This study was supported by the Christian Doppler Research Foundation. We would like to thank Sabine Rauscher from the Core Facility Imaging at the Medical University of Vienna, Austria for support throughout the project. Primary antibodies were developed by Schiaffino, S., obtained from the Developmental Studies Hybridoma Bank, created by the NICHD of the NIH and maintained at The University of Iowa, Department of Biology, Iowa City, IA.

All procedures including animal subjects were conducted in compliance with the principles of laboratory animal care as recommended by FELASA26. Approval was obtained prior to the study by the institutional review board of the Medical University of Vienna and the Austrian Ministry for Research and Science (BMWF: Bundesministerium fuer Wissenschaft und Forschung, reference number: BMWF-66.009/0222-WF/II/3b/2014).
1. Muscle Harvest
NOTE: A previous publication by Meng et al.27 is available describing the correct freezing of muscle specimens in great detail.
<ol>
	<li>Obtain entire muscles or sufficient tissue to acquire entire cross sections immediately after euthanasia of the animals.
	<ol>
		<li>Remove the entire muscle from an anesthetized or euthanized rat. Remove all connective tissue and tendons surrounding the muscle with forceps and scissors. For anesthesia or euthanasia, follow the international guidelines of FELASA28.</li>
	</ol>
	</li>
	<li>Weigh muscle using a calibrated precision scale. This quick step allows comparative analyses between muscle samples, especially after interventional procedures.</li>
	<li>Put muscle into a vessel, filled entirely with O.C.T. compound, according to the size of the muscle with a safety margin of approximately 1 mm to all sides (e.g., made of aluminum foil).</li>
	<li>Freeze it in approximately 2 min precooled isopentane using liquid nitrogen. The isopentane must begin to show white crystals at the bottom of the container. 
	NOTE: This step is essential for preserving the correct architecture of the muscle and thereby allows the correct staining and fiber analyses27.</li>
	<li>Store samples conserved in O.C.T. for at least 24 h at -80 &#176;C. Samples can, however, be stored at -80 &#176;C for several months or even years, if correctly cooled.</li>
	<li>Cut 10 &#181;m transverse sections from the midportion of the muscle at -20 &#176;C using a cryotome. If cutting at 10 &#181;m is not feasible, increase the thickness in steps of 2 &#181;m up to a maximum of 20 &#181;m. Sufficiently sharp cutting blades are essential for this step.</li>
	<li>Apply the sections to adhesive slides by approximating the slides to the cross sections. The cross sections will automatically stick to the adhesive slides. Store the slides at -20 &#176;C overnight before staining.</li>
</ol>
2. Staining
NOTE: A large number of antibodies against myosin heavy chain proteins are available; however, high quality antibodies are essential for automated acquisition and analyses. For dilutions and reference to antibodies, see Table 1. For a spreadsheet file to calculate the correct dilutions and see the number of required solution quantities, see <a href="https://cloudfront-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/55441/Calculator_for_muscle_fiber_typing.xlsx">Supplemental File 1</a>.
<ol>
	<li>Thaw and air-dry the frozen muscle sections for 10 min before the staining procedure.</li>
	<li>Wash slides carefully with phosphate buffered saline (PBS) + 0.05% Triton X for 10 min and then for 5 min. Triton has been shown to improve the staining quality significantly. See the discussion for further details.</li>
	<li>Let sections air-dry for 2 min and outline individual cross sections on each slide using a hydrophobic pen (to minimize required amount of antibody) and allow additional drying for 15 min.</li>
	<li>Block all slides with PBS + 0.05% Triton X (PBST) containing 10% goat serum for 1 h at room temperature.</li>
	<li>Dilute all primary antibodies (Table 1) in a cocktail of PBS + 0.05% Triton X (PBST) containing 10% goat serum. Mix sufficiently before application. Triton X is essential for equal staining of the whole cross sections. Calculate approximately 30 &#181;L of primary antibody cocktail per cross section.</li>
	<li>For steps 2.7-2.13 ensure sufficient protection from light. Dimmed room lights and covered staining trays provide adequate protection. Additionally, fill the staining tray with fluid at the bottom to provide a humid environment during staining and prevent drying.</li>
	<li>Apply primary antibody (Table 1) cocktails to the slides for 1 h in a humid staining tray on room temperature.</li>
	<li>Wash slides with PBS + 0.05% Triton X for 10 min and then with new PBST for 5 min.</li>
