Name: Unveiling Xenobiotic Transport and Effects in Isolated Mitochondria: Insights from Respirometric and Enzymatic Assays (Video) | JoVE Journal
Uploaded: 2025-03-07T12:20:00.000+00:00
Duration: 8 min 3 s
Description: Discover respirometric and enzymatic assays to evaluate xenobiotic effects on mitochondrial function in rats, mosquitoes, and ticks. This method supports bio-safe pesticide development by identifying mitochondrial bioenergetics disruptions.

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5 minutes

The authors thank the &#34;Vicerrector&#237;a de Investigaci&#243;n y Extensi&#243;n VIE&#34; of the Universidad Industrial de Santander for financing the grant number 3739 corresponding to the project &#34;Formulation of an ixodicide and ixorepellent designed in a laboratory and validated in the field to prevent tick infestation in cattle.&#34;

All methods described in this protocol have been approved by the Ethics Committee &#34;Comit&#233; de &#233;tica en investigaci&#243;n cient&#237;fica CEINCI&#34;: Scientific Research Ethics Committee of the Industrial University of Santander. Furthermore, these assays complied with the Arrive guidelines and followed the 1989 Colombian Law 84 (Chapter IV, Art. 23-26) and Resolution 8430 (1993, Title IV, Art. 83-93) that regulates animal research in Colombia.
NOTE: It is strongly recommended to read the oxidative phosphorylation chapter of any biochemistry book before starting this protocol.
1. Reagents and solutions
<ol>
	<li>To isolate rat liver mitochondria, prepare 250 mL of mitochondria isolation buffer: 250 mM D-mannitol, 10 mM HEPES pH 7.2, and 1 mM EGTA. Take out 50 mL of isolation buffer (this will be called isolation buffer without BSA) and then add 0.2 g of BSA to the other 200 mL (this will be called isolation buffer with BSA).</li>
	<li>To homogenize Aedes aegypti larvae, prepare 100 mL of buffer: 250 mM sucrose, 10 mM HEPES, 1 mM EGTA pH 7.2.</li>
	<li>Prepare reaction buffer: 80 mM Sodium Phosphate Buffer solution, pH 7.4, 50 &#181;M EDTA.</li>
	<li>Prepare a stock of reagents used for enzymatic activities at pH 7.2 as described below: 25 mM NADH (freshly prepared for each assay), 2 M sodium succinate, 4 mM cytochrome c (freshly prepared for each assay), 200 &#181;M rotenone, 0.5 mM KCN, 6 mM 2,6-dichlorophenolindophenol (DCPIP), 100 mM phenazine methosulfate (PMS), and 60 mM potassium ferricyanide. 
	NOTE: Ultrapure water with a resistivity of 18.2 M&#937;&#183;cm at 25 &#176;C was used for all experiments and reagents preparation. This water was generated using a Milli-Q purification system, ensuring the removal of organic, inorganic, and microbial contaminants to levels below detectable limits, making it suitable for high-sensitivity biochemical and molecular biology applications.</li>
</ol>
2. Liver dissection, homogenization, and mitochondria isolation from rat liver
<ol>
	<li>Before starting, place all materials and buffers on ice, ensuring all materials are ice-cold during the entire assay to avoid mitochondrial damage.</li>
	<li>Place 10 mL of mitochondria isolation buffer in an ice-cold conic tube.</li>
	<li>Euthanize the male Wistar rat by cervical decapitation, avoiding CO2 use that can interfere with respiration assays. For this, the Committee on Ethics in Scientific Research of the Industrial University of Santander (CEINCI) approved the methods on resolution 13 of 2022. 
	NOTE: Rats should weigh no more than 170 g; fat can also affect respiration analyses.</li>
	<li>Make an incision (approx 4-5 cm) with a sterile scalpel through the abdominal rat section.</li>
	<li>Dissect the liver completely once the abdominal area is visible. The rat liver is a multilobulated organ, dark red approx 10 cm x 5 cm.</li>
	<li>Carefully put the liver within the ice-cold isolation buffer and rinse twice with this medium.</li>
	<li>Finely mince the liver (size less than 1 cm) with sharp scissors in a Petri dish with 10 mL of isolation medium.</li>
	<li>Put the solution into the Potter Elvehjem tissue homogenizer tube and carefully homogenize at least ten times, low velocity (~390 rpm).</li>
	<li>Transfer the solution into a conical tube and centrifuge at 600 x g for 10 min at 4 &#176;C.</li>
	<li>After centrifugation, carefully pour the supernatant into a new conical tube. Discard the pellet with part of the tissue, blood, and intact cellular components.</li>
	<li>Fill the tube with more isolation buffer (~35 mL) and centrifuge at 6000 x g for 10 min at 4 &#176;C.</li>
	<li>After centrifugation, discard the supernatant and carefully resuspend the pellet (mitochondria) in a new isolation buffer without BSA. 
	NOTE: For mitochondrial resuspension, it&#39;s recommended to use a soft brush.</li>
