The authors acknowledge Mark Nelms and John Tennant from the Electrical and Computer Engineering department of the Samuel Ginn College of Engineering at Auburn University for helping with the structural and electrical outfitting of the AU MitoMobile. Additionally, the authors acknowledge the funding to outfit the AU MitoMobile and research from an Auburn University Presidential Awards for Interdisciplinary Research (PAIR) grant.

The following sections describe the mitochondrial laboratory methods. All animal handling and tissue collection procedures were approved by the Auburn University Institutional Animal Care and Use Committee (#2019-3582).
1. Description of buffers used for data collection
NOTE: These buffers can be prepared in a stationary laboratory and moved to the mobile laboratory prior to the field trip (unless otherwise noted below).
<ol>
	<li>Prepare the skeletal muscle mitochondrial isolation buffer with bovine serum albumin (BSA), as seen in Table 1.
	<ol>
		<li>Dissolve chemicals in deionized water (~ 90% volume) except for the fatty acid-free BSA. Place the buffer in the refrigerator until the temperature is 4 &#176;C.</li>
		<li>Adjust the solution to a pH of 7.5 while maintaining the temperature at 4 &#176;C.</li>
		<li>Add the fatty acid-free BSA and bring up the volume to 100%. Aliquot the solution into 50 mL conical tubes. Store this solution at -20 &#176;C until use.</li>
	</ol>
	</li>
	<li>Prepare the skeletal muscle mitochondrial isolation buffer without BSA as seen in Table 1.
	<ol>
		<li>Dissolve the chemicals in deionized water (~ 90% volume). Place the buffer in the refrigerator until the temperature is 4 &#176;C.</li>
		<li>Adjust the solution to a pH of 7.5 while maintaining the temperature at 4 &#176;C.</li>
		<li>Bring up the volume to 100%. Aliquot the solution into 50 mL conical tubes. Store this solution at -20 &#176;C until use.</li>
	</ol>
	</li>
	<li>Prepare the skeletal muscle resuspension buffer as seen in Table 1.
	<ol>
		<li>Dissolve the chemicals in deionized water (~ 90% volume). Place the buffer in the refrigerator until the temperature is 4 &#176;C.</li>
		<li>Adjust the solution to a pH of 7.4 while maintaining the temperature at 4 &#176;C.</li>
		<li>Bring up the volume to 100%. Aliquot the solution into 50 mL conical tubes. Store this solution at -20 &#176;C until use.</li>
	</ol>
	</li>
	<li>Prepare the skeletal muscle respiration buffer as seen in Table 2.
	<ol>
		<li>Dissolve the chemicals in deionized water (~ 90% volume) except for the fatty acid-free BSA. Heat the buffer until the temperature is 37 &#176;C.</li>
		<li>Adjust the solution to a pH of 7.0 while maintaining the temperature at 37 &#176;C.</li>
		<li>Add the fatty acid-free BSA and bring up the volume to 100%. Aliquot the solution into 50 mL conical tubes. Store this solution at -20 &#176;C until use.</li>
	</ol>
	</li>
	<li>Prepare the respiration substrates as seen in Table 2.
	<ol>
		<li>Ensure that these substrates are made fresh on the day of data collection in 100 mM Tris-HCl, pH 7.4. Store on ice until use. 
		&#8203;NOTE: The provided values are to make a sufficiently concentrated solution for enough substrate to be taken up by the mitochondria. The final concentrations of the substrates are 2 mM pyruvate, 2 mM malate, 10 mM glutamate, and 5 mM succinate.</li>
	</ol>
	</li>
</ol>
2. Performing mitochondrial isolation (Figure 2)
NOTE: Mitochondrial isolation and mitochondrial respiration measurements are performed in the laboratory bench area of the mobile laboratory, and all solutions should be kept at 4 &#176;C unless otherwise noted.
<ol>
	<li>Park the mobile laboratory on flat ground. Turn on the generator and level the vehicle. Extend the slide and set up the equipment.</li>
	<li>Thaw the desired amounts of buffers. 
	NOTE: Generally, 30 mL of skeletal muscle isolation buffer and 10 mL of skeletal muscle isolation buffer without BSA are needed per muscle.</li>
	<li>Set up and calibrate the mitochondrial respiration chambers to the desired temperature of experiments and current barometric pressure per manufacturer&#39;s instructions. See the Table of Materials for specific chambers used in experiments.</li>
	<li>Euthanize the animal via decapitation. 
	NOTE: The current study used decapitation for euthanasia. Some gases, such as carbon dioxide and isoflurane, affect mitochondrial function18,19,20; these effects should be considered when selecting the best method of euthanasia for each study. Which method should be performed for each study will be determined by the scientific question being asked.</li>
