed, then careful standardization is necessary. Stirring and temperature stability affect the signal of the polarographic oxygen sensor, generating error in respiratory measurements using an oxygraph. Therefore, the correct calibration of the chamber constitutes an important step to reduce errors during the respiratory measurements.
This method for assessing mitochondrial function in fly samples offers several advantages over alternative approaches. One of the main strengths of HRR is its ability to provide direct and accurate measurements of oxygen consumption, allowing for a detailed analysis of mitochondrial function and cellular metabolism. Therefore, HRR is often used in research focused on understanding mitochondrial dysfunction, energy production, and cellular responses to different substrates or conditions. Furthermore, it is versatile - allowing the use of a wide variety of sample types, including isolated mitochondria - and requires small amounts of biological samples, which is useful when the sample amount is limited1,2,3. Methods that use mitochondrial isolates, for instance, typically involve a substantial number of flies, ranging from 50 to 200 individuals, which makes the study with mutants difficult, as obtaining a large number of mutants for certain disease models may not be practical.
Similar to the HRR method, the Seahorse Bioscience Extracellular Flux Analyzer is a scientific tool used to measure oxygen consumption and extracellular acidification. However, they have different approaches and applications. The Seahorse Bioscience Extracellular Flux Analyzer quantifies instantaneous alterations in the oxygen consumption rate (OCR) and extracellular acidification rate (ECAR) of cells. OCR is indicative of mitochondrial respiration and energy generation, whereas ECAR offers insights into glycolytic activity. Its principal function lies in the assessment of cellular metabolic dynamics under diverse physiological conditions, rendering it particularly instrumental in elucidating the intricate metabolic modulations associated with pathological contexts. In addition, Seahorse is designed for ease of use, allowing researchers to perform rapid metabolic assays37,38,39. In contrast, the HRR requires specialized training and expertise to operate and analyze data effectively. It involves the use of an oxygen electrode to directly measure oxygen consumption by cells or mitochondria13,14,40. An advantage of HRR is its versatility in designing different respirometry protocols, which is impracticable when using Seahorse. This versatility to design protocols associated with low cost makes the HRR method a viable choice for laboratories with limited funding. Another disadvantage of the respirometry system used in the present protocol is the impossibility of working with several samples simultaneously, which hampers the high-throughput analysis. Thus, experimental groups must be well-designed to allow the correct comparison between them. However, the advantages of this system lie in the fact that samples of any kind can be tested, from isolated cells to whole live organisms, such as C. elegans, for example40.
In summary, HRR is a reliable method for studying mitochondrial function under physiological and pathological conditions41. Each experimental model has different characteristics and restrictions, requiring methodology and sample preparation adjustments to ensure reliable and meaningful data acquisition in mitochondrial respiration evaluation. This protocol offers researchers a reliable method to assess the effects of environmental factors, experimental interventions, or genetic mutations on mitochondrial function in D. melanogaster.

Mitochondria are cellular organelles that control important functions, including apoptotic regulation, calcium homeostasis, and participation in biosynthetic pathways. By possessing autonomous genetic material, they are capable of contributing to cellular maintenance and repair processes. Their structure houses the electron transport chain and oxidative phosphorylation, both crucial for cellular energy1,2,3. In particular, energy control is achieved through adenosine triphosphate (ATP) production via oxidative phosphorylation (OXPHOS)2. Disruption of energy metabolism with impairment of mitochondrial functions occurs both in cell survival and death4,5, frequently associated with a wide range of human pathologies, such as cancer, and neurodegenerative diseases such as Parkinson&#39;s Disease (PD)3,6.
PD is a chronic, progressive, and neurological disorder. The primary cause of this disease is the death of brain cells, especially in the substantia nigra, which are responsible for the production of the neurotransmitter dopamine, which controls movement6,7,8. The earliest observation that linked Parkinsonism to mitochondrial dysfunction was made in 1988, in experimental models using toxins that inhibit the respiratory chain Complex I9.
Currently, there are several methods to evaluate mitochondrial dysfunction10,11,12,13; however, compared to conventional approaches, high-resolution respirometry (HRR) presents superior sensitivity and advantages13,14. For example, HRR protocols allow the evaluation of the whole mitochondrial respiratory chain function or the activity of specific mitochondrial complexes14,15. Mitochondrial dysfunctions can be assessed in intact cells, isolated mitochondria, or even ex vivo10,11,13,14.
Mitochondrial dysfunctions are closely associated with many pathological and physiological processes. It is therefore important to study mitochondrial physiology and bioenergetics using genetically and experimentally tractable model systems. In this regard, research on Drosophila melanogaster, the fruit fly, has several advantages. This model shares fundamental cellular characteristics and processes with humans, including the use of DNA as genetic material, common organelles, and conserved molecular pathways involved in development, immunity, and cell signaling. In addition, fruit flies have a rapid life cycle, easy maintenance, low cost, high throughput, and fewer ethical concerns, thus constituting an invaluable tool for dissecting complex cellular processes16,17,18,19,20.
Furthermore, a homolog of the PTEN-induced putative kinase 1 (pink1) gene is expressed in D. melanogaster. It plays a crucial role in the removal of damaged mitochondria through the process of mitophagy8. In humans, mutations in this gene predispose individuals to an autosomal recessive familial form of PD associated with mitochondrial dysfunction8,21,22,23. Consequently, the fruit fly is a powerful animal model for studies on the pathophysiology of PD and screening of drug candidates focusing on mitochondrial dysfunction and bioenergetics. Therefore, the present work explains how to analyze mitochondrial function in a model of PD from D. melanogaster using the HRR technique in the OROBOROS with the Substrate-Uncoupler-Inhibitor-Titration (SUIT) protocol.