	<li>Then, centrifuge at 7000 x g for 10 min at 4 &#176;C, discard the supernatant and resuspend the isolated mitochondria in 2 mL of isolation buffer without BSA.</li>
	<li>Determine protein concentration using the Bradford method or similar14.</li>
</ol>
3. Aedes aegypti colony and larvae homogenization.
NOTE: For this procedure Ae. aegypti colony was kept inside breeding cages (40 cm &#215; 40 cm &#215; 40 cm) under insectary conditions like humidity (70% &#177; 5%), photoperiod (12:12), and temperature (27 &#177; 5 &#176;C). The eggs were collected on filter paper.
<ol>
	<li>Ensure that the eggs are hatched in trays with filtered and dechlorinated water one week before the experiment. Identify the instar by the molting process in the larvae cycle. Observe each molting process to verify its status L1, L2, and L3. When the larva enters L3, wait a maximum of 12 h to collect the material.</li>
	<li>Collect at least 100 Ae. aegypti larvae at instar L3-L4 in chlorine-free water (~30 mL of water). Take the larvae with a Pasteur pipette.</li>
	<li>Filter the larvae using filter paper, rinse the collected larvae, and resuspend them in the Aedes aegypti homogenization buffer (~2 mL).</li>
	<li>Transfer the solution with larvae into the Potter Elvehjem tissue homogenizer tube and carefully homogenize at least ten times at ~390 rpm.</li>
	<li>Transfer the homogenized tissue and filter it through a glass wool syringe to remove large exoskeleton residues.</li>
	<li>Determine the protein concentration by using the Bradford method or similar15. 
	NOTE: It is not necessary to isolate mitochondria from mosquito larvae; it is possible to evaluate the effect of xenobiotics using the protein extract obtained in this step.</li>
</ol>
4. Determination of xenobiotics effects on mitochondrial electron transport chain
NOTE: This protocol was designed to evaluate the effect of xenobiotics on mitochondrial bioenergetics on isolated mitochondria from rat liver and insect tissue homogenized.
The assays are divided into two: polarographic (NADH and Succinate oxidase) and spectrophotometric (NADH dehydrogenase - complex I, succinate dehydrogenase - complex II, cytochrome c reductase - complex III and cytochrome c oxidase - complex IV).
<ol>
	<li>Polarographic assays: For this, use high-resolution respirometry to measure the complete oxidation of NADH and succinate by O2 consumption, and report results as NADH and succinate oxidase.
	<ol>
		<li>Calibrate the instrument every day before starting assays. This step may vary depending on the oxygraph used. Follow the manufacturer&#39;s instructions.
		<ol>
			<li>For Oroboros calibration, add reaction medium into the chambers with an excess of at least 0.2 mL above the experimental chamber volume (2 mL), to fill the chamber and injection capillary with medium.</li>
		</ol>
		</li>
		<li>Switch on the stirrers after medium addition.</li>
		<li>Wait until the oxygen concentration signal stabilizes and record it.</li>
		<li>For zero oxygen calibration, add mitochondria, cell suspensions, baker&#39;s yeast, or dithionite (Baker&#39;s yeast is recommended) that allow the oxygen depletion, and record it. Wait for the signal to reach background levels (typically &#177; 2 pmol/mL/s for the O2k system).</li>
		<li>Perform NADH and succinate oxidase activities by monitoring the oxygen consumption when adding reaction buffer, 0.1 mg of protein (mitochondrial or larvae homogenized), and the substrate (0.17 mM NADH or 10 mM sodium succinate) to the chamber.</li>
		<li>Use electron transport protein inhibitors to confirm that the observed oxygen consumption is due to the mitochondrial respiratory chain. The common ones are rotenone 1 &#181;M and KCN 1mM16. In the oxygraph cell, add the reagents according to Table 1&#160;(the sequence is important).</li>
	</ol>
	</li>
	<li>Spectrophotometric assays 
	NOTE: To assess electron transport through the mitochondrial complexes, enzymatic assays were conducted and measured using a spectrophotometer and artificial electron acceptors such as PMS, DCPIP, and cytochrome c. It is recommended to measure one enzyme per day. Prepare buffers and reagents at least one day before the assays. Some compounds are photosensible; see Table 2 for more recommendations.
	<ol>
		<li>For NADH dehydrogenase activity, the reaction systems contain 50 mM phosphate buffer solution pH 7.4, 2 mM EDTA, 0.1 mg/mL mitochondrial protein, 1 &#181;M rotenone, and 1 mM KCN. In the spectrophotometer cell, add the reagents according to Table 2&#160;(the sequence is important)17. The recommended steps are as follows:
		<ol>
			<li>Prepare 100 mL of buffer phosphate mixed with the EDTA, and double-check that the pH does not change. Keep the buffer in the refrigerator, but it is mandatory to keep it at room temperature (RT; approximately 25 &#176;C) during assays.</li>
