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.

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			<td>1.7 mL centrifuge tubes</td>
			<td>VWR</td>
			<td>87003-294</td>
			<td></td>
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		<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>
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		<tr>
			<td>ADP</td>
			<td>VWR</td>
			<td>97061-104</td>
			<td></td>
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			<td>ATP</td>
			<td>VWR</td>
			<td>700009-070</td>
			<td></td>
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			<td>Bradford</td>
			<td>VWR</td>
			<td>7065-020</td>
			<td></td>
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		<tr>
			<td>Clear 96 well plate</td>
			<td>VWR</td>
			<td>82050-760</td>
			<td>Greiner Bio-One</td>
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			<td>Dounce homogenizer</td>
			<td>VWR</td>
			<td>22877-284</td>
			<td>Corning</td>
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			<td>EGTA</td>
			<td>VWR</td>
			<td>EM-4100</td>
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			<td>Filter paper</td>
			<td></td>
			<td></td>
			<td>Included with Hansatech OxyGraph</td>
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		<tr>
			<td>Free-fatty acid BSA</td>
			<td>VWR</td>
			<td>89423-672</td>
			<td></td>
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			<td>Glucose</td>
			<td>VWR</td>
			<td>BDH8005-500G</td>
			<td></td>
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		<tr>
			<td>Glutamate</td>
			<td>VWR</td>
			<td>A12919</td>
			<td></td>
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		<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 --&#62; Electrode Control Units</td>
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		<tr>
			<td>KH2PO4</td>
			<td>VWR</td>
			<td>97062-350</td>
			<td></td>
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			<td>Malate</td>
			<td>VWR</td>
			<td>97062-140</td>
			<td></td>
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			<td>Mannitol</td>
			<td>VWR</td>
			<td>97061-052</td>
			<td></td>
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		<tr>
			<td>Membrane</td>
			<td></td>
			<td></td>
			<td>Included with Hansatech OxyGraph</td>
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		<tr>
			<td>MgCl2</td>
			<td>VWR</td>
			<td>97063-152</td>
			<td></td>
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			<td>MOPS</td>
			<td>VWR</td>
			<td>80503-004</td>
			<td></td>
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			<td>Policeman</td>
			<td>VWR</td>
			<td>470104-462</td>
			<td></td>
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			<td>Polytron</td>
			<td>Thomas Scientific</td>
			<td>11090044</td>
			<td></td>
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			<td>Potassium chloride (KCl)</td>
			<td>VWR</td>
			<td>97061-566</td>
			<td></td>
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			<td>Protease</td>
			<td>VWR</td>
			<td>97062-366</td>
			<td>Trypsin is commonly used; however, other proteases can be used.</td>
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			<td>Pyruvic acid</td>
			<td>VWR</td>
			<td>97061-448</td>
			<td></td>
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			<td>Sodium Dithionite</td>
			<td>VWR</td>
			<td>AA33381-22</td>
			<td></td>
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			<td>Succinate</td>
			<td>VWR</td>
			<td>89230-086</td>
			<td></td>
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			<td>Sucrose</td>
			<td>VWR</td>
			<td>BDH0308-500G</td>
			<td></td>
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			<td>Tris-Base</td>
			<td>VWR</td>
			<td>97061-794</td>
			<td></td>
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		<tr>
			<td>Tris-HCl</td>
			<td>VWR</td>
			<td>97061-258</td>
			<td></td>
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</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...

Watch this Scientific Journal Video about Development of a Mobile Mitochondrial Physiology Laboratory for Measuring Mitochondrial Energetics in the Field at JoVE.com

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

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JoVE Journal

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1.4K Views.  Auburn University. 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. 

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

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 ...

Near Infrared (NIr) Light Increases Expression of a Marker of Mitochondrial Function in the Mouse Vestibular Sensory Epithelium

Strategies for attenuating decline in balance function with increasing age are predominantly focused on physical therapies including balance tasks and exercise. However, these approaches do not address the underlying causes of balance decline. Using mice, the impact of near infrared light (NIr) on the metabolism of cells in the vestibular sensory epithelium was assessed. Data collected shows that this simple and safe intervention may protect these vulnerable cells from the deleterious effects of natural aging. mRNA was extracted from the isolated peripheral vestibular sensory epithelium (crista ampullaris and utricular macula) and subsequently transcribed into a cDNA library. This library was then probed for the expression of ubiquitous antioxidant (SOD-1). Antioxidant gene expression was then used to quantify cellular metabolism. Using transcranial delivery of NIr in young (4 weeks) and older (8 - 9 months) mice, and a brief treatment regime (90 sec/day for 5 days), this work suggests NIr alone may be sufficient to improve mitochondrial function in the vestibular sensory epithelium. Since there are currently no available, affordable, non-invasive methods of therapy to improve vestibular hair cell function, the application of external NIr radiation provides a potential strategy to counteract the impact of aging on cellular metabolism inthe vestibular sensory epithelium.

