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>
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			<td>2.0 mL centrifuge tubes</td>
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			<td>Nalgene Oak Ridge Centrifuge Tube</td>
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			<td>97061-104</td>
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			<td>Clear 96 well plate</td>
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			<td>Included with Hansatech OxyGraph</td>
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			<td>Free-fatty acid BSA</td>
			<td>VWR</td>
			<td>89423-672</td>
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			<td>Glucose</td>
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			<td>BDH8005-500G</td>
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			<td>Glutamate</td>
			<td>VWR</td>
			<td>A12919</td>
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			<td>Hamilton Syringes</td>
			<td>VWR</td>
			<td>60373-985</td>
			<td>Gaslight 1700 Series Syringes</td>
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			<td>Hansatech OxyGraph</td>
			<td>Hansatech Instruments Ltd</td>
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			<td>VWR</td>
			<td>97062-350</td>
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			<td>VWR</td>
			<td>97062-140</td>
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			<td>97061-052</td>
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			<td>MgCl2</td>
			<td>VWR</td>
			<td>97063-152</td>
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			<td>Polytron</td>
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			<td>Potassium chloride (KCl)</td>
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			<td>97062-366</td>
			<td>Trypsin is commonly used; however, other proteases can be used.</td>
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			<td>VWR</td>
			<td>97061-448</td>
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			<td>VWR</td>
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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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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

Probing for Mitochondrial Complex Activity in Human Embryonic Stem Cells

Mitochondria are cytoplasmic organelles that have a primary role in cellular metabolism and homeostasis, regulation of the cell signaling network, and programmed cell death. Mitochondria produce ATP, regulate the cytoplasmic redox state and Ca2+ balance, catabolize fatty acids, synthesize heme, nucleotides, steroid hormones, amino acids, and help assemble iron-sulfur clusters in proteins. Mitochondria also have an essential role in heat production. Mutations of the mitochondrial genome cause several types of human disorder. The accumulation of mtDNA mutations correlates with aging and is suspected to have an important role in the development of cancer. Due to their vitally important role in all cell types, the function of mitochondria must also be critical for stem cells. Key advances have been made in our understanding of stem cell viability, proliferation, and differentiation capacity. But the functional activity of stem cells, in particular their energy status, was not yet been studied in detail. Almost nothing is known about the mitochondrial properties of human embryonic stem cells (hESCs) and their differentiated precursor progeny. One way to understand and evaluate the role of mitochondria in hESC function and developmental potential is to directly measure the activity of mitochondrial respiratory complexes. Here, we describe high resolution clear native gel electrophoresis and subsequent in gel activity visualization as a method for analyzing the five respiratory chain complexes of hESCs.

In this video, we will show you how the mitochondrial respiratory chain complexes of human embryonic stem cells can be analyzed using in gel activity assays.

Mitochondria are cytoplasmic organelles that have a primary role in cellular metabolism and homeostasis, regulation of the cell signaling network, and ...

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 ...

A Screenable In Vivo Assay for Mitochondrial Modulators Using Transgenic Bioluminescent Caenorhabditis elegans

The multicellular model organism Caenorhabditis elegans is a small nematode of approximately 1 mm in size in adulthood that is genetically and experimentally tractable. It is economical and easy to culture and dispense in liquid medium which makes it well suited for medium-throughput screening. We have previously validated the use of transgenic luciferase expressing C. elegans strains to provide rapid in vivo assessment of the nematode&#8217;s ATP levels.1-3 Here we present the required materials and procedure to carry out bioassays with the bioluminescent C. elegans strains PE254 or PE255 (or any of their derivative strains). The protocol allows for in vivo detection of sublethal effects of drugs that may identify mitochondrial toxicity, as well as for in vivo detection of potential beneficial drug effects. Representative results are provided for the chemicals paraquat, rotenone, oxaloacetate and for four firefly luciferase inhibitory compounds. The methodology can be scaled up to provide a platform for screening drug libraries for compounds capable of modulating mitochondrial function. Pre-clinical evaluation of drug toxicity is often carried out on immortalized cancerous human cell lines which derive ATP mostly from glycolysis and are often tolerant of mitochondrial toxicants.4,5 In contrast, C. elegans depends on oxidative phosphorylation to sustain development into adulthood, drawing a parallel with humans and providing a unique opportunity for compound evaluation in the physiological context of a whole live multicellular organism.

A Screenable In Vivo Assay for Mitochondrial Modulators Using Transgenic Bioluminescent Caenorhabditis elegans

A protocol is described for in vivo detection of effects of mitochondrial inhibitors in the model organism Caenorhabditis elegans and for identification of potential enhancing compounds. This protocol can be used to screen drug libraries for compounds modulating mitochondrial function.

The multicellular model organism Caenorhabditis elegans is a small nematode of approximately 1 mm in size in adulthood that is genetically 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 ...

