The authors would like to acknowledge the generous support of the John and Debbie Oxley Endowed Chair for Equine Sports Medicine and the Grayson Jockey Club Research Foundation.

This study was approved by the Oklahoma State University Institutional Animal Care and Use Committee. Four Thoroughbred geldings (17.5 &#177; 1.3 years, 593 &#177; 45 kg) were used in this study to generate the representative results.
1. Obtaining skeletal muscle biopsy specimen
<ol>
	<li>Obtain skeletal muscle biopsies (follow sterile technique) from the center of the semitendinosus muscle (or other muscle of interest), using a 12 G University College Hospital (UCH) biopsy needle (see Table of Materials) while under light sedation, and using local anesthesia following a previously published report11.</li>
	<li>Transfer the biopsy specimens immediately into vials with ice-cold biopsy transport media solution (2.77 mM CaK2-EGTA, 7.23 mM K2-EGTA, 20 mM imidazole, 20 mM taurine, 50 mM K-MES, 0.5 mM dithiothreitol, 6.56 mM MgCl2, 5.77 mM ATP, and 15 mM phosphocreatine, adjusted to pH 7.1; see Table of Materials). Transport to the laboratory for analysis. 
	NOTE: Refer to Doerrier et al.15&#160;for specific instructions on preparing the biopsy transport media.</li>
	<li>Isolate mitochondria from the biopsy samples using a commercial kit, according to the manufacturer&#39;s instructions (see Table of Materials). The final storage buffer should not contain substrates such as ADP, ATP, and succinate, that are part of the respirometry assay.</li>
	<li>Resuspend the final pellet of the isolated mitochondria using 80 &#181;L of suspension media (225 mM mannitol, 75 mM sucrose, and 1 mM EGTA; see Table of Materials) per 100 mg of muscle used for mitochondria isolation.</li>
	<li>Minimize the interval from the time of the biopsy procedure to the isolation of mitochondria, and keep the samples at 0-4 &#176;C throughout the processing steps until added to the high-resolution respirometers. 
	&#8203;NOTE: Preliminary studies have found that suspensions of isolated mitochondria begin to lose functional capacity after approximately 2 h when maintained at 0-4 &#176;C.</li>
</ol>
2. Setting up of the high-resolution respirometer
<ol>
	<li>High-resolution respirometers are used to quantify mitochondrial respiration and associated processes. Use the software control program provided by the manufacturer (see Table of Materials) to control the respirometer, calibrate the sensors, and collect raw data. Analyze the samples in duplicate for each test condition. 
	NOTE: In all cases, the recommended titrations and final reagent concentrations described in this protocol are based on a 2 mL respirometry chamber.</li>
	<li>Determine the background O2 flux and zero point of the O2 sensor every 2-4 weeks through the serial titration of dithionite, following the manufacturer&#39;s instructions. Retain these calibration constants for subsequent assays until the calibration is repeated. 
	NOTE: In contrast to previously published procedures using permeabilized muscle fibers11,12,13,16, in which hyperoxia (250-500 &#181;M) is necessary to avoid diffusion limitation of O2 to the mitochondria, isolated mitochondria can be evaluated using a range of 50-200 &#181;M O2. This can be achieved simply by allowing the respiration media to equilibrate with room air prior to adding the mitochondrial suspension (i.e., following daily single-point calibration). Similarly, reoxygenation of the respiration chamber can be accomplished by simply opening the chamber until sufficient ambient oxygen has dissolved into the respirometry media to raise the oxygen concentration to the desired level.</li>
	<li>Fill the respirometer chambers with magnesium-free media (0.5 mM EGTA, 60 mM K-lactobionate, 20 mM taurine, 10 mM KH2PO4, 20 mM HEPES, 110 mM sucrose, and 1 g/L bovine serum albumin [BSA] essentially fatty acid-free, adjusted to pH 7.1; see Table of Materials). 
	NOTE: Refer to Komlodi et al.17 for specific instructions on preparing the respiration media.
	<ol>
		<li>Set the instrument incubation temperature to 38 &#176;C to represent the basal temperature of equine skeletal muscle, and set the mixing of the respiration media to 800 rpm using a magnetic stirrer turning in the bottom of the respirometer chamber.</li>
		<li>Turn chamber illumination off to avoid interference with fluorescent sensors.</li>
		<li>Energize the oxygen electrode with an 800 mV polarization voltage, and amplify the resulting signal with a gain setting of 1.</li>
		<li>Record the oxygen concentration every 2 s, calculate the oxygen flux as the negative slope of the oxygen measurement over the preceding 40 s (20 data points), and report as pmol x s-1 x mL of incubation solution.</li>
	</ol>
	</li>
	<li>Calibrate the oxygen sensor by allowing the media to equilibrate with room air. Calculate the reference oxygen partial pressure, based on barometric pressure measured by the high-resolution respirometer and standard atmospheric oxygen concentration.</li>
</ol>
3. Measurement of mitochondrial membrane potential using TMRM
<ol>
	<li>Use &#34;Green&#34; fluorescent sensors (530 nm dominant wavelength) to quantify the fluorescent signal from the respiration chamber. Energize the sensors at 400-500 mV; the resulting signal gets amplified with a gain of 1:1,000. 
	NOTE: Specific settings must be optimized for individual instruments to capture the expected signal within the linear range of the sensor.</li>
	<li>Add TMRM (4 &#181;L of 1 mM solution for a final concentration of 2 &#181;M; see Table of Materials) before the addition of mitochondria.</li>
	<li>Calibrate the fluorescent signal using a simple two-point calibration of fluorescent signal (voltage) versus the amount of added fluorophore (mM), prior to the addition of the mitochondria. 
	NOTE: Use the fluorescent signal calibration function in the machine control software to calibrate this signal. The raw signal from the fluorescent probe can require 30 min or more to stabilize in order to record the second point of the calibration.</li>
	<li>Perform the final calibration of the TMRM signal after completion of the respirometry titration protocol by delivering several titrations of uncoupling agent (carbonyl cyanide m-chlorophenyl hydrazone; 2 &#181;L per titration) until no further increases in TMRM fluorescent signal are observed, indicating the complete collapse of mitochondrial membrane potential (Figure 1). 
	&#8203;NOTE: This fluorescent signal value is considered equal to 0 mV transmembrane potential, and used as the reference point for relative membrane potential values recorded during the respirometry titration protocol.</li>
</ol>
4. Measurement of ATP production using magnesium green (MgG)
<ol>