Near Infrared NIr Light Increases Expression of a Marker of Mitochondrial Function in the Mouse Vestibular Sensory Epithelium

Mitochondrial dysfunction is a hallmark of cellular senescence. This paper uses non-invasive near-infrared (NIr) treatment to improve mitochondrial function in the aging mouse vestibular sensory epithelium.

Strategies for attenuating decline in balance function with increasing age are predominantly focused on physical therapies including balance tasks and ...

Design and Development of Aptamer&#8211;Gold Nanoparticle Based Colorimetric Assays for In-the-field Applications

The design and development of an aptamer&#8211;gold nanoparticle (AuNP) colorimetric assay for the detection of small molecules for in-the-field applications was examined. Target selective AuNP based color assays have been developed in controlled proof-of-concept laboratory settings. However, these schemes have not been exerted to a point of failure to determine their practical use beyond laboratory settings. This work describes a generic approach to design, develop, and troubleshoot an aptamer-AuNP colorimetric assay for small molecule analytes and using the assay for in-the-field settings. The assay is advantageous because adsorbed aptamers passivate the nanoparticle surfaces and provide a means to reduce and eliminate false positive responses to non-target analytes. Transitioning this system to practical uses required defining not only the shelf-life of the aptamer-AuNP assay, but establishing methods and procedures for extending the long-term storage capabilities. Also, one of the recognized concerns with colorimetric readout is the burden placed on analysts to accurately identify often subtle changes in color. To lessen the responsibility on analysts in the field, a color analysis protocol was designed to perform the color identification duties without the need for performing this task on laboratory grade equipment. The method for creating and testing the data analysis protocol is described. However to understand and influence the design of adsorbed aptamer assays, the interactions associated with the aptamer, target, and AuNPs require further study. The knowledge gained could lead to tailoring aptamers for improved functionality.

The design and development of an aptamer&#8211;gold nanoparticle colorimetric assay for the detection of small molecules for in-the-field applications was examined. Additionally, a smart-device colorimetric application (app) was validated and long-term storage of the assay was established for use in the field.

The design and development of an aptamer&#8211;gold nanoparticle (AuNP) colorimetric assay for the detection of small molecules for in-the-field ...

Simultaneous Measurement of Mitochondrial Calcium and Mitochondrial Membrane Potential in Live Cells by Fluorescent Microscopy

Apart from their essential role in generating ATP, mitochondria also act as local calcium (Ca2+) buffers to tightly regulate intracellular Ca2+ concentration. To do this, mitochondria utilize the electrochemical potential across their inner membrane (&#916;&#936;m) to sequester Ca2+. The influx of Ca2+ into the mitochondria stimulates three rate-limiting dehydrogenases of the citric acid cycle, increasing electron transfer through the oxidative phosphorylation (OXPHOS) complexes. This stimulation maintains &#916;&#936;m, which is temporarily dissipated as the positive calcium ions cross the mitochondrial inner membrane into the mitochondrial matrix.
We describe here a method for simultaneously measuring mitochondria Ca2+ uptake and &#916;&#936;m in live cells using confocal microscopy. By permeabilizing the cells, mitochondrial Ca2+ can be measured using the fluorescent Ca2+ indicator Fluo-4, AM, with measurement of &#916;&#936;m using the fluorescent dye tetramethylrhodamine, methyl ester, perchlorate (TMRM). The benefit of this system is that there is very little spectral overlap between the fluorescent dyes, allowing accurate measurement of mitochondrial Ca2+ and &#916;&#936;m simultaneously. Using the sequential addition of Ca2+ aliquots, mitochondrial Ca2+ uptake can be monitored, and the concentration at which Ca2+ induces mitochondrial membrane permeability transition and the loss of &#916;&#936;m determined.

Mitochondria can utilize the electrochemical potential across their inner membrane (&#916;&#936;m) to sequester calcium (Ca2+), allowing them to shape cytosolic Ca2+ signaling within the cell. We describe a method for simultaneously measuring mitochondria Ca2+ uptake and &#916;&#936;m in live cells using fluorescent dyes and confocal microscopy.

Apart from their essential role in generating ATP, mitochondria also act as local calcium (Ca2+) buffers to tightly regulate intracellular Ca2+ ...