	<li>Excise skeletal muscle, quickly trim away fat and connective tissue, weigh, and place the muscle in skeletal muscle isolation buffer with BSA (at least 1/10 w/v) (e.g., 1 g of skeletal muscle to 10 mL of buffer).</li>
	<li>Mince the skeletal muscle with scissors on ice.</li>
	<li>Transfer the minced tissue to a 50 mL centrifuge tube using a cut 5 mL pipet tip. Homogenize it with a blade (see the Table of Materials) at 50% power for 5 s. Add protease (5 mg/g wet muscle) and digest for 7 min, mixing the solution every 30 s. Terminate the reaction by adding an equal volume of isolation buffer with BSA.</li>
	<li>Centrifuge the homogenate at 500 &#215; g for 10 min. Transfer the supernatant through double-layered cheesecloth using a cut 5 mL pipet tip into a clean 50 mL centrifuge tube. Centrifuge the supernatant at 3,500 &#215; g for 10 min to precipitate a brown mitochondrial pellet.</li>
	<li>Pour out the remaining supernatant. Add the same volume of isolation buffer with BSA to the centrifuge tube. Resuspend the mitochondrial pellet with a flexible scraper (policeman) by gently working the mitochondrial pellet off the walls of the centrifuge tube. Centrifuge at 3,500 &#215; g for 10 min.</li>
	<li>Pour out the remaining supernatant. Add the same volume of isolation buffer without BSA to the centrifuge tube. Resuspend the mitochondrial pellet by gently working the mitochondrial pellet off the walls of the centrifuge tube with a clean policeman. Centrifuge at 3,500 &#215; g for 10 min.</li>
	<li>Decant the supernatant and resuspend the mitochondrial pellet in resuspension buffer by gently working the mitochondrial pellet off the walls of the centrifuge tube with a clean policeman. 
	NOTE: The volume of the resuspension buffer will depend on the size of the mitochondria pellet.</li>
	<li>Transfer the resuspended mitochondria to a Dounce homogenizer with a cut 1 mL pipet tip. Using the Dounce homogenizer, carefully homogenize the suspension with 4-5 passes.</li>
	<li>Place the mitochondrial suspension in a labeled 2 mL microcentrifuge tube using another cut 1 mL pipet tip.</li>
</ol>
3. Mitochondrial respiration measurements (Figure 3)
<ol>
	<li>Complex I substrates
	<ol>
		<li>Add 945 &#181;L of respiration buffer to the chamber. Ensure that the stirrer is spinning, and the buffer temperature is maintained at 37 &#176;C. Start the recording of the data collection.</li>
		<li>After the oxygen concentration has stabilized, add 20 &#181;L of the mitochondria and place the lid on the chamber. In the software, denote that mitochondria were added to the chamber.</li>
		<li>Add 10 &#181;L of 1 M glutamate, 10 &#181;L of 200 mM malate, and 10 &#181;L of 200 mM pyruvate to the chamber with individual syringes and wait until the signal stabilizes. In the software, denote that substrates have been added. 
		NOTE: These substrates are typically used to measure carbohydrate-driven respiration. For other combinations of substrates to be used to measure fat-driven respiration, see21.</li>
		<li>Add 5 &#181;L of ADP with a separate syringe and observe the rapid oxygen consumption (state 3). In the software, denote that ADP was added. 
		NOTE: Following the phosphorylation of the added ADP, the oxygen consumption rate will plateau to state 4.</li>
		<li>After 4 min of state 4 data collection, terminate the recording. Save the data file.</li>
	</ol>
	</li>
	<li>Complex II substrates
	<ol>
		<li>Add 963 &#181;L of the respiration buffer to the chamber. Ensure that the stirrer is spinning, and the buffer temperature is maintained at 37 &#176;C. Start the recording of the data collection.</li>
		<li>After the oxygen concentration has stabilized, add 20 &#181;L of mitochondria and place the lid on the chamber. In the software, denote that mitochondria were added to the solution.</li>
		<li>Add 2 &#181;L of 4 &#181;g/&#181;L rotenone followed by 10 &#181;L of 500 mM succinate to the chamber using separate syringes and wait until the signal stabilizes. In the software, denote that substrates have been added.</li>
		<li>Add 5 &#181;L of ADP using a separate syringe and observe the rapid oxygen consumption (state 3). In the software, denote that ADP was added. 
		NOTE: Following the phosphorylation of the added ADP, the oxygen consumption rate will plateau to state 4.</li>
		<li>After 4 min of state 4 data collection, terminate the recording. Save the data file.</li>
	</ol>
	</li>
</ol>