			<li>Prepare fresh stock solutions of rotenone, KCN, NADH, and potassium ferricyanide (stable until 4 h). The suggested volume is 1 mL of concentrated solutions (see the first column of the table).</li>
			<li>If the tested compound has low solubility in water, include an additional control for enzyme activity in the experimental design using just the solvent of the compound.</li>
			<li>First, monitor the NADH dehydrogenase activity by the reduction of potassium ferricyanide at 420 nm. Calculate its concentration by its molar extinction coefficient of 1.04 mM-1&#183;cm-1. The spectrophotometer has an option to calculate the maximum activity in a reaction; check the user manual. Subsequently, divide the reported activity by the concentration of protein used to obtain the specific activity of the enzyme.</li>
		</ol>
		</li>
		<li>Evaluate succinate dehydrogenase activity by adding 20 mM sodium succinate and 1 mM phenazine methosulphate (PMS) with 60 &#181;M 2,6-dichlorophenolindophenol (DCPIP) as artificial electron acceptors. In the spectrophotometer cell, add the reagents according to Table 3&#160;(the sequence is important)17. The recommendations are as follows:
		<ol>
			<li>Prepare the same buffer as used for NADH dehydrogenase. Maintain it in the same conditions.</li>
			<li>Prepare fresh stock solutions of Succinate, PMS, and DCPIP. The suggested volume is 1 mL of concentrated solutions. Prepare PMS and DCPIP solutions on the same day of the assay.</li>
			<li>If the tested compound has low solubility in water, include an additional control for enzyme activity in the experimental design.</li>
			<li>Monitor the succinate oxidation by the reduction of DCPIP at 600 nm. Calculate its concentration by its molar extinction coefficient of 21.0 mM&#8722;1&#183;cm&#8722;1, applying Beer&#39;s law. Subsequently, divide the reported activity by the concentration of protein used to obtain the specific activity of the enzyme.</li>
		</ol>
		</li>
		<li>Measure the Cytochrome c reductase (complex III) activity using either NADH or succinate as oxidizable substrates. With this enzyme, the electron transport from complex I (using NADH) or complex II (using succinate) passes to ubiquinone and then to complex III, which passes the electrons to cytochrome c. This reaction is followed by the reduction of cytochrome c. Prepare the reaction system containing 50 mM phosphate buffer solution pH 7.4, 2 mM EDTA, 0.1 mg of mitochondrial protein, 50 &#181;M NADH, and 0.1 mM KCN. In the spectrophotometer cell, add the reagents according to Table 4&#160;(the sequence is important)18. The recommendations are as follows:
		<ol>
			<li>Prepare the same buffer as used for NADH dehydrogenase. Maintain it in the same conditions.</li>
			<li>Use the same NADH and KCN from NADH dehydrogenase.</li>
			<li>If the tested compound has low solubility in water, include an additional control for enzyme activity in the experimental design.</li>
			<li>Monitor the cytochrome c reduction at 550 nm and calculate its concentration by its molar extinction coefficient of 19.8 mM&#8722;1&#183;cm&#8722;1, applying Beer&#39;s law. Subsequently, divide the reported activity by the concentration of protein used to obtain the specific activity of the enzyme.</li>
		</ol>
		</li>
		<li>Measure cytochrome c oxidase (complex IV) expressed as cytochrome c oxidase activity. For this, prepare a system reaction containing 50 mM phosphate buffer solution, pH 7.4, 2 mM EDTA, and 0.1 mg of mitochondrial protein (Table 5). For these enzymatic assays, reduce cytochrome c by mixing equimolar amounts of cytochrome c and sodium dithionite, ensuring complete reduction of the cytochrome c.</li>
		<li>Then, pass the resulting mixture through a cross-linked dextran gel column using the same phosphate-buffered solution as the mobile phase. Subsequently, quantify the collected fractions using a spectrophotometer at 550 nm19. The recommendations are as follows:
		<ol>
			<li>Use the same buffer used for NADH dehydrogenase. Maintain it in the same conditions.</li>
			<li>If the tested compound has low solubility in water, an additional control for enzyme activity should be included in the experimental design.</li>
			<li>Monitor the cytochrome c oxidation at 550 nm and calculate its concentration by its molar extinction coefficient of 19.8 mM&#8722;1&#183;cm&#8722;1, applying Beer&#39;s law. Subsequently, divide the reported activity by the concentration of protein used to obtain the specific activity of the enzyme. 
			NOTE: A spreadsheet file showing the process of calculating the specific activity of enzymes, measured by oxygen consumption and spectrophotometry, is included as Supplementary File 1.</li>
		</ol>
		</li>
	</ol>
	</li>
</ol>