<table><tbody><tr><td>ADP</td><td>Sigma-Aldrich</td><td>A5285</td><td>Adenosine 5&amp;prime;-diphosphate sodium sal (CAS number 72696-48-1); &amp;ge;95%; molecular weight = 501.31 g/mol.</td></tr><tr><td>&amp;Aacute;gar</td><td>Kasv</td><td>K25-1800</td><td>For bacteriologal use</td></tr><tr><td>Antimycin-A</td><td>Sigma-Aldrich</td><td>A8674</td><td>Antimycin A from Streptomyces sp. (CAS number 1397-94-0); molecular weight&amp;nbsp; 540 g/mol;</td></tr><tr><td>Bovine Serum Albumin (BSA)</td><td>Sigma-Aldrich</td><td>A7030</td><td>Bovine Serum Albumin (CAS number 9048-46-8); pH 7,0 &amp;ge; 98%</td></tr><tr><td>Datlab software</td><td>Oroboros Instruments, Innsbruck, Austria</td><td>20700</td><td>Software for data acquisition and analysis</td></tr><tr><td>Digitonin</td><td>Sigma-Aldrich</td><td>D 5628</td><td>CAS number 11024-24-1</td></tr><tr><td>Distilled water</td><td></td><td></td><td></td></tr><tr><td>&lt;em&gt;Drosophila melanogaster &lt;/em&gt;strain w[*] Pink1[B9]/FM7i, P{w[+mC]=ActGFP}JMR3</td><td></td><td></td><td>Obtained from Bloomington Drosophila stock center</td></tr><tr><td>&lt;em&gt;Drosophila melanogaster&lt;/em&gt; strain&amp;nbsp;&lt;em&gt;w&lt;sup&gt;1118&lt;/sup&gt;&lt;/em&gt;</td><td></td><td></td><td>Obtained&amp;nbsp; from the Federal University of Santa Maria</td></tr><tr><td>EGTA</td><td>Sigma-Aldrich</td><td>E8145</td><td>Ethylene glycol-bis(2-aminoethylether)-N,N,N&#039;,N&#039;-tetraacetic acid (CAS number 13638-13-3); &amp;ge;97%; molecular weight =468.28 g/mol</td></tr><tr><td>FCCP</td><td>Sigma-Aldrich</td><td>C2920</td><td>Carbonyl cyanide 4- (trifluoromethoxy)phenylhydrazone&amp;nbsp; (CAS number 370-86-5); &amp;ge;98% (TLC), powder&amp;nbsp;</td></tr><tr><td>GraphPad Prism version 8.0.1.</td><td></td><td></td><td>Software for data acquisition and analysis</td></tr><tr><td>Hepes</td><td>Sigma-Aldrich</td><td>H4034</td><td>4-(2-Hydroxyethyl)piperazine-1-ethanesulfonic acid (CAS number 7365-45-9); &amp;ge;99,5% (titration), cell cultured tested; molecular weight = 238.30 g/mol</td></tr><tr><td>High-resolution respirometer Oxygraph O2K</td><td>Oroboros Instruments, Innsbruck, Austria</td><td>10022-02</td><td>Startup O2K respirometer kit</td></tr><tr><td>KH&lt;sub&gt;2&lt;/sub&gt;PO4</td><td>Sigma-Aldrich</td><td>P5379</td><td>Monopotassium