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The authors have no conflicts of interest to declare.

The mobile mitochondrial physiology laboratory enables researchers to isolate mitochondria and measure mitochondrial respiration rates within 2 h of tissue collection at remote field sites. The results presented herein suggest that measurements of mitochondrial respiration made in the AU MitoMobile are comparable to measurements made in a university research laboratory. Specifically, the values for state 3, state 4, and RCR for wild-derived Mus musculus presented here are comparable with previously published res...

To date, studies designed to measure mitochondrial energetics have been limited to laboratory animals or animals captured near established physiology laboratories, which precluded scientists from performing mitochondrial bioenergetic studies in tissues collected from animals during such activities as migration, diving, and hibernation1,2,3,4,5,6. While many investigators have successfully measured the basal and peak metabolic rates and daily energy expenditures of wild animals7,8, the capacity of researchers to measure the performance of mitochondria has remained limited (but see1,4,9). This is partly due to the need for fresh tissue for isolating mitochondria and a laboratory facility to perform the isolations within about 2 h of obtaining the fresh tissue. Once the mitochondria have been isolated, the mitochondrial respiration measurements should also be completed within ~1 h.
Isolated mitochondrial respiration rates are usually performed by measuring oxygen concentration in a sealed container connected to a Clark electrode. The theory behind this method is founded on the basic observation that oxygen is the last electron acceptor of mitochondrial respiration during oxidative phosphorylation. Therefore, as oxygen concentration falls during an experiment, it is assumed that adenosine triphosphate (ATP) production occurs10. Consumed oxygen is a proxy for produced ATP. Researchers can create specific experimental conditions using different substrates and initiate adenosine diphosphate (ADP)-stimulated respiration (state 3) by adding predetermined amounts of ADP to the chamber. Following the phosphorylation of the exogenous ADP to ATP, the oxygen consumption rate decreases, and state 4 is reached and can be measured. Furthermore, the addition of specific inhibitors allows information regarding leak respiration and uncoupled respiration to be obtained10. The ratio of state 3 to state 4 determines the respiratory control ratio (RCR), which is the indicator of overall mitochondrial coupling10,11. Lower values of RCR indicate overall mitochondrial dysfunction, whereas higher RCR values suggest a greater extent of mitochondrial coupling10.
As previously stated, the collection of biological material, mitochondrial isolation, and measurement of respiration rates must be completed within 2 h of obtaining tissue. To accomplish this task without transporting animals over large distances to established laboratories, a mobile mitochondrial physiology laboratory was constructed to be taken to field locations where these data can be collected. A 2018 Jayco Redhawk recreational vehicle was converted into a mobile molecular physiology laboratory and named the Auburn University (AU) MitoMobile (Figure 1A). A recreational vehicle was selected because of the built-in refrigerator, freezer, water storage tank and plumbing, electricity powered by 12-volt batteries, gas generator, propane tank, and self-leveling system. Further, the recreational vehicle provides the capability of staying at remote sites overnight for data collection. The front of the vehicle was not altered and provides the driving and sleeping quarters (Figure 1B). Previously installed bedroom amenities (bed, TV, and cabinet) in the rear of the vehicle and the stovetop were removed.
Custom-made stainless-steel shelving and a custom quartz countertop supported by 80/20 aluminum framing were installed in place of the bedroom amenities and stovetop (Figure 1C). The laboratory benches provide adequate space for data collection (Figure 1D). Power consumption of each piece of equipment (i.e., refrigerated centrifuge, mitochondrial respiration chambers, plate readers, computers, homogenizers, scales, portable ultra-freezer, and other general laboratory supplies) was taken into consideration. To support the large voltage and current demands of the centrifuge, the electrical system was upgraded to that of aircraft-grade equipment. An external compartment in the rear of the vehicle was converted into a liquid nitrogen storage bay, which meets the United States Department of Transportation&#39;s guidelines for liquid nitrogen storage and transport. This storage unit was constructed with stainless steel and has proper venting to keep any expanding nitrogen gas from leaking into the passenger compartment of the vehicle.
To confirm that the mobile laboratory can be used in mitochondrial bioenergetic studies, mitochondria were isolated, and mitochondrial respiration rates from wild-derived house mice (Mus musculus) hindlimb skeletal muscle were measured. Because Mus musculus is a model organism, the mitochondrial respiration rates of this species are well-established12,13,14. Although previous studies have documented mitochondrial isolation via differential centrifugation15,16,17, a brief overview of the methods used in the mobile mitochondrial physiology laboratory methods is described below.

