This work was supported by RO1- AG028614 to JM, RO1- AR063084to NLW, and RO1-AR057404 to JZ.

1. Setting up Osmotic-stress Perfusion System
Figure 1 is a schematic protocol of calcium sparks assessment in intact skeletal muscle fibers.
<ol>
	<li>Set up a three-axis (xyz) micromanipulator capable of positioning the outlet tip of the perfusion system containing a minimum of two channels. This could be made with disposable Luer-syringe barrel with attached three-way Luer-Lok&#160;stopcock to switch on and/or off of the flow of perfusion solutions. These perfusion channels should be capable of delivering &#62;1 ml/min of solution through a single perfusion tip with a &#62;0.2 mm diameter.</li>
	<li>Load two channels of the perfusion system, one with Isotonic Tyrode Solution and the other with Hypotonic Tyrode Solution.</li>
</ol>
2. Preparing Intact Single Flexor Digitorum Brevis (FDB) Muscle Fibers from Mouse
<ol>
	<li>Remove the foot from a mouse euthanized following NIH and IACUC guidelines using a pair of heavy dissecting scissors by cutting through the leg above the ankle joint.</li>
	<li>Pin the foot onto a dissection chamber filled with Minimal Ca2+ Tyrode Solution with the plantar surface facing up.</li>
	<li>Dissect FDB by cutting the tendon and gently pulling up on the tendon to separate the muscle from the surrounding tissue.</li>
	<li>Finish dissection by cutting the distal tendon where it branches into individual digits in the deep layers of the muscle.</li>
	<li>Transfer the FDB muscle by picking up with forceps via its tendons to avoid additional damage to the muscle fibers and place into a tube with a thawed aliquot of Collagenase Digestion Solution prewarmed to 37 &#176;C.</li>
	<li>Incubate the tube containing FDB upright on an orbital shaker at 37 &#176;C for 60-90 min with a speed of 160 rpm. This muscle bundles dissociation protocol needs to be experimentally examined for different strains of animals, especially for those disease/mutant fibers vulnerable to membrane damage.</li>
	<li>Transfer the digested FDB into tube with 700 &#956;l of Isotonic Tyrode Solution and gently triturate to release the fibers by drawing the muscle several times through a series of 200 ml-micropipette tips of gradually decreasing diameter via cutting with a clean razor blade to remove the tips of 200 ml&#160;micropipette. To avoid damaging fibers in particularly weak muscles from disease, make sure the tip is just large enough to allow the fiber bundle to pass through without sticking. Do not force the bundle through too small an opening.</li>
	<li>Plate out the FDB fibers by gently tapping and resuspending FDB fibers in the tube and then drawing out 70 ml (1/10 of total fibers) with a cut 200 ml&#160;micropipette into the center of a 35&#160;mm Delta TPG dish containing 1 ml of Isotonic Tyrode Solution to reduce diffusion across the dish.</li>
	<li>Determine the number of intact single fibers in the dish using a dissection microscope. Add additional aliquots of FDB fibers to obtain 3-4 intact fibers on the dish.</li>
	<li>Store additional isolated muscle fibers at 4 &#176;C and use within 6 hr.</li>
</ol>
3. Fluo-4 AM Dye Loading and Ca2+ Imaging (Sparks Measurement)
<ol>
	<li>Transfer 500 ml of Isotonic Tyrode Solution from dish with plated FDB fibers into a tube with 10 ml Fluo-4 AM stock (1 mM), mix and add back to the dish to a final Fluo-4 AM concentration of 10 mM. Alternatively, pluronic acid can be used to increase dye loading efficiency.</li>
	<li>Load for 60 min at room temperature in the dark.</li>
	<li>Wash the fibers by carefully drawing out 500 ml of Fluo-4 AM-containing Isotonic Tyrode Solution from the dish with an uncut 200 ml-plastic micropipette tip and replace with equal volume of fresh Isotonic Tyrode Solution.</li>
	<li>Repeat this process 3 more times.</li>
	<li>To visualize mitochondrial function, transfer 500 ml of Isotonic Tyrode Solution from dish with FDB fibers into a tube with 5 ml TMRE stock (1 mM), mix, and add back to the dish for a final concentration of 50 nM.</li>
	<li>Incubate at room temperature for 10 min.</li>
	<li>Wash the fibers by carefully drawing out 500 ml of TMRE-containing Isotonic Tyrode Solution from the dish with an uncut 200 ml&#160;plastic micropipette tip and replace with equal volume of fresh Isotonic Tyrode Solution.</li>
	<li>Repeat this process 3 more times.</li>