Changing Mitochondrial Function for Therapeutic and Bioinsecticide Applications

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<li>Valmas, N., Zuryn, S., Ebert, P. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Mitochondrial+uncouplers+act+synergistically+with+the+fumigant+phosphine+to+disrupt+mitochondrial+membrane+potential+and+cause+cell+death%5BTitle%5D)+AND+%22Toxicology%22%5BJournal%5D)">Mitochondrial uncouplers act synergistically with the fumigant phosphine to disrupt mitochondrial membrane potential and cause cell death.</a> Toxicology. 252 (1-3), 33-39 (2008).</li>
<li>Slocińska, M., Rosinski, G., Jarmuszkiewicz, W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Activation+of+mitochondrial+uncoupling+protein+4+and+ATP-sensitive+potassium+channel+cumulatively+decreases+superoxide+production+in+insect+mitochondria%5BTitle%5D)+AND+%22Protein+Pept+Lett%22%5BJournal%5D)">Activation of mitochondrial uncoupling protein 4 and ATP-sensitive potassium channel cumulatively decreases superoxide production in insect mitochondria.</a> Protein Pept Lett. 23 (1), 63-68 (2016).</li>
<li>Borrero Landazabal, M. A., Carreño Otero, A. L., Kouznetsov, V. V., Duque Luna, J. E., Mendez-Sanchez, S. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Alterations+of+mitochondrial+electron+transport+chain+and+oxidative+stress+induced+by+alkaloid-like+%CE%B1-aminonitriles+on+Aedes+aegypti+larvae%5BTitle%5D)+AND+%22Pestic+Biochem+Physiol%22%5BJournal%5D)">Alterations of mitochondrial electron transport chain and oxidative stress induced by alkaloid-like α-aminonitriles on Aedes aegypti larvae.</a> Pestic Biochem Physiol. 144, 64-70 (2018).</li>
<li>Castillo-Morales, R. M., Carreño Otero, A. L., Mendez-Sanchez, S. C., Da Silva, M. A. N., Stashenko, E. E., Duque, J. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Mitochondrial+affectation%2C+DNA+damage+and+AChE+inhibition+induced+by+Salvia+officinalis+essential+oil+on+Aedes+aegypti+larvae%5BTitle%5D)+AND+%22Comp+Biochem+Physiol+C+Toxicol+Pharmacol%22%5BJournal%5D)">Mitochondrial affectation, DNA damage and AChE inhibition induced by Salvia officinalis essential oil on Aedes aegypti larvae.</a> Comp Biochem Physiol C Toxicol Pharmacol. 221, 29-37 (2019).</li>
<li>Montagut, D. C., Bueno, Y., Vesga, L. C., Stashenko, E. E., Mendez-Sanchez, S. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Cymbopogon+flexuosus+%28nees+ex+steud.%29+w.+watson+essential+oil+effect+on+mitochondrial+bioenergetics%5BTitle%5D)+AND+%22J+Essent+Oil+Res%22%5BJournal%5D)">Cymbopogon flexuosus (nees ex steud.) w. watson essential oil effect on mitochondrial bioenergetics.</a> J Essent Oil Res. 34 (3), 233-239 (2022).</li>
</ol>

All authors declare that they have no conflicts of interest.

