This work was supported by R01AR075773 to GAM.

The care and sacrifice of mice were performed in accordance with the National Institutes of Health Guide for the Use and Care of Laboratory Animals. All work was approved by the Animal Studies Committee at Washington University in the St. Louis School of Medicine. Male C57BL/6J mice aged 2-3 months (see Table of Materials) were used to generate the example images included in this protocol. All steps described below are performed at room temperature.
1. Muscle decellularization
<ol>
	<li>Prepare a 1% w/v sodium dodecyl sulfate (SDS; see Table of Materials) solution in phosphate-buffered saline (PBS). Stir the solution until completely mixed. 
	NOTE: 1% SDS can be prepared in bulk and stored at room temperature for 1 month.</li>
	<li>Dissect the muscle(s) of interest as described previously38,39.
	<ol>
		<li>Expose the muscles of interest by wetting the hair and skin with 70% ethanol solution, then making a transcutaneous incision of approximately 2 cm followed by removing the covering skin with forceps and spring scissors. 
		NOTE: The muscle(s) of interest here include the tibialis anterior (TA), extensor digitorum longus (EDL), and diaphragm. Some muscles may lie deep in other musculature, requiring dissection of other muscles (e.g., dissecting the EDL requires removal of the TA).</li>
		<li>Dissect the muscle(s) of interest using sharp scissors or blades (see Table of Materials) to ensure that the entire extent of the muscle is obtained and the edges are smooth. 
		NOTE: This is ideally done under a dissecting microscope to improve visualization.</li>
		<li>Inspect the muscles to ensure that no bone chips or ragged edges are included, then trim these away with sharp scissors as needed.</li>
		<li>Weigh the muscle using an analytical balance and record the weight.</li>
	</ol>
	</li>
	<li>Place the dissected muscle in at least 0.1 mL of 1% SDS per milligram of weight; 6-, 12- or 24-well plates work well for the diaphragm, TA, and EDL mouse muscles, respectively. Typical volumes are 6 mL, 3 mL, and 1.5 mL for 6-, 12- and 24-well plates, respectively. 
	NOTE: The SDS solution will sometimes cause the muscles to deform, so ensure that the muscle is flat/extended at initial immersion.</li>
	<li>Place the well plate (or other vessels) on a rocking shaker set to 50-80 Hz.</li>
	<li>Visually inspect the SDS solution periodically. When the solution becomes cloudy, remove the solution with a pipette (taking care not to aspirate the muscle) and replace it with an equal volume of fresh 1% SDS. When the solution remains clear for 24 h, decellularization is complete. 
	NOTE: The frequency required for solution changes varies by muscle size and the initial solution volume. Larger muscles like the TA need new SDS within a few hours, and smaller muscles like the EDL can go overnight in the original solution.</li>
	<li>Remove the final SDS solution from the decellularized muscles with a pipette (taking care not to aspirate the muscle) and replace it with an equal volume of PBS.</li>
	<li>Visually inspect the decellularized muscles under a dissecting microscope and use forceps and scissors to remove any hair or debris that is stuck to the muscle. 
	NOTE: Also, note the clarity of the decellularized muscle in this step. If the decellularized muscle is not fully transparent and the SDS solution does not increase in cloudiness over 24 h, then the decellularization was not efficient. This creates an artifact in qualitative and quantitative assessments, and so decellularization should always be optimized on practice samples before proceeding with experimental muscles.</li>
	<li>Carefully remove the PBS, replace it with an equal volume of 3.7% formaldehyde or 4% paraformaldehyde solution, and return the plate to the rocking shaker for 24 h. 
	&#8203;NOTE: Ensure that the formaldehyde solution fully covers the decellularized muscle, otherwise staining will be uneven.</li>
</ol>
2. Visualization of IMAT with oil red O
<ol>
	<li>Prepare a solution of ORO.
	<ol>
		<li>Dissolve 0.5 g of ORO powder (see Table of Materials) in 100 mL of isopropanol to generate a stock solution. 
		NOTE: Add gentle heat to the solution to fully dissolve it. This stock can be stored at room temperature for 1 month.</li>
		<li>Combine the ORO stock solution and deionized water at a 60:40 ratio to obtain the working solution needed for all muscles using a volume of at least 0.1 mL/ mg muscle weight. Typical volumes are 6 mL, 3 mL, and 1.5 mL for 6-, 12- and 24-well plates, respectively. 
		NOTE: Inspect the stock solution prior to mixing. If the stock solution contains substantial particulate, make fresh stock.</li>