	<li>Transfer the dish to the confocal microscope and select an intact fiber with clear striations and a smooth sarcolemmal membrane for experimentation.</li>
	<li>Position the tip of the perfusion system 400-500 &#181;m away from the target muscle fiber and off center using micromanipulator controls.</li>
	<li>Begin flow of the Isotonic Tyrode Solution to ensure the FDB fiber stays in place.</li>
	<li>Record the fluorescence signal from Fluo-4 AM with a 40X oil immersion objective and excite Fluo-4 AM with an argon laser (excitation wavelength 488 nm) and record emission signals (wavelength 510-580 nm). The intensity of the fluorescent signal is proportional to the relative level of intracellular Ca2+ concentration ([Ca2+]i).</li>
	<li>Induce Ca2+ transients by changing the osmotic pressure of the extracellular solution in order to produce osmotic stress on fiber. Initially perfuse the fibers with Isotonic Tyrode solution (290 mOsM) (baseline collection for 60 sec) and then with Hypotonic Tyrode solution (170 mOsM) for 100 sec to induce swelling. Perfuse the FDB fibers back with Isotonic Tyrode solution. Generally, only one osmotic shock is applied to a single intact fiber in one delta TPG dish and the spark events last 10 min for data acquisition.</li>
	<li>Record spatial localization of Ca2+ sparks (Figure 2) using a time-series of xy two-dimensional images with scan speed of 166 lps (line per second, and each line comprised of 512 pixels resulting in 3.08 sec/frame ) for each condition of isotonic (1 min), hypotonic stress (100 sec) and post-hypotonic stress (5 min). Store signals for offline analysis to evaluate the distribution and frequency of Ca2+ sparks following completion of the experiment.</li>
	<li>Find a new fiber on a new dish and follow the same osmotic shock protocol to induce Ca2+ transients. Use a confocal line scan (xt) mode (512 pixels in length, sample rate of 2 msec/line scan to record Ca2+ transients for one minute (i.e. 30,000 lines) with the confocal scan line placed directly underneath sarcolemmal membrane (Figure 3). Change the line scan to different focal planes to record three sequential series (at 1-min intervals) of linescans for analysis of the magnitude and kinetics of individual Ca2+ sparks.</li>
</ol>
4. Data Analysis
<ol>
	<li>Collect a large number of xt line scan traces to evaluate the morphology and kinetics of individual Ca2+ sparks parameters, i.e.&#160;the amplitude (F/F0; where F0 is the resting Ca2+ fluorescence), rise time, time to the peak, duration (FDHM, full duration at half magnitude), and spatial width (FWHM, full width at half magnitude) of the recorded sparks. These parameters represent the basic gating properties of RYR Ca2+ channels such as the Ca2+ flux (amplitude), and gating kinetics of RYRs (duration).</li>
	<li>Analyze digital image data with a custom-developed, semi-automatic algorithm which was created with image-processing language IDL (Interactive Data Language), as described previously11,16. Because the unique kinetics of Ca2+ release in the case of Ca2+ sparks induced in FDB fibers by osmotic stress, it is better to combine the manual and automated features of Ca2+ spark events identification and processing with these custom-build image analysis programs11. This algorithm is very useful when it's necessary to manually identify ROI (region of interest) in some muscle disease models. Once a ROI is defined, the algorithm could automatically derive the above mentioned spark parameters which could then be exported to Excel file. Alternatively, the public available image processing software, e.g. SparkMaster in ImageJ, can be used to define the kinetic properties of Ca2+ sparks and their spatial distribution in skeletal muscle17.</li>
</ol>

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J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Ca2++sparks+as+a+plastic+signal+for+skeletal+muscle+health,+aging,+and+dystrophy.">Ca2+ sparks as a plastic signal for skeletal muscle health, aging, and dystrophy.</a> Acta. 27 (7), 791-798 (2006).</li><li>Weisleder, N., Zhou, J., Ma, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Detection+of+calcium+sparks+in+intact+and+permeabilized+skeletal+muscle+fibers.">Detection of calcium sparks in intact and permeabilized skeletal muscle fibers.</a> Methods Mol. 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No conflicts of interest declared.