	<li>Dilute secondary antibodies in a cocktail of PBS + 0.05% Triton X (PBST) + 10% goat serum. Calculate 30 &#181;L of primary antibody cocktail per cross section.</li>
	<li>Apply secondary antibody cocktail to the slides for 1 h.</li>
	<li>Wash slides with PBS + 0.05% Triton X (PBST) for 10 min and with new PBST for another 5 min.</li>
	<li>Apply a DAPI nuclear staining kit (preferably read-to-use solutions) for 15 min or according to the manufacturer&#39;s instructions to stain cell nuclei.</li>
	<li>Wash slides briefly with PBS + 0.05% Triton X (PBST) for 1 min. Afterward, let slides air-dry briefly.</li>
	<li>Apply fluorescent mounting medium and coverslips. Store at 4 &#176;C protected from light and perform imaging optimally within 24-48 h.</li>
</ol>
3. Microscopy
<ol>
	<li>Load slides into slide scanner. A maximum of 12 slides is feasible depending on the slide scanner.</li>
	<li>Start a new project. For preview set the DAPI channel and the 2.5X lens. For the detailed analysis use DAPI, GFP, FITC and Cy5 channels and the 20X objective. Autofocus is acquired throughout the project in the DAPI channel.</li>
	<li>Use the following colors for each channel: DAPI -blue, FITC: green, Texas Red: red, Cy5: yellow.</li>
	<li>Create previews of each slide using the DAPI channel. For this purpose, apply manual focus on the first specimen and use it throughout the preview analysis to acquire preview images of every specimen.</li>
	<li>Outline each cross-section that should be included for the detailed acquisition process. Do not draw outlines close to the edges of the specimen, to ensure caption of the entire area of interest.</li>
	<li>Set exposure times for each channel. In most cases, exposure parameters of the different channels are identical for all cross-sections.</li>
	<li>Run automatic image acquisition. Check correct acquisition for the first field of views, to prevent incorrect acquisition early in the automated process.</li>
	<li>If required, establish remote control using desktop mirroring software. This allows the remote monitoring of the image acquisition process by mirroring the computer screen of the slide scanner to a different computer via any network or internet connection. Although, no changes can be made to the physical setup regarding the used slides, the virtual environment of the image acquisition software allows monitoring the correct focus or exposure times of the images. In addition, the estimated time for completion can be checked regularly.</li>
	<li>After completion of the image acquisition, control the correct autofocus of all images by manually controlling if the fiber architecture, individual fibers and the various fiber types are distinguishable.</li>
	<li>Re-acquire incorrectly focused image areas for single fields of view or entire areas of interest. For reacquisition of single areas, mark areas or images throughout the entire project and reacquire areas with manual focus. In most cases, an additional image detail in a different focus level leads to the incorrectly focused images</li>
	<li>For every cross section that should be included in the final analyses, export each channel (excluding DAPI) separately as jpeg files. Name and sort files according to the exposed channels in folders named Texas Red, FITC and Cy5.</li>
</ol>
4. Automated Fiber Analysis
NOTE: The macro can be obtained from the following web page: https://www.meduniwien.ac.at/hp/bionicreconstruction/macro/
<ol>
	<li>Preview each image using a conventional image editing software before analyses and check for sufficient contrast between stained fibers and background. If necessary, adjust contrast and brightness to increase the difference, using brightness and contrast adjustment commands.</li>
	<li>Open ImageJ or Fiji. Open the macro using the command &#34;Plugins - Macros - Edit.&#34; This shows the macros source code and allows quick adaption of parameters. If necessary, change folder directory for each channel in the source code of the macro. 
	NOTE: Values for muscle fiber analyses are based on rat biceps and lumbrical muscles, other species or muscles may require different values.</li>
	<li>Use the &#34;run&#34; command to start the macro. All images of the folder are now loaded into the macro and quantified in a consecutive order. The results are shown in a new window. This step may take from seconds to several hours, depending on the amount of data being analyzed.</li>
	<li>Export the results window as a spreadsheet file. Identify values for positively counted Cy5 (slow), FITC (intermediate) and Texas Red (fast) fibers and sum up for total fiber number.</li>
</ol>