<table><tbody><tr><td>1.7 mL centrifuge tubes</td><td>VWR</td><td>87003-294</td><td></td></tr><tr><td>2.0 mL centrifuge tubes</td><td>VWR</td><td>87003-298</td><td></td></tr><tr><td>50 mL centrifuge tubes</td><td>VWR</td><td>21009-681</td><td>Nalgene Oak Ridge Centrifuge Tube</td></tr><tr><td>ADP</td><td>VWR</td><td>97061-104</td><td></td></tr><tr><td>ATP</td><td>VWR</td><td>700009-070</td><td></td></tr><tr><td>Bradford</td><td>VWR</td><td>7065-020</td><td></td></tr><tr><td>Clear 96 well plate</td><td>VWR</td><td>82050-760</td><td>Greiner Bio-One</td></tr><tr><td>Dounce homogenizer</td><td>VWR</td><td>22877-284</td><td>Corning</td></tr><tr><td>EGTA</td><td>VWR</td><td>EM-4100</td><td></td></tr><tr><td>Filter paper</td><td></td><td></td><td>Included with Hansatech OxyGraph</td></tr><tr><td>Free-fatty acid BSA</td><td>VWR</td><td>89423-672</td><td></td></tr><tr><td>Glucose</td><td>VWR</td><td>BDH8005-500G</td><td></td></tr><tr><td>Glutamate</td><td>VWR</td><td>A12919</td><td></td></tr><tr><td>Hamilton Syringes</td><td>VWR</td><td>60373-985</td><td>Gaslight 1700 Series Syringes</td></tr><tr><td>Hansatech OxyGraph</td><td>Hansatech Instruments Ltd</td><td></td><td>No Catalog Number, but can be found under Products --&gt; Electrode Control Units</td></tr><tr><td>KH&lt;sub&gt;2&lt;/sub&gt;PO4</td><td>VWR</td><td>97062-350</td><td></td></tr><tr><td>Malate</td><td>VWR</td><td>97062-140</td><td></td></tr><tr><td>Mannitol</td><td>VWR</td><td>97061-052</td><td></td></tr><tr><td>Membrane</td><td></td><td></td><td>Included with Hansatech OxyGraph</td></tr><tr><td>MgCl&lt;sub&gt;2&lt;/sub&gt;</td><td>VWR</td><td>97063-152</td><td></td></tr><tr><td>MOPS</td><td>VWR</td><td>80503-004</td><td></td></tr><tr><td>Policeman</td><td>VWR</td><td>470104-462</td><td></td></tr><tr><td>Polytron</td><td>Thomas Scientific</td><td>11090044</td><td></td></tr><tr><td>Potassium chloride (KCl)</td><td>VWR</td><td>97061-566</td><td></td></tr><tr><td>Protease</td><td>VWR</td><td>97062-366</td><td>Trypsin is commonly used; however, other proteases can be used.</td></tr><tr><td>Pyruvic acid</td><td>VWR</td><td>97061-448</td><td></td></tr><tr><td>Sodium Dithionite</td><td>VWR</td><td>AA33381-22</td><td></td></tr><tr><td>Succinate</td><td>VWR</td><td>89230-086</td><td></td></tr><tr><td>Sucrose</td><td>VWR</td><td>BDH0308-500G</td><td></td></tr><tr><td>Tris-Base</td><td>VWR</td><td>97061-794</td><td></td></tr><tr><td>Tris-HCl</td><td>VWR</td><td>97061-258</td><td></td></tr></tbody></table>