		<li>Cover the working solution for 10 min to allow the particulate to settle.</li>
		<li>Filter the working solution through a 40 &#181;m mesh, followed by a 0.22 &#181;m syringe filter. 
		NOTE: The 0.22 &#181;m syringe filter clogs rapidly and must be exchanged if the working solution does not push easily through.</li>
	</ol>
	</li>
	<li>Remove the formaldehyde or paraformaldehyde solution and wash the decellularized muscles with three solution changes of an equal volume of PBS.</li>
	<li>Replace the PBS with an equal volume of 60% isopropanol solution and incubate on a rocking shaker for 5 min.</li>
	<li>Replace the 60% isopropanol solution with ORO working solution and incubate on the rocking shaker for 10 min. 
	NOTE: Ensure that the ORO working solution fully covers the decellularized muscle, otherwise staining will be uneven.</li>
	<li>Replace the ORO working solution with an equal volume of 1% SDS. Visually inspect the SDS solution periodically. When the solution becomes noticeably pink, remove the solution with a pipette (taking care not to aspirate the muscle) and replace it with fresh 1% SDS.</li>
	<li>When the solution remains clear for 24 h, replace the 1% SDS with PBS and inspect the stained muscle under a dissecting/stereo microscope at 4x magnification. Remove any obvious debris or particulate stuck to the outside of the muscle. If this is extensive, the decellularized muscle can be rolled gently on a cleaning tissue to pull off the debris/particulate.</li>
	<li>If the staining is satisfactory (bright red spheres floating in a transparent matrix), acquire images of the staining as desired using a camera attached to the stereo microscope (see Table of Materials). 
	&#8203;NOTE: If the dissecting microscope doesn&#39;t have an attached camera, a phone camera can be used to take pictures through the eyepiece.</li>
</ol>
3. Visualization of IMAT lipid droplets with BODIPY
NOTE: Confocal imaging is most effective with thin muscles like the EDL or diaphragm (~2 mm thickness). Alternatively, comparable thickness strips of muscles like the TA could be used.
<ol>
	<li>Prepare a 1:200 solution of fluorescent BODIPY 493/503 (see Table of Materials) in PBS to obtain a working solution volume of at least 0.1 mL/ mg muscle weight. Typical volumes are 6 mL, 3 mL, and 1.5 mL for 6-, 12- and 24-well plates, respectively.</li>
	<li>Remove the formaldehyde, paraformaldehyde, or 1% SDS and wash the decellularized muscles with three solution changes of an equal volume of PBS.</li>
	<li>Replace the PBS with the BODIPY working solution and incubate on the rocking shaker for 20 min.</li>
	<li>Wash the decellularized muscles with three solution changes of an equal volume of PBS.</li>
	<li>Place the muscle in a clear-bottomed vessel that is compatible with an available confocal microscope. Ideally, the dish will have a recessed bottom to place a coverslip over the muscle without deforming it (see Table of Materials).</li>
	<li>Obtain image stacks with a standard confocal microscope (see Table of Materials) using the 488 laser. Typical stack sizes for an EDL are 0.5-1 mm with a 10 &#181;m slice thickness, yielding 50-100 images per stack. 
	NOTE: To quantify total lipid volume, the total number of IMAT lipid droplets, and the nearest neighbor index of clustering, image through the entire muscle volume, taking care to leave some overlap for image registration. To quantify the average lipid droplet volume, one stack may be sufficient.</li>
	<li>If desired, stain the muscles with ORO, as per section 2, for a complimentary whole-muscle image of IMAT distribution. 
	&#8203;NOTE: Because BODIPY is fluorescent, it won&#39;t be visible under the light microscope when images of ORO are acquired.</li>
</ol>
4. Estimation of total lipid volume by lipid extraction
<ol>
	<li>Following image acquisition, transfer the muscles to 200 &#181;L of isopropanol in individual wells of a 96-well plate, or adapt the well/volume if the muscle is too large to fit.</li>
	<li>Agitate the solution by tapping the plate, pipetting the solution up and down, and/or mechanically crushing the muscle with a pipette tip until no red/fluorescent spheres can be seen in the decellularized muscle under a microscope. 
	NOTE: Treat all the muscles with the same combination of tapping, pipetting, and crushing for the best reliability. Also, take care to limit the time spent tapping, pipetting, and crushing, as the isopropanol will evaporate rapidly.</li>
	<li>Mix the solution in each well by pipetting up and down, then transfer 75 &#181;L to two clean wells of the plate.</li>
	<li>Cover the plate and read the absorbance of the duplicate 75 &#181;L wells using a spectrophotometer or plate reader. If the construct was stained with ORO, read the solution at 500 nm; if it was stained with BODIPY 493/503, read the plate at 493 nm. 