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The authors have nothing to disclose.

Here, we demonstrate a widely accessible methodology to study and automatically quantify the muscle fiber populations of rat cross sections via immunohistochemistry in a time efficient manner. For reproducibility, we present a detailed step by step description and potential modifications for applications in other species not described in this study. Furthermore, we discuss the advantages of the procedure, prerequisites for optimal function and its limitations.
Currently, a number of staining methods exist to identify muscle fiber populations and a number of methods have been described for manual or semi-automatic quantification of muscle fibers17. However, no protocols exist that combine both and describe a standardized approach to reliably stain muscle fiber populations and automatically acquire and quantify muscle fibers, using freely available analyses software. Based on previous publications24,29, we optimized existing myosin heavy chain immunohistochemistry staining procedures for automated analyses of fiber populations in murine muscle cross sections. Thereby, we recently showed that the automated analysis of whole cross sections is feasible, reliable compared to manual analyses, and more time-efficient27. In addition, the method is user-independent thus reducing human error and thereby providing an objective comparison as for example between different treatment groups.
The whole protocol is based on providing detailed and high-quality images of whole cross sections for the automated quantification process. ImageJ as an open source software provides a number of powerful yet simple commands, which arranged in a macro allow muscle fiber identification. However, quality differences within or between cross section need to be minimized to achieve reliable results. We have identified some critical steps that are essential to achieving the required quality. This includes the use of optimal primary antibodies. Similarly, the use of appropriate secondary antibodies providing distinct fluorescent signals was essential, and we doubled concentrations for improved signal quality. Additionally, we observed a higher and more even staining quality with the use of Triton X in the antibody cocktails. Importantly, this was not observed with the use of Tween instead. Images with suboptimal staining quality were manually adjusted for better contrast to allow automatic image acquisition. Most important for the acquisition process was the correct function of the autofocus system. For this purpose, we used a DAPI-kit to stain the cell nuclei of satellite cells, which provided a reliable landmark for the autofocus system.
For the ImageJ quantification process, optimal values were determined by comparing manual to automated results and consecutive fine-tuning of the parameters25: Threshold and Analyze particles. The resulting values were sufficient for biceps and lumbrical muscles and therefore potentially other rat muscles. Additionally, Bloemberg et al.24 have shown that the staining is applicable for other species such as humans or mice. Therefore, the entire protocol can theoretically be adapted for the use in other species, but would, however, require identifying the necessary parameters. The relevant commands are highlighted in the source code.
Further possible modifications of the procedure include the setup of the slide scanner. In our setup, we used a slide scanner system, with a 2.5X lens for the preview and 20X lens for the actual acquisition. Whereas changing the lens for the preview process will provide no significant benefit in terms of quality or speed, the acquisition can also be conducted using a 10X lens to increase the speed (and consequently decrease the resolution) or instead a 40X lens for increasing the resolution and consequently acquisition time. As is shown in Figure 2, increasing the resolution is most likely not necessary for rat muscles but may be considered for other species. Other modifications that are suitable in this setup are staining for MHC-IIX fibers or laminin to visualize the muscle&#39;s extracellular matrix. These modifications, however, require changing the acquisition setup for two additional reflectors (for example Cy7) and modifying the ImageJ macro to include additional analyses with the correct parameters to the script code.
A limitation of this technique is that the absolute fiber counts were significantly higher compared to manual counts in a previous analysis, although relative counts remained similar25. This is most likely the result of inadequate separation and thus repeated quantification of fibers. This effect is more present in fibers that are inadequately stained or show other types of artifacts. However, the number of relative fiber counts and thus fiber populations remained highly accurate. In comparison to the common practice of quantifying a defined sub-portion of a cross section, e.g. 40%, we believe that analyses of the entire cross section by the ImageJ are more accurate as a result of the heterogenic muscle fiber architecture (Figure 2). A potential limitation of the procedure is the requirement to use expensive slide scanners for automated acquisition of entire cross section, which are not generally available. An alternative approach is to acquire single field of views and quantify each individually by the macro.
In conclusion, we believe this protocol is widely accessible and allows the fast and reliable quantification of whole muscle cross sections using freely available analyses software. Therefore, the introduced procedure provides a useful contribution to the analyses of muscle fiber morphology.