phosphate (CAS number 7778-77-0); Reagente Plus, molecular weigt = 136.09 g/mol</td></tr><tr><td>KOH</td><td>Sigma-Aldrich</td><td>211473</td><td>Potassium hydroxide (CAS number 1310-58-3); ACS reagent, &amp;ge;85%, pellets</td></tr><tr><td>Malate</td><td>Sigma-Aldrich</td><td>M1296</td><td>Malonic acid (CAS number 141-82-2); 99%, molecular weight = 104.06 g/mol). A solution is pH adjusted to approximately 7.0.</td></tr><tr><td>Malic acid</td><td>Sigma-Aldrich</td><td>M1000</td><td>(S)-(&amp;minus;)-2-Hydroxysuccinic acid (CAS number 97-67-6); &amp;ge;95% ; molecular weight = 134.09 g/mol</td></tr><tr><td>MES</td><td>Sigma-Aldrich</td><td>M3671</td><td>2-(N-Morpholino)ethanesulfonic acid (CAS number 4432-31-9); &amp;ge;99% (titration); molecular weight = 195.24 g/mol</td></tr><tr><td>MgCl&lt;sub&gt;2&lt;/sub&gt;</td><td>Sigma-Aldrich</td><td>M8266</td><td>Magnesium chloride (CAS number 7786-30-3); anhydrous, &amp;ge;98%, molecular weight = 95.21 g/mol</td></tr><tr><td>Microcentrifuge tubes</td><td>Eppendorf</td><td></td><td></td></tr><tr><td>O2K-Titration Set</td><td>Oroboros Instruments, Innsbruck, Austria</td><td>20820-03</td><td>Hamilton syringes with different volumes</td></tr><tr><td>Oligomycin</td><td>Sigma-Aldrich</td><td>O 4876</td><td>Oligomycin from&amp;nbsp;Streptomyces diastatochromogenes (CAS number&amp;nbsp; 1404-19-9); &amp;ge;90% total oligomycins basis (HPLC)</td></tr><tr><td>Pistil to homogenization</td><td></td><td></td><td></td></tr><tr><td>Proline</td><td>Sigma-Aldrich</td><td>P0380</td><td>L-Proline (CAS number 147-85-3); powder; 99%; molecular weight = 115.13 g/mol</td></tr><tr><td>Pyruvate</td><td>Sigma-Aldrich</td><td>P2256</td><td>Sodium pyruvate (CAS number 113-24-6), &amp;ge;99%; molecular weight = 110.04 g/mol</td></tr><tr><td>Rotenone</td><td>Sigma-Aldrich</td><td>R8875</td><td>Rotetone (CAS number 83-79-4); &amp;ge;95%, molecular weight 394.42 g/ mol</td></tr><tr><td>Succinate</td><td>Sigma-Aldrich</td><td>S 2378</td><td>Sodium succinate dibasic hexahydrate (CAS number 6106-21-4); &amp;ge;99%</td></tr><tr><td>Sucrose</td><td>Merck</td><td>107,651,000</td><td>Sucrose for microbiology use (CAS number 57-50-1)</td></tr><tr><td>Taurine</td><td>Sigma-Aldrich</td><td>T0625</td><td>CAS number 107-35-7</td></tr></tbody></table>