	NOTE: If the construct was stained with both BODIPY and ORO, it is recommended to read the plate at 500 nm, as 500 nm reads give similar results between constructs stained with ORO only and ORO plus BODIPY.</li>
	<li>Divide the absorbance reading by the recorded weight of the muscle if desired to correct for size differences between samples.</li>
</ol>
5. Quantification of IMAT lipid droplet metrics from confocal images
NOTE: This section require access to ImageJ version 1.47 (see Table of Materials) or later and basic ImageJ skills40.
<ol>
	<li>Open confocal stacks in ImageJ. 
	NOTE: Different confocal microscopes may save confocal images in different formats. ImageJ may require an additional plugin or variant, such as Fiji, to open the stacks41. The algorithms used are part of the ImageJ software package.</li>
	<li>Run a thresholding algorithm in ImageJ to identify BODIPY positive pixels. This converts the current image to a binary image.
	<ol>
		<li>Open the threshold user interface by selecting Image &#62; Adjust &#62; Threshold. In the user interface, select Intermodes as the thresholding type and ensure Dark background is selected. No other options need to be selected. Then click on Apply.</li>
		<li>A dialog box opens to convert the stack to binary. Ensure that it has the following options selected: Method: Intermodes; Background: Dark. Calculate the threshold for each image and black background (of binary masks). This generates a binary stack, where white indicates BODIPY positive pixels and black indicates BODIPY negative pixels. 
		NOTE: The Intermodes thresholding algorithm may not be the best choice for all users. Subjective inspection of the built-in thresholding options of ImageJ helps to select the optimal algorithm for separating BODIPY positive and negative pixels.</li>
	</ol>
	</li>
	<li>Run the Watershed algorithm in ImageJ to separate touching lipid droplets. Select Process &#62; Binary &#62; Watershed. A dialog box opens asking whether all images in the stack should be processed. Select Yes. This adds thin black lines dividing larger areas of solid white.</li>
	<li>Identify ROIs with the Analyze Particles algorithm in ImageJ.
	<ol>
		<li>Select Analyze &#62; Analyze Particles. A dialog box opens to set the selection settings. 
		NOTE: Selecting the size range is critical to achieving the best estimate of lipid droplet ROIs. The lower bound eliminates regions that are likely too small to be IMAT-derived lipid droplets (background artifact), and the upper bound eliminates regions that are likely too large to be single IMAT-derived lipid droplets (touching lipid droplets that weren&#39;t separated by the Watershed algorithm).</li>
		<li>To set these values, open the original image and outline the smallest and largest lipid droplet in view using the Oval tool. Then, add these shapes to the ROI manager by typing &#34;t&#34;. Select Analyze, then Set Measurements. A dialog box opens for selection of the settings.
		<ol>
			<li>Check Area only and click OK. Then, select Measure from the ROI Manger. Use the two area measures from this Results window as the size range in Analyze Particles.</li>
		</ol>
		</li>
		<li>Ensure Add to Manager is checked. No other options are needed.</li>
	</ol>
	</li>
	<li>Overlay the ROIs on the confocal stack by checking Show All in the ROI Manager. Use the slider bar to inspect each stack slice individually and add missing regions by hand using the Oval tool (located in the ImageJ toolbar).</li>
	<li>Output the ROI measurements. Select Analyze &#62; Set Measurements and select Area, Centroid, Fit ellipse and Stack position. Click on OK. Then, select Measure from the ROI Manger. The data in the Results table can be selected and copied into Matlab or Excel for further analysis.</li>
	<li>Refine the ROIs in each stack to a single ROI using the Refine.m Matlab code37 or a similar algorithm. 
	NOTE: This step is required as steps 5.2-5.4 identify the same lipid droplet in adjacent slices as distinct ROIs. However, ROIs can alternatively be identified by hand only, or duplicate ROIs can be eliminated by hand in ImageJ to avoid the need for Matlab.</li>
	<li>Obtain the summary statistics in the ROI using Matlab or Excel.
	<ol>
		<li>Estimate the total number of lipid droplets as the total number of ROIs.</li>
		<li>Estimate lipid droplet volume as the volume of the ellipse fitted to each 2D ROI, assuming the depth of the shape is the average of the major and minor axes of the ellipse.</li>
		<li>Estimate the total lipid volume, which is the sum of individual estimated lipid droplet volumes.</li>
	</ol>
	</li>
</ol>