Skeletal muscle composition undergoes profound changes during physiological processes such as aging1,2, exercise3,4,5,6,7, or pathophysiological processes such as disease8,9,10 or trauma11. Hence, several fields of research concentrate on the structural effects of these processes to understand functional changes. One of the key aspects determining muscle function is the composition of muscle fibers. Muscle fibers express different myosin heavy chain (MHC) proteins and are thereby classified into slow, intermediate, or fast fibers7,12,13,14,15,16,17. Physiologically, muscles have different muscle fiber compositions depending on their function in the body. Using muscle fiber typing, fiber populations can be quantified to identify adaption to physiological or pathophysiological processes7,17. Historically, a number of time-consuming methods have been applied to differentiate between muscle fiber types. For this purpose, muscle fibers were classified either by reactivity of myosin ATPase at various pH levels or muscle enzyme activity. As different fiber qualities could not be assessed in a single section, multiple cross sections were required to identify all muscle fibers and allow manual quantification14,16,17,18,19,20,21,22. In contrast, recent publications used immunohistochemistry (IHC) against myosin heavy chain protein to rapidly stain multiple fiber types in a single cross sections. Based on the advantages of this procedure, it is now considered the gold standard in muscle fiber population analysis19,23,24. Using improved IHC staining protocols, we were recently able to show that the fully automatic acquisition of whole muscle cross sections and subsequent automatic muscle fiber quantification is feasible using the open source platform ImageJ. Compared to manual quantification, our procedure provided a significant decrease in time (approximately 10% of manual analyses) required per slide while being accurate to &#177;4%25.
The overall goal of this method is to describe a rapid, reliable, user-independent guide to automatic muscle fiber quantification in whole rat muscles using an open source platform. In addition, we describe potential modifications that would permit its use for other specimens such as mice or human muscles.

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	<tbody>
		<tr height="21">
			<td height="21" style="height:21px;width:303px;">O.C.T compound</td>
			<td style="width:397px;">Tissue-Tek, Sakura, Netherlands</td>
			<td style="width:119px;"></td>
			<td style="width:167px;">For embedding of muscle tissue</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Isopentane</td>
			<td></td>
			<td></td>
			<td>for adequate freezing of muscle tissue</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Superfrost Ultra Plus slides</td>
			<td>Thermo Scientific, Germany</td>
			<td align="right">1014356190</td>
			<td>adhesive slides</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">phosphate buffered saline&#160;</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Triton X-100</td>
			<td>Thermo Scientific, Germany</td>
			<td align="right">85112</td>
			<td>Detergent Soluation</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Goat serum</td>
			<td>Thermo Scientific, Germany</td>
			<td>50197Z</td>
			<td>Goat Serum</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">DAKO Fluorescent Mounting Medium</td>
			<td>Dako Denmark</td>
			<td>S3023</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Dako pen</td>
			<td>Dako Denmark</td>
			<td>S200230-2</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">TissueFAXSi plus&#160;</td>
			<td>TissueGnostics, Vienna, Austria</td>
			<td></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;"></td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Primary antibodies</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">MHC-I (Cat# BA-F8, RRID: AB_10572253)</td>
			<td>Developmental Studies Hybridoma Bank (DSHB, Iowa, USA)</td>
			<td></td>
			<td>Supernatant</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">MHC-IIa (Cat# SC-71, RRID: AB_2147165)</td>
			<td>Developmental Studies Hybridoma Bank (DSHB, Iowa, USA)</td>
			<td></td>
			<td>Supernatant</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">MHC-IIb (Cat# BF-F3, RRID: AB_2266724)</td>
			<td>Developmental Studies Hybridoma Bank (DSHB, Iowa, USA)</td>
			<td></td>
			<td>Supernatant</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Secondary antibodies</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Alexa&#160;Fluor&#160;633&#160;Goat Anti-Mouse IgG2b&#160;</td>
			<td>Thermo Scientific, Germany</td>
			<td>A-21146</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Alexa Fluor 488 Goat Anti-Mouse IgG1&#160;(&#947;1)</td>
			<td>Thermo Scientific, Germany</td>
			<td>A-21121</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Alexa Fluor 555 Goat Anti-Mouse IgM (&#181; chain),</td>
			<td>Thermo Scientific, Germany</td>
			<td>A-21426</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;"></td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">NucBlue Fixed Cell ReadyProbes Reagent</td>
			<td>Thermo Scientific, Germany</td>
			<td>R37606</td>
			<td></td>
		</tr>
	</tbody>
</table>

a rapid automated protocol for muscle fiber population analysis in rat muscle cross sections using myosin heavy chain immunohistochemistry

Quantification of muscle fiber populations provides a deeper insight into the effects of disease, trauma, and various other influences on skeletal muscle composition. Various time-consuming methods have traditionally been used to study fiber populations in many fields of research. However, recently developed immunohistochemical methods based on myosin heavy chain protein expression provide a quick alternative to identify multiple fiber types in a single section. Here, we present a rapid, reliable and reproducible protocol for improved staining quality, allowing automatic acquisition of whole cross sections and automatic quantification of fiber populations with ImageJ. For this purpose, embedded skeletal muscles are cut in cross sections, stained using myosin heavy chains antibodies with secondary fluorescent antibodies and DAPI for cell nuclei staining. Whole cross sections are then scanned automatically using a slide scanner to obtain high-resolution composite pictures of the entire specimen. Fiber population analyses are subsequently performed to quantify slow, intermediate and fast fibers using an automated macro for ImageJ. We have previously shown that this method can identify fiber populations reliably to a degree of &#177;4%. In addition, this method reduces inter-user variability and time per analyses significantly using the open source platform ImageJ.