development of a mobile mitochondrial physiology laboratory for measuring mitochondrial energetics in the field

Mitochondrial energetics is a central theme in animal biochemistry and physiology, with researchers using mitochondrial respiration as a metric to investigate metabolic capability. To obtain the measures of mitochondrial respiration, fresh biological samples must be used, and the entire laboratory procedure must be completed within approximately 2 h. Furthermore, multiple pieces of specialized equipment are required to perform these laboratory assays. This creates a challenge for measuring mitochondrial respiration in the tissues of wild animals living far from physiology laboratories as live tissue cannot be preserved for very long after collection in the field. Moreover, transporting live animals over long distances induces stress, which can alter mitochondrial energetics.
This manuscript introduces the Auburn University (AU) MitoMobile, a mobile mitochondrial physiology laboratory that can be taken into the field and used on-site to measure mitochondrial metabolism in tissues collected from wild animals. The basic features of the mobile laboratory and the step-by-step methods for measuring isolated mitochondrial respiration rates are presented. Additionally, the data presented validate the success of outfitting the mobile mitochondrial physiology laboratory and making mitochondrial respiration measurements. The novelty of the mobile laboratory lies in the ability to drive to the field and perform mitochondrial measurements on the tissues of animals captured on site.

The current manuscript investigated the mitochondrial respiration of wild-derived Mus musculus (n = 7, male = 5, female = 2; age = 1.30 &#177; 0.2 years) in a mobile mitochondrial physiology laboratory (Figure 1). To measure skeletal muscle mitochondrial respiration, the entire hindlimb, thus aerobic and anaerobic muscle, was used for mitochondrial isolation (Figure 2). Examples of raw mitochondrial respiration data are shown in Fig...

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Development of a Mobile Mitochondrial Physiology Laboratory for Measuring Mitochondrial Energetics in the Field

We designed and constructed a mobile laboratory to measure respiration rates in isolated mitochondria of wild animals captured at field locations. Here, we describe the design and outfitting of a mobile mitochondrial laboratory and the associated laboratory protocols.

Mitochondrial energetics is a central theme in animal biochemistry and physiology, with researchers using mitochondrial respiration as a metric to ...

development-mobile-mitochondrial-physiology-laboratory-for-measuring

Research

JoVE Journal

Biology

1.4K Views.  Auburn University. De ontwikkeling van een mobiel mitochondriaal laboratorium stelt fysiologen in staat om hun werk in het veld te brengen, en als gevolg daarvan kunnen ze een fascinerende diversiteit aan energetische uitdagingen evalueren die dieren vertonen. Gevangenschap is stressvol voor veel dieren, dus met een mobiel lab kunnen we nu het lab naar de dieren brengen in plaats van de dieren terug te brengen naar het laboratorium. Met de mogelijkheid om mitochondriale ademhaling in het veld te isoleren en te ...