This method of assessing Ca2+ sparks in intact skeletal muscle is a useful tool for muscle physiology and disease research. We showed that the Ca2+&#160;spark response was altered in different conditions, including muscular dystrophy11, aging18,19,&#160;as well as in amyotrophic lateral sclerosis20. Our recent study also revealed functional coupling between IP3 receptor and RyR representing a critical component that contributes to Ca<...

Intracellular free Ca2+ ([Ca2+]i) is a versatile and important secondary messenger that regulates multiple cellular functions in excitable cells such as neurons, cardiac, skeletal and smooth muscles (for review see&#160;Stutzmann&#160;and Mattson1). Regulated Ca2+ mobilization and cross-talk between sarcoplasmic reticulum (SR) and T-tubule (TT) membranes are fundamental regulators of muscle physiology. Furthermore, changes in Ca2+ signaling had been shown to be an underlying mechanism of contractile dysfunction in various muscle diseases.
Ca2+ sparks are localized elementary Ca2+ release events originating from opening of the ryanodine receptor (RyR) channel on the sarcoplasmic reticulum (SR) membrane2. In cardiac muscle, sparks occur spontaneously through opening of the RyR2 channel by a Ca2+-induced Ca2+&#160;release (CICR) mechanism 3-5. In skeletal muscle, RyR1 is strictly controlled by the voltage sensor at the TT membrane6,7. Thus Ca2+ sparks are suppressed and rarely detected in resting conditions in intact skeletal muscle fibers. Until recently, the sarcolemmal membrane needed to be disrupted by various chemical or mechanical &#34;skinning&#34; methods to uncouple the suppression of the voltage sensor on RyR1 and allowed for Ca2+ spark events to be detected in skeletal muscle8,9. One method previously described required permeabilization of the membrane of muscle fibers by saponin detergent10.
In 2003, we discovered that either transient hypoosmotic stress or hyperosmotic stress could induce peripheral Ca2+ sparks adjacent to the sarcolemmal membrane in intact muscle fibers11. This method has since been modified to study the molecular mechanism and modulation of Ca2+ release and dynamics12-16. Here we outline the details of our experimental approach for induction, detection and analysis of Ca2+ sparks in intact skeletal muscle. We also present our custom-built spark analysis program that can be used to quantify the elemental properties of individual Ca2+ sparks in skeletal muscle, e.g. spark frequency and amplitude (&#916; F/F0, which reflects the open probability of the RyR channels and the Ca2+ load inside the SR); time to peak (rise time) and duration (FDHM, full duration at half-maximal amplitude) of sparks, as well as the spatial distribution of Ca2+ sparks. In addition, we present evidence that links altered Ca2+ sparks to the various pathophysiological states in skeletal muscle, such as muscular dystrophy and amyotrophic lateral sclerosis.
The advantage of this technique involves the ability to measure Ca2+ in relatively undamaged cells, instead of stripping the muscle fibers, allowing recording of Ca2+ sparks in conditions closer to physiologic. Additionally, our custom-designed program provides more accurate calculations of the properties of the spark in relation to muscle fibers.