Whole rat muscle cross sections were stained rapidly using immunohistochemistry to identify MHC I, IIA and IIB muscle fibers. Using a fluorescent microscope slide scanner, entire cross sections were then automatically acquired for automated muscle fiber analyses with ImageJ. The concept of the procedure is based on providing a simple, reliable and time-efficient workflow for quantification of muscle fibers.
The procedure&#39;s workflow (Figure 1) starts with the correct removal and freezing of muscle samples, as previously described in a previous tutorial by Meng et al.27. This is essential for adequate staining quality. In the next step, the muscle samples are cut and stained using primary antibodies against myosin heavy chain proteins class I, IIA and IIB and appropriate secondary fluorescent antibodies. For optimal staining quality, the manufacturer&#39;s recommended concentrations for the secondary antibodies were doubled and Triton X used for the antibody cocktails to provide even staining quality. The immunofluorescent staining of muscle fibers is quick with minor cross-reactivity and can be accomplished in approximately 5 h for batches of up to 26 slides in our setup. Resulting images showed high contrast between positive fibers and surrounding tissue and a strong distinction between different fiber types (Figure 2). Next, image acquisition is performed using a slide scanner that permits automatic scanning of entire muscle cross sections. Here, up to 12 slides are scanned automatically, for example, overnight. Hereby, overview images of the cross sections are created in high detail with a resolution that permits inspection of single fiber morphology (Figure 2).
In the final step, individual channels of each cross section are exported for quantification of muscle fiber populations using a specifically designed macro for ImageJ. The automatic analysis takes approximately 5-10 min for a rat biceps cross section containing between 3,000-6,000 fibers, whereas the manual analyses took approximately 60 min25. Automatic analyses depend strongly on computing power of the workstation and file size. As previously shown, the macro is able to detect relative muscle fiber populations with an accuracy of &#177; 4% compared to manual analysis (Figure 3). Overall absolute counts are higher compared to manual analyses, however, relative counts remained similar within a &#177;4% range, as previously shown.
<img alt="Figure 1" src="/files/ftp_upload/55441/55441fig1.jpg" /> 
Figure 1: Workflow of the Muscle Fiber Quantification Protocol. First, muscle samples are obtained from the animal and frozen correctly (see JoVE video by Meng et al.27). Then cut muscle cross sections and conduct immune staining. Acquire images of stained cross sections using a slide scanner for detailed whole cross sections images. Run the quantification macro for ImageJ for muscle fiber population analyses. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/55441/55441fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" src="/files/ftp_upload/55441/55441fig2.jpg" /> 
Figure 2: Rat Biceps Muscle Cross Section. Overview of a stained rat biceps muscle cross section: The cross section shows slow MHC I fibers in yellow, intermediate MHC IIA fibers in green and fast MHC IIB fibers in red. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/55441/55441fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" src="/files/ftp_upload/55441/55441fig3.jpg" /> 
Figure 3: Muscle Fiber Analyses Results of Rat Cross Section in Figure 2. Analyses of muscle MHC I, MHC IIA and MHC IIB fibers. Top: Absolute counts of muscle fibers by manual or automatic count. Middle: Relative portions of muscle fibers populations after manual quantification. Bottom: Relative portions of muscle fibers populations after absolute quantification. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/55441/55441fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Table 1" src="/files/ftp_upload/55441/55441table1.jpg" /> 
Table 1: Immunohistochemical Staining of MHC Fibers. Description of primary and secondary antibodies including dilutions used for the staining procedure. (Modified from Bergmeister et al.25)

Watch this Scientific Journal Video about A Rapid Automated Protocol for Muscle Fiber Population Analysis in Rat Muscle Cross Sections Using Myosin Heavy Chain Immunohistochemistry at JoVE.com

A Rapid Automated Protocol for Muscle Fiber Population Analysis in Rat Muscle Cross Sections Using Myosin Heavy Chain Immunohistochemistry

Here, we present a protocol for rapid muscle fiber analyses, which allows improved staining quality, and thereby automatic acquisition and quantification of fiber populations using the freely available software ImageJ.

