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

Summary

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.

Abstract

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 ±4%. In addition, this method reduces inter-user variability and time per analyses significantly using the open source platform ImageJ.

Introduction

Skeletal muscle composition undergoes profound changes during physiological processes such as aging^1,2, exercise^3,4,5,6,7, or pathophysiological processes such as disease^8,9,10 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 fibers^{7,12,13,14,15,16,17}. 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 processes^7,17. 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 quantification^{14,16,17,18,19,20,21,22}. 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 analysis^19,23,24. 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 ±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.

Protocol

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.

1. Obtain entire muscles or sufficient tissue to acquire entire cross sections immediately after euthanasia of the animals.
   1. 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.
2. Weigh muscle using a calibrated precision scale. This quick step allows comparative analyses between muscle samples, especially after interventional procedures.
3. 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).
4. 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.
5. Store samples conserved in O.C.T. for at least 24 h at -80 °C. Samples can, however, be stored at -80 °C for several months or even years, if correctly cooled.
6. Cut 10 µm transverse sections from the midportion of the muscle at -20 °C using a cryotome. If cutting at 10 µm is not feasible, increase the thickness in steps of 2 µm up to a maximum of 20 µm. Sufficiently sharp cutting blades are essential for this step.
7. 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 °C overnight before staining.

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 Supplemental File 1.

1. Thaw and air-dry the frozen muscle sections for 10 min before the staining procedure.
2. 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.
3. 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.
4. Block all slides with PBS + 0.05% Triton X (PBST) containing 10% goat serum for 1 h at room temperature.
5. 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 µL of primary antibody cocktail per cross section.
6. 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.
7. Apply primary antibody (Table 1) cocktails to the slides for 1 h in a humid staining tray on room temperature.
8. Wash slides with PBS + 0.05% Triton X for 10 min and then with new PBST for 5 min.
9. Dilute secondary antibodies in a cocktail of PBS + 0.05% Triton X (PBST) + 10% goat serum. Calculate 30 µL of primary antibody cocktail per cross section.
10. Apply secondary antibody cocktail to the slides for 1 h.
11. Wash slides with PBS + 0.05% Triton X (PBST) for 10 min and with new PBST for another 5 min.
12. Apply a DAPI nuclear staining kit (preferably read-to-use solutions) for 15 min or according to the manufacturer's instructions to stain cell nuclei.
13. Wash slides briefly with PBS + 0.05% Triton X (PBST) for 1 min. Afterward, let slides air-dry briefly.
14. Apply fluorescent mounting medium and coverslips. Store at 4 °C protected from light and perform imaging optimally within 24-48 h.

3. Microscopy

1. Load slides into slide scanner. A maximum of 12 slides is feasible depending on the slide scanner.
2. 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.
3. Use the following colors for each channel: DAPI -blue, FITC: green, Texas Red: red, Cy5: yellow.
4. 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.
5. 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.
6. Set exposure times for each channel. In most cases, exposure parameters of the different channels are identical for all cross-sections.
7. Run automatic image acquisition. Check correct acquisition for the first field of views, to prevent incorrect acquisition early in the automated process.
8. 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.
9. 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.
10. 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
11. 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.

4. Automated Fiber Analysis

NOTE: The macro can be obtained from the following web page: https://www.meduniwien.ac.at/hp/bionicreconstruction/macro/

1. 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.
2. Open ImageJ or Fiji. Open the macro using the command "Plugins - Macros - Edit." 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.
3. Use the "run" 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.
4. 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.

Results

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's w...

Discussion

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 m...

Materials

Name	Company	Catalog Number	Comments
O.C.T compound	Tissue-Tek, Sakura, Netherlands		For embedding of muscle tissue
Isopentane			for adequate freezing of muscle tissue
Superfrost Ultra Plus slides	Thermo Scientific, Germany	1014356190	adhesive slides
phosphate buffered saline
Triton X-100	Thermo Scientific, Germany	85112	Detergent Soluation
Goat serum	Thermo Scientific, Germany	50197Z	Goat Serum
DAKO Fluorescent Mounting Medium	Dako Denmark	S3023
Dako pen	Dako Denmark	S200230-2
TissueFAXSi plus	TissueGnostics, Vienna, Austria
Primary antibodies
MHC-I (Cat# BA-F8, RRID: AB_10572253)	Developmental Studies Hybridoma Bank (DSHB, Iowa, USA)		Supernatant
MHC-IIa (Cat# SC-71, RRID: AB_2147165)	Developmental Studies Hybridoma Bank (DSHB, Iowa, USA)		Supernatant
MHC-IIb (Cat# BF-F3, RRID: AB_2266724)	Developmental Studies Hybridoma Bank (DSHB, Iowa, USA)		Supernatant
Secondary antibodies
Alexa Fluor 633 Goat Anti-Mouse IgG2b	Thermo Scientific, Germany	A-21146
Alexa Fluor 488 Goat Anti-Mouse IgG1 (γ1)	Thermo Scientific, Germany	A-21121
Alexa Fluor 555 Goat Anti-Mouse IgM (µ chain)	Thermo Scientific, Germany	A-21426
NucBlue Fixed Cell ReadyProbes Reagent	Thermo Scientific, Germany	R37606