<table border="0" cellpadding="0" cellspacing="0" width="790">
	<colgroup>
		<col />
		<col />
		<col />
		<col />
	</colgroup>
	<tbody>
		<tr height="21">
			<td height="21" style="height:21px;width:195px;">Dissecting microscope</td>
			<td style="width:111px;"></td>
			<td style="width:119px;"></td>
			<td style="width:367px;">Dissect out fibers and check muscle integrity</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:195px;">Dissection tools -scissors, and fine tip forceps</td>
			<td style="width:111px;">Fine Science Tools</td>
			<td style="width:119px;"></td>
			<td></td>
		</tr>
		<tr height="84">
			<td height="84" style="height:84px;width:195px;">Sylgard 184 Silicone Elastomer Kit</td>
			<td>DOW CORNING</td>
			<td style="width:119px;">184 SIL ELAST KIT</td>
			<td style="width:367px;">&#160;Make ahead of time for fiber dissection. Mix 5 ml liquid Sylgard components and pour into a 60-mm glass Petri dish and waiting 48 hr to let the Sylgard compound become clear, colorless solid substrate.&#160;</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">60-mm glass petri dish&#160;</td>
			<td style="width:111px;">Pyrex</td>
			<td style="width:119px;">3160</td>
			<td>Fiber dissection</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Isotonic Tyrode Solution&#160;</td>
			<td>Sigma-Aldrich</td>
			<td style="width:119px;"></td>
			<td></td>
		</tr>
		<tr height="23">
			<td height="23" style="height:23px;">Minimal Ca2+ Tyrode Solution</td>
			<td>Sigma-Aldrich</td>
			<td style="width:119px;"></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Hypotonic Tyrode Solution&#160;</td>
			<td>Sigma-Aldrich</td>
			<td style="width:119px;"></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:195px;">collagenase type I&#160;</td>
			<td>Sigma-Aldrich</td>
			<td>C-5138</td>
			<td>Fiber digestion&#160;</td>
		</tr>
		<tr height="23">
			<td height="23" style="height:23px;">Fluo-4 AM&#160;</td>
			<td style="width:111px;">Invitrogen</td>
			<td>F14201</td>
			<td>Fluorescent Ca2+ imaging dyes&#160;</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">TMRE</td>
			<td style="width:111px;">Invitrogen</td>
			<td>T669</td>
			<td>mitochondrial membrane potential fluorescent dyes</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Delta TPG dishes&#160;</td>
			<td>Bioptechs</td>
			<td style="width:119px;">0420041500C</td>
			<td>35 mm glass bottom dish for imaging</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:195px;">Spark Fit software</td>
			<td style="width:111px;"></td>
			<td style="width:119px;"></td>
			<td>data analysis software</td>
		</tr>
		<tr height="84">
			<td height="84" style="height:84px;width:195px;">Radiance 2100 laser scanning confocal microscope or equivalent microscope</td>
			<td style="width:111px;">Bio-Rad</td>
			<td style="width:119px;"></td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:195px;">3 axis micromanipulator</td>
			<td style="width:111px;">Narishige International</td>
			<td style="width:119px;">MHW-3</td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:195px;">temperature controllable orbital shaker</td>
			<td style="width:111px;">New Brunswick scientific</td>
			<td style="width:119px;"></td>
			<td>Fiber dissociation</td>
		</tr>
	</tbody>
</table>

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).

Earlier studies showed that transient hypoosmotic stress induced peripheral Ca2+ sparks adjacent to the sarcolemmal membrane in intact muscle fibers11. Figure 1 shows the images of intact single muscle fibers with smooth sarcolemmal membrane and characteristic clear striations. Figure 2 shows typical Ca2+ sparks (as xy images) were induced by transient (100 sec) treatment with a hypoosmotic solution that swells the FDB muscle fibers from young, h...

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Assessment of Calcium Sparks in Intact Skeletal Muscle Fibers

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 ...

assessment-of-calcium-sparks-in-intact-skeletal-muscle-fibers

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JoVE Journal

Biology

15.1K 조회수.  The Ohio State University Wexner Medical Center.  직접 칼슘 불꽃을 측정하는 방법이 여기에 설명, 칼슘 2 +의 기본 단위는 그대로 골격 근육 섬유의 근질 세망에서 놓습니다. 이 방법은 고립 된 근육 섬유의 ryanodine 수용체에서 칼슘 2 + 릴리스의 삼투 스트레스 매개 트리거를 사용합니다. 역학과 내 Ca 2 + 신호 전달의 항상성 용량은 건강과 질환에서 근육 기능을 평가하기 위해 사용될 ?...