A spectrophotometric and polarographic method with high-resolution O2 was employed to investigate the effects of xenobiotics on mitochondrial function. The study specifically assessed the impact of Cymbopogon flexuosus essential oil and carvacrol on NADH and succinate oxidation, as well as electron transport through the mitochondrial electron transport chain, using two different biological models: isolated mitochondria from rat liver and homogenized Aedes aegypti larvae. Additionally, potenti...

Mitochondria have always been called the power plant of cells due to their chemical energy generation during the oxidative phosphorylation process1,2. Beyond their main core metabolic functions, mitochondria are also implicated in many other cellular processes, ranging from intracellular Ca2+ regulation levels, inflammatory processes, and reactive oxygen species production to cell death induction processes3. Mitochondrial bioenergetics and their dysregulation have been associated with intracellular processes that affect cellular function and overall health. This has sparked a growing interest in researching the pharmaceutical potential of targeting mitochondrial function to develop innovative therapies and safe bioproducts4,5,6. Additionally, understanding the intricate interplay of mitochondrial processes and their impact on cellular function is crucial for developing targeted therapies and presenting opportunities for designing bioproducts to control vector-borne diseases and pest infestations7,8.
Targeting specific components of the electron transport chain with xenobiotics has emerged as a strategy to modulate mitochondrial function for biopesticide generation. Natural and synthetic compounds included within pesticide formulations, such as nanoparticle-based mitochondria-targeted pesticides, have been shown to affect mitochondrial function by decreasing mitochondrial potential and ATP synthesis in S. frugiperda9.
Additionally, most of the carbamates and pyrethroid-based products have been reported as complex I inhibitors in mosquitoes; however, their extended exposure has created some pyrethroid-resistant populations in Anopheles gambiae, often related to elevated levels of P450s that can metabolize these substrates10,11. Similarly, plants and their essential oils, extracts, and secondary metabolites have been documented as potential insecticides having a wide range of mortality in different insect stages12,13. The insecticide activity of these natural compounds may be related to the electron transport inhibition in a specific point in the mitochondrial respiratory chain by modulating the reactive oxygen species production, mitochondrial potential &#916;&#936;m, and ATP synthesis that can trigger cell death processes6,8.
Natural compounds, as sources for pesticide formulation, and their effects on mitochondria play a crucial role in the design of new and safe biopesticides. In this context, this manuscript describes a methodology to identify the capacity of xenobiotics to activate or inhibit electron transport through mitochondrial complexes. This comprehensive approach allows precise measurement of xenobiotics&#39; effects on mitochondrial function using a variety of substrates, employing both respirometry and spectrophotometry methods. This methodology has been applied to analyze the mitochondrial function of key vector species, including Aedes aegypti, leading to theorizing the presence of uncoupler proteins and distinct oxidizable substrate preferences specific to each species.