Quantification of Skeletal Muscle Fatty Infiltration via Decellularization

The authors have no conflicts of interest to disclose.

This manuscript describes methods to qualitatively visualize and quantify skeletal muscle fatty infiltration in small animal models that can be applied to further understanding the pathogenesis of intramuscular adipose tissue (IMAT) development and pathological expansion. The use of whole-muscle decellularization and lipid-soluble staining allows for a cost-effective, reproducible, and simple methodology to comprehensively assess the presence of IMAT in whole muscles. 
The basis for this protocol is that decellularization of muscle with SDS removes the cellular components of myofibers, including the small lipid droplets of IMCL, but spares the large lipid droplets in intramyocellular adipocytes. SDS has been used extensively42 in tissue engineering to decellularize matrices. Tissues such as adipose and skeletal muscle typically require additional mechanical dissociation and/or alcohol extraction to remove the residual adipocyte lipid42,43. We have previously shown that this is because while decellularization with SDS eliminates IMCL, it spares the large lipid droplet in adipocytes37. Imaging of osmium tetroxide-stained intact muscle pre- and post-decellulariztion with &#181;CT verified that the spatial pattern of IMAT was not disrupted by decellularization. Further, intramuscular triglyceride quantification in a decellularized muscle with negligible IMAT was ~5% of the intact muscle values, verifying the removal of IMCL. Therefore, this methodology retains IMAT lipid droplets in their original anatomical distribution through a semi-transparent muscle matrix.
Proper decellularization is the most critical step in this protocol. If the decellularization is incomplete, IMAT lipid droplets will be difficult to visualize and residual IMCL will cause high background staining with either ORO or BODIPY (Figure 2). Common errors by inexperienced users are inadequate SDS coverage per muscle (within each well), such that each muscle is not completely covered in SDS solution, not using a rocker to agitate the solution during decellularization, and not performing solution changes frequently enough. In this manuscript, we have recommended the amount of SDS needed per unit muscle mass, but the user will still need to ensure that muscles are completely covered by solutions, as each muscle has a unique geometry. Users are also recommended to change the solutions liberally (as much as twice per day) to ensure that decellularization is complete. Good quality staining of IMAT lipid droplets has been achieved after as many as 4 days of SDS treatment. For high-quality ORO staining results, adequate fixation and ORO solution prep are also important. Similar to the SDS treatment described above, adequate coverage of 3.7% formaldehyde solution for each muscle sample is needed. If the muscle is removed from the fixative too early, lipid droplets will only weakly stain with ORO. A total of 1-2 h should be sufficient, but overnight fixation is recommended to ensure the fixative penetrates to the center of the muscle and fully fixes all lipid droplets. An additional challenge with ORO staining is that when the alcohol concentration is reduced to 60%, a particulate begins to form. This particulate can settle on the surface and become stuck on the border of muscle. The best way to avoid this is to make a fresh working solution for each staining and use both 40 mesh &#181;m and 0.22 &#181;m filters. Then, maintaining agitation with the rocker and limiting the staining time to 10 min will help keep any particulate that forms from settling. If the problem persists, making a fresh ORO stock solution may help. If some artifact remains stuck to the decellularized muscle surface, a stereo microscope, forceps, and surgical scissors can be used to remove this artifact. Failing to eliminate artifacts will impact the image quality of muscles and overestimate IMAT content during the lipid extraction portion in preparation for OD reading. 
Overall, this technique is straightforward and offers several advantages over gold standard methods for visualizing and quantifying skeletal muscle fatty infiltration. Noninvasive techniques, such as CT, MRI and US, which are used extensively in humans and sometimes in animal models, have limited spatial resolution and are unable to distinguish lipid droplets from muscle fibers. Thus, a pixel or voxel of intermediate signal intensity is assigned as "muscle" or "fat", while in actuality it is likely a mix of myofibers and adipocytes. More commonly, fatty infiltration in animal muscle is assessed by histology, most frequently by ORO in muscle cryosections. However, this is typically only performed in a single representative section and is difficult to quantify due to