비디오: Assessment of Calcium Sparks in Intact Skeletal Muscle Fibers

Respirometric Oxidative Phosphorylation Assessment in Saponin-permeabilized Cardiac Fibers

Investigation of mitochondrial function represents an important parameter of cardiac physiology as mitochondria are involved in energy metabolism, oxidative stress, apoptosis, aging, mitochondrial encephalomyopathies and drug toxicity. Given this, technologies to measure cardiac mitochondrial function are in demand. One technique that employs an integrative approach to measure mitochondrial function is respirometric oxidative phosphorylation (OXPHOS) analysis.
The principle of respirometric OXPHOS assessment is centered around measuring oxygen concentration utilizing a Clark electrode. As the permeabilized fiber bundle consumes oxygen, oxygen concentration in the closed chamber declines. Using selected substrate-inhibitor-uncoupler titration protocols, electrons are provided to specific sites of the electron transport chain, allowing evaluation of mitochondrial function. Prior to respirometric analysis of mitochondrial function, mechanical and chemical preparatory techniques are utilized to permeabilize the sarcolemma of muscle fibers. Chemical permeabilization employs saponin to selectively perforate the cell membrane while maintaining cellular architecture.
This paper thoroughly describes the steps involved in preparing saponin-skinned cardiac fibers for oxygen consumption measurements to evaluate mitochondrial OXPHOS. Additionally, troubleshooting advice as well as specific substrates, inhibitors and uncouplers that may be used to determine mitochondria function at specific sites of the electron transport chain are provided. Importantly, the described protocol may be easily applied to cardiac and skeletal tissue of various animal models and human samples.

Saponin-permeabilized fiber preparation in conjunction with respirometric oxidative phosphorylation analysis provides integrative assessment of mitochondrial function. Mitochondrial respiration in physiological and pathological states can reflect various regulatory influences including mitochondrial interactions, morphology and biochemistry.

Investigation of mitochondrial function represents an important parameter of cardiac physiology as mitochondria are involved in energy metabolism, ...

Fluorescence-based Measurement of Store-operated Calcium Entry in Live Cells: from Cultured Cancer Cell to Skeletal Muscle Fiber

Store operated Ca2+ entry (SOCE), earlier termed capacitative Ca2+ entry, is a tightly regulated mechanism for influx of extracellular Ca2+ into cells to replenish depleted endoplasmic reticulum (ER) or sarcoplasmic reticulum (SR) Ca2+ stores1,2. Since Ca2+ is a ubiquitous second messenger, it is not surprising to see that SOCE plays important roles in a variety of cellular processes, including proliferation, apoptosis, gene transcription and motility. Due to its wide occurrence in nearly all cell types, including epithelial cells and skeletal muscles, this pathway has received great interest3,4. However, the heterogeneity of SOCE characteristics in different cell types and the physiological function are still not clear5-7.
The functional channel properties of SOCE can be revealed by patch-clamp studies, whereas a large body of knowledge about this pathway has been gained by fluorescence-based intracellular Ca2+ measurements because of its convenience and feasibility for high-throughput screening. The objective of this report is to summarize a few fluorescence-based methods to measure the activation of SOCE in monolayer cells, suspended cells and muscle fibers5,8-10. The most commonly used of these fluorescence methods is to directly monitor the dynamics of intracellular Ca2+ using the ratio of F340nm and F380nm (510 nm for emission wavelength) of the ratiometric Ca2+ indicator Fura-2. To isolate the activity of unidirectional SOCE from intracellular Ca2+ release and Ca2+ extrusion, a Mn2+ quenching assay is frequently used. Mn2+ is known to be able to permeate into cells via SOCE while it is impervious to the surface membrane extrusion processes or to ER uptake by Ca2+ pumps due to its very high affinity with Fura-2. As a result, the quenching of Fura-2 fluorescence induced by the entry of extracellular Mn2+ into the cells represents a measurement of activity of SOCE9. Ratiometric measurement and the Mn+2 quenching assays can be performed on a cuvette-based spectrofluorometer in a cell population mode or in a microscope-based system to visualize single cells. The advantage of single cell measurements is that individual cells subjected to gene manipulations can be selected using GFP or RFP reporters, allowing studies in genetically modified or mutated cells. The spatiotemporal characteristics of SOCE in structurally specialized skeletal muscle can be achieved in skinned muscle fibers by simultaneously monitoring the fluorescence of two low affinity Ca2+ indicators targeted to specific compartments of the muscle fiber, such as Fluo-5N in the SR and Rhod-5N in the transverse tubules9,11,12.