	<li>Use &#34;Blue&#34; fluorescent sensors (465 nm dominant wavelength) to quantify the fluorescent signal from the respiration chamber. Energize these sensors for individual instruments to capture the expected signal within the linear range of the sensor.</li>
	<li>Perform the chemical setup and calibration of the MgG fluorescent signal after the addition of the mitochondria, but prior to the addition of any substrates. Add 8 &#181;L of 2 mM ethylenediaminetetraacetic acid (EDTA) to the respirometry chamber to chelate cations (particularly Ca2+) that would compete with Mg2+ for binding to MgG, then add 4 &#181;L of 1 mM MgG (1.1 &#181;M) to the respiration chamber.</li>
	<li>Calibrate the raw fluorescence signal with 10 x 2 &#181;L sequential titrations of 100 mM MgCl2, allowing 1 min between titrations for stabilization of the fluorescent signal (Figure 2). 
	NOTE: This process, which is completed offline following completion of the assay and using templates provided by the manufacturer of the machine and associated software, produces a second-order curve of magnesium concentration to fluorescent signal (expected r2 &#62; 0.98), that will be used with a known amount of ADP added to the respirometry chamber and the previously determined Kd values to determine the corresponding concentration of ATP11.</li>
	<li>Determine the rate of ATP synthesis, which is the slope of the concentration of ATP over time throughout the protocol (Figure 3).</li>
</ol>
5. Measurement of mitochondrial production of ROS&#160;using Amplex UltraRed (AmR)
<ol>
	<li>Use &#34;Green&#34; fluorescent sensors (530 nm dominant wavelength) to quantify the fluorescent signal from the respiration chamber. Energize the sensors at 300-400 mV; the resulting signal gets amplified with a gain of 1:1,000. Optimize specific settings for individual instruments to capture the expected signal within the linear range of the sensor. 
	NOTE: If using respiration media without MgCl2, then this should be added (20-60 &#181;L of 100 mM MgCl2 to produce 1-3 &#181;M MgCl2) prior to adding reagents for the AmR assay.</li>
	<li>Perform the chemical setup and initial calibration of the AmR assay prior to the addition of the mitochondria. Add 30 &#181;moles of DTPA (6 &#181;L of 5 mM solution) to chelate cations that might interfere with the reaction, then add superoxide dismutase (2 &#181;L of a 5,000 U/mL stock solution to convert superoxide anions to H2O2 for a more comprehensive detection of reactive oxygen generation), horseradish peroxidase (5 &#181;L of a 500 U/mL stock solution), and Amplex UltraRed (2 &#181;L of a 10 mM stock solution) (see Table of Materials) to produce 5 U/mL, 1 U/mL, and 10 &#181;M in the respirometry chamber, respectively.</li>
	<li>Allow the fluorescent signal to stabilize, then add 0.2 &#181;moles of hydrogen peroxide (5 &#181;L of a 40 &#181;M solution made fresh daily) twice, approximately 5 min apart. 
	NOTE: The fluorescent signal before and after the two titrations of H2O2 provides a three-point linear calibration curve of the system (expected r2 &#62; 0.95), the slope of which reflects the overall responsiveness of the system in reporting the relationship between the fluorescent signal and the production (or addition) of H2O2. Use the fluorescent signal calibration function in the machine control software to calibrate this signal.</li>
	<li>Perform additional two-point calibrations (5 &#181;L of a 40 &#181;M solution made fresh daily) throughout the assay, to allow adjustment of the responsiveness of the assay as the chemistry of the respirometry changes throughout the assay, with the specific timing of these calibration points at the discretion of the investigator (Figure 4).</li>
</ol>
6. Measurement of mitochondrial respiration
<ol>
	<li>Add 15 &#181;L of isolated mitochondria suspension (step 1.4) to each 2 mL incubation chamber, so that the results represent the mitochondrial yield of 18.75 mg of muscle. Seal the incubation chamber. Vortex the sample between each titration to maintain a uniform suspension of the sample.</li>
	<li>Measure the residual oxygen consumption (ROX) prior to the addition of any substrates. Subtract this value (typically less than 0.2 pmol O2 x s-1 x mL-1)&#160;from the oxygen consumption values of each step in the substrate/uncoupler/inhibitor titration (SUIT) protocol11 after completion of the protocol. 
	NOTE: It is critical that the signals from both the oxygen sensor and the fluorescence sensor are allowed to stabilize for at least 1 min (as assessed by the stable calculated slope of the primary sensor signals), as the overall state of respiration is changed by the titrations in order to obtain reliable results. This applies to all titration steps.</li>
	<li>Use a general-purpose SUIT that allows for the initial characterization of equine skeletal muscle mitochondrial function. Start with sequential titrations of pyruvate (5 &#181;L of 2 M aqueous solution), glutamate (10 &#181;L of 2 M aqueous solution), and malate (10 &#181;L of 0.4 M aqueous solution) (see Table of Materials) into each chamber to produce Nicotinamide adenine dinucleotide (NADH) and stimulate non-phosphorylating (leak) respiration supported by NADH oxidized through Complex I (LN) (Figure 1, Figure 3, and Figure 4).</li>
	<li>Add ADP (20 &#181;L of 500 mM aqueous solution; see Table of Materials) to stimulate phosphorylating respiration through Complex I (PN).</li>
	<li>Add succinate (20 &#181;L of 1 M aqueous solution; see Table of Materials) to produce phosphorylating respiration through the combination of Complex I and Complex II (PN+S). 
	NOTE: With the combination of respiration through both Complex I and Complex II, oxygen consumption may be high enough to consume most of the oxygen dissolved in the incubation media. If the O2 concentration decreases below 50 &#181;M, reoxygenate the incubation media by unsealing the incubation chamber until the measured O2 concentration has increased above 150 &#181;M. Repeat this step as necessary to maintain sufficient O2 for mitochondrial respiration.</li>
	<li>Add rotenone (2 &#181;L of 0.1 mM ethanol solution; see Table of Materials) to block Complex I. The resulting oxygen flux represents the capacity of Complex II to support mitochondrial oxygen consumption via the oxidation of succinate alone (PS). 
	CAUTION: Rotenone is a poisonous substance. Use standard laboratory safety and avoid ingestion or inhalation.</li>
	<li>Calculate the mean value for a given experimental condition across individual respirometry chambers containing aliquots of a single biopsy. 
	NOTE: Derived calculations, such as flux control ratios (FCRs), are valuable in identifying relative changes in different pathways, and include FCRLeak (LN/PN), FCRN (PN/PN+S), and FCRS (PS/PN+S). Calculated oxidative phosphorylation efficiency (1-LN/PN+S) provides an estimate of the overall impact of leak respiration adjusted for total respiratory capacity.</li>
</ol>