lipid scatter over the section. By contrast, ORO staining of an entire decellularized muscle provides a comprehensive assessment of IMAT with similar costs and effort as intact morphology. Furthermore, in addition to enhancing visualization, ORO staining of decellularization enables the quantification of fatty infiltration by lipid extraction. For a deeper dive into the features of fatty infiltration, a fluorescent stain, BODIPY, can be used in conjunction with confocal microscopy. This enables the reconstruction of individual IMAT lipid droplets to map the 3D landscape, which is not possible with histology unless sections are analyzed over the length of the muscle. While a confocal microscope is not standard lab equipment, it is more likely to be accessible in a university or industry setting than small animal MRI or CT. Furthermore, much of this process can be automated, reducing the time cost compared with sequential histology. Optimizing the settings on the confocal microscope is an additional consideration for BODIPY staining. These are unique to each microscope. The critical value is laser intensity, which must be high enough to detect the lipid droplets on the distant surface of the muscle while also not saturating the signal from the lipid droplets on the near side. Because of this, it is suggested that using BODIPY staining with confocal microscopy is best suited on thinner muscles, including the EDL or diaphragm. 
Several limitations of this approach warrant discussion. First, while it is anticipated that this technique has broad applicability beyond injury models (cardiotoxin and glycerol) in mice presented here, new applications (e.g., the mdx model) may require optimization, as the size and composition of the muscle (e.g., fibrosis) could affect decellularization, requiring increased SDS concentration or incubation times. Other disease models with altered muscle mass would also require analysis of both absolute and normalized (to muscle mass) metrics of fatty infiltration to determine the absolute amount of lipid or percentage of lipid relative to the muscle volume to provide a more meaningful outcome measure. Furthermore, this technique is anticipated to be broadly applicable to larger animal models and human biopsies, but this may require optimization for each new application. Second, in this strategy, the entire muscle must be dedicated to this assay and cannot be used to assess another pathological feature. Studies that aim to assess longitudinal changes in IMAT are better served with noninvasive imaging techniques and studies whose primary aim requires the muscle for other purposes (histology, quantitative polymerase chain reaction, western blotting) are better served by histological assessment, as the remainder of the frozen muscle can be allocated to other assays. However, this assay is well suited to pair with in vivo testing, such as treadmill running, or ex vivo contractile testing ,since these measures can be made before decellularization44. Third, although the use of BODIPY stain with confocal microscopy provides high-resolution visualization and quantification of lipid droplets, it cannot conclusively identify lipid droplets as individual adipocytes, as the cell membrane is removed and endogenous adipocyte proteins are lost. Multilocular adipocytes, representing immature adipocytes or a "brown/beige" phenotype, may be identified as multiple lipid droplets. Finally, the protocol does not work well on previously frozen muscle. These limitations are probably most profound for human biopsies, as while the entire biopsy can be decellularized, the spatial distribution of IMAT in the biopsy is not likely to be more representative of the whole muscle than a histological slice. However, since this technique is relatively insensitive to unfrozen biopsy handling conditions (e.g., hours on ice in PBS), the biopsy could be divided later for various assays, including a portion for decellularization, which would provide a better resolution of individual lipid droplets.
In conclusion, a novel method for qualitative and quantitative analysis of skeletal muscle fatty infiltration has been developed by staining and imaging the retained lipid of decellularized constructs. This methodology offers improvements over gold-standard approaches, in that it enables comprehensive imaging of three-dimensional fatty infiltration within muscle and quick, cheap quantification with ORO staining. For more detailed measures, a second lipid-soluble BODIPY stain provides a more detailed quantification of lipid droplet number, volume, and distribution pattern, as imaged by confocal microscopy. Together, these measures provide researchers with a way to precisely measure skeletal muscle fatty infiltration at the level of the individual lipid droplets without sampling or expensive noninvasive imaging.