<table><tbody><tr><td>2,6-dichlorophenolindophenol</td><td>Merck</td><td>119814</td><td>Prepare on the same day of use.</td></tr><tr><td>Bradford reagent</td><td>Merck</td><td>B6916</td><td></td></tr><tr><td>BSA</td><td>Merck</td><td>A7030</td><td></td></tr><tr><td>Carvacrol</td><td>Merck</td><td>W224511</td><td></td></tr><tr><td>Cytochrome c</td><td>Merck</td><td>C2506</td><td>Photosensitive conver cover it with aluminum foil. In solution it should be stored in the freezer.</td></tr><tr><td>D-mannitol</td><td>Merck</td><td>M4125</td><td></td></tr><tr><td>EGTA</td><td>Merck</td><td>&gt;E3889</td><td></td></tr><tr><td>Ethylenediaminetetraacetic acid tetrasodium salt dihydrate</td><td>Merck</td><td>E6511</td><td></td></tr><tr><td>HEPES</td><td>Merck</td><td>H3375</td><td></td></tr><tr><td>KCN</td><td>Merck</td><td>60178</td><td>In solution works weel for 2 weeks.</td></tr><tr><td>KH&lt;sub&gt;2&lt;/sub&gt;PO&lt;sub&gt;4&lt;/sub&gt;</td><td>Merck</td><td>P5655</td><td></td></tr><tr><td>Multiskan FC Microplate Photometer</td><td>Thermo Scientific&amp;nbsp;</td><td>51119100</td><td></td></tr><tr><td>Na&lt;sub&gt;2&lt;/sub&gt;HPO&lt;sub&gt;4&lt;/sub&gt;</td><td>Merck</td><td>567550</td><td></td></tr><tr><td>Oroboros O2k&amp;nbsp;</td><td>Oroboros Instruments</td><td>10203-03</td><td>https://www.oroboros.at/index.php/product/o2k-startup/</td></tr><tr><td>Phenazine methosulphate&amp;nbsp;</td><td>Merck</td><td>P9625</td><td>Prepare on the same day of use.</td></tr><tr><td>Potassium ferricyanide&amp;nbsp;</td><td>Merck</td><td>244023</td><td>Prepare on the same day of use.</td></tr><tr><td>Potter Elvehjem tissue homogenizer tube</td><td></td><td></td><td></td></tr><tr><td>Rotenone</td><td>Merck</td><td>557368</td><td>Soluble in ethanol. Photosensitive conver cover it with aluminum foil. In solution it should be stored in the freezer.</td></tr><tr><td>Sodium succinate</td><td>Merck</td><td>S9637</td><td>In solution it should be stored in the freezer.</td></tr><tr><td>Sucrose</td><td>Merck</td><td>S0389</td><td></td></tr><tr><td>&amp;beta;-Nicotinamide adenine dinucleotide, reduced Disodium salt hydrate</td><td>Merck</td><td>N8129</td><td>Photosensitive conver cover it with aluminum foil. In solution it should be stored in the freezer.</td></tr></tbody></table>

unveiling xenobiotic transport and effects in isolated mitochondria: insights from respirometric and enzymatic assays

Unveiling Xenobiotic Transport and Effects in Isolated Mitochondria: Insights from Respirometric and Enzymatic Assays

Mitochondria are often referred to as the cell's powerhouse due to their crucial role in energy production through the electron transport chain (ETC). However, their significance extends far beyond energy production. Dysregulation of mitochondrial bioenergetics can trigger intracellular cascades that impact health and cellular function. Given their physiological importance, there is growing interest in exploring the pharmacological potential of mitochondrial function for developing new therapies and bioproducts. Modulating mitochondrial bioenergetics could provide innovative approaches to address challenges such as neurodegenerative disorders, metabolic diseases, and cancer, as well as to develop bioproducts for controlling pests and vector-borne diseases.
This work presents a method for assessing the effects of xenobiotics on the electron transport chain (ETC) using various substrates, including glutamate and NADH for complex I, succinate for complex II, and cytochrome c (both oxidized and reduced) for complexes III and IV. This methodology allows the activation or inhibition of electron transport through mitochondrial complexes to be evaluated using a respirometer and spectrophotometer. The mitochondria can be sourced from isolated mitochondria, fragmented cells, or homogenized tissue from various species.
In the laboratory, mitochondrial function has been analyzed in Aedes aegypti and Wistar rats, but this method is also applicable to other species, such as Rhipicephalus microplus and Rhodnius prolixus. This approach provides the basis for theorizing about the existence of uncoupler proteins and species-specific oxidizable substrate preferences influenced by their unique energetic demands. Recent findings offer valuable insights into innovative bioinsecticide design strategies that target mitochondrial function, holding significant potential for effectively controlling vector-borne diseases and pest infestations.

Before analyzing the results obtained with this protocol, it is essential to remember a little about the biochemistry of the electron transport chain in the mitochondria. First, NADH supplies a pair of electrons to the oxidoreduction centers located in complex I. There, it begins the transport of electrons that pass from complex I to ubiquinone, subsequently to complex III, which donates them to cytochrome c, and finally to complex IV, where oxygen is the final electron acceptor, reducing it to water. On the other hand, ...