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A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Measurement+of+ADP-ATP+exchange+in+relation+to+mitochondrial+transmembrane+potential+and+oxygen+consumption.">Measurement of ADP-ATP exchange in relation to mitochondrial transmembrane potential and oxygen consumption.</a> Methods in Enzymology. 542, 333-348 (2014).</li><li>Krumschnabel, G., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Simultaneous+high-resolution+measurement+of+mitochondrial+respiration+and+hydrogen+peroxide+production.">Simultaneous high-resolution measurement of mitochondrial respiration and hydrogen peroxide production.</a> Methods in Molecular Biology. 1264, 245-261 (2015).</li><li>Lemieux, H., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mitochondrial+function+is+altered+in+horse+atypical+myopathy.">Mitochondrial function is altered in horse atypical myopathy.</a> Mitochondrion. 30, 35-41 (2016).</li><li>Houben, R., Leleu, C., Fraipont, A., Serteyn, D., Votion, D. 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E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Submaximal+exercise+training+improves+mitochondrial+efficiency+in+the+gluteus+medius+but+not+in+the+triceps+brachii+of+young+equine+athletes.">Submaximal exercise training improves mitochondrial efficiency in the gluteus medius but not in the triceps brachii of young equine athletes.</a> Scientific Reports. 7 (1), 14389 (2017).</li><li>White, S. H., Wohlgemuth, S., Li, C., Warren, L. K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Rapid+communication:+Dietary+selenium+improves+skeletal+muscle+mitochondrial+biogenesis+in+young+equine+athletes.">Rapid communication: Dietary selenium improves skeletal muscle mitochondrial biogenesis in young equine athletes.</a> Journal of Animal Science. 95 (9), 4078-4084 (2017).</li><li>Doerrier, C., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=High-resolution+FluoRespirometry+and+OXPHOS+protocols+for+human+cells,+permeabilized+fibers+from+small+biopsies+of+muscle,+and+isolated+mitochondria.">High-resolution FluoRespirometry and OXPHOS protocols for human cells, permeabilized fibers from small biopsies of muscle, and isolated mitochondria.</a> Methods in Molecular Biology. 1782, 31-70 (2018).</li><li>Li, C., White, S. H., Warren, L. K., Wohlgemuth, S. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Effects+of+aging+on+mitochondrial+function+in+skeletal+muscle+of+American+American+Quarter+Horses.">Effects of aging on mitochondrial function in skeletal muscle of American American Quarter Horses.</a> Journal of Applied Physiology. 121 (1), 299-311 (2016).</li><li>Komlodi, T., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Comparison+of+mitochondrial+incubation+media+for+measurement+of+respiration+and+hydrogen+peroxide+production.">Comparison of mitochondrial incubation media for measurement of respiration and hydrogen peroxide production.</a> Methods in Molecular Biology. 1782, 137-155 (2018).</li><li>Gnaiger, E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mitochondrial+physiology.">Mitochondrial physiology.</a> Bioenergetic Communications. , (2020).</li><li>Li Puma, L. 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The authors have no conflicts of interest related to this manuscript.

The addition of fluorescent signals to the standard output of the high-resolution respirometer provides valuable information regarding mitochondrial physiology, but meticulous calibration of the fluorescent signal is critical for quality data. The original protocols for the use of MgG suggest that the calibration curves generated while calculating magnesium-adenylate dissociation constants could be applied to subsequent assays4; however, the fluorescent signal from the MgG may not be not sufficien...