Video: Development of a Mobile Mitochondrial Physiology Laboratory for Measuring Mitochondrial Energetics in the Field

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

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

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

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

Developmental Biology

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

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

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

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

Biochemistry

Stable Isotope Tracing & LC-MS Analysis to Measure DHODH and SDH Activities

Oxygen-Independent Assays to Measure Mitochondrial Function in Mammals

The flow of electrons in the mitochondrial electron transport chain (ETC) supports multifaceted biosynthetic, bioenergetic, and signaling functions in mammalian cells. As oxygen (O2) is the most ubiquitous terminal electron acceptor for the mammalian ETC, the O2 consumption rate is frequently used as a proxy for mitochondrial function. However, emerging research demonstrates that this parameter is not always indicative of mitochondrial function, as fumarate can be employed as an alternative electron acceptor to sustain mitochondrial functions in hypoxia. This article compiles a series of protocols that allow researchers to measure mitochondrial function independently of the O2 consumption rate. These assays are particularly useful when studying mitochondrial function in hypoxic environments. Specifically, we describe methods to measure mitochondrial ATP production, de novo pyrimidine biosynthesis, NADH oxidation by complex I, and superoxide production. In combination with classical respirometry experiments, these orthogonal and economical assays will provide researchers with a more comprehensive assessment of mitochondrial function in their system of interest.

Author Spotlight: Oxygen-Independent Assays to Measure Mitochondrial Function in Mammals  

Here, we present a compilation of assays to directly measure mitochondrial function in mammalian cells independently of their ability to consume molecular oxygen.

The flow of electrons in the mitochondrial electron transport chain (ETC) supports multifaceted biosynthetic, bioenergetic, and signaling functions in ...

Mitochondrial Isolation from Skeletal Muscle

Mitochondria are organelles controlling the life and death of the cell. They participate in key metabolic reactions, synthesize most of the ATP, and regulate a number of signaling cascades2,3. Past and current researchers have isolated mitochondria from rat and mice tissues such as liver, brain and heart4,5. In recent years, many researchers have focused on studying mitochondrial function from skeletal muscles.
Here, we describe a method that we have used successfully for the isolation of mitochondria from skeletal muscles 6. Our procedure requires that all buffers and reagents are made fresh and need about 250-500 mg of skeletal muscle. We studied mitochondria isolated from rat and mouse gastrocnemius and diaphragm, and rat extraocular muscles. Mitochondrial protein concentration is measured with the Bradford assay. It is important that mitochondrial samples be kept ice-cold during preparation and that functional studies be performed within a relatively short time (~1 hr). Mitochondrial respiration is measured using polarography with a Clark-type electrode (Oxygraph system) at 37&deg;C7. Calibration of the oxygen electrode is a key step in this protocol and it must be performed daily. Isolated mitochondria (150 &mu;g) are added to 0.5 ml of experimental buffer (EB). State 2 respiration starts with addition of glutamate (5mM) and malate (2.5 mM). Then, adenosine diphosphate (ADP) (150 &mu;M) is added to start state 3. Oligomycin (1 &mu;M), an ATPase synthase blocker, is used to estimate state 4. Lastly, carbonyl cyanide p-[trifluoromethoxy]-phenyl-hydrazone (FCCP, 0.2 &mu;M) is added to measurestate 5, or uncoupled respiration 6. The respiratory control ratio (RCR), the ratio of state 3 to state 4, is calculated after each experiment. An RCR &ge;4 is considered as evidence of a viable mitochondria preparation.
In summary, we present a method for the isolation of viable mitochondria from skeletal muscles that can be used in biochemical (e.g., enzyme activity, immunodetection, proteomics) and functional studies (mitochondrial respiration).

This protocol describes a procedure to study the respiration of mitochondria isolated from skeletal muscles. This method was adapted from Scorrano et al. (2007). The mitochondrial isolation procedure requires about 2 hours. The mitochondrial respiration can be completed in about 1 hour.