The accumulation of adipocytes between myofibers within the skeletal muscle is a prominent feature of disparate conditions, from type 2 diabetes to sarcopenia to musculoskeletal injury1,2,3,4,5,6,7. Comprehensive assessment of this intramuscular adipose tissue (IMAT) is critical to understanding the pathogenesis of these conditions, as IMAT deposition is strongly correlated with insulin resistance3,8,9,10 and poor skeletal muscle function11,12,13,14,15. Although these associations have been noted for decades, the mechanisms associated with and the origin of IMAT still remain an area of intense investigation. This is partly because most studies assessing skeletal muscle fatty infiltration have been performed in humans, where mechanistic investigations are limited16,17. However, more recently, small animal models, including mice, have been utilized to help pinpoint cellular regulation of IMAT development and signaling18,19,20. This work aims to provide a new tool for use with small animal models for qualitatively visualizing and quantifying skeletal muscle fatty infiltration.
Clinically in human populations, fatty infiltration is assessed using noninvasive methods, including computed tomography (CT)6,21, magnetic resonance imaging (MRI)16,17,22,23, and ultrasound (US)17,24. These imaging techniques typically identify a defined region of interest (ROI) in a muscle and acquire image slices within that region, although comprehensive approaches have also been employed25,26,27. These image slices are subjected to qualitative grading6 and quantified via pixel thresholding28. Similar approaches have been utilized in animals previously29,30; however, they are costly and require access to small animal imaging systems. Spatial resolution via CT and MRI use also presents a major issue, as they are unable to delineate IMAT adipocytes from skeletal muscle fibers within a voxel and instead rely on subjective separation of primarily muscle regions and primarily IMAT regions31,32. As such, the inability to accurately identify fat or muscle tissue also presents inaccurate quantification of representative amounts of these tissues.
Due to these limitations, current techniques to assess skeletal muscle fatty infiltration in small animal models most commonly rely on histology as an inexpensive and accessible alternative33,34. Standard staining procedures, including hematoxylin and eosin (H&#38;E), oil red O (ORO), and immunostaining for adipocyte markers such as perilipin, allow for simple detection and visualization of adipocytes comprising fatty infiltration within the muscle. However, histology approaches are rarely comprehensive, and typically, IMAT qualitative or quantitative assessment is limited to a single section34. Lipid extraction has also been used to quantify total muscle lipids35; however, this technique fails to distinguish between intramyocellular lipid (IMCL) and intramuscular adipose tissue (IMAT) stores36. In summary, current methodologies to visualize and quantify fat in muscle remain limited either by financial costs or the specific detection of IMAT.
Here, we describe a detailed method for assessing skeletal muscle fatty infiltration both by qualitative visualization and multi-scale quantification. This methodology employs a simple decellularization technique that removes myocellular structures, including IMCL, but keeps the larger IMAT adipocyte-derived lipid droplets intact. Validation of the specificity of this technique has been published37, including using lipid extraction to show the depletion of IMCL with decellularization, &#181;CT to show the retention of IMAT patterning with decellularization, and histology to show the similar size distribution of IMAT lipid droplets compared with those identified with decellularization. Once decellularized, muscles can be stained with lipid-soluble dyes for qualitative visualization of the pattern and the extent of fatty infiltration and/or quantitative imaging of individual IMAT lipid droplets. Dyes can subsequently be extracted with isopropanol, and the optical density (OD) of the resulting solution can be used to estimate the IMAT lipid volume. The stringent validation of this technique has been published elsewhere37. This article provides a detailed protocol to use this methodology with mouse muscles and provides troubleshooting tips to support the adoption of this method in other applications, such as muscles from other species or other tissues.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>0.22 &#181;m Syringe Filter</td>
			<td>Fisher Scientific</td>
			<td>SLGP033RS</td>
			<td></td>
		</tr>
		<tr>
			<td>1 mL LuerLock Syringes</td>
			<td>Fisher Scientific</td>
			<td>14823434</td>
			<td></td>
		</tr>
		<tr>
			<td>12 mm Coverslips</td>
			<td>Fisher Scientific</td>
			<td>12545F</td>
			<td></td>
		</tr>
		<tr>
			<td>12 well plates</td>
			<td>Fisher Scientific</td>
			<td>08-772-29</td>
			<td></td>
		</tr>
		<tr>
			<td>24 well plates</td>
			<td>Fisher Scientific</td>
			<td>08-772-1H</td>
			<td></td>
		</tr>
		<tr>
			<td>2-Propanol (Isopropanol)</td>
			<td>Sigma Aldrich</td>
			<td>I9516</td>
			<td>0.5 mg/mL stock solution can be stored at room temperature for 1 month.&#160; Working solution must be made fresh.</td>
		</tr>
		<tr>
			<td>37% Formaldehyde Solution</td>
			<td>Sigma Aldrich</td>
			<td>8187081000</td>
			<td></td>
		</tr>
		<tr>
			<td>40 &#181;m Mesh Filter</td>
			<td>Fisher Scientific</td>
			<td>87711</td>
			<td></td>
		</tr>
		<tr>
			<td>6 well plates</td>
			<td>Fisher Scientific</td>
			<td>08-772-1B</td>
			<td></td>
		</tr>
		<tr>
			<td>96 well plates</td>
			<td>Fisher Scientific</td>
			<td>08-772-2C</td>
			<td></td>
		</tr>
		<tr>
			<td>BODIPY 493/503</td>
			<td>Fisher Scientific</td>
			<td>D-3922</td>
			<td></td>
		</tr>
		<tr>
			<td>C57BL/6J Mice</td>
			<td>Jackson Laboratory</td>
			<td>000664</td>
			<td></td>
		</tr>
		<tr>
			<td>Confocal Imaging Dish</td>
			<td>VWR</td>
			<td>734-2905</td>
			<td></td>
		</tr>
		<tr>
			<td>Confocal Microscope</td>
			<td>Leica</td>
			<td>TCS SPEII</td>
			<td></td>
		</tr>
		<tr>
			<td>Dissecting/stereo Microscope</td>
			<td>Zeiss</td>
			<td>4107009123001000</td>
			<td></td>
		</tr>
		<tr>
			<td>Dissection scissors</td>
			<td>Fine Science Tools</td>
			<td>14060-09</td>
			<td></td>
		</tr>
		<tr>
			<td>Dumont #5 forceps</td>
			<td>Fine Science Tools</td>
			<td>11254-20</td>
			<td></td>
		</tr>
		<tr>
			<td>Ethanol</td>
			<td>Fisher Scientific</td>
			<td>033361.K2</td>
			<td></td>
		</tr>
		<tr>
			<td>ImageJ</td>
			<td>NIH</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Matlab</td>
			<td>Mathworks</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Oil Red O Powder</td>
			<td>Sigma Aldrich</td>
			<td>O0625</td>
			<td></td>
		</tr>
		<tr>
			<td>Plate reader</td>
			<td>Bio-tek</td>
			<td>Synergy II&#160;</td>
			<td></td>
		</tr>
		<tr>
			<td>Rocker/Shaker</td>
			<td>Reliable Scientific</td>
			<td>55D</td>
			<td></td>
		</tr>
		<tr>
			<td>Sodium Dodecyl Sulfate (SDS)</td>
			<td>Sigma Aldrich</td>
			<td>L3771</td>
			<td>1% Solution can be stored at room temperature for 1 month</td>
		</tr>
		<tr>
			<td>Transfer pipettes</td>
			<td>Fisher Scientific</td>
			<td>137119D</td>
			<td></td>
		</tr>
		<tr>
			<td>Vannas spring scissors</td>
			<td>Fine Science Tools</td>
			<td>15000-00</td>
			<td></td>
		</tr>
	</tbody>
</table>