The mitochondria of eukaryotic cells produce the majority of the ATP used by the cells for work and maintenance1. A key step in the mitochondrial production of ATP is the conversion of oxygen to water, and thus the metabolic capacity of mitochondria and the associated cells is frequently quantified through the measurement of oxygen consumption2. However, mitochondrial physiology is more complex than the simple process of oxygen consumption, and reliance on this endpoint exclusively provides an incomplete assessment of the impact of mitochondrial function and dysfunction on cellular health. Full characterization of mitochondrial function requires the assessment of not only oxygen consumption, but also the production of ATP as well as reactive oxygen species (ROS).
Additional measures of key mitochondrial functions can be accomplished concurrently with the measurement of respiration through the use of specific fluorophores. Tetramethylrhodamine methylester (TMRM) is a cationic fluorophore that accumulates in the mitochondrial matrix in proportion to the mitochondrial transmembrane voltage potential, resulting in a decrease in fluorescent intensity due to this accumulation3. TMRM can be used as an indicator of relative changes in mitochondrial membrane potential, or can be used to quantify precise changes in transmembrane voltage with additional experiments to determine constants that allow conversion of the fluorescent signal to mV. Magnesium green (MgG) is a fluorophore that fluoresces when bound with Mg2+, and is used for measurements of ATP synthesis based on the differential affinity of ADP and ATP for magnesium divalent cation4. Investigators must determine the specific affinity/dissociation constants (Kd) for both ADP and ATP under specific analytical conditions to convert the changes in MgG fluorescence to a change in ATP concentration. Amplex UltraRed (AmR) is the fluorophore used to measure the production of hydrogen peroxide and other ROS&#160;during mitochondrial respiration5. The reaction between H2O2 and AmR (which is catalyzed by horseradish peroxidase) produces resorufin, which is detectable through fluorescence at 530 nM. Each of these assays can be added individually to assays of real-time mitochondrial respiration, to provide concurrent measurements of the respective aspects of mitochondrial physiology, thus providing a direct link between respiration and mitochondrial output.
Horses are capable of very high rates of mass-specific oxygen consumption, due in part to the very high mitochondrial content of equine skeletal muscle, making this tissue highly relevant for studying mitochondrial physiology. With the development of high-resolution respirometry, studies using this novel technology have helped define the contributions of equine skeletal muscle mitochondria to both the remarkable exercise capacity of horses and the pathophysiology of skeletal muscle diseases6,7,8,9,10,11,12,13,14. Studies of equine skeletal muscle mitochondrial function are particularly advantageous, as obtaining large amounts of this tissue is non-terminal. Thus, equine subjects can not only provide sufficient tissue for the complete characterization of mitochondrial function, but also serve as longitudinal controls for high-quality, mechanistic studies into mitochondrial physiology. For this reason, additional assays to quantify mitochondrial membrane potential, ATP synthesis, and the production of ROS&#160;that complement the measurement of oxygen consumption in this tissue have been developed, in order to provide a more robust characterization of mitochondrial physiology in equine skeletal muscle.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>ADP</td>
			<td>Sigma-Aldrich (MilliporeSigma)</td>
			<td>A5285</td>
			<td></td>
		</tr>
		<tr>
			<td>Amplex UltraRed</td>
			<td>Life Technologies</td>
			<td>A36006</td>
			<td></td>
		</tr>
		<tr>
			<td>ATP</td>
			<td>Sigma-Aldrich (MilliporeSigma)</td>
			<td>A2383</td>
			<td></td>
		</tr>
		<tr>
			<td>BSA</td>
			<td>Sigma-Aldrich (MilliporeSigma)</td>
			<td>A6003</td>
			<td></td>
		</tr>
		<tr>
			<td>Calcium carbonate</td>
			<td>Sigma-Aldrich (MilliporeSigma)</td>
			<td>C4830</td>
			<td></td>
		</tr>
		<tr>
			<td>CCCP</td>
			<td>Sigma-Aldrich (MilliporeSigma)</td>
			<td>C2759</td>
			<td></td>
		</tr>
		<tr>
			<td>DatLab 7.0</td>
			<td>Oroboros Inc</td>
			<td></td>
			<td>Software to operate O2K fluororespirometer</td>
		</tr>
		<tr>
			<td>Dithiothreitol</td>
			<td>Sigma-Aldrich (MilliporeSigma)</td>
			<td>D0632</td>
			<td></td>
		</tr>
		<tr>
			<td>DTPA</td>
			<td>Sigma-Aldrich (MilliporeSigma)</td>
			<td>D1133</td>
			<td></td>
		</tr>
		<tr>
			<td>EGTA</td>
			<td>Sigma-Aldrich (MilliporeSigma)</td>
			<td>E4378</td>
			<td></td>
		</tr>
		<tr>
			<td>Glutamate</td>
			<td>Sigma-Aldrich (MilliporeSigma)</td>
			<td>G1626</td>
			<td></td>
		</tr>
		<tr>
			<td>HEPES</td>
			<td>Sigma-Aldrich (MilliporeSigma)</td>
			<td>H7523</td>
			<td></td>
		</tr>
		<tr>
			<td>Horseradish peroxidase</td>
			<td>Sigma-Aldrich (MilliporeSigma)</td>
			<td>P8250</td>
			<td></td>
		</tr>
		<tr>
			<td>Hydrogen peroxide</td>
			<td>Sigma-Aldrich (MilliporeSigma)</td>
			<td>516813</td>
			<td>Must be made fresh daily prior to assay</td>
		</tr>
		<tr>
			<td>Imidazole</td>
			<td>Sigma-Aldrich (MilliporeSigma)</td>
			<td>I2399</td>
			<td></td>
		</tr>
		<tr>
			<td>K-MES</td>
			<td>Sigma-Aldrich (MilliporeSigma)</td>
			<td>M8250</td>
			<td></td>
		</tr>
		<tr>
			<td>Magnesium chloride hexahydrate</td>
			<td>Sigma-Aldrich (MilliporeSigma)</td>
			<td>M9272</td>
			<td></td>
		</tr>
		<tr>
			<td>Magnesium Green</td>
			<td>Thermo Fisher Scientific</td>
			<td>M3733</td>
			<td></td>
		</tr>
		<tr>
			<td>Malate</td>
			<td>Sigma-Aldrich (MilliporeSigma)</td>
			<td>M1000</td>
			<td></td>
		</tr>
		<tr>
			<td>Mannitol</td>
			<td>Sigma-Aldrich (MilliporeSigma)</td>
			<td>M9647</td>
			<td></td>
		</tr>
		<tr>
			<td>Mitochondrial isolation kit</td>
			<td>Sigma-Aldrich (MilliporeSigma)</td>
			<td>MITOISO1</td>
			<td></td>
		</tr>
		<tr>
			<td>O2K fluororespirometer</td>
			<td>Oroboros Inc</td>
			<td></td>
			<td>Multiple units required to run full spectrum of assays concurrently.</td>
		</tr>
		<tr>
			<td>Phosphocreatine</td>
			<td>Sigma-Aldrich (MilliporeSigma)</td>
			<td>P7936</td>
			<td></td>
		</tr>
		<tr>
			<td>Potassium hydroxide</td>
			<td>Sigma-Aldrich (MilliporeSigma)</td>
			<td>P1767</td>
			<td></td>
		</tr>
		<tr>
			<td>Potassium lactobionate</td>
			<td>Sigma-Aldrich (MilliporeSigma)</td>
			<td>L2398</td>
			<td></td>
		</tr>
		<tr>
			<td>Potassium phosphate</td>
			<td>Sigma-Aldrich (MilliporeSigma)</td>
			<td>P0662</td>
			<td></td>
		</tr>
		<tr>
			<td>Pyruvate</td>
			<td>Sigma-Aldrich (MilliporeSigma)</td>
			<td>P2256</td>
			<td>Must be made fresh daily prior to assay</td>
		</tr>
		<tr>
			<td>Rotenone</td>
			<td>Sigma-Aldrich (MilliporeSigma)</td>
			<td>R8875</td>
			<td></td>
		</tr>
		<tr>
			<td>Succinate</td>
			<td>Sigma-Aldrich (MilliporeSigma)</td>
			<td>S2378</td>
			<td></td>
		</tr>
		<tr>
			<td>Sucrose</td>
			<td>Sigma-Aldrich (MilliporeSigma)</td>
			<td>84097</td>
			<td></td>
		</tr>
		<tr>
			<td>Superoxide dismutase</td>
			<td>Sigma-Aldrich (MilliporeSigma)</td>
			<td>S8160</td>
			<td></td>
		</tr>
		<tr>
			<td>Taurine</td>
			<td>Sigma-Aldrich (MilliporeSigma)</td>
			<td>T0625</td>
			<td></td>
		</tr>
		<tr>
			<td>Titration pump</td>
			<td>Oroboros Inc</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Titration syringes</td>
			<td>Oroboros Inc</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>TMRM</td>
			<td>Sigma-Aldrich (MilliporeSigma)</td>
			<td>T5428</td>
			<td></td>
		</tr>
		<tr>
			<td>UCH biopsy needle</td>
			<td>Millenium Surgical Corp</td>
			<td>72-238067</td>
			<td>Available in a range of sizes</td>
		</tr>
	</tbody>
</table>