Quantification of muscle fiber populations provides a deeper insight into the effects of disease, trauma, and various other influences on skeletal muscle ...

a-rapid-automated-protocol-for-muscle-fiber-population-analysis-rat

Nanyang Technological University

Research

JoVE Journal

Biology

11.2K Views.  Medical University of Vienna. Here, we present a protocol for rapid muscle fiber analyses, which allows improved staining quality, and thereby automatic acquisition and quantification of fiber populations using the freely available software ImageJ.

Video: A Rapid Automated Protocol for Muscle Fiber Population Analysis in Rat Muscle Cross Sections Using Myosin Heavy Chain Immunohistochemistry

Immunohistochemistry on Paraffin Sections of Mouse Epidermis Using Fluorescent Antibodies

In the epidermis, immunohistochemistry is an efficient means of localizing specific proteins to their relative expression compartment; namely the basal, suprabasal, and stratum corneum layers. The precise localization within the epidermis of a particular protein lends clues toward its functional role within the epidermis. In this chapter, we describe a reliable method for immunolocalization within the epidermis modified for both frozen and paraffin sections that we use very routinely in our laboratory. Paraffin sections generally provide much better morphology, hence, superior results and photographs; however, not all antibodies will work with the harsh fixation and treatment involved in their processing. Therefore, the protocol for frozen sectioning is also included. Within paraffin sectioning, two fixation protocols are described (Bouin's and paraformaldehyde); the choice of fixative will be directly related to the antibody specifications and may require another fixing method.

We describe a reliable method for immunolocalization within the epidermis modified for both frozen and paraffin sections that we use very routinely in our laboratory.

In the epidermis, immunohistochemistry is an efficient means of localizing specific proteins to their relative expression compartment; namely the basal, ...

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 ...

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 ...

Analysis of Skeletal Muscle Defects in Larval Zebrafish by Birefringence and Touch-evoke Escape Response Assays

Zebrafish (Danio rerio) have become a particularly effective tool for modeling human diseases affecting skeletal muscle, including muscular dystrophies1-3, congenital myopathies4,5, and disruptions in sarcomeric assembly6,7, due to high genomic and structural conservation with mammals8. Muscular disorganization and locomotive impairment can be quickly assessed in the zebrafish over the first few days post-fertilization. Two assays to help characterize skeletal muscle defects in zebrafish are birefringence (structural) and touch-evoked escape response (behavioral).
Birefringence is a physical property in which light is rotated as it passes through ordered matter, such as the pseudo-crystalline array of muscle sarcomeres9. It is a simple, noninvasive approach to assess muscle integrity in translucent zebrafish larvae early in development. Wild-type zebrafish with highly organized skeletal muscle appear very bright amidst a dark background when visualized between two polarized light filters, whereas muscle mutants have birefringence patterns specific to the primary muscular disorder they model. Zebrafish modeling muscular dystrophies, diseases characterized by myofiber degeneration followed by repeated rounds of regeneration, exhibit degenerative dark patches in skeletal muscle under polarized light. Nondystrophic myopathies are not associated with necrosis or regenerative changes, but result in disorganized myofibers and skeletal muscle weakness. Myopathic zebrafish typically show an overall reduction in birefringence, reflecting the disorganization of sarcomeres.
The touch-evoked escape assay involves observing an embryo&#39;s swimming behavior in response to tactile stimulation10-12. In comparison to wild-type larvae, mutant larvae frequently display a weak escape contraction, followed by slow swimming or other type of impaired motion that fails to propel the larvae more than a short distance12. The advantage of these assays is that disease progression in the same fish type can be monitored in vivo for several days, and that large numbers of fish can be analyzed in a short time relative to higher vertebrates.

The zebrafish is now an established and powerful tool for modeling muscular dystrophies, congenital myopathies, and related neuromuscular diseases. Birefringence and touch-evoked escape behavior are two common noninvasive assays used to determine the degree of muscular disorganization and locomotive impairment of zebrafish embryos during early development.

Zebrafish (Danio rerio) have become a particularly effective tool for modeling human diseases affecting skeletal muscle, including muscular ...

Optimized Protocol for Retinal Wholemount Preparation for Imaging and Immunohistochemistry

Working with delicate tissue can be a complicating factor when performing immunohistochemical assessment. Here, we present a method that utilizes a ring-supported hydrophilized PTFE membrane to provide structural support to both living and fixed tissue during immunohistochemical processing, which allows for the use of a variety of protocols that would otherwise cause damage to the tissue. First, this is demonstrated with bolus loading of fluorescent markers into living retinal tissue. This method allows for quick visualization of targeted structures, while the membrane support maintains tissue integrity during the injection and allows for easy transfer of the preparation for further imaging or processing.
Second, a procedure for antibody staining in tissue fixed with carbodiimide is described. Though paraformaldehyde fixation is more common, carbodiimide fixation provides a superior substrate for the visualization of synaptic proteins. A limitation of carbodiimide is that the resulting fixed tissue is relatively fragile; however, this is overcome with the use of the supporting membrane. Retinal tissue is used to demonstrate these techniques, but they may be applied to any fragile tissue.