A Protocol for Laboratory Housing of Turquoise Killifish (Nothobranchius furzeri)

The development of husbandry practices in non-model laboratory fish used for experimental purposes has greatly benefited from the establishment of reference fish model systems, such as zebrafish and medaka. In recent years, an emerging fish &#8211; the turquoise killifish (Nothobranchius furzeri) &#8211; has been adopted by a growing number of research groups in the fields of biology of aging and ecology. With a captive life span of 4 - 8 months, this species is the shortest-lived vertebrate raised in captivity and allows the scientific community to test &#8211; in a short time &#8211; experimental interventions that can lead to alterations of the aging rate and life expectancy. Given the unique biology of this species, characterized by embryonic diapause, explosive sexual maturation, marked morphological and behavioral sexual dimorphism - and their relatively short adult life span -&#160;ad hoc husbandry practices are in urgent demand. This protocol reports a set of key husbandry measures that allow optimal turquoise killifish laboratory care, enabling the scientific community to adopt this species as a powerful laboratory animal model.

A Protocol for Laboratory Housing of Turquoise Killifish Nothobranchius furzeri

Laboratory housing of turquoise killifish can be scaled up&#160;to house and efficiently raise thousands of individual fish in a centralized water filtration system, employing the same infrastructure used for standard zebrafish facilities. Here we detail a list of standardized procedures that allow efficient killifish maintenance.

The development of husbandry practices in non-model laboratory fish used for experimental purposes has greatly benefited from the establishment of ...

Measuring Mitochondrial Substrate Flux in Recombinant Perfringolysin O-Permeabilized Cells

Mitochondrial substrate flux is a distinguishing characteristic of each cell type, and changes in its components such as transporters, channels, or enzymes are involved in the pathogenesis of several diseases. Mitochondrial substrate flux can be studied using intact cells, permeabilized cells, or isolated mitochondria. Investigating intact cells encounters several problems due to simultaneous oxidation of different substrates. Besides, several cell types contain internal stores of different substrates that complicate results interpretation. Methods such as mitochondrial isolation or using permeabilizing agents are not easily reproducible. Isolating pure mitochondria with intact membranes in sufficient amounts from small samples is problematic. Using non-selective permeabilizers causes various degrees of unavoidable mitochondrial membrane damage. Recombinant perfringolysin O (rPFO) was offered as a more appropriate permeabilizer, thanks to its ability to selectively permeabilize plasma membrane without affecting mitochondrial integrity. When used in combination with microplate respirometry, it allows testing the flux of several mitochondrial substrates with enough replicates within one experiment while using a minimal number of cells. In this work, the protocol describes a method to compare mitochondrial substrate flux of two different cellular phenotypes or genotypes and can be customized to test various mitochondrial substrates or inhibitors.

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.

Mitochondrial substrate flux is a distinguishing characteristic of each cell type, and changes in its components such as transporters, channels, or ...

Analyzing Mitochondrial Morphology Through Simulation Supervised Learning

The quantitative analysis of subcellular organelles such as mitochondria in cell fluorescence microscopy images is a demanding task because of the inherent challenges in the segmentation of these small and morphologically diverse structures. In this article, we demonstrate the use of a machine learning-aided segmentation and analysis pipeline for the quantification of mitochondrial morphology in fluorescence microscopy images of fixed cells. The deep learning-based segmentation tool is trained on simulated images and eliminates the requirement for ground truth annotations for supervised deep learning. We demonstrate the utility of this tool on fluorescence microscopy images of fixed cardiomyoblasts with a stable expression of fluorescent mitochondria markers and employ specific cell culture conditions to induce changes in the mitochondrial morphology.

This article explains how to use simulation-supervised machine learning for analyzing mitochondria morphology in fluorescence microscopy images of fixed cells.

The quantitative analysis of subcellular organelles such as mitochondria in cell fluorescence microscopy images is a demanding task because of the ...

Improved Protein Quantification Using Two-Step Mitochondrial Isolation

Two-Step Tag-Free Isolation of Mitochondria for Improved Protein Discovery and Quantification

Most physiological and disease processes, from central metabolism to immune response to neurodegeneration, involve mitochondria. The mitochondrial proteome is composed of more than 1,000 proteins, and the abundance of each can vary dynamically in response to external stimuli or during disease progression. Here, we describe a protocol for isolating high-quality mitochondria from primary cells and tissues. The two-step procedure comprises (1) mechanical homogenization and differential centrifugation to isolate crude mitochondria, and (2) tag-free immune capture of mitochondria to isolate pure organelles and eliminate contaminants. Mitochondrial proteins from each purification stage are analyzed by quantitative mass spectrometry, and enrichment yields are calculated, allowing the discovery of novel mitochondrial proteins by subtractive proteomics. Our protocol provides a sensitive and comprehensive approach to studying mitochondrial content in cell lines, primary cells, and tissues.