high-resolution fluoro-respirometry of equine skeletal muscle

Mitochondrial function-oxidative phosphorylation and the generation of reactive oxygen species-is critical in both health and disease. Thus, measuring mitochondrial function is fundamental in biomedical research. Skeletal muscle is a robust source of mitochondria, particularly in animals with a very high aerobic capacity, such as horses, making them ideal subjects for studying mitochondrial physiology. This article demonstrates the use of high-resolution respirometry with concurrent fluorometry, with freshly harvested skeletal muscle mitochondria, to quantify the capacity to oxidize substrates under different mitochondrial states and determine the relative capacities of distinct elements of mitochondrial respiration. Tetramethylrhodamine methylester is used to demonstrate the production of mitochondrial membrane potential resulting from substrate oxidation, including calculation of the relative efficiency of the mitochondria by calculating the relative membrane potential generated per unit of concurrent oxygen flux. The conversion of ADP to ATP results in a change in the concentration of magnesium in the reaction chamber, due to differing affinities of the adenylates for magnesium. Therefore, magnesium green can be used to measure the rate of ATP synthesis, allowing the further calculation of the oxidative phosphorylation efficiency (ratio of phosphorylation to oxidation [P/O]). Finally, the use of Amplex UltraRed, which produces a fluorescent product (resorufin) when combined with hydrogen peroxide, allows the quantification of reactive oxygen species production during mitochondrial respiration, as well as the relationship between ROS production and concurrent respiration. These techniques allow the robust quantification of mitochondrial physiology under a variety of different simulated conditions, thus shedding light on the contribution of this critical cellular component to both health and disease.