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.

Here, we that O2 flux in OXPHOS CI (P = 0.0341) and OXPHOS CI&#38;II (P = 0.0392) states is reduced in PINK1B9&#160;null flies when compared to control flies (Figure 4). This result was also observed in previous findings from our group29,30.
CI and CII are key components of the electron transport system (ETS), in which CI is responsible for the transfer of electrons from NADH to ubiquinone, while CII transfers electrons from succinate to ubiquinone31,32,33. PINK1B9 null flies showed lower O2 flux in ETS CI (P = 0.0338), ETS CII (P = 0.0457), and ETS CI&#38;II (P = 0.0247) states (Figure 5). These results indicate that the electron transfer system is impaired in flies lacking the pink1 gene and the O2 flux in both OXPHOS and ETS stages are dependent on CI and CI&#38;CII, consistent with other works demonstrating reduced CI activity in pink1-/- models33,34,35.
Furthermore, the proton gradient across the mitochondrial inner membrane is essential for the synthesis of ATP28. The decrease in O2 flux in ETS CI and ETS CII indicates a disruption in the flux of electrons along the ETS. This disruption in the flux of electrons affects the OXPHOS process leading to reduced ATP synthesis. There was also a significant decrease in O2 consumption related to ATP synthesis (P = 0.0280) in PINK1B9 null flies when compared to control flies (Figure 6B). A decrease in ATP synthesis in D. melanogaster can have significant effects on energy metabolism, cellular processes, and overall physiological functions. In addition, the efficiency of the OXPHOS process can be quantified by an index known as RCR, which reflects the tightness of the coupling between respiration and phosphorylation. Therefore, RCR reduction (P = 0.0432) indicates mitochondrial uncoupling, which may affect the OXPHOS process suggesting that the mitochondria are less efficient at utilizing oxygen and producing ATP (Figure 6C). These results may impact the fruit fly&#39;s growth, development, locomotion, reproduction, and overall health and contribute to the pathogenesis of certain neurodegenerative diseases, including PD8,28,29,32.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/65664/65664fig01.jpg" /> 
Figure 1: Layouts in OROBOROS Dat.Lab software. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/65664/65664fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/65664/65664fig02.jpg" /> 
Figure 2: Steps of calibration of chambers in OROBOROS Dat.Lab software. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/65664/65664fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/65664/65664fig03.jpg" /> 
Figure 3: SUIT protocol demonstrating the main points of substrate and inhibitor addition. Firstly, digitonin (DIG) is added followed by complex I specific substrates: malate, pyruvate, and proline (MPP), ATP synthase substrate (ADP), and then, substrate for complex II: succinate (S). Subsequently, the ATP synthase inhibitor: oligomycin (OMY) is added, followed by the uncoupler Carbonyl-4-(trifluoromethoxy) phenylhydrazone cyanide (F), and complex I, II, and III inhibitors: rotenone (R), malonate (MNA), and antimycin (AMA)36. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/65664/65664fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/65664/65664fig04.jpg" /> 
Figure 4: High-resolution respirometry comparing w1118 and PINK1B9 flies. (A) OXPHOS CI, (B) OXPHOS CII, and (C) OXPHOS CI&#38;CII. Data are presented as mean &#177; S.E.M. and analyzed using t-test. n = 5-9. p &#60; 0.05. Abbreviations: OXPHOS = oxidative phosphorylation; CI = complex I; CII = complex II. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/65664/65664fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/65664/65664fig05.jpg" /> 
Figure 5: High-resolution respirometry comparing w1118 and PINK1B9&#160;flies. (A) ETS CI, (B) ETS CII, and (C) ETS CI&#38;CII. Data are presented as mean &#177; S.E.M. and analyzed by t-test. n = 5-9. p &#60; 0.05. Abbreviations: ETS = electron transport system; CI = complex I; CII = complex II. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/65664/65664fig05large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 6" class="xfigimg" src="/files/ftp_upload/65664/65664fig06.jpg" /> 
Figure 6: High-resolution respirometry comparing w1118 and PINK1B9 flies. (A) LEAK, (B) ATP synthesis, and (C) respiratory control ratio. Data are presented as mean &#177; S.E.M. and analyzed by t-test. n = 5-9. p &#60; 0.05. Abbreviation: RCR = respiratory control ratio. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/65664/65664fig06large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Exploring Mitochondrial Function and Chemical Toxicity Using Drosophila melanogaster at JoVE.com

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.

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 ...

analyzing-mitochondrial-function-drosophila-melanogaster-pink1b9-null

Nanyang Technological University

Research

JoVE Journal

Biochemistry

859 Views.  Federal University of Santa Maria. 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.

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

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 ...

Preparation of Mitochondrial Enriched Fractions for Metabolic Analysis in Drosophila

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

Preparation of Mitochondrial Enriched Fractions for Metabolic Analysis in Drosophila

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

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

Developmental Biology

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

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

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

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

A Colorimetric Assay of Citrate Synthase Activity in Drosophila Melanogaster

Mitochondria play the most prominent roles in cellular metabolism by producing ATP through oxidative phosphorylation and regulating a variety of physiological processes. Mitochondrial dysfunction is a primary cause of a number of metabolic and neurodegenerative diseases. Intact mitochondria are critical for their proper functioning. The enzyme citrate synthase is localized in the mitochondrial matrix and thus can be used as a quantitative enzyme marker of intact mitochondrial mass. Given that many molecules and pathways that have important functions in mitochondria are highly conserved between humans and Drosophila, and that an array of powerful genetic tools are available in Drosophila, Drosophila serves as a good model system for studying mitochondrial function. Here, we present a protocol for fast and simple measurement of citrate synthase activity in tissue homogenate from adult flies without isolating mitochondria. This protocol is also suitable for measuring citrate synthase activity in larvae, cultured cells, and mammalian tissues.