This study introduces a methodology for studying xenobiotic effects on mitochondrial function via respirometric and enzymatic assays. It examines electron transport chain modulation in various species, offering insights for targeted biopesticide development against insect vector-borne diseases.

Mitochondria are often referred to as the cell's powerhouse due to their crucial role in energy production through the electron transport chain (ETC). ...

unveiling-xenobiotic-transport-effects-isolated-mitochondria-insights

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JoVE Journal

Biochemistry

Comparison of mitochondrial membrane potential between samples yields valuable information about cellular status. Detailed steps for isolating mitochondria and assessing response to inhibitors and uncouplers using fluorescence are described. The method and utility of this protocol are illustrated by use of a cell culture and animal model of cellular stress.

Isolation and Functional Analysis of Mitochondria from Cultured Cells and Mouse Tissue

Methods for biopsy of Vastus lateralis, preparation of purified mitochondria, and respirometric profiling are described. The use of small muscle volume makes this technique suitable for clinical research applications.

Preparation and Respirometric Assessment of Mitochondria Isolated from Skeletal Muscle Tissue Obtained by Percutaneous Needle Biopsy

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

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

Here, we present a modification of a previously reported method that allows for the isolation of high quality and purified mitochondria from smaller quantities of mouse skeletal muscle. This procedure results in highly coupled mitochondria that respire with high function during microplate based respirometirc assays.

Isolation of Mitochondria from Minimal Quantities of Mouse Skeletal Muscle for High Throughput Microplate Respiratory Measurements

High-resolution respirometry is used to determine mitochondrial oxygen consumption. This is a straightforward technique to determine mitochondrial respiratory chain complexes' (I-IV) respiratory rates, maximal mitochondrial electron transport system capacity, and mitochondrial outer membrane integrity.

High-resolution Respirometry to Assess Mitochondrial Function in Permeabilized and Intact Cells

Here, a quadriceps muscle specimen is taken from an anaesthetized pig and mitochondria are isolated by differential centrifugation. Then, the respiratory rates of mitochondrial respiratory chain complexes I, II and IV are determined using high-resolution respirometry.

Isolation of Intact Mitochondria from Skeletal Muscle by Differential Centrifugation for High-resolution Respirometry Measurements

In this work, we describe a modified protocol to test mitochondrial respiratory substrate flux using recombinant perfringolysin O  in combination with microplate-based respirometry. With this protocol, we show how metformin affects mitochondrial respiration of two different tumor cell lines.

Measuring Mitochondrial Substrate Flux in Recombinant Perfringolysin O-Permeabilized Cells

Assessing oxidative phosphorylation using high-resolution respirometers has become an integral part of the functional analysis of mitochondria and cellular energy metabolism. Here, we present protocols for the analysis of cellular energy metabolism using chamber and microplate-based high-resolution respirometers and discuss the key benefits of each device.

High-Resolution Respirometry to Assess Bioenergetics in Cells and Tissues Using Chamber- and Plate-Based Respirometers

Here, we describe a detailed method for mitochondria isolation from mouse skeletal muscle and the subsequent analysis of respiration by Oxygen Consumption Rate (OCR) using microplate-based respirometric assays. This pipeline can be applied to study the effects of multiple environmental or genetic interventions on mitochondrial metabolism.

Isolation of Mitochondria from Mouse Skeletal Muscle for Respirometric Assays

These protocols will help users probe mitochondrial energy metabolism in 3D cancer cell-line-derived spheroids using Seahorse extracellular flux analysis.

Exploring Mitochondrial Energy Metabolism of Single 3D Microtissue Spheroids Using Extracellular Flux Analysis

Mitochondrial respiration in yeast whole cells is a valuable indicator of cell bioenergetics. Here, we present a protocol to quantify this phenotype applicable to different yeast species.

Mitochondrial Respiration Quantification in Yeast Whole Cells

Discover respirometric and enzymatic assays to evaluate xenobiotic effects on mitochondrial function in rats, mosquitoes, and ticks. This method supports bio-safe pesticide development by identifying mitochondrial bioenergetics disruptions.

Unveiling Xenobiotic Transport and Effects in Isolated Mitochondria: Insights from Respirometric and Enzymatic Assays

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