The proposed reference state is that of a healthy sedentary Thoroughbred (no increased fitness due to compulsory exercise) and a fresh muscle sample collected from the center of a postural muscle, containing a high percentage of mitochondria-rich type I skeletal muscle fibers and incubated under conditions approximating resting metabolism (i.e., 38 &#176;C and pH 7.0). Under these conditions, the investigator can expect LN values of 2.71 &#177; 0.90, PN values of 62.40 &#177; 26.22, PN+S ...

Watch this Scientific Journal Video about High-Resolution Fluoro-Respirometry of Equine Skeletal Muscle at JoVE.com

High-Resolution Fluoro-Respirometry of Equine Skeletal Muscle

Horses have an exceptional aerobic exercise capacity, making equine skeletal muscle an important tissue for both the study of equine exercise physiology as well as mammalian mitochondrial physiology. This article describes techniques for the comprehensive assessment of mitochondrial function in equine skeletal muscle.

Mitochondrial function-oxidative phosphorylation and the generation of reactive oxygen species-is critical in both health and disease. Thus, measuring ...

high-resolution-fluoro-respirometry-of-equine-skeletal-muscle

Research

JoVE Journal

Biology

1.1K Views.  Oklahoma State University. Horses have an exceptional aerobic exercise capacity, making equine skeletal muscle an important tissue for both the study of equine exercise physiology as well as mammalian mitochondrial physiology. This article describes techniques for the comprehensive assessment of mitochondrial function in equine skeletal muscle.

Video: High-Resolution Fluoro-Respirometry of Equine Skeletal Muscle

Adult and Embryonic Skeletal Muscle Microexplant Culture and Isolation of Skeletal Muscle Stem Cells

Cultured embryonic and adult skeletal muscle cells have a number of different uses. The micro-dissected explants technique described in this chapter is a robust and reliable method for isolating relatively large numbers of proliferative skeletal muscle cells from juvenile, adult or embryonic muscles as a source of skeletal muscle stem cells. The authors have used micro-dissected explant cultures to analyse the growth characteristics of skeletal muscle cells in wild-type and dystrophic muscles. Each of the components of tissue growth, namely cell survival, proliferation, senescence and differentiation can be analysed separately using the methods described here. The net effect of all components of growth can be established by means of measuring explant outgrowth rates. The micro-explant method can be used to establish primary cultures from a wide range of different muscle types and ages and, as described here, has been adapted by the authors to enable the isolation of embryonic skeletal muscle precursors.
Uniquely, micro-explant cultures have been used to derive clonal (single cell origin) skeletal muscle stem cell (SMSc) lines which can be expanded and used for in vivo transplantation. In vivo transplanted SMSc behave as functional, tissue-specific, satellite cells which contribute to skeletal muscle fibre regeneration but which are also retained (in the satellite cell niche) as a small pool of undifferentiated stem cells which can be re-isolated into culture using the micro-explant method.

The micro-dissected explants technique is a robust and reliable method for isolating proliferative skeletal muscle cells from juvenile, adult or embryonic muscles as a source of skeletal muscle stem cells. Uniquely, these cells have been clonally derived to produce skeletal muscle stem cell lines used for in vivo transplantation.

Cultured embryonic and adult skeletal muscle cells have a number of different uses. The micro-dissected explants technique described in this chapter is a ...

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 ...

Purification of Progenitors from Skeletal Muscle

Skeletal muscle contains multiple progenitor populations of distinct embryonic origins and developmental potential. Myogenic progenitors, usually residing in a "satellite cell position" between the myofiber plasma membrane and the laminin-rich basement membrane that ensheaths it, are self-renewing cells that are solely committed to the myogenic lineage1,2. We have recently described a second class of vessel associated progenitors that can generate myofibroblasts and white adipocytes, which responds to damage by efficiently entering proliferation and provides trophic support to myogenic cells during tissue regeneration3,4. One of the most trusted assays to determine the developmental and regenerative potential of a given cell population relies on their isolation and transplantation5-7. To this end we have optimized protocols for their purification by flow cytometry from enzymatically dissociated muscle, which we will outline in this article. The populations obtained using this method will contain either myogenic or fibro/adipogenic colony forming cells: no other cell types are capable of expanding in vitro or surviving in vivo delivery. However, when these populations are used immediately after the sort for molecular analysis (e.g qRT-PCR) one must keep in mind that the freshly sorted subsets may contain other contaminant cells that lack the ability of forming colonies or engrafting recipients.

Method for the enzymatic dissociation, surface labeling and purification by flow cytometry of fibro/adipogenic and myogenic progenitors from murine skeletal muscle.

Skeletal muscle contains multiple progenitor populations of distinct embryonic origins and developmental potential. Myogenic progenitors, usually residing ...

Ex Vivo Assessment of Contractility, Fatigability and Alternans in Isolated Skeletal Muscles

Described here is a method to measure contractility of isolated skeletal muscles. Parameters such as muscle force, muscle power, contractile kinetics, fatigability, and recovery after fatigue can be obtained to assess specific aspects of the excitation-contraction coupling (ECC) process such as excitability, contractile machinery and Ca2+ handling ability. This method removes the nerve and blood supply and focuses on the isolated skeletal muscle itself. We routinely use this method to identify genetic components that alter the contractile property of skeletal muscle though modulating Ca2+ signaling pathways. Here, we describe a newly identified skeletal muscle phenotype, i.e., mechanic alternans, as an example of the various and rich information that can be obtained using the in vitro muscle contractility assay. Combination of this assay with single cell assays, genetic approaches and biochemistry assays can provide important insights into the mechanisms of ECC in skeletal muscle.