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.

Qualitative visualization of skeletal muscle fatty infiltration 
Properly decellularized muscles are white and semi-transparent (section 1; Figure 1). When decellularized muscles are stained with ORO to visualize IMAT (section 2), IMAT lipid droplets are apparent within the clear muscle structures as red spheres (Figure 1). Healthy mouse hindlimb muscles has little natural IMAT, evidenced by little to no red, ORO positive lipid (Figure 1A). By comparison, hindlimb muscles injected with cardiotoxin (CTX; Figure 1B) or glycerol (GLY; Figure 1C) 14 days prior to decellularization have an increased accumulation of IMAT, with a larger concentration of IMAT following CTX compared with GLY as previously noted37.
Incomplete decellularization can be identified immediately following initial SDS treatment or after washout of the ORO staining as semi-opaque, light pink fibers (Figure 2B compared with Figure 2A). Incomplete clearance of ORO can be identified following ORO washout as a pink or red uniform background, rather than distinct fiber lines (Figure 2C). Figure 2A,B also contains epimuscular adipose (asterisks), a clump of lipid droplets outside the decellularized muscle. Figure 2C also demonstrates the muscle folding that can occur if muscles are not spread out during decellularization. Incomplete decellularization, incomplete ORO clearance, and residual epimuscular adipose all artifactually increase the (OD) of extracted lipid, but do not necessarily impede qualitative assessment of fatty infiltration if they are recognized as an artifact.
Quantitative imaging of skeletal muscle fatty infiltration 
BODIPY fluorescently labeled lipid droplets can be imaged via confocal microscopy for a more detailed assessment of individual lipid droplet metrics and distribution (Figure 3). This process is semi-automated, as previously described, including Matlab code37. Good BODIPY staining results in bright elliptical shapes delineated from neighboring shapes when imaged in plane (Figure 3A). Thresholding and shape segmentation provides a good first pass for generating ROIs for each lipid droplet (Figure 3B), but manual edits are needed to correct errors. The most pervasive error is lipid droplets that are deep in the tissue and thus are not bright on any slices (Figure 3B; pink double arrows). This can be remedied by adding ROIs by hand using the Oval tool in ImageJ (Figure 3C). The second is identifying a group of lipid droplets as a single ROI (Figure 3B; red asterisk). This can be corrected by deleting and replacing the original ROI with multiple new ROIs (Figure 3D). Finally, a single lipid droplet is identified as a unique ROI in multiple slices, so the duplicate ROIs must be consolidated into a single ROI (Figure 3E; blue arrow). This is most easily done using a data processing tool such as Matlab, but could also be done by hand by identifying the largest ROI and deleting the rest. Average values for each metric in the C57BL6/J and 129Sv mouse can be found in Biltz et al.37.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/65461/65461fig01.jpg" /> 
Figure 1: Example oil red O (ORO) staining of decellularized muscles. ORO-stained muscles 14 days post-injection with saline (SAL), cardiotoxin (CTX), or glycerol (GLY). Mouse extensor digitorum longus (EDL) and tibialis anterior (TA) muscles have little IMAT (red spheres) with SAL treatment (A), but accumulate IMAT in response to CTX (B) and GLY (C) treatment. There is a complete decellularization and ORO washout, evidenced by distinct ORO positive lipid droplets in a transparent white muscle background. Scale bars = 500 &#181;m. <a href="https://jove.unicartagenaproxy.elogim.com/files/ftp_upload/65461/65461fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/65461/65461fig02.jpg" /> 
Figure 2: Examples of poor ORO staining results. Incomplete decellularization or incomplete ORO clearance leads to a semi-opaque pink/red background. Compared with the transparent white background of fully decellularized mouse diaphragm muscle (A), incomplete decellularization is characterized by light pink/red fiber tracks (B), and incomplete ORO clearance is characterized by a diffuse pink/red background (C). Scale bars: upper panels = 1 mm; lower panels = 500 &#181;m. <a href="https://jove.unicartagenaproxy.elogim.com/files/ftp_upload/65461/65461fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/65461/65461fig03.jpg" /> 
Figure 3: Examples of individual lipid droplet identification with fluorescent BODIPY staining and confocal microscopy Individual BODIPY stained lipid droplets can be identified and quantified in decellularized muscles using confocal microscopy. Individual slices through a confocal stack show lipid droplets in plane as bright green ellipses, and lipid droplets out of plane as fainter shapes (A; blue arrows). Thresholding combined with watershed object segmentation and ROI identification can map BODIPY-stained ROIs (B). Thresholding may miss some fainter lipid droplets (B; pink double arrow), requiring identification by hand (C). Watershed segmentation may group several lipid droplets together (B; red asterisk), requiring deletion of the ROI and re-estimation by hand (D). The same lipid droplet is identified in multiple slices requiring image registration (E) to delete the duplicate ROIs. Scale bars = 100 &#181;m. <a href="https://jove.unicartagenaproxy.elogim.com/files/ftp_upload/65461/65461fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Decellularization-Based Quantification of Skeletal Muscle Fatty Infiltration at JoVE.com

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.

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 ...

decellularization-based-quantification-skeletal-muscle-fatty

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2.8K Views.  Washington University in St. Louis. 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.

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

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 ...

Procedures for Rat in situ Skeletal Muscle Contractile Properties

There are many circumstances where it is desirable to obtain the contractile response of skeletal muscle under physiological circumstances: normal circulation, intact whole muscle, at body temperature. This includes the study of contractile responses like posttetanic potentiation, staircase and fatigue. Furthermore, the consequences of disease, disuse, injury, training and drug treatment can be of interest. This video demonstrates appropriate procedures to set up and use this valuable muscle preparation.
To set up this preparation, the animal must be anesthetized, and the medial gastrocnemius muscle is surgically isolated, with the origin intact. Care must be taken to maintain the blood and nerve supplies. A long section of the sciatic nerve is cleared of connective tissue, and severed proximally. All branches of the distal stump that do not innervate the medial gastrocnemius muscle are severed. The distal nerve stump is inserted into a cuff lined with stainless steel stimulating wires. The calcaneus is severed, leaving a small piece of bone still attached to the Achilles tendon. Sonometric crystals and/or electrodes for electromyography can be inserted. Immobilization by metal probes in the femur and tibia prevents movement of the muscle origin. The Achilles tendon is attached to the force transducer and the loosened skin is pulled up at the sides to form a container that is filled with warmed paraffin oil. The oil distributes heat evenly and minimizes evaporative heat loss. A heat lamp is directed on the muscle, and the muscle and rat are allowed to warm up to 37&deg;C. While it is warming, maximal voltage and optimal length can be determined. These are important initial conditions for any experiment on intact whole muscle. The experiment may include determination of standard contractile properties, like the force-frequency relationship, force-length relationship, and force-velocity relationship.
With care in surgical isolation, immobilization of the origin of the muscle and alignment of the muscle-tendon unit with the force transducer, and proper data analysis, high quality measurements can be obtained with this muscle preparation.