The protocol described here is for structural assessment of a wholemount retinal preparation. This includes descriptions of tissue dissection, mounting onto a hydrophilized polytetrafluoroethylene (PTFE) membrane insert, bolus loading with fluorescent markers, and a comparison of fixation with carbodiimide and paraformaldehyde for immunohistochemical analysis of cellular and synaptic components.

Working with delicate tissue can be a complicating factor when performing immunohistochemical assessment. Here, we present a method that utilizes a ...

Generation of Myospheres From hESCs by Epigenetic Reprogramming

Generation of a homogeneous and abundant population of skeletal muscle cells from human embryonic stem cells (hESCs) is a requirement for cell-based therapies and for a "disease in a dish" model of human neuromuscular diseases. Major hurdles, such as low abundance and heterogeneity of the population of interest, as well as a lack of protocols for the formation of three-dimensional contractile structures, have limited the applications of stem cells for neuromuscular disorders. We have designed a protocol that overcomes these limits by ectopic introduction of defined factors in hESCs - the muscle determination factor MyoD and SWI/SNF chromatin remodeling complex component BAF60C - that are able to reprogram hESCs into skeletal muscle cells. Here we describe the protocol established to generate hESC-derived myoblasts and promote their clustering into tridimensional miniaturized structures (myospheres) that functionally mimic miniaturized skeletal muscles7.

Here, we describe a protocol based on epigenetic reprogramming of human embryonic stem cells (hESCs) toward generating a homogeneous population of skeletal muscle progenitors that under permissive culture conditions form three-dimensional clusters of contractile myofibers (myospheres), which recapitulate biological features of human skeletal muscles.

Generation of a homogeneous and abundant population of skeletal muscle cells from human embryonic stem cells (hESCs) is a requirement for cell-based ...

Visualization and Quantification of the Cell-free Layer in Arterioles of the Rat Cremaster Muscle

The cell-free layer is defined as the parietal plasma layer in the microvessel flow, which is devoid of red blood cells. The measurement of the in vivo cell-free layer width and its spatiotemporal variations can provide a comprehensive understanding of hemodynamics in microcirculation. In this study, we used an intravital microscopic system coupled with a high-speed video camera to quantify the cell-free layer widths in arterioles in vivo. The cremaster muscle of Sprague-Dawley rats was surgically exteriorized to visualize the blood flow. A custom-built imaging script was also developed to automate the image processing and analysis of the cell-free layer width. This approach enables the quantification of spatiotemporal variations more consistently than previous manual measurements. The accuracy of the measurement, however, partly depends on the use of a blue filter and the selection of an appropriate thresholding algorithm. Specifically, we evaluated the contrast and quality of images acquired with and without the use of a blue filter. In addition, we compared five different image histogram-based thresholding algorithms (Otsu, minimum, intermode, iterative selection, and fuzzy entropic thresholding) and illustrated the differences in their determination of the cell-free layer width.

This study demonstrates the surgical preparation of the rat cremaster muscle for the visualization of the in vivo cell-free layer. Considerable factors affecting the accuracy of the cell-free layer width measurement are discussed in this study.

The cell-free layer is defined as the parietal plasma layer in the microvessel flow, which is devoid of red blood cells. The measurement of the in vivo ...

Optical Cross-Sectional Muscle Area Determination of Drosophila Melanogaster Adult Indirect Flight Muscles

Muscle mass wasting, known as muscle atrophy, is a common phenotype in Drosophila models of neuromuscular diseases. We have used the indirect flight muscles (IFMs) of flies, specifically the dorso-longitudinal muscles (DLM), as the experimental subject to measure the atrophic phenotype brought about by different genetic causes. In this protocol, we describe how to embed fly thorax muscles for semi thin sectioning, how to obtain a good contrast between muscle and the surrounding tissue, and how to process optical microscope images for semiautomatic acquisition of quantifiable data and analysis. We describe three specific applications of the methodological pipeline. First, we show how the method can be applied to quantify muscle degeneration in a myotonic dystrophy fly model; second, measurement of muscle cross-sectional area can help to identify genes that either promote or prevent muscle atrophy and/or muscle degeneration; third, this protocol can be applied to determine whether a candidate compound is able to significantly modify a given atrophic phenotype induced by a disease-causing mutation or by an environmental trigger.