The extent of store-operated Ca2+ entry (SOCE) can be monitored using fluorescent Ca2+ indicators. Mn2+ quenching of such indicators assays SOCE in cultured cells and skeletal muscle fibers. A technique allowing spatial and temporal resolution of SOCE by confocal imaging of mechanically skinned muscle fibers is also described.

Store operated Ca2+ entry (SOCE), earlier termed capacitative Ca2+ entry, is a tightly regulated mechanism for influx of extracellular Ca2+ into cells to ...

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, ...

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 ...

High-Throughput Contractile Measurements of Hydrogel-Embedded Intact Mouse Muscle Fibers Using an Optics-Based System

In vitro cell culture is a powerful tool to assess cellular processes and test therapeutic strategies. For skeletal muscle, the most common approaches involve either differentiating myogenic progenitor cells into immature myotubes or the short-term ex vivo culture of isolated individual muscle fibers. A key benefit of ex vivo culture over in vitro is the retention of the complex cellular architecture and contractile characteristics. Here, we detail an experimental protocol for the isolation of intact flexor digitorum brevis muscle fibers from mice and their subsequent ex vivo culture. In this protocol, muscle fibers are embedded in a fibrin-based and basement membrane matrix hydrogel to immobilize the fibers and maintain their contractile function. We then describe methods to assess the muscle fiber contractile function using an optics-based, high-throughput contractility system. The embedded muscle fibers are electrically stimulated to induce contractions, after which their functional properties, such as sarcomere shortening and contractile velocity, are assessed using optics-based quantification. Coupling muscle fiber culture with this&#160;system allows for high-throughput testing of the effects of pharmacological agents on contractile function and ex vivo studies of genetic muscle disorders. Finally, this protocol can also be adapted to study dynamic cellular processes in muscle fibers using live-cell microscopy.

Skeletal muscle function can be assessed by quantifying the contractility of isolated muscle fibers, traditionally using laborious, low-throughput approaches. Here, we describe an optics-based, high-throughput method to quantify the contractility of hydrogel-embedded muscle fibers. This approach has applications for drug screening and therapeutic development.

In vitro cell culture is a powerful tool to assess cellular processes and test therapeutic strategies. For skeletal muscle, the most common approaches ...

Functional Site-Directed Fluorometry in Native Skeletal Muscle Cells

Functional Site-Directed Fluorometry in Native Cells to Study Skeletal Muscle Excitability

Functional site-directed fluorometry has been the technique of choice to investigate the structure-function relationship of numerous membrane proteins, including voltage-gated ion channels. This approach has been used primarily in heterologous expression systems to simultaneously measure membrane currents, the electrical manifestation of the channels&#39; activity, and fluorescence measurements, reporting local domain rearrangements. Functional site-directed fluorometry combines electrophysiology, molecular biology, chemistry, and fluorescence into a single wide-ranging technique that permits the study of real-time structural rearrangements and function through fluorescence and electrophysiology, respectively. Typically, this approach requires an engineered voltage-gated membrane channel that contains a cysteine that can be tested by a thiol-reactive fluorescent dye. Until recently, the thiol-reactive chemistry used for the site-directed fluorescent labeling of proteins was carried out exclusively in Xenopus oocytes and cell lines, restricting the scope of the approach to primary non-excitable cells. This report describes the applicability of functional site-directed fluorometry in adult skeletal muscle cells to study the early steps of excitation-contraction coupling, the process by which muscle fiber electrical depolarization is linked to the activation of muscle contraction. The present protocol describes the methodologies to design and transfect cysteine-engineered voltage-gated Ca2+ channels (CaV1.1) into muscle fibers of the flexor digitorum brevis of adult mice using in vivo electroporation and the subsequent steps required for functional site-directed fluorometry measurements. This approach can be adapted to study other ion channels and proteins. The use of functional site-directed fluorometry of mammalian muscle is particularly relevant to studying basic mechanisms of excitability.