Procedures for Rat in situ Skeletal Muscle Contractile Properties

This video demonstrates the surgical preparation and procedures needed to study the contractile responses of the rat medial gastrocnemius muscle preparation in situ. This preparation allows measurement of skeletal muscle contractile properties under physiological conditions. The animal is anesthetized and the muscle is separated from surrounding tissue at its distal end. The Achilles tendon is attached to a force transducer, allowing measurement of the muscle&#x2019;s contractile response at 37 degrees C with an intact circulation.

There are many circumstances where it is desirable to obtain the contractile response of skeletal muscle under physiological circumstances: normal ...

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 ...

Tissue Triage and Freezing for Models of Skeletal Muscle Disease

Skeletal muscle is a unique tissue because of its structure and function, which requires specific protocols for tissue collection to obtain optimal results from functional, cellular, molecular, and pathological evaluations. Due to the subtlety of some pathological abnormalities seen in congenital muscle disorders and the potential for fixation to interfere with the recognition of these features, pathological evaluation of frozen muscle is preferable to fixed muscle when evaluating skeletal muscle for congenital muscle disease. Additionally, the potential to produce severe freezing artifacts in muscle requires specific precautions when freezing skeletal muscle for histological examination that are not commonly used when freezing other tissues. This manuscript describes a protocol for rapid freezing of skeletal muscle using isopentane (2-methylbutane) cooled with liquid nitrogen to preserve optimal skeletal muscle morphology. This procedure is also effective for freezing tissue intended for genetic or protein expression studies. Furthermore, we have integrated our freezing protocol into a broader procedure that also describes preferred methods for the short term triage of tissue for (1) single fiber functional studies and (2) myoblast cell culture, with a focus on the minimum effort necessary to collect tissue and transport it to specialized research or reference labs to complete these studies. Overall, this manuscript provides an outline of how fresh tissue can be effectively distributed for a variety of phenotypic studies and thereby provides standard operating procedures (SOPs) for pathological studies related to congenital muscle disease.

The analysis of skeletal muscle tissues to determine structural, functional, and biochemical properties is greatly facilitated by appropriate preparation. This protocol describes appropriate methods to prepare skeletal muscle tissue for a broad range of phenotyping studies.

Skeletal muscle is a unique tissue because of its structure and function, which requires specific protocols for tissue collection to obtain optimal ...

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 ...

Preparation of Rat Skeletal Muscle Homogenates for Nitrate and Nitrite Measurements

Nitrate ions (NO3-) were once thought to be inert end products of nitric oxide (NO) metabolism. However, previous studies demonstrated that nitrate ions can be converted back to NO in mammals through a two-step reduction mechanism: nitrate being reduced to nitrite (NO2-) mostly by oral commensal bacteria, then nitrite being reduced to NO by several mechanisms including via heme- or molybdenum-containing proteins. This reductive nitrate pathway contributes to enhancing NO-mediated signaling pathways, particularly in the cardiovascular system and during muscular exercise. The levels of nitrate in the body before such utilization are determined by two different sources: endogenous NO oxidation and dietary nitrate intake, principally from plants. To elucidate the complex NO cycle in physiological circumstances, we have examined further the dynamics of its metabolites, nitrate and nitrite ions, which are relatively stable compared to NO. In previous studies skeletal muscle was identified as a major storage organ for nitrate ions in mammals, as well as a direct source of NO during exercise. Therefore, establishing a reliable methodology to measure nitrate and nitrite levels in skeletal muscle is important and should be helpful in extending its application to other tissue samples. This paper explains in detail the preparation of skeletal muscle samples, using three different homogenization methods, for nitrate and nitrite measurements and discusses important issues related to homogenization processes, including the size of the samples. Nitrate and nitrite concentrations have also been compared across four different muscle groups.

We present protocols for three different methods for the homogenization of four different muscle groups of rat skeletal muscle tissue to measure and compare the levels of nitrate and nitrite. Furthermore, we compare different sample weights to investigate whether tissue sample size affects the results of homogenization.

Nitrate ions (NO3-) were once thought to be inert end products of nitric oxide (NO) metabolism. However, previous studies demonstrated that nitrate ions ...

Decellularization-Based Quantification of Skeletal Muscle Fatty Infiltration

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

Mitochondrial Isolation from Skeletal Muscle

Purification of Progenitors from Skeletal Muscle

Procedures for Rat in situ Skeletal Muscle Contractile Properties

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

Laser-inflicted Injury of Zebrafish Embryonic Skeletal Muscle

Assessment of Calcium Sparks in Intact Skeletal Muscle Fibers

Tissue Triage and Freezing for Models of Skeletal Muscle Disease

Isolation and Characterization of Exosomes from Skeletal Muscle Fibroblasts

Preparation of Rat Skeletal Muscle Homogenates for Nitrate and Nitrite Measurements