Optical Cross-Sectional Muscle Area Determination of Drosophila Melanogaster Adult Indirect Flight Muscles

We report a method to quantify muscle area, which is an indirect method to determine muscle mass in Drosophila adults. We demonstrate the application of our methodology by analyzing the indirect flight muscles in a Drosophila model of Myotonic Dystrophy disease.

Muscle mass wasting, known as muscle atrophy, is a common phenotype in Drosophila models of neuromuscular diseases. We have used the indirect flight ...

Sequential Immunofluorescence and Immunohistochemistry on Cryosectioned Zebrafish Embryos

Investigation of intercellular interactions often requires discrete labeling of specific cell populations and precise protein localization. The zebrafish embryo is an excellent tool for examining such interactions with an in vivo model. Whole-mount immunohistochemical and immunofluorescence assays are frequently applied in zebrafish embryos to assess protein expression. However, it can be difficult to achieve accurate mapping of co-localized proteins in three-dimensional space. In addition, some studies may require the use of two antibodies that are not compatible with the same technique (e.g., antibody 1 is only suitable for immunohistochemistry and antibody 2 is only suitable for immunofluorescence). The purpose of the method described herein is to perform sequential immunofluorescence and/or immunohistochemistry on individual cryosections derived from early-stage zebrafish embryos. Here we describe the use of sequential rounds of immunofluorescence, imaging, immunohistochemistry, imaging for a single cryosection in order to achieve precise identification of protein expression at the single-cell level. This methodology is suitable for any study in early-stage zebrafish embryos that requires accurate identification of multiple protein targets in individual cells.

This protocol demonstrates sequential immunofluorescence and immunohistochemistry on cryosections from early-stage zebrafish embryos enabling precise colocalization analyses in specific cell populations.

Investigation of intercellular interactions often requires discrete labeling of specific cell populations and precise protein localization. The zebrafish ...

Isolation of Human Ventricular Cardiomyocytes from Vibratome-Cut Myocardial Slices

The isolation of ventricular cardiac myocytes from animal and human hearts is a fundamental method in cardiac research. Animal cardiomyocytes are commonly isolated by coronary perfusion with digestive enzymes. However, isolating human cardiomyocytes is challenging because human myocardial specimens usually do not allow for coronary perfusion, and alternative isolation protocols result in poor yields of viable cells. In addition, human myocardial specimens are rare and only regularly available at institutions with on-site cardiac surgery. This hampers the translation of findings from animal to human cardiomyocytes. Described here is a reliable protocol that enables efficient isolation of ventricular myocytes from human and animal myocardium. To increase the surface-to-volume ratio while minimizing cell damage, myocardial tissue slices 300 &#181;m thick are generated from myocardial specimens with a vibratome. Tissue slices are then digested with protease and collagenase. Rat myocardium was used to establish the protocol and quantify yields of viable, calcium-tolerant myocytes by flow-cytometric cell counting. Comparison with the commonly used tissue-chunk method showed significantly higher yields of rod-shaped cardiomyocytes (41.5 &#177; 11.9 vs. 7.89 &#177; 3.6%, p &#60; 0.05). The protocol was translated to failing and non-failing human myocardium, where yields were similar as in rat myocardium and, again, markedly higher than with the tissue-chunk method (45.0 &#177; 15.0 vs. 6.87 &#177; 5.23 cells/mg, p &#60; 0.05). Notably, with the protocol presented it is possible to isolate reasonable numbers of viable human cardiomyocytes (9&#8211;200 cells/mg) from minimal amounts of tissue (&#60;50 mg). Thus, the method is applicable to healthy and failing myocardium from both human and animal hearts. Furthermore, it is possible to isolate excitable and contractile myocytes from human tissue specimens stored for up to 36 h in cold cardioplegic solution, rendering the method particularly useful for laboratories at institutions without on-site cardiac surgery.

Presented is a protocol for the isolation of human and animal ventricular cardiomyocytes from vibratome-cut myocardial slices. High yields of calcium-tolerant cells (up to 200 cells/mg) can be obtained from small amounts of tissue (&#60;50 mg). The protocol is applicable to myocardium exposed to cold ischemia for up to 36 h.

The isolation of ventricular cardiac myocytes from animal and human hearts is a fundamental method in cardiac research. Animal cardiomyocytes are commonly ...