Ex Vivo Assessment of Contractility, Fatigability and Alternans in Isolated Skeletal Muscles

We describe a method to directly measure muscle force, muscle power, contractile kinetics and fatigability of isolated skeletal muscles in an in vitro system using field stimulation. Valuable information on Ca2+ handling properties and contractile machinery of the muscle can be obtained using different stimulating protocols.

 
Described here is a method to measure contractility of isolated skeletal muscles. Parameters such as muscle force, muscle power, contractile kinetics, ...

Laser-inflicted Injury of Zebrafish Embryonic Skeletal Muscle

Various experimental approaches have been used in mouse to induce muscle injury with the aim to study muscle regeneration, including myotoxin injections (bupivacaine, cardiotoxin or notexin), muscle transplantations (denervation-devascularization induced regeneration), intensive exercise, but also murine muscular dystrophy models such as the mdx mouse (for a review of these approaches see 1). In zebrafish, genetic approaches include mutants that exhibit muscular dystrophy phenotypes (such as runzel2 or sapje3) and antisense oligonucleotide morpholinos that block the expression of dystrophy-associated genes4. Besides, chemical approaches are also possible, e.g. with Galanthamine, a chemical compound inhibiting acetylcholinesterase, thereby resulting in hypercontraction, which eventually leads to muscular dystrophy5. However, genetic and pharmacological approaches generally affect all muscles within an individual, whereas the extent of physically inflicted injuries are more easily controlled spatially and temporally1. Localized physical injury allows the assessment of contralateral muscle as an internal control. Indeed, we recently used laser-mediated cell ablation to study skeletal muscle regeneration in the zebrafish embryo6, while another group recently reported the use of a two-photon laser (822 nm) to damage very locally the plasma membrane of individual embryonic zebrafish muscle cells7.
Here, we report a method for using the micropoint laser (Andor Technology) for skeletal muscle cell injury in the zebrafish embryo. The micropoint laser is a high energy laser which is suitable for targeted cell ablation at a wavelength of 435 nm. The laser is connected to a microscope (in our setup, an optical microscope from Zeiss) in such a way that the microscope can be used at the same time for focusing the laser light onto the sample and for visualizing the effects of the wounding (brightfield or fluorescence). The parameters for controlling laser pulses include wavelength, intensity, and number of pulses.
Due to its transparency and external embryonic development, the zebrafish embryo is highly amenable for both laser-induced injury and for studying the subsequent recovery. Between 1 and 2 days post-fertilization, somitic skeletal muscle cells progressively undergo maturation from anterior to posterior due to the progression of somitogenesis from the trunk to the tail8, 9. At these stages, embryos spontaneously twitch and initiate swimming. The zebrafish has recently been recognized as an important vertebrate model organism for the study of tissue regeneration, as many types of tissues (cardiac, neuronal, vascular etc.) can be regenerated after injury in the adult zebrafish10, 11.

The method presented here comprises the precise injury of live zebrafish embryos with high-energy laser pulses and the subsequent analysis of these injuries and their recovery with time. We also show how genetically labeled single or groups of skeletal muscle cells can be tracked during and after laser light induced damage.

Various experimental approaches have been used in mouse to induce muscle injury with the aim to study muscle regeneration, including myotoxin injections ...

Assessment of Calcium Sparks in Intact Skeletal Muscle Fibers

Maintaining homeostatic Ca2+ signaling is a fundamental physiological process in living cells. Ca2+ sparks are the elementary units of Ca2+ signaling in the striated muscle fibers that appear as highly localized Ca2+ release events mediated by ryanodine receptor (RyR) Ca2+ release channels on the sarcoplasmic reticulum (SR) membrane. Proper assessment of muscle Ca2+ sparks could provide information on the intracellular Ca2+&#160;handling properties of healthy and diseased striated muscles. Although Ca2+ sparks events are commonly seen in resting cardiomyocytes, they are rarely observed in resting skeletal muscle fibers; thus there is a need for methods to generate and analyze sparks in skeletal muscle fibers.
Detailed here is an experimental protocol for measuring Ca2+ sparks in isolated flexor digitorm brevis (FDB) muscle fibers using fluorescent Ca2+ indictors and laser scanning confocal microscopy. In this approach, isolated FDB fibers are exposed to transient hypoosmotic stress followed by a return to isotonic physiological solution. Under these conditions, a robust Ca2+ sparks response is detected adjacent to the sarcolemmal membrane in young healthy FDB muscle fibers. Altered Ca2+ sparks response is detected in dystrophic or aged skeletal muscle fibers. This approach has recently demonstrated that membrane-delimited signaling involving cross-talk between inositol (1,4,5)-triphosphate receptor (IP3R) and RyR contributes to Ca2+ spark activation in skeletal muscle. In summary, our studies using osmotic stress induced Ca2+ sparks showed that this intracellular response reflects a muscle signaling mechanism in physiology and aging/disease states, including mouse models of muscle dystrophy (mdx mice) or amyotrophic lateral sclerosis (ALS model).

Described here is a method to directly measure calcium sparks, the elementary units of Ca2+ release from sarcoplasmic reticulum&#160;in intact skeletal muscle fibers. This method utilizes osmotic-stress-mediated triggering of Ca2+ release from ryanodine receptor in isolated muscle fibers. The dynamics and homeostatic capacity of intracellular Ca2+ signaling can be employed to assess muscle function in health and disease.