Mitochondria are organelles controlling the life and death of the cell. They participate in key metabolic reactions, synthesize most of the ATP, and ...

Bioenergetic Profile Experiment using C2C12 Myoblast Cells

The ability to measure cellular metabolism and understand mitochondrial dysfunction, has enabled scientists worldwide to advance their research in understanding the role of mitochondrial function in obesity, diabetes, aging, cancer, cardiovascular function and safety toxicity.
Cellular metabolism is the process of substrate uptake, such as oxygen, glucose, fatty acids, and glutamine, and subsequent energy conversion through a series of enzymatically controlled oxidation and reduction reactions. These intracellular biochemical reactions result in the production of ATP, the release of heat and chemical byproducts, such as lactate and CO2 into the extracellular environment.
Valuable insight into the physiological state of cells, and the alteration of the state of those cells, can be gained through measuring the rate of oxygen consumed by the cells, an indicator of mitochondrial respiration - the Oxygen Consumption Rate - or OCR. Cells also generate ATP through glycolysis, i.e.: the conversion of glucose to lactate, independent of oxygen. In cultured wells, lactate is the primary source of protons. Measuring the lactic acid produced indirectly via protons released into the extracellular medium surrounding the cells, which causes acidification of the medium provides the Extra-Cellular Acidification Rate - or ECAR.
In this experiment, C2C12 myoblast cells are seeded at a given density in Seahorse cell culture plates. The basal oxygen consumption (OCR) and extracellular acidification (ECAR) rates are measured to establish baseline rates. The cells are then metabolically perturbed by three additions of different compounds (in succession) that shift the bioenergetic profile of the cell.
This assay is derived from a classic experiment to assess mitochondria and serves as a framework with which to build more complex experiments aimed at understanding both physiologic and pathophysiologic function of mitochondria and to predict the ability of cells to respond to stress and/or insults.

A description of a method for profiling mitochondrial function in cells is provided. The mitochondrial profile generated provides four parameters of mitochondrial function that can be measured in one experiment: basal respiration rate, ATP-linked respiration, proton leak, and reserve capacity.

The ability to measure cellular metabolism and understand mitochondrial dysfunction, has enabled scientists worldwide to advance their research in ...

Mitochondrial Functional Assessment in Brush Lizard Liver Mitochondria

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

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

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

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

Real-Time Measurement of the Mitochondrial Bioenergetic Profile of Neutrophils

Neutrophils are the first line of defense and the most abundant leukocytes in humans. These effector cells perform functions such as phagocytosis and oxidative burst, and create neutrophil extracellular traps (NETs) for microbial clearance. New insights into the metabolic activities of neutrophils challenge the early concept that they primarily rely on glycolysis. Precise measurement of metabolic activities can unfold different metabolic requirements of neutrophils, including the tricarboxylic acid (TCA) cycle (also known as the Krebs cycle), oxidative phosphorylation (OXPHOS), pentose phosphate pathway (PPP), and fatty acid oxidation (FAO) under physiological conditions and in disease states. This paper describes a step-by-step protocol and prerequirements to measure oxygen consumption rate (OCR) as an indicator of mitochondrial respiration on mouse bone marrow-derived neutrophils, human blood-derived neutrophils, and the neutrophil-like HL60 cell line, using metabolic flux analysis on a metabolic extracellular flux analyzer. This method can be used for quantifying the mitochondrial functions of neutrophils under normal and disease conditions.

We describe stepwise protocols measuring the mitochondrial respiration of mouse and human neutrophils and HL60 cells using the metabolic extracellular flux analyzer.

Neutrophils are the first line of defense and the most abundant leukocytes in humans. These effector cells perform functions such as phagocytosis and ...

Immunology and Infection

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

Comparison between two or more distinct groups, such as healthy vs. disease, is necessary to determine cellular status. Mitochondria are at the nexus of cell heath due to their role in both cell metabolism and energy production as well as control of apoptosis. Therefore, direct evaluation of isolated mitochondria and mitochondrial perturbation offers the ability to determine if organelle-specific (dys)function is occurring. The methods described in this protocol include isolation of intact, functional mitochondria from HEK cultured cells and mouse liver and spinal cord, but can be easily adapted for use with other cultured cells or animal tissues. Mitochondrial function assessed by TMRE and the use of common mitochondrial uncouplers and inhibitors in conjunction with a fluorescent plate reader allow this protocol not only to be versatile and accessible to most research laboratories, but also offers high throughput.