A Colorimetric Assay of Citrate Synthase Activity in Drosophila Melanogaster

We present a protocol for a colorimetric assay of citrate synthase activity for quantification of intact mitochondrial mass in Drosophila tissue homogenates.

Mitochondria play the most prominent roles in cellular metabolism by producing ATP through oxidative phosphorylation and regulating a variety of ...

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

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 ...

Measurement of Carbon Dioxide Production from Radiolabeled Substrates in Drosophila melanogaster

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

Measurement of Carbon Dioxide Production from Radiolabeled Substrates in Drosophila melanogaster

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

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

Biology

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 ...

Real-Time Quantification of Mitochondrial Respiration in Muscle Fibers

Measurement of Mitochondrial Respiration in Human and Mouse Skeletal Muscle Fibers by High-Resolution Respirometry

Mitochondrial function, a cornerstone of cellular energy production, is critical for maintaining metabolic homeostasis. Its dysfunction in skeletal muscle is linked to prevalent metabolic disorders (e.g., diabetes and obesity), muscular dystrophies, and sarcopenia. While there are many techniques to evaluate mitochondrial content and morphology, the hallmark method to assess mitochondrial function is the measurement of mitochondrial oxidative phosphorylation (OXPHOS) by respirometry. Quantification of mitochondrial OXPHOS provides insight into the efficiency of mitochondrial oxidative energy production and cellular bioenergetics. A high-resolution respirometer provides highly sensitive, robust measurements of mitochondrial OXPHOS in permeabilized muscle fibers by measuring real-time changes in mitochondrial oxygen consumption rate. The use of permeabilized muscle fibers, as opposed to isolated mitochondria, preserves mitochondrial networks, maintains mitochondrial membrane integrity, and ultimately allows for more physiologically relevant measurements. This system also allows for the measurement of fuel preference and metabolic flexibility - dynamic aspects of muscle energy metabolism. Here, we provide a comprehensive guide for mitochondrial OXPHOS measurements in human and mouse skeletal muscle fibers using a high-resolution respirometer. Skeletal muscle groups are composed of different fiber types that vary in their mitochondrial fuel preference and bioenergetics. Using a high-resolution respirometer, we describe methods for evaluating both aerobic glycolytic and fatty acid substrates to assess fuel preference and metabolic flexibility in a fiber-type-dependent manner. The protocol is versatile and applicable to both human and rodent muscle fibers. The goal is to enhance the reproducibility and accuracy of mitochondrial function assessments, which will improve our understanding of an organelle important to muscle health.

Author Spotlight: Unveiling Mitochondrial Function and Cellular Metabolic Adaptation in Metabolic Diseases

Here, we describe a comprehensive method for measuring mitochondrial oxidative phosphorylation in fresh permeabilized skeletal muscle fibers from either human or mouse muscle. This method allows for the real-time quantification of mitochondrial respiration and the assessment of fuel preference and metabolic flexibility while preserving existing mitochondrial networks and membrane integrity.

Mitochondrial function, a cornerstone of cellular energy production, is critical for maintaining metabolic homeostasis. Its dysfunction in skeletal muscle ...

Analyzing Mitochondrial Function in a Drosophila melanogaster PINK1^B9-Null Mutant Using High-resolution Respirometry

Summary

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Chapters in this video

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

Preparation of Mitochondrial Enriched Fractions for Metabolic Analysis in Drosophila

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

A Colorimetric Assay of Citrate Synthase Activity in Drosophila Melanogaster

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

Metabolic Analysis of Drosophila melanogaster Larval and Adult Brains

Measuring O₂ Consumption in Drosophila melanogaster Using Coulometric Microrespirometry

Measurement of Carbon Dioxide Production from Radiolabeled Substrates in Drosophila melanogaster

Studying Mitochondrial Structure and Function in Drosophila Ovaries

Measurement of Mitochondrial Respiration in Human and Mouse Skeletal Muscle Fibers by High-Resolution Respirometry