Author Spotlight: Functional Site-Directed Fluorometry in Native Cells to Study Skeletal Muscle Excitability  

Functional site-directed fluorometry is a method to study protein domain motions in real time. Modification of this technique for its application in native cells now allows the detection and tracking of single voltage-sensor motions from voltage-gated Ca2+ channels in murine isolated skeletal muscle fibers.

Functional site-directed fluorometry has been the technique of choice to investigate the structure-function relationship of numerous membrane proteins, ...

Quantification of Skeletal Muscle Fatty Infiltration via Decellularization

Decellularization-Based Quantification of Skeletal Muscle Fatty Infiltration

Fatty infiltration is the accumulation of adipocytes between myofibers in skeletal muscle and is a prominent feature of many myopathies, metabolic disorders, and dystrophies. Clinically in human populations, fatty infiltration is assessed using noninvasive methods, including computed tomography (CT), magnetic resonance imaging (MRI), and ultrasound (US). Although some studies have used CT or MRI to quantify fatty infiltration in mouse muscle, costs and insufficient spatial resolution remain challenging. Other small animal methods utilize histology to visualize individual adipocytes; however, this methodology suffers from sampling bias in heterogeneous pathology. This protocol describes the methodology to qualitatively view and quantitatively measure fatty infiltration comprehensively throughout intact mouse muscle and at the level of individual adipocytes using decellularization. The protocol is not limited to specific muscles or specific species and can be extended to human biopsy. Additionally, gross qualitative and quantitative assessments can be made with standard laboratory equipment for little cost, making this procedure more accessible across research laboratories.

Author Spotlight: Decellularization-Based Quantification of Skeletal Muscle Fatty Infiltration  

The present study describes decellularization-based methodologies for visualizing and quantifying intramuscular adipose tissue (IMAT) deposition through intact muscle volume, as well as quantifying metrics of individual adipocytes that comprise IMAT.

Fatty infiltration is the accumulation of adipocytes between myofibers in skeletal muscle and is a prominent feature of many myopathies, metabolic ...

Construct Design for Transfecting Cysteine-Engineered Voltage-Gated Ca2+ Channels in Murine Isolated Skeletal Muscle Fibers

Construct Design for Transfecting Cysteine-Engineered Voltage-Gated Ca2+ Channels in Murine Isolated Skeletal Muscle Fibers  

To begin, generate a wild-type, fluorescently-tagged CaV1.1 CDNA construct. Here, a plasmid coating for the enhanced green fluorescent protein or EGFP-tagged alpha subunit of rabbit Cav1.1 is used. The presence of the cytomegalovirus promoter allows strong transfection efficiency in muscle fibers.Introduce cystine substitution in the channel that would be tracked with fluorometry using a commercially available, site-directed mutagenesis kit. Here the cysteines are inserted at a position corresponding to the membrane extracellular interface of one of the four S4 of the channel. With the help of GFP observation and ionic current recording, confirm that neither the inserted cysteines nor the presence thiol-reactive dye impacts the voltage dependence and the conductance of the channel.

Decellularization of Skeletal Muscle 

To begin, inspect the muscles of interest dissected from the sacrificed mouse for the absence of bone chips or ragged edges and trim them away with sharp scissors if needed. Weigh the cleaned muscles using an analytical balance and record their weight. After placing the muscle in at least 0.1 milliliters of 1%SDS per milligram of weight, place the well plate on a rocking shaker set to 50 to 80 Hertz.Visually inspect the SDS solution periodically. When the solution becomes cloudy, remove the solution with a pipette without aspirating the muscles and replace it with an equal volume of fresh 1%SDS. When the solution remains clear for 24 hours, remove the final SDS solution from the decellularized muscles and replace it with an equal volume of PBS.Inspect the decellularized muscles under a stereo microscope and carefully remove any hair or debris stuck to the muscle using forceps. Carefully remove the PBS and replace it with an equal volume of 3.7%formaldehyde before returning the plate to the rocking shaker for 24 hours.

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 ...