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.

Comparison between two or more distinct groups, such as healthy vs. disease, is necessary to determine cellular status. Mitochondria are at the nexus of ...

Measuring Mitochondrial Function of Na&#239;ve and Effector CD8 T Cells

Understanding how immunometabolism impacts the function, differentiation, and fate of lymphocytes has garnered significant interest and attention. Lymphocyte biology has been explored using bioenergetic analysis and has now become a critically import tool in the field. Thus, we sought to optimize a bioenergetic analysis assay that can be adapted with pretreatments and acute injection for receptor stimulations. Here, we evaluated CD8 T cell ex vivo metabolism using the Cell Mito Stress Test to assess rates of oxygen consumption and extracellular acidification in na&#239;ve and effector CD8 T cells. Antigen-specific effector CD8 T cells were derived via ex vivo stimulation, and na&#239;ve CD8 T cells harvested from splenocytes and isolated with magnetic bead column separation. 
Pretreatments are performed in microplates and we detail how to prepare sensor cartridges. We show how injection ports can loaded with drugs to indirectly measure metabolic capacities and with metabolic modulators, this protocol can be used to study specific enzyme activity. T-cell receptor stimulations can be studied in real time with acute injection and stimulation with anti-CD3/CD28 using the injection ports. Instrument analyzers are used for measurements and data collection and data visualization is done with software programs to interpret cellular metabolism. This strategy produces an extensive amount of data on immune cell biology and mitochondrial bioenergetics allowing researchers to customize the protocol in numerous ways to explore CD8 T cell metabolism.

CD8 T cell bioenergetics can be interrogated using the Mito Stress Test. This methodology can be used to study acute and chronic metabolic programming. This protocol describes approaches to examine the relationships between T cell receptor biology and bioenergetic analysis.

Understanding how immunometabolism impacts the function, differentiation, and fate of lymphocytes has garnered significant interest and attention. ...

Measurement of Mitochondrial Membrane Potential In Vivo using a Genetically Encoded Voltage Indicator

Mitochondrial membrane potential (MMP, &#916;&#936;m) is critical for mitochondrial functions, including ATP synthesis, ion transport, reactive oxygen species (ROS) generation, and the import of proteins encoded by the nucleus. Existing methods for measuring &#916;&#936;m typically use lipophilic cation dyes, such as Rhodamine 800 and tetramethylrhodamine methyl ester (TMRM), but these are limited by low specificity and are not well-suited for in vivo applications. To address these limitations, we have developed a novel protocol utilizing genetically encoded voltage indicators (GEVIs). Genetically encoded voltage indicators (GEVIs), which generate fluorescent signals in response to membrane potential changes, have demonstrated significant potential for monitoring plasma membrane and neuronal potentials. However, their application to mitochondrial membranes remains unexplored. Here, we developed protein-based mitochondrial-targeted GEVIs capable of detecting &#916;&#936;m fluctuations in cells and the motor cortex of living animals. The mitochondrial potential indicator (MPI)offers a non-invasive approach to study &#916;&#936;m dynamics in real-time, providing a method to investigate mitochondrial function under both normal and pathological conditions.

This protocol describes the application of mitochondria-targeted genetically encoded voltage indicators (GEVIs). These GEVIs offer a significant advantage over traditional mitochondrial membrane potential dyes by enabling specific, in vivo, and real-time monitoring of mitochondrial membrane potential.

Mitochondrial membrane potential (MMP, &#916;&#936;m) is critical for mitochondrial functions, including ATP synthesis, ion transport, reactive oxygen ...

Development of a Mobile Mitochondrial Physiology Laboratory for Measuring Mitochondrial Energetics in the Field

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