Maintaining homeostatic Ca2+ signaling is a fundamental physiological process in living cells. Ca2+ sparks are the elementary units of Ca2+ signaling in ...

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

Mitochondria are involved in cellular energy metabolism and use oxygen to produce energy in the form of adenosine triphosphate (ATP). Differential centrifugation at low- and high-speed is commonly used to isolate mitochondria from tissues and cultured cells. Crude mitochondrial fractions obtained by differential centrifugation are used for respirometry measurements. The differential centrifugation technique is based on the separation of organelles according to their size and sedimentation velocity. The isolation of mitochondria is performed immediately after tissue harvesting. The tissue is immersed in an ice-cold homogenization medium, minced using scissors and homogenized in a glass homogenizer with a loose-fitting pestle. The differential centrifugation technique is efficient, fast and inexpensive and the mitochondria obtained by differential centrifugation are pure enough for respirometry assays. Some of the limitations and disadvantages of isolated mitochondria, based on differential centrifugation, are that the mitochondria can be damaged during the homogenization and isolation procedure and that large amounts of the tissue biopsy or cultured cells are required for the mitochondrial isolation.

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

Mitochondria are involved in cellular energy metabolism and use oxygen to produce energy in the form of adenosine triphosphate (ATP). Differential ...

Isolation and Characterization of Exosomes from Skeletal Muscle Fibroblasts

Exosomes are small extracellular vesicles released by virtually all cells and secreted in all biological fluids. Many methods have been developed for the isolation of these vesicles, including ultracentrifugation, ultrafiltration, and size exclusion chromatography. However, not all are suitable for large scale exosome purification and characterization. Outlined here is a protocol for establishing cultures of primary fibroblasts isolated from adult mouse skeletal muscles, followed by purification and characterization of exosomes from the culture media of these cells. The method is based on the use of sequential centrifugation steps followed by sucrose density gradients. Purity of the exosomal preparations is then validated by western blot analyses using a battery of canonical markers (i.e., Alix, CD9, and CD81). The protocol describes how to isolate and concentrate bioactive exosomes for electron microscopy, mass spectrometry, and uptake experiments for functional studies. It can easily be scaled up or down and adapted for exosome isolation from different cell types, tissues, and biological fluids.

This protocol illustrates the 1) the isolation and culture of primary fibroblasts from the adult mouse gastrocnemius muscle as well as 2) purification and characterization of exosomes using a differential ultracentrifugation method combined with sucrose density gradients followed by western blot analyses.

Exosomes are small extracellular vesicles released by virtually all cells and secreted in all biological fluids. Many methods have been developed for the ...

Construction of a Realistic, Whole-Body, Three-Dimensional Equine Skeletal Model using Computed Tomography Data

Therapies based upon whole-body biomechanical assessments are successful for injury prevention and rehabilitation in human athletes. Similar approaches have rarely been used to study equine athletic injury. Degenerative osteoarthritis caused by mechanical stress can originate from chronic postural dysfunction, which, because the primary dysfunction is often distant from the site of tissue injury, is best identified through modeling whole-body biomechanics. To characterize whole-body equine kinematics, a realistic skeletal model of a horse was created from equine computed tomography (CT) data that can be used for functional anatomical and biomechanical modeling. Equine CT data were reconstructed into individual three-dimensional (3D) data sets (i.e., bones) using 3D visualization software and assembled into a complete 3D skeletal model. The model was then rigged and animated using 3D animation and modeling software. The resulting 3D skeletal model can be used to characterize equine postures associated with degenerative tissue changes as well as to identify postures that reduce mechanical stress at the sites of tissue injury. In addition, when animated into 4D, the model can be used to demonstrate unhealthy and healthy skeletal movements and can be used to develop preventative and rehabilitative individualized therapies for horses with degenerative lamenesses. Although the model will soon be available for download, it is currently in a format that requires access to the 3D animation and modeling software, which has quite a learning curve for new users. This protocol will guide users in (1) developing such a model for any organism of interest and (2) using this specific equine model for their own research questions.

The purpose of this protocol is to describe the method of creation of a realistic, whole-body, skeletal model of a horse that can be used for functional anatomical and biomechanical modeling to characterize whole-body mechanics.

Therapies based upon whole-body biomechanical assessments are successful for injury prevention and rehabilitation in human athletes. Similar approaches ...

Electroporation of Plasmid DNA into Mouse Skeletal Muscle

Transient gene expression modulation in murine skeletal muscle by plasmid electroporation is a useful tool for assessing normal and pathological physiology. Overexpression or knockdown of target genes enables investigators to manipulate individual molecular events and, thus, better understand the mechanisms that impact muscle mass, muscle metabolism, and contractility. In addition, electroporation of DNA plasmids that encode fluorescent tags allows investigators to measure changes in subcellular localization of proteins in skeletal muscle in vivo. A key functional assessment of skeletal muscle includes the measurement of muscle contractility. In this protocol, we demonstrate that whole muscle contractility studies are still possible after plasmid DNA injection, electroporation, and gene expression modulation. The goal of this instructional procedure is to demonstrate the step-by-step method of DNA plasmid electroporation into mouse skeletal muscle to facilitate uptake and expression in the myofibers of the tibialis anterior and extensor digitorum longus muscles, as well as to demonstrate that skeletal muscle contractility is not compromised by injection and electroporation.

Electroporation of plasmid DNA into skeletal muscle is a viable method to modulate gene expression without compromising muscle contractility in mice.