Measuring the Stiffness of Ex Vivo Mouse Aortas Using Atomic Force Microscopy

Arterial stiffening is a significant risk factor and biomarker for cardiovascular disease and a hallmark of aging. Atomic force microscopy (AFM) is a versatile analytical tool for characterizing viscoelastic mechanical properties for a variety of materials ranging from hard (plastic, glass, metal, etc.) surfaces to cells on any substrate. It has been widely used to measure the stiffness of cells, but less frequently used to measure the stiffness of aortas. In this paper, we will describe the procedures for using AFM in contact mode to measure the ex vivo elastic modulus of unloaded mouse arteries. We describe our procedure for isolation of mouse aortas, and then provide detailed information for the AFM analysis. This includes step-by-step instructions for alignment of the laser beam, calibration of the spring constant and deflection sensitivity of the AFM probe, and acquisition of force curves. We also provide a detailed protocol for data analysis of the force curves.

Measuring the Stiffness of Ex Vivo Mouse Aortas Using Atomic Force Microscopy

We present detailed protocols for isolation of aortas from mouse and measurement of their elastic modulus using atomic force microscopy.

Arterial stiffening is a significant risk factor and biomarker for cardiovascular disease and a hallmark of aging. Atomic force microscopy (AFM) is a ...

Studying Mitochondrial Structure and Function in Drosophila Ovaries

Analysis of the mitochondrial structure-function relationship is required for a thorough understanding of the regulatory mechanisms of mitochondrial functionality. Fluorescence microscopy is an indispensable tool for the direct assessment of mitochondrial structure and function in live cells and for studying the mitochondrial structure-function relationship, which is primarily modulated by the molecules governing fission and fusion events between mitochondria. This paper describes and demonstrates specific methods for studying mitochondrial structure and function in live as well as in fixed tissue in the model organism Drosophila melanogaster. The tissue of choice here is the Drosophila ovary, which can be isolated and made amenable for ex vivo live confocal microscopy. Furthermore, the paper describes how to genetically manipulate the mitochondrial fission protein, Drp1, in Drosophila ovaries to study the involvement of Drp1-driven mitochondrial fission in modulating the mitochondrial structure-function relationship. The broad use of such methods is demonstrated in already-published as well as in novel data. The described methods can be further extended towards understanding the direct impact of nutrients and/or growth factors on the mitochondrial properties ex vivo. Given that mitochondrial dysregulation underlies the etiology of various diseases, the described innovative methods developed in a genetically tractable model organism, Drosophila, are anticipated to contribute significantly to the understanding of the mechanistic details of the mitochondrial structure-function relationship and to the development of mitochondria-directed therapeutic strategies.

Studying Mitochondrial Structure and Function in Drosophila Ovaries

Analysis of the mitochondrial structure-function relationship is required for a thorough understanding of the regulatory mechanisms of mitochondrial functionality. Specific methods for studying mitochondrial structure and function in live and fixed Drosophila ovaries are described and demonstrated in this paper.

Analysis of the mitochondrial structure-function relationship is required for a thorough understanding of the regulatory mechanisms of mitochondrial ...

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+ ...

Simultaneous Mapping and Quantitation of Ribonucleotides in Human Mitochondrial DNA

Established approaches to estimate the number of ribonucleotides present in a genome are limited to the quantitation of incorporated ribonucleotides using short synthetic DNA fragments or plasmids as templates and then extrapolating the results to the whole genome. Alternatively, the number of ribonucleotides present in a genome may be estimated using alkaline gels or Southern blots. More recent in vivo approaches employ Next-generation sequencing allowing genome-wide mapping of ribonucleotides, providing the position and identity of embedded ribonucleotides. However, they do not allow quantitation of the number of ribonucleotides which are incorporated into a genome. Here we describe how to simultaneously map and quantitate the number of ribonucleotides which are incorporated into human mitochondrial DNA in vivo by Next-generation sequencing. We use highly intact DNA and introduce sequence specific double strand breaks by digesting it with an endonuclease, subsequently hydrolyzing incorporated ribonucleotides with alkali. The generated ends are ligated with adapters and these ends are sequenced on a Next-generation sequencing machine. The absolute number of ribonucleotides can be calculated as the number of reads outside the recognition site per average number of reads at the recognition site for the sequence specific endonuclease. This protocol may also be utilized to map and quantitate free nicks in DNA and allows adaption to map other DNA lesions that can be processed to 5&#180;-OH ends or 5&#180;-phosphate ends. Furthermore, this method can be applied to any organism, given that a suitable reference genome is available. This protocol therefore provides an important tool to study DNA replication, 5&#180;-end processing, DNA damage, and DNA repair.

Here we describe a method amenable to simultaneously quantitate and genome-wide map ribonucleotides in highly intact DNA at single-nucleotide resolution, combining enzymatic cleavage of genomic DNA with its alkaline hydrolysis and subsequent 5&#180;-end sequencing.

Established approaches to estimate the number of ribonucleotides present in a genome are limited to the quantitation of incorporated ribonucleotides using ...

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.