Author Spotlight: Two-Step Tag-Free Isolation of Mitochondria for Improved Protein Discovery and Quantification  

We present a two-step protocol for high-quality mitochondria isolation that is compatible with protein discovery and quantification at a proteome scale. Our protocol does not require genetic engineering and is thus suitable for studying mitochondria from any primary cells and tissues.

Most physiological and disease processes, from central metabolism to immune response to neurodegeneration, involve mitochondria. The mitochondrial ...

Imaging of Mitochondrial Peroxide in Living Yeast Cells Using mtHyper7

Imaging of mtHyPer7, a Ratiometric Biosensor for Mitochondrial Peroxide, in Living Yeast Cells

Mitochondrial dysfunction, or functional alteration, is found in many diseases and conditions, including neurodegenerative and musculoskeletal disorders, cancer, and normal aging. Here, an approach is described to assess mitochondrial function in living yeast cells at cellular and subcellular resolutions using a genetically encoded, minimally invasive, ratiometric biosensor. The biosensor, mitochondria-targeted HyPer7 (mtHyPer7), detects hydrogen peroxide (H2O2) in mitochondria. It consists of a mitochondrial signal sequence fused to a circularly permuted fluorescent protein and the H2O2-responsive domain of a bacterial OxyR protein. The biosensor is generated and integrated into the yeast genome using a CRISPR-Cas9 marker-free system, for&#160;more consistent expression compared to plasmid-borne constructs.
mtHyPer7 is quantitatively targeted to mitochondria, has no detectable effect on yeast growth rate or mitochondrial morphology, and provides a quantitative readout for mitochondrial H2O2 under normal growth conditions and upon exposure to oxidative stress. This protocol explains how to optimize imaging conditions using a spinning-disk confocal microscope system and perform quantitative analysis using freely available software. These tools make it possible to collect rich spatiotemporal information on mitochondria both within cells and among cells in a population. Moreover, the workflow described here can be used to validate other biosensors.

Author Spotlight: Advancing Mitochondrial Research - mtHyper7 Biosensor for Subcellular Analysis 

Hydrogen peroxide (H2O2) is both a source of oxidative damage and a signaling molecule. This protocol describes how to measure mitochondrial H2O2 using mitochondria-targeted HyPer7 (mtHyPer7), a genetically encoded ratiometric biosensor, in living yeast. It details how to optimize imaging conditions and perform quantitative cellular and subcellular analysis using freely available software.

Mitochondrial dysfunction, or functional alteration, is found in many diseases and conditions, including neurodegenerative and musculoskeletal disorders, ...

Confocal Microscopic Analysis of Rat Muscle Mitochondria

Analysis of the Mitochondrial Density and Longitudinal Distribution in Rat Live-Skeletal Muscle Fibers by Confocal Microscopy

The mitochondrion is an organelle that can be elongated, fragmented, and renovated according to the metabolic requirements of the cells. The remodeling of the mitochondrial network allows healthy mitochondria to meet cellular demands; however, the loss of this capacity has been related to the development or progression of different pathologies. In skeletal muscle, mitochondrial density and distribution changes are observed in physiological and pathological conditions such as exercise, aging, and obesity, among others. Therefore, the study of the mitochondrial network may provide a better understanding of mechanisms related to those conditions.
Here, a protocol for mitochondria imaging of live-skeletal muscle fibers from rats is described. Fibers are manually dissected in a relaxing solution and incubated with a fluorescent live-cell imaging indicator of mitochondria (tetramethylrhodamine ethyl ester, TMRE). The mitochondria signal is recorded by confocal microscopy using the XYZ scan mode to obtain confocal images of the intermyofibrillar mitochondrial (IMF) network. After that, the confocal images are processed by thresholding and binarization. The binarized confocal image accounts for the positive pixels for mitochondria, which are then counted to obtain the mitochondrial density. The mitochondrial network in skeletal muscle is characterized by a high density of IMF population, which has a periodic longitudinal distribution similar to that of T-tubules (TT).&#160;The Fast Fourier Transform (FFT) is a standard analysis technique performed to evaluate the distribution of TT that allows finding the distribution frequency and the level of their organization. In this protocol, the implementation of the FFT algorithm is described for the analysis of the longitudinal mitochondrial distribution in skeletal muscle.

Author Spotlight: Mitochondrial Remodeling in Skeletal Muscle 

Here, we present a protocol to analyze changes in mitochondrial density and longitudinal distribution by live-skeletal muscle imaging using confocal microscopy for mitochondrial network scanning.

The mitochondrion is an organelle that can be elongated, fragmented, and renovated according to the metabolic requirements of the